Impact of Silver-Containing Wound Dressings on Bacterial Biofilm Viability and Susceptibility to Antibiotics during Prolonged Treatment

Transcription

1 ANTIMICROBIAL AGENTS AND CHEMOTHERAPY, Dec. 2010, p Vol. 54, No /10/$12.00 doi: /aac Copyright 2010, American Society for Microbiology. All Rights Reserved. Impact of Silver-Containing Wound Dressings on Bacterial Biofilm Viability and Susceptibility to Antibiotics during Prolonged Treatment Victoria Kostenko, 1,2 Jeffrey Lyczak, 3 Katherine Turner, 4 and Robert John Martinuzzi 2 * Calgary Center for Innovative Technology, University of Calgary, Calgary, Alberta T2N 1N4, Canada 1 ; Biomedical Engineering, University of Calgary, Calgary, Alberta T2N 1N4, Canada 2 ; Shire Human Genetic Therapies, 700 Main Street Cambridge, Massachusetts ; and Attogen Inc., 100 Barber Avenue, Worcester, Massachusetts Received 16 June 2010/Returned for modification 14 August 2010/Accepted 9 September 2010 The long-term antimicrobial efficacy of silver dressings against bacterial biofilms was investigated in a 7-day treatment in vitro model where the protein-rich medium was refreshed daily in order to mimic the conditions found in a wound bed. The use of plate-to-plate transfer assays demonstrated measurable differences in the effectivenesses of several silver dressings on the viability of biofilm bacteria and their susceptibility to antibiotics. Whereas after the first day of treatment, all dressings used resulted in a significant reduction in the number of viable cells in the biofilms and disruption of the biofilm colonies, during prolonged treatment, the efficacy of dressings with hydrophilic base materials diminished with daily transfers, and bacterial populations recovered. For dressings with hydrophobic base materials, the level of efficacy correlated with the silver species loaded. Biofilm bacteria, which survived the initial silver treatment, were susceptible to tobramycin, ciprofloxacin, and trimethoprim-sulfamethoxazole, in contrast to untreated biofilms, which were highly tolerant to the same antibiotics. This acquired susceptibility was unaffected by the longevity of pretreatment with the silver dressings but depended on the dressing used. The antimicrobial efficacy of the dressings correlated with the type of the dressing base material and silver species loaded. * Corresponding author. Mailing address: Department of Mechanical and Manufacturing Engineering, 2500 University Drive NW, University of Calgary, Calgary, Alberta T2N 1N4, Canada. Phone: (403) Fax: (403) rmartinu@ucalgary.ca. Published ahead of print on 20 September Bacterial biofilms are a predominant challenge to wound healing (31). Biofilms are generally characterized as mono- or multispecies communities of microbial cells attached to surfaces and/or each other and embedded in self-produced extracellular polymeric substances (EPS) (36). Unlike planktonic populations, bacterial cells embedded in biofilms exhibit intrinsic resistance to antibiotics due to several specific defense mechanisms conferred by the biofilm environment, including the inactivation of antimicrobial agents by EPS, overexpression of stress-responsive genes, oxygen gradients within the biofilm matrix, and differentiation of a subpopulation of biofilm cells into resistant dormant cells (10, 17). The intrinsic resistance of bacterial cells within biofilms to conventional antimicrobials has motivated new approaches for the treatment of biofilm-associated infections, including the use of silver preparations. Unlike antibiotics, silver interferes with multiple components of bacterial cell structures and functions, including cell membrane integrity, respiratory chains, transmembranous energy and electrolyte transport, enzyme activities, and cell proliferation (20), and, hence, is less affected by specific biofilm microenvironmental variations (3). Moreover, silver is known to decrease bacterial adhesion and destabilize the biofilm matrix by compromising intermolecular forces (5, 18). Several silver-containing dressings are recommended for long-term decontamination and wound healing based on silver s broad-spectrum, high-level antimicrobial activity (21). Surprisingly, data on the effectiveness of silver dressings on bacterial biofilms during prolonged treatment are extremely limited. Percival et al. (28) previously reported the in vitro bactericidal activity of silver-containing Hydrofiber dressings against bacterial biofilms after 24 h of treatment, with total kill within 48 h, and recommended this dressing for healing recalcitrant wound infections. However, their data do not correlate well with in vivo observations reported previously by Hegger et al. (12), where silver-containing dressings reduced biofilm populations in an animal infection model by less than 90% over a week of treatment. Similarly, while silver coatings were reported to prevent biofilm formation on the surface of endotracheal tubes and catheters for a few days (2, 34), longer use results in device contamination (25). Device colonization has been linked to the inactivation of silver ions by the protein-rich deposit on the device surface or by wound bed compounds thus depleting antimicrobial activity (1). Unfortunately, existing in vitro protocols for antimicrobial efficacy of silver preparations (37, 40, 43), including the biofilm model reported by Percival et al. (28), do not reflect these microenvironmental conditions in the wound bed (16, 22). Recently, Hill et al. (13) attempted to mimic the wound bed environment in an in vitro model. In their experiment, biofilms of wound pathogens were exposed to silver dressings moistened with protein-rich medium. Thus, this model addresses the issue of reduced silver bioavailability in wound beds, since medium compounds compete with bacterial cells for silver absorption. The biofilms were transferred daily onto fresh dressings to simulate daily dressing replacement, a practice widely used in wound management. The results of this study indicated that most of the tested dressings, including the silver- 5120

2 VOL. 54, 2010 SILVER DRESSINGS EFFICACY AGAINST BACTERIAL BIOFILMS 5121 containing Hydrofiber dressings previously assessed by Percival et al. (28), were ineffective against bacterial biofilms after the seventh day of exposure. This conclusion was based only on endpoint data. Hence, it is difficult to reconcile these results with those of earlier studies focused on short-term silver dressing activity. Hill et al. (13) attributed this ineffectiveness to a limited release of silver from the dressings. However, it should be noted that the effective release of Ag from the dressings generally requires a higher level of hydration (19) than that provided in the experimental procedure described by Hill et al. (13). High hydration is typical for chronic wounds contaminated with bacterial pathogens due to a high rate of release of wound fluid as an inflammatory response (38). Thus, the low level of hydration in the model of Hill et al. likely limits silver release from the dressings and, hence, underestimates the dressings efficacies. Moreover, the dressings were refreshed daily, whereas these are designed for long-term treatment without replacement (21). The present study aimed to design a in vitro model where bacterial biofilms and silver dressings were transferred daily into fresh protein-rich medium over a week. This approach is hypothesized to more closely mimic the wound environment during prolonged treatment without dressing replacement by addressing (i) the silver dressing-to-medium ratio, which allows sufficient Ag release; (ii) daily refreshment of the protein levels to control silver bioavailability; and (iii) biofilm bacteriumsilver dressing interactions similar to those in wound beds. This model was used to study five commercially available silver wound dressings with different properties, attributed to the type of silver species and base materials, for their capacity to destroy biofilms formed by common wound pathogens methicillin-resistant Staphylococcus aureus (MRSA), Pseudomonas aeruginosa, and Escherichia coli; diminish the viability of biofilm bacterial cells; and increase bacterial cell susceptibility to conventional antibiotics during short-term and prolonged treatments. MATERIALS AND METHODS Bacterial strains and silver-containing dressings. Nosocomial isolates with known biofilm-forming capacities, P. aeruginosa PAO1, MRSA UC18, and E. coli BSC11380, were kindly provided by the Biofilm Research Group (University of Calgary, Canada). The dressings, Acticoat nanocrystalline silver on polyethylene mesh (NCPE), Silverlon metallic silver on nylon core (MSN), Aquacel Ag silver carboxymethylcellulose (SCMC), SilverCel metallic silver with alginate (MSAL), and PolyMem Silver metallic silver with starch copolymers on a polyurethane membrane (MSPU), were tested in blind assays. Biofilm daily plate-to-plate transfer assay. Biofilms were grown in an MBEC device (Innovotech, Canada), a 96-peg lid compatible with a standard 96-well microplate, according to the manufacturer s instructions and as previously described (4). Briefly, a bacterial suspension of 10 7 CFU/ml was incubated in MBEC devices on an orbital shaker at 0.25 g for 24 h. The lids with biofilms were submerged in the challenge plates with 0.2-cm 2 dressing samples and tryptic soy broth (TSB) to a final volume of 0.2 ml. For all tests, a ratio of 1 ml medium volume to 1 cm 2 of silver-releasing area was maintained. This ratio yields a silver concentration sufficient to kill planktonic bacteria after2hofexposure (data not shown). The plates were incubated for 24 h at 37 C. After that, dressing samples and lids with biofilms were transferred into microplates with fresh medium and incubated for an additional 24 h. This procedure was repeated daily for a week. Daily, a series of pegs with biofilms were treated with 0.4% sodium thioglycolate (STG) for 15 min (determined to be sufficient to neutralize remaining silver accumulated within biofilms), washed by submerging biofilm-covered pegs into sterile phosphate-buffered saline (PBS) for 5 min, and sonicated in TSB at 40 khz for 10 min as described previously by Ceri et al. (4). The numbers of bacterial cells resuspended from biofilms were determined with plate counts. The efficacy of silver dressings was determined as the log reduction in viable cell numbers after 1 or 7 days of exposure. The silver concentration needed to kill 90% (1 log 10 ) of biofilm cells was determined as the concentration of silver released from the dressing divided by the corresponding log reduction in the viable biofilm population. Effect of antibiotics on viability of silver-pretreated biofilm bacteria. Biofilms were grown and treated in daily transfer assays as described above. After the first and seventh days of treatment, the biofilms were treated with 0.4% STG for 15 min to neutralize silver accumulated in the biofilms and washed with PBS. A series of pegs were sonicated in TSB at 40 khz for 10 min. The number of cells resuspended from biofilms was determined with plate counts. The remaining pegs were transferred into challenge plates with 0.2 ml of Mueller-Hinton broth supplemented with 10 the MIC of tobramycin (TOB), ciprofloxacin (CIP), or trimethoprim-sulfamethoxazole (SXT). MICs were determined according to Clinical and Laboratory Standards Institute guidelines (6). The plates were incubated for 24 h. After that, viable cell numbers within biofilms were determined as described above. Viability was represented as the percentage of viable cell numbers within the biofilms pretreated with silver dressings and exposed to antibiotics compared to the number of cells that survived exposure to silver dressings alone. Scanning electron microscopy and energy-dispersive X-ray analysis of the biofilms. Biofilms were grown and treated in the daily transfer assays as described above. After the first and seventh days of treatment, the biofilms were fixed with 5% glutaraldehyde, dehydrated in serial ethanol dilutions, and viewed with an FEI ESME XL30 scanning electron microscope (SEM) (Microscopy and Imaging Facility, University of Calgary) to observe biofilm morphology and distribution. Biofilm coverage of a peg surface and colony sizes were analyzed with ImageJ image analysis software. Elemental analysis of the biofilms was performed with energy-dispersive X-ray spectrometry and a data analysis system (EDAX). The relative silver deposition in the biofilms was determined as the silver-to-carbon weight ratio, where carbon was used as an indicator of overall biomass deposition. Calcofluor white assay. Biofilms were grown and treated in the daily transfer assays as described above. After the desired intervals, biofilms were stained with 0.1% calcofluor white (CFW) for 15 min and then washed to remove stain unbound to EPS (13). The calcofluor white fluorescence intensity (488 nm/543 nm) was monitored by a Perkin-Elmer Victor V5 multilabel counter. To determine the overall biofilm matter (cells and EPS) deposited on the peg, biofilmcovered pegs were stained with crystal violet for 15 min, washed to remove unbound stain, and exposed to ethanol for 15 min to dissolve crystal violet bound to biofilms. Optical densities at 570 nm (OD 570 ) were measured with a Beckman Coulter AD340C absorbance detector. The amount of EPS was determined as CFW units, which are the CFW fluorescence intensities normalized by the OD 570. Confocal scanning laser microscopy of the biofilms. Biofilms for confocal scanning laser microscopy (CSLM) were grown on glass coverslips (VWR Scientific) and treated in daily plate-to-plate transfer assays. Briefly, sterile coverslips were submersed into six-well multidishes with 4 ml of a bacterial suspension of 10 7 CFU/ml and incubated on the orbital shaker at 0.25 g for 24 h. Silver dressing samples of 4 cm 2 were added to the biofilm-covered coverslips and submerged in six-well multidishes with 4 ml of fresh TSB. The plates were incubated for 24 h. After that, the dressing samples and the biofilm-coated slips were transferred into multidishes with fresh medium and incubated for an additional 24 h. This procedure was repeated daily for a week. After the first and seventh days of incubation, slips with biofilms were stained by using the Live/ Dead BacLight bacterial viability kit (L7012; Molecular Probes, Invitrogen) according to the manufacturer s instructions and examined for bacterial cell viability and biofilm morphology with an Olympus FV1000 confocal scanning laser microscope. Additionally, bacterial cells/aggregates resuspended from biofilms following 4hofexposure to silver dressings were stained with a viability kit and examined with CSLM. Cells surviving the treatment were identified by green fluorescence staining due to the uptake of Syto 9, while cells with compromised membranes absorbed propidium iodide and were identified by red fluorescence. The live/dead cell ratio and average thickness of the biofilms were analyzed with Fluorview image analysis software. Silver dressing absorption capacity, silver content in the dressings, and silver release dynamics. Dressing samples of 1 cm 2 were submerged in 1 ml of TSB in 24-well multidishes. Plates were incubated for 24 h, and the dressings were then transferred into fresh medium. This process was repeated daily for a week. Daily, the concentration of silver was analyzed in 0.5 ml of solution by atomic absorption spectrometry (SpectrAA 200; Varian Inc.). The volume of medium absorbed by the dressing was determined as the initial volume of medium minus the volume after exposure to the dressing. The relative silver content in the dressings

3 5122 KOSTENKO ET AL. ANTIMICROB. AGENTS CHEMOTHER. TABLE 1. Characteristics of biofilms exposed to silver dressings for 1 and 7 days in a daily transfer assay Mean value SD for groupa Organism and parameter b NCPE MSN 1st day 7th day 1st day 7th day P. aeruginosa Log 10 reduction in CFU/peg % reduction in surface coverage % reduction in biofilm thickness P/C ratio CFW units % live cells in surface (detached) biofilms ( ) ( ) ( ) ( ) MRSA Log 10 reduction in CFU/peg % reduction in surface coverage % reduction in biofilm thickness P/C ratio CFW units % live cells in surface (detached) biofilms ( ) ( ) ( ) ( ) E. coli Log 10 reduction in CFU/peg % reduction in surface coverage % reduction in biofilm thickness P/C ratio CFW units % live cells in surface (detached) biofilms ( ) (0 0) ( ) ( ) a NCPE, Acticoat nanocrystalline silver on polyethylene mesh; MSN, Silverlon metallic silver on nylon core; SCMC, Aquacel Ag silver carboxymethylcellulose; MSAL, SilverCel metallic silver with alginate; MSPU, PolyMem Silver metallic silver with starch copolymers on a polyurethane membrane. b P/C ratio, ratio of phosphorus to carbon weight as determined by energy-dispersive X-ray spectrometry; CFW units, calcofluor white florescence intensity normalized by the biofilm biomass concentration. was determined from the intensity of the characteristic atom radiation with EDAX. The data were normalized by the initial silver content in the corresponding dressing. Silver dressing contamination. Dressing samples of 1 cm 2 were exposed to 1 ml of bacterial suspensions of 10 7 CFU/ml in 24-well multidishes. Plates were incubated for 24 h, and the dressing samples were then transferred into the plates with fresh bacterial suspensions of 10 7 CFU/ml. This process was repeated daily for a week. Daily, the silver dressing samples were removed from the plates, treated with 0.4% STG for 15 min, washed with PBS, and sonicated in TSB at 40 khz for 10 min to resuspend bacterial cells from the dressings. The number of viable cells was determined by plate counts. To determine the overall number of cells embedded, the dressing samples were fixed in 5% glutaraldehyde, dehydrated in ethanol serial dilutions, and examined for bacterial contamination with an FEI ESME XL30 SEM. Cell numbers were evaluated with ImageJ image analysis software. Statistical analysis. All experiments were performed a minimum of six times. Results are reported in this paper in terms of means standard deviations over multiple measurements. A one-way analysis of variance (ANOVA) was used to determine statistical significance. A P value of 0.05 indicates a significant difference in the efficacy of the silver dressings. Spearman s rank correlation coefficient, r, was used to characterize the relationship between silver load and accumulation within biofilms and biofilm suppression. RESULTS Time-dependent effects of silver-containing dressings on biofilm bacterial viability and colony morphology. The timedependent effect of silver dressings on bacterial biofilms was assessed from the daily plate-to-plate transfer assays. In general, the antibiofilm efficacy varied according to the dressing used (P 0.05) and longevity of treatment (P 0.05), although all dressings significantly decreased the number of cells in biofilms after 24 h of exposure (Table 1). In silver-free medium, P. aeruginosa PAO1 formed biofilm carpets of log 10 CFU/peg embedded in profuse EPS covering 84.6% 4.2% of peg surfaces, as observed with an SEM (Fig. 1) and confirmed by calcofluor white binding to EPS (CFW units of ) (Table 1). MRSA UC18 biofilms of log 10 CFU/peg covered 38.2% 6.8% of the peg surfaces and consisted of isolated EPS-containing (CFW units of ) colonies of 45 to 600 m in diameter (Fig. 2). E. coli BSC11380 formed colonies of 10 to 70 m in diameter covering 80.9% 6.4% of the peg surfaces with log 10 CFU/peg. EPS within E. coli biofilms was not detected by either an SEM (Fig. 3) or calcofluor white ( units). The average thickness of colonies, determined by CSLM, varied from 54 to 88 m for both P. aeruginosa and MRSA and from5to12 m for E. coli. CSLM observation of the biofilms stained with the viability kit demonstrated the presence of mostly live cells within the biofilms (data not shown). The nanocrystalline silver-containing dressing (NCPE) dramatically decreased viable cell numbers (by 4 to 5 log 10 CFU/ peg) within the tested biofilms after the first day of treatment and maintained less than 3 log 10 CFU of viable cells over a week (Table 1). A 1-day exposure to the hydrophilic base material dressings SCMC, MSAL, and MSPU resulted in a 2- to 3.5-log 10 reduction of viable cells within biofilms. However, further treatment in the daily transfer assay resulted in biofilm bacterial regrowth (P 0.05). The metallic silver on hydrophobic base dressing, MSN, demonstrated relatively low antibiofilm activity (approximately a 1-log 10 reduction) during the first day of exposure but decreased viable cell numbers within biofilms by 2 to 3 log 10 CFU by the seventh day. Thus, the dressings categorized in the three groups demonstrated different biofilm-killing efficacies (P 0.05) independent of the microorganisms treated (P 0.05). However, the effect of the dressing on the peg surface coverage by biofilms and the thickness of the biofilm colonies differed between the microbial strains tested (P 0.05), as described below. These differences

4 VOL. 54, 2010 SILVER DRESSINGS EFFICACY AGAINST BACTERIAL BIOFILMS 5123 TABLE 1 Continued Mean value SD for group a SCMC MSAL MSPU 1st day 7th day 1st day 7th day 1st day 7th day ( ) ( ) ( ) ( ) ( ) ( ) ( ) ( ) ( ) ( ) ( ) ( ) ( ) ( ) ( ) ( ) ( ) ( ) reflected variations in microbial biofilm properties such as EPS production and residual attachment of dead cells (see below). SEM observations indicate that NCPE, SCMC, MSAL, and MSPU reduced the surface coverage by P. aeruginosa biofilms by 16 to 30% (Table 1). No difference (P 0.05) in the surface coverage was observed after 1 or 7 days of exposure. In contrast, MSN, which hardly decreased surface coverage by P. aeruginosa biofilms after the first day, caused a 22% reduction after 7 days. The decrease in surface area covered by the MRSA biofilm after a 1-day exposure to silver dressings was 15 to 35%. Prolonged treatment with NCPE and MSN provided a further reduction in the surface coverage by MRSA biofilms, whereas the area on the peg surface colonized by biofilms increased during prolonged treatment with SCMC, MSAL, and MSPU (P 0.05). The surface coverage by E. coli biofilms was reduced by the tested dressings by approximately 20% after the first day. The 7-day exposure to silver dressings resulted in a further reduction of the area covered by E. coli biofilms (P 0.05), except for biofilms treated with MSAL, which showed an increase in the surface coverage by the seventh day (P 0.05). The average thickness of all biofilms exposed to NCPE was dramatically reduced (P 0.05) after both 1 and 7 days, as determined by CSLM (Table 1). SCMC, MSAL, and MSPU significantly (P 0.05) reduced the biofilm thickness after the first day. However, prolonged treatment allowed colony regrowth (P 0.05). MSN showed low destructive activity after the first day of treatment (P 0.05) but significantly reduced the biofilm thickness by the seventh day (P 0.05). CSLM observations revealed the presence of large (more than 20 cells) aggregates detached from the biofilms and increased surface roughness of colonies treated with silver dressings compared to untreated biofilms (data not shown), which suggested an induced sloughing of biofilm sections rather than the erosion of cells. Both detached and surface populations consisted of both live and dead cells (Table 1), indicating that the biofilm sloughing results from the weakening of the biofilm structure via silver activity in addition to the death of bacterial cells within biofilms. SEM images of P. aeruginosa biofilms after exposure to the dressings showed a profuse matrix regardless of treatment longevity (Fig. 1). No significant changes (P 0.05) in calcofluor white absorption (CFW units) were observed (Table 1), indicating that 1 day of silver dressing treatment did not impact the EPS amount. However, after prolonged treatment with SCMC, MSAL, and MSPU, numbers of CFW units increased (P 0.05), indicating increased EPS production by recovering biofilm cells. A significant portion of the cells within the matrix were killed, as evidenced by propidium iodide uptake (see % live cells in Table 1). For the biofilms treated with SCMC, MSAL, and MSPU, approximately one-half of the biofilm bacteria were dead after the first day of treatment, but the proportion of live cells increased with prolonged treatment. P. aeruginosa biofilms contained 50% and 97% compromised cells after the first day of treatment with MSN and NCPE, respectively, with further increases in the number of dead cells with longer exposures. MRSA lost EPS after treatment with NCPE, SCMC, and MSPU for 1 day, as observed with an SEM (Fig. 2) and confirmed with reduced numbers of CFW units (Table 1). In contrast, SCMC and MSPU during prolonged treatment, as well as MSAL and MSN after both 1 and 7 days of treatment, stimulated EPS production by MRSA. Most of the cells remaining attached within the MRSA biofilms after treatment were alive (Table 1). E. coli biofilms treated with dressings for a day contained a significant portion of cells with compromised membranes but small amounts of EPS (Fig. 3), as confirmed by low live/dead

5 5124 KOSTENKO ET AL. ANTIMICROB. AGENTS CHEMOTHER. FIG. 1. SEM images of P. aeruginosa biofilms developed on the peg surface before exposure (left) to Acticoat nanocrystalline silver on polyethylene mesh (NCPE), Silverlon metallic silver on nylon core (MSN), Aquacel Ag silver carboxymethylcellulose (SCMC), SilverCel metallic silver with alginate (MSAL), and PolyMem Silver metallic silver with starch copolymers on a polyurethane membrane (MSPU) and after 1 day (middle) and 7 days (right) of the treatment in a daily transfer assay. Bar, 5 m.

6 VOL. 54, 2010 SILVER DRESSINGS EFFICACY AGAINST BACTERIAL BIOFILMS 5125 FIG. 2. SEM images of MRSA biofilms developed on the peg surface before exposure (left) to NCPE, MSN, SCMC, MSAL, and MSPU and after 1 day (middle) and 7 days (right) of the treatment in a daily transfer assay. Bar, 5 m.

7 5126 KOSTENKO ET AL. ANTIMICROB. AGENTS CHEMOTHER. FIG. 3. SEM images of E. coli biofilms developed on the peg surface before exposure (left) to NCPE, MSN, SCMC, MSAL, and MSPU and after 1 day (middle) and 7 days (right) of the treatment in a daily transfer assay. Bar, 5 m.

8 VOL. 54, 2010 SILVER DRESSINGS EFFICACY AGAINST BACTERIAL BIOFILMS 5127 TABLE 2. Effects of antibiotics on viability of biofilm cells pretreated with silver dressings for 1 and 7 days in a daily transfer assay Organism and antibiotic b Mean % viable cells SD a NCPE MSN SCMC MSAL MSPU 1st day 7th day 1st day 7th day 1st day 7th day 1st day 7th day 1st day 7th day P. aeruginosa TOB CIP SXT MRSA TOB CIP SXT E. coli TOB CIP SXT a Percentage of viable cell numbers within the biofilms pretreated with silver dressings and exposed to antibiotics compared to the number of cells that survived exposure to silver dressings alone. b TOB, tobramycin; CIP, ciprofloxacin; SXT, trimethoprim-sulfamethoxazole. cell ratios and low numbers of CFW units (Table 1). However, similarly to MRSA, the regrowth of E. coli during prolonged treatment with SCMC, MSAL, and MSPU was accompanied by EPS production. Antibiotic susceptibilities of biofilms pretreated with silver dressings. Table 2 summarizes the percent survival of bacterial cells within biofilms pretreated with silver dressings in the daily transfer assay after exposure to 10 the MICs of tobramycin (TOB), ciprofloxacin (CIP), and trimethoprim-sulfamethoxazole (SXT). Planktonic cells were sensitive to the antibiotics, with the exception of P. aeruginosa, which is intrinsically resistant to SXT. The MICs of TOB, CIP, and SXT were 0.5 g/ml, 0.25 g/ml, and 85 g/ml for P. aeruginosa, respectively; 2.0 g/ml, 0.5 g/ml, and 0.1 g/ml for E. coli, respectively; and 1.0 g/ml, 0.25 g/ml, and 0.04 g/ml for MRSA, respectively. Bacteria in untreated biofilms, however, were resistant to all of the antibiotics used and survived with 10 the MICs for at least 24 h (data not shown). In contrast, biofilm bacteria that had survived silver pretreatment were significantly more susceptible (P 0.01) to antibiotic attack. Pretreatment with silver dressings resulted in 70 to 97% killing of bacterial cells within biofilms. It is remarkable that pretreatment with silver dressings decreased the resistance of P. aeruginosa to SXT. Susceptibility of cells within silver-treated biofilms depended on the dressings used (P 0.05) but not on the longevity of treatment and the microbial strain exposed (P 0.05). Accumulation of silver in the biofilms. Bacterial biofilms treated with silver dressings in daily transfer assays were subjected to EDAX in order to determine the amount of silver accumulated in the biofilms. Large amounts of silver (8 to 15% of the biomass carbon weight) were found to accumulate in biofilms exposed to NCPE (Table 3). Silver accumulations in biofilms treated with MSN, SCMC, MSAL, and MSPU were less than 7% for P. aeruginosa and MRSA biofilms and less than 3% for the E. coli biofilm. The silver content in biofilms after prolonged treatment with NCPE and MSN increased nearly 2-fold. In contrast, prolonged treatment with other dressings resulted in reduced silver contents in the biofilms compared to 1-day-treated biofilms. Thus, silver accumulation in biofilms ranged according to the dressing type, longevity of treatment, and strain treated (P 0.05). Variability in the accumulation of silver between different biofilms is attributed to biofilm composition, and in particular EPS content (r 0.78), whereas the difference between dressings effects is associated with the silver species used in the dressing (r 0.92). Absorption capacity, silver release into medium, and silver content in wound dressings. SCMC, MSAL, and MSPU demonstrated high absorption capacities, 0.5 to 0.8 ml/cm 2 sample, while NCPE and MSN absorbed approximately 0.2 ml/cm 2 (Table 4). Concentrations of silver released into medium from MSAL and MSPU after the first day of exposure ranged from 10 to 15 g/ml and then dropped down to 3 to 4 g/ml (Table 4). NCPE and MSN maintained silver concentrations of 8 to 14 g/ml during the entire experiment. SCMC released about 30 g/ml of silver in medium after the first day of exposure, but TABLE 3. Silver contents in biofilms after exposure to silver dressings for 1 and 7 days in a daily transfer assay Mean silver/carbon wt ratio SD Organism NCPE MSN SCMC MSAL MSPU 1st day 7th day 1st day 7th day 1st day 7th day 1st day 7th day 1st day 7th day P. aeruginosa MRSA E. coli

9 5128 KOSTENKO ET AL. ANTIMICROB. AGENTS CHEMOTHER. TABLE 4. Silver dressing characteristics after 1 and 7 days of treatment in a daily transfer assay Mean value for group SD a Parameter NCPE MSN 1st day 7th day 1st day 7th day Medium absorption (ml/cm 2 ) NT NT Silver released ( g/ml) % silver content in dressing compared to initial value Overall/viable cell no. in dressing (log 10 bacterial cells/cm 2 ) P. aeruginosa ( ) ( ) ( ) ( ) MRSA ( ) ( ) ( ) ( ) E. coli ( ) ( ) ( ) ( ) a NT, nontested. the silver concentration decreased to 1.5 g/ml by the end of the seventh day. The silver contents at the first and seventh days of the experiments were normalized to data observed for brand-new dressing samples and are shown in Table 4. After the first day of exposure, SCMC and MSPU still contained 50 to 60% silver loaded in the dressing. MSAL contained approximately 70%. The further loss of silver from these types of dressings was insignificant, indicating a slow release of silver into the medium. NCPE and MSN retained 80% to 90% of the initial silver after the first day of exposure, but only 40% of the initial silver remained by the end of the experiment. While tested dressings provided similar silver releases after 1 day of exposure (P 0.05), their long-term activity significantly differed (P 0.05), which correlated well with medium absorption by dressings (r 0.96) but not with silver load (r 0.32). Silver dressing contamination. The potential absorbance of wound pathogens by the dressings was assessed with an SEM. The number of viable cells attached to the dressing substrate was determined by plate counts. The hydrophobic base material dressings (NCPE and MSN) entrapped between 10 and 100 bacterial cells/cm 2. These values did not change (P 0.05) over a week of exposure (Table 4). MSAL entrapped approximately 10 3 bacterial cells/cm 2 after the first day, with no change over a week. SCMC and MSPU were contaminated with 10 3 to 10 4 cells/cm 2 after the first day of exposure. However, the number of entrapped cells increased 5- to 10-fold by the seventh day. Thus, pathogen absorption depended on the dressing type and longevity of treatment (P 0.05) but not on the microbial strain used (P 0.05). The viability of entrapped bacterial cells was also independent of the microbial strain (P 0.05) but depended on the type of dressing (P 0.05). Fewer than 1% of bacterial cells entrapped in NCPE survived during the experiment. MSN and MSAL allowed the survival of 2 to 4% of entrapped cells after the first day of exposure, but the proportion of viable cells doubled by the seventh day. SCMC and MSPU allowed the survival of 3 to 5% and 7.5 to 10% of entrapped cells, respectively, after 1 day of treatment, but the number of viable cells increased (22 to 28%) after prolonged exposure. DISCUSSION In this study, the long-term antimicrobial efficacy of silver dressings was investigated in an in vitro model, which addressed two important issues complicating wound healing: the development of recalcitrant bacterial biofilms (31) and the inactivation of antimicrobials by wound bed compounds (19). The diagnostic model designed for this study uses daily refreshment of protein-rich media to reduce the bioavailability of silver. Nutrient media were observed to decrease the bioavailability of silver by at least 2 orders of magnitude compared to water or PBS (32), which is still approximately 5-fold lower than chelation by biological fluids (11). Thus, protein-rich TSB medium was refreshed daily to increase the chelation capacity of the experimental medium. In traditional batch assays, silver released from the dressings is absorbed by medium to a certain saturation level above which it will be free to interact with bacterial cells. In contrast, exudate production in the chronic wound bed is continuous, providing a continuous supply of proteins to bind silver ions (38). If TSB is refreshed daily, fresh proteins will be provided daily to consume a certain amount of silver ions from every portion of silver released daily from the dressing. This approach fits well the wound bed scenario. Observing that exudate is partially absorbed by the dressing and partially accumulated between the dressing and underlying tissue (15), biofilm bacteria in the wound bed can interact directly with the silver dressings or be exposed to silver accumulated in wound fluid (38). To bring the biofilm into direct contact with the dressings and silver released into the medium, biofilms were grown in MBEC devices, a lid with 96 pegs submerged into a 96-well microplate. The biofilms were grown on the pegs coated with serum and submerged into medium supplemented with dressing samples. The optimal ratio of medium volume to dressing area was determined from the dressings absorptions and antimicrobial efficacies (Table 4). The MBEC device allows the nondestructive transfer of the biofilms into fresh medium and, hence, does not impact viability and susceptibility assessments. The density of bacterial cells within biofilms was maintained at a level comparable to that determined for biofilms formed in wound beds, 10 5 to 10 8 CFU/ml (8). This experimental design permitted the highthroughput quantitative assessment of the silver released from

10 VOL. 54, 2010 SILVER DRESSINGS EFFICACY AGAINST BACTERIAL BIOFILMS 5129 TABLE 4 Continued Mean value for group SD a SCMC MSAL MSPU 1st day 7th day 1st day 7th day 1st day 7th day NT NT NT ( ) ( ) ( ) ( ) ( ) ( ) ( ) ( ) ( ) ( ) ( ) ( ) ( ) ( ) ( ) ( ) ( ) ( ) the dressings and its effect on biofilm bacterial cell viability and susceptibility to antibiotics on a daily basis. The assessment efficacy of nonsilver wound dressings is beyond the scope of this paper. The long-term silver dressings antimicrobial activities observed with this model correlate better with clinical observations than data obtained with traditional batch assays. After the first day of treatment, all tested dressings exhibited marked antibiofilm efficacy, which correlated well with results of shortterm wound management (9, 21). Further treatment of the biofilms without daily transfers (i.e., the batch experiments) resulted in a gradual reduction in viable cell numbers within the biofilms, as independently observed by our group (data not shown) and Percival et al. (28), but were in disagreement with previously reported clinical observations (12, 33). In contrast, prolonged treatment with daily transfer decreased the efficacies of most tested dressings, resulting in a recovery of the biofilms after 7 days that was consistent with clinical observations (12, 41, 42). Thus, measuring and predicting the antimicrobial effectivenesses of different silver dressings can be improved by the use of the daily transfer assay to more closely mimic the actual wound bed environment. Although all presently tested silver dressings killed bacterial cells within biofilms and destroyed biofilm colonies, the antimicrobial efficacies differed between dressings, especially during prolonged treatment. The factors underlying these differences are unclear. The antimicrobial efficacy of the silver dressings is attributed to the ability of silver cations, Ag,to damage cell membranes, suppress transmembranous transport and enzyme activity, inactivate DNA, and, thus, result in bacterial cell death (35). Silver ions bound to bacterial cells and EPS also interfere with intermolecular forces in biofilms and facilitate biofilm dispersion (5). Consequently, the antimicrobial effects of silver products would be expected to be related directly to the amount and rate of Ag release (20). However, previously reported data showed a lack of a consensus in supporting this conclusion. For example, Thomas and McCubbin (40) postulated that Acticoat produced the most rapid antimicrobial effect against S. aureus, E. coli, and Candida albicans compared to those of Contreet-H (Coloplast) and Actisorb Silver 220 (Johnson & Johnson), due to the rapid release of a relatively large concentration of silver ions. In contrast, Parsons et al. (27) showed that a larger amount of silver released from a dressing did not lead to a higher rate or degree of antimicrobial activity. In the present study, NCPE, which was observed to provide the highest antibiofilm efficacy, maintained 8 to 12 g/ml of silver in the medium. In contrast, SCMC and MSN maintained higher and similar silver concentrations compared to those of NCPE, respectively, but demonstrated lower antimicrobial efficacy. The correlation between silver concentration and biofilm suppression was as low as an r value of 0.36 when considering all the dressings tested but increased to an r value of 0.89 when restricted to the group of dressings loaded with metallic silver. Thus, the silver concentration alone cannot account for the antibiofilm efficacy of the silver dressings. The type of silver species present also plays a role. The dressings tested in the present study release Ag by replacing silver ions in silver carboxymethylcellulose, as in SCMC, or by gradually generating Ag from elemental silver particles, as in other dressings tested in this study (9, 21). The immediate release of silver by SCMC provided a high concentration of Ag, which, however, was quickly inactivated by medium compounds. Under these conditions, more than 9 g/ml of silver has to be released in order to kill 90% (1 log 10 ) of the biofilm cells. In contrast, the gradual release of Ag from metallic silver particles sustains the dressings activities for longer periods, and smaller amounts of silver (2.5 to 6.5 g/ml) are required to be released into the medium in order to kill 90% of the biofilm cells. The wide range of effective concentrations observed for dressings loaded with metallic silver can be explained by particle size variations. The reduction of the silver particle to the nanoscale level increases the relative surface area, which provides higher Ag release rates than for elemental silver particles (9). Moreover, nanoparticles have a higher capacity to attach to and penetrate bacterial membranes and accumulate inside cells, providing a continuous release of silver ions inside the cell (30; this study). The higher silver accumulations within biofilms treated with NCPE observed in this study correlate well with NCPE s higher efficacy (r 0.92). Nevertheless, the sustained release of sufficient amounts of silver is required for the long-term antibiofilm activity of the silver dressings. The prolonged release of silver concentrations greater than 8 g/ml (although the effective concentration may be lower, as described above) by NCPE and MSN supports their long-term activity. A reduction in

11 5130 KOSTENKO ET AL. ANTIMICROB. AGENTS CHEMOTHER. the silver concentration to below 3 g/ml, as determined for SCMC, MSAL, and MSPU after a 7-day exposure, resulted in the regrowth of bacteria embedded in biofilms. Silver levels above 5 g/ml were also reported previously by Bjarnsholt et al. (3) to be sufficient to suppress the growth of biofilm bacteria. The rate of Ag release is driven by the oxidation of elemental silver or the displacement of silver ions in the polymer fiber due to hydration and diffusion. For hydrophilic materials during prolonged exposure, however, the polymer is plasticized by water, which limits the silver ion release rate (7), and medium or wound fluid, absorbed in the dressing base material, binds silver ions entrapped in the dressing (29). As a result, silver ion release from hydrophilic dressing base materials is more efficient than that from more hydrophobic ones only in the short term (19). Hydrophilic dressings (SCMC, MSAL, and MSPU) generate high silver concentrations in medium just after hydration but have low residual silver release despite the high silver content in the dressings. Hence, the hydrophilic dressings lose their antimicrobial efficacy faster and become contaminated more easily than hydrophobic dressings, such as NCPE and MSN, which provide lower but sustained silver release. Limited medium absorption also decreases the inactivation of silver within the dressing material. The hydrophobic dressings immobilize fewer cells and kill them even during prolonged treatment. The high barrier capacity of NCPE observed in this study was also reported independently by Holder et al. (14) and Strohal et al. (39). However, the barrier function and antimicrobial activity of MSN were lower than those observed for NCPE despite the higher silver content. The lower efficacy of MSN might be explained by differences in the silver species present in these dressings: conventional metallic silver in MSN and nanocrystalline silver in NCPE. Although the tested dressings did not allow the complete eradication of the biofilms, the remaining bacterial cells, which had survived silver treatment, were susceptible to antibiotics, including tobramycin, ciprofloxacin, trimethoprim, and sulfamethoxazole. In contrast, bacterial cells in untreated biofilms were highly tolerant to the same doses of antibiotics. The combination of silver sulfadiazine or colloidal silver nanoparticles with some antibiotics was previously reported to increase antimicrobial activity against planktonic bacteria in in vitro assays and in a burned-mouse model (23, 24, 37). The mechanisms driving the sensitization of bacteria via silver exposure are not understood. One possibility is that the binding of silver to biofilm bacterial cells and matrix might prevent the chelation of antibiotics and increase their interaction with target cells. Another possible mechanism is that cell damage caused by silver treatment might promote an increased uptake of antibiotics and facilitate their activity. Silver ions and silver nanoparticles are known to mediate intracellular reactive oxygen species generation that could enhance the antimicrobial activity of antibiotics in the presence of silver (26). The present study revealed that silver accumulation within the biofilms was the leading factor (r 0.77) in enhancing susceptibility to antibiotics compared to the silver concentration in the medium (r 0.26). Conclusion. The long-term antimicrobial efficacy of silver wound dressings against bacterial biofilms was investigated in a 7-day treatment in vitro model, in which silver dressing samples and biofilms grown on serum-coated MBEC devices were transferred daily in fresh protein-rich medium in order to mimic the conditions found in a wound bed. Specifically, this model addresses conditions for the dressing s hydration, silver interaction with wound fluid and bacterial biofilms, the dressing s absorptiveness and contamination, and silver bioavailability during prolonged treatment. The results obtained with this biofilm plate-to-plate transfer model agree well with those observed for infected wounds and should predict the long-term activity of silver preparations more reliably than traditional batch assays. Five commercially available silver wound dressings tested using this model have demonstrated measurable differences in their antimicrobial efficacies, which is attributed to the type of silver species and base materials. Dressings employing hydrophilic base materials and loaded with ionic or metallic silver showed short-term efficacy. Reduced silver release by hydrophilic dressings after long exposure resulted in biofilm recovery and enhanced EPS production. Dressings with hydrophobic base material demonstrated sustained antibiofilm activity for at least 7 days when loaded with metallic or nanocrystalline silver. These results were independent of the strain used. The observed differences in antimicrobial efficacy can be explained by the diversity of silver species and the base materials comprising the dressings. The base material controls the concentration and the duration of silver release, while the types of silver species released from the dressings are responsible for the interaction with target cells within the biofilm. In particular, hydrophobic materials provide a sustained release of a sufficient amount of silver, while the minimization of silver particle size (to the nanoscale level) provides better penetration and accumulation of silver within biofilms and bacterial cells, thus contributing to the increased efficacy of the silver formulation. The application of silver dressings can also improve wound healing via antibiotic therapy since the interaction of silver released from the dressings significantly increases the susceptibility of bacterial cells within biofilms to the effects of antibiotics. ACKNOWLEDGMENT This work was supported by the National Sciences and Engineering Research Council of Canada through the Industrial Research Chair program for R.J.M. REFERENCES 1. 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