This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowReprints and Permissions
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Nett, J.
Right arrow Articles by Andes, D.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Nett, J.
Right arrow Articles by Andes, D.

 Previous Article  |  Next Article 

Antimicrobial Agents and Chemotherapy, February 2007, p. 510-520, Vol. 51, No. 2
0066-4804/07/$08.00+0     doi:10.1128/AAC.01056-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Putative Role of ß-1,3 Glucans in Candida albicans Biofilm Resistance{triangledown}

Jeniel Nett,1 Leslie Lincoln,1 Karen Marchillo,1 Randall Massey,2 Kathleen Holoyda,1 Brian Hoff,1 Michelle VanHandel,1 and David Andes1*

Departments of Medicine and Medical Microbiology and Immunology,1 University of Wisconsin Electron Microscopy Facility, University of Wisconsin, Madison, Wisconsin2

Received 22 August 2006/ Returned for modification 17 October 2006/ Accepted 12 November 2006


arrow
ABSTRACT
 
Biofilms are microbial communities, embedded in a polymeric matrix, growing attached to a surface. Nearly all device-associated infections involve growth in the biofilm life style. Biofilm communities have characteristic architecture and distinct phenotypic properties. The most clinically important phenotype involves extraordinary resistance to antimicrobial therapy, making biofilm infections very difficulty to cure without device removal. The current studies examine drug resistance in Candida albicans biofilms. Similar to previous reports, we observed marked fluconazole and amphotericin B resistance in a C. albicans biofilm both in vitro and in vivo. We identified biofilm-associated cell wall architectural changes and increased ß-1,3 glucan content in C. albicans cell walls from a biofilm compared to planktonic organisms. Elevated ß-1,3 glucan levels were also found in the surrounding biofilm milieu and as part of the matrix both from in vitro and in vivo biofilm models. We thus investigated the possible contribution of ß-glucans to antimicrobial resistance in Candida albicans biofilms. Initial studies examined the ability of cell wall and cell supernatant from biofilm and planktonic C. albicans to bind fluconazole. The cell walls from both environmental conditions bound fluconazole; however, four- to fivefold more compound was bound to the biofilm cell walls. Culture supernatant from the biofilm, but not planktonic cells, bound a measurable amount of this antifungal agent. We next investigated the effect of enzymatic modification of ß-1,3 glucans on biofilm cell viability and the susceptibility of biofilm cells to fluconazole and amphotericin B. We observed a dose-dependent killing of in vitro biofilm cells in the presence of three different ß-glucanase preparations. These same concentrations had no impact on planktonic cell viability. ß-1,3 Glucanase markedly enhanced the activity of both fluconazole and amphotericin B. These observations were corroborated with an in vivo biofilm model. Exogenous biofilm matrix and commercial ß-1,3 glucan reduced the activity of fluconazole against planktonic C. albicans in vitro. In sum, the current investigation identified glucan changes associated with C. albicans biofilm cells, demonstrated preferential binding of these biofilm cell components to antifungals, and showed a positive impact of the modification of biofilm ß-1,3 glucans on drug susceptibility. These results provide indirect evidence suggesting a role for glucans in biofilm resistance and present a strong rationale for further molecular dissection of this resistance mechanism to identify new drug targets to treat biofilm infections.


arrow
INTRODUCTION
 
Biofilms are structured microbial communities in which organisms irreversibly attach to a surface and become embedded in a matrix of extracellular polymeric substances produced by these cells (6, 15, 17, 20, 21, 24, 41, 46, 50). Biofilms are the most common form of microbial growth in nature, and the National Institutes of Health estimate that biofilms cause the majority of infections in humans (21, 28, 45, 46). Most times, an implanted device, such as an intravascular catheter, is associated with biofilm infections (48). C. albicans is the fourth and third leading cause of hospital-acquired bloodstream and urinary tract infections, respectively. Up to 70 to 80% of Candida bloodstream infections are associated with central venous catheters, and the majority of Candida urinary tract infections are associated with bladder catheters. Among central venous catheter-related infections, those due to Candida spp. are associated with mortality rates approaching 40% (44).

Biofilms have characteristic architectural and phenotypic properties distinct from their planktonic counterparts. Perhaps the most clinically relevant biofilm-specific property is the development of antimicrobial resistance that can be up to 1,000-fold greater than planktonic cells (2, 6, 9-12, 15, 25, 30, 31, 37, 47-49). This characteristic of biofilms makes them extremely difficult, if not impossible, to control in the medical setting. Current guidelines recommend device removal for patients with candidemia, as patients with biofilm-infected devices are rarely cured with only antifungal therapy (44).

Candida biofilms have been shown to be resistant to antifungals, including amphotericin B, triazoles, and flucytosine (12, 15, 25, 30, 31). Numerous mechanisms have been proposed to explain Candida biofilm resistance. Studies have examined the impact of drug diffusion, rate of growth, cell wall efflux pumps, and plasma membrane changes (2, 3, 9-11, 37, 47, 49). Each of these has been shown to impact biofilm cell susceptibility. Yet, these studies suggest another unidentified mechanism(s) is important for the drug resistance of Candida cells growing in a biofilm environment. Previous work by Douglas and colleagues demonstrated secreted carbohydrate differences between biofilm and planktonic cells (2, 10, 26). The current studies further explore cell wall and secreted carbohydrate differences associated with the biofilm mode of growth. Drug binding assays and cell wall component modification studies suggest these cell wall changes may contribute to antifungal biofilm drug resistance.


arrow
MATERIALS AND METHODS
 
Organisms and culture conditions. A clinical isolate of C. albicans, designated K1, was used for all studies. The strain was stored in 15% (vol/vol) glycerol stock at –80°C and maintained on yeast extract-peptone-dextrose medium (1% yeast extract, 2% peptone, and 2% dextrose) prior to experiments. Fresh colonies were used for inoculation of RPMI 1640 with morpholinepropanesulfonic acid (MOPS), and cultures were allowed to incubate for 16 h at 37°C with orbital shaking at 250 rpm prior to inoculation.

In vitro biofilm model. Two in vitro biofilm models were used for these studies. Studies for collection of a large amount of organism cell wall or culture supernatant utilized a circular piece of sterile medical-grade silicone disk as a substrate. The C. albicans inoculum was prepared by growth in RPMI-MOPS overnight at 37°C, followed by dilution to 1 x 106 in RPMI-MOPS based on hemacytometer counts. Prior to inoculation, the substrates were coated with mouse serum and incubated at 37°C for 30 min. Following this conditioning period, 150 µl of the inoculum was placed on the surface of the silicone disk. The disks were then incubated with 1 ml of RPMI-MOPS for 45 min at 37°C. After the adherence period, they were washed and placed in 10 ml of fresh medium for further incubation with gentle shaking (80 rpm) for 48 h. The disks were then rinsed with sterile saline to remove nonadherent cells, and biofilm formation was confirmed by visual inspection or by confocal microscopy with FUN-1 and concanavalin A fluorescent staining as previously described (6, 15). To harvest cells, disks were rinsed and placed in 30 ml sterile water, where cells were dislodged manually and by gentle sonication (water bath sonicator; 40-kHz transducer) for 10 min. After removal of the disks, the cells were collected by centrifugation and washed with sterile water prior to additional study.

Studies designed to assess the impact of drug therapy on biofilm cell viability used a polystyrene substrate in a 96-well format as previously described by Ramage et al. (49). For the polystyrene assay, 100 µl of the inoculum was added to each well of a 96-well flat-bottom plate. After a 24-h incubation at 37°C, the wells were washed with phosphate-buffered NaCl (PBS) three times to remove any nonadherent cells. Fresh medium and various drugs were added followed by additional periods of incubation. Quantitation of biofilm cell viability utilized the tetrazolium salt 2,3-bis-(2-methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-carboxanilide inner salt (XTT) reduction assay as previously described (49). Briefly, 90 µl of XTT (1 mg/ml) and 10 µl phenazine methosulfate (320 µg/ml) were added to each well and the plate was incubated at 37°C for 2 h. Absorbance at 490 nm was measured using an automated plate reader. Each of the above growth assays was performed in triplicate for each of the experiments described below.

In vivo C. albicans venous catheter biofilm and disseminated infection models. A rat central venous catheter infection model was selected for in vivo biofilm studies (6). Briefly, polyethylene tubing (inner diameter, 0.76 mm; outer diameter, 1.52 mm) was chosen. Specific-pathogen-free Sprague-Dawley rats weighing 400 g were used (Harlan Sprague-Dawley, Indianapolis, Ind.). A heparinized (100 U/ml) catheter was surgically inserted into the external jugular vein and advanced to a site above the right atrium (2-cm length). The catheter was secured to the vein, and the proximal end was tunneled subcutaneously to the midscapular space and externalized through the skin. The catheters were placed 24 h prior to infection to allow a conditioning period for deposition of host protein on the catheter surface. Infection was achieved by intraluminal instillation of 500 µl C. albicans K1 cells (106 cells/ml). After a dwelling period of 4 h, the catheter volume was withdrawn and the catheter was flushed with heparinized 0.15 M NaCl. Two animals were used per time and treatment point. Blood was drawn from the catheter and tail vein of the rats at 12 h for glucan measurement as described below. At the end of the observation period, the catheters were removed and processed by examination of the cell wall and biofilm architecture using electron microscopy, as described below. The viable burden of Candida cells in the kidney was used as an estimate of total body Candida burden, as previously described (6, 7). The kidneys of rats were removed, homogenized, and plated on Saboraud's dextrose agar for plate counts.

A disseminated candidiasis model, without a biofilm catheter infection, was used as a comparison (7). The comparator represents a systemic Candida infection without a device biofilm infection. The same rat species was injected with a similar C. albicans inoculum via the tail vein. Blood was collected from the tail vein for glucan measurement as described below. The kidneys of animals were removed for estimation of total viable organism burden as described above.

Biofilm scanning electron microscopy (SEM). Catheters were harvested, gently flushed with 0.15 M NaCl, and transected. Segments were then placed in fixative (4% formaldehyde, 1% glutaraldehyde in PBS) for 20 h, washed for 5 min in PBS, and placed in 1% osmium tetroxide for 30 min. After a series of alcohol washes, final drying was performed by critical point drying. Catheter segments were mounted and gold coated. Samples were imaged in a scanning electron microscope (JEOL JSM-6100) in the high-vacuum mode at 10 kV. The images were assembled using Adobe Photoshop 7.0.1. Images at x50 were examined to assess biofilm coverage of the entire catheter lumen for each catheter segment. Images at x1,000 were used to discern biofilm cellularity, amount of matrix, and biofilm thickness.

Biofilm cell TEM. The cell wall architecture of C. albicans cells isolated from the in vivo venous catheter model was examined and compared to cells grown in stationary phase in suspension using transmission electron microscopy (TEM). The planktonic cells were grown at 37°C with shaking at 250 rpm in RPMI-MOPS. Cells from the in vivo model were collected after 48 h of growth. Following collection, cells were fixed in 4% formaldehyde and 2% glutaraldehyde, postfixed with 1% osmium tetroxide and 1% potassium ferricyanide, stained with 1% uranyl acetate, dehydrated in a graded series of ethanol solutions, and embedded in Spurr's resin. Section (70 nm) were cut and placed on copper grids, poststained with 8% uranyl acetate in 50% methanol and Reynolds' lead citrate, and analyzed with a transmission electron microscope (Philips CM 120). Cell wall thickness was assessed for 50 cells from the biofilm and planktonic models using AnalySIS Docu software.

Cell wall isolation. Biofilm cells from the silicone disk assay described above and planktonic cells were collected for cell wall carbohydrate composition analysis as previously described (18, 19, 40). Briefly, biofilm cells were grown until a mature biofilm had formed (24 h, confirmed with microscopy). Planktonic cells were collected in both the log (6 h) and stationary (24 h) phases of growth. Both cultures used RPMI-MOPS and an incubation temperature of 37°C on an orbital shaker at 250 rpm. Washed cells were broken with glass beads, and cell walls were isolated as previously described (18, 19, 27, 32, 55). Cells (5 mg [dry cell weight] each) were washed twice with PBS, resuspended in PBS, and broken with glass beads five times for 1 min each. The cell walls were harvested by centrifugation and washed 10 times with buffer. Cell walls were further extracted by boiling in 0.5 ml sodium dodecyl sulfate buffer (50 mM Tris-HCl, pH 8.0, 0.1 M EDTA, 2% sodium dodecyl sulfate, 10 mM dithiothreitol) for 10 min. The cell walls were then washed 10 times with 0.5 ml ice-cold 0.1 M sodium acetate, pH 5.5, with 1 mM phenylmethylsulfonyl fluoride, and then five times with 0.5 ml ice-cold water (38, 51, 55).

Component fractionation and carbohydrate composition analysis. Cell wall components were fractionated by a series of chemical and enzymatic digestions, as previously described (19). Isolated cell walls (5 mg, dry cell weight) were alkali extracted for 60 min with 500 µl 0.7 M NaOH at 75°C. The alkali-soluble supernatant was collected by centrifugation, and alkali extraction was repeated on the pellet two more times. The alkali-soluble supernatants were neutralized with 250 µl glacial acetic acid. The alkali-insoluble fraction was neutralized with an initial 500 µl 0.1 M Tris-HCl (pH 7.5) wash, followed by a 500-µl 10 mM Tris-HCl (pH 7.5) wash and was then resuspended in 1 ml 10 mM Tris-HCl (pH 7.5) with 0.01% sodium azide. The alkali-insoluble pellet was then digested with glucanase (100 units zymolyase 20T; Arthrobacter luteus) at 37°C with gentle shaking for 16 h. The zymolyase-insoluble fraction was collected by centrifugation. The soluble supernatant was further treated with 1 unit chitinase (Streptomyces griseus; 713 units/g; Sigma) and incubated for 16 h at ambient temperature. One-half of the glucanase-soluble fraction (both ß-1,3 and ß-1,6 glucans) was dialyzed (Slide-A-Lyzer dialysis cassette; 7,000 molecular weight cutoff; Pierce) three times against 200 ml water to yield a ß-1,6 glucan fraction. The ß-1,3 glucan was calculated as the difference between total glucanase-soluble glucan and ß-1,6 glucan.

Carbohydrate concentrations of each fraction were measured as hexoses by the phenol-sulfuric method as previously described (18, 22, 40). Briefly, 2.5 µl phenol (80%) and 250 µl concentrated sulfuric acid were added to 100 µl of each cell wall fraction. Following incubation at 30°C for 30 min, color was detected by an automated plate reader at 490 nm. Carbohydrate concentration was determined using glucose controls (37.5, 75, 150, and 300 µg/ml). Two biologic and three assay replicates were performed for each cell population. The statistical significance of any differences observed among biofilm and the two planktonic growth environments was determined using analysis of variance.

Secreted ß-1,3 glucan measurement from biofilm and planktonic cell growth in vitro and in vivo. Supernatants from C. albicans in vitro biofilm and planktonic cells were collected, and ß-1,3 glucan concentrations were compared using a limulus lysate assay (1, 39). Cultures were grown on silicone disks or in suspension as described above. Viable count growth curves were conducted to assure the cultures contained a similar number of cells and were in a similar phase of growth. Culture supernatants were collected from both the planktonic and biofilm systems at 12 h for glucan measurement. The supernatants were centrifuged at 3,000 x g for 10 min, and the supernatants were removed and stored at –20°C until glucan analysis. Glucan concentrations were determined using the Glucatell (1,3)-ß-D-glucan detection reagent kit (Associates of Cape Cod, Cape Cod, MA) per the manufacturer's directions.

The catheter biofilm and disseminated candidiasis models were used as described above. After 12 h of growth in these models, serum was collected from the tail vein in both models and from the venous catheter in the biofilm model. Rat kidneys were then removed and processed for viable counts as an estimate of total body Candida burden. ß-1,3 Glucan was measured in serum using the Fungitell (1,3)-ß-D-glucan detection reagent kit (Associates of Cape Cod) per the manufacturer's directions (43). The statistical significance of observed differences between measurements from biofilm and planktonic or biofilm and disseminated conditions was determined using Student's t test.

Ability of cell walls and culture supernatant to bind fluconazole. In vitro C. albicans biofilm and planktonic (stationary-phase) cells were grown, and cell walls were isolated as described above. Cell walls (3 mg, dry cell weight) from each condition were incubated with 100 µg/ml fluconazole (1-ml volume) for 1 h at ambient temperature. We then separated bound from unbound fluconazole using size exclusion dialysis cassettes (7,000 molecular weight cutoff) for 2 h against 300 ml saline. We measured the fluconazole remaining in the dialysis cassette with a previously described bioassay using Candida kefyr as the test organism (53). The lower limit of detection of the assay was 0.12 µg/ml. The interday assay variation was less than 5%. Zones of inhibition were measured and compared to fluconazole standards.

Supernatants were collected from in vitro biofilm and planktonic cultures in stationary phase as described above. Dry cell weights of cultures were measured to ensure that the cultures contained similar numbers of cells. Samples (6 ml) were concentrated to 1 ml. The concentrated supernatants were then incubated with fluconazole, and the fluconazole bound to supernatant components was separated and measured by bioassay as described above.

Biofilm matrix material was harvested from in vitro biofilms as previously described (2, 36). Briefly, biofilm cells from the silicone disk assay were placed in 10 ml sterile water and sonicated for 10 min. The cells were then removed from the supernatant containing the matrix by two centrifugations at 4,500 x g for 20 min. A planktonic culture in stationary phase was similarly processed to serve as a control. The matrix was then incubated with fluconazole, and the bound compound was isolated and measured as described above. The statistical significance of any observed differences between samples was examined using Student's t test.

Impact of antifungal and cell wall-modifying enzyme exposures on in vitro and in vivo biofilm viability. In vitro biofilms using the 96-well polystyrene substrate were used as described above (49). Mature (24-h) biofilms were used for all experiments. The cell wall-modifying enzymes included three ß-glucanase preparations, zymolyase 20T from A. luteus, lysing enzyme from Rhizoctonia solani, and lysing enzyme from Trichoderma harzianum. Eight twofold compound dilutions were examined. The highest concentrations of each enzyme were as follows: zymolyase 20T, 5 units/ml; Rhizoctonia lysing enzyme, 5 units/ml; Trichoderma lysing enzyme, 20 units/ml. Because the enzyme preparations contain small amounts of other enzyme activities, we examined the impact of these components, mannanase and protease, alone on biofilm cell viability. The concentration range for each enzyme (mannanase, 0 to 50 units/ml; proteinase K, 0 to 2.5 units/ml) included the concentrations above and below that measured in the zymolyase preparation. The effect of each of these enzymes was examined at 48 h after incubation at 37°C with RPMI-MOPS, buffered at pH 7. Zymolyase, heat inactivated by 100°C for 10 min, served as a control.

The impacts of the antifungals drugs, fluconazole and amphotericin B, on biofilm growth were similarly examined over concentration ranges of 0.0625 to 1,000 µg/ml and 0.00975 to 0.156 µg/ml, respectively. Each of the antifungals was examined alone and in combination with zymolyase in a 96-well checkerboard format. For each drug exposure, mature biofilms were incubated in the presence of drug and zymolyase at 37°C for 48 h. Following exposure, the drug and enzyme were removed and wells were washed with PBS three times. Biofilm cell viability was then assessed using the XTT assay described above. We determined the drug concentration associated with both a 50% effective dose (ED50) and ED80 reduction in optical density compared to the no-drug control wells. Four replicates were performed for each drug and each drug combination. The impact of the compounds in combination was assessed using the fractional inhibitory concentration (FIC) index {FIC index = [(ED50 or ED80 of drug A in combination)/(ED50 or ED80 of drug A alone)] + [(ED50 or ED80 of drug B in combination)/(ED50 or ED80 drug B alone)]; values below 0.5 are suggestive of an enhanced interaction } (5, 52). For a planktonic control, cells were grown in RPMI-MOPS at 37°C to log phase. The cells were exposed to the same enzymes and concentrations. Following a 24-h exposure, cells were collected for microbiologic enumeration by plating serial dilutions on Saboraud's dextrose agar plates.

A mature biofilm from the catheter model was used for in vivo studies. The impacts of intraluminal zymolyase and fluconazole exposures alone and in combination were examined. A single fluconazole concentration was used and was chosen based on the highest concentration used in the in vitro studies (1,000 µg/ml). Two zymolyase concentrations (2.5 units/ml and 1.25 units/ml) were chosen in order to include the lowest effective concentration from the in vitro study and a twofold-lower concentration that was ineffective in this study. Both zymolyase concentrations and fluconazole were examined separately and in combination. Following a 24-h exposure, the catheters were removed and processed for SEM to discern the extent of remaining biofilm on the catheter surface.

Impact of biofilm-secreted material and exogenous ß-1,3 glucan on fluconazole activity against planktonic cells. A modification of the CLSI M27 A microbroth susceptibility method was used to examine the activity of fluconazole against planktonic Candida in the presence of biofilm matrix material or commercial ß-1,3 glucan (laminarin) (16). C. albicans strain K1 was used for these experiments. The fluconazole concentrations included in these studies ranged from 0.125 µg/ml to 128 µg/ml. The in vitro silicone disk assay was used to collect biofilm material. Biofilm matrix was collected from mature biofilms as described above. The matrix material was concentrated from 10 ml to 1 ml by vacuum centrifugation. Microscopy was used to ensure the biofilm materials were cell free. The biofilm-secreted materials were added to the 96-well plates in addition to drug, medium, and C. albicans K1 inoculum. In addition to the standard CLSI turbidity endpoint, quantitative cultures of each well were undertaken. These experiments were undertaken on two occasions, and the assays were performed in duplicate. Results were compared to standard fluconazole susceptibility testing performed at the same time.

A similar susceptibility experiment was performed using exogenous ß-1,3 glucan (laminarin; Sigma). A concentration of 0.25 mg/ml of laminarin was used. The concentration of laminarin was chosen based on the carbohydrate concentrations from matrix examination experiments described above. Similar endpoints and comparisons were performed as described above. The significance of differences among the experiments was assessed using analysis of variance.


arrow
RESULTS
 
Cell wall ultrastructure of in vivo biofilm and planktonic C. albicans. TEM was used to screen for ultrastructural differences between the cell wall of biofilm and planktonic cells. Yeast cells from both conditions were of similar diameter. Imaging of the cell walls from the two conditions, however, demonstrated quantitative and qualitative differences. The cell walls of the in vivo biofilm cells were up to two times thicker than planktonic cells (Fig. 1). Cell walls from both groups were layered in appearance, with an outer electron-dense component and an inner electron-lucent layer. However, there were apparent ultrastructural differences between the two cell types. The electron-dense material was more prominent and fibrillar in appearance in the biofilm cells. The more electron-lucent component was scattered, with electron-dense granules. In addition, the periplasmic layer was more prominent in the biofilm cells. These images suggest that biofilm growth induces dramatic changes in the cell wall architecture of C. albicans.


Figure 1
View larger version (27K):
[in this window]
[in a new window]

  		FIG. 1. Transmission electron microscopy of biofilm (right) and planktonic (left) cell walls. The images are representative of cell walls examined in more than 10 fields (50 cells from each condition). Brackets indicate cell walls. The magnification for the two images is x110,000.

Cell wall carbohydrate composition of in vitro biofilm compared to planktonic C. albicans cells. To investigate cell wall content associated with the microscopy findings, biofilm cells walls were fractionated by chemical and enzymatic methods. In vitro biofilm cells, grown on medical-grade silicone in RPMI-MOPS, were compared to planktonic cells in stationary and log phases. The cell walls from biofilm cells were found to contain significantly more total carbohydrate and ß-1,3 glucan compared to stationary- or log-phase planktonic cells (P < 0.001) (Fig. 2). These data suggest that the cell wall imaging changes using TEM may, in part, be explained by quantitative differences in cell wall glucan.


Figure 2
View larger version (15K):
[in this window]
[in a new window]

  		FIG. 2. Biofilm and planktonic cell wall carbohydrate composition. Biofilm cells were collected from the in vitro silicone disk assay. Planktonic cells were collected in both log and stationary phases of growth. Each bar color represents a different cell wall fraction, with means and standard deviations from three biologic and three technical assay replicates. The data are expressed as micrograms of carbohydrate per 5 mg of dry cell weight. Carbohydrate concentrations of various fractions were estimated using the phenol-sulfuric acid methods with a glucose standard curve. *, biofilm cells contained significantly more ß-1,3 glucan compared to planktonic cells in both stationary and log phases (P < 0.001).

Comparison of extracellular ß-1,3 glucan from biofilm and planktonic C. albicans. Given the greater concentration ß-1,3 glucan observed in the cell wall of biofilm cells, we reasoned that the unidentified matrix carbohydrate could be a ß-glucan molecule. To test this hypothesis, we compared supernatants of C. albicans in vitro biofilm cultures and planktonic cultures. Viable count growth curves were conducted to ensure that the cultures contained a similar number of cells and were in a similar phase of growth. At 12 h, both the planktonic culture and the biofilm culture (combination of biofilm and free-floating cells) contained 105 CFU/ml. Using a limulus lysate-based assay, ß-1,3 glucan was detected in the supernatants of both biofilm and planktonic cultures. However, biofilm culture supernatants were found to contain significantly higher concentrations of ß-1,3 glucan than planktonic culture supernatants (P < 0.001) (Fig. 3). The supernatants from the biofilm system contained at least twofold more ß-1,3 glucan.


Figure 3
View larger version (10K):
[in this window]
[in a new window]

  		FIG. 3. Extracellular ß-1,3 glucan concentrations from C. albicans biofilm and planktonic C. albicans in vitro and in vivo. The silicone disk assay was used for in vitro biofilm formation and for matrix isolation. The extracellular sample from the in vitro model represents cell-free culture supernatant. The rat central venous catheter model was used for in vivo biofilm formation. Biofilm formation was confirmed with microscopy. A disseminated candidiasis model was used as a control. All samples were collected after 12 h of biofilm formation. Viable cell counts were determined by plate counts and were similar for the biofilm and planktonic cultures at the time of sample collection. Each bar represents the mean and standard deviation from two biologic and three technical replicates. The black bars represent data from biofilm cultures, and the gray bars represent data from planktonic cultures. ß-1,3 Glucan content was measured using the limulus lysate assay. Data are expressed as picograms of ß-1,3 glucan per milliliter of sample. Both biofilm systems contained significantly more ß-1,3 glucan compared to planktonic conditions (P < 0.001).

We used a matrix isolation protocol and assayed ß-1,3 glucan content as described above. A planktonic cell culture was similarly processed to serve as a control group. As shown in Fig. 3, ß-1,3 glucan was also detected in the isolated matrix material.

Our next experiment was intended to determine if the enhanced release of ß-1,3 glucan from biofilm cells observed in vitro occurs in vivo. We measured ß-1,3 glucan similarly using the limulus lysate assay in serum from animals with an in vivo biofilm catheter infection or disseminated candidiasis. We used viable kidney counts as an estimate of total body Candida burden, since it is widely accepted as the most significantly impacted end organ in disseminated models (7, 42). Furthermore, our previous examination of the kidney in the current biofilm model demonstrated marked dissemination to this organ (6). Over the time course of growth examined in both models, the burden of viable organisms was significantly higher in the disseminated candidiasis model. At 12 h, the kidneys of animals with a disseminated infection contained 104 CFU/kidney, while those with a catheter infection only contained 102 CFU/kidney. Despite the in vivo presence of fewer cells in the biofilm model, the sera from the biofilm catheter infections contained nearly 10-fold more ß-1,3 glucan than serum from rats with disseminated candidiasis at the 12-h time point (P < 0.001). Not only did serum obtained from catheter infection animals have more ß-glucan (259 pg/ml), but serum obtained from the tail vein in rats with biofilm infections also had a higher concentration of glucan (248 pg/ml). These results corroborate findings from the in vitro biofilm model.

Biofilm component antifungal drug binding assay. Previous studies in a bacterial biofilm model demonstrated sequestration of antimicrobial compounds by a cell wall component, specifically, the ß-1,3 glucan component of biofilm cells (33, 34). The objective of our next line of experimentation was to determine if a similar interaction may occur in C. albicans biofilms. We exploited the relatively large molecular weight of the cell wall components and the secreted ß-1,3 glucans relative to the antifungal fluconazole in a size exclusion experimental design. The cell wall from both biofilm and planktonic cells bound a measurable concentration of fluconazole (Fig. 4). However, the biofilm cell walls bound four- to fivefold more compound per cell wall weight (P < 0.001). Similar examination of culture supernatants from both growth conditions identified binding with only the biofilm culture. The cell burdens from both cultures were similar based on dry cell weight (biofilm, 21.1 mg; planktonic, 24.7 mg). Finally, we utilized a biofilm matrix isolated by the method described above in the binding assay. Fluconazole binding was observed at a concentration near that found from biofilm culture supernatant.


Figure 4
View larger version (9K):
[in this window]
[in a new window]

  		FIG. 4. Fluconazole binding to biofilm and planktonic cell wall, culture supernatant, and biofilm matrix. The silicone disk assay was used for biofilm formation and for matrix isolation. Planktonic cells were collected in the log phase of growth. All samples were collected after 24 h of biofilm formation. For the cell wall study, 3 mg of dry cell wall weight from each condition was used. Viable cell counts were determined by plate counts and were similar for the biofilm and planktonic cultures at the time of supernatant collection. Following incubation of the culture component with fluconazole, unbound drug was separated using size exclusion dialysis. Fluconazole concentration was measured using a microbiologic assay. Each data value represents the mean and standard deviation from two biologic and three assay replicates. The black vertical bars represent data from the biofilm culture, and the gray bar represents data from the planktonic culture. Biofilm cell walls, supernatant, and matrix were associated with increased fluconazole binding compared to respective planktonic controls (P < 0.001).

Effect of glucanase on Candida biofilms in vitro. We next examined the impact of enzymatic modification of the cell wall component changes observed in biofilm cells on their viability in mature C. albicans biofilms. To test this, we first compared in vitro biofilm cell viability to planktonic cell viability in the presence of various concentrations of zymolyase, a ß-1,3 glucanase. The highest concentration of ß-1,3 glucanase used in these studies, 5 units/ml, was chosen to be more than fivefold lower than that used in cell lysis protocols (4). We observed a concentration-dependent reduction in biofilm cell viability over the 512-fold range of concentrations (Fig. 5). The same drug exposure did not impact planktonic cell viability even at the highest concentration examined. Heat-inactivated zymolyase had no effect on biofilm growth (data not shown).


Figure 5
View larger version (17K):
[in this window]
[in a new window]

  		FIG. 5. Impact of glucanase on biofilm cell viability, determined using three ß-glucanase preparations and two additional components (proteinase K and mannanase) of the predominant ß-glucanase preparation (zymolyase). The 96-well polystyrene in vitro biofilm model was used. Cell viability was assessed using the XTT reduction assay. Viability is expressed as the optical density (OD) at 490 nm. The assays were performed in triplicate, and results are expressed as the mean and standard deviation. All drug exposures were of 48-h duration. (A) Biofilm cell viability following exposure to ß-glucanase from Rhizoctonia over a concentration range from 0 to 5 units/ml. (B) Viability following exposure to ß-1,3 glucanase from Arthrobacter luteus (zymolyase) over a concentration range from 0 to 5 units/ml. (C) Cell viability following exposure to ß-1,3 glucanase from Tricoderma over a concentration range from 0 to 20 units/ml. (D) Biofilm cell viability following exposure to mannanase over a concentration range from 0 to 50 units/ml. (E) Viability following exposure to proteinase K over a concentration range from 0 to 2.5 units/ml.

In an attempt to discern if the activity of zymolyase was due to the ß-1,3 glucanase component, we examined the activity of two additional ß-glucanase preparations from distinct microbial sources. ß-Glucanases from Rhizoctonia and Trichoderma species also inhibited biofilm growth at concentrations similar to zymolyase, suggesting that the biofilm inhibitory effects are likely due to ß-1,3 glucanase activity. To ensure that the glucanase component was responsible for the difference seen, we examined the activity of the other available components of the zymolyase preparation, mannanase and protease. Mannanase and proteinase K did not impact biofilm or planktonic cell viability, even at concentrations exceeding those present in the glucanase preparations (Fig. 5).

We next considered the possibility that ß-glucan modification may impact the effect of traditional antifungals on biofilm cells. We chose to examine fluconazole and amphotericin B, as numerous prior investigations and the current study have demonstrated drug resistance in C. albicans biofilm cells. We exposed mature C. albicans biofilms to various concentrations of zymolyase and antifungal alone and in combination. Similar to previous reports, fluconazole exposures at concentrations as high as 1,000 µg/ml had no impact on biofilm cell viability. (The planktonic MICs for C. albicans K1 are 0.5 µg/ml and 0.25 µg/ml for fluconazole and amphotericin B, respectively [2, 6, 7, 9, 11, 15, 25, 30, 31, 47].) Amphotericin B did reduce viable cell burden, however, only at concentrations considerably higher than those associated with the killing of planktonic cells. Over the fluconazole (0.0625 to 1,000 µg/ml) and ß-1,3 glucanase (0.0195 to 5 units/ml) concentrations studied, the FIC indices suggested a synergistic interaction. The FIC indices for fluconazole and ß-1,3 glucanase associated with a 50% (ED50) and 80% (ED80) reduction in optical density were 0.31 ± 0.06 and 0.10 ± 0.04, respectively. Similarly, the FIC indices for the amphotericin B and ß-1,3 glucanase combination were also low, 0.20 ± 0.12 and 0.21 ± 0.09, respectively. The potentiation of fluconazole and amphotericin B activity was notably observed at glucanase concentrations which had no impact on biofilm cell viability.

We used the rat catheter model to determine if these in vitro findings were meaningful in vivo. Initial studies examined the impact of two concentrations of ß-1,3 glucanase alone. The highest concentration, 2.5 units/ml, was chosen based on efficacy in the in vitro studies. The second concentration was twofold lower, 1.25 units/ml. ß-1,3 Glucanase at 2.5 units/ml essentially eliminated the biofilm process. The lower concentration, 1.25 units/ml, produced little if any discernible impact on the biofilm extent or architecture (Fig. 6). The high fluconazole concentration, 1,000 µg/ml (1,000 times the planktonic MIC) had no discernible impact on the extent of biofilm formation, similar to our previous findings in this model (6).However, when this fluconazole concentration was exposed to the mature in vivo biofilm in the presence of the lower concentration of ß-1,3 glucanase, the biofilm was eliminated (Fig. 6). These results are congruent with findings from the in vitro C. albicans model.


Figure 6
View larger version (85K):
[in this window]
[in a new window]

  		FIG. 6. Impact of fluconazole and ß-1,3 glucanase alone and in combination as lock therapy against C. albicans biofilm cells in an in vivo catheter model. The SEM image perspective is from above the biofilm, imaging from the inside of the catheter lumen. The biofilms were grown for 24 h followed by instillation of the compound(s) into the lumen of the catheter for 24 h. Following compound exposure, the catheters were removed for SEM processing. For each of the panel groups (A, B, C, and D), the top row represents a x50 magnification, and the bottom row represents a x1,000 magnification. (A) Images are from a 48-h control biofilm with extensive biofilm formation. (B) Images are after a 1,000-µg/ml fluconazole exposure at 1,000 times the planktonic MIC. (C) Images are from catheters exposed to zymolyase at 1.25 units/ml alone. (D) Images are from a combination of zymolyase and fluconazole at 1.25 units/ml and 1000 µg/ml, respectively.

Impact of biofilm matrix and exogenous ß-1,3 glucan on fluconazole activity against planktonic cells. The final experiments were designed to determine if the biofilm-secreted material (matrix) could impact the activity of fluconazole against Candida growing in planktonic culture. Furthermore, since we observed markedly higher concentrations of extracellular ß-1,3 glucan from biofilm cells, we also examined the impact of adding exogenous commercial ß-1,3 glucan (laminarin) on fluconazole activity against planktonic cells. The planktonic CLSI MIC for fluconazole was 0.5 µg/ml. The MIC in the presence of biofilm material and laminarin was >128 µg/ml. Results of quantitative cultures are shown in Fig. 7. Quantitative cultures of the planktonic wells in the presence of fluconazole revealed growth in wells with visible turbidity as well as low levels of growth in the next three wells without visible turbidity (likely due to the previously described trailing growth associated with in vitro triazole exposures). However, all of the wells with either laminarin or the biofilm-secreted material demonstrated high levels of growth (P < 0.001). These quantitative culture results were congruent with the turbidity endpoint data.


Figure 7
View larger version (25K):
[in this window]
[in a new window]

  		FIG. 7. Impact of biofilm matrix and ß-1,3 glucan (laminarin) on susceptibility of planktonic C. albicans K1 to fluconazole. Each vertical bar represents the log10 CFU/ml following 48 h of incubation in the presence of twofold-escalating concentrations of fluconazole (range, 0.13 to 128 µg/ml) using the CLSI M27A methodology. Each value represents the mean from experiments performed on two separate days.


arrow
DISCUSSION
 
C. albicans is a major human pathogen which can grow as a biofilm and inhabit the surface of many medical devices (28, 44). Cells in a biofilm have altered phenotypes and are extraordinarily resistant to many antifungals, making eradication of this process difficult. C. albicans biofilm resistance is not entirely understood and likely involves a multifactorial mechanism. It has been suggested that resistance may be related to contributions from the extracellular matrix (preventing active drug diffusion), the physiological state of the cell (slow growth), activation of drug efflux pumps in the cell membrane, sterol perturbations, or differential gene expression patterns by biofilm cells (10, 37, 47).

A number of molecular mechanisms have been implicated in nonbiofilm antifungal resistance in C. albicans (54). For example, investigators have characterized several efflux pumps that are transcriptionally upregulated in cells resistant to triazole compounds. The relevance of these mechanisms for C. albicans biofilm cells has also been examined. Recent studies have demonstrated that genes for two types of efflux pump (CDR and MDR) are upregulated during biofilm formation (6, 37, 47). However, strains carrying single or double mutations of these efflux genes retained a resistant phenotype during biofilm growth, demonstrating that additional yet-unidentified resistance mechanisms are in play. In addition, both transcriptional upregulation and point mutations in the triazole target 14-{alpha} demethylase (ERG11) have been shown to play a role in triazole resistance in nonbiofilm cells. Investigations to determine the role of these mechanisms in biofilm cells have not found the ERG11 gene to be differentially expressed in biofilm cells. However, sterol changes have been associated with mature (late) biofilm growth and have been hypothesized to contribute to biofilm drug resistance (37). More in depth examination of the sterol synthesis pathway in relation to biofilm resistance has not been undertaken.

Several approaches have been used to examine the impact of the relatively slower growth rate in Candida biofilms (9, 11). Transition from exponential to slow growth is generally accompanied by an increase in antimicrobial resistance. Slow growth of microorganisms has been observed in mature biofilms, presumably due to some form of nutrient limitation, and has been suggested to account for biofilm resistance. By closely monitoring the growth phase of planktonic and biofilm C. albicans cells using nutrient limitation, studies have been able to examine the contribution of growth rate to biofilm cell survival against antifungals. In these studies, cell resistance to the polyene antifungal amphotericin B was identical for each of the growth rates studied, suggesting that some determinant other than growth rate is responsible for biofilm resistance.

Finally, a number of studies have experimentally addressed the contribution of the extracellular matrix in modulating biofilm cell susceptibility (2, 10, 26, 36). Results from these studies have been mixed, with initial lines of evidence suggesting a minimal role. However, more recent investigation utilizing a different experimental approach suggests the matrix may impact drug activity in Candida biofilms (2). The first studies examined the ability of various antifungal compounds to penetrate the matrix. For the most part, each of the drugs was able to readily diffuse through the entire thickness of the biofilm. The second approach examined the relationship between the extent of matrix and drug resistance. The matrix extent was experimentally controlled by varying the flow characteristics across the substrate surface. The cells were similarly susceptible to the antifungal amphotericin B in these initial studies, despite visible differences in the amount of matrix material. However, in more recent studies, these investigators altered the flow characteristics and demonstrated a relationship between matrix extent and amphotericin B activity in an in vitro model. Furthermore, the investigators identified a strain of Candida tropicalis with extensive matrix production and marked associated antifungal drug resistance. Despite these important and thorough studies, each of these investigators agrees that additional mechanisms likely contribute to the extensive resistance phenotype associated with growth in the Candida biofilm milieu.

The results of the current set of experiments suggest that changes in a cell wall and secreted carbohydrate (including those within the matrix material) are associated with Candida biofilm growth and contribute to the biofilm resistance phenotype. The cell walls of fungi, plants, and bacteria provide a number of essential functions, including acting as a permeability barrier, providing cell shape, and interacting with host cells (14, 29, 35). Approximately 80 to 90% of the cell wall of C. albicans is carbohydrate. The C. albicans cell wall is composed primarily of three components interconnected by covalent bonds: ß-1,3 and ß-1,6 glucans (50 to 60%), mannoproteins (30 to 40%), and chitin (0.6 to 9%). ß-1,3 glucans are also thought to be the main component of the three-dimensional matrix surrounding biofilm cells. ß-1,3 Glucans are synthesized by a plasma membrane-bound glucan synthase complex which uses UDP-glucose as a substrate and extrudes ß-1,3 glucan through the periplasmic space. The ß-1,3 glucan generated remains unorganized until it is covalently linked to other cell wall components. The cell wall is a dynamic structure constantly changing in response to environmental signals, and the data presented here suggest that biofilm growth signals cell wall changes. It is perhaps not surprising to find dramatic cell wall changes associated with the biofilm environment. These studies identified predominantly a change in the quantity of ß-1,3 glucan. It is possible the change in cell wall content is transient and associated with transfer of this glucan molecule to the extracellular portion of the biofilm. The finding of marked ß-1,3 glucan concentrations in the biofilm milieu and specifically as a component of the matrix supports this hypothesis. Prior study of Candida matrix composition has identified increased concentrations of glucose and an unidentified carbohydrate. We think it is likely that the unidentified molecule is a ß-1,3 glucan, as recently hypothesized by those authors as well (2).

The hypothesis that cell wall and specifically glucan changes may be associated with drug resistance is not novel. In the 1970s, studies suggested that glucan changes associated with the stationary phase of growth in C. albicans were responsible for reduced susceptibility to the polyene amphotericin B (13, 23). More recent study of glucan-associated proteins also suggests an association between glucan-associated protein content and fluconazole resistance in C. albicans planktonic cells (6). Interestingly, results from a Pseudomonas biofilm model demonstrated a role for periplasmic ß-glucans in modulation of biofilm-associated antibacterial resistance to several drug classes (33, 34). The studies suggest that the glucan molecules limit penetration of an antimicrobial into the cell cytoplasm.

The current studies suggest that cell wall changes involving ß-1,3 glucan in C. albicans biofilm cells may contribute to antifungal drug resistance in this important environment. First, the experiments identified a cell wall and matrix ß-glucan change associated with biofilm growth of Candida albicans in vitro and in vivo. Biofilm cell walls contain significantly higher concentrations of ß-1,3 glucan than their planktonic counterparts, and these glucans can be found in the supernatant surrounding the biofilm and in the matrix. Second, enzymatic manipulation of the glucans at high concentrations destroys biofilms but has virtually no effect on planktonic cells under the same enzymatic conditions. The impact of ß-1,3 glucanase on biofilm growth is similar to that recently observed by Al-Fattani et al. using a different C. albicans strain and biofilm model (2). The enzyme concentration used by this group was twofold greater than the highest concentration examined in the current investigation. At lower concentrations, modification of ß-1,3 glucan did not affect biofilm cell viability. However, the addition of these lower glucanase concentrations markedly enhanced the activities of two antifungal drugs, fluconazole and amphotericin B, against biofilm cells. These results suggest that a cell wall component, such as glucan, may physically interact with the antifungal and inhibit penetration to the site of action. Another possibility is that the cell wall changes in biofilm cells structurally protect the cell from external insults, including antifungals. For example, the fungal cell wall has been shown to provide a critical defense against osmotic and oxidative attack. While the fluconazole binding assay results do not confirm a specific binding interaction, the data do support a physical interaction or sequestration of the antifungal.

The current lines of evidence provide a strong rationale for further genetic and biochemical investigation of the mechanistic role of ß-glucan molecules in the drug-resistant phenotype of biofilm cells. Further understanding of this process is likely to identify potential targets for this important infection environment.


arrow
FOOTNOTES
 
* Corresponding author. Mailing address: 600 Highland Ave., Room H4.572, Madison, WI 53792. Phone: (608) 263-1545. Fax: (608) 263-4464. E-mail: dra{at}medicine.wisc.edu. Back

{triangledown} Published ahead of print on 27 November 2006. Back


arrow
REFERENCES
 
    1
  1. Aketagawa, J., S. Tanaka, H. Tamura, Y. Shibata, and H. Saito. 1993. Activation of limulus coagulation factor G by several (1->3)-beta-D-glucans: comparison of the potency of glucans with identical degree of polymerization but different conformations. J. Biochem. 113:683-686.[Abstract/Free Full Text]
    2. 2
  2. Al-Fattani, M. A., and L. J. Douglas. 2006. Biofilm matrix of Candida albicans and Candida tropicalis: chemical composition and role in drug resistance. J. Med. Microbiol. 55:999-1008.[Abstract/Free Full Text]
    3. 3
  3. Al-Fattani, A. M., and L. J. Douglas. 2004. Penetration of Candida biofilms by antifungal agents. Antimicrob Agents Chemother. 48:3291-3297.[Abstract/Free Full Text]
    4. 4
  4. Amberg, D. C., D. J. Burke, J. N. Strathern, and D. Burke. 2005. Methods in yeast genetics: a Cold Spring Harbor Laboratory course manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
    5. 5
  5. American Society for Microbiology. 2006. Antimicrobial Agents and Chemotherapy 2006 instructions to authors. Antimicrob. Agents Chemother. 50:1-21.[Free Full Text]
    6. 6
  6. Andes, D., J. Nett, P. Oshel, R. Albrecht, K. Marchillo, and A. Pitula. 2004. Development and characterization of an in vivo central venous catheter Candida albicans biofilm model. Infect. Immun. 72:6023-6031.[Abstract/Free Full Text]
    7. 7
  7. Andes, D. 2005. Use of an animal model of disseminated candidiasis in the evaluation of antifungal therapy. Methods Mol. Med. 118:111-128.[Medline]
    8. 8
  8. Angiolella, L., M. M. Micoci, S. D'Alessio, A. Girolamo, B. Maras, and A. Cassone. 2002. Identification of major glucan-associated cell wall proteins of C. albicans and their role in fluconazole resistance. Antimicrob Agents Chemother. 46:1688-1694.[Abstract/Free Full Text]
    9. 9
  9. Baillie, G. S., and L. J Douglas. 1998. Effect of growth rate on resistance of Candida albicans biofilms to antifungal agents. Antimicrob Agents Chemother. 42:1900-1905.[Abstract/Free Full Text]
    10. 10
  10. Baillie, G. S., and L. J. Douglas. 2000. Matrix polymers of Candida biofilms and their possible role in biofilm resistance to antifungal drugs. J Antimicrob Chemother. 46:397-403.[Abstract/Free Full Text]
    11. 11
  11. Baillie, G. S., and L. J. Douglas. 1998. Iron-limited biofilms of Candida albicans and their susceptibility to amphotericin B. Antimicrob Agents Chemother. 42:2146-2149.[Abstract/Free Full Text]
    12. 12
  12. Baillie, G. S., and L. J. Douglas. 1999. Candida biofilms and their susceptibility to antifungal agents. Methods Enzymol 310:644-656.[Medline]
    13. 13
  13. Cassone, A., D. Kerridge, and E. F. Gale. 1979. Ultrastructural changes in the wall of Candida albicans following cessation of growth and their possible relationship to the development of drug resistance. J Gen. Microbiol. 110:339.[Abstract/Free Full Text]
    14. 14
  14. Chaffin, W. L. J., J. L. Lopez-Ribot, M. Casanova, D. Gozalbo, and J. P. Martinez. 1998. Cell wall and secreted proteins of Candida albicans: identification, function, and expression. Microbiol. Mol. Biol Rev. 62:130-180.[Abstract/Free Full Text]
    15. 15
  15. Chandra, J., D. M. Kuhn, P. K. Mukherjee, L. L. Hoyer, T. McCormick, and M. A. Ghannoum. 2001. Biofilm formation by the fungal pathogen Candida albicans: development, architecture, and drug resistance. J. Bacteriol. 183:5385-5394.[Abstract/Free Full Text]
    16. 16
  16. Clinical and Laboratory Standards Institute (formly NCCLS). 1997. Reference method for broth dilution antifungal susceptibility testing of yeasts; approved standard. M27A, Wayne, PA.
    17. 17
  17. Costerton, J. W., P. S. Stewart, and E. P. Greenberg. 1999. Bacterial biofilms: a common cause of persistent infections. Science 284:1318-1322.[Abstract/Free Full Text]
    18. 18
  18. Dallies, N., J. Fancois, and V. Paquet. 1998. A new method for quantitative determination of polysaccharides in the yeast cell wall. Application to the cell wall defective mutants of Saccharomyces cerevisiae. Yeast. 14:1297-1306.[CrossRef][Medline]
    19. 19
  19. Dijkgaraff, G. J. P., J. L. Brown, and H. Bussey. 1996. The KHN1 gene of Saccharomyces cerevisiae is a functional homolog of KRE9. Yeast. 12:683-692.[CrossRef][Medline]
    20. 20
  20. Donlan, R. M., and J. W. Costerton. 2002. Biofilms: survival mechanisms of clinically relevant microorganisms. Clin. Microbiol. Rev. 15:167-193.[Abstract/Free Full Text]
    21. 21
  21. Douglas, L. J. 2003. Candida biofilms and their role in infection. Trends Microbiol. 11:30-36.[CrossRef][Medline]
    22. 22
  22. Dubois, M., K. A. Gilles, J. K. Hamilton, P. A. Rebers, and F. Smith. 1956. Colorometric method for determination of sugars and related substances. Anal. Chem. 28:350-356.[CrossRef]
    23. 23
  23. Gale, E. F., J. Ingram, D. Kerridge, V. Notario, and F. Wayman. 1980. Reduction of amphotericin B resistance in stationary phase cultures of Candida albicans by treatment with enzymes. J Gen. Microbiol. 117:383-391.[Abstract/Free Full Text]
    24. 24
  24. Hawser, S. P., and L. J. Douglas. 1994. Biofilm formation by Candida species on the surface of catheter materials in vitro. Infect. Immun. 62:915-921.[Abstract/Free Full Text]
    25. 25
  25. Hawser, S. P., and L. J. Douglas. 1995. Resistance of Candida albicans biofilms to antifungal agents in vitro. Antimicrob Agents Chemother. 39:2128-2131.[Abstract]
    26. 26
  26. Hawser, S. P., G. S. Baillie, and L. J. Douglas. 1998. Production of extracellular matrix by Candida albicans biofilms. J. Med. Microbiol. 47:253-256.[Abstract/Free Full Text]
    27. 27
  27. Herrero, A. B., P Magnelli, M. K. Mansour, S. M. Levitz, H. Bussey, and C. Abeijon. 2004. KRE5 gene null mutant strains of Candida albicans are avirulent and have altered cell wall composition and hypha formation properties. Eukaryot Cell. 3:1423-1431.[Abstract/Free Full Text]
    28. 28
  28. Kojic, E. M., and R. O. Darouiche. 2004. Candida infections of medical devices. Clin. Microbiol. Rev. 17:255-267.[Abstract/Free Full Text]
    29. 29
  29. Kollar, R., B. B. Reinhold, E. Petrakova, H. J. C. Yeh, G. Ashwell, J. Drgonava, J. C. Kapteyn, F. M. Klis, and E. Cabib. 1997. Architecture of the yeast cell wall. J. Biol. Chem. 272:17762-17775.[Abstract/Free Full Text]
    30. 30
  30. Kuhn, D. M., and M. A. Ghannoum. 2004. Candida biofilms: antifungal resistance and emerging therapeutic options. Curr. Opin. Investig Drugs 5:186-197.[Medline]
    31. 31
  31. Lewis, R. E., D. P. Kontoyiannis, R. O. Darouiche, and I. I. Raad, R. A. Prince. 2002. Antifungal activity of amphotericin B, fluconazole, and voriconazole in an in vitro model of Candida catheter-related bloodstream infection. Antimicrob Agents Chemother. 46:3499-3505.[Abstract/Free Full Text]
    32. 32
  32. Magnelli, P., J. F. Cipollo, and C. Abeijon. 2002. A refined method for the determination of Saccharomyces cerevisiae cell wall composition and ß-1,6-glucan fine structure. Anal. Biochem. 301:136-150.[CrossRef][Medline]
    33. 33
  33. Mah, T. F., B. Pitts, B. Pellock, G. C. Walker, P. S. Stewart, and G. A. O'Toole. 2003. A genetic basis for Pseudomonas aeruginosa biofilm antibiotic resistance. Nature 426:306-310.[CrossRef][Medline]
    34. 34
  34. Mah, T. F., and G. A. O'Toole. 2001. Mechanisms of biofilm resistance to antimicrobial agents. Trends in Microbiol. 9:34-38.[CrossRef][Medline]
    35. 35
  35. Masuoka, J. 2004. Surface glycans of Candida albicans and other pathogenic fungi: physiological roles, clinical uses, and experimental challenges. Clin. Microbiol. Rev. 17:281-310.[Abstract/Free Full Text]
    36. 36
  36. McCourtie, J., and L. J. Douglas. 1985. Extracellular polymer of Candida albicans: isolation, analysis and role in adhesion. J Gen. Microbiol. 131:495-503.[Abstract/Free Full Text]
    37. 37
  37. Mukherjee, P. K., J. Chandra, D. M. Kuhn, and M. A. Ghannoum. 2003. Mechanism of fluconazole resistance in Candida albicans biofilms: phase-specific role of efflux pumps and membrane sterols. Infect. Immun. 71:4333-4340.[Abstract/Free Full Text]
    38. 38
  38. Munro, C. A., S. Bates, E. T. Buurman, H. B. Hughes, D. M. MacCallum, G. Bertram, A. Atrih, M. A. J. Ferguson, J. M. Bain, A. Brand, S. Hamilton, C. Westerwater, L. M. Thomson, A. J. P. Brown, F. C. Odds, and N. A. R. Gow. 2005. Mnt1 and Mnt2p of Candida albicans are partially redundant {alpha}-1,2-mannosylation and are required for adhesion and virulence. J. Biol. Chem. 280:1051-1060.[Abstract/Free Full Text]
    39. 39
  39. Muta, T., N. Seki, Y. Takaki, R. Hashimoto, T. Oda, A. Iwanaga, F. Tokunaga, and S. Iwanaga. 1995. Purified horseshoe crab factor G: reconstitution and characterization of the (1,3) B-D-glucan sensitive protease cascade. J. Biol. Chem. 270:892-897.[Abstract/Free Full Text]
    40. 40
  40. Nemecek, J. C., M. Wuthrich, and B. S. Klein. 2006. Global control of dimorphism and virulence in fungi. Science 312:583-588.[Abstract/Free Full Text]
    41. 41
  41. Nett, J., and D. Andes. 2006. Candida albicans biofilm development, modeling of a host pathogen interaction. Curr. Opin. Microbiol. 9:340-345.[CrossRef][Medline]
    42. 42
  42. Odds, F. C., L. Van Nuffel, and N. Gow. 2000. Survival in experimental Candida albicans infections depends on inoculum growth conditions as well as animal host. Microbiology 146:1881-1889.[Abstract/Free Full Text]
    43. 43
  43. Ostrosky-Zeichner, L., B. D. Alexander, D. H. Kett, J. Vazquez, P. G. Pappas, F. Saeki, F., P. A. Ketchum, J. Wingard, R. Schiff, H. Tamura, M. A. Finkelman, and J. Rex. 2005. Multicenter clinical evaluation of the (1->3) beta-D-glucan assay as an aid to diagnosis of fungal infections in humans. Clin. Infect. Dis. 41:654-659.[CrossRef][Medline]
    44. 44
  44. Pappas, P. G., J. H. Rex, J. D. Sobel, S. G. Filler, W. E. Dismukes, T. J. Walsh, and J. E. Edwards. 2003. Guidelines for treatment of candidiasis. Clin. Infect. Dis. 38:161-189.
    45. 45
  45. Potera, C. 1999. Forging a link between biofilms and disease. Science 283:1837-1838.[Free Full Text]
    46. 46
  46. Ramage, G, S. P. Saville, D. P. Thomas, and J. L. Lopez-Ribot. 2005. Candida biofilms: an update. Eukaryot Cell. 4:633-638.[Free Full Text]
    47. 47
  47. Ramage, G., S. Bachmann, T. F. Patterson, B. L. Wickes, and J. L. Lopez-Ribot. 2002. Investigation of multidrug efflux pumps in relation to fluconazole resistance in Candida albicans biofilms. J. Antimicrob Chemother. 49:973-980.[Abstract/Free Full Text]
    48. 48
  48. Ramage, G., K. Vande Walle, S. P. Bachmann, B. L. Wickes, and J. L. Lopez-Ribot. 2002. In vitro pharmacodynamic properties of three antifungal agents against preformed Candida albicans biofilms determined by time-kill studies. Antimicrob Agents Chemother. 46:3634-3636.[Abstract/Free Full Text]
    49. 49
  49. Ramage, G., K. Vande Walle, B. L. Wickes, and J. L. Lopez-Ribot. 2001. Standardized method for in vitro antifungal susceptibility testing of Candida albicans biofilms. Antimicrob Agents Chemother. 45:2475-2479.[Abstract/Free Full Text]
    50. 50
  50. Ramage, G., K. Vande Walle, B. L. Wickes, and J. L. Lopez-Ribot. 2001. Characteristics of biofilm formation by Candida albicans. Rev. Iberoam Micol. 18:163-170.[Medline]
    51. 51
  51. Shimoi, H., H. Kitagaki, H. Ohmori, Y. Iimura, and K. Ito. 1998. Sed1p is a major cell wall protein of Saccharomyces cerevisiae in the stationary phase and is involved in lytic enzyme resistance. J. Bacteriol. 180:3381-3387.[Abstract/Free Full Text]
    52. 52
  52. Te Dorsthorst, D. T., P. E. Verweij, J. F. Meis, N. C. Punt, and J. W. Mouton. 2004. In vitro interactions between amphotericin B, itraconazole, and flucytosine against 21 clinical Aspergillus isolates determined by two drug interaction models. Antimicrob Agents Chemother. 48:2007-2013.[Abstract/Free Full Text]
    53. 53
  53. Warnock, D. W., E. M. Johnson, and D. A. White. 1999. Antifungal drug measurements, p. 221-223. In D. S. Reeves, R. Wise, J. M. Andrews, L. O. White, and D. Speller (ed.), Clinical antimicrobial assays. Oxford University Press, Oxford, United Kingdom.
    54. 54
  54. White, T. C., K. A. Marr, and R. A. Bowden. 1998. Clinical, cellular, and molecular factors that contribute to antifungal drug resistance. Clin. Microbiol. Rev. 11:382-402.[Abstract/Free Full Text]
    55. 55
  55. Yin Q. Y., P. W. J. de Groot, H. L. Dekker, L. de Jong, F. M. Klis, and C. G. de Koster. 2005. Comprehensive proteomic analysis of Saccharomyces cerevisiae cell walls. J. Biol. Chem. 280:20894-20901.[Abstract/Free Full Text]

Antimicrobial Agents and Chemotherapy, February 2007, p. 510-520, Vol. 51, No. 2
0066-4804/07/$08.00+0     doi:10.1128/AAC.01056-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.



This article has been cited by other articles:

  • ten Cate, J.M., Klis, F.M., Pereira-Cenci, T., Crielaard, W., de Groot, P.W.J. (2009). Molecular and Cellular Mechanisms That Lead to Candida Biofilm Formation. JDR 88: 105-115 [Abstract] [Full Text]  
  • Nett, J. E., Guite, K. M., Ringeisen, A., Holoyda, K. A., Andes, D. R. (2008). Reduced Biocide Susceptibility in Candida albicans Biofilms. Antimicrob. Agents Chemother. 52: 3411-3413 [Abstract] [Full Text]  
  • Al-Dhaheri, R. S., Douglas, L. J. (2008). Absence of Amphotericin B-Tolerant Persister Cells in Biofilms of Some Candida Species. Antimicrob. Agents Chemother. 52: 1884-1887 [Abstract] [Full Text]  
  • Cowen, L. E., Steinbach, W. J. (2008). Stress, Drugs, and Evolution: the Role of Cellular Signaling in Fungal Drug Resistance. Eukaryot Cell 7: 747-764 [Full Text]  
  • Angiolella, L., Stringaro, A. R., De Bernardis, F., Posteraro, B., Bonito, M., Toccacieli, L., Torosantucci, A., Colone, M., Sanguinetti, M., Cassone, A., Palamara, A. T. (2008). Increase of Virulence and Its Phenotypic Traits in Drug-Resistant Strains of Candida albicans. Antimicrob. Agents Chemother. 52: 927-936 [Abstract] [Full Text]  
  • Uppuluri, P., Nett, J., Heitman, J., Andes, D. (2008). Synergistic Effect of Calcineurin Inhibitors and Fluconazole against Candida albicans Biofilms. Antimicrob. Agents Chemother. 52: 1127-1132 [Abstract] [Full Text]  
  • Hiller, E., Heine, S., Brunner, H., Rupp, S. (2007). Candida albicans Sun41p, a Putative Glycosidase, Is Involved in Morphogenesis, Cell Wall Biogenesis, and Biofilm Formation. Eukaryot Cell 6: 2056-2065 [Abstract] [Full Text]  

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowReprints and Permissions
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Nett, J.
Right arrow Articles by Andes, D.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Nett, J.
Right arrow Articles by Andes, D.

Copyright © 2009 by the American Society for Microbiology.   For an alternate route to Journals.ASM.org, visit: http://intl-journals.asm.org | More Info»