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Biofilms are a protected niche for microorganisms, where they are safe from antibiotic treatment and can create a source of persistent infection. Using two clinically relevant Candida albicans biofilm models formed on bioprosthetic materials, we demonstrated that biofilm formation proceeds through three distinct developmental phases. These growth phases transform adherent blastospores to well-defined cellular communities encased in a polysaccharide matrix. Fluorescence and confocal scanning laser microscopy revealed that C. albicans biofilms have a highly heterogeneous architecture composed of cellular and noncellular elements. In both models, antifungal resistance of biofilm-grown cells increased in conjunction with biofilm formation. The expression of agglutinin-like (ALS) genes, which encode a family of proteins implicated in adhesion to host surfaces, was differentially regulated between planktonic and biofilm-grown cells. The ability of C. albicans to form biofilms contrasts sharply with that of Saccharomyces cerevisiae, which adhered to bioprosthetic surfaces but failed to form a mature biofilm. The studies described here form the basis for investigations into the molecular mechanisms of Candida biofilm biology and antifungal resistance and provide the means to design novel therapies for biofilm-based infections.
Biofilms are studied in a wide range of scientific disciplines including biomedicine, water engineering, and evolutionary biology (3, 10, 14, 15, 22, 23, 33). Biofilms are the most common mode of bacterial growth in nature and are also important in clinical infections, especially due to the high antibiotic resistance associated with them (4, 11, 46). In contrast to the extensive literature describing bacterial biofilms (consult references 34, 35, and 42 for excellent reviews on bacterial biofilms), little attention has been paid to medically relevant fungal biofilms. Transplantation procedures, immunosuppression, the use of chronic indwelling devices, and prolonged intensive care unit stays have increased the prevalence of fungal disease. Fungi most commonly associated with such disease episodes are in the genus Candida, most notably Candida albicans, which causes both superficial and systemic disease. Even with current antifungal therapy, mortality of patients with invasive candidiasis can be as high as 40% (43). Candidiasis is usually associated with indwelling medical devices (e.g., dental implants, catheters, heart valves, vascular bypass grafts, ocular lenses, artificial joints, and central nervous system shunts), which can act as substrates for biofilm growth. In a multicenter study of 427 consecutive patients with candidemia, the mortality rate for patients with catheter-related candidemia was found to be 41% (31). Forty percent of patients with microbial colonization of intravenous catheters develop occult fungemia, with consequences ranging from focal disease to severe sepsis and death (2, 31). The tenacity with which Candida infects indwelling biomedical devices necessitates their removal to effect a cure. Biofilm formation is also critical in the development of denture stomatitis, a superficial form of candidiasis that affects 65% of edentulous individuals (8, 9). Despite the use of antifungal drugs to treat denture stomatitis, infection is often reestablished soon after treatment (28). These clinical observations emphasize the importance of biofilm formation to both superficial and systemic candidiasis and the inability of current antifungal therapy to cure such diseases.
Our initial work on fungal biofilms involved development and characterization of C. albicans biofilms formed on two common bioprosthetic materials: polymethylmethacrylate, which is used in construction of dentures, and silicone elastomer, a model material used for indwelling devices including catheters. The availability of well-characterized, reproducible biofilm models is essential to understanding the nature of Candida biofilms and performing studies of biofilm formation and antifungal drug resistance. Recently, using the polymethylmethacrylate biofilm model, we showed that biofilm-grown C. albicans cells are highly resistant to antifungal agents such as fluconazole, nystatin, amphotericin B, and chlorhexidine (11), similar to previous observations reported for catheter-associated C. albicans biofilms (5, 21). Here, we extend our studies of the model biofilms with emphasis on identifying biofilm growth phases, architectural organization, and correlating antifungal resistance with biofilm development. The use of physiologic parameters and comparison to biofilms from patient specimens demonstrated the clinical relevance of our observations. We also compared C. albicans biofilm formation with that of Saccharomyces cerevisiae and performed an initial assessment of differential gene expression between planktonic and biofilm-grown Candida cells. Based on our results, we propose that biofilm formation is a highly complex phenomenon, distinct from fungal adhesion. Our data support the conclusion that true biofilms involve both the production of specific extracellular materials and special cellular functions. The information derived from these studies will further our understanding of Candida biofilm biology as well as antifungal resistance and may lead to novel therapies for biofilm-based diseases.
C. albicans strain GDH-2346 was isolated from a denture stomatitis patient (obtained from L. Julia Douglas, University of Glasgow, Glasgow, United Kingdom). C. albicans strain M-61 was obtained from an infected intravascular catheter. S. cerevisiae strain M-20 was obtained from the stool of a patient on immunosuppressive therapy and admitted to University Hospitals of Cleveland, while strain MRL-138 was a bronchoscopy specimen provided by Lynn Steele More at Christiana Care Health Services, Infectious Disease Laboratory, Wilmington, Del. These fungal strains were grown overnight at 37°C in yeast nitrogen base medium (YNB; Difco Laboratories, Detroit, Mich.) supplemented with 50 mM glucose. The identities of the C. albicans and S. cerevisiae isolates were determined using the api20C-AUX system, germ tube formation, and urea and nitrate assimilation tests. In these assays, both strains of S. cerevisiae were found to be negative for the germ tube and urea and potassium nitrate assimilation tests. Furthermore, these strains were found to be positive for glucose, galactose, maltose, saccharose, and raffinose, but they were negative for 2-keto-d-gluconate, arabinose, xylose, adonitol, xylitol, inositol, sorbitol, N-acetyl-d-glucosamine, cellobiose, and lactose assimilation tests.
Biofilms were formed on 1.5-cm2 polymethylmethacrylate strips (Dentsply Intl., York, Pa.) as described previously (11). The method for growing biofilms on silicone elastomer disks is described below. A standard inoculum of 1 × 107 cells from overnight cultures of the fungal strains was used to form biofilms in both models. For filamentation experiments, the standard inoculum was added to polymethylmethacrylate strips and then incubated in RPMI 1640 medium (Cellgro; Mediatech). For biofilms grown on silicone elastomer, 1.5-cm-diameter disks of the material (Cardiovascular Instrument Corp., Wakefield, Mass.) were placed in 12-well tissue culture plates and incubated in fetal bovine serum (FBS) for 24 h at 37°C on a rocker. After this pretreatment, disks were washed with phosphate-buffered saline (PBS) to remove residual FBS. To ensure uniform biofilm formation across the strongly hydrophobic disk surface, disks were immersed in 3 ml of standardized cell suspension (1 × 107 cells/ml) and incubated for 90 min at 37°C. Wells were gently washed with PBS to remove nonadherent cells. Subsequently, disks were immersed in YNB medium with 50 mM glucose and incubated for various durations at 37°C on a rocker. Biofilms formed on both polymethylmethacrylate and silicone elastomer materials were quantitated using a tetrazolium XTT [2,3-bis(2-methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-carboxanilide] reduction assay and dry weight measurement as described previously (11). Disks containing no Candida cells served as controls. Assays were carried out in four replicates and were repeated on different days.
Polymethylmethacrylate strips or silicone elastomer disks with biofilms were transferred to microscope slides and stained for 1 min with 50 μl of Calcofluor-White (0.05% [vol/vol]; Sigma Chemical Co., St. Louis, Mo.), which fluoresces in the UV range (λmax = 432 nm). Stained biofilms were examined under a fluorescence microscope (ZVS-47E microscope; Carl Zeiss, Inc., Oberkochen, Germany).
At various time points during biofilm formation, polymethylmethacrylate strips or silicone elastomer disks on which biofilms were developing were transferred to a 12-well plate and incubated for 45 min at 37οC in 4 ml of PBS containing the fluorescent stains FUN-1 (10 μM) and concanavalin A-Alexa Fluor 488 conjugate (ConA; 25 μg/ml). FUN-1 (excitation wavelength = 543 nm and emission = 560 nm long-pass filter) is converted to orange-red cylindrical intravacuolar structures by metabolically active cells, while ConA (excitation wavelength = 488 nm and emission = 505 nm long-pass filter) binds to glucose and mannose residues of cell wall polysaccharides with green fluorescence.
After incubation with the dyes, polymethylmethacrylate strips or silicone elastomer disks were flipped and placed on a 35-mm-diameter glass-bottom petri dish (MatTek Corp., Ashland, Mass.). Stained biofilms were observed with a Zeiss LSM510 confocal scanning laser microscope equipped with argon and HeNe lasers and mounted on a Zeiss Axiovert100 M microscope (Carl Zeiss, Inc.). The objective used was a water immersion C-apochromat lens (40×; numerical aperture of 1.2). Depth measurements were taken at regular intervals across the width of the device. To determine the structure of the biofilms, a series of horizontal (xy) optical sections with a thickness of 0.9 μm, at 0.44-μm intervals, were taken throughout the full length of the biofilm. Confocal images of green (ConA) and red (FUN-1) fluorescence were conceived simultaneously using a multitrack mode. For Calcofluor staining, silicone elastomer disks containing biofilms were gently washed with fresh PBS. Any excess liquid was carefully blotted from the side of the disk, and 50 μl of Calcofluor-White was added to the biofilm surface. Disks were then observed under the confocal microscope as described above.
Biofilms were grown on polymethylmethacrylate strips as described previously (11). To measure antifungal susceptibility of C. albicans cells grown in developing biofilms, strips were transferred to wells containing different concentrations (ranging from 0.5 to 256 μg/ml) of fluconazole, amphotericin B, nystatin, or chlorhexidine. Strips were incubated for 48 h and metabolic activities of biofilms were measured using the XTT reduction assay as described previously (11). The antifungal susceptibility of planktonic cells was measured using the National Committee for Clinical Laboratory Standards standard M-27A (30) and XTT methods (11) as described previously.
To determine whether expression of C. albicans genes is altered as a result of growth as a biofilm on polymethylmethacrylate strips, both C. albicans biofilms and planktonic cells were grown in YNB as previously described (11). Biofilm material was scraped from the surface of denture acrylic strips, resuspended in PBS, and centrifuged (3,000 × g) to obtain a pellet containing biofilm matrix. Planktonic cells were similarly collected. Total RNA was extracted from biofilm and planktonic cells according to the procedure described by Collart and Oliviero (13) and analyzed by Northern blot analysis as described previously (26). The blot was probed with a fragment encoding the tandem repeat of ALS1, which recognizes multiple genes in the ALS family (25). A fragment of the C. albicans TEF1 gene (41) was used as a control for equal loading as previously described (25).
We previously developed a model for denture fungal biofilm growth of C. albicans on strips of polymethylmethacrylate (11). In the present study, we investigated temporal development of fungal biofilms using the polymethylmethacrylate biofilm model, as well as one based on silicone elastomer disks. Biofilms grown on polymethylmethacrylate strips were examined by fluorescence microscopy using Calcofluor-White, a UV-excitable dye that binds chitin and beta-glucan and has long been used to highlight fungal cell walls (1). Figure Figure11 shows that C. albicans biofilm formation on polymethylmethacrylate strips progresses in three distinct developmental phases: early (≈0 to 11 h), intermediate (≈12 to 30 h), and maturation (≈38 to 72 h). Initially (0 to 2 h), the majority of C. albicans cells were present as blastospores (yeast forms) adhering to the surface of the polymethylmethacrylate strips. At 3 to 4 h, distinct microcolonies appeared on the surface of the strips (Fig. (Fig.1a).1a). By 11 h, C. albicans communities appeared as thick tracks of fungal growth, due to cell growth and aggregation along areas of surface irregularities. The intermediate developmental phase was characterized by the emergence of predominantly noncellular material (at ≈12 to 14 h), which appeared as a haze-like film covering the fungal microcolonies (Fig. (Fig.1b).1b). The hazy appearance was due to diffuse staining of the extracellular material with Calcofluor and implied that this material was composed mainly of cell-wall-like polysaccharides. During the maturation phase, the amount of extracellular material increased with incubation time, until C. albicans communities were completely encased within this material. At this stage it was difficult to focus on the basal blastospore communities covered by the matrix (Fig. (Fig.1c).1c). Fungal communities and the extracellular material in which they are embedded constitute the biofilm.
These studies were performed on C. albicans cells growing in YNB medium, which supports growth of blastospores. C. albicans can exist as yeast cells, pseudohyphae, or hyphae. The hyphal forms of C. albicans are believed to play an important role in fungal infection (18, 48, 49). Therefore, we repeated our experiments using a different medium, RPMI 1640, which induces hyphal formation in C. albicans (26). To initiate biofilm formation, C. albicans yeast cells (1 × 107) were added to polymethylmethacrylate strips and incubated in RPMI 1640 medium. After 72 h, biofilms formed in this medium showed no significant difference in terms of dry biomass and metabolic activity, compared to biofilms grown in YNB. In this regard, metabolic activity and dry biomass (optical density at 492 nm [mean ± standard error of the mean], 0.285 ± 0.009; 2.707 ± 0.005 mg/denture piece) of biofilms grown in RPMI 1640 were similar to those grown in YNB (optical density at 492 nm, 0.366 ± 0.054; 2.750 ± 0.150 mg/denture piece) (P > 0.05 for both comparisons). Unlike the YNB-grown biofilms, which contained mainly yeast forms, the RPMI-grown biofilms consisted mostly of C. albicans filaments (data not shown). These data suggested that both yeast and hyphal forms of C. albicans were capable of biofilm formation, indicating that biofilm growth was not morphology specific.
We found a similar pattern of biofilm growth on the silicone elastomer substrate model. Fluorescence microscopy showed a much more confluent blastospore layer early in development, followed by the production of extracellular material. The resulting biofilm matrix, although grown in YNB medium, had an abundant component of hyphal elements. However, this silicone elastomer biofilm model incorporates FBS, which is known to promote hyphal formation in C. albicans (32). In order to determine if the abundant hyphae observed in silicone elastomer biofilms are induced by FBS, we repeated these experiments by replacing FBS with PBS. Our data showed that biofilms grown in the presence of PBS also contained hyphae, albeit to a lesser extent than that observed in FBS-grown biofilms.
To determine whether our in vitro model biofilms mimic those formed on bioprosthetic devices in vivo, an infected central venous catheter was obtained from a patient with C. albicans fungemia. The catheter was removed and several distal centimeters were cut off and placed in a specimen cup. After the tip had been rolled on a blood agar plate for culture, the distal 1 cm of the catheter was cut off, placed on a glass microscope slide, stained with Calcofluor, and observed under fluorescence microscopy. This examination revealed that the biofilm formed on the infected intravenous catheter was similar in structure to biofilms grown using our silicone elastomer model (data not shown). These data suggest that our in vitro model system is analogous to in vivo events and may be clinically relevant. However, to conclude that the in vitro and in vivo biofilms share the same properties, it will be necessary to study multiple in vivo-derived biofilms and compare the biomass per surface area of the in vitro and in vivo biofilms and to determine the antifungal resistance of the biofilm-grown fungi versus planktonically grown fungi recovered from implants.
It is important to note that similar biofilm morphological patterns were produced in our systems under a variety of environmental parameters (substrates, substrate preconditioning solutions, and growth media). While environmental conditions affect biofilm production, it is possible that certain pathogenic strains of fungi such as C. albicans have a special ability to initiate biofilm formation under a wide range of conditions, similar to that seen in some bacteria such as Pseudomonas fluorescens (16).
Biofilm development was further explored using CSLM. We chose this technique instead of scanning electron microscopy because the fixation and dehydration required for scanning electron microscropy severely distort biofilm architecture and shrink any aqueous phase, while CSLM preserves the intact structure of biofilms (36, 44, 45). CSLM examination of C. albicans biofilms used a combination of the fluorescent dyes FUN-1 and ConA (both from Molecular Probes, Inc., Eugene, Oreg.). In addition to localizing cells within a specimen, combinations of FUN-1 (a cytoplasmic fluorescent probe for cell viability) and ConA (which selectively binds to mannose and glucose residues present in cell wall polysaccharides) can be used to assess cell viability (20, 29).
Biofilm images were either displayed individually or reconstructed in three-dimensional (3-D) projections. In addition, vertical (xz) sections or side views of the 3-D reconstructed images were used to determine biofilm thickness and architecture. Figure Figure22 shows 3-D reconstruction of images of the early biofilm development phase with individual yeast cells adhering to acrylic strips (Fig. (Fig.2a).2a). Intense green fluorescence resulting from ConA binding to polysaccharides outlined the cell walls of the yeast, while the red color due to FUN-1 staining localized in dense aggregates in the cytoplasm of metabolically active cells. Thus, areas of red fluorescence represent metabolically active cells and green fluorescence indicates cell-wall-like polysaccharides, while yellow areas represent dual staining. By 8 h, C. albicans cells increased in density and tended to aggregate (Fig. (Fig.2b).2b). Microcolonies of predominantly yeast forms were visible by 11 h (Fig. (Fig.2c),2c), while mature biofilms showed fungal cells embedded within the extracellular material (Fig. (Fig.2d).2d). These analyses revealed a highly heterogeneous architecture of mature C. albicans biofilms in terms of the distribution of fungal cells (indicated by red color) and extracellular material (green coloration). Importantly, no extracellular material could be detected at the early biofilm stage. The lack of extracellular material was confirmed by orthogonal presentation of the data (Fig. (Fig.2e2e and f). As shown in Fig. Fig.2e,2e, yeast cells in early biofilm development were separated by regions lacking fluorescence, indicating the absence of extracellular material. In contrast, C. albicans cells in mature biofilms were encased in extracellular material which appeared as diffuse green fluorescence separate from cell bodies (Fig. (Fig.2f).2f). The fact that the biofilm extracellular material stained with ConA, in parallel to the staining pattern of Calcofluor, reinforced the concept that biofilm extracellular material was composed of polysaccharide material. Projection analysis as well as vertical (xz) sectioning (side view) of 3-D reconstructed images also revealed that mature C. albicans biofilms have a heterogeneous matrix structure (25 to 30 μm thick), with thin areas of metabolically active cells interwoven with extracellular polysaccharide material (Fig. (Fig.2g2g and h). This appearance is similar to that seen with bacterial biofilm systems (16).
Because silicone elastomer disks have a uniformly flat surface, unlike the irregular surface of polymethylmethacrylate, planar imaging is readily obtainable. C. albicans cells grown on silicone elastomer produced a nearly uniform confluent layer of adherent blastospores, which at maturation were several cells thick (approximately 10 to 12 μm; Fig. Fig.3).3). Above this layer of cells, profuse matrix (at least ≈450 μm thick) was apparent which consisted of extracellular material and hyphal elements (Fig. (Fig.3a3a and b). Hyphal elements originating at the base layer pervaded the extracellular material, both in proximity to the blastospores and through the entire thickness (Fig. (Fig.3c3c to f). Graininess of the images is not an artifact, but rather is due to binding of Calcofluor or ConA (depending on the protocol used) to extracellular material, as demonstrated by the projections in Fig. Fig.3.3. If matrix was physically removed (either by rubbing or vigorous washing of the biofilm surface), a basal layer of blastospores remained and the granularity disappeared (Fig. (Fig.3e3e and f). Based on our studies with the polymethylmethacrylate as well as the silicone elastomer models, a schematic representation of biofilm development on these surfaces is shown in Fig. Fig.4.4.
S. cerevisiae is a model microbe for the study of eukaryotic organisms, including fungi like C. albicans. Although S. cerevisiae has been isolated from clinical conditions (12), its presence is quite rare and the organism is considered relatively nonpathogenic. We sought to determine whether the property of biofilm formation is a characteristic of pathogenic fungi or a general property that can be extended to species such as S. cerevisiae. To do this, we compared biofilm formation by C. albicans and S. cerevisiae using both the polymethylmethacrylate and silicone elastomer models. In the polymethylmethacrylate model, biofilm formed by S. cerevisiae had significantly less growth than biofilm formed by C. albicans, as determined by dry weight measurements (P = 0.0008; Fig. Fig.5a).5a). To rule out the possibility that differences in growth rates may account for the varying ability of these species to form biofilms, we compared their growth profiles, which appeared quite similar (Fig. (Fig.5b).5b). The superior ability of C. albicans to form biofilms compared to S. cerevisiae was confirmed by fluorescence microscopy, which revealed that C. albicans formed extracellular material-rich biofilms on polymethylmethacrylate strips (Fig. (Fig.5c).5c). In contrast, S. cerevisiae adhered to the strips, but its growth was limited to thin colonies which failed to produce appreciable extracellular material (Fig. (Fig.5d).5d). A similar observation was noted for biofilms grown on silicone elastomer disks. Dry weight measurement (3.7 ± 0.001 mg/disk for C. albicans and 1.6 ± 0.004 mg/disk for S. cerevisiae) as well as fluorescence microscopy analysis of biofilms growing on silicone elastomer disks also showed that while S. cerevisiae adhered to the silicone elastomer surface (Fig. (Fig.5e),5e), it was unable to form mature biofilms compared to C. albicans.
Previous studies have shown that fungal biofilms grown on denture and catheter material become resistant to antifungals (11, 21). A similar resistance pattern was seen with our silicone elastomer model where MICs of fluconazole were 1 and >128 μg/ml for planktonic and biofilm-grown C. albicans cultures, respectively. Since it is plausible that antifungal resistance evolves as the biofilm grows to maturation, we investigated correlations between biofilm development and antifungal susceptibility. MICs of amphotericin B, nystatin, fluconazole, and chlorhexidine were determined for early, intermediate, or mature biofilms. C. albicans exhibited low MICs at the early biofilm phase. MICs during this phase were 0.5, 1, 8, and 16 μg/ml for amphotericin B, fluconazole, nystatin, and chlorhexidine, respectively (Fig. (Fig.6).6). As the biofilms developed, MICs progressively increased (Fig. (Fig.6).6). By 72 h, C. albicans cells were highly resistant, with MICs of 8, 128, 32, and 256 μg/ml for amphotericin B, fluconazole, nystatin, and chlorhexidine, respectively. The progression of drug resistance was associated with the concomitant increase in metabolic activity of developing biofilms (Fig. (Fig.6).6). This indicated that the observed increase in drug resistance was not simply a reflection of lower metabolic activity of cells in maturing biofilms but that drug resistance develops over time, coincident with biofilm maturation. To determine whether S. cerevisiae develops antifungal resistance during growth on polymethylmethacrylate strips, MICs of amphotericin B, fluconazole, nystatin, and chlorhexidine were measured at early (0 h) and late (72 h) phases. Our results showed that the susceptibilities of these two phases were similar (Table (Table1).1). This was in contrast to C. albicans biofilms, for which the MICs increased dramatically between the two time points. Thus, unlike C. albicans biofilms, resistance of S. cerevisiae to antifungals does not evolve significantly over time when grown on polymethylmethacrylate strips from 0 to 72 h (Table (Table1).1).
Since adhesion to bioprosthetic surfaces and cell aggregation are precursors to biofilm formation, it is logical to assume expression of genes involved in these processes changes during the transition from planktonic to biofilm growth. To begin addressing this issue, we investigated the expression profile of C. albicans genes belonging to the ALS family, which encode proteins implicated in adhesion of C. albicans to host surfaces (24). Northern blot analysis of total RNA from planktonic and biofilm-grown C. albicans cells showed that there was differential expression of genes between the two growth forms, with additional gene(s) expressed in biofilms (Fig. (Fig.7).7). Further experimentation is required to assess the role of individual Als proteins in biofilm formation.
C. albicans biofilm formation proceeds in an organized fashion through the early, intermediate, and maturation phases of development. Similar distinct developmental phases have been reported for biofilm formation by many bacterial species (16, 34, 42). Thus, microorganisms appear to share common basic steps during biofilm formation. Development of biofilm is closely associated with the generation of matrix, the majority of which is extracellular material. Microscopy strongly suggests that extracellular material is predominantly composed of cell-wall-like polysaccharides containing mannose and glucose residues, based on staining with dyes that specifically bind these carbohydrates. Mature C. albicans biofilms have a highly heterogeneous architecture in terms of distribution of fungal cells and extracellular material. In addition, compared to biofilms grown on the irregular surface of polymethylmethacrylate, those grown on flat hydrophobic surfaces such as silicone elastomer have a distinct biphasic structure composed of an adherent blastospore layer covered by sparser hyphal elements embedded in a deep layer of extracellular material. A similar biphasic distribution was suggested for C. albicans biofilms grown on polyvinyl chloride disks (6). Formation of this biphasic architecture could be in response to environmental conditions, such as differences in pH, oxygen availability, and redox potential, prevailing within the biofilm (32, 37, 40, 44, 47). Heterogeneity in the biofilms is another characteristic shared among microorganisms. A “heterogeneous mosaic model” for biofilms has been described, containing stacks of bacterial microcolonies held together by extracellular polymeric substances. Below the stacks is an underlying layer of cells (≈5 μm thick) attached to the substratum (44).
Like their bacterial counterparts, biofilm-grown C. albicans cells are highly resistant to antimicrobials. Although drug resistance has been shown in C. albicans (11, 21) and bacterial biofilms (7, 15), this is the first report correlating the emergence of antifungal resistance with the development of biofilms. Developing C. albicans biofilms are associated with an increasing presence of extracellular material. Extracellular polymeric substance in bacterial biofilms is known to physically interact with antibiotics and is believed to contribute to resistance against these drugs (19, 27). It is unclear if the increase in drug resistance in C. albicans biofilms is due to production of extracellular material or due to genetic and biochemical alterations in fungal cells; this is an area for future study. An alternative explanation proposed for antifungal resistance in biofilms is metabolic quiescence of cells. However, this possibility is not likely since biofilm-embedded cells actively metabolize substrates, including XTT and FUN-1.
The biofilm-forming ability of the pathogen C. albicans is markedly different from that of S. cerevisiae. While the latter is capable of adherence, it fails to progress to a mature biofilm characterized by the presence of extracellular material. A recent report suggested that S. cerevisiae forms biofilms in vitro (38). However, the putative S. cerevisiae biofilms described in that study do not resemble those formed by C. albicans in our standardized model. We believe that these results are in agreement with ours and indicate that, although S. cerevisiae adheres to surfaces in a limited way, it fails to form extracellular material-encased biofilms similar to those formed by C. albicans. Additionally, our results show that antifungal resistance of adherent S. cerevisiae cells did not increase with time. This result was in contrast to C. albicans biofilms, where a significant jump in the MICs of drugs was observed between early and mature phase biofilms.
ALS gene expression is differentially regulated during the transition of Candida from a planktonic to biofilm-associated organism. This documentation of differential gene expression between the two growth forms represents a small number of the transcriptional changes that are likely to occur during biofilm formation. The formation of extracellular material associated with C. albicans biofilms suggests that genes encoding enzymes involved in carbohydrate synthesis are differentially regulated during biofilm growth. It is also possible that increased expression of drug resistance genes is responsible for the increased MICs observed for C. albicans biofilms. Finally, the appearance of a well-defined basal layer of yeast followed by extracellular material production also suggests that genes involved in quorum sensing are important, as is the case with Pseudomonas spp. (17, 39). There may also be increased expression of drug resistance genes such as CDR1, CDR2, and MDR. Relevant gene regulation can likely only be determined using pathogenic organisms. The well-characterized genome of C. albicans and the availability of deletion mutants should allow for rapid evaluation of such possibilities.
Fungal biofilm formation is a complex phenomenon distinct from adhesion. It is best studied using pathogenic species grown on relevant bioprosthetic materials under near-physiologic conditions. Study of such systems will reveal the true nature of fungal biofilms and their biology. Demonstration of common biofilm features (distinct developmental phases, heterogeneous architecture, and drug resistance phenotypes) across different taxa extends the implication of this study beyond fungi to other organized cellular communities. The impact of this information will be widespread, ranging from new environmental microbiology insights to the development of antimicrobials specifically targeted against biofilm-associated infections.
We thank Anna-Liisa Nieminen for helpful discussions and invaluable suggestions.
This work was supported by grants from the U.S. National Institutes of Health (NIH grant nos. AI35097–03 and RO1-DE13992), the Steris Corporation Award for Emerging/Nosocomial Infections (no. 1–88-8225), the Center for AIDS Research at Case Western Reserve University (grant no. AI-36219), an NIH-Infectious Diseases/GeoMed training grant (no. AIO7024; D.M.K.), and a Dermatology Foundation/Janssen Pharmaceutical Research Fellowship (to P.K.M.).