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Journal of Bacteriology, September 2001, p. 5385-5394, Vol. 183, No. 18
0021-9193/01/$04.00+0 DOI: 10.1128/JB.183.18.5385-5394.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
Biofilm Formation by the Fungal Pathogen
Candida albicans: Development, Architecture, and
Drug Resistance
Jyotsna
Chandra,1
Duncan M.
Kuhn,1,2
Pranab
K.
Mukherjee,1
Lois L.
Hoyer,3
Thomas
McCormick,4 and
Mahmoud A.
Ghannoum1,*
Center for Medical Mycology, University
Hospitals of Cleveland, and Department of Dermatology, Case Western
Reserve University,1 Division of
Infectious Diseases, University Hospitals of
Cleveland,2 and Department of
Dermatology, Case Western Reserve University,4
Cleveland, Ohio 44106, and Department of Veterinary
Pathobiology, University of Illinois at Urbana-Champaign, Urbana,
Illinois 618023
Received 14 May 2001/Accepted 27 June 2001
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ABSTRACT |
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.
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INTRODUCTION |
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.
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MATERIALS AND METHODS |
Strains.
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.
Biofilm formation.
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.
Fluorescence microscopy.
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 (
max = 432 nm). Stained biofilms
were examined under a fluorescence microscope (ZVS-47E microscope; Carl
Zeiss, Inc., Oberkochen, Germany).
Confocal scanning laser microscopy (CSLM).
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
37oC 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.
Antifungal susceptibility.
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.
Northern blot analysis.
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).
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RESULTS |
C. albicans biofilm formation proceeds in three
distinct developmental phases.
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
1 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. 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. 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. 1c). Fungal communities and the
extracellular material in which they are embedded constitute the
biofilm.
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FIG. 1.
Development of C. albicans biofilm on
polymethylmethacrylate strips. Fluorescence microscopy images show the
three distinct developmental phases of C. albicans
biofilms over a 72-h period: early (a), intermediate (b), and
maturation (c) phases. Magnification, ×10.
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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 × 10
7) 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.
Model biofilms are morphologically similar to those formed in
vivo.
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).
C. albicans biofilm has a highly heterogeneous
structure.
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
2 shows 3-D reconstruction of images of
the early biofilm development
phase with individual yeast cells
adhering to acrylic strips (Fig.
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.
2b). Microcolonies of
predominantly yeast forms were visible by 11 h (Fig.
2c), while
mature biofilms showed fungal cells embedded within
the extracellular
material (Fig.
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.
2e and f). As shown in Fig.
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.
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.
2g and h).
This appearance is similar to that seen with
bacterial biofilm systems
(
16).
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FIG. 2.
CSLM images of a C. albicans biofilm
grown on denture acrylic surface. (a to d) Horizontal
(xy) view of reconstructed 3-D images at 0 (a), 8 (b),
11 (c), and 48 (d) h. Bar, 20 µm. (e and f) Orthogonal images of
C. albicans biofilms grown to early and maturation
phases show that early-phase (0 h) biofilm consisted of mostly yeast
cells separated by blank spaces (arrows) (e), while maturation-phase
(48 h) biofilm showed metabolically active (red, FUN1-stained) cells
embedded in the polysaccharide extracellular material (green,
ConA-stained, arrows) (f). (g and h) Thickness of the biofilm (25
µm) can be observed in the side view of the reconstruction (g), while
a horizontally tilted image (with false 3-D cubes) shows the
heterogeneity of the biofilm (h). Magnification, ×40.
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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.
3).
Above this layer of cells, profuse matrix
(at least
450 µm thick)
was apparent which consisted of extracellular
material and hyphal
elements (Fig.
3a 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.
3c 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.
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.
3e 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.
4.
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FIG. 3.
CSLM images of Calcofluor-stained mature C.
albicans biofilms formed on silicone elastomer surface. (a and
b) Projection (xz or side view) of 3-D reconstructed
images showing an approximately 450-µm-thick biofilm with a basal
layer (10 to 12 µm thick) consisting of yeast cells and a top layer
consisting of hyphal elements (arrow). The extracellular material (ECM)
is stained with ConA, resulting in the green color. (c and d)
Orthogonal images of the basal (c) and upper layers (d). The
ECM-derived haziness seen in mature biofilm (e) is absent when the
extracellular material is removed (f). Magnification, ×20.
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FIG. 4.
Schematic representation of biofilm development in
C. albicans. (a and b) Biofilm grown on
polymethylmethacrylate (PMA) strips. (c and d) Biofilm grown on
silicone elastomer (SE) disks. Panels a and c represent the substrate
seen from the top, while panels c and d show the view from the sides of
the PMA strip and SE disk, respectively. ECM, extracellular material.
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C. albicans has a greater ability to form biofilms
than S. cerevisiae
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.
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. 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. 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. 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. 5e), it was unable to
form mature biofilms compared to C. albicans.
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FIG. 5.
Comparison of the abilities of C.
albicans and S. cerevisiae to form biofilms. (a)
Quantitative measurement of dry weight of biofilms formed by C.
albicans and S. cerevisiae. (b) Growth profiles
of planktonic C. albicans and S.
cerevisiae. Fluorescence microscopy images of C.
albicans (c) and S. cerevisiae (d) grown on
polymethylmethacrylate strips. (e) Fluorescence micrograph showing
S. cerevisiae growing on silicone elastomer disk (3
to 4 µm in thickness). Magnification, ×10.
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Antifungal resistance increases during biofilm development.
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.
6). As the biofilms developed, MICs
progressively increased (Fig. 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. 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 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 1).
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FIG. 6.
Correlation of biofilm development and metabolic
activity with antifungal resistance. The susceptibilities of C.
albicans at different stages of biofilm development to
fluconazole (a), amphotericin B (b), nystatin (c), and chlorhexidine
(d) are represented as histograms. The line curves show percent
metabolic activity of growing C. albicans biofilms
exposed to fluconazole (64 µg/ml), amphotericin B (4 µg/ml),
nystatin (8 µg/ml), or chlorhexidine (64 µg/ml). Metabolic activity
was normalized to the control without drugs, which was taken as
100%.
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TABLE 1.
MICs of different agents for S. cerevisiae
strain MRL-138 and C. albicans strain GDH-2346 grown on
polymethylmethacrylate stripsa
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Differential gene expression under planktonic and biofilm
conditions.
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.
7). Further experimentation is required
to assess the role of individual Als proteins in biofilm formation.
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|
FIG. 7.
Northern blot analysis of total RNA extracted from
planktonic and biofilm-grown C. albicans cells. Total
RNA from planktonic and biofilm-grown cells was loaded in various
quantities (20, 30, 40, and 50 µg) on a formaldehyde-agarose gel. The
resulting Northern blot was probed with a fragment of the C.
albicans ALS1 tandem repeats, which hybridized several genes in
the family. Probing with a fragment of the TEF1 gene
served as loading control. Molecular size markers (kb) are shown on the
left.
|
|
| |
DISCUSSION |
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.
| |
ACKNOWLEDGMENTS |
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.).
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Center for
Medical Mycology, University Hospitals of Cleveland and Department of
Dermatology, Case Western Reserve University, 11100 Euclid Ave.,
Cleveland, OH 44106-5028. Phone: (216) 844-8580. Fax: (216) 844-1076. E-mail: mag3{at}po.cwru.edu.
| |
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Journal of Bacteriology, September 2001, p. 5385-5394, Vol. 183, No. 18
0021-9193/01/$04.00+0 DOI: 10.1128/JB.183.18.5385-5394.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
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Abstract
Introduction
