| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
1 Department of Biological and Biomedical Sciences, Glasgow Caledonian University, Glasgow, UK
2 Microbiology Department, Yorkhill Hospital, Glasgow, UK
3 Section of Infection and Immunity, Glasgow University Dental School and Hospital, Glasgow, UK
Correspondence
Gordon Ramage
g.ramage{at}dental.gla.ac.uk
Received 21 February 2007
Accepted 14 May 2007
Abbreviations: ABPA allergic bronchopulmonary aspergillosis; AmpB, amphotericin B; CF, cystic fibrosis; CLSM, confocal laser scanning microscopy; IA, invasive aspergillosis; PMIC, planktonic cell MIC; SMIC, sessile cell MIC; XTT, 2,3-bis(2-methoxy-4-nitro-5-sulfo-phenyl)-2H-tetrazolium-5-carboxanilide.
 |
INTRODUCTION |
|---|
 TOP INTRODUCTION METHODSRESULTS AND DISCUSSIONREFERENCES |
|---|
In recent times, the frequency of disseminated fungal infections has increased dramatically. Overall, A. fumigatus is now the second most common cause of fungal infection found in hospitalized patients, after Candida albicans (Ellis et al., 2000). Aspergillus conidia are usually eliminated efficiently by the innate and acquired immune systems. However, in immunocompromised patients, such as transplant, leukaemia and human immunodeficiency virus (HIV)-positive patients, A. fumigatus can cause a range of systemic diseases with mortality rates ranging from 30 to 90 % (Brakhage, 2005; Denning et al., 1998; Herbrecht et al., 2002). Pulmonary infection may also occur in other patients such as those with cystic fibrosis (CF). In these patients, infection with A. fumigatus may cause allergic bronchopulmonary aspergillosis (ABPA), a mycetoma (fungus ball) or invasive aspergillosis (IA) (de Almeida et al., 2006; Shibuya et al., 2004).
The initial establishment of chronic A. fumigatus infection involves the germination of conidia and subsequent hyphal invasion of the lung tissue (Filler & Sheppard, 2006). Histological and microscopic examination of bronchopulmonary lavage samples has revealed the presence of numerous A. fumigatus hyphae in the form of a complex multicellular structure (mycetoma), which is similar to the biofilms formed by Candida species (Ramage et al., 2001c, 2005). In contrast to the biofilms formed by Candida species, very limited information is currently available on the development and behaviour of A. fumigatus adherent multicellular communities and their response to antifungal treatment. To date, there are only two reports suggesting that Aspergillus species are able to grow and form biofilms (Beauvais et al., 2007; Villena & Gutierrez-Correa, 2006).
The purpose of this study was to investigate the growth characteristics of filamentous A. fumigatus multicellular communities through the development of an in vitro model that could be utilized to screen the growth characteristics of clinical isolates or mutants, and to examine the antifungal susceptibility profiles of complex biofilm-like structures (Ramage et al., 2002a, c).
|
METHODS |
|---|
|
TOPINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
|---|
Growth conditions and standardization of conidial inoculum. A. fumigatus was grown on Sabouraud dextrose agar at 37 °C for 72 h. Conidia were harvested by flooding the surface of the agar plates with 5 ml PBS (Oxoid) containing 0.025 % (v/v) Tween 20 and rocking gently. The conidial suspension was recovered and dispensed into a 5 ml sterile glass bottle. The conidia were counted using a Neubauer haemocytometer and adjusted to the required concentration in RPMI 1640 (Sigma) buffered to pH 7.0 with 0.165 M MOPS. All procedures were carried out in a HERASafe laminate flow cabinet (model K515; Kendro).
Biofilm formation. A. fumigatus biofilms were formed on commercially available, pre-sterilized, polystyrene, flat-bottomed, 96-well microtitre plates (Corning). Biofilms were formed by adding 200 µl of a standardized cell suspension in MOPS-buffered RPMI 1640 to each well for selected time periods (4, 8, 12, 24 and 48 h), and incubating statically at 37 °C. A minimum of 12 replicates was performed for each experimental parameter, plus suitable controls. At each selected time point, the medium was aspirated and the biofilms were washed thoroughly three times with sterile PBS by repeated pipetting to remove non-adherent cells.
Confocal laser scanning microscopy (CLSM). A. fumigatus biofilms were formed as described above on the surface of 13-mm-diameter sterile Thermanox plastic cell culture coverslips (Nunc) in 24-well tissue culture plates (Nunc). After incubation at 37 °C for various time periods (0, 2, 4, 8, 10, 12, 16, 18 and 24 h), the coverslips were washed in sterile PBS and stained using the LIVE/DEAD fluorescent stain (Molecular Probes), according to the manufacturer's instructions. The FUN1 component of the kit was used, which is a bright green fluorescent intracellular stain. This was applied to the washed biofilms for 20 min in the dark. The biofilms were then washed in PBS and mounted on a slide. The fluorescent filamentous biomass was examined using a Zeiss Axiovert LSM510 confocal microscope attached to an LSM510 laser scanning system with a 488 argon ion laser at x200 magnification. Sections of the xy plane were taken at 1 µm intervals along the z-axis to determine the depth of the biofilms and overall physical ultrastructure. Three-dimensional images were obtained using computer software.
Biofilm quantification. Biofilm biomass was assessed using a modified version of a protocol first developed by Christensen et al. (1985) and subsequently modified by O'Toole & Kolter (1998). At each time interval, the spent culture medium was removed from each well and the adherent cells were washed three times with PBS. These were air-dried and 100 µl of 0.5 % (w/v) crystal violet solution was added for 5 min. The solution was then removed by carefully rinsing the biofilms under running water until excess stain was removed. The biofilms were destained by adding 100 µl 95 % ethanol to each well. The ethanol was gently pipetted to completely solubilize the crystal violet for 1 min, the ethanol was transferred to a clean 96-well microtitre plate and the A570 was read (FLUO Star Optima fluorescence microplate reader; BMG Labtech). The absorbance values are proportional to the quantity of biofilm biomass, which comprises hyphae and extracellular polymeric material (the greater the quantity of biological material, the higher the level of staining and absorbance).
2,3-bis(2-Methoxy-4-nitro-5-sulfo-phenyl)-2H-tetrazolium-5-carboxanilide (XTT) reduction assay. A semi-quantitative measure of each biofilm was calculated using an XTT reduction assay, adapted from previous studies (Hawser et al., 1998). This is a metabolic reduction assay that measures the activity of cells and can be used to compare untreated cells with cells treated with antimicrobial agents, and has been used previously in studies to evaluate the antifungal sensitivities of filamentous fungi (Antachopoulos et al., 2006; Meletiadis et al., 2001b, c). Briefly, XTT (Sigma) was prepared in a saturated solution at 0.5 g l–1 in PBS. The solution was filter-sterilized through a 22 µm pore size filter, aliquoted and stored at –80 °C. Prior to each assay, an aliquot of stock XTT was thawed and menadione (10 mM prepared in acetone; Sigma) was added to a final concentration of 10 µM. A 100 µl aliquot of XTT/menadione solution was added to each well and to appropriate control wells to measure background XTT reduction levels. The plates were incubated in the dark for 3 h at 37 °C and the colour change was measured using a 490 nm filter in a microplate reader (FLUO Star Optima; BMG Labtech). The colorimetric change in the XTT reduction assay directly correlates with the metabolic activity of the biofilm.
End-point susceptibility testing
M38-A broth microdilution testing.
Planktonic cell MICs (PMICs) were evaluated using the CLSI (formerly NCCLS) M38-A standard methodology (NCCLS, 2002). The antifungal agents itraconazole (Sigma), voriconazole (Pfizer), amphotericin B (AmpB; Bristol Myers Squibb) and caspofungin (Merck) were prepared as stock solutions in DMSO (Sigma) and diluted in MOPS-buffered RPMI 1640 to working concentrations. Microtitre plates containing 100 µl of each antifungal were serially diluted twofold to produce a concentration range of 0.03–16 mg l–1. A range of conidial suspensions was then prepared in MOPS-buffered RPMI 1640 to a final concentration of 0.4–5x104 conidia ml–1 and 100 µl was added to each well. The plates were incubated for 48 h at 35 °C. The PMIC for each antifungal was defined as the lowest concentration that produced complete visible inhibition of growth. Testing of these isolates was performed in quadruplicate.
Modified M38-A broth microdilution testing. For antifungal susceptibility testing of biofilms [sessile cell MICs (SMICs)], conidia were prepared as described above and standardized to a density of 1x106 conidia ml–1 in MOPS-buffered RPMI 1640. Biofilms were formed by pipetting standardized conidial suspensions into selected wells of a microtitre plate and incubating for 24 h at 35 °C, as described above. After growth, the medium was aspirated and non-adherent cells were removed by thorough washing of the cells (three times) using sterile PBS and gentle pipetting. Residual PBS from each well was removed by blotting with paper towels. All antifungals (itraconazole, voriconazole, AmpB and caspofungin) were prepared as described above to provide a working concentration of 512 mg l–1 in MOPS-buffered RPMI 1640. These were serially diluted twofold (1–256 mg l–1) directly into adjacent wells and the challenged cells were incubated statically for a further 48 h at 35 °C. A number of antifungal-free wells and biofilm-free wells were also included to serve as positive and negative controls, respectively. SMICs were determined as 50 and 90 % reduction in metabolism compared with the untreated control using the XTT reduction assay described above. Testing of these isolates was performed in quadruplicate.
Statistical analysis. The absorbance values of individual biofilms were compared by one-way analysis of variance and using the Bartlett's test for homogeneity of variances and the Bonferroni's multiple comparison post-test. A value of P <0.05 was considered to be significant. Analyses were performed using SPSS 13.0 for Windows.
|
RESULTS AND DISCUSSION |
|---|
|
TOPINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
|---|
Standardizing an in vitro model
To date, there have been no published accounts establishing that filamentous A. fumigatus grows as a biofilm. Histopathological evidence has indicated the presence of mycelial plugs and proliferation of hyphae with acute-angle dichotomous branching (Shibuya et al., 2004), which are complicit with the definition of a biofilm. There are many forms of aspergillosis, ranging from ABPA to IA, and despite different clinical presentations, morphological characteristics remain relatively similar and are typified by intricate mycelial networks. Formation of A. fumigatus mycelial aggregates that exhibit classic biofilm characteristics is reported here, i.e. an adherent microbial population, adherent to each other and/or surfaces or interfaces (Costerton et al., 1995). To our knowledge, this study is the first to examine submerged cultures of A. fumigatus in vitro in this growth modality.
A key factor that we noticed early in our studies was the critical importance of conidial seeding density. Clearly, the structural morphology and integrity of these multicellular structures was dependent on the concentration of conidia (ml medium)–1 (Figs 1
and 2), a phenomenon previously identified with C. albicans biofilm development (Ramage et al., 2001c). To produce adherent aggregated multicellular populations (biofilms) with a similar morphology to those seen during in vivo aspergillosis lung infection, we developed a model, initially examining conidial seeding density of A. fumigatus NCPF 7367 and four clinical isolates. First, we examined a serial dilution of conidial densities, ranging from 10 to 1x106 conidia ml–1, and examined the resultant multicellular structures after a 24 h inoculation using semi-quantitative metabolic and biomass assays. Fig. 1 illustrates that both the metabolic activity and the biomass of the biofilms exhibited a positive correlation with the conidial seeding density, although at the highest concentrations of conidia the biomass appeared to decline. We subsequently used CLSM, a non-invasive technique that enabled three-dimensional structural imaging and depth measurements of intact multicellular structures on Thermanox coverslips. The mature biofilms formed using a series of standardized conidial suspensions (1x104, 1x105 and 1x106 conidia ml–1) were evaluated by microscopy to investigate the optimal conidial inoculum concentration. Fig. 2(a) illustrates complex multicellular aggregates formed from three different conidial seeding densities after growth for 24 h, which ranged in depth from 117 to 360 µm. All of the filamentous multicellular structures exhibited acute-angle dichotomous branching to a varying extent. However, the conidial seeding density played an important role in the overall structural integrity of the biofilm structure. Biofilm stability was assessed by shear mechanical force by serially pipetting the biofilms with PBS during the washing procedure (results not shown). Either increasing or decreasing the conidial seeding density 10-fold led to significant differences in the depth of the resultant biofilms (P >0.05). At the highest conidial concentration (1x106 conidia ml–1), the biofilms were relatively thin (117 µm) and development of filamentous growth was severely restricted, resulting in less overall biomass (Fig. 2). Conversely, biofilms formed by fewer seeded conidia (1x104 conidia ml–1) were observed to be thicker (360 µm) and have longer mycelial frameworks. These were easily disrupted and removed by mechanical forces and were therefore not reproducible. The conidial concentration of 1x105 conidia ml–1 produced robust filamentous structures that were resistant to mechanical disruption (Fig. 2). Therefore, this concentration of conidia was selected, as the resultant biofilms exhibited reproducible characteristics that were amenable to the high-throughput testing required for screening of clinical isolates and defined mutants, or for testing the susceptibility of antifungal agents.
|
|
). A correlation between metabolism, biomass and hyphal development was demonstrated. The metabolic activity and the biomass of the biofilms were shown to increase over time. Microscopic analysis revealed that A. fumigatus did not form hyphae until approximately 8 h, in agreement with the results of Meletiadis et al. (2001a), who demonstrated germination at approximately 6 h using a similar microtitre plate method while examining filamentous growth characteristics of A. fumigatus. When the kinetics of biofilm formation was measured, their results were comparable to those in this study, with a lag phase of approximately 10 h. Hyphae began to intertwine forming a monolayer (10–16 h), followed by increased structural complexity over the subsequent 4–8 h (Fig. 4a). This was shown more clearly from the depth measurements taken over a 24 h period. Fig. 4(b) showed an exponential rise in the depth of the biofilm between 10 and 18 h, which then reached a plateau as development ceased and a steady state occurred. These biofilm development characteristics were observed for all isolates tested in this study. The development of the biofilm was slower than that of C. albicans biofilms; nevertheless, the overall characteristics of development were similar (Chandra et al., 2001; Ramage et al., 2001c). The slower initial development of mycelia may be related to the overall density of the cells, indicating a potential role for quorum sensing in multicellular A. fumigatus populations. This phenomenon has been demonstrated in C. albicans with the secreted molecule farnesol, which prevents hyphal development and subsequent biofilm formation (Ramage et al., 2002b).
|
|
).
|
In this study, five strains of A. fumigatus were examined for their susceptibility to a range of antifungal agents when grown as a complex mycelial structure, using a methodology previously employed for C. albicans biofilms (Ramage et al., 2001a). These agents included two azoles (itraconazole and voriconazole), a polyene (AmpB) and an echinocandin (caspofungin), which were serially diluted and used to challenge planktonic and sessile cells. Table 1
illustrates the results from both conventional planktonic CLSI M38-A susceptibility testing and the modified sessile susceptibility testing. The results indicated that caspofungin had a PMIC of 0.25 mg l–1, itraconazole 0.25–0.5 mg l–1, voriconazole 0.25–1 mg l–1 and AmpB 0.25–1 mg l–1. Testing of the sessile cells showed that AmpB was the most effective antifungal agent, with SMIC50 values ranging from <0.125 to 1 mg l–1 and SMIC90 values from 8 to 32 mg l–1. The other azole, voriconazole, exhibited increased efficacy (SMIC50 of 16–128 mg l–1). Caspofungin showed poor overall activity against adherent multicellular A. fumigatus, with SMIC50 and SMIC90 values of 64–128 and >256 mg l–1, respectively. Itraconazole was ineffective against all adherent multicellular populations (SMIC50 and SMIC90 values of >256 mg). Overall, AmpB was the most effective against sessile cells at the lowest concentrations, followed by voriconazole, caspofungin and itraconazole. It has been shown previously that azole antifungals were ineffective against C. albicans and Candida dubliniensis biofilms, whereas echinocandins were the most effective (Ramage et al., 2001a, b, c). Hawser et al. (2001) reported that A. fumigatus isolates were more susceptible to echinocandins using conidial concentrations equivalent to the CLSI M38-A method combined with XTT measurements. However, in this study, with an increased cell biomass, caspofungin was ineffective. The inability to successfully treat IA patients with caspofungin has been reported elsewhere, with only 41 % responding during salvage therapy (Maertens et al., 2004). We note, however, that both voriconazole and AmpB had the ability to reduce cellular viability by over 50 % at relatively low concentrations (Table 1). Therefore, although total death was not achieved, there was a certain degree of efficacy against these tenacious multicellular structures, which has been reported previously for other in vitro fungal biofilms (Ramage et al., 2002c). This may be an important observation when these agents are used empirically, as they may have a role in preventing the establishment of a mature biofilm. More work will be required to elucidate this.
How do these results relate to clinical practice? A recent review of empirical antifungal therapy in neutropenic patients compared 13 studies. The success rates of the treatments reported showed variations from 31 to 86 % (Martino & Viscoli, 2006). This variability in outcome may be as much to do with the patient population, the weakness of the indications for treatment and the consequent difficulty in establishing objective and reproducible end points for comparisons as the effectiveness of the antifungal agents. The time that treatment is started in relation to the development of the mature biofilm may also impact on the outcome. The optimal time for starting antifungal therapy in neutropenic patients remains undetermined, although most experts recommend waiting until day 5 or 7 of persistent fever (Bennett et al., 2003). It may be more prudent to start treatment before the multicellular structure has established, but this will require further in vitro studies. In CF, there is recognition that A. fumigatus in sputum cultures in the absence of ABPA may be a pathogen that can directly cause respiratory exacerbations. Antifungal therapy should be considered when deteriorating respiratory function is not responding to antibacterial therapy. Treatment with antifungal agents has been evaluated in these patients and an improvement in clinical condition observed (Shoseyov et al., 2006). Nevertheless, more studies are required before the effectiveness of antifungal agents in this group of patients can be evaluated fully.
Overall, this study demonstrated that A. fumigatus grows as a complex, multicellular biofilm and that the concentration of antifungal drug required for the effective treatment of these biofilm-related infections is distinct from assessment by the standard CLSI M38-A assay. This standard method of susceptibility testing does not give a true evaluation of the susceptibility of the disease-causing organism to a given antifungal agent. This may have implications for the diagnosis and management of patients with both invasive and non-invasive infections with this organism. Future studies to examine changes in gene expression of these biofilm-associated organisms may provide ways of elucidating new therapeutic options for controlling A. fumigatus biofilms in immunocompromised individuals.
|
ACKNOWLEDGEMENTS |
|---|
|
REFERENCES |
|---|
|
TOPINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
|---|
This article has been cited by other articles:
|
|
 |
|
  J. Chen, X. Song, P. Yang, and J. Wang Appearance of Anaphylactic Shock after Long-Term Intravenous Itraconazole Treatment Ann. Pharmacother., March 1, 2009; 43(3): 537 - 541. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. T. Cushion, M. S. Collins, and M. J. Linke Biofilm Formation by Pneumocystis spp. Eukaryot. Cell, February 1, 2009; 8(2): 197 - 206. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 E. Mowat, S. Lang, C. Williams, E. McCulloch, B. Jones, and G. Ramage Phase-dependent antifungal activity against Aspergillus fumigatus developing multicellular filamentous biofilms J. Antimicrob. Chemother., December 1, 2008; 62(6): 1281 - 1284. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. J. Seidler, S. Salvenmoser, and F.-M. C. Muller Aspergillus fumigatus Forms Biofilms with Reduced Antifungal Drug Susceptibility on Bronchial Epithelial Cells Antimicrob. Agents Chemother., November 1, 2008; 52(11): 4130 - 4136. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 A. S. Shehata, P. K. Mukherjee, and M. A. Ghannoum Comparison between the Standardized Clinical and Laboratory Standards Institute M38-A2 Method and a 2,3-Bis(2-Methoxy-4-Nitro-5-[(Sulphenylamino)Carbonyl]-2H-Tetrazolium Hydroxide- Based Method for Testing Antifungal Susceptibility of Dermatophytes J. Clin. Microbiol., November 1, 2008; 46(11): 3668 - 3671. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 K. Smith, A. Perez, G. Ramage, D. Lappin, C. G. Gemmell, and S. Lang Biofilm formation by Scottish clinical isolates of Staphylococcus aureus J. Med. Microbiol., August 1, 2008; 57(8): 1018 - 1023. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| INT J SYST EVOL MICROBIOL | J MED MICROBIOL | MICROBIOLOGY | J GEN VIROL | ALL SGM JOURNALS |
