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Yarrowia lipolytica is an ascomycetous dimorphic fungus that exhibits biofilm mode of growth. Earlier work has shown that biosurfactants such as rhamnolipids are efficient dispersants of bacterial biofilms. However, their effectiveness against fungal biofilms (particularly Y. lipolytica) has not been investigated. The aim of this study was to determine the effect of rhamnolipid on a biofilm forming strain of Y. lipolytica. Two chemical surfactants, cetyl-trimethyl ammonium bromide (CTAB) and sodium dodecyl sulphate (SDS) were used as controls for comparison.
The methylene blue dye exclusion assay indicated an increase in fungal cell permeability after rhamnolipid treatment. Microtiter plate assay showed that the surfactant coating decreased Y. lipolytica biofilm formation by 50%. Rhamnolipid treatment disrupted pre-formed biofilms in a more effective manner than the other two surfactants. Confocal laser scanning microscopic studies showed that biofilm formation onto glass surfaces was decreased by 67% after sub-minimum inhibitory concentration (sub-MIC) treatment with rhamnolipids. The disruption of biofilms after rhamnolipid treatment was significant (P<0.05) when compared to SDS and CTAB.
The results indicate a potential application of the biological surfactant to disrupt Y. lipolytica biofilms.
Yarrowia lipolytica earlier referred to as Endomycopsis lipolyticaSaccharomycopsis lipolytica or Candida lipolytica is a hemiascomycetous fungus belonging to the Saccharomycetales order. It is often isolated from environments that are rich in hydrophobic substrates . The organism inhabits soil , seawater  and refrigerated meat products . The fungus is found in the oral cavity, pulmonary tract and intestines of healthy individuals. It is also an opportunistic pathogen that causes oral candidiasis, candidemia and catheter related infections . From biomedical point of view, the eradication of this organism thus becomes important. The fungus forms biofilms on different surfaces in the presence of a variety of substrates . It is well known that microorganisms in the biofilm mode of growth often resist a variety of antimicrobial agents. There is thus a need to explore alternative means of disrupting biofilms. A variety of chemicals including biocides and surfactants have been used to control biofilms . Chemical surfactants find applications in areas of medical care. For example, cetyl trimethyl ammonium bromide (CTAB) is used as a disinfectant in medical settings . Sodium dodecyl sulfate (SDS) is effective by mediating leakage of cellular material from microorganisms .
Widespread use of chemical surfactants is discouraged due to their inherent toxicity. In this context, biosurfactants are being favored . The latter group of surfactants offer several advantages in being relatively non-toxic, effective under different environmental conditions and in being biocompatible [11,12]. Biosurfactants have been used to disrupt bacterial biofilms [13,14]. However the reports on the efficacy of biosurfactants on fungal biofilms are limited . We hypothesized that rhamnolipids may be effective against biofilms of Y. lipolytica. The yeast strain used in the current investigation forms biofilms on a variety of water-soluble and -insoluble substrates. The objective of this work was therefore to test the effectiveness of rhamnolipids in (i) preventing biofilm formation and (ii) in disrupting pre-established biofilms of Y. lipolytica. The results have been compared with two chemical surfactants.
The chemical and biological surfactants displayed antifungal activity against the cells of Y. lipolytica NCIM 3589. Rhamnolipid and CTAB displayed a minimum inhibitory concentration (MIC) of 5%±0.1w/v and minimum fungicidal concentration (MFC) value >10%±0.1w/v, while SDS showed MIC and MFC values of 0.62%±0.05w/v. SDS was more effective as an antifungal agent compared to rhamnolipids and CTAB. This anionic surfactant is known to possess detergent and antimicrobial properties . The surfactant permeabilizes cells by targeting the cytoplasmic membranes and by affecting membrane-bound enzymes . Rhamnolipids are anionic biosurfactants that disrupt cells by interacting with the phospholipid components of the biological membranes [17,18]. Rhamnolipids derived from Pseudomonas aeruginosa are known to possess antifungal activity against some plant pathogenic fungi such as Cercospora kikuchii, Cladosporium cucumerinum, Colletotrichum orbiculare, Cylindrocarpon destructans, Magnaporthe grisea and Phytophthora capsici. The rhamnolipids inhibited spore germination and prevented hyphal growth in P. capsici at concentrations of 50μgml−1. A cationic surfactant, CTAB displayed lower antifungal activity towards Y. lipolytica as compared to SDS or rhamnolipids. In the present investigation, lower antifungal activity of CTAB could be a result of reversal of fungal cell surface charge and not due to cell lysis, as observed with SDS .
Rhamnolipids display antimicrobial and surfactant properties . They are known to increase cell permeability of P. aeruginosaEscherichia coli and Bacillus subtilis. In the present case, rhamnolipids were found to be less effective in increasing the cell permeability of Y. lipolytica, as compared to SDS. SDS displayed higher permeability even at concentrations lower than MIC values (Figure1, Table1). In Figure1, white arrows point towards non-permeabilized cells and the black arrows depict permeabilized cells. The increase in permeability observed after treatment with SDS may be due to the possible formation of molecular aggregates in the membrane and creation of trans-membrane pores . CTAB was least effective in permeabilizing cells. This reduced permeability of the dye could be due to the robust nature of the fungal cell walls .
Anti-adhesive activity of microbial surfactants has been reported earlier [11,14,23]. Pre-coating of microtiter plate wells with the surfactants effectively reduced the development of Y. lipolytica biofilms. Adhesion of Y. lipolytica cells to the microtiter plate wells was inhibited to 50% with rhamnolipids at MIC concentration (5%) (Figure2, asterisk). Rhamnolipids showed significant anti-adhesive ability (P<0.05) as compared to CTAB that inhibited 29% at MIC value of 5% (Figure2, black arrow). With SDS at MIC (0.625%) the inhibition was less than 10% (Figure2, black triangle). Rhamnolipids are known to decrease Listeria monocytogenes attachment when preconditioned onto Polytetrafluoroethylene (PTFE) surfaces . We have also recently demonstrated the ability of rhamnolipids in disrupting Bacillus pumilus biofilms by removing exopolymeric substances . SDS is known to display anti-adhesive ability by affecting the hydrophobic bonds that help in attachment of cells to the surfaces . In the present study however, SDS showed lower anti-adhesive ability than rhamnolipids. Although CTAB is reported to bind to the negatively charged microbial surfaces, alter their surface charge and prevent the binding of the cells to the surfaces , this surfactant was not as effective in preventing Y. lipolytica adhesion. In the present study, the anti-adhesive effect of rhamnolipids was found to be significantly higher as compared to SDS and CTAB suggesting the potential of rhamnolipids as anti-adhesive agents in the treatment of fungal biofilms.
The preformed biofilms of Y. lipolytica were treated with the surfactants for 1, 2 or 3h. Biofilms of Y. lipolytica formed for 3days in microtiter plate wells were disrupted effectively (55% with rhamnolipid, 35% with CTAB and 40% with SDS at respective MIC values) within 1h of treatment with the surfactants (Figure3a asterisk, arrow and triangle, respectively). At concentrations of surfactants lower than MIC, rhamnolipid displayed effective dispersion of biofilms (46%), followed by SDS (38%) and CTAB (25%). At higher concentrations (>2.5%), the effect of SDS was slightly better than that of the rhamnolipid. However, over a period of time, the efficacy of rhamnolipids was found to be similar to that of SDS (Figure3b and and3c).3c). CTAB was less effective in controlling biofilms, possibly due to its chemical interaction with fungal proteins present in the exopolymeric matrix . There are also reports that show the efficacy of CTAB over SDS in controlling bacterial biofilms . The present study showed that Y. lipolytica biofilms could be removed effectively by SDS compared to CTAB.
Biofilms of Y. lipolytica formed on glass slides for 3days were disrupted by using sub-MIC concentrations of the surfactants. The concentrations of surfactants were selected on the basis of the microtiter plate results. Rhamnolipids effectively disrupted Y. lipolytica biofilms on glass surfaces upto 76% with SDS, 53% and with CTAB 38% disruption was observed. The disruption with rhamnolipids was found to be statistically significant (P<0.05) as compared to SDS and CTAB. Rhamnolipids have earlier shown to be effective against bacterial biofilms , however there are limited reports on their effect on fungal biofilms. In the present study rhamnolipids were found to be effective in disrupting biofilms of Y. lipolytica compared to SDS and CTAB, suggesting their potential application as biofilm disrupting agents (Figure4). The effectiveness of rhamnolipids against biofilms even at low concentrations makes them a good candidate for therapeutic applications.
Rhamnolipids have potential to disrupt Y. lipolytica biofilms as compared to chemical surfactants, SDS and CTAB. The results suggest potential of rhamnolipid biosurfactants as anti-adhesive and preformed biofilm disrupting agents and their possible role in the treatment of fungal infections.
A biofilm forming strain of Y. lipolytica NCIM 3589 was used in the experiments . Cells were grown in 50ml YEPD broth (yeast extract 0.3%, peptone 0.5% and dextrose 1%) in 250ml Erlenmeyer flasks at 30°C on shaker at 150rpm for 24h.
Rhamnolipid biosurfactant, a mixture of mono and di-rhamnolipids was used . CTAB and SDS were obtained from HiMedia, India. Stock solutions of the surfactants were prepared in sterile distilled water; filter sterilized through 0.22μ filters and used further for experimentation.
MIC of surfactants against Y. lipolytica was determined by broth microdilution assay in sterile 96 well microtiter plates (Tarsons, India) . Briefly, pre-grown (36h) cells of Y. lipolytica were added to the microtiter plate wells containing YEPD medium to achieve the final cell numbers of 1×107 cells ml-1. Surfactants (rhamnolipid, SDS and CTAB) were added to these wells at varying concentrations (0.005 to 10%w/v). The final volume in the microtiter plate wells was maintained to 200μl. The plates were incubated at 30°C for 48h and after the incubation period, growth in presence of the surfactants was estimated as OD600 using a microtiter plate reader (Multiskan, Thermo Lab systems). Wells without surfactants and those lacking the cells were used as controls. The MFC was determined by streaking the culture grown in presence of different concentrations of the biosurfactant onto YEPD agar plates. The plates were incubated for 48h and growth was recorded. Minimum inhibitory concentration (MIC) was determined as the lowest concentration without visible growth and the minimum fungicidal concentration (MFC) as the lowest concentration showing no growth on the agar surface. The experiments were performed in triplicate and the mean values were obtained.
In order to determine the effect of surfactants on yeast cell permeability, the method of Hammer et al. (2004) was followed with few modifications . Y. lipolytica cells were grown in YEPD broth for 24h. The cells were harvested, washed twice with sterile distilled water and resuspended in PBS to achieve 107 cells ml-1. Aliquots were distributed equally in sterile flasks containing rhamnolipids, SDS or CTAB at MIC and sub-MIC concentrations. The flasks were incubated at 30°C under shaking conditions. After 1h, 100μl of these samples were withdrawn and to this 20μl of 0.05%w/v methylene blue (prepared in sterile distilled water) was mixed and incubated for 1min at room temperature. Methylene blue stained samples were placed onto the glass slides and the cells were examined microscopically by using an Axio Scope-A1 microscope with a photographic attachment (ProgRes ® Capture Pro 2.7) at a magnification of 400×. A minimum of 500 cells in consecutive visual fields were examined and the percentage stained cells was calculated manually . Cells untreated with the surfactants were used as controls for the experiment.
Surfactants (100μl containing 0.3-10%w/v concentration) were added to wells of the polystyrene microtiter plates and incubated for 12h at 4°C to facilitate effective coating . After the incubation period, the wells were emptied of residual surfactants, rinsed with sterile distilled water and air dried in a laminar air flow for 5min. Cells (100μl containing ~107 cells ml-1) of Y. lipolytica were added to the microtiter plate wells and incubated for 24h at 30°C. After the incubation period, the microtiter plate wells were emptied of the non-adherent cells and the plates were rinsed with sterile distilled water. The adherent cells were quantified by using the crystal violet assay . All experiments were carried out in triplicates with two biological replicates and average values indicating standard deviation are presented here.
Y. lipolytica biofilms were allowed to form in sterile polystyrene 96 well microtiter plate wells for 3days . After the incubation period, planktonic cells were removed and varying concentrations (0.3-10%w/v) of rhamnolipid, SDS and CTAB were individually added to the wells. The plates were further incubated at 30°C for 1, 2 or 3h. After each time interval, the microtiter plate wells were emptied of the non-adherent cells and rinsed with sterile distilled water. The residual biofilms were quantified by using the standardized crystal violet assay . All experiments were performed in triplicates with two biological replicates and the data is presented as average values indicating standard deviation.
Y. lipolytica biofilms were formed on sterile microscopic glass slides as described earlier . Cells were inoculated in sterile petriplates containing the growth medium (YEPD) to reach a cell density of 1×107 cells ml-1. Sterile microscopic glass slides were placed in the petriplates and incubated on a rocker at 30°C for 3days. After the growth period, glass slides were removed and placed in another petri dish containing growth medium supplemented with 0.3%w/v concentration of rhamnolipids, SDS or CTAB. Biofilms were treated for 1h in presence of the surfactants. Untreated biofilms were used as controls and the biofilm coverage thus obtained was considered to be 100%. The slides were removed, rinsed twice with sterile distilled water and stained with LIVE/DEAD BacLight staining kit containing SYTO9 and Propidium Iodide (PI) (Molecular Probes, Eugene, Oregon, US) as per the manufacturer’s instructions. The slides were observed under a CLSM (SP2 AOBS, Leica Microsystems, Germany). A 63× 1.2 NA water immersion objective was used with 488nm Ar laser excitation and 500–640nm band pass emission setting. Multiple (20) images were scanned and analyzed using the image processing software; ImageJ (http://rsb.info.nih.gov/ij). The observations were made in triplicates and representative images are presented here.
The effect of rhamnolipid on biofilms was analyzed statistically by the Students t-test and treatments were considered significantly different if P≤0.05.
CTAB, Cetyl-trimethyl ammonium bromide; SDS, Sodium-dodecyl-sulphate; MIC, Minimum inhibitory concentration; MFC, Minimum fungicidal concentration; PTFE, Polytetrafluoroethylene; NCIM, National collection of industrial microorganisms; YEPD, Yeast extract peptone dextrose; OD, Optical density; CLSM, Confocal laser scanning microscopy.
The authors declare that they have no competing interests.
DHD and SD performed the experimental work and acquired the data. YVN performed the CSLM analysis. VPV and SSZ made contributions to conception and design. ARK analyzed and interpreted the data. DHD, SD, VPV and SSZ have been involved in drafting the manuscript and revising it critically for important intellectual content. All authors have given final approval for the version to be published.
DHD is a currently a postdoctoral fellow at McGill University. SD is a post-graduate student at Institute of Bioinformatics and Biotechnology (IBB). YVN and VPV are Scientists at Biofouling and Biofilm Processes Section, Chemistry Group, Bhabha Atomic Research Centre, BARC Facilities, Kalpakkam, Tamil Nadu 603102 India. ARK and SSZ are Associate Professors at IBB, University of Pune, Pune 411007 India.
DD would like to acknowledge the research fellowship provided under the Bhabha Atomic Research Centre (BARC) and University of Pune collaborative research programme.