Dusane et al. Aquatic Biosystems 2012, 8:17
http://www.aquaticbiosystems.org/content/8/1/17
AQUATIC BIOSYSTEMS
RESEARCH
Open Access
Disruption of Yarrowia lipolytica biofilms by
rhamnolipid biosurfactant
Devendra H Dusane1,3, Sushovan Dam1, Yarlagadda V Nancharaiah2, Ameeta Ravi Kumar1,
Vayalam P Venugopalan2 and Smita S Zinjarde1*
Abstract
Background: 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.
Results: 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.
Conclusion: The results indicate a potential application of the biological surfactant to disrupt Y. lipolytica biofilms.
Keywords: Biofilm, Biosurfactant, CTAB, Rhamnolipid, SDS, Yarrowia lipolytica
Background
Yarrowia lipolytica earlier referred to as Endomycopsis lipolytica, Saccharomycopsis 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 [1]. The organism inhabits soil [2], seawater [3] and refrigerated meat products
[4]. 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 [5]. 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 [6]. 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
* Correspondence: smita@unipune.ac.in
1
Institute of Bioinformatics and Biotechnology, University of Pune, Pune 411
007, India
Full list of author information is available at the end of the article
been used to control biofilms [7]. Chemical surfactants find
applications in areas of medical care. For example, cetyl trimethyl ammonium bromide (CTAB) is used as a disinfectant in medical settings [8]. Sodium dodecyl sulfate (SDS) is
effective by mediating leakage of cellular material from
microorganisms [9].
Widespread use of chemical surfactants is discouraged
due to their inherent toxicity. In this context, biosurfactants are being favored [10]. 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 [15]. 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.
© 2012 Dusane et al.; licensee BioMed Central Ltd. This is an Open Access article distributed under the terms of the Creative
Commons Attribution License (http://creativecommons.org/licenses/by/2.0), which permits unrestricted use, distribution, and
reproduction in any medium, provided the original work is properly cited.
Dusane et al. Aquatic Biosystems 2012, 8:17
http://www.aquaticbiosystems.org/content/8/1/17
Results and discussion
Minimum inhibitory concentration (MIC) and minimum
fungicidal concentration (MFC) values of 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% ± 0.1 w/v and minimum fungicidal concentration (MFC) value >10% ± 0.1 w/v, while
SDS showed MIC and MFC values of 0.62% ± 0.05 w/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
[16]. The surfactant permeabilizes cells by targeting the
cytoplasmic membranes and by affecting membranebound enzymes [16]. 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 [19]. The rhamnolipids inhibited spore germination and prevented hyphal growth in P. capsici at concentrations of 50 μg ml−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 [9].
Increase in cell permeability after treatment with surfactants
Rhamnolipids display antimicrobial and surfactant properties [14]. They are known to increase cell permeability of
P. aeruginosa, Escherichia coli and Bacillus subtilis [20]. In
the present case, rhamnolipids were found to be less effective in increasing the cell permeability of Y. lipolytica, as
Page 2 of 7
compared to SDS. SDS displayed higher permeability even
at concentrations lower than MIC values (Figure 1, Table 1).
In Figure 1, 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 [21]. 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 [22].
Effect of surfactant pre-coating on biofilm growth
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%) (Figure 2,
asterisk). Rhamnolipids showed significant anti-adhesive
ability (P<0.05) as compared to CTAB that inhibited 29%
at MIC value of 5% (Figure 2, black arrow). With SDS at
MIC (0.625%) the inhibition was less than 10% (Figure 2,
black triangle). Rhamnolipids are known to decrease
Listeria monocytogenes attachment when preconditioned
onto Polytetrafluoroethylene (PTFE) surfaces [24]. We
have also recently demonstrated the ability of rhamnolipids in disrupting Bacillus pumilus biofilms by removing
exopolymeric substances [14]. SDS is known to display
anti-adhesive ability by affecting the hydrophobic bonds
that help in attachment of cells to the surfaces [25]. 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 [26], 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
Sub-MIC
MIC
Figure 1 Morphological features of Y. lipolytica NCIM 3589 cells. Control cells (A and B); after treatment with rhamnolipid (C, D); SDS (E, F);
and CTAB (G, H) at sub-MIC and MIC concentrations, respectively, for 1 h. The sub-MIC and MIC concentrations of 2.5% and 5% w/v respectively
for rhamnolipid and CTAB and 0.3% and 0.62% w/v concentration of SDS were used. Bar indicates 10 μm. The upper panel indicates sub-MIC
concentrations and lower panel shows MIC concentrations of the surfactants tested. White arrow shows non-permeability of methylene blue
within the yeast cells whereas the black arrow depicts increase in cell permeability and uptake of the dye after treatment with surfactants.
Dusane et al. Aquatic Biosystems 2012, 8:17
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Table 1 Methylene blue dye exclusion by Y. lipolytica
cells post treatment with ½ MIC and MIC of surfactants
for 1 h
Surfactants
Cells stained at ½ MIC (%)
Cells stained at MIC (%)
Rhamnolipid
20 ± 2
25 ± 2
SDS
70 ± 4
95 ± 2
CTAB
7±2
14 ± 4
significantly higher as compared to SDS and CTAB suggesting the potential of rhamnolipids as anti-adhesive
agents in the treatment of fungal biofilms.
Disruption of preformed biofilms of Y. lipolytica
The preformed biofilms of Y. lipolytica were treated with
the surfactants for 1, 2 or 3 h. Biofilms of Y. lipolytica
formed for 3 days in microtiter plate wells were disrupted
effectively (55% with rhamnolipid, 35% with CTAB and
40% with SDS at respective MIC values) within 1 h of
treatment with the surfactants (Figure 3a 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 (Figure 3b and
3c). CTAB was less effective in controlling biofilms, possibly due to its chemical interaction with fungal proteins
present in the exopolymeric matrix [27]. There are also
reports that show the efficacy of CTAB over SDS in controlling bacterial biofilms [27]. The present study showed
that Y. lipolytica biofilms could be removed effectively by
SDS compared to CTAB.
Page 3 of 7
Biofilms of Y. lipolytica formed on glass slides for 3 days
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 [14], 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 (Figure 4). The effectiveness of rhamnolipids against biofilms even at low concentrations makes
them a good candidate for therapeutic applications.
Conclusions
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.
Methods
Culture and growth conditions
A biofilm forming strain of Y. lipolytica NCIM 3589 was
used in the experiments [6]. Cells were grown in 50 ml
YEPD broth (yeast extract 0.3%, peptone 0.5% and dextrose 1%) in 250 ml Erlenmeyer flasks at 30°C on shaker
at 150 rpm for 24 h.
Figure 2 Inhibition of Y. lipolytica NCIM 3589 biofilms by surfactants. Observations in microtiter plate wells after pre-coating with different
concentrations of rhamnolipid, SDS or CTAB. Control indicates biofilm formation by Y. lipolytica in microtiter plates untreated with the surfactants.
The OD values are normalized with reference to control (considered as 100%). Where asterisk (*), triangle (▲) and arrow (#) indicates MIC
concentrations of rhamnolipid, SDS and CTAB respectively.
Dusane et al. Aquatic Biosystems 2012, 8:17
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Page 4 of 7
Figure 3 Effect of surfactants on preformed biofilms of Y. lipolytica NCIM 3589. Observations with rhamnolipid, SDS and CTAB after (A) 1 h
(B) 2 h and (C) 3 h of incubation. Where, star (*), triangle (▲) and arrow (#) indicates MIC concentrations of rhamnolipid, SDS and CTAB
respectively.
Chemical agents
Determination of MIC and MFC values
Rhamnolipid biosurfactant, a mixture of mono and dirhamnolipids was used [14]. CTAB and SDS were obtained
from HiMedia, India. Stock solutions of the surfactants
were prepared in sterile distilled water; filter sterilized
through 0.22 μ 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) [14]. Briefly, pre-grown (36 h)
cells of Y. lipolytica were added to the microtiter plate
wells containing YEPD medium to achieve the final cell
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Page 5 of 7
Figure 4 Representative CLSM images of Y. lipolytica NCIM 3589 biofilms. Observations of control cells and cells treated with sub-MIC concentration
of rhamnolipid (2.5% w/v), SDS (0.3% w/v) and CTAB (2.5% w/v) for 1 h, showed live cells stained with SYTO9 and dead cells stained with PI. Bar indicates
30 μm.
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 200 μl. The
plates were incubated at 30°C for 48 h 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 48 h 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.
Fungal cell permeability analysis by methylene blue
dye exclusion
In order to determine the effect of surfactants on yeast cell
permeability, the method of Hammer et al. (2004) was followed with few modifications [28]. Y. lipolytica cells were
grown in YEPD broth for 24 h. 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
1 h, 100 μl of these samples were withdrawn and to this
20 μl of 0.05% w/v methylene blue (prepared in sterile distilled water) was mixed and incubated for 1 min 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 W 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 [28]. Cells untreated
with the surfactants were used as controls for the
experiment.
Effect of surfactant pre-coating on biofilm formation
Surfactants (100 μl containing 0.3-10% w/v concentration) were added to wells of the polystyrene microtiter
plates and incubated for 12 h at 4°C to facilitate effective
coating [23]. 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 5 min.
Cells (100 μl containing ~107 cells ml-1) of Y. lipolytica
were added to the microtiter plate wells and incubated
for 24 h 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 [14]. All experiments were carried out in triplicates with two biological replicates and average values
indicating standard deviation are presented here.
Disruption of preformed biofilms
Y. lipolytica biofilms were allowed to form in sterile polystyrene 96 well microtiter plate wells for 3 days [6]. 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 3 h. 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 [14]. All experiments
were performed in triplicates with two biological replicates
and the data is presented as average values indicating
standard deviation.
Confocal laser scanning microscopy (CLSM)
Y. lipolytica biofilms were formed on sterile microscopic
glass slides as described earlier [29]. 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
Dusane et al. Aquatic Biosystems 2012, 8:17
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and incubated on a rocker at 30°C for 3 days. 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 1 h 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 488 nm Ar laser excitation and 500–
640 nm 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.
Statistical analysis
The effect of rhamnolipid on biofilms was analyzed statistically by the Students t-test and treatments were considered significantly different if P≤0.05.
Abbreviations
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.
Competing interests
The authors declare that they have no competing interests.
Authors’ contribution
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.
Authors’ information
DHD is a currently a postdoctoral fellow at McGill University. SD is a postgraduate 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.
Acknowledgements
DD would like to acknowledge the research fellowship provided under the
Bhabha Atomic Research Centre (BARC) and University of Pune collaborative
research programme.
Author details
1
Institute of Bioinformatics and Biotechnology, University of Pune, Pune 411
007, India. 2Biofouling and Biofilm Processes Section, BARC Facilities,
Kalpakkam 603 102, India. 3Present address: Biocolloids and Surfaces
Laboratory, Department of Chemical engineering, McGill University, Montreal,
QC, Canada.
Page 6 of 7
Received: 29 May 2012 Accepted: 27 July 2012
Published: 27 July 2012
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doi:10.1186/2046-9063-8-17
Cite this article as: Dusane et al.: Disruption of Yarrowia lipolytica
biofilms by rhamnolipid biosurfactant. Aquatic Biosystems 2012 8:17.
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