hamnolipid mediated disruption of marine Bacillus pumilus biofilms
evendra H. Dusanea, Y. Venkata Nancharaiahb, Smita S. Zinjardea, Vayalam P. Venugopalanb,∗
Institute of Bioinformatics and Biotechnology, University of Pune, Pune 411 007, IndiaBiofouling and Biofilm Processes Section, Water and Steam Chemistry Division, BARC Facilities, Kalpakkam 603 102, India
r t i c l e i n f o
rticle history:eceived 4 February 2010eceived in revised form 17 June 2010ccepted 7 July 2010vailable online 13 July 2010
eywords:
a b s t r a c t
Removal of detrimental biofilms from surfaces exposed in the marine environment remains a challenge.A strain of Bacillus pumilus was isolated from the surface of titanium coupons immersed in seawater in thevicinity of Madras Atomic Power Station (MAPS) on the East coast of India. The bacterium formed exten-sive biofilms when compared to species such as Bacillus licheniformis, Pseudomonas aeruginosa PAO1 andPseudomonas aureofaciens. A commercially available rhamnolipid was assessed for its ability to inhibitadhesion and disrupt pre-formed B. pumilus biofilms. The planktonic growth of B. pumilus cells was inhib-
ited by concentrations >1.6 mM. We studied the effect of various concentrations (0.05–100 mM) of therhamnolipid on adhesion of B. pumilus cells to polystyrene microtitre plates, wherein the effectivenessvaried from 46 to 99%. Biofilms of B. pumilus were dislodged efficiently at sub-MIC concentrations, sug-gesting the role of surfactant activity in removing pre-formed biofilms. Scanning electron microscopy(SEM) confirmed the removal of biofilm–matrix components and disruption of biofilms by treatmentwith the rhamnolipid. The results suggest the possible use of rhamnolipids as efficient anti-adhesive and
with
biofilm-disrupting agents
. Introduction
Several species of Bacillus inhabit coastal and marine environ-ents, where they form important members of the marine bacterial
ommunities [1]. Members of this genus have been observed onitanium surfaces that are generally used for the manufacture ofondensers and other heat exchangers in power plants [2]. Theacteria that belong to Bacillus are resistant to environmental con-itions, such as low nutrient availability, irradiation and chemicalisinfectants [3]. They are known to produce exopolysaccharidesnd organic acids that accelerate corrosion of steel [4]. Micro-ial adhesion is the first step in the establishment of microbialommunities on surfaces, commonly termed as biofilms. Suchiofilms can potentially cause enormous damage by leading to theormation of more complex biofouling [5]. In industrial applica-ions that use seawater, problems such as reduced heat transfercross heat exchanger surfaces, mechanical blockages in coolingater pipes and enhanced corrosion of structural materials occurue to the growth of biofilms [6]. Biofilms and biofouling can be
ontrolled by mechanical, chemical and thermal treatments. Con-entional antifouling biocides have limited effectiveness againstiofilm bacteria in industrial settings [7]. Bacteria in biofilms areenerally protected against the onslaught of biocides [8]. Biofilm
formation is initiated when a group of primary colonizing bacteriaattach to a surface freshly exposed to the aqueous environment.The control of primary colonizing bacteria in particular is thusimportant in finding an effective solution. Along with the killingof biofilm bacteria, their removal from the surface is desirable[8]. Biosurfactants are quite effective for such combined applica-tions.
Biosurfactants of microbial origin are reported to have anti-adhesive and biofilm disruption abilities [9,10]. Fungal andbacterial biofilms have shown to be disrupted by enzymaticallysynthesized surfactants such as lauroyl glucose [11]. In recent years,rhamnolipids derived from Pseudomonas aeruginosa have emergedas an important group of biosurfactants with several applications;they have also been produced on a commercial scale [12]. More-over, they display diversity in structure, are environment-friendlyand are effective at low concentrations and under extreme condi-tions. They are safe and hence are being considered as an alternativeto conventional antimicrobial agents in a wide variety of indus-trial areas [13]. Despite their potential, there are only a few studieson the interactions of biosurfactants with bacterial cells and theirrole in anti-adhesion [14,12,15] and biofilm dispersion [10,16]. Todate, the applications of rhamnolipids have been studied mainly
with respect to disruption of clinically significant biofilms, and tothe best of our knowledge, information on rhamnolipid mediatedgrowth inhibition and removal of biofilms associated with marineenvironment either alone or in combination with biocides is lack-ing.
The objective of the present study was to assess the effectf rhamnolipid on the adhesion and biofilm formation by aotent biofilm forming bacterium isolated from condenser mate-ial exposed to seawater. The biofilm forming bacterium used in theresent study was isolated from titanium surfaces exposed near theeawater intake point of Madras Atomic Power Station (MAPS) andharacterized using 16S rRNA gene sequencing.
. Materials and methods
.1. Titanium coupon preparation and isolation of marineacteria
Titanium coupons (2.5 cm × 2.5 cm) were polished upto 400 gritnd immersed in seawater at Kalpakkam (12◦33′′ N and 80◦11′′ E)ear the cooling water intake point of Madras Atomic Power StationMAPS) on the Bay of Bengal coast of India. After 2 days, the couponsere removed, rinsed twice with phosphate buffer (0.1 M, pH 7.0)
o remove loosely bound cells. The biofilm was scraped into knownolume of phosphate buffer with a sterile nylon brush. The biofilmas disrupted by vortexing, serially diluted in phosphate buffer
nd suitable dilutions were plated on Zobell marine agar mediumHiMedia, India). The plates were incubated for 48 h at 30 ◦C. Sin-le bacterial colonies were transferred to flasks containing Zobellarine broth (HiMedia, India). The flasks were incubated at 30 ◦C
n a shaker at 120 rpm for 24 h.
.2. Biochemical characterization of the bacterial strain
Among the bacterial strains, a dominant one was selectedased on colony morphology. The selected strain was inoculated
n Zobell marine broth and incubated at 30 ◦C. Selected biochem-cal tests (such as glucose utilization, catalase, oxidase activity)
ere carried out using standard procedures. The tolerance of theacterial cells to different salt concentrations (1–5%, w/v) waserformed using NaCl supplemented in nutrient broth. The pro-uction of enzymes such as protease, lipase and amylase wasstimated on respective growth media. The bacterial cell surfaceydrophobicity was estimated by the salt aggregation test (SAT)17]. The susceptibility of B. pumilus to various antibiotics likemikacin, ciprofloxacin, carbenicillin, deoxycycline hydrochloride,entamycin, kanamycin, moxifloxacin, norfloxacin, piperacillin,pectinomycin, tobramycin, tetracycline, streptomycin, ampicillin,ethicillin, erythromycin, cefprime and penicillin-G was deter-ined using paper disks (HiMedia, India) on Muller Hinton Agar
MHA).
.3. Bacterial DNA isolation, amplification of 16s rRNA gene andhylogenetic analysis
As mentioned earlier, the marine bacterial isolate (BI304R) thatormed extensive biofilms was chosen and further characterizedy using biochemical and molecular techniques. Single isolatedolonies of the bacterial culture were taken from agar plate anduspended in lysis buffer. The bacterial genomic DNA was iso-ated using phenol chloroform method [18]. The purity of the DNA
as checked using agarose gel electrophoresis (Horizon 58, MD,SA). The 16S rRNA gene was amplified using universal forward
16F27N) (5′ CCA GAG TTT GAT CMT GGC TCA G 3′) and reverse16R1525XP) (5′TTC TGC AGT CTA GAA GGA GGT GWT CCA GCC′) primers. DNA amplification was carried out using PCR (Eppen-
orf, Germany). The PCR conditions used were: initial denaturationt 94 ◦C for 2 min, followed by 35 cycles of denaturation at 95 ◦Cor 1 min, annealing at 55 ◦C for 1 min, and extension at 72 ◦C formin, and a final extension at 72 ◦C for 10 min [19]. Amplifica-
ion of rRNA gene was confirmed by agarose gel electrophoresis.
: Biointerfaces 81 (2010) 242–248 243
The PCR product was purified using MinElute PCR purification kit(Qiagen, Germany) and the sequence was analysed at the Ribo-somal Database Project (RDP II, Michigan State University, EastLansing, MI) and National Centre for Biotechnology Information(Bathesda, MD) (http://www.ncbi.nlm.nih.gov/BLAST). Similar-ity matrix was prepared using the similarity matrix calculatoravailable at the RDP II site. The alignment of all the 16SrRNA gene sequences was done using CLUSTALW programat the European Bio Informatics server (European Bioinfor-matics Institute, Wellcome Trust Genome Campus, Hinxton,Cambridge, United Kingdom) (http://www.ebi.ac.uk/clustalw).The analysed sequence obtained was submitted to GenBank(http://www.ncbi.nlm.nih.gov/Genbank/), for which an accessionnumber (FJ938166) was obtained.
2.4. Adhesion and biofilm formation in microtitre plates
The adhesion and biofilm formation by B. pumilus BI304R wasperformed using pre-sterilized flat bottom polystyrene 96-wellmicrotitre plates. Two known biofilm forming strains, Ps. aeruginosaPAO1, Ps. aureofaciens, and a marine isolate, B. licheniformis, wereused for comparative purposes [11]. These cultures were grown inLuria Bertani (LB) broth and the overnight cultures (20 �l; effec-tively containing 5 × 106 cells) were added to each well of themicrotitre plate (Tarsons, India) containing 180 �l of LB medium.The plates were incubated at 30 ◦C for 4 h and 24 h. After incubation,the planktonic cells were discarded and weakly adherent cells wereremoved by gently rinsing thrice with phosphate buffer. Adhesionof cells and biofilm formation were quantified by using the crystalviolet assay as described by us earlier [11]. Briefly, before stain-ing, planktonic cells were removed from the wells of microtiterplates. The wells were washed with phosphate buffer, air dried andstained with crystal violet (0.2%, w/v; 200 �l) for 5 min. After incu-bation, the wells were washed with phosphate buffer, air-dried and200 �l of 95% (v/v) ethanol was added. The plates were read atOD600 using microtiter plate reader (Thermo Scientific, India). Allthe experiments were carried out in triplicate with suitable controls(uninoculated microtitre plates).
2.5. Biofilm formation on glass surfaces and confocal laserscanning microscopy (CLSM)
Biofilm formation by B. pumilus was studied on pre-sterilizedmicroscopic glass surfaces. The adhesion and biofilm formationwere analyzed after incubation for 4 and 24 h. Aliquots (200 �l)of overnight culture of B. pumilus were inoculated in sterile petriplates containing 20 ml of LB medium. Pre-sterilized microscopicglass slides were immersed into the medium as surface for biofilmformation [20]. The petri plates were incubated at 30 ◦C for 4 and24 h on a rocker. The glass slides were removed after 4 and 24 h,rinsed twice with phosphate buffer and stained with 0.01% acri-dine orange for 3 min. The slides were then rinsed twice withphosphate buffer and observed under a confocal laser scanningmicroscope (TCS SP2 AOBS) equipped with DM IRE 2-invertedmicroscope (Leica Microsystems, Germany) as described earlier[21]. A 40 × 1.25 NA water immersion objective was used forobtaining all images. The 488 nm Ar laser and a 500–640 nmemission detection bandwidth were used to excite and detect,respectively, the stained cells. Percentage area of colonization wasestimated using the digital image analysis freeware ImageJ1.29x,downloadable from the site http://rsb.info.nih.gov/ij. The grayscale
images were selected, spatially calibrated and segmented by inter-active thresholding, following which they were converted intobinary images. About 20 image stacks were collected from 24 h oldbiofilms and x–z sagital sections of biofilms were obtained usingthe Leica confocal software.
and 99.22% with B. pumilus strain CN1-3 (EU102277.1). Surfaceproperties, attachment behaviour and biofilm formation abilityof this bacterium on polystyrene and glass surfaces were furtherinvestigated.
Table 1Phenotypic characteristics of B. pumilus BI304R isolate.
Phenotype BI304R
Gram’s character PositiveShape Short rodsGrowth on 1–10% NaCl +Growth at temperature (◦C)
4 ◦C −30 ◦C, 45 ◦C +
Oxidase, catalase +Utilization of d-glucose, sucrose, galactose, fructose +Haemolysis of human RBCs +Production of enzymes (protease, lipase, amylase) +Salt aggregation test (SAT) HydrophobicSusceptibility to antibiotics (�g per disc)
The rhamnolipid containing 50% (w/v) mono-rhamnolipidC26H48O9, MW: 504, CMC: 10−4 M at neutral pH) and 50% (w/v)irhamnolipid (C32H58O13, MW: 650, CMC: 1.5 × 10−4 M at neutralH) was obtained from Jeneil Biosurfactant Co. (Saukville, Wiscon-in). Both the rhamnolipids are anionic biosurfactants produced bys. aeruginosa. These rhamnolipids have two head groups and twodentical hydrocarbon tails [22].
.7. Determination of minimum inhibitory concentration (MIC)f rhamnolipids
The antibacterial activity of rhamnolipids against the tropicalarine isolate of B. pumilus was determined by using the stan-
ard broth microdilution method. An overnight grown culture of B.umilus was diluted to 0.5 McFarland with the growth medium andhe wells of microtitre plates were inoculated. Rhamnolipid con-entrations (0.05–100 mM) were prepared in LB and added to theicrotitre plate wells. The plates were incubated at 30 ◦C for 24 h.fter the incubation period, growth in presence of rhamnolipid wasstimated using microtiter plate reader (Thermo Scientific, India)t OD600. Wells without rhamnolipids and those lacking the cellsere used as controls.
.8. Anti-adhesive action of rhamnolipids towards B. pumilusells on polystyrene surface
The anti-adhesive activity of rhamnolipids against the cells of. pumilus was estimated in 96-well microtitre plates. The cellsere allowed to adhere for 4 h at 30 ◦C as described earlier. After
he incubation period, different concentrations of rhamnolipids0.05–100 mM) were added and the plates were further incubatedt 30 ◦C for 1 h. Cells without rhamolipids served as control forhe experiment. After 1 h incubation period, the medium was aspi-ated, the wells were gently rinsed with phosphate buffer, air-driednd stained with (0.2%, w/v; 200 �l) crystal violet for 5 min [11].he microtiter plate wells were washed with sterile phosphateuffer, air-dried and 200 �l of 95% (v/v) ethanol was added. Thelates were read at OD600 using microtiter plate reader. The resultsere expressed in terms of percent cell adhesion compared to
hamnolipid-untreated wells, which were used as controls.
.9. Rhamnolipid-mediated disruption of pre-formed biofilms
Biofilms of B. pumilus were allowed to form in 96-well microtitrelates as described earlier. After 24 h incubation period, planktonicells were removed and the medium was replaced with LB con-aining different concentrations of rhamnolipids (0.05–100 mM).hese plates were further incubated at 30 ◦C for 24 h. The plank-onic cells were discarded at the end of incubation and the biofilmsere quantified by using the crystal violet assay as mentioned ear-
ier. The wells untreated with rhamnolipid served as control for thisxperiment. The values are expressed in terms of percent biofilmormed in comparison to untreated biofilms as control [11].
.10. Scanning electron microscopy (SEM)
Cells of B. pumilus were grown overnight in LB at 30 ◦C. Aliquotsf 200 �l of the culture (∼106 cells ml−1) were added to sterile petrilates containing 20 ml of LB medium and pre-sterilized micro-
copic glass slides as surfaces for biofilm formation. The platesere incubated at 30 ◦C for 24 h. After the incubation period, the
lass slides were removed and placed in LB containing rhamnolipid1.6 mM) and incubated further at 30 ◦C for 1 h and 24 h to deter-
ine the effect of rhamnolipids on the pre-formed biofilm. After
: Biointerfaces 81 (2010) 242–248
the incubation period, the glass slides were removed, rinsed twicewith phosphate buffer, air dried and treated with 4% gluteraldehyde(Sigma, USA) prepared in 1 mM PBS. The treated slides were keptin dark for 2 h. The slides were successively dehydrated through analcohol series (30, 50, 75% and absolute alcohol for 30 min each).The samples were air-dried and coated with platinum vapor (to athickness of ∼30 nm) and observed under a scanning laser electronmicroscope (Jeol 6360A LV, Japan). Biofilms without rhamnolipidtreatment served as controls.
2.11. Statistical analysis
The differences in biofilm formation between the selected bac-terial strains were estimated by one-way analysis of variance(ANOVA) using Origin 6.0 software. Biofilm disruption values wereevaluated statistically using Student’s t-test and treatments wereconsidered significantly different if p ≤ 0.05.
3. Results and discussion
3.1. Identification of the dominant marine biofilm bacterium
A surface, when exposed to seawater, is approached and col-onized by microbial cells. They establish microcolonies with thehelp of sticky exopolymeric substances (EPS), which further facil-itate association of other organisms. It is believed that the initialsettlers known as primary colonizing bacteria have a significantrole in the subsequent development of complex multi-speciesbiofilm communities, which ultimately leads to biofouling on sur-faces [23]. Different species of Bacillus are predominant in marineenvironments [1,2]. In the present study, a species of Bacillus wasisolated during early stages of biofilm formation in the sea. It wassubjected to morphological, biochemical and phenotypic charac-terization (Table 1). 16S rRNA sequencing of this bacterium showedits identity as Bacillus pumilus (Fig. 1). The gene sequence has beensubmitted to GenBank with the accession number FJ938166. The B.pumilus strain BI304 showed 99.23% sequence match to the closestrelative of B. pumilus clone 007.2 (accession number: EU515205.1)
(30), Tetracycline (30), Streptomycin (10)Ampicillin (25), Methicillin (30), Erythromycin (15) ICefprime (30), Penicillin G (10) R
(+) indicates positive, (−) indicates negative and S, I and R (sensitive, intermediateand resistant, respectively).
D.H. Dusane et al. / Colloids and Surfaces B
Fic
3a
s
Rhamnolipids are amphipathic anionic glycolipids which pos-
F2f
ig. 1. Phylogenic tree of Bacillus pumilus BI304R (GenBank accession No. FJ938166)solated from titanium surfaces immersed in seawater. Phylogenetic analysis wasarried out at RDP and NCBI site with the sequence alignment done using CLUSTALW.
.2. Adhesion and biofilm formation by B. pumilus on polystyrene
nd glass surfaces
The biofilm formation by B. pumilus was compared with twotandard biofilm forming bacteria (Ps. aeruginosa PAO1 and Ps.
ig. 2. (A) Adhesion and biofilm formation by B. pumilus (�); Ps. aeruginosa PAO1 ( ); P4 h, respectively, as estimated by the crystal violet assay. Error bars = ± SD. (B). Confocalormation after 24 h (b) on glass surfaces. Saggital section of the 24 h biofilm shows typic
: Biointerfaces 81 (2010) 242–248 245
aureofaciens) and a marine bacterium B. licheniformis. The initialadhesion after 4 h for all the isolates was more or less compara-ble (p > 0.05). However, after 24 h, the extent of biofilm formationby B. pumilus was higher as compared to Ps. aureofaciens or B.licheniformis [Fig. 2(A)]. There was a significant difference when24 h biofilm of B. pumilus was compared with that of Ps. aureo-faciens (p = 0.0001), B. licheniformis (p < 0.0001) and Ps. aeruginosa(p = 0.02).
CSLM images showed individual bacterial cells adhering to theglass surface after 4 h of growth and subsequently, developmentof microcolonies (24 h) [Fig. 2(B)]. The three-dimensional struc-ture of microcolonies formed by B. pumilus during 24 h of growthwas observed under CSLM. The x–z sagittal section of these micro-colonies showed internal voids necessary for nutrient and watertransport within biofilms. The biofilm formation as estimated bymicrotitre plate assay suggests better biofilm forming ability of B.pumilus on polystyrene surfaces as compared to the other knownbiofilm forming strains.
3.3. Minimum inhibitory concentration (MIC) of rhamnolipids
sess antimicrobial and surfactant properties. They have beenreported to show antimicrobial activity towards B. subtilis, Staphy-lococcus epidermidis and Propiniobacterium acnes at low MIC levels(<1.6 mM) [24]. The antimicrobial property of rhamnolipids is
s. aureofaciens 30–84 ( ) and B. licheniformis ( ) in microtitre plate wells at 4 andscanning laser microscopic images of B. pumilus cell adhesion at 4 h (a) and biofilmal microcolony formation. Bar represents 25 �m.
246 D.H. Dusane et al. / Colloids and Surfaces B
Fp
aotcptG
3
ta(s
ig. 3. Minimum inhibitory concentration (MIC) of rhamnolipid towards cells of B.umilus after 24 h at 30 ◦C. Error bars are ± SD.
ttributed to their effect on cell permeability, as reported in casef Ps. aeruginosa, E. coli and B. subtilis [25]. In the present case,he growth of B. pumilus was inhibited by rhamnolipids at con-entrations >1.6 mM (Fig. 3). These results showed resistance of B.umilus cells towards rhamnolipids at concentrations higher thanhose required to inhibit the growth of other Gram positive andram negative bacterial cells reported earlier [24].
.4. Anti-adhesive effect of rhamnolipids
Rhamnolipids, a class of glycolipid biosurfactants, are knowno decrease adhesive interactions [26]. We tested the anti-dhesion property of rhamnolipids at different concentrations0.05–100 mM) against B. pumilus. The attachment of B. pumilus wasignificantly inhibited (46–99%) at low concentrations of rhamno-
Fig. 4. (A) Percent cell adhesion (�); and (B) biofilm formation of B. pu
: Biointerfaces 81 (2010) 242–248
lipids (Fig. 4A). About 80% inhibition of B. pumilus cells attachmentto polystyrene surfaces was observed after 1 h of treatment withsub-MIC concentrations of rhamnolipid. In an earlier report, co-incubation of Ps. aeruginosa with 0.25 mM rhamnolipid showedprevention of adhesion and subsequent biofilm formation [27].In another report, the pretreatment of silicone rubber with asurfactant derived from Streptococcus thermophilus inhibited theadhesion of Candida albicans by about 85% [28]. Similarly, the sur-factant surlactin, produced by Lactobacillus also blocked the initialadhesion onto glass surfaces by the uropathogenic bacteria Ente-rococcus faecalis, E. coli and S. epidermidis by 77% [14]. Viewed inconjunction with the earlier reports, the present results highlightthe potential anti-adhesive property of rhamnolipids.
3.5. Disruption of pre-formed biofilm and removal of EPS
Biofilm mode of bacterial growth poses serious threat due toincreased resistance to antibiotics and biocides [6]. The presenceof EPS secreted by bacterial biofilms may help in neutralizing theantimicrobial agents. The disruption of pre-formed biofilms byeffective removal of exopolymeric substances using biosurfactants,either alone or in combination with antimicrobial agents, can beused as a strategy to treat detrimental biofilms.
In the present study, preformed biofilms of B. pumilus inpolystyrene microtitre plates were disrupted upto the extent of93% with rhamnolipids at a concentration of 100 mM (Fig. 4B).The preformed biofilms were effectively removed at concentra-
tions greater than the MIC values. At lower concentrations (0.4 mMor less), the biofilm disruption by the rhamnolipid was not quiteevident. At higher concentrations there was a dose-dependentincrease in biofilm disruption caused by the rhamnolipid (Fig. 4B).Statistical analysis using Student’s t-test showed significant differ-
milus ( ) in presence of different concentrations of rhamnolipid.
D.H. Dusane et al. / Colloids and Surfaces B: Biointerfaces 81 (2010) 242–248 247
F ed (24t the efa eprese
ewsbdStse
evbwabltric(Poctattmct
ig. 5. Representative SEM images depicting the effect of rhamnolipid on pre-formration of rhamnolipid for 24 h. Scale = 5 �m, High magnification images depictingfter treatment with 1.6 mM concentration of rhamnolipid for 1 h. [In C and D bar r
nce (p < 0.05) at higher concentrations of rhamnolipid (>1.6 mM)hen compared with untreated biofilms. Earlier studies also have
hown that rhamnolipids can bring about disruption of preformediofilms of Ps. aeruginosa [9]. Another biosurfactants, surfactin pro-uced by B. subtilis has been shown to inhibit biofilm formation byalmonella enterica, E. coli, and Proteus mirabilis [29]. Interestingly,he surfactant produced by Streptococcus thermophilus has also beenhown to be effective industrially for the control of fouling of heatxchanger plates in pasteurizers [30].
The removal of exopolymeric substances (EPS), as mentionedarlier, could be beneficial in the treatment of biofilms. Our obser-ations on disruption of preformed biofilm were further supportedy SEM images, which showed removal of EPS after 1 h of treatmentith the rhamnolipid (Fig. 5). The untreated B. pumilus cells grew
s biofilms (Fig. 5A) and showed the presence of cells tightly boundy EPS (Fig. 5C). However, treatment of the biofilms with rhamno-
ipid for 1 h resulted in removal of the EPS, revealing cells withinheir polymer cover (Fig. 5D). Treatment with rhamnolipids for 24 hesulted in the destruction of microcolonies (Fig. 5B and D). The SEMmages clearly show that biofilms and associated EPS of B. pumilusan be effectively removed after the treatment with rhamnolipidFig. 5). Such removal of the EPS matrix has been reported whens. aeruginosa was treated with mono-rhamnolipids. The authorsbserved that the outer cell membrane lipopolysaccharide proteinomplexes were removed by the rhamnolipid [31]. Interestingly,he removal of the EPS cover is known to enhance the effect ofntimicrobial agents and disinfectants [32]. The EPS removal in
he present case was probably aided by the surfactant activity ofhe rhamnolipid. Apparently, treatment of biofilms for 1 h would
ake them significantly more susceptible to antibacterial chemi-als. This would make them an attractive choice as chemical agentshat can enhance the efficacy of antibacterial agents, for example, in
h) biofilms of B. pumilus. (A) Control and (B) after treatment with 1.6 mM concen-fect on exopolymeric substances (EPS) produced by B. pumilus (C) Control and (D)nts 1 �m].
cooling water systems. One would expect that rhamnolipid treat-ment of cooling tower basin prior to biocide dosing would increasecell death and biofilm removal. Apart from external addition ofbiosurfactants for disruption of pre-existing biofilms, surface con-ditioning or coating could prove effective in preventing bacterialadhesion [14,12,29,30]. The results of the present study indicatethat rhamnolipids have potential to be used individually or in com-bination with biocides or antimicrobial agents for efficient removaland killing of detrimental biofilms.
4. Conclusions
A marine isolate from titanium surfaces exposed to seawaterwas identified as B. pumilus on the basis of phenotypic, biochemi-cal and 16S rRNA sequencing. The bacterium formed biofilms moreefficiently than Ps. aureofaciens and B. licheniformis in polystyrenemicrotitre plates and on glass surfaces. Rhamnolipid at sub-MICconcentrations was effective as an anti-adhesive and biofilm-disrupting agent against the B. pumilus isolate. Scanning electronmicroscopic observations confirmed the role of the rhamnolipidin removal of EPS and microcolonies from pre-formed biofilms.The results show that rhamnolipids are promising compounds forinhibition/disruption of marine biofilms and their potential forapplication in industrial systems (e.g. cooling water circuits) needsto be studied in detail.
Acknowledgements
DD would like to thank BARC-University of Pune collaborativeresearch programme for financial support. The authors would liketo thank Dr. Pattanathu KSM Rahman for providing the rhamno-lipids.
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