Bacterial extracellular polymeric substances and their effect on settlement of zoospore of Ulva fasciata

Colloids and Surfaces B: Biointerfaces 103 (2013) 223–230

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Bacterial extracellular polymeric substances and their effect on settlement of zoospore of Ulva fasciata Ravindra Pal Singh, Mahendra K. Shukla, Avinash Mishra, C.R.K. Reddy ∗ , Bhavanath Jha Discipline of Marine Biotechnology and Ecology, CSIR-Central Salt and Marine Chemicals Research Institute, Bhavnagar 364 002, India

a r t i c l e

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Article history: Received 15 September 2012 Received in revised form 13 October 2012 Accepted 22 October 2012 Available online 30 October 2012 Keywords: Biofilm EPS MALDI TOF–TOF MS Ulva fasciata Zoospores

a b s t r a c t The extracellular polymeric substances (EPSs) secreted by Bacillus flexus (GU592213) were estimated to have the molecular weight of approximately 1528 and 33,686 kDa with the elemental composition of Na, P, Mg, C, O, Cl and S. The 1 H NMR and FT-IR analysis of EPS confirmed the presence of different aliphatic and aromatic groups. The EPS was amorphous in nature with an average particle size of 13.969 ␮m (d 0.5) and roughness of 193 nm. The GC–MS analysis has revealed different monosaccharides such as fucose, ribose, xylose, galactose, mannose and glucose. Oligo and polysaccharides were detected with MALDI TOF–TOF MS. The bacterial EPS for the first time tested as a natural substratum for settle of zoospores of Ulva fasciata by incubating for various durations ranging from 2 h to 48 h. The zoospore settlement on EPS coated cover slips progressively increased with incubation time in axenic cultures over controls. The EPS, thus investigated in this study was found to facilitate the primary settlement of spores that play crucial role in recruitment of macroalgal communities in coastal environment including intertidal regions. © 2012 Elsevier B.V. All rights reserved.

1. Introduction Extracellular polymeric substances (EPS) can be referred to as a network of organic compounds (carbohydrate, proteins and nucleic acids) bound with cation and/or anion, and can be either loosely attached to the cell surface or tightly associated with the cells of producers [1]. EPS helps to hold the marine aggregates and keep their networks intact that eventually promote aggregate formation and subsequently leads to the biofilms formation [2]. Microbial biofilm forming communities provide primary biotic-substrata for settlement of different fouling prokaryotic and eukaryotic organisms. The normal morphological growth of many foliaceous green macroalgae has been reported to be controlled by the macroalgal associated bacteria [3,4]. Biofilm formation is a process of succession following in a sequential pattern. Initially, single cells attach to the surface and differentiate into complex of closed microcolonies separated by a network of open water channels [5]. These aggregates are centers of high microbial activity and are presumed to play a significant role in the carbon cycle [6]. EPS alter the surface properties of the bacteria themselves to either promote or prevent initial attachment to surface or cell aggregation [7]. Recently, it has been reported that EPS secreted by marine microbes

∗ Corresponding author. Tel.: +91 278 256 5801/256 3805x6140; fax: +91 278 256 6970/256 7562. E-mail address: [email protected] (C.R.K. Reddy). 0927-7765/$ – see front matter © 2012 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.colsurfb.2012.10.037

has enhanced the growth of marine eukaryotic flora [1]. In addition, the EPS produced by marine bacteria has been utilized as ingredient in food, pharmaceutical and petrochemical industry. It can also be used as bioremediation agents in environment management system [8,9]. Although the research on seaweed-bacterial association has been recently on raise, the ecological significance of EPS secreted by epi- and endophytic bacterial communities has not been well investigated. Further, the bacterial species involved in this process in many cases have also not been identified. A few studies dealing with bacterial biofilms have reported that they enhanced the settlement of zoospore but the role of EPS in settlement of zoospore is not understood [3]. In case of fouling organisms such as phytoplankton, benthic algae and larvae, the source material must find a suitable surface to settle and adhere to a substratum in a reasonable time frame, otherwise they will perish without completing their life cycle processes. Spore settlement is one of the most important developmental phases in the life cycle of fouling marine organisms [10]. The previous studies on zoospore colonization on bacterial biofilms have reported that the settlement takes place in three steps i.e. contact, temporary and irreversible adhesion [11]. Commonly, the key factor determining the strength of zoospore settlement process in different fouling microorganisms is a sticky material having either permanent or temporary adhesive properties [12]. Furthermore, zoospores of the marine macroalgae respond to a number of physicochemical characteristics of substratum including wettability, charge, surface chemistry and topography [12,13].

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Ulva fasciata is one of the most prevalent green alga with worldwide distribution. The propagation of this alga is mainly through tiny motile asexual zoospores. These spores often settle gregariously in order to form groups or colonies of cells. The biochemical studies investigated by several researchers across the world have shown its possible utilization as a promising food supplement [14] and recently have been projected for its possible utilization in biofuel production [15]. In this study, the EPS produced by an epiphytic marine bacterium Bacillus flexus was studied from the context of physicochemical properties employing a variety of analytical tools and techniques and, thereafter investigated its possible effect on settlement of zoospore of U. fasciata.

desiccators for overnight. Dried samples were mixed with pyridine and acetic anhydride (1:1, v/v) and refluxed at 100 ◦ C. On appearance of yellow color, mixture was added to ice water and extracted with ethyl acetate. The extracted solution was washed with Milli-Q, Na2 CO3 , and saturated CuSO4 to remove excess acetic anhydride and pyridine. Thereafter, anhydrous Na2 SO4 was added to remove water from organic layer. Organic layer was separated and kept in vacuum desiccators for overnight. For gas chromatography mass spectroscopy (GC–MS) analysis, a pinch of powder was dissolved in 15 ml of dichloromethane, filtered with Whatmann filter paper and injected into GC–MS (Shimadzu, QP2010). 2.3. Chemical properties of EPS

2. Materials and methods 2.1. Growth curve and EPS production B. flexus (NCBI accession number GU592213) an endophytic bacterium, was isolated from U. lactuca [4]. EPS production medium was prepared according to Singh et al. [9]. Sub samples of 5 ml aliquots were drawn at every 6 h interval and reading was taken at 600 nm for growth curve. Three replicates were performed for each test. The supernatant was obtained in stationary phase by centrifuging bacterial culture at 15,000 × g, 4 ◦ C for 30 min. Thereafter, supernatants were filtered with 0.45 and 0.25 ␮m pore size (Millipore filters, Bangalore, India) and the resultant bacterial cell pellet were freeze-dried and weighed. Precipitation of supernatant to obtain EPS and then, obtained EPS was washed following the protocol of Singh et al. [9]. Thereafter, EPS was collected, dried and dialyzed at 4 ◦ C for 24 h against Milli-Q water for desalting. The desalted EPS was recollected by centrifuging at 15,000 × g, 4 ◦ C for 30 min and lyophilized. The lyophilized EPS was stored in pure form for subsequent chemical and physical analysis. 2.2. Determination of molecular mass and their mass analysis For molecular weight determination, 50 ␮g (2% EPS in Milli-Q, w/w) EPS was applied to gel permeation chromatography (GPC, Water Allaince, model 2695) calibrated with ultrahydrogel columns 120 and 500 at 40 ◦ C and elution was monitored by a refractive index detector (2414). The molecular weight of EPS was calculated with standard Dextran (molecular weight; 5200–668,000 kDa) procured from PSS, USA. The desalted EPS was dissolved in acetonitrile (5%, w/v) and mixed with equal volume of matrix ␣-cyano-4-hydroxycinnamic acid (5 mg ml−1 ) for matrix assisted laser desorption–ionization (MALDI)-time-of-flight (TOF)–TOF mass spectroscopy analysis. It was performed on an Applied Biosystem 4800 MALDI TOF–TOF analyzer and conditions were maintained as previously described [8]. The spectrum obtained from 12 spot-sets in positive ion mode was taken for mass analysis of EPS. Centroids and de-isotoping spectra were analyzed using Data explorer software (Applied Biosystem, USA). Further, monosaccharide contents of EPS were estimated by alditol-acetate method [9,16]. In brief, the purified EPS (100 mg) was hydrolyzed in 5 ml of 2 N H2 SO4 in a sealed vial (Teflon-lined cap) at 100 ◦ C for 6 h. Thereafter, the acidic solution was neutralized by adding of BaCO3 and sample was concentrated upto 4–5 ml followed by addition of NaBH4 then reaction mixture was kept at 25 ◦ C for overnight. Subsequently, solution was passed through cation exchange resin column at elute rate of 5 ml min−1 and solution was again concentrated with methanol. This step was repeated thrice to remove completely borohydride in the form of methyl borate. The solution was evaporated, dried and kept in

For chemical analyses, lyophilized EPS were hydrolyzed and processed as previously described [17]. Total carbohydrate, protein and sulfate contents of EPS were calculated according to anthrone [18], Bradford [19] and Dodgson & Price methods [20] respectively. Three replicates were used for each assay. The standard curve of sulfate was established in the range of 2–20 ␮g ml−1 H2 SO4 . The protein present in the EPS was analyzed with 1-dimensional 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDSPAGE) according to Laemmli [21]. Protein bands were visualized by silver staining. Noise-decoupled 1 H nuclear magnetic resonance (NMR) spectrum was recorded at 200 MHz on a Bruker Avance II 200 spectrometer (Switzerland). About 50 mg of NaOH pellets were dissolved in 1 ml of D2 O followed by addition of 50 ␮g EPS. The spectra of the EPS solution were recorded at 25 ± 1 ◦ C. The conditions for 1 H NMR were 5000–5200 accumulations, pulse duration 5.9 ␮s, acquisition time 1.2 s and relaxation delay 6 ␮s. Fourier transformed-infrared spectroscopy (FT-IR) spectrum was recorded on Perkin Elmer (Spectrum GX) with a resolution of 4 cm−1 in 4000–400 cm−1 region. For FT-IR analysis, pellets of 6 mg of EPS were prepared with 500 mg KBr followed by pressing the mixture into a 16 mm diameter mold. The inorganic elements of EPS were analyzed with energy dispersive X-ray spectroscopy (EDX; Oxford Instruments, UK). X-rays emitted by matter in response to bombardment with charged particles present in the matter was examined using SEM-EDX [22]. 2.4. Physical properties of EPS An amount of 1 g sample was used for X-ray diffraction analysis (XRD) and performed on X-ray powder diffractometer (Philips X’pert MPD, The Netherlands) instrument equipped with ˚ Cona PW3123/00 curved Ni-filtered CuK␣ source ( = 1.54056 A). ditions were maintained as described previously [8]. The dried EPS sample was mounted on a quartz substrate and diffraction peaks were plotted as degrees of 2 value (half of the scattering angle measured from the incident beam). The peaks of diffracted X-rays were calculated with Bragg’s law d=

 2 sin 

crystallinity index (CIxrd ) was calculated according to Ricou, Pinel, & Juhasz [23]. CIxrd =

˙Acrystal ˙Acrystal + ˙Aamorphous

Thermal gravimetric (TG) analysis of EPS was carried with Mettler Toledo TGA/SDTA System (Greifensee, Switzerland). Approximately, 5 mg of dried samples was applied to TGA and thermogram was obtained in the range of 30–500 ◦ C at rate of

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10 ◦ C min−1 under nitrogen atmosphere. The thermogram was plotted with weight (percentage) loss vs. temperature. The particle size distributions were measured by laser diffraction (Malvern Mastersizer 2000, Malvern Ltd., Worcestershire, UK). The wettability of EPS measured with the DataPhysics Tensiometer of DCAT series connected with Software Module SCAT 32. The contact angle of EPS was calculated according to following formula:  powder contact angle : M2 T= Cp2  cos  where, T – time after contact,  – viscosity, C – material const., p – density,  – surface tension,  – contact angle, M – mass of liquid adsorbed on solid. Wettability of EPS was checked against seawater with three replicates. 2.5. AFM analysis AFM is a powerful tool to detect topographical variation and reveal more detailed surface properties than a morphological image. Glass cover slips were sterilized with mixture of 5 ml of 50% HCl and 5 ml of 50% of H2 SO4 (v/v) for 2 h followed by rinsing with autoclaved Milli-Q water. Thereafter, treated with ethanol and again washed with autoclaved Milli-Q water and stored in same for further uses. AFM analysis was carried out with NT-MDT-NTegra Aura (Russia) in semi-contact mode for EPS coated cover slips and zoospore settled onto EPS coated cover slips. 2.6. Settlement and adhesion of zoospores on EPS film Fertile thalli of U. fasciata were collected from Veraval (20.55◦ N and 70.20◦ E) Southern west coast of India during November 2010 and conditions for zoospores released from a single population were maintained according to Callow et al. [24]. The cover slips (22 mm × 22 mm) were submerged in a 50% methanol/50% HCl (v/v) mixture, concentrated HCl (2 h in each) and followed by washing thoroughly with autoclaved Milli-Q water for sterilization. Thereafter, 2 cover slips were submersed in 60 mm sterilized Petri plates with 4 ml of 1% EPS suspension and kept in desiccator to form complete dry film of 0.35 ± 3 mm. There were 7 replicates of this experiment which were successively used for the zoospore settlement assay having an incubation time of 2, 6, 24 and 48 h. About 25 ml aliquots (1.5 × 106 spore ml−1 ) of zoospore suspension were mixed to 90 mm sterilized Petri plates having 3 replicates of cover slips coated with EPS and kept in dark at 25 ± 1 ◦ C for settlement of zoospores. The control was also considered as cover slips without coated with EPS. There were different 90 mm sterilized Petri plates having 3 replicates of cover slips coated with EPS for different incubation time of interval. Unattached zoospores on coverslips coated with EPS and without EPS were removed by gentle washing with autoclaved seawater. Zoospores were stained with dilute carbol fuschin and counted with Olympus inverted microscope (model IX70) fitted with DP 72 camera after 2, 6, 24 and 48 h of incubation time interval. Settled spores on the glass slide were counted in 15 fields of view located at 100 mm intervals along the diagonal of each square, starting at the central corner with 3 replicates of each cover slips. The mean number of attached spores per square centimeter was calculated with 95% confidence of one way ANOVA. 3. Results and discussion 3.1. Growth and optimization of EPS production Although B. flexus showed an overall linear growth up to 36 h, there was an exponential growth between 3 and 36 h, and

Fig. 1. Bacterial growth and EPS produced by B. flexus.

thereafter it sharply declined (Fig. 1). The EPS produced by B. flexus was granular in texture and yield varied with duration of time. The EPS yield was found maximum during the late log phase. Unlike bacterial growth, the EPS yield showed a linear increase throughout the study and ranged from 7.66 ␮g mg−1 to 77.33 ␮g mg−1 dry cell weights at 6 and 48 h, respectively (Fig. 1). While EPS secreted by another closely related Bacillus licheniformis was ranged from 11.5 ␮g mg−1 to 60.2 ␮g mg−1 dry cell weight at 6 and 48 h, respectively [9]. The bacterial species of Vibrio also produced maximum EPS during log phage [8]. The methods employed for extraction of EPS from microorganisms depends on the nature of the EPS produced [1,25,26]. However, the amount of EPS produced by the organism depend on a number of complex controlling factors like nutrient depletion, cell physiology and changes in growth conditions and/or bacterial–seaweed interactions. 3.2. Determination of molecular mass and their analysis GP chromatogram of purified EPS produced by B. flexus showed two molecular mass peaks corresponding to 1528 and 33,686 kDa with 2.21 and 1.246 polydispersity, respectively (Fig. S1). GPC pattern of present EPS was somewhat similar to that of previously reported B. licheniformis having approximately 1540 and 44,565 kDa with 1.92 and 1.41 polydispersity [9]. While, EPS of dinoflagellate, Amphidinium carterae consisted of low molecular weight approximately 233 and 1354 kDa with 1.195 and 1.107 polydispersity, respectively. However, a single fraction of EPS with GPC has also been reported from different bacteria. It has been observed that EPS secreted by Cronobacter sakazakii showed single chromatographic peak with 3760 kDa approximately with 1.017 polydispersity [27]. Additionally, higher molecular masses have also been reported for EPS produced by Idiomarina spp. (1.5 × 104 –1.5 × 106 kDa) and Bacillus strain B3-15 (6 × 105 kDa) [28]. It has been reported that MALDI TOF–TOF MS [26] is convenient method for rapid and sensitive structural analysis of oligo and polysaccharides. MALDI TOF–TOF MS spectra of EPS represented a series of masses in low range mode which correspond to deprotonated pentose (150 m/z), deoxy hexose and hexose (180 m/z) sugars with inorganic (Na, P, Mg, C, O, Cl and S) and organic groups (aliphatic and aromatic functional groups). A series of mass peaks 175.1294, 197.1588 and 251.1529 were obtained in low range mode for monosaccharide of this EPS and deviation from their actual mass value revealed that sugars were attached with different inorganic (Na, P, Mg, C, O, Cl and S) and organic groups (aliphatic and aromatic functional groups). Apart from these, a mass peak 343.2768 m/z was also observed in positive ion mid

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Fig. 2. (A) The fragmentation mass peaks of oligo and polysaccharides contained of purified EPS were detected with positive ion mode in MALDI TOF–TOF MS analysis and (B) monosaccharide contained of the purified EPS determined with GC–MS analysis.

range mode corresponding to that of hexose and pentose sugars while 440.4741 m/z peak revealed the presence of disaccharide (Fig. S2a and S2b). Another mass peak of 679.5378 consisted of 2 hexose and 2 pentose sugars. The mass peaks of 861.8578, 900.4803, 964.5788, 980.7354, 1056.5901, 1110.5988, 1145.7754, 1221.6525, 1316.9216, 1340.5688, 1362.2107, 1375.9656 and 1386.7 reveal the presence of oligosaccharides in the present EPS. Moreover, higher mass peaks of 4367.53 and 5397.9 revealed the presence of polysaccharides in EPS (Fig. S2c). In contrast, the higher molecular mass peaks were not found in EPS produced by C. sakazakii and Vibrio parahaemolyticus [8,27]. Additionally, hexose (glucose, galactose and mannose), deoxy hexose (fucose) and pentose sugars (ribose and xylose) were confirmed with GC–MS analysis (Fig. 2B). The amounts of sugars present in EPS were 14.01, 16.43, 3.35, 23.52, 37.09, and 5.6% corresponding to fucose, ribose, xylose, mannose, galactose and glucose respectively. The monosaccharide compositions of present EPS were found to be apparently variable from other EPS produced by a number of marine bacteria such as Flavobacterium, C. sakazakii, V. parahaemolyticus and Pseudoalteromonas sp. [29]. The uronic acid such as galacturonate was not found in the present EPS while found in the EPS secreted by Flavobacterium and Pseudoalteromonas species. High contained of the glucose sugar was present in the EPS of B. licheniformis [9], Flavobacterium and Pseudoalteromonas sp. [29] while mannose was in the content of EPS of V. parahaemolyticus [8] and C. sakazakii [27]. In contrast, only two monosaccharides sugars (galactose, 73.13% and glucose, 26.87%) were observed in the EPS of A. carterae [1]. Similar to A. carterae, present EPS also contained high ratio of galactose than other sugars. Thus, monosaccharide composition determined in this study was found to be quite contrasting from those of bacterial and cyanobacterial EPS reported till date [1,8,26,30]. Nielsen and Jahn [30] described that sugar play an essential role in EPS synthesis as an activated precursor.

thiocyanate ( SCN) and isothiocyanate ( NCS) groups respectively [32]. The thiocyanate group was not found in the EPS secreted from other sources such as V. parahaemolyticus [8], C. sakazakii [27] and A. carterae [1]. A peak at 1649 cm−1 indicated the characteristic IR absorption of alkenyl stretched (C C) and/or aryl substituted (C C) of polysaccharides [33]. Absorbance band 1418 cm−1 was representative of functional groups such as methylene and methyl anti-symmetrical and/or symmetrical bending [32]. The absorption peak at 1138 cm−1 was detected due to both sulfate ion and C-O stretching bend in polysaccharides [31]. The absorption at 996 cm−1 was identified as C C and C H vibrations of polysaccharides and nucleic acids [33]. The carbohydrate ring peaks were also detected at 847 and 620 cm−1 . Further, disulfide bond (S S) of the protein was indicated at 620 cm−1 (Fig. 3). The FT-IR spectrum of EPS in this study confirms the presence of various different functional groups as compared to previously reported EPS [1,8,9,26,27]. The 1 H NMR spectrum exhibited various anomeric signals for chemical shift of functional group present in the EPS [34]. The chemical shift at ı 1.0–1.258 ppm arouses from the methyl protons of 6 deoxy sugars of polysaccharides. The chemical shift of functional group (R2 CHOR) was observed at 3.2–4.3 ppm. On the other hand, stretching of N H group of protein was observed

3.3. Chemical analysis of EPS FT-IR spectroscopy was used to characterize functional groups within the EPS and showed a broad peak at 3417 cm−1 , a characteristic of polymeric OH stretching [31]. Relatively strong absorption peaks at 2334, 2184 and 2098 cm−1 represented cyanate ion,

Fig. 3. FT-IR analysis of purified EPS produced by B. flexus.

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Fig. 4.

1

H NMR analysis purified EPS produced by B. flexus.

at 1.3 ppm while 0.8–1.2 and 1.1–1.5 ppm represent the alkanes and alkenes respectively (Fig. 4). These chemical shift of different groups confirmed glycocalyx nature of EPS produced by B. flexus. Similar characteristic spectral peaks of 1 H NMR were also observed in biopolymers obtained from different sources including bacteria, diatom and dinoflagellate [1,8,9,27]. The ratio of total carbohydrates, protein and sulfate content in EPS secreted by B. flexus was found to be 62, 23 and 15 mg g−l dry cell weight respectively. Protein and sulfate content of the present EPS is higher than the previously reported EPS from B. licheniformis and C. sakazakii [9,27]. Nevertheless, the carbohydrate content of EPS was higher than sulfate and proteins, a feature that corroborates with previous report of Zhenming & Yan [35]. There are also studies wherein stated that protein content has been higher than carbohydrate content in EPS [30]. Protein moiety of EPS involved in exopolysaccharides production and/or enhanced the production of EPS [36]. Protein loosely associated to EPS was not well resolved because of interference with polysaccharides considering the much higher carbohydrate/protein ratio. Although, Cao & Hu [36] reported that predominant proteins in bound EPS with micro-organisms were estimated in the range of 30–40 and/or 60–90 kDa. In this study, the 1-dimensional 10% SDS-PAGE revealed that protein content in EPS has consisted of two different polypeptides chains with approximately 12 and 42 kDa (Fig. S3). It has been reported that extracellular matrixes (carbohydrate and protein) play an important role in stabilizing biofilm structure by forming electrostatic bonds with multivalent cations [2]. The previous reports especially revealed that extracellular protein might have more involved than polysaccharides in electrostatic bonds inside biofilms because it has relatively high content of negatively charged amino acids and also help to bind various cation (Na and Mg) [37]. Elemental qualitative and quantitative analysis by SEM-EDX revealed the weight and atomic percentage of seven elements (S, Na, P, Mg, Cl, C and O) present in EPS (Table S1). The distribution of cations such as Na and Mg in the EPS suggest their bonding to negative charge of sulfate groups and/or thiol group of the protein and thus, better formation of biofilm. The sulfate was present as a functional group in the polysaccharides, confirmed its anionic character in the marine environment [38]. The cation Mg was not found in the EPS of closely related other Bacillus species [9] and C. sakazakii [27] while contained the cation Ca which might be played similar role in those EPS. Moreover, in this study various different masses were observed by the MALDI TOF–TOF MS may be due to pentose and hexose sugars of EPS attached with different elements that detected with SEM-EDX (Fig. 2A). The results obtained from MALDI TOF–TOF MS, FT-IR, 1 H NMR and SEM-EDX for present EPS are significantly different from the previously reported from other sources [26,39]. 3.4. Physical properties of EPS Particle size distribution is related to flowability, moldability, compressibility and die-filling characteristics of a powder that

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defines the relative amounts of particles present in the EPS. The EPS in this study constituted varied particle sizes from 2.856 (d 0.1) to 40.845 (d 0.9) ␮m with an average size from 12.245 ␮m (d 0.5) to 1.0005 m2 g−1 specific surface areas (Fig. S4). While EPS produced from B. licheniformis was constituted of particle size from 5.274 (d 0.1) to 68.447 (d 0.9) ␮m with an average size from 24.977 ␮m (d 0.5) to 0.63771 m2 g−1 specific surface areas [9,26]. It is revealed that particle size of the present EPS was smaller than previously reported EPS [9,26]. Besides this, particle size approximately similar to EPS produced by dinoflagellate A. carterae [1] while quiet distinct from those EPS produced from other bacteria such as C. sakazakii and V. parahaemolyticus [8,27]. The XRD profile of EPS obtained from B. flexus exhibited the characteristic diffraction peaks at 32.1◦ , 33.8◦ , 38.6◦ and 48.8◦ with 2.8, 2.7, 2.3 and 1.8 A˚ d-spacing respectively (Fig. S5). The XRD pattern revealed the amorphous nature of EPS with CIxrd 0.304 crystallinity area. The ratio between sharp narrow diffraction and broad peaks were used to calculate the amounts of crystallinity [40]. XRD pattern of the present EPS somewhat found similar to previously reported EPS from different source [9,26] which were confirmed that EPS always amorphous in nature. TGA showed degradation pattern of EPS from B. flexus that took place in three steps. Initially 6.44% loss in the weight of EPS was observed between 100 and 200 ◦ C then approximately 52.6% and 41% of degradation was observed at 270 ◦ C and 400 ◦ C in second and third phase respectively (Fig. S6). This result corroborates with EPS of Bacillus sp. I-450 where degradation was taken place in three steps [34]. However, Kavita et al. [8] reported that EPS produced from V. parahaemolyticus followed two degradation patterns in TGA analysis. High level of carboxyl group in the EPS increased degradation of first phase as it is bound to more water molecules [41]. Additionally, the detection of S O, C O S and carboxyl groups in the present EPS could be responsible for binding of more water molecules and contributed to more weight loss in the TGA analysis. 3.5. Atomic force microscopy (AFM) analysis Physical properties of EPS chain can be determined by measuring parameters like shape, persistent length and end-to-end distances of the polysaccharide chain using AFM. Now, AFM analysis is extensively being used for the topological characterization of EPS from different source [1,9,42]. The 3D view of EPS can be analyzed in live condition using AFM which is being more advantageous over SEM. An AFM micrograph of the 2.0 ␮g ml−1 solution of the EPS following deposition on cover slips showed that cover slips surface was covered with irregular EPS aggregates of varying dimensions (as represented in two and three dimensional in Fig. 5A and B, respectively). Although, these compounds appear nearly homogeneous, closer inspection (Fig. 5C) revealed that each compound consists of L shaped with 1.5 ␮m length. Recently, pearl necklace shaped EPS was obtained from mixed culture aerobic sludge granules [42]. Average roughness was 193 nm and histograms of the AFM demonstrating the topological distribution pattern of particle of EPS onto cover slips before and after 48 h of incubation (Fig. S7). Zoospore trapped with EPS was depicted in the Fig. 5D. The results obtained from GPC collaborated with homogeneous distribution of EPS onto the cover slips and particle size distribution agreed with AFM. 3.6. Settlement and adhesion of zoospores on EPS film It has been reported that settlement of zoospores of Ulva and Enteromorpha spp. increased toward bacterial biofilm [3,24,43,44]. Moreover, it is observed that not only AHLs producing bacterial biofilms but also topology of the co-polymers of different organic coating enhance or reduce settlement of the Ulva zoospore [45].

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Fig. 5. AFM analysis of [two (A) and three dimensional view (B) before experiment set up] purified EPS. (C) Amplification of the square section from (A) and (D) incubation of zoospore and EPS after 48 h of incubation. Zoospore stained with carbol fuschin and settled manually counting with Olympus inverted microscope. (E) Control and (F) incubation of zoospore with EPS.

Ederth et al. [46] prepared methylated-galactoside-terminated alkanethiol self-assembled monolayers which reduced the settlement of zoospore. Similarly, Cho et al. [47] made fluorine-free, amphiphilic and nonionic surface active block copolymers (SABCs) with antifouling properties. In addition to this, surface charge of the co-polymer also enhances or reduces settlement of the Ulva zoospore. Ederth et al. [48] used cationic oligopeptide selfassembled monolayers (SAMs) for zoospore settlement assay. The U. linza zoospores interacted strongly with lysine- and argininerich SAMs as compared to acid-washed glass. In another study, Krishnan et al. [49] used hydrophobic fluorinated and hydrophilic PEGylated block co-polymer and reported that surface wettability of the material decreased the zoospore settlement as hydrophobic surface gave strength of adhesion of cells to substrates. Despite, these many studies have been carried out but effect of the bacterial EPS on the settlement of Ulva zoospore has not been elucidated yet. Hence, present study provides clear evidence that

U. fasciata zoospores respond to EPS coated cover slip. It was first time observed that relative increase in the duration of incubation with zoospore successively enhanced zoospores settlement. Maximum settlement of zoospore (310,911 per cm2 ) was observed with EPS coated cover slips after 48 h of incubation over control (41,882 per cm2 ). There were significant differences (p ≤ 0.05) in the effect of EPS on spore settlement based on one-way ANOVA and the Tukey HSD test as compared to control (Fig. 5E and F, Fig. 6). Decrease in mobility of spore was observed after 2 h and after 48 h of incubation nearly all the zoospores were settled down. These spores could be counted using direct, manual counting rather than image analysis. Contact angle () 90 ± 0.2 of present EPS revealed that EPS contained hydrophobic as well as hydrophilic moiety. Similarly, Callow and coworker reported a positive correlation between numbers of spores attached and increasing contact angle (increased in hydrophobicity). It is reported that most pronounced response of spore settlement to wettability for the contact angle between 40◦

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Marine Laboratory, UK for his valuable suggestions on zoospore staining and their settlement assay. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/ j.colsurfb.2012.10.037. References

Fig. 6. The effect of EPS on zoospore settlement was significant at p > 0.05 (one way ANOVA), in comparison to control. Significant denoted as +.

and 80◦ which is agreement with our findings [24,49]. Different analytical methods such as SEM-EDX, FT-IR and NMR also confirmed that present EPS is charged in nature which further gave strength to spore settlement. The present study, for the first time demonstrated 48 h of incubation as compared to previously report where 60–70 min has been given for zoospore settlement assay [24,44–49]. Considering the fact that in their natural environment biofouling bacteria are continuously secreting EPS by which spore of different fouling species are settle with respect to different time interval [24,43]. The incubation of zoospore with EPS enhanced settlement might be due to sticky nature of the former. Present EPS was also rich in various elements and other polymeric compounds (polysaccharides and protein) as detected with analytical methods. The organic and inorganic contents of the present EPS may provide nutrients to phytoplankton and seaweed for their better survival and growth [1,9]. The presence of hydroxyl and carboxyl groups in EPS could be responsible for hydrophilic polysaccharide formation which serves as binding sites for divalent cations [1]. In a natural marine environment, the nutrients can interact with EPS in order to increase the rate of elements uptake. It also concentrates the dissolved organic compounds and making them readily available for microbial growth and their surroundings flora. EPS also facilitates a mechanism for “gliding” motility by the adhesion complex of a range of connector molecules that link the extracellular adhesive strands through the plasma membrane to an actin–myosin system [50]. 4. Conclusion The findings of this study revealed that EPS produced by B. flexus was distinct in terms of chemical composition, size, monosaccharide, oligo and polysaccharide contents, even from those of produced by closely related bacterial strains. The zoospore settlement assays conducted with EPS coated cover slips revealed higher colonization of spores than control. The studies exploring the effect of interaction between settlement of zoospore and bacterial EPS may provide deeper insights into ecological significance of such polymers.

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The financial support received from the Council of Scientific and Industrial Research (RSP 0016), New Delhi is gratefully acknowledged. The first author (Ravindra Pal Singh) gratefully acknowledges the CSIR, New Delhi (India) for awarding the Senior Research Fellowship. Special thanks to Dr. Ian Joint, Plymouth

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