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A Molecular Mechanistic Study on Bacterial Biofouling Using a Surface Forces Apparatus (SFA) Q. Lu*,**, X. Yang*, F. Teng**, J. Wang**, Y. Liu* and H. Zeng** * Department of Civil and Environmental Engineering, ** Department of Chemical and Material Engineering, University of Alberta, Edmonton, AB, T6G2W2, Canada (E-mail: [email protected]; [email protected]; [email protected]; [email protected]; [email protected]; [email protected]) Abstract The impact of bacterial surface polymers (i.e., extracellular polymeric substances (EPS), lipopolysaccharides (LPS)) on biofouling was investigated using a surface forces apparatus (SFA) combined with macroscopic bacterial adhesion experiments and zeta potential measurements. EPS were extracted from Burkholderia cepacia strain (PC184) while LPS were from Escherichia coli strain (JM109). The SFA force measurements showed that both EPS and LPS have bridging adhesion and growth ability on all the substrates tested. Two types of bacteria were tested for macroscopic adhesion: B. cepacia (PC184, PC184 rml, PC184 bcek) and E. coli (JM109), of which the three B. cepacia strains have different EPS-producing capacities and E. coli JM109 has significant amount of LPS but with very limited EPS on its surface. It is evident from this work that (1) the adhesion abilities of B. cepacia strains on polycarbonate substrate were positively related to their EPS producing capacities, and (2) the adhesion efficiency of E. coli JM109 was dependent on the interaction forces between its surface LPS and different substrates (i.e., polystyrene, mica and aminopropyltriethoxysilane-modified mica). Our results provide new insights into the fundamental understanding of molecular interaction mechanism of bacterial surface polymers and their role in biofouling. Keywords Biofouling; extracellular polymeric substances; lipopolysaccharides; surface forces apparatus INTRODUCTION The undesirable accumulation of microorganisms (e.g., bacteria) on wetted surfaces by adhesion, growth, and biofilm formation is known as biofouling. Biofouling is very detrimental to our lives and many engineering systems (Hori & Matsumoto 2010; Camesano et al. 2007; Liao et al. 2004; Flemming et al. 1996; Flemming 1993). For example, biofouling on ship hulls causes extra energy consumption, high maintenance costs, and increased corrosion (Cao et al. 2011). Biofouling on membranes and water distribution systems demands costly replacement, decreases water quality and threats public health (Ivnitsky et al. 2007). Therefore, it is very critical to elucidate the biofouling mechanism for better control and development of new materials and strategies for antifouling. Bacterial surface lipopolysaccharides (LPS) and extracellular polymeric substances (EPS) are both believed to play very important roles in biofouling (Hori & Matsumoto 2010; Camesano et al. 2007; Tsuneda et al. 2003). Lipolysacaccharides are the most common moieties present in Gram-negative bacterial outer membranes; and bacterial adhesion was observed to decrease after the LPS was removed from bacterial surfaces (Abu-Lail & Camesano 2003). In addition, most bacteria secrete EPS, and EPS are the major mass constituents of biofilms (Cao et al. 2011; Flemming & Wingender 2001). An increase in EPS concentration was observed to automatically increase bacterial adhesion and biofilm formation on solid surfaces (Tsuneda et al. 2003). LPS and EPS are the major contributors to biofouling, however, the molecular interaction mechanism controlling biofouling still remains unclear. The surface forces apparatus (SFA) provides extremely sensitive measurements of surface-surface interaction forces as a function of absolute separation distance with a distance resolution down to <0.1 nm (Israelachvili et al. 2010; Leckband 1995; Israelachvili & Adams 1976; Lu et al. 2012), and thus it provides a unique and effective way to elucidate the molecular interaction mechanism during bacterial adhesion. In this study, SFA combined with macroscopic bacterial adhesion experiments, atomic force microscopy, and quartz crystal microbalance were applied to elucidate the interaction mechanism for the control of bacterial biofouling (Lu et al. 2011; Yang et al. 2012). MATERIALS AND METHODS Chemicals LPS extracted from Escherichia coli JM109 was obtained commercially from Invitrogen (Montréal, Québec, Canada). EPS components were isolated from the Burkholderia cepacia PC184 by ethanol extraction following the reported procedure (Gong et al. 2009). Mica sheets (ruby mica blocks, grade 1) were purchased from S&J Trading Inc. (Floral park, NY, USA). Aqueous solutions were prepared in 18.2 M∙cm deionized water (Millipore, Mississauga, ON, Canada) and filtered through 0.2 µm filters (Nalgene, Rochester, NY, USA) prior to use. SFA force measurements The interaction force-distance profiles between surfaces were determined using an SFA (Surforce LLC, Santa Barbara, CA) (Israelachvili et al. 2010; Leckband 1995; Israelachvili & Adams 1976; Lu et al. 2012). A schematic of the SFA experimental setup is shown in Figure 1. Two thin mica sheets were glued onto cylindrical silica disks, and mounted in the SFA chamber in a crossedcylinder configuration which is locally equivalent to a sphere near a flat surface. The interaction Figure 1. Schematic of SFA experimental setup: (a) symmetric configuration where two curved mica surfaces close to each other in EPS or LPS solution, (b) asymmetric configuration where polystyrene (PS)-supported LPS and another surface (mica, PS-, or aminopropyltriethoxysilane (APTES)-modified mica) close to each other in NaCl solution. The interactions between two surfaces were monitored by SFA in real-time by using multiple beam interferometry (MBI) employing fringes of equal chromatic order (FECO). force F between the two curved mica surfaces was measured as a function of absolute surface separation distance D, which was monitored in real-time using multiple beam interferometry (MBI) employing fringes of equal chromatic order (FECO). SFA force measurements were carried out at room temperature (23 °C) in two different configurations (Figure 1 panel a and b): symmetric and asymmetric cases. For adsorption experiments (symmetric case), EPS or LPS solution were injected in between two mica surfaces (Figure 1a) (Lu et al. 2011; Yang et al. 2012). For asymmetric case, LPS were deposited on polystyrene (PS) coated mica surface and the opposing surface is mica or PS- or aminopropyltriethoxysilane (APTES)-modified mica surface (Figure 1b) (Lu et al. 2011). Each measurement was repeated at least three times. Bacterial culture B. cepacia complex strains PC184, PC184rml, and PC184bceK were obtained from Dr. Jonathan Dennis (Department of Biological Science, University of Alberta, Canada). PC184 is a wild-type cepacian EPS producer. PC184rml is an isogenic mutant strain of PC184 in the rml region of the genome and produces a reduced amount of EPS (Vinion-Dubiel & Goldberg 2003). PC184bceK is an isogenic mutation of PC184 in the bceK gene and has no EPS producing capacity (Moreira et al. 2003). Of the three strains tested, PC184 produces the highest amount of EPS while PC184bceK produces the least. E. coli JM109 was kindly provided by Dr Mavanur Suresh (Faculty of Pharmacy and Pharmaceutical Sciences, University of Alberta, Canada). E. coli JM109 expresses a full LPS chain that consists of lipid A, core polysaccharides, and O-antigen and rarely contains EPS secretion. Bacteria were cultured in Luria-Bertani (LB) broth (Miller) at 37ºC on a rocking incubator (Excella E24 incubator shaker series, New Brunswick Scientific, Edison, New Jersey, USA) and harvested at mid-log-growth phase (16 hours). Cells were harvested by centrifugation (Avanti J-20I, Beckman Coulter, CA) at 3000 g at 4 °C for 10 min. The growth medium was then decanted and pellets were resuspended in buffer. The centrifugation-resuspension process was repeated three times to remove traces of growth media. Prior to macroscopic adhesion experiments, a final cell density of approximately 108 CFU/mL was determined by optical density (OD) of 0.1 using a UV/visible spectrophotometer (Varian, Inc., CA) at a wavelength of 600 nm. Bacterial adhesion Bacterial initial adhesion was examined either by flowing cells or by a static soaking method by putting targeted substrates in a bacterial suspension. Bacteria were counted either by plate counting method or bacterial staining with Syto 9 (5 µM, Invitrogen, Burlington, ON, Canada) for imaging and counting under a fluorescence microscope (Leica DMRXA microscope, Nikon Digital Camera – DXM1200, Nikon ACT-1 software 2.70). Each measurement was repeated at least three times. Zeta potential measurements The zeta potentials of B. cepacia complex strains in 0.01 M PBS were measured at room temperature (23 °C) with a zeta potential analyzer (ZetaPALS, Brookhaven Instruments Corp, NY). Each measurement was repeated using at least three samples for 10 times with 30 cycles for each assay. RESULTS AND DISCUSSION EPS and LPS adsorption on mica The SFA was employed to investigate the molecular interactions by injecting EPS or LPS solution between two mica surfaces in a geometry shown in Figure 1a. The two mica surfaces were first brought close to reach a “hard wall” distance where the thickness of confined polymers becomes asymptotic with increasing normal load or pressure, and then the mica surfaces were separated. The typical force-distance profiles determined during approach (■) and separation (●) are shown in Figure 2. Figure 2. Normal forces F (normalized by the radius of local curvature R) measured between two mica surfaces as a function of surface separation D during approach (■) and separation (●): (a) and (b) in EPS (20 µg/mL in 0.01 M PBS) at time t = 6 min and t = 90 min after injecting the EPS solution, respectively; (c) and (d) in LPS (10 µg/mL in 0.1 M NaCl) at time t = 6 min and t = 90 min after injecting the LPS solution, respectively. Inset in Figure 2d shows the “hard wall” distance as a function of adsorption time between two mica surfaces (n.b. the “hard wall” is defined as the mica-mica separation at which the thickness of the confined polymers becomes asymptotic with increasing the normal load or pressure) for EPS and LPS, respectively. Both EPS and LPS showed bridging adhesion and accumulative growth on mica. For EPS, an adhesion Fad/R = -0.50 mN/m (or adhesion energy Wad = Fad/1.5R ~ -0.11 mJ/m2) was measured 6 min after injecting the solution (Figure 2a). Successive measurements showed similar adhesion as the first measurement. The hard wall increased from ~5 nm for the initial measurement (t = 6 min) to ~30 nm after 90 min adsorption, while the adhesion force remained almost unchanged with time (Figure 2b). For LPS solution, an adhesion Fad/R = -0.70 (± 0.04) mN/m was measured 6 min after injecting the solution, as shown in Figure 2c. Successive measurements showed smaller adhesion than the first measurement, possibly due to conformation changes of LPS after the previous measurement. The hard wall distance shifted from ~12 nm for the initial measurement (t = 6 min) to ~20 nm after 90 min adsorption with the adhesion decreased to Fad/R = -0.30 (± 0.02) mN/m (Figure 2d). The SFA measurements indicate that both bacterial surface polymers EPS and LPS can adhesively interact with mica. The measured adhesion forces arose from bridging interactions between the two mica surfaces due to the adsorbed EPS or LPS, which is supported by the sustained adhesion associated with increased hard wall distance with longer adsorption time, where further adsorption of EPS or LPS onto pre-adsorbed polymers on mica occurs. EPS are highly hydrated mixtures and mainly consist of polysaccharides as well as other macromolecules such as protein, DNA, lipids and humic substances (Hori & Matsumoto 2010; Yang et al. 2012). The LPS in this work contains a full chain of hydrophobic lipid A, core polysaccharides, and highly hydrated O-antigen (Hori & Matsumoto 2010; Lu at al. 2011). The accumulative growth of hard wall distance was possibly driven by the hydrophobic and hydrogen-bonding attraction among the polymers. The low growth rates (0.3 nm/min for EPS, 0.09 nm/min for LPS) are possibly due to the steric repulsion among polymers due to the presence of highly hydrated polysaccharide segments and electrostatic repulsion due to the negatively charged groups on the polymer chains. Impact of EPS on bacterial adhesion The above SFA adsorption experiments clearly showed that the cepacia EPS can adhere and grow on mica surface and thus play a role during bacterial adhesion. The adhesion energy (-0.11 mJ/m2) is close to the adhesion energy measured using atomic force microscopy (AFM) between EPS from P. atlantica cells and polymer membrane surfaces reported (Fang et al. 2000; Frank & Belfort 2003; Li & Logan 2004). The macroscopic bacterial adhesion experiments using B. cepacia strains with different EPS-producing ability further confirmed the impact of EPS during adhesion (Figure 3a). The presence of EPS on wild type strain PC184 increased bacterial adhesion over the mutant strains PC184rml with little EPS and PC184bceK with no EPS on polycarbonate (PC) surfaces (Figure 3a). The close relationship between B. cepacia strains adhesion on PC and their EPS producing ability is consistent with the inherent interactions from the dominant electrostatic repulsion and EPS attraction to PC depending on bacterial EPS producing ability. PC surface is negatively charged in the buffer solution in this work (Kirby & Hasselbrink 2004). All B. cepacia strains exhibited negative zeta potentials (Figure 3a inset), i.e., negative surface charges under the conditions tested. With decreased bacterial EPS-producing ability or reduced EPS coverage on bacterial surfaces, the bacterial zeta potential became more negative and the electrostatic repulsion between bacteria and PC increased. The cepacian EPS of B. cepacia is mainly composed of neutral and hydrophilic sugars (Cescutti et al. 2000; Lagatolla et al. 2002) which can reduce the spatial negative charge density of the EPS producing bacterial cells by partially shielding bacterial surface charges. Therefore, it can be concluded that the reduced adhesion of PC184rml and PC184bceK is through two interactions: increased electrostatic repulsion and reduced EPS attraction to PC. Impact of LPS on bacterial adhesion The above SFA adsorption experiments confirmed that LPS can adsorb to different substrate surfaces. LPS is largely anchored in live Gram-negative bacterial membrane surfaces and normally plays a dominant role when EPS is not significantly secreted and covers bacterial surface. E. coli JM109 is Gram-negative bacterial strain with not much EPS secretion on the surface. LPS chains cover most of E. coli JM109 surface and determine its surface properties and adhesion behaviours. The macroscopic adhesion E. coli JM109 showed obvious substrate dependency, as shown in Figure 3b. E. coli JM109 shows the highest adhesion on APTES and lowest on PS among the three substrates tested. Figure 3. Macroscopic bacterial adhesion behaviours: (a) three B. cepacia bacteria (PC184, PC184 rml, PC184 bcek) on polycarbonate disks in flow cells flowed by bacterial suspension in 0.01 M PBS for 10 min, followed by plate counting; (b) E. coli bacteria on three different substrate surfaces, polystyrene (PS)-coated mica, mica, and aminopropyltriethoxysilane (APTES)-coated mica by static soaking in bacterial suspensions in 0.1 M NaCl for 30 min, followed by cell counting using fluorescent microscopy. Inset in Figure 3a shows the zeta potential of three B. cepacia bacteria. Inset in Figure 3b shows the interaction force-distance profiles between LPS and three different substrates. The macroscopic bacterial adhesion is positively related to the interaction forces between LPS layer and three substrate surfaces (Figure 3b inset). By depositing LPS on hydrophobic PS surface by attaching hydrophobic lipid A onto PS and extending hydrophilic polysaccharides into surrounding solution, the LPS layer mimics the LPS on bacterial surface (Lu et al. 2011). Therefore, the interaction between PS-supported LPS and a substrate (Figure 1b asymmetric configuration) mimics the interaction between bacterial surface LPS and the substrate. The SFA measurements showed that PS-supported LPS have strong adhesion to APTES, weak adhesion to mica, and repulsion to PS (Figure 3b inset). The close relationship between macroscopic E. coli JM109 adhesion and PS-supported LPS interaction with the three substrates indicates that in addition to EPS, LPS plays a very important role in bacterial adhesion. E. coli JM109 is negatively charged under the tested solution condition (Li et al. 2004; Burks 2003), and LPS is negatively charged mainly due to the phosphate and carboxyl groups (Lu et al. 2011). PS, mica and APTES are negatively charged, nearly neutral, and positively charged, respectively (Lu et al. 2011). Therefore, the dominant interaction forces between E. coli JM109 or PS-supported LPS and the three substrates are electrostatic interactions, and the LPS contributes most to the E. coli JM109 adhesion measured. CONCLUSIONS Using a molecular force probe technique SFA and macroscopic/microscopic adhesion characterization, we have studied the impact of bacterial surface polymers, EPS and LPS, on bacterial adhesion. Both EPS and LPS contribute significantly to bacterial adhesion and affect the biofilm growth. The surface properties of bacteria strains and solid substratum play important roles in controlling the bacterial adhesion. 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