Environ. Sci. Technol. 2009, 43, 2308–2314
Influence of Extracellular Polymeric Substances (EPS) on Deposition Kinetics of Bacteria G U O Y U L O N G , † P I N G T I N G Z H U , †,‡ Y U N S H E N , † A N D M E I P I N G T O N G * ,†,‡ The Key Laboratory of Water and Sediment Sciences, Ministry of Education; Department of Environmental Engineering, Peking University, Beijing, 100871, P. R. China, and School of Environment and Urban Studies, Shenzhen Graduate School of Peking University, Shenzhen, 518055, P. R. China
Received September 1, 2008. Revised manuscript received February 5, 2009. Accepted February 7, 2009.
The significance of extracellular polymer substances (EPS) on cell deposition on silica surfaces was examined by direct comparison of the deposition kinetics of untreated “intact” bacteria versus those from the same strain but with EPS removal via cation exchange resin (CER) treatment using a quartz crystal microbalance with dissipation (QCM-D). Four bacterial strains, mutant Escherichia coli BL21 (gram-negative, nonmotile), Pseudomonas sp QG6 (gram-negative, motile), Rhodococcus sp QL2 (gram-positive, nonmotile), and Bacillus subtilis (grampositive, motile), were employed to determine the influence of EPS on cell deposition. Experiments were conducted in both monovalent (NaCl) and divalent (CaCl2) solutions under a variety of environmentally relevant ionic strength ranging from 1 to 100 mM at pH 6.0. The effectiveness of EPS removal via CER method was ensured by biochemical composition analysis of EPS solutions and further confirmed by FTIR analysis. Comparable zeta potentials were observed for untreated and CER treated bacterial cells in both NaCl and CaCl2 solutions, indicating that removal of EPS from cell surfaces via CER treatment did not affect the electrokinetic properties of the cell surfaces for all four strains. However, observed deposition efficiencies (R) were greater for untreated cells relative to those with CER treated cells across the entire ionic strength range examined in both NaCl and CaCl2 solutions for all four bacterial strains. These results strongly demonstrated that the removal of EPS from cell surfaces for all four strains decreased the deposition of bacteria on silica surfaces. This study clearly showed that the enhancement of cell deposition on silica surfaces due to the presence of EPS on cell surfaces was relevant to all bacterial strains examined regardless of cell types and motility.
Introduction Deposition of bacteria on mineral surface has great significance in the formation of bacterial biofilms (1-3), microorganism (e.g., pathogen) transport in porous media (4, 5), and in situ bioremediation (6-9). A number of biophysical and biochemical factors have been proposed to govern the deposition of bacteria on surfaces, such as bacterial cell type * Corresponding author phone: (86) 10-6275-6491; fax: (86) 106275-6526; e-mail:
[email protected]. † The Key Laboratory of Water and Sediment Sciences. ‡ School of Environment and Urban Studies. 2308
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(e.g., gram negative or gram positive) (10, 11), motility (e.g., the presence or absence of flagella and fimbriae) (12-16), hydrophobic interactions (11, 17-19), outer membrane proteins (20, 21), and lipopolysaccharides (LPS) (22-26). By examining the cell deposition behaviors of different heterotrophic bacterial strains with various amount of extracellular polymeric substances (EPS) in packed glass bead beds, Tsuneda et al. (27) proposed that the deposition of bacteria on surfaces can be enhanced by the presence of a larger amount of EPS. Very recently, Liu et al. (28) compared the deposition behaviors of three genetically mutant Pseudomonas aeruginosa strains with different EPS secretion capability and EPS composition in packed glass bead columns. The authors found that although the zeta potential measurements showed surface charges were similar for these three isogenic strains, the deposition rate coefficients of EPSproducing strains PAO1 and PDO300 were greater than those of EPS deficient PAO1 psl pel strain under the same conditions. This study also suggested that the presence of EPS on cell surfaces can significantly increase cell adhesion to porous media. However, the significance of EPS on cell adhesion to surfaces has never been tested by the direct comparison of bacterial deposition behaviors of “intact” cell (presence of EPS on cell surface) with those of bacteria from the same strain but with EPS removed from cell surfaces, and it requires investigation. EPS, comprised of a wide variety of organic compounds including proteins, polysaccharides, humic acid substances, lipid, DNAs (DNA), and so on, can be separated from cell surfaces by combining centrifugation with cation exchange resin (CER) extraction processes (29-34). With minor induced cell lysis, these extraction processes have been reported as the most effective methods to remove EPS from cell surfaces (30-34). Therefore, with the employment of centrifugation and CER extraction processes, direct comparison of the deposition behaviors of CER treated bacteria versus those of bacteria from the same strain but without CER treatment could be achieved to test the hypothesis that the presence of EPS on cell surfaces can contribute to the greater bacterial deposition to surfaces relative to those absent of EPS. Furthermore, the few previous investigations (27, 28) regarding the significance of EPS on cell deposition, e.g., Liu et al. (28) only concerned strains of one type (gram-negative and nonmotile). The relevance of the significance of EPS on cell deposition to strains of other cell types, e.g., gram-negative and motile, gram-positive, and nonmotile, and gram-positive and motile has not been explored and required examination. The objective of this paper is to fully examine the significance of EPS on bacterial deposition on silica surfaces via comparison of the deposition kinetics of untreated “intact” bacteria versus those from the same strain but with EPS removed from cell surfaces. Four bacterial strains, mutant Escherichia coli BL21 (gram-negative, nonmotile), Pseudomonas sp. QG6 (gram-negative, motile), Rhodococcus sp. QL2 (gram-positive, nonmotile), and Bacillus subtilis (BST) (grampositive, motile), respectively, were employed to study the influence of EPS on cell deposition. A quartz crystal microbalance with dissipation (QCM-D) was utilized to determine the initial deposition rates for all four untreated and CER treated bacteria strains in both NaCl and CaCl2 solutions over a wide range of ionic strengths (ranging from 1 to 100 mM) at pH 6.0. Our results demonstrated that for all four bacterial strains, regardless of cell types and motility, EPS on cell surfaces could improve cell deposition on silica surfaces. 10.1021/es802464v CCC: $40.75
2009 American Chemical Society
Published on Web 03/04/2009
Materials and Methods Cell Culture and Preparation. Four bacterial strains: mutant Escherichia coli BL21 (E.coli), Pseudomonas sp. QG6 (QG6), Rhodococcus sp. QL2 (QL2), and Bacillus subtilis (BST) were used in this study. E. coli BL21, a gram-negative and flagellar biosynthesis protein flgN insertion inactivation mutant nonmotile strain, was provided by Professor Zhang Chuanmao at school of life sciences, Peking University, China. QG6, a gram-negative and motile bacterial strain, and QL2, a gram-positive and nonmotile strain, were quinolinedegrading bacterial strains isolated from activated sludge from coke plant wastewater biological treatment process by Dr. Zhu Shunni at department of environmental engineering, Peking University, China. BST, a gram-positive and motile bacteria strain was obtained from Dr. Yao Maosheng at college of environmental sciences and engineering, Peking University, China. These four strains were grown according to the protocol described in the Supporting Information (SI). Cells were harvested by centrifugation (4000g for 10 min at 4 °C) upon reaching early stationary phase. Following centrifugation, the growth medium was decanted, and the pellets were washed three times in Milli-Q water to remove any residual growth medium (35, 36). The cell pellets were then resuspended in Milli-Q water. The centrifugation-resuspension process was repeated one more time. After that, each bacterial cell suspension was divided into two portions with the same volume. One portion of cell suspension was then repeated the centrifugation-resuspension process once more (8000g for 20 min at 4 °C). This portion of cell suspension was used as stock of untreated bacteria (with EPS). The other portion of cell suspension was used to prepare stock of treated bacteria (with EPS removed). This portion of cell suspension was transferred to a sterilized extraction beaker to release EPS via cation exchange resin (CER) technique developed by Frolund et al. (29). CER (Dowex Marathon C, 20-50 mesh, sodium form, Fluka 91973), which was soaked in Milli-Q water overnight prior to use, was added to the extraction beaker with a dosage of 2.5 g/g bacterial mass. The bacteria-CER suspension was then stirred at 600 rpm for 2.5 h at 4 °C. This was followed by a settlement of the suspension for 3 min to separate CER. The cell suspension was then transferred to centrifugation tube and the CER treated bacteria were collected by centrifugation at 8000g for 20 min at 4 °C. The CER treated bacterial pellets were then resuspended in Milli-Q water. Cell concentration and viability was determined according to the methods descried in the SI. To ensure the effectiveness of the EPS removal from cell surfaces via CER treatment, infrared spectra of untreated bacterial cells and those treated with CER technique were determined by using a Fourier transformed infrared spectroscopy (FTIR, Nicolet magna-IR750, U.S.) with resolution of 2 cm-1 and scan range of 650-4000 cm-1. To avoid KBr commonly used to homogenize samples might contribute to the possible shift of absorption band, cells predried in a freeze drier (FD-1, Beijing Detianyou Ltd., China) was transferred into a diamond chamber to directly acquire infrared spectra under the microspectroscopy mode. All spectra were collected in absorbance mode. The biochemical components of EPS solution collected from centrifugation after CER treatment, including dry weight (DW), volatile dry weight (VDW), proteins, polysaccharides, and humic acids were determined according to the methods provided in the SI. The electrophoretic mobilities of all untreated and CER treated bacterial strains were determined using a Zetasizer Nano ZS90 (Malvern Instruments, UK) at cell concentration of ∼107 cells mL-1. Measurements were performed in both NaCl and CaCl2 solutions with ionic strength ranging from
1 to 100 mM at room temperature (25 °C) and at pH of 6. The electrophoretic mobility measurements were repeated 9-12 times. Quartz Crystal Microbalance with Dissipation (QCMD). A QCM-D E1 system (Q-Sense AB, Gothenburg, Sweden) was utilized to examine the deposition kinetics of bacteria on silica surface. QCM-D experiments were preformed with 5 MHz AT-cut quartz sensor crystals with silica-coated surface (Batch 070625). Before each measurement, the crystals were soaked 30 min in a 2% SDS solution, rinsed thoroughly with Milli-Q water, dried with ultrahigh-purity N2 gas, and then oxidized for 30 min in a UV/O3 chamber (Bioforce nanosciences, Inc., Ames, IA). Deposition Experiment. The bacterial deposition experiments were performed in a flow-through mode, using a peristaltic pump (ISMATEC, Switzerland) operating in clockwise mode. Specifically, the pump was connected to the sensor crystal outlet, and the studied solutions, stored in a sterilized 50 mL polypropylene conical tube (Becton Dickinson, Franklin Lakes, NJ) connected to the sensor crystal inlet, were fed through the crystal sensor chamber at a flow rate of 0.1 mL-min-1. Salt solutions for the QCM-D experiments were degassed by sonicating for at least 15 min and subsequently kept in a water bath at 27 °C before being introduced into the E1 system thus to prevent the formation of air bubble in the crystal sensor chamber during the deposition experiments (37). Surfaces of cell (of all four strains) and bare silica crystal display bulk negative, yielding an overall repulsive electric double layer energy barrier between them (12-14, 38). To eliminate the repulsive energy barrier, silica crystals can be precoated with a layer of positively charged poly-L-lysine (PLL) hydrobromide (molecular weight of 70 000-150 000) (P-1274, Sigma-Aldrich, St. Louis, MO). Detailed protocol of silica surfaces modification with PLL is given by Chen and Elimelech (37) and can also be found in the SI. Bacteria (untreated and CER treated cells) deposition was examined both on bare silica surface (in the presence of an energy barrier) and positively charged PLL-coated silica surface (in the absence of an energy barrier). Experiments in the presence of an energy barrier (unfavorable condition) were conducted in both NaCl and CaCl2 solutions at six ionic strengths (ranging from 1 to 100 mM) at pH 6.0 (adjusted with HCl). For all experiments performed in the presence of an energy barrier, the QCM-D system was pre-equilibrated with salt solution at desired ionic strength (NaCl or CaCl2) for a minimum of 30 min to establish a stable baseline (the drift of average normalized frequency was less than 0.2 Hz within 30 min) prior to the injection of the bacterial electrolyte suspension. After pre-equilibration, cell suspension at desired ionic strength and pH at desired cell concentration (the influent concentration for bacterial strains E.coli and QL2 was set to be ∼2.5 × 107 cells mL-1, whereas, the influent concentration for bacterial strains QG6 and BST was set to be ∼7.5 × 107 cells mL-1) was injected into the crystal chamber. Experiments in the absence of an energy barrier (favorable condition) were conducted on PLL-coated silica surface at a 1 mM ionic strength solution (either NaCl or CaCl2) at pH of 6 (adjusted with HCl) at same cell concentration as that of unfavorable conditions. Details about favorable bacteria deposition experiments are provided in the SI. QCM-D Data Analysis. For the experiments performed both in the presence and absence of an energy barrier, the deposition rate can be determined from the slope of the initial (linear) portion of the change in normalized frequency ∆f3 versus time curve (37, 39), since a perfect linear curve (R2 ) 0.9998) of average number of deposited bacteria /view area versus frequency shift (f3) was observed (see detailed curve in SI Figure S1): VOL. 43, NO. 7, 2009 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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kf )
d∆f3
/dt
(1)
The deposition rate at different solution conditions in the presence of an energy barrier is then presented in terms of the adsorption efficiency (R), which is the ratio of the deposition rate (kfp) in the presence relative to the absence of an energy barrier (kfa). R)
kfp kfa
(2)
On the basis of four replicates, the average favorable deposition rate coefficient (kfa) of untreated bacterial strains E.coli, QG6, QL2, and BST (with intact EPS) were 2.28 ( 0.23, 14.20 ( 0.81, 1.72 ( 0.32, and 17.30 ( 0.63 Hz-min-1, respectively, whereas the values for the CER treated bacterial strains E.coli, QG6, QL2, and BST were determined to be 2.34 ( 0.17, 13.06 ( 0.44, 1.43 ( 0.01, and 16.60 ( 0.53 Hz-min-1, respectively.
Results and Discussion The influence of solution ionic composition and ionic strength on the electrophoretic mobilities and zeta potentials of untreated and CER treated bacterial cells are examined and presented in SI Figure S2. Zeta potentials of all bacterial strains (both with and without CER treatment) were negative at pH 6.0 and became less negative with increasing solution ionic strength in both NaCl (square) and CaCl2 (triangle) solutions due to compression of the electrostatic double layer. Under the same ionic strength condition, zeta potentials of all four bacterial strains in CaCl2 solutions (triangle) were less negative relative to those in NaCl solutions (square) (SI Figure S2). This was true over entire examined ionic strengths range. This observation was possibly due to the adsorption of calcium ions to the cell surface resulting in the neutralization of surface charge (40). More importantly, SI Figure S2 showed that for all four bacterial strains, zeta potentials of untreated bacterial cells (without EPS removal) (solid symbol) were comparable with those of CER treated bacterial cells (with EPS removal) (open symbol) in both NaCl (square) and CaCl2 (triangle) solutions. The results clearly show that removal of EPS from cell surfaces via CER treatment does not have large influence the electrokinetic properties of the cell surfaces for all four strains (28). To ensure EPS has been effectively removed from cell surfaces via the employment of CER treatment, analysis of biochemical components of solutions collected from centrifugation after CER treatment was performed. Three major EPS components: proteins, polysaccharides, and humic acids were found to be present in the above-mentioned collected solutions. The amount of these three major EPS components were also determined and presented in SI Table S1. The contents of proteins, polysaccharides, and humic acids in collected solutions varied from 472.7 ( 1.5 to 810.7 ( 3.6, 23.6 ( 0.1 to 154.6 ( 1.3, and 2.8 ( 1.3 to 21.8 ( 0.6 mg g-1 of EPS VDW, respectively. It can be seen from SI Table S1 that proteins was the dominant component of EPS for all four examined bacterial strains, which agreed with many previous studies (19, 21, 41). The biochemical components of solutions collected from the last centrifugation process for untreated cells were also analyzed and found that proteins, polysaccharides, and humic acid were negligible in these solutions. Images of EPS extracted from cell surfaces via the method of CER were also captured by a CCD camera DSRi1-U2 (Nikon, Japan) connected to Ti-E microscopy (Nikon, Japan) and representative image is presented in SI Figure S3. Size measurement via Zetasizer Nano ZS90 (Malvern Instruments, UK) showed that the effective size of EPS in Milli-Q water is around 150∼200 nm. These results indicated that 2310
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FIGURE 1. FTIR spectra of CER-treated E.coli (bottom).
untreated
E.coli
(top)
and
EPS were successfully extracted from cell surfaces by the utilization of CER treatment. FTIR spectra of both untreated and CER treated cells were also acquired for four bacterial strains. A number of absorption peaks representing the presence of various functional groups on cell surface were observed in the FTIR spectra of both untreated and CER treated E.coli (Figure 1). Close inspection of the absorption peaks of CER treated E.coli (Figure 1, bottom) showed shift of bands relative to those observed in untreated cells (Figure 1, top). After treated with CER technique, the bands at 3426.4 cm-1, which reflected stretching vibrations of OsH (the functional group in polysaccharides and proteins) (42) and 1083.2 cm-1, which presented sCOCs wagging (the functional groups in polysaccharides) (42), shifted to 3325.0 cm-1 and 1075.0 cm-1, respectively. Meanwhile, the bands at 1651.7 cm-1, which was the result of CdO and CsN (Amide I) stretching, and 1544.8 cm-1, which was a combination of NsH bending and CsN (Amide II) stretching (Amide I and Amide II corresponded to the functional groups in proteins) (42), shifted to 1608.2 cm-1 and 1512.5 cm-1, respectively. It should be noted that even the smallest shift (sCOCs) in all these absorption bands was 8.2 cm-1, which was 4 times greater than the resolution of FTIR (2 cm-1). The obvious shift of these bands suggested that polysaccharides and proteins (major components of EPS) were involved in the process of CER treatment, which was consistent with above-mentioned biochemical analysis of EPS solutions. Cell size distributions of the untreated and CER treated bacterial strains were monitored on an inverted fluorescence microscope (Ti-E, Nikon, Japan). Measurements of at least 60 cells were taken for each sample and showed that the sizes of CER treated cells were equivalent as those of untreated cells, the cell sizes of bacterial strains E.coli (gram-negative, nonmotile), QG6 (gram-negative, motile), QL2 (gram-positive, nonmotile), and BST (gram-positive, motile) were 1.15 ( 0.19 × 0.60 ( 0.10 µm, 1.45 ( 0.07 × 0.60 ( 0.08 µm, 1.35 ( 0.15 × 0.90 ( 0.02 µm, and 2.50 ( 0.20 × 0.75 ( 0.07 µm, respectively. The results indicate removal of EPS from cell surfaces via CER treatment does not affect the size of bacterial cells. The above results showed that insignificant change of cell size and electrokinetic properties of the cell surfaces was induced for all four bacterial strains by employment of CER treatment to remove EPS, thus negligible differences in
FIGURE 2. Deposition efficiencies (r) of untreated (2) and CER treated (4) bacterial strains E.coli (a), QG6 (b), QL2 (c), and BST (d) on silica surface in NaCl solutions at pH 6.0 (adjusted with HCl) as a function of solution ionic strength. Error bars represent standard deviations of duplicate measurements. The lines are meant to guide the eye. deposition behaviors between untreated and CER treated bacterial cells would be expected according to DerjaguinLandau-Verwey-Overbeek (DLVO) theory, which was commonly used to elucidate the total interaction between two surfaces (22, 43, 44). However, previous studies (27, 28) demonstrated that larger deposition efficiencies were observed for strains with larger amount of EPS relative to strains with less EPS (or without EPS) on cell surfaces and suggested that EPS could enhance cell deposition to surfaces. To further test the significance of EPS on bacterial deposition behavior, deposition kinetics of four untreated bacteria strains (with EPS present) and CER treated strains (with EPS absent) were examined in both NaCl and CaCl2 solutions over a wide range of ionic strength from 1 to 100 mM at pH 6.0 (adjusted with HCl). SI Figure S4 presents representative normalized frequency shifts at the third overtone (∆f3) as bacterial strain E. coli deposition occurred in the presence of 1 and 10 mM NaCl on silica surfaces, and under favorable (fast) deposition conditions onto a PLL-coated silica surfaces in 1 mM NaCl. The number of cells deposited on the surfaces is linearly proportional to the frequency shift (SI Figure S1). Therefore, the increase in the magnitude of frequency shift with elapsed time indicates the number of cells deposit with time elapses. The initial slopes of deposition curves were employed to derive the deposition efficiency (R) (eq 2) since our study focuses on the early stage of deposition kinetics. In most cases, linear normalized frequency shift versus time curves were observed. Deposition efficiencies (R) of untreated and CER treated bacterial cells as a function of NaCl and CaCl2 ionic strength at pH 6.0 were presented in Figures 2 and 3, respectively. In NaCl solutions, R of all untreated bacterial strains (without EPS removal) (solid triangle) increased with increasing solution ionic strength over the whole examined ionic strength range (Figure 2). The same trend was also observed for the CER treated bacterial strains (with EPS removal) (Figure 2, open triangle). This deposition behavior is consistent with the observed trends of zeta potentials versus
ionic strength (SI Figure S2) and generally agreed with classic DLVO theory. An increase in solution ionic strength results in compression of the electrostatic double layer, and thus results in greater deposition of bacteria. In agreement with the obvious increases of deposition efficiencies of bacteria over the entire range of ionic strength in NaCl solutions (Figure 2), R of untreated nonmotile bacterial strains E. coli and QL2 in CaCl2 solutions also increased with increasing solution ionic strength (Figure 3a and c), whereas R of untreated motile bacterial strains QG6 and BST in CaCl2 solutions first increased, and then kept constant, with increasing of ionic strength (Figure 3b and d). Similar trends as untreated cells in CaCl2 solutions were also observed for the CER treated bacterial strains. The different trend of deposition efficiencies over the whole of ionic strength in CaCl2 solutions observed for nonmotile and motile strains was possible due to the involvement of calcium cations in biochemical pathways of cell chemotaxis of motile cells (45-47). Calcium cations could activate the flagellar rotor mechanism and promote motile cells swimming ability at high calcium cations concentrations (45, 47), thus result in the different deposition behaviors from those of nonmotile cells at high ionic strengths. Close comparison of R in CaCl2 solutions with those in NaCl solutions yielded that R in CaCl2 solutions (Figure 3) were, for most cases (for both untreated and treated cells from all four bacterial strains), larger than those in NaCl solutions (Figure 2) across the entire examined ionic strength range. Exceptions included both untreated and treated QG6 at high ionic strength (100 mM), and treated BST at ionic strength of both 50 and 100 mM. These results generally agreed with the observed less negative zeta potentials of both untreated and treated bacterial strains in CaCl2 solutions relative to those in NaCl solutions under all examined ionic strength (SI Figure S2), thus generally agreed with DLVO prediction. Close inspections of Figure 2 and Figure 3 showed that deposition efficiency did not reach the theoretical maximum of 1 even at the highest ionic strength conditions for all four bacterial strains, which can be VOL. 43, NO. 7, 2009 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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FIGURE 3. Deposition efficiencies (r) of untreated (2) and CER treated (4) bacterial strains E.coli (a), QG6 (b), QL2 (c), and BST (d) on silica surface in CaCl2 solutions at pH 6.0 (adjusted with HCl) as a function of solution ionic strength. Error bars represent standard deviations of duplicate measurements. The lines are meant to guide the eye. explained by the presence of an electrostatic interaction energy barrier to cell deposition for all four bacterial strains at the highest ionic strength (100 mM) at pH of 6 both in NaCl and CaCl2 solutions (SI Figure S5). More noteworthy observation is that the observed values of R were greater for the untreated bacterial strains (Figure 2, 2) relative to those with CER treated bacterial strains (Figure 2, 4) across the entire ionic strength range examined in NaCl solutions. This observation held true for all four bacterial strains examined. The same results were also observed for all four bacterial strains at a wide range of ionic strengths (from 1 to 100 mM) in CaCl2 solutions (exception was QL2 at low ionic strength) (Figure 3). These results strongly demonstrated that the removal of EPS from cell surfaces for all four strains decreased the deposition of bacteria onto silica surface. The observed decrease of deposition to surfaces by removal of EPS from cell surfaces agreed with previous studies (27, 28). EPS enhancement of cell deposition is possibly due to the formation of attractive polymeric interaction between silica surfaces and major EPS components like polysaccharides and proteins (27, 28, 48-50), thus results in greater deposition of bacteria. Hydrophobic groups such as nonpolar groups in proteins may allow cell to approach silica surface closely. This may be followed by conformational changes in surface polymers allowing other functional groups to come close enough to the surface for the formation of short-range attractive polymeric interactions (51). It should be noted that previous investigations e.g. Liu et al. (27) only concerned strains of one type (gram-negative and nonmotile) and have not addressed the relevance of the significance of EPS on cell deposition to strains of other cell types (e.g., gram-negative and motile, gram-positive and nonmotile, gram-positive and motile). The four bacterial strains E.coli, QG6, QL2, and BST examined in this study were gram-negative and nonmotile, gram-negative and motile, gram-positive and nonmotile, and gram-positive and 2312
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motile cells, respectively, which have relevance to a wide range of bacteria in natural environment. Our results showed that the deposition efficiencies were greater for untreated cells relative to CER treated cells for all four bacterial strains examined. Hence, it is reasonable to conclude that the enhancement of cell deposition on silica surfaces with the presence of EPS on cell surfaces might be relevant to a wide range of bacterial strains regardless of cell types and motility. The results of this study also have important environmental implications, such as in some in situ bioremediation processes in which bacteria with novel metabolic properties was expected to travel far enough to clean up contaminants; the removal of EPS from cell surfaces can be employed prior to the injection of cells to the contaminated sites.
Acknowledgments This work was supported by the National Basic Research Program of China under grant No. 2006BAB04A14 and the National Natural Science Foundation of China-Young Scientists Fund under grant No.40802054. We acknowledge the editor and three anonymous reviewers for their very positive and helpful comments. We are also grateful to Prof. Bengt Kasemo from department of Physics at Chalmers University of Technology, Gothenburg for his useful comments and to Professor Alistair Borthwick from the University of Oxford for his kind help in English editing.
Supporting Information Available Characteristics of EPS extracted from E.coli, QG6, QL2, and BST via the employment of CER treatment (Table S1); Relationship of the deposited cells number versus frequency shift (Figure S1); Electrophoretic mobilities and zeta potentials of untreated and CER treated bacterial strains E. coli, QG6, QL2, and BST as a function of ionic strength in both NaCl and CaCl2 solutions (Figure S2); Image of EPS extracted from cell surfaces via the employment of CER method (Figure S3); Deposition profiles of untreated and treated E.coli at 1
and 10 mM NaCl, and E.coli and QL2 under favorable conditions (Figure S4); DLVO interaction force profiles for E.coli, QG6, QL2, and BST at 100 mM in both NaCl and CaCl2 solutions (Figure S5). Also presented are details about the growth protocol for all four bacterial strains, determination of cell concentration and viability, biochemical component analysis of EPS solutions, and modification of silica surfaces with PLL and favorable bacteria deposition experiments on PLL-coated silica surfaces. This material is available free of charge via the Internet at http://pubs.acs.org.
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