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Role of Nonadsorbing Polymers in Bacterial Aggregation K. E. Eboigbodin, J. R. A. Newton, A. F. Routh, and C. A. Biggs* Department of Chemical and Process Engineering, The University of Sheffield, Mappin Street, Sheffield S1 3JD, U.K. Received June 28, 2005. In Final Form: October 10, 2005 Bacteria exhibit properties similar to those of nonbiological colloids and can display pairwise attractions when in close proximity. This interaction is governed by the surface chemistry of the cells. We seek to understand bacterial aggregation at the cellular level using Escherichia coli (E. coli) AB1157. Aggregation studies were carried out using 0.5 to 2.5 wt % E. coli AB1157 harvested in different growth phases with varying concentrations of a nonadsorbing polymer, sodium polystyrene sulfonate (SPS). The electrophoretic mobility of E. coli AB1157 in different growth phases was determined using phase-amplitude light scattering. E. coli AB1157 was found to be negatively charged, and the cell surface properties changed in different growth phases. The electrokinetic results correlated well with the different concentrations of nonadsorbing polymer needed to induce depletion aggregation. This shows that a difference in aggregation properties is due to changes in the bacteria electrokinetic properties during their growth.
1. Introduction Bacterial aggregation is the cell-to-cell adhesion of bacterial species or strains in order to perform a specialized function under certain physiological conditions.1 When aggregated, bacteria are able to communicate between themselves and uniformly perform metabolic functions, which are greater than their planktonic counterparts. The effect of bacterial aggregation can be beneficial or detrimental to its surroundings. For example, microorgansims can colonize on medical devices, which may result in a dangerous source of infection.2 Conversely, some bioreactors are developed to enhance bacterial aggregation that is very effective for treating wastewater and contaminated soil.3 For example, a common and popular method of biologically degrading domestic and industrial wastewater is the activated sludge process, and the sucessful operation of this process is largely dependent on the ability of a mixed consortium of microorganisms to aggregate.4 Like inert colloids, bacteria can display pairwise attractions when in close proximity. This interaction is governed by the surface chemistry of the bacterial surface (some van der Waals forces but mainly electrostatic forces due to the ionic groups on the cell surface), and not surprisingly their surface properties have been shown to play a significant role in bacterial aggregation.5-14 The electrostatic repulsive potential is crucial in determining * Corresponding author. E-mail:
[email protected]. Tel: (+44) 01142227510. Fax: (+44) 01142227501. (1) Marshall, K. C. Microbial Adhesion and Aggregation; SpringerVerlag: New York, 1984. (2) Jucker, B. A.; Harms, H.; Zehnder, A. J. J. Bacteriol. 1996, 178, 5472-5479. (3) Okunuki, S.; Kawaharasaki, M.; Tanaka, H.; Kanagawa, T. Water Res. 2004, 38, 2433-2439. (4) Biggs, C. A.; Lant, P. A. Water Res. 2000, 34, 2542-2550. (5) Hermansson, M. Colloids Surf., B 1999, 14, 105-119. (6) Rutter, P. R.; Vincent, B.; Marshall, K. C., Ed.; Springer-Verlag: New York, 1984; p 21. (7) van Loosdrecht, M. C.; Lyklema, J.; Norde, W.; Schraa, G.; Zehnder, A. J. Appl. Environ. Microbiol. 1987, 53, 1893-1897. (8) van Loosdrecht, M. C.; Lyklema, J.; Norde, W.; Schraa, G.; Zehnder, A. J. Appl. Environ. Microbiol. 1987, 53, 1898-1901. (9) Smyth, C. J.; Jonsson, P.; Olsson, E.; et al. Infect. Immun. 1978, 22, 462-472. (10) Zita, A.; Hermansson, M. Appl. Environ. Microbiol. 1997, 63, 1168-1170. (11) Kos, B.; Suskovic, J.; Vukovic, S.; Simpraga, M.; Frece, J.; Matosic, S. J. Appl. Microbiol. 2003, 94, 981-987.
stability and is determined by the cell because it is largely influenced by the cell surface potential.15 In addition, other forces, such as those exerted by extracellular polymeric substances (EPS) produced by the microorganisms, may affect dispersion stability. Quantification of the surface electrokinetic properties of bacteria appears to be more difficult in comparison with that of nonbiological colloids as a result of the chemical and structural complexity of bacterial cell surfaces. A wellknown method for measuring surface electric potentials is the measurement of electrophoretic mobility to obtain the zeta potential.8,16-19 The zeta potential is usually calculated from the electrophoretic mobility,2 but this calculation is not suitable for bacteria because of their surface complexity and nonspherical shape. Ohshima’s soft-particle electrophoresis theory, developed by Ohshima and Kondo,20-23 is suitable for colloids with a soft outer layer such as bacteria, although a spherical shape is still assumed. The model assumes the presence of an ionpenetrable layer of finite thickness around a core spherical particle, and this approach has been found to be useful in estimating the surface charge of several biological systems.16-18 Tsuneda et al.18 used Ohshima’s theory in investigating the effect of extracellular polymeric substances produced by bacterial cells on the ability to adhere to solid surfaces. Hayashi et al.16 were able to demonstrate that the growth phase of bacteria alters their cell surface (12) Liu, Y.; Yang, S.-F.; Liu, Q.-S.; Tay, J.-H. Curr. Microbiol. 2003, 46, 270-274. (13) Bos, R.; van der Mei, H. C.; Busscher, H. J. FEMS Microbiol. Rev. 1999, 23, 179-229. (14) Liu, Y.; Yang, S.-F.; Tay, J.-H.; Liu, Q.-S.; Qin, L.; Li, Y. Enzyme Microb. Technol. 2004, 34, 371-379. (15) Hayashi, H.; Tsuneda, S.; Hirata, A.; Sasaki, H. Colloids Surf., B 2001, 22, 149-157. (16) Hayashi, H.; Seiki, H.; Tsuneda, S.; Hirata, A.; Sasaki, H. J. Colloid Interface Sci. 2003, 264, 565-568. (17) Sonohara, R.; Muramatsu, N.; Ohshima, H.; Kondo, T. Biophys. Chem. 1995, 55, 273-277. (18) Tsuneda, S.; Aikawa, H.; Hayashi, H.; Hirata, A. J. Colloid Interface Sci. 2004, 279, 410-417. (19) van der Mei, H. C.; Busscher, H. J. Appl. Environ. Microbiol. 2001, 67, 491-494. (20) Ohshima, H.; Kondo, T. J. Colloid Interface Sci. 1989, 130, 281282. (21) Ohshima, H.; Kondo, T. J. Colloid Interface Sci. 1987, 116, 305311. (22) Ohshima, H. Colloids Surf., A 1995, 103, 249-255. (23) Ohshima, H. Adv. Colloid Interface Sci. 1995, 62, 189-235.
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properties, using Ohshima’s theory to deduce the cell surface potential of bacteria. Sonohara et al.17 also used Ohshima’s theory to detect differences in surface properties between gram-negative and gram-positive bacteria. Their findings revealed that gram-negative bacteria are more negatively charged and have a less soft surface than gram-positive bacteria. Other existing methods of investigating colloidal surface changes include pH titration.24 Extracellular polymeric substances (EPS) produced by bacteria have also been shown to be involved in biological aggregation.25-32 Extracellular polymeric substances (EPS) are biopolymers produced by microorgansims during their growth. The highest production of EPS occurs at the onset of the stationary phase.33,34 EPS contains polysaccharides, proteins, nucleic acid, lipids, and other biological polymers such as humic substances.35 Although EPS have been shown to keep microbial aggregates together by providing a cohesive force,25,35-37 their role in aggregation still remains unclear. Detailed knowledge of bacteria surface electrokinetic properties and the precise role of EPS are crucial for understanding bacterial flocculation, aggregation, and adhesion. From a physical point of view, bacteria can be considered to be a dispersion of colloids surrounded by nonadsorbing polyelectrolytes, EPS. Bacteria are negatively charged,8,17 and a repulsive bacteria-EPS interaction results in an attractive interaction between bacterial cells commonly referred to as the depletion attraction.38,39 It is generally accepted that a phase separation occurs in a suspension of colloids and nonadsorbing polymers as a result of this depletion interaction.40-42 The role of nonadsorbing polymers in inducing aggregation can be dated as far back as Traube,43 where he was able to demonstrate that the addition of a water-soluble polymer to natural rubber resulted in a phase separation. However, it was not until 1954 that a satisfactory explanation of this depletion interaction was given by Asakura and Oosawa.38,39 They postulated that phase separation in a mixture of colloids and nonadsorbing polymers is due to an imbalance in osmotic pressure when the nonadsorbing polymers are excluded from the region between particles. As the (24) Mikkelsen, L. H. Water Res. 2003, 37, 2458-2466. (25) Kreft, J. U.; Wimpenny, J. W. Water Sci. Technol. 2001, 43, 135-141. (26) Flemming, H. C.; Wingender, J. Water Sci. Technol. 2001, 43, 9-16. (27) Tsuneda, S.; Aikawa, H.; Hayashi, H.; Yuasa, A.; Hirata, A. FEMS Microbiol. Lett. 2003, 223, 287-292. (28) Liu, Y.; Fang, H. H. P. Crit. Rev. Environ. Sci. Technol. 2003, 33, 237-273. (29) Tsuneda, S.; Jung, J.; Hayashi, H.; Aikawa, H.; Hirata, A.; Sasaki, H. Colloids Surf., B 2003, 29, 181-188. (30) Hammer, B. K.; Bassler, B. L. Mol. Microbiol. 2003, 50, 101104. (31) Miller, M. B.; Bassler, B. L. Annu. Rev. Microbiol. 2001, 55, 165-199. (32) Whitehead, N. A.; Barnard, A. M.; Slater, H.; Simpson, N. J.; Salmond, G. P. FEMS Microbiol. Rev. 2001, 25, 365-404. (33) Petry, S.; Furlan, S.; Crepeau, M.-J.; Cerning, J.; Desmazeaud, M. Appl. Environ. Microbiol. 2000, 66, 3427-3431. (34) Wolfstein, K.; Stal., L. Mar. Ecol.: Prog.s Ser. 2002, 236, 13-22. (35) Flemming, H. C.; Wingender, J. Water Sci. Technol. 2001, 43, 1-8. (36) Wingender, J.; Flemming, H.-C. In Biotechnology; Winter, J., Ed., 1999; Vol. 8, pp 63-86. (37) Sutherland, I. W. Water Sci. Technol. 2001, 43, 77-86. (38) Asakura, S.; Oosawa, F. J. Chem. Phys. 1954, 22, 1255-1256. (39) Asakura, S.; Oosawa, F. I. J. Polym. Sci 1958, 183-192. (40) Jenkins, P.; Snowden, M. Adv. Colloid Interface Sci. 1996, 68, 57-96. (41) Tuinier, R.; Rieger, J.; de Kruif, C. G. Adv. Colloid Interface Sci. 2003, 103, 1-31. (42) Cerda, J. J.; Sintes, T.; Sorensen, C. M.; Chakrabarti, A. Phys. Rev. E: Stat. Nonlinear, Soft Matter Phys. 2004, 70, 011405. (43) Traube, J. Gummi-Ztg. 1925, 39, 434-435.
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particles are drawn closer, the distance between the particles becomes less than the effective diameter of the nonadsorbing polymers. The nonadsorbing polymers are unable to penetrate this region, and pushing the particles together leads to an increase in the configurational entropy of the system. This leads to an effective attraction between each particle known as the depletion interaction. Vrij44 developed a model for estimating the depletion attraction potential between two hard spherical particles induced by nonadsorbing polymers. Vincent and co-workers45 later developed a method to analyze the depletion interaction of soft spheres (colloids with a layer of adsorbed polymer chains). They analyzed this interaction by taking into account the penetration and compression effects between the nonadsorbing polymer and the steric layer on the particles. Recent developments in colloid science, such as atomic force microscopy (AFM), have allowed the direct measurement of depletion and structural forces.46-50 Milling and Biggs47 analyzed the depletion forces between silica surfaces and free neutral polymer poly(dimethylsiloxane).47 Yan et al.50 studied the structure and strength of aggregates from latex particles induced by the addition of poly(acrylic) acid. Their findings revealed that the concentration of poly(acrylic) acid affects the structural denseness and strength of latex aggregates. Several nonadsorbing EPS have been shown to induce depletion attractions in oil-in-water emulsions51 and in dispersions of casein micelles.52,53 Tuinier et al.41 studied the depletion attraction in a mixture of casein micelles from skim milk and extracellular polysaccharides produced by lactic acid bacteria, Lactococcus lactis subsp. cremoris strain NIZO B40. They analyzed the depletion attraction between the casein micelles in the presence of polymers using Vrij’s depletion model.44 Their findings reveal that upon addition of EPS to casein micelles depletion attraction occurred. Similarly, phase separation was also found in casein and amylopectin mixtures, which was analyzed on the basis of Vrij’s depletion theory.54 The laws, kinetics, and mechanism of depletion aggregation of inert colloids have been described in the literature.49,55,56 However, the depletion of nonadsorbing polymers as a mechanism for aggregation in bacterial suspensions has not been well exploited. Only a handful of papers are available on the subject. In this study, we seek to understand the role of nonadsorbing polymers in the aggregation of Escherichia coli AB1157. We show that a nonadsorbing polymer can induce a depletion attraction (44) Vrij, A. Pure Appl. Chem. 1976, 48, 471-483. (45) Vincent, B.; Edwards, J.; Emmett, S.; Jones, A. Colloids Surf. 1986, 18, 261-281. (46) Richetti, P.; Kekicheff, P. Phys. Rev. Lett. 1992, 68, 1951-1954. (47) Milling, A.; Biggs, S. J. Colloid Interface Sci. 1995, 170, 604606. (48) Mondain-Monval, O.; Leal-Calderon, F.; Phillip, J.; Bibette, J. Phys. Rev. Lett. 1995, 75, 3364-3367. (49) Burns, J. L.; Yan, Y.-d.; Jameson, G. J.; Biggs, S. J. Colloid Interface Sci. 2002, 247, 24-32. (50) Yan, Y.-d.; Burns, J. L.; Jameson, G. J.; Biggs, S. Chem. Eng. J. 2000, 80, 23-30. (51) Tuinier, R.; de Kruif, C. G. J. Colloid Interface Sci. 1999, 218, 201-210. (52) Tuinier, R.; ten Grotenhuis, E.; Holt, C.; Timmins, P. A.; de Kruif, C. G. Phys. Rev. E: Stat. Phys., Plasmas, Fluids, Relat. Interdiscip. Top. 1999, 60, 848-856. (53) Tuinier, R.; de Kruif, C. G. J. Chem. Phys. 1999, 110, 92969304. (54) de Bont, P. W.; van Kempen, G. M. P.; Vreeker, R. Food Hydrocolloids 2002, 16, 127-138. (55) Anderson, V. J.; de Hoog, E. H. A.; Lekkerkerker, H. N. W. Phys. Rev. E 2002, 65, 011401-011408. (56) Jenkins, P.; Vincent, B. Langmuir 1996, 12, 3107-3113.
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between bacterial cells and that the tendency of the cells to aggregate is influenced by its growth phase. 2. Materials and Methods 2.1. Bacterial Strains and Culture Conditions. Nonpathogenic, freeze-dried E. coli AB1157 (DSM number 9036) was purchased from Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH (DSMZ) and resuspended in Luria-Bertani (LB) media (tryptone 10.0 g/L, yeast extract 5.0 g/L, NaCl 10.0 g/L, adjusted to pH 7.0). E. coli was grown at 30 °C overnight with aeration in LB broth supplemented with 0.5 wt % glucose. The culture was then used to inoculate fresh LB broth containing 0.5 wt % glucose at 1:100 dilution and grown at 30 °C with aeration. The optical density at wavelength 600 nm (OD600) was measured using a spectrophotometer (ThermoSpectronic, U.K.). The cell number was determined using a counting chamber. A correlation between optical density and cell number was determined and used to generate the growth curve of E. coli under batch conditions. Cells were sampled in triplicate at intervals between 0 and 24 h and harvested by centrifugation at 5000g for 10 min. The cell pellets were used in the following experiments. 2.2. Electrophoretic Mobility Measurement. Electrophoretic mobility was measured using a phase-amplitude light scattering (PALS) zeta potential analyzer (Brookhaven Zeta PALS, U.K.) following the technique developed by Hayashi et al.16 After harvesting, the cell pellets were washed by resuspending in distilled water followed by centrifugation at 5000g for 10 min. This washing step was repeated four times to eliminate residual substrates and extracellular polymers, which were produced by E. coli AB1157 during growth. The washed cell suspensions were then dispersed in an ultrasonic bath prior to measurement. Measurements were conducted using an electric field of 4 V cm-1 at a frequency of 2.0 Hz. 2.3. Aggregation Measurement. For the aggregation measurements, the cells were harvested in different growth phases: 3.5 h (early exponential phase), 6 h (mid exponential phase), 8 h (late-exponential phase) 14 h, 18 h, and 24 h (stationary phase). The pellet was washed three times and resuspended in distilled water. Different concentrations of nonadsorbing polymer sodium polystyrene sulfonate (Acros Organics; average molecular weight 70 000) were added to cuvettes containing 0.5 to 2.5 wt % E. coli AB1157. The cuvettes were visually inspected after 24 h to determine if the solutions were stable (i.e., no aggregation) or unstable (i.e., aggregation and settling of solution). An aggregation assay was also carried out according to Shen et al.57 The percentage aggregation was the difference in OD600 readings taken at time 0 and 24 h according to eq 1
Figure 1. Increase in cell number over time for E. coli AB1157. (Typical phases for batch growth are indicated.)
Gram-negative, nonpathogenic E. coli AB1157 was chosen as the model system to investigate bacterial aggregation because of its fast growth rate and the fact that the genome has already been sequenced.58 The information from the bacteria genome can be analyzed to provide structural and functional information about unknown genes and proteins that might be implicated in bacteria aggregation. Figure 1 shows the increase in cell number over time for E. coli AB1157 grown under batch conditions. This follows a typical growth curve under batch
conditions with three clearly distinct phases (i.e., lag phase 0-2 h, exponential 2-8 h, and onset of stationary phase after 10 h). The generation time and the growth rate for the organism in a batch culture are calculated during the exponential growth phase and were found to be about 51.67 min and 0.0134 min-1 respectively. The previously documented values for generation time ranges from as fast as 17 min to several hours.59,60 Figure 2 shows the effect of the addition of nonadsorbing polymer sodium polystyrene sulfonate (SPS) to a constant concentration of E. coli AB1157 (2.0 wt %) that were harvested during the stationary phase. With the addition of no SPS (Figure 2i), the 2.0 wt % E. coli solution is turbid and represents a stable (unaggregated) solution. However, the addition of 2.0 wt % SPS to the E. coli solution (Figure 2viii) results in a clear solution with the E. coli settled on the bottom of the cuvette. This represents an unstable (aggregated) solution. Therefore, from Figure 2 it can be seen that as the concentration of SPS increases (from 0 to 2.0 wt %) the upper part of the cuvette becomes clearer and the lower phase becomes very turbid. That is, the solution becomes unstable, and the E. coli settles to the bottom of the cuvette. A sharp interface is evident, which is characteristic of phase separation. The upper phase is rich in SPS whereas the lower is E. coli-rich. The kinetics of the phase separation is dependent on the concentration of SPS used. For example, for the 2.0 wt % E. coli solution shown in Figure 2, phase separation is observed after 4-6 h for solutions with 2.0 wt % SPS. However, when phase separation occurs for lower concentrations of SPS, this occurs after approximately 16 h. However, irrespective of the amount of SPS used, each phase (either stable or unstable) remained constant after 24 h. We also investigated whether depletion stabilization reoccurred in an E. coli suspension containing a higher concentration of SPS, as previously reported for an inert colloidal system.61 However, this was not observed for our E. coli suspension containing up to 10.0 wt % SPS. Another way of representing the results in Figure 2 is shown in Figure 3. In this case, the stability of suspensions of varying concentrations of E. coli is determined for different concentrations of SPS. In each case, the cells are harvested during the stationary growth phase. The circles represent a stable solution (i.e., no aggregation) whereas the diamonds indicate an unstable solution (i.e., aggrega-
(57) Shen, S.; Samaranayake, L. P.; Yip, H.-K. Arch. Oral Biol. 2005, 50, 23-32. (58) Blattner, F. R.; Plunkett, G., III; Bloch, C. A.; Perna, N. T.; Burland, V.; Riley, M.; Collado-Vides, J.; Glasner, J. D.; Rode, C. K.; Mayhew, G. F.; Gregor, J.; Davis, N. W.; Kirkpatrick, H. A.; Goeden, M. A.; Rose, D. J.; Mau, B.; Shao, Y. Science 1997, 277, 1453-1474.
(59) Olsson, J.; Dasgupta, S.; Berg, O. G.; Nordstrom, K. Mol. Microbiol. 2002, 44, 1429-1440. (60) Olsson, J. A.; Nordstrom, K.; Hjort, K.; Dasgupta, S. J. Mol. Biol. 2003, 334, 919-931. (61) Feigin, R. I.; Napper, D. H. J. Colloid Interface Sci. 1980, 75, 525-541.
% aggregation )
OD0 - OD24 h × 100 OD0
(1)
where OD0 is the optical density at 600 nm of E. coli AB1157 immediately after adding polymer and OD24 is the optical density after 24 h. The percentage aggregation was determined for E. coli AB1157 harvested in different growth phases.
3. Results and Discussion
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Figure 2. Visual inspection of constant E. coli AB1157 concentration (2.0 wt %) solution harvested in the late stationary phase (24 h) with 0.0-2.0 wt % sodium polystyrene sulfonate (SPS) after 24 h of SPS addition. The picture clearly shows the difference between stable (unaggregated) and unstable (aggregated) solutions. The blank SPS solution in cuvette i shows a suspension of E. coli in water. The remaining solutions (cuvettes ii-viii) becoming increasingly clear with an increase in SPS because of aggregation and settling of cells at the bottom of the cuvettes.
Figure 3. Stability of E. coli AB1157 harvested in the stationary phase in the presence of nonabsorbing polymer sodium polystyrene sulfonate (SPS). Stable means nonaggregating, and unstable means aggregating.
tion). The bold line indicates the transition from a stable to an unstable solution. From Figure 3, it is evident that the degree of depletion aggregation is dependent on the concentration of the nonadsorbing polymers used. The same experiment was repeated for cells harvested in different growth phases to investigate the possible biological interactions that may influence the observed results. The transition line between a stable and an unstable solution in different growth phases (similar to Figure 3) can be seen in Figure 4. These results reveal that different amounts of polymer are needed for a constant weight percent of E. coli to induce the aggregation of cells harvested in different growth phases. The least amount of polymer is required for cells harvested during the stationary phase, whereas as the cells move from the early to late exponential growth phase an increase in polymer is needed to create an unstable (i.e., aggregation) solution. Therefore, phase separation due to depletion interaction is dependent on the growth phase of the cells.
Figure 4. Transition from stable to unstable solutions for E. coli AB1157 harvested in different growth phases in the presence of nonabsorbing polymer sodium polystyrene sulfonate (SPS).
Similarly, as shown in Figure 5, the percentage of cells that have aggregated is also dependent on the growth phase of the cells. Again, the highest percentage of aggregation occurs for cells harvested during the stationary phase for a constant SPS concentration. These results were also conducted for different concentrations of E. coli (0.5 to 2.5 wt %) with the same trend observed (results not shown) (i.e., greater aggregation during the stationary growth phase than during the exponential growth phase). To investigate this further, Figure 6 shows the percentage aggregation in different growth phases for a constant concentration of E. coli AB1157 (2.0 wt %) and a constant concentration of SPS (0.2 wt %). The most marked increase in percentage aggregation occurs during the stationary growth phase (at 18 h), which supports the results in Figure 5. Similar trends were also found for different concentrations of E. coli and SPS (results not shown).
Nonadsorbing Polymers in Bacterial Aggregation
Figure 5. Percentage aggregation of 2.0 wt % E. coli harvested in different growth phases with varying concentrations of nonadsorbing polymer sodium polystyrene sulfonate (SPS).
Figure 6. Percentage aggregation of E. coli AB1157 harvested in different growth phases with the addition of 0.2 wt % nonadsorbing polymer sodium polystyrene sulfonate (SPS).
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U.K.) in different growth phases. E. coli AB1157 displays a negative electrophoretic motility value throughout the growth period, suggesting that the cell surface is negatively charged. A similar negative surface charge was also found by Hayashi et al.16 for gram-negative bacteria. This may be due to the major components of the outer membraneslipopolysaccharides (i.e., the carboxyl group carries a net negative charge), sialic acids, and/or proteins.62 The electrophoretic mobility of the cells steadily increases in magnitude from the early to mid exponential phase and then decreases in magnitude to reach a maximum at -2.5 (µm s-1)/(V/cm) at 18 h (stationary phase) and remains relatively constant until the final measurement at 24 h (again in the stationary phase). This shows that the surface charge of the cells is not constant during the growth phase and that the magnitude of the charge is also dependent on the stage in the growth phase, hence the electrokinetic properties are influenced by the growth phase. Also, the electrostatic contribution of the bacterial surface predicts the amount of polymer needed to induce depletion attraction, and this is dependent on the growth phase. As the electrophoretic mobility of E. coli AB1157 increases in magnitude, the electrostatic repulsion between the cells will increase, thereby increasing the amount of SPS needed for aggregation to occur. The above findings are consistent with recent findings by Hayashi et al.16 and suggest that differences in aggregation properties are due to changes in the electrokinetic properties of bacteria surfaces during their growth. The correlation between high aggregation during the stationary phase (at 18 h in Figure 6) and the electrophoretic mobility (Figure 7) is also in full agreement with previous findings that state that cellular aggregation or adhesiveness increases as surface charge decreases.18,29 The strength of induced interaction between cells increases as the concentration of SPS increases. This observation can be related back to the production of EPS by bacteria. As cells transit from the exponential to the stationary phase, the bacteria tend to produce more EPS,34 which are released to its surroundings. When the production of EPS gets to a certain threshold, the bacteria and the EPS will come into close proximity, which will trigger a depletion attraction between bacterial cells. Conclusions
Figure 7. Electrophoretic mobility of E. coli AB1157 in different growth phases.
On the basis of the results thus far, it is evident that the growth phase of E. coli has a significant effect on the percentage aggregation and the amount of polymer necessary to induce this aggregation. Therefore, a characteristic of E. coli is changing during the growth phase in order to elicit this response. Figure 7 shows the electrophoretic mobility of E. coli AB1157 measured using the PALS zeta potential analyzer (Brookhaven Zeta PALS,
Phase separation occurs in a mixture of bacteria and nonadsorbing polymers because of a depletion interaction. Different concentrations of nonadsorbing polymer are needed to induce depletion aggregation from cells harvested in different growth phases. Cells tend to aggregate more easily during the stationary phase than in the exponential phase. The magnitude of the electrophoretic mobility of E. coli AB1157 decreased from the early to mid exponential phase to the stationary phase, and this correlates very closely with differences in aggregation properties during growth. These findings suggest that depletion interactions induced by extracellular polymeric substances are involved in bacterial aggregation. Acknowledgment. This research is funded by BBSRC through grant number BB/C505391/14 and an EPSRC studentship. LA051740U (62) Torimura, M.; Ito, S.; Kano, K.; Ikeda, T.; Esaka, Y.; Ueda, T. J. Chromatogr., B 1999, 721, 31-37.
