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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/1.5R ~ -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. EPS and LPS can affect bacterial adhesion in following ways:
(1) the adsorption of free EPS and LPS onto substratum modifies substratum surface properties, (2)
the coverage of EPS and LPS on bacterial surface impact the adhesive properties of bacterial
surfaces, (3) EPS and LPS on bacterial surface interacts directly with substrate surfaces. Our results
provide new insights into the fundamental understanding of the molecular mechanisms of bacterial
biofouling.
ACKNOWLEDGEMENTS
The authors appreciate the financial support of an NSERC (Natural Sciences and Engineering
Research Council of Canada) Discovery Grant Award and an RTI Award (for a Surface Forces
Apparatus, SFA 3).
REFERENCES
Journal: Abu-Lail, N.I. & Camesano, T.A. (2003). Role of lipopolysaccharides in the adhesion,
retention, and transport of Escherichia coli JM109. Environmental Science and Technology, 37
(10), 2173–2183.
Burks, G. A., Velegol, S. B., Paramonova, E. B., Lindenmuth, E., Feick, J. D. & Logan, B. E.
(2003). Macroscopic and nanoscale measurements of the adhesion of bacteria with varying outer
layer surface composition. Langmuir, 19 (6), 2366–2371.
Camesano, T.A., Liu, Y.T. & Datta, M. (2007). Measuring bacterial adhesion at environmental
interfaces with single-cell and single-molecule techniques. Advances in Water Resources, 30 (6-7),
1470-1491.
Cao, S., Wang, J.D., Chen, H.S. & Chen D.R. (2011). Progress of marine biofouling and antifouling
technologies. Chinese Science Bulletin, 56 (7), 598-612.
Fang, H.H.P., Chan, K.Y. & Xu, L.C. (2000). Quantification of bacterial adhesion forces using
atomic force microscopy (AFM). Journal of Microbiological Methods, 40 (1), 89-97.
Flemming, H.-C. (1993). Biofilms and environmental-protection. Water Science and Technology,
27 (7-8), 1-10.
Flemming, H.-C., Griebe, T., Schaule, G. (1996). Antifouling strategies in technical systems - A
short review. Water Science and Technology, 34 (5-6), 517-524.
Flemming, H.-C. & Wingender, J. (2001). Relevance of microbial extracellular polymeric
substances (EPSs) - Part I: Structural and ecological aspects. Water Science and Technology, 43 (6),
1–8.
Frank, B.P. & Belfort, G. (2003). Polysaccharides and sticky membrane surfaces: critical ionic
effects. Journal of Membrane Science, 212 (1-2), 205-212.
Gong, A. S., Bolster, C. H., Benavides, M. & Walker, S. L. (2009) Extraction and analysis of
extracellular polymeric substances: comparison of methods and extracellular polymeric substance
levels in salmonella pullorum SA 1685. Environmental Engineering Science, 26 (10), 1523-1532.
Hori, K. & Matsumoto, S. (2010). Bacterial adhesion: From mechanism to control. Biochemical
Engineering Journal, 48 (3), 424-434.
Israelachvili, J.N. & Adams, G.E. (1976) Direct measurement of long-range forces between 2 mica
surfaces in aqueous KNO3 solutions. Nature, 262 (5571), 773-776.
Israelachvili, J., Min, Y., Akbulut, M., Alig, A., Carver, G., Greene, W., Kristiansen, K., Meyer, E.,
Pesika, N., Rosenberg, K. & Zeng, H. (2010). Recent advances in the surface forces apparatus
(SFA) technique. Reports on Progress in Physics, 73 (3), 036601:1-16.
Ivnitsky, H., Katz, I., Minz, D., Volvovic, G., Shimoni, E., Kesselman, E.,Semiat, R. & Dosoretz,
C. G. (2007) Bacterial community composition and structure of biofilms developing on
nanofiltration membranes applied to wastewater treatment. Water Research, 41 (17) 3924-3935.
Kirby, B.J. & Hasselbrink, E.F. (2004). Zeta potential of microfluidic substrates: 2. Data for
polymers. Electrophoresis, 25 (2), 203–213.
Leckband, D. (1995). The surface force apparatus — a tool for probing molecular protein
interactions. Nature, 376 (6541), 617–618.
Liao, B.Q., Bagley, D.M., Kraemer, H.E., Leppard, G.G., Liss, S.N. (2004) A review of biofouling
and its control in membrane separation bioreactors. Water Environment Research, 76 (5), 425-436.
Li, B.K. & Logan, B.E. (2004). Bacterial adhesion to glass and metal-oxide surfaces. Colloids
Surface B Biointerfaces 36 (2), 81-90.
Lu, Q., Hwang, D. S., Liu, Y., & Zeng, H. (2012) Molecular interactions of mussel protective
coating protein, mcfp-1, from Mytilus Californianus. Biomaterials, 33 (6) 1903-1911.
Lu, Q., Wang, J., Faghihnejad, A., Zeng, H. & Liu, Y. (2011). Understanding the molecular
interactions of lipopolysaccharides during E. coli initial adhesion with a surface forces apparatus.
Soft Matter, 7 (19), 9366-9379.
Moreira, L.M., Videira, P.A., Sousa, S.A., Leitao, J.H., Cunha, M.V. & Sa-Correia, I. (2003).
Identification and physical organization of the gene cluster involved in the biosynthesis of
Burkholderia cepacia complex exopolysaccharide. Biochemical and Biophysical Research
Communications, 312 (2), 323-333.
Tsuneda, S., Aikawa, H., Hayashi, H., Yuasa, A. & Hirata, A. (2003). Extracellular polymeric
substances responsible for bacterial adhesion onto solid surface. Fems Microbiology Letters, 223
(2), 287-292.
Vinion-Dubiel, A.D., & Goldberg, J.B. (2003). Lipopolysaccharide of Burkholderia cepacia
complex. Journal of Endotoxin Research, 9 (4), 201-213.
Yang, X., Teng, F., Zeng, H., & Liu, Y. (2012). Impact of cranberry juice on initial adhesion of the
EPS producing bacterium Burkholderia cepacia. Biofouling, in press.