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Applied and Environmental Microbiology, April 2009, p. 2346-2353, Vol. 75, No. 8
0099-2240/09/$08.00+0     doi:10.1128/AEM.02671-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

Variation in Bacterial ATP Level and Proton Motive Force Due to Adhesion to a Solid Surface

Yongsuk Hong and Derick G. Brown^*

Department of Civil and Environmental Engineering, Lehigh University, Bethlehem, Pennsylvania

Received 21 November 2008/ Accepted 2 February 2009

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Bacterial adhesion to natural and man-made surfaces can be beneficial or detrimental, depending on the system at hand. Of vital importance is how the process of adhesion affects the bacterial metabolic activity. If activity is enhanced, this may help the cells colonize the surface, whereas if activity is reduced, it may inhibit colonization. Here, we report a study demonstrating that adhesion of both Escherichia coli and Bacillus brevis onto a glass surface resulted in enhanced metabolic activity, assessed through ATP measurements. Specifically, ATP levels were found to increase two to five times upon adhesion compared to ATP levels in corresponding planktonic cells. To explain this effect on ATP levels, we propose the hypothesis that bacteria can take advantage of a link between cellular bioenergetics (proton motive force and ATP formation) and the physiochemical charge regulation effect, which occurs as a surface containing ionizable functional groups (e.g., the bacterial cell surface) approaches another surface. As the bacterium approaches the surface, the charge regulation effect causes the charge and pH at the cell surface to vary as a function of separation distance. With negatively charged surfaces, this results in a decrease in pH at the cell surface, which enhances the proton motive force and ATP concentration. Calculations demonstrated that a change in pH across the cell membrane of only 0.2 to 0.5 units is sufficient to achieve the observed ATP increases. Similarly, the hypothesis indicates that positively charged surfaces will decrease metabolic activity, and results from studies of positively charged surfaces support this finding.

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Bacterial adhesion and biofilm formation are important to a wide array of fields, such as environmental, chemical, and biomedical engineering; food processing; materials science; marine science; and ecology and environmental sciences. A key consideration in the development of a biofilm is the initial interaction between the bacterium and the substrata and the way in which these interactions affect the metabolic activity of the cells. If the metabolic activity increases, this would help the cells colonize the surface, whereas if the metabolic activity decreases, then the cells may become inactive or die.

There have been a number of studies that have examined the effects of attachment on bacterial metabolic activity. The first reported observation of this effect was made in 1943, where it was found that bacterial activity increased in the presence of glass surfaces, particularly with low nutrient concentrations (54). Since this first study, researchers have examined how various materials, including glass and polymer surfaces (10, 11, 24-26, 40, 44, 47), silicone surfaces (52), ceramic surfaces (32), dialysis membranes (18), activated carbon (7), clays (43), sands (31, 48), estuary particles (19), and ion exchange resins (46), affect the metabolic activity of attached bacteria.

Different mechanisms have been examined in an attempt to explain how surfaces affect bacterial metabolic activity. For example, one study found that activity increased with bacterial hydrophobicity, presumably due to firmer adhesion on the surface (24). Other researchers found that bacterial activity both increased (11) and decreased (40) with decreasing hydrophobicity of the solid surface. A study of bacterial attachment to clays showed that most 2:1 clays enhanced respiration, while 1:1 clays had minimal effect (43). It has also been observed that positively charged surfaces can reduce cell viability (13, 26, 44, 47). Unfortunately, given these disparate results, the prediction of how a specific surface may affect bacterial metabolic activity remains elusive.

Here, we propose a hypothesis on how surfaces may affect bacterial metabolic activity. This hypothesis is based on a link between the physiochemical charge regulation effect, which occurs as a surface with acid/base functional groups approaches another surface, and cellular bioenergetics, where ATP is produced from pH and electrostatic charge gradients (pH and , respectively) set up across the bacterial cytoplasmic membrane, the sum of which is termed proton motive force (p). The charge regulation effect results in a variation in the cell surface pH as the cell approaches another surface, and the hypothesis we propose is that this variation in cell surface pH will affect pH, ultimately affecting p, with a concomitant variation in the cellular ATP level.

In order to explore this hypothesis, the objectives of the current study were to (i) demonstrate the direct link between bacterial adhesion to a surface and an increase in cellular ATP concentration and (ii) calculate the increase in p required to achieve the observed ATP increase. The gram-negative strain Escherichia coli K-12 and the gram-positive organism Bacillus brevis were the focus of this study. Two different solution chemistries were used, one containing phosphate buffer solution (PBS) (a phosphate-based buffer) and another containing piperazine-N,N'-bis(2-ethanesulfonic acid) (PIPES) (a nitrogen- and sulfur-based buffer). Through the use of vials containing a fixed mass of one of three different-sized glass beads and also vials without glass beads as controls, various glass surface areas were available for cellular adhesion. These experiments were based on the concept that the total ATP concentration in vials with different surface areas for adhesion would vary as a function of surface area if adhesion affects ATP, whereas the total ATP concentration would be invariant with surface area if adhesion did not affect ATP. The number of adhered cells and total ATP per vial were quantified, allowing calculation of the ATP concentrations for both planktonic and adhered bacteria. To verify that bacterial adhesion, rather than just the presence of the glass beads, affected metabolic activity, an identical series of experiments was conducted with the nonionic surfactant Tween 80 present in solution. Tween 80 reduced bacterial adhesion to the glass beads and allowed the verification that the presence of the glass beads themselves did not have any effect on cellular ATP concentrations. Used in combination, these experiments allowed quantification of the effects of cell adhesion on the bacterial ATP concentration and p.

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Bacteria and cultivation.
The gram-negative strain E. coli K-12 (ATCC 29181) and the gram-positive organism B. brevis (ATCC 9999) were used for this study. Bacterial cultures were grown in a minimal medium (16), harvested in the late exponential phase, and then stored in 15% glycerol at –86°C using the glass-bead method (20) to maintain a uniform bacterial stock. For each experiment, bacteria from the frozen stock were grown in 500 ml of minimal medium in which glucose served as the sole carbon source. Cultures were incubated at 30°C on a rotary shaker at 500 rpm to late exponential phase.

Glass-bead preparation.
Solid surfaces used were spherical, soda lime glass beads with three different mean diameters of 0.5, 1.0, and 2.5 mm (Biospec, Inc., Bartlesville, OK). These glass beads were negatively charged in solution (15) and had surface areas of 48, 24, and 9.6 cm²/g, respectively. The beads were cleaned by being soaked in 1 M HCl solution for 12 h, followed by a rinse under flowing tap water for 1 h and a final rinse in deionized water (DI) for 10 min. They were then dried at 60°C and stored in a polypropylene container prior to use.

Buffer solutions.
PBS consisted of 0.258 g KH₂PO₄ and 0.470 g K₂HPO₄ in 1 liter of DI, providing a pH of 7.2. While ATP extracted in nucleotide releasing agent (NRB) is stable (42), in order to ensure that the phosphate present in PBS did not impact the ATP measurements, a second buffer solution, PIPES, was also used independently of PBS. PIPES consisted of 1.2307 g PIPES in 1 liter DI, adjusted to pH 7.2 with 1 N NaOH.

Experimental procedures.
At the late exponential phase, bacteria were harvested by centrifugation at 3,500 x g for 15 min at room temperature, washed twice in either PBS or PIPES, and then incubated at 25°C on a rotary shaker for 12 h in either buffer solution. After 12 h, the bacteria were harvested and washed once more. One half of the washed bacteria was resuspended in buffer, and the other half was resuspended in 0.12 mM Tween 80 in buffer. The final concentration of bacteria in both suspensions was approximately 2.0 x 10⁸/ml for E. coli or 4.7 x 10⁷/ml for B. brevis, determined through acridine orange direct counts.

Eight series of the experiments were conducted for each experimental condition. Four of the series contained the bacterial suspension with 0.5-, 1.0-, and 2.5-mm-diameter glass beads and without glass beads, and the other four series were similar but contained the nonionic surfactant Tween 80 at a final concentration of 0.12 mM. Tween 80 was used because it reduces bacterial attachment to the glass surface (27) and minimizes cell clumping (1, 41), and it has been shown to have no effect on bacterial metabolic activity at concentrations up to 0.24 mM (53). For each experimental condition, a total of 240 sealed, round-bottomed glass vials (16 by 100 mm; volume of 12 cm³) were prepared. The vials were first filled with 8 g of glass beads and 4 ml of bacterial suspension. The same volume of bacterial suspension was also added to the vials without glass beads. All vials were sealed and placed horizontally on an Oribitron shaker (Boekel Scientific) in an incubator at 25°C. The vials were orbitally rotated at 8 to 10 rpm with a tray tilt angle of 13° to provide a gentle mixing of the bacterial suspension without moving the glass beads in the vials. At specified times, three vials were sampled from each experimental series; two vials were used for ATP analysis and one vial was used for cell counting.

For the ATP analysis, 4 ml of NRB was added to each vial to extract the bacterial ATP. NRB consisted of a 0.05% solution of alkyldimethylbenzylammonium chloride in Tris-Mg²⁺ buffer (20 mM Tris, 2 mM EDTA, and 10 mM Mg²⁺ [added as Mg acetate]; adjusted to pH 7.75 with acetic acid) (9, 29, 30). NRB was freshly prepared before each use. The vials were thoroughly mixed and then sonicated for 2 min in an ultrasonic cleaning bath. Extract (200 µl) from each vial was carefully transferred to polystyrene tubes in duplicate and stored frozen at –15°C until the ATP measurement was performed.

For the planktonic cell counts, 1 ml of bacterial suspension was taken from each vial, transferred to a tube containing 1 ml of 4% formaldehyde solution, and then stored at 4°C until the cell number was counted. The bacteria were stained using acridine orange and then counted via epifluorescence microscopy using a minimum of 20 fields. The number of attached bacteria was calculated by subtracting the number of planktonic bacteria from the total number of bacteria added to the vial.

ATP analysis.
For the ATP analysis, a luciferin-luciferase enzyme solution was freshly prepared before each use (4, 29, 30). Luciferase was prepared by adding 1 mg of luciferase (Sigma) to 1 ml of Tris buffer (20 mM Tris and 2 mM EDTA; adjusted to pH 7.75 with acetic acid) and stored frozen in 25-µl aliquots. Prior to use, one aliquot of luciferase was dissolved in 5 ml of Tris-albumin buffer (60 mM Tris, 2 mM EDTA, 150 mM Mg acetate, 50 µM dithiothreitol, and 1.0 g bovine serum albumin; adjusted to pH 7.75 with acetic acid). Luciferin was prepared by adding 1 mg of luciferin (Sigma) to 5 ml of Tris-albumin buffer. To prepare the combined luciferin-luciferase solution, the two solutions were gently mixed together in an amber glass vial and protected from light. This solution was kept at room temperature for at least 30 min before use.

To measure the total ATP level per vial, the frozen ATP extract samples were thawed at room temperature. Then, 100 µl of Tris-Mg²⁺ buffer was added to each sample and the tubes were mixed thoroughly for 3 s using a vortex mixer. Each tube was placed in a Sirius luminometer (Berthold Detection System, Germany), and 100 µl of luciferin-luciferase enzyme was added to the sample. The relative light units from the luminometer were converted to ATP by use of standard curves prepared with ATP standard (Sigma) in a manner identical to that for the experimental samples.

To ensure that the experimental conditions did not have an effect on the ATP measurements, standard curves were prepared for each experimental condition. This resulted in the development of 16 standard curves (prepared in duplicate) for the following combinations: (i) PBS or PIPES, (ii) with and without Tween 80, and (iii) with each of the three different sizes of glass beads and without glass beads. Seven different ATP concentrations were used to develop the standard curves. Results from the standard curve analysis showed a linear relationship between ATP and the luminometer response, with minimal differences in the curves between the different experimental conditions.

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Planktonic and adhered cell counts.
Examples of results for the adhesion of E. coli onto the glass beads in PBS are presented in Fig. 1. The total numbers of bacteria per control vial (no glass beads) were constant over the 144-h experimental period. In the absence of Tween 80, the planktonic cell concentration decreased with the increasing surface area of the glass beads, indicating that the number of adhered cells increased. All cell counts in the presence of the glass beads were statistically different from the cell counts without beads (P < 0.05, n = 20). Conversely, the planktonic cell counts in the presence of Tween 80 were very similar to the counts for the control vials, indicating that the surfactant reduced bacterial adhesion onto the glass beads. Data analysis indicated that in the presence of Tween 80 there was no statistical difference between the cell counts with beads and those without beads (P > 0.05, n = 20), with the exception of the 0.5-mm beads at 48 and 96 h. The results of the other adhesion tests for E. coli in PIPES and for B. brevis in both buffers revealed similar trends (data not shown).

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FIG. 1. Numbers of planktonic E. coli cells in vials containing 8 g of glass beads with diameters of 0.5 mm, 1.0 mm, and 2.5 mm and vials without glass beads, in PBS and PBS with Tween 80. Error bars depict ±1 standard deviation (n = 20).

ATP levels for adhered and planktonic bacteria.
The total levels of ATP per vial for E. coli and B. brevis were determined as a function of the glass-bead surface area, and examples of results are presented in Fig. 2 for E. coli in PBS. The total ATP levels per vial increased rapidly and then slowly decreased over the course of the experiment, while for the control vials, the total ATP levels remained relatively constant. Data analysis indicated that all samples with glass beads present were statistically different from the corresponding controls without glass beads (P < 0.05, n = 4). The presence of Tween 80 had no effect on the control vials, indicating that the surfactant did not affect the cellular ATP levels. For the vials with beads, Tween 80 reduced the total ATP level per vial, in correlation with the reduced adhesion seen in Fig. 1. Similar trends for ATP levels per vial for E. coli in PIPES and B. brevis in PBS and PIPES were also observed (data not shown).

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FIG. 2. Total level of ATP per vial containing 8 g of 0.5-mm, 1.0-mm, and 2.5-mm glass beads and no glass beads, for E. coli in PBS and PBS with Tween 80. Error bars depict ±1 standard deviation (n = 4; for each data point, duplicate vials were utilized, and ATP measurements were conducted in duplicate for each vial).

The cell adhesion and total ATP results indicate that the observed increase in the total ATP per vial was related to the number of adhered cells. From these data, the ATP per adhered cell was calculated as follows: ATP/adhered cell = [(total ATP/vial) – (planktonic ATP/vial)]/(number of adhered cells/vial), where the planktonic ATP per vial was obtained from the control vials. The results of this analysis are shown in Fig. 3, and it can clearly be seen that adhesion to the glass beads resulted in a substantial increase in cellular ATP and that the enhanced ATP levels persisted for 4 or more days under these experimental conditions.

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FIG. 3. ATP levels for planktonic cells and adhered cells on 0.5-mm, 1.0-mm, and 2.5-mm glass beads, for E. coli and B. brevis in both PBS and PIPES.

p for adhered and planktonic cells.
These ATP levels for adhered and planktonic cells can be used to calculate the associated p values. To do this, ADP and AMP levels must also be known. While the three adenylates—ATP, ADP, and AMP—are interconverted during metabolism, their total sum within a cell, known as the adenylate pool (AP), remains fairly constant (3, 8, 36) (AP = [ATP] + [ADP] + [AMP]). The relative distribution of the adenylates is expressed as the adenylate energy charge (EC_A), where EC_A = ([ATP] + 1/2[ADP])/([ATP] + [ADP] + [AMP]). The EC_A can vary from 0 (adenine nucleotides are fully discharged and AP = [AMP]) to 1 (adenine nucleotides are fully charged and AP = [ATP]) (2). The EC_A of bacteria during growth is 0.8 (3, 5, 12, 21), and this declines to a constant value of 0.2 to 0.3 over 1 week of starvation (3, 5). While the EC_A varies depending on the growth phase, the fraction of ADP in the AP remains relatively constant at 25 to 36% of the AP (2). For the analysis performed here, it was assumed that the EC_A at the beginning of the experiment (t = 0) was 0.3. It was also assumed that ADP was 35% of the AP. By using these values, along with the experimental ATP values measured at t = 0, the initial values of ADP and AMP were calculated from the equations for AP and EC_A above. This allowed the calculation of AP, which was then used for the remainder of the analysis. Then, given the experimental ATP values, the AMP concentration and the EC_A were determined as a function of time by using the equations for AP and EC_A above.

Examples of relative concentrations of ATP, ADP, and AMP as a function of EC_A for planktonic and adhered cells of E. coli in PBS are shown in Fig. 4. The EC_A values of the planktonic cells ranged from 0.25 to 0.3, while the EC_A values of the adhered cells ranged from 0.4 to over 0.7. Correspondingly, ATP was 10% of the AP for the planktonic cells and ranged from 20% to 60% for the adhered cells. The calculated values are in good agreement with the theoretical curves of Atkinson and Walton (2), suggesting that the assumptions used in the analysis presented here were reasonable.

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FIG. 4. Relative percent concentrations of ATP, ADP, and AMP and corresponding ECA values for E. coli in PBS. The lines represent theoretical concentrations (2), and the symbols represent the calculated values for planktonic cells (open circles) and for adhered cells on 0.5-mm (filled circles), 1.0-mm (filled squares), and 2.5-mm (filled triangles) glass beads.

Now, p is calculated directly from the phosphorylation potential, G_p (kJ/mol), which is defined as follows: G_p = G^o + RT · ln([ATP]/[ADP][P_i]), where G^o is the standard free energy for ATP hydrolysis, with a reported value of 30.1 kJ/mol (23), R is the gas constant (8.314 J/K · mol), T is the temperature (298 K), and P_i is the intracellular inorganic phosphate concentration (mol/liter). The P_i of starved E. coli is 14 mM, and that of actively growing bacteria is 5 mM (23). Thus, the value of P_i at the beginning of the experiments for the starved bacterial cultures used in this study was assumed to be 14 mM. This allowed the calculation of the total intracellular phosphate concentration (C_T,PO4), where C_T,PO4 = 3[ATP] + 2[ADP] + [AMP] + [P_i]. Then, given the ATP, ADP, and AMP concentrations as a function of time, G_p was calculated via the equations for G_p and C_T,PO4. Finally, p was calculated from G_p as follows: –np = G_p/F, where n is the number of protons translocated by the ATP synthase enzyme per molecule of ATP synthesized and F is the Faraday constant. The value of n is typically reported in the range of two to three for bacteria (23, 28, 35, 45, 50), and the results of these studies indicate that n may vary as an inverse function of p, i.e., n decreases as p increases (45). This suggests that for the current study, where ATP levels increased upon adhesion to the glass beads, n may lie near the lower end of the reported range. Thus, in the current analysis, the value of n = 2 was used to calculate p.

The results from this analysis are shown in Fig. 5, where p is plotted as a function of time for E. coli and B. brevis in PBS. For the planktonic cells, p decreased from –200 mV to approximately –190 mV over the course of the experiment, whereas for the adhered cells, p increased rapidly to approximately –220 mV and then decreased slowly over the course of the experiment. These values are within reported ranges of p for bacteria, which extend from –140 mV to over –220 mV (22, 51). Ultimately, the results shown in Fig. 5 suggest that only a 10% to 15% increase in p is necessary to account for the two- to fivefold increase in ATP observed upon cell adhesion to the glass beads.

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FIG. 5. p for planktonic and adhered cells on 0.5-mm, 1.0-mm, and 2.5-mm glass beads, for E. coli and B. brevis in PBS.

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Several possibilities for the high ATP levels of adhered bacteria compared to those of planktonic bacteria may exist. One possibility may be the enhanced availability of growth substrate at the solid surface. However, in this study the bacteria were washed three times and starved for 12 h before use, and thus the opportunity for residual glucose in solution was minimal. Furthermore, since glucose has negligible sorption onto surfaces, any residual glucose in solution would have affected both planktonic and adhered bacteria similarly (7). Another possibility is that adhesion to a solid surface prevents the diffusional loss of protons from the cell membrane, resulting in a higher concentration of protons near the surface and ultimately increasing p (10, 25). However, as pointed out by van Loosdrecht et al. (49), this specific mechanism is unlikely because (i) this diffusional loss of protons from bacterial cells has never been observed and (ii) even if the loss does occur, the creation of a localized increase in protons in this manner is highly unlikely due to the rapid diffusion of protons through both the periplasmic space and the bulk medium.

Here, we propose a mechanism where a localized enhancement in p occurs at the adhesion interface (e.g., the interface between the cell and glass surfaces). However, rather than being due to the prevention of a loss of protons by the solid surface, it is due to an increase in the local proton concentration resulting from electrostatic interactions.

Biological surfaces, such as the bacterial cell surface, acquire a charge in water through the dissociation of acidic and basic groups at the cell surface. As a surface containing ionizable groups approaches another surface, electroneutrality requires the counterion concentration to increase in the solution between the surfaces to offset the decrease in volume. As H⁺ is a counterion for negatively charged surfaces, its concentration next to a negatively charged surface tends to increase, decreasing the surface pH, which shifts the dissociation equilibrium in favor of fewer dissociated acid groups and a lower surface charge density () (17, 38). Simultaneously, the increased counterion concentration changes the magnitude of the surface potential () (17, 37). Because of these processes, neither nor remains fixed during the approach of these surfaces. This is the well-known charge regulation effect (6, 14, 34, 37, 39), and the effects of the functional groups on and can be determined by solving the Poisson-Boltzmann equation using boundary conditions that consider the multiple dissociation equilibria of ionizable groups on each surface (34, 37, 39).

The hypothesis proposed here is that bacteria can take advantage of a link between cellular bioenergetics (p and ATP formation) and the physiochemical charge regulation effect. As the bacterium approaches the surface, the charge regulation effect causes both and pH at the cell surface to vary as a function of separation distance (6, 14, 34, 37, 39). If the surface that the bacterium is approaching is negatively charged, such as the glass beads used in this study, the charge regulation effect results in a drop in pH at the cell surface. The hypothesis is that this drop in pH at the cell surface enhances the proton gradient across the cytoplasmic membrane, with a resulting increase in p and cellular ATP. Conversely, if the cell approaches a positively charged surface, this results in an increase in pH at the cell surface and a corresponding drop in p and ATP. Observations in support of the former have been shown here, where adhesion to the negatively charged glass surface resulted in enhanced ATP concentrations. Supporting observations for the latter have been reported previously, where positively charged surfaces resulted in a loss in cell viability and increased cell death (13, 26, 44, 47).

Following the chemiosmotic theory, when cells are actively utilizing growth substrate, protons are extruded across the cell membrane to the periplasmic space as electrons are passed from the growth substrate to the terminal electron acceptor via membrane-bound enzymes. This process results in the generation across the cell membrane of pH and , which combine to form p (33). At 25°C, this relationship can be written as follows (33): p = – 59pH. These protons are then allowed back into the cell in a controlled manner to form ATP. This overall process is called oxidative phosphorylation. In contrast, under starvation conditions, planktonic bacteria cannot generate ATP via oxidative phosphorylation because the extrusion of extra protons does not occur. In this case, bacteria attempt to maintain p by using ATP to drive protons across the cell membrane (33). However, following the proposed hypothesis, when starving cells approach a negatively charged surface, the local pH at the cell surface decreases (proton concentration increases), which in turn enhances the proton gradient across the cell membrane and enhances the formation of ATP (Fig. 6). In this manner, bacteria can take advantage of the charge regulation effect and maintain an ATP level higher than that associated with the corresponding planktonic cells.

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FIG. 6. Proposed hypothesis for a gram-negative bacterial cell adhering to a negatively charged surface (e.g., E. coli adhering to the glass surface used in this study). The charge regulation effect results in an increase in local proton concentration between the cell and the negatively charged surface. This increase in proton concentration enhances p, which increases the ATP level within the cell.

The question arises as to whether the increase in proton concentration affects p through changes to either pH or or both simultaneously and whether the changes are sufficient to result in the increase in p shown in Fig. 5 for adhered bacteria. The extrusion of protons across the cell membrane can affect both pH and , depending on whether the extrusion is done electroneutrally (which affects pH) or via charge separation (which effects ). Both processes occur during oxidative phosphorylation, and for neutrophilic bacteria, accounts for 70 to 80% of p, with pH contributing 20 to 30% (51). When bacterial adhesion is considered, the formation of an enhanced proton gradient via the charge regulation effect (Fig. 6) would be electroneutral, since the protons are present as counterions to the ionized surface functional groups rather than due to charge separation across the periplasmic membrane. In this case, only pH would be altered.

To determine the change in pH necessary to provide the observed ATP increases, it was assumed that was equal to –150 mV, i.e., 75% (51) of the p value of –200 mV at t = 0 (Fig. 5), and that the subsequent variation in p over time was due solely to changes in pH via the charge regulation effect. The difference in pH between adhered and planktonic cells was then calculated using the p data from Fig. 5 and the equation p = – 59pH. The results of this analysis, presented in Fig. 7, show that the difference in pH between adhered and planktonic cells [(pH)_adhered – (pH)_planktonic] is approximately 0.2 to 0.5 pH units. Thus, only a minor shift in pH at the membrane surface is required to achieve the ATP levels observed for adhered bacteria (Fig. 3).

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FIG. 7. Difference in pH between adhered and planktonic cells for E. coli and B. brevis in PBS.

In summary, it was demonstrated here that for bacteria under starvation conditions, adhesion to the glass surface resulted in cellular ATP levels that were two to five times higher than those for the corresponding planktonic bacteria. It is proposed that this elevated ATP is due to a link between cellular bioenergetics (p and ATP formation) and the charge regulation effect, which occurs at the cell surface. Following the hypothesis, the charge regulation effect increases the local proton concentration at the interface between the bacterium and the negatively charged glass surface, which enhances p and ATP formation. Similarly, the hypothesis suggests that positively charged surfaces should reduce p and ATP formation, and this is supported by previous results. Ultimately, if this hypothesis is shown to be true, it will provide a framework for interpreting interactions of bacteria with surfaces and may be used to design or select surfaces for desired effects on bacterial metabolic activity.

 
We gratefully acknowledge the National Science Foundation's valuable support for this work through CAREER grant 0134362.

 
* Corresponding author. Mailing address: Department of Civil and Environmental Engineering, Lehigh University, 13 East Packer Avenue, Bethlehem, PA 18015. Phone: (610) 758-3543. Fax: (610) 758-6405. E-mail: dgb3{at}lehigh.edu

Published ahead of print on 13 February 2009.

Present address: Department of Chemical and Environmental Engineering, University of California, Riverside, Riverside, CA 92521.

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Applied and Environmental Microbiology, April 2009, p. 2346-2353, Vol. 75, No. 8
0099-2240/09/$08.00+0     doi:10.1128/AEM.02671-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

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