Environ. Sci. Technol. 2005, 39, 3116-3120

Effect of Chemical Speciation on Toxicity of Mercury to Escherichia coli Biofilms and Planktonic Cells ISAAC NAJERA, CHU-CHING LIN, GOLENAZ ADELI KOHBODI, AND JENNIFER A. JAY* Civil and Environmental Engineering Department, University of CaliforniasLos Angeles, 5732 H Boelter Hall, Los Angeles, California 90095-1593

While it is known that microbial uptake of mercury (Hg) by planktonic cultures is influenced by the extracellular speciation of mercury in aquatic systems, Hg uptake in biofilm cultures is understudied. We compared the importance of Hg(II) speciation in toxicity to both planktonic and biofilm cultures of the Gram-negative bacterium Escherichia coli O55. Variable chloride chemistry experiments were carried out to modify mercury speciation. Biofilms were observed to be more resistant to Hg than planktonic cells. In both planktonic and biofilm cultures, the toxicity of Hg increased and then decreased along the chloride gradient. The percent reduction in cell viability was linearly related to the concentration of HgCl20 when Hg-chloro complexes dominated the speciation, consistent with a passive diffusion model. However, toxicity to both planktonic cells and biofilms at low salinities could not be explained by passive diffusion alone, which suggests that microbial uptake of Hg in both planktonic cells and biofilms may occur by both passive diffusion of neutral species and facilitated uptake. The relationship between toxicity and chloride concentration was similar in the presence and absence of a biofilm, indicating that the presence of the biofilm does not drastically change the relative availability of the dominant mercury species.

Introduction In aquatic systems, inorganic mercury is microbially transformed to methylmercury, a very toxic form of Hg(II) that is readily biomagnified through food webs (1, 2). The degree to which the biological, chemical, and physical parameters in an aquatic system promote conditions favoring microbial methylation are strong determinants of methylmercury accumulation in fish (3, 4). Microbial uptake of mercury is a key step in the methylation, bioaccumulation, and reduction of Hg(II) to elemental mercury in aquatic systems. Current uptake models propose that mercury permeates the cell membrane either by passive diffusion of lipophilic uncharged complexes, such as HgCl2 and HgS(aq) (2, 5-7), or by facilitated uptake (8-10). To date, all of the work concerning uptake of specific species of Hg(II) by pure cultures has been carried out with batch planktonic cultures (2, 8, 9, 11, 12). However, it is now well-established that the majority of bacteria in the environment live in attached communities, or biofilms (13). Recently, * Corresponding author phone: (310) 267-5365; fax: (310) 2062222; e-mail: [email protected]. 3116

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attached bacterial communities, or biofilms, have been hypothesized to be a critical site for methylation of mercury in the Florida Everglades (14, 15), an area which experiences high mercury contamination in the aquatic food web. Biofilms may greatly affect the cycling of mercury in the environment by providing an anoxic niche for methylating organisms in environments that would be characterized as oxic at the bulk scale (16-19). Steep redox and chemical gradients within biofilms will result in rapid cycling of sulfur species to drive the metabolism of sulfate reducers (14, 16, 20) and may change the speciation of mercury at equilibrium in different regions of the biofilm (21, 22). Also, mercury transformations within microbial biofilms may be strongly influenced by both physiological changes within the bacteria and a barrier effect of extracellular polymeric substances (EPS) (23, 24). Various species of mercury might be expected to diffuse through biofilms at different rates, as the effective diffusion coefficient would be dependent on sorptive affinity to the EPS and the size of the permeant. The relatively small body of literature on the interaction of Hg with biofilms indicates that accumulation of both inorganic and methylHg can occur in biofilms (25) and that Hg is available for reduction at a wide range of salinities in attached communities (26). The goals of this work are as follows: (I) to test the hypothesis that uncharged mercury species are the main cell permeants in both biofilm and planktonic cultures of Gram-negative bacteria by testing mercury toxicity along a chloride gradient and (II) to compare the relative mercury uptake rate of planktonic cultures and biofilms. Escherichia coli (E. coli) O55 was selected as a model Gram-negative organism to study Hg(II) uptake in biofilms and planktonic cultures since many of the methylating bacteria (i.e., sulfate reducing bacteria (SRB)) are also Gram-negative. Data from this study demonstrate that the toxicity, and presumably uptake, of Hg(II) to E. coli is strongly influenced by the Hg chemical speciation in the growth medium for both planktonic and biofilm cultures. In particular, our data suggest that both passive diffusion of lipophilic uncharged complexes and facilitated uptake of positively charged species may cause reduced cell viability in E. coli cells over a wide range of salinities. Further, we report that biofilm cultures of E. coli were more resistant than planktonic cells to Hg(II) at every chloride concentration. These findings improve our understanding of mercury uptake mechanisms of gramnegative bacteria in both planktonic cultures and biofilms.

Materials and Methods Bacterial Strain and Growth Conditions. The bacterial strain used in this study was E. coli O55 (ATCC 12014). Cultures were grown in glucose minimal salts medium (GMM), pH 6.8, with 7 g L-1 glucose as the sole carbon source for the planktonic cultures. Casamino acids (CAA) were added to a final concentration of 0.5% (w/v) as the sole carbon source for biofilm growth but not for toxicity experiments (27). The media contained 7 g L-1 K2HPO4, 2 g L-1 KH2PO4, 1 g L-1 (NH4)2SO4, 200 mg L-1 MgSO4‚7H2O, 20 mg L-1 CaCl2‚2H2O, and 2-10 µg L-1 trace elements (Fe, Co, Mn, Zn, Cu, Ni, and Mo) (28). Media were sterilized by autoclaving. Experiments were conducted at room temperature, which was 25 ( 1 °C. Since E. coli forms biofilms in a nutrient-dependent fashion (29), we studied biofilm formation in different growth media at different dilution rates. Half-strength CAA medium (50% (v/v) CAA medium/Milli-Q water) was found to be the optimum medium for E. coli under our experimental conditions. 10.1021/es048549b CCC: $30.25

 2005 American Chemical Society Published on Web 03/25/2005

Thermodynamic Speciation. The USGS geochemical equilibrium speciation model PHREEQC ver. 2.0 (30), with the constants from MINTEQ database, was used to calculate the speciation of Hg(II) in the experimental medium (in the absence of CAA). The model took into account all ingredients of the medium except trace elements, and all Hg complexes for which published formation constants exist. Equilibrium modeling of Hg(II) was carried out at varying levels of chloride (0.27, 1, 10, 100, and 200 mM; with their corresponding measured pH), strict aerobic conditions (pe )10), and T ) 25 °C. Uptake Rate Estimation. The maximum uptake rate of a species crossing the cell membrane by passive diffusion, V (attomol cell-1 day-1), can be estimated as V ) PCaqA, where P is the permeability of the cell membrane to a particular species (cm s-1), Caq is its extracellular aqueous concentration (attomol cm-3; the intracellular concentration is neglected), and A is the area of a cell (cm2 cell-1) (2, 31). P can be defined as the rate at which a complex crosses cell membranes by passive diffusion, and P for a given species is a function of both Kow and molar volume. An empirical relationship exists between Kow and P* (2, 31), in which P* is defined as the theoretical limit for permeability of a molecule. P can then be estimated from the empirical relationship: log P ) log P* - mvv, where mv is the size selectivity of membranes (0.0546 mol cm-3) and v is the estimated molar volume of the permeant (cm3 mol-1) (31). This approach has been used in the case of uptake of neutral mercury species by phytoplankton (2) and sulfate-reducing bacteria (32, 33). Relative Resistance of Planktonic and Biofilm Cells. Bioassay medium was prepared by adding NaCl to GMM to a final concentration of 0.27, 1, 10, 100, and 200 mM of chloride. Different salinities showed slightly different pH values (6.82, 6.65, 6.64, 6.58, and 6.57, respectively). A Hg(II) stock solution was added to a final concentration of 250 ppb, or 1.25 µM, Hg(II)T. This high concentration was chosen so that toxicity to the cells would be observed in biofilms. Biofilms were grown on single frosted microscope slides (Corning 2948). Sterile, 50 mL Falcon tubes were filled with 18 mL of 50% CAA medium and inoculated with 2 mL of overnight culture (8.5 × 105 cells mL-1). Sterile microslides were placed into Falcon tubes and incubated under static conditions for 24 h at 25 ( 1 °C (27). Biofilms of 104 cells cm-2 (as measured by viable plate counts) were formed after 24 h, with a total biofilm surface (two sides) of 15 cm2. Biofilms were rinsed three times with sterile phosphate-buffered saline (PBS) to wash out loosely attached cells before exposure to mercury. Slides were placed into new Falcon tubes containing bioassay medium (which did not contain CAA). Biofilms were exposed to mercury for 24 h and then rinsed three times in sterile PBS. Washed biofilms were scraped with stainless steel disposable scalpels and collected into 30 mL of sterile phosphate buffer. As previously described by Costerton et al. (13) and Chen and Stewart (34), detached bacterial aggregates were dispersed by mechanical shaking (vortexed for 30 s) and ultrasonication (5 min). Dilutions of the suspension containing biofilm cells were plated in quadruplicate for colony counting after a 24 h incubation on solid GMM medium (containing 15 g/L agar). Aliquots were kept in ice during viable plate counts processing. Colony morphology was monitored to check for contamination. Batch planktonic stationary-phase cultures were prepared by inoculating 1 mL of 18 h overnight culture (106 cells mL-1) into 50 mL sterile Falcon tubes containing 19 mL of GMM and incubating at 25 ( 1 °C for 24 h. Samples were incubated with orbital shaking (speed of 200 rpm) to prevent biofilm formation. Incubated samples, containing E. coli cells in the stationary phase, were centrifuged at 4000 rpm for 5 min and resuspended in 20 mL of fresh glucose medium containing 250 ppb of Hg(II)T and variable NaCl concentra-

FIGURE 1. Calculated speciation of main dissolved Hg(II) species in glucose medium as a function of chloride using PHREEQC (where, for example, 1.0E-05 represents 1.0 × 10-5). Calculations were carried out at 250 ppb Hg(II)T, pe ) 10, and T ) 25 °C, at the measured pH for each solution. tions. Exposed cells were incubated under the same conditions for an additional 24 h. After exposure, samples were vortexed for 30 s and sonicated for 5 min. Viable cells were processed as previously described.

Results Resistance of Stationary-Phase Planktonic Cells versus Biofilm. Chloride chemistry in the glucose growth medium was varied between 0.27 and 200 mM, while Hg(II)T was held constant at 250 ppb in order to test the hypothesis of increasing bioavailability, and hence toxicity, of mercury at increasing molar fraction of HgCl2 in both planktonic and biofilm cultures. Salinity changes in the medium drastically affected the speciation of mercury. All Hg complexes with a molar fraction above 1% are represented in Figure 1. At low chloride (0.27 and 1 mM), the speciation is dominated by the charged Hg(NH3)22+, accounting for the 97 and 95.2% of the total Hg, respectively (Figures 1 and 4). It should be noted that the artificial nature of the growth medium results in the dominance of the diammonia complex, which is likely unimportant in nature. The only significant uncharged complex is Hg(OH)2 totaling around 2.7% in both cases. The dominant complex of mercury becomes HgCl2 at 10 and 100 mM with a respective abundance of 52.1 and 47.2%. Further increase in the salinity of the medium results in a shift in mercury speciation to charged forms HgCl3- and HgCl42(Figure 1). Figure 2 depicts the maximum predicted uptake rate, V (attomol cell-1 day-1), calculated from our thermodynamic speciation data and P values for each uncharged species. Values of Kow for HgCl2 (3.33), HgClOH (1.20), and Hg(OH)2 (0.05) and molar volumes of 51 cm3 mol-1 for HgCl2 and 43 cm3 mol-1 for Hg(OH)2 were obtained from the literature (1, 2). These Kow values were confirmed by Mason et al. (2) across a chloride gradient of relevance for our study (10-4 to 10-1 M). A molar volume of 47 cm3 mol-1 for HgClOH was estimated (35). The resulting estimated P values were 1.70 × 10-3, 7.25 × 10-4, and 1.77 × 10-5 cm s-1 for HgCl2, HgClOH, and Hg(OH)2, respectively. For maximum uptake rate calculations a measured P value for HgCl2 of 7.4 × 10-4 cm s-1 was used instead of the estimated one (2). Due to their much lower molar fraction (at salinities above 1 mM) and lower Kow, Hg(OH)2 and HgClOH are expected to have a smaller contribution to diffusive mercury uptake when compared to HgCl2. E. coli biofilms were first grown on microslides in the absence of mercury before the experiment at variable VOL. 39, NO. 9, 2005 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

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FIGURE 2. Observed toxicity on planktonic and biofilm cultures as a function of the calculated maximum uptake rate of neutral chloride complexes (where, for example, 1.0E-01 represents 1.0 × 10-1)).

FIGURE 3. Observed toxicity on planktonic and biofilms cultures as a function of the chloride gradient (where, for example, 1.E+09 represents 1.0 × 109). Samples were incubated at 25 ( 1 °C during 24 h in glucose medium containing 250 ppb Hg(II)T. Bars represent standard deviations of quadruplicates. chemistry. The biofilms completely coated the submerged portion of the microslide, showing a very pronounced growth in the air-liquid interface. Microscopic examination of the biofilms confirmed that the highest cell density was in the air-liquid interface in the form of a multilayer biofilm. This is a representative biofilm formation phenotype of E. coli in CAA medium (27). Controls of both stationary-phase and biofilm cells showed decreasing cell counts when exposed to increasing levels of NaCl (Figure 3). This effect was much more pronounced in the biofilm cultures. At 200 mM chloride, detachment of biofilm cultures led to much less cell density in both the presence and absence of Hg due to osmotic stress (Figure 3). In the presence of Hg in stationary-phase planktonic cells, there was a decrease in survival as the chloride concentration increased from 0.27 to 10 mM (Figures 2-4) (the molar fraction of HgCl2 increases from less than 0.1 to 52.1% in this range). Similar decreases in cell counts were recorded at 10 and 100 mM, 59.5 and 58.9%, respectively, concentrations at which HgCl2 is the dominant species. At 200 mM a noticeable reduction in toxicity was accompanied by a smaller mole fraction of HgCl2 (25.8%). The viability reduction in exposed cells versus controls was 17.6, 42.2, 59.5, 58.9, and 17.3% respectively (Figures 3 and 4). In planktonic cells, the reduction in cell viability was not linearly related to the HgCl2 estimated maximum uptake rate in the range of 0.27-200 mM with a R2 ) 0.484. This linear relationship slightly improved (R2 ) 0.504) when the combined uptake rate of the neutral species was evaluated. A 3118

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FIGURE 4. Observed toxicity on planktonic and biofilm cultures as a function of the calculated concentrations of Hg(NH3)22+ and HgCl2 (where, for example, 1.4E-06 represents 1.4 × 10-6). very strong linear relationship between the cell viability reduction and estimated HgCl2 uptake rate was reached when mercury chloride complexes were dominant (10-200 mM), with a R2 ) 0.976 (Figure 3). Biofilm E. coli cells were found to be more resistant than stationary cultures when exposed to 250 ppb Hg(II)T (Figures 3 and 4). In biofilms, cell viability reduction was never greater than 30%. Further, the reduction again increases and then decreases along the chloride gradient. At the lowest chloride concentration, 0.27 mM, as well as the two highest concentrations, 100 and 200 mM, Hg treatments were not significantly different from the control. Significant reduction in cell viability was seen at midrange chloride concentrations (29.2% at 1 mM and 29.5% at 10 mM). In the range 10-200 mM, assuming no reduction in cell viability at 200 mM, a good correlation (R2 ) 0.717) was obtained between the combined estimated uptake rate of the lipophilic uncharged species and the viability reduction (Figure 2).

Discussion Evidence for Both Passive Diffusion and Facilitated Uptake in Stationary-Phase Planktonic Cells. The speciation of mercury has been shown to be an important determinant of its biological uptake. In planktonic cultures, a strong linear relationship between cells viability reduction and HgCl2 estimated uptake was reached when Hg-chloro complexes were the dominant species, from 10 to 200 mM, with a R2 ) 0.976 for stationary-phase cells. Maximum reduction in cell viability in planktonic cells was reached at 10 and 100 mM, concentrations at which HgCl2 clearly dominates the speciation. The maximum uptake, and hence toxicity, in planktonic cells occurred between 1 and 100 mM, salinity range in which the Hg-permeability to biological systems is maximum (5, 6) showing a clear effect of Cl- on Hg(II) bioavailability (Figure 2). Our data show a marked decrease in the toxicity of mercury at 200 mM chloride, a concentration at which the speciation of mercury shifts such that the dominant species are negatively charged Hg-chloro complexes (HgCl3-, HgCl42-). These species have been shown to have a reduced bioavailability to E. coli cells (6) and artificial cell membranes (5). This reduced uptake of mercury to bacteria may be one reason to explain why lower methylation rates and methyl-Hg accumulation are generally reported in estuarine and marine systems (36). Thus, uptake at high chloride concentrations seems to have the characteristics of passive diffusion. However, if lipophilic uncharged Hg-chloro complexes were solely responsible for the reduction in cell viability, the toxicity would exclusively depend on the neutral mercury

complexes permeability and concentration gradient across the membrane. This is not the case in our experiments. A significant toxicity was also noted when the charged species Hg(NH3)22+ dominated the Hg speciation with 97 and 95.2%, respectively (Figure 4). Thus, uptake of Hg as a positively charged complex appears important and may indicate a facilitated uptake of this species (8). Farrell et al. (11) reported that cationic metal-ligand species are considered to be more toxic than anionic or neutral complexes, with Hg2+ being the most bioavailable form of inorganic mercury. Our results suggest that Hg(NH3)22+, with a concentration at least 102fold higher than Hg(OH)2 and 104-fold higher than HgCl2, seems to play a dominant role in mercury toxicity at 0.27 and 1 mM. Laporte et al. (10) also suggest that uptake of mercury in crab tissue can involve both active diffusion and passive mechanisms. They reported that uptake is reduced by temperature and in the presence of Na+K+ATPase inhibitor, showing a metabolic influence in the uptake that cannot be explained by diffusive flux into the cells. Also, experiments with Vibrio anguillarum genetically engineered with a merlux bioreporter plasmid showed a strong pH dependency of mercury uptake between pH 6.3 and 7.3 which could not be explained by differences in the concentration of neutral species (9). Another study indicated that neutral or positively charged complexes such as Hg(NH3)2+ and Hg complexed with small organic ligands appeared equally toxic (8). Our results suggest that both mechanisms, facilitated uptake of species such as Hg(NH3)22+ and passive diffusion of neutral species such as HgCl2, HgClOH, and Hg(OH)2, play an important role in toxicity and, hence, in the biological uptake in planktonic cells. Role of Biofilms in Mercury Uptake and Resistance. Saline stress was observed in both planktonic cells and biofilms (Figure 2). Chen and Stewart (34) reported that 0.3 M NaCl can cause more than a 25% removal of Pseudomonas aeruginosa and Klebsiella pneumoniae biofilm biomass, accompanied by more than 30% reduction in viable cell numbers, within a few hours. Therefore, the exponential dieoff observed in biofilms may be a consequence of both EPS chemical detachment and cell inhibition due to osmotic stress. Our data showed that E. coli biofilms were more resistant to 250 ppb Hg(II) than planktonic cells at every salinity level, from 0.27 to 200 mM (Figures 3 and 4). These findings are consistent with the current evidence regarding biofilms resistance to antimicrobials and heavy metals. Biofilms generally exhibit an increased tolerance to antibiotics if compared to planktonic cells (37-39). This resistance has also been shown for heavy metals (Cu2+, Zn2+, and Pb2+) (40) and metalloid oxyanions (41). Similar resistance to antimicrobials of stationary-phase planktonic and biofilms cells may be observed (42). In some cases, increased resistance of biofilm cells compared to planktonic organisms may be explained by slow aqueous diffusive transport in biofilms (19, 43). However, in our experiments, the observed increased resistance to Hg in biofilms is probably not due simply to a decrease in the diffusivity of the surrounding layer. The maximum Hg uptake rate per cell (VHg) based on aqueous layer diffusion limitation can be calculated according to VHg ) 4πRDaqCbulk (Fenchel et al. (44)), where VHg ) uptake in mol cell-1 s-1, Cbulk ) concentration of the Hg species of interest in the bulk medium, Daq ) diffusion coefficient for solute in water, and R ) radius of the cell (assumed ∼10-4 cm). The diffusion coefficient (Daq) was taken as 2 × 10-5 cm2 s-1, based on the observed linear relationship for molar volume of neutral molecules and the diffusion coefficient in water (45), recently applied to mercury compounds (46). Under our experimental conditions, this estimated Hg uptake rate is at least 6 orders

of magnitude faster than the estimated uptake by passive diffusion of HgCl2 through cell membranes (Figure 3). The effective diffusion coefficient in a biofilm, De, is commonly 20-80% of Daq (19). Even at the low end of this range, we would still not expect aqueous or biofilm diffusion to be rate limiting compared to passive diffusive uptake. Another possible explanation for increased resistance to mercury in biofilms is that EPS has been shown to effectively bind metal cations from the bulk solution due to the negatively charged nature of the extracellular polysaccharides (47-52), which can result in increased heavy metal resistance (40). Also, local variations in the concentration of cells, nutrients, and substrates is the hallmark of biofilm growth (13, 19), and some microzones in the biofilm may be less susceptible to mercury toxicity due to their metabolic state (19, 38). In both planktonic and biofilm cultures, the toxicity of Hg increased and then decreased along the chloride gradient. This trend has been previously shown for planktonic cells and provides insight into the availability of Hg as ionic strength increases in a natural setting. In these experiments, a positively charged Hg-ammonia complex appears to be taken up by facilitated transport; however, dominance of a positively charged complex is not commonly important in nature. The similarity in the relationships in the presence and absence of a biofilm indicates that the existence of the biofilm does not drastically change the relative availability of the dominant mercury species. While E. coli is not a methylating bacterium, this work provides valuable insight into the factors controlling Hg availability in biofilms. Understanding the mechanisms that control not only mercury uptake but accumulation and methylation rate in biofilms (species composition and metabolic rates, chemical gradients, EPS characteristics) is crucial to predicting mercury transformation rates in the environment.

Acknowledgments We thank Dianne Newman, Anand Patel, and Tracy Teal for helpful discussion regarding biofilm cultures, and we are grateful to three anonymous reviewers for substantial improvements to the manuscript. This work was funded by a CAREER grant (BES-0348783) from the National Science Foundation.

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(31) (32)

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Received for review September 16, 2004. Revised manuscript received February 11, 2005. Accepted February 15, 2005. ES048549B