Organomercurial Lyase (MerB)-Mediated Demethylation Decreases Bacterial Methylmercury Resistance in the Absence of Mercuric Reductase (MerA)

ABSTRACT

The mer operon encodes enzymes that transform and detoxify methylmercury (MeHg) and/or inorganic mercury [Hg(II)]. Organomercurial lyase (MerB) and mercuric reductase (MerA) can act sequentially to demethylate MeHg to Hg(II) and reduce Hg(II) to volatile elemental mercury (Hg⁰) that can escape from the cell, conferring resistance to MeHg and Hg(II). Most identified mer operons encode either MerA and MerB in tandem or MerA alone; however, microbial genomes were recently identified that encode only MerB. However, the effects of potentially producing intracellular Hg(II) via demethylation of MeHg by MerB, independent of a mechanism to further detoxify or sequester the metal, are not well understood. Here, we investigated MeHg biotransformation in Escherichia coli strains engineered to express MerA and MerB, together or separately, and characterized cell viability and Hg detoxification kinetics when these strains were grown in the presence of MeHg. Strains expressing only MerB are capable of demethylating MeHg to Hg(II). Compared to strains that express both MerA and MerB, strains expressing only MerB exhibit a lower MIC with MeHg exposure, which parallels a redistribution of Hg from the cell-associated fraction to the culture medium, consistent with cell lysis occurring. The data support a model whereby intracellular production of Hg(II), in the absence of reduction or other forms of demobilization, results in a greater cytotoxicity than the parent MeHg compound. Collectively, these results suggest that in the context of MeHg detoxification, MerB must be accompanied by an additional mechanism(s) to reduce, sequester, or redistribute generated Hg(II).

IMPORTANCE Mercury is a globally distributed pollutant that poses a risk to wildlife and human health. The toxicity of mercury is influenced largely by microbially mediated biotransformation between its organic (methylmercury) and inorganic [Hg(II) and Hg⁰] forms. Here, we show in a relevant cellular context that the organomercurial lyase (MerB) enzyme is capable of MeHg demethylation without subsequent mercuric reductase (MerA)-mediated reduction of Hg(II). Demethylation of MeHg without subsequent Hg(II) reduction results in a greater cytotoxicity and increased cell lysis. Microbes carrying MerB alone have recently been identified but have yet to be characterized. Our results demonstrate that mer operons encoding MerB but not MerA put the cell at a disadvantage in the context of MeHg exposure, unless subsequent mechanisms of reduction or Hg(II) sequestration exist. These findings may help uncover the existence of alternative mechanisms of Hg(II) detoxification in addition to revealing the drivers of mer operon evolution.

INTRODUCTION

Mercury (Hg) is a prevalent environmental contaminant and toxicant. The global distribution of mercury over the last century has increased due to anthropogenic activities, including the burning of fossil fuels, small-scale mining operations, municipal waste combustion, and industrial processing of chlor-alkali products (1). Once in the atmosphere, elemental mercury (Hg⁰) can be oxidized to Hg(II) and returned to terrestrial and aquatic ecosystems through wet deposition (2), where it can be converted to methylmercury (MeHg) by anaerobic microorganisms (3–7). Methylmercury is known to biomagnify in aquatic and terrestrial food chains, leading to its accumulation in diverse biotas, ranging from soil microenvironments to the human body (8, 9). The toxicity associated with Hg(II) and MeHg is attributed to their high affinity for amino acid thiols and their resulting perturbation of protein structure and function (10–12). In addition, Hg(II) and MeHg can disrupt the intracellular glutathione and thiol pools (13, 14). These properties call for a strong selective pressure for organisms to detoxify their local environment of Hg compounds or to acquire mechanisms to resist their toxic effects.

Several diverse microbial species of environmental and clinical relevance have been identified that are resistant to Hg compounds (15–18). An even broader array of microbes carrying the mer operon have been discovered and characterized based on genomic data (19). The mer operon can exist integrated in chromosomal DNA or on mobile genetic elements such as plasmids, transposons, and integrons (20–22). The mer operon exists in one of two forms in Bacteria and Archaea, conferring either “narrow-spectrum” or “broad-spectrum” Hg resistance based on the ability of cells to detoxify Hg(II) only or both MeHg and Hg(II), respectively (23, 24). Broad-spectrum resistance is conferred when MerB protonolytically cleaves the carbon-mercury bond in MeHg, yielding Hg(II) and methane, with subsequent reduction of Hg(II) to Hg⁰ by MerA (19, 20). In addition to detoxification enzymes, mer operons encode regulatory proteins such as MerR and MerD and transport proteins such as MerP, MerT, and MerE (18, 19, 25, 26).

A recent phylogenic analysis of mer homologs among ∼84,000 available archaeal and bacterial genomes suggested that MerB was recruited to a MerA-encoding operon relatively recently, supporting the hypothesis of a functional dependence of MerB on the MerA enzyme (27). Unexpectedly, the same study identified genomes that encode homologs of MerB alone, without MerA. In vitro studies of MerB enzymes have confirmed the ability to demethylate MeHg under cell-free conditions, albeit with relatively low catalytic efficiency (28, 29). MerB exhibits much higher efficiency when coupled with MerA (28, 29), whereby turnover is thought to increase when MerA extracts Hg(II) from MerB for reduction (30). However, isolated MerB activity in a relevant whole-cell environment is less well characterized (31). Prokaryotic and eukaryotic systems demonstrate poor cellular uptake of Hg(II) compared to MeHg, which has obscured comparisons of the relative toxicity of Hg(II) versus MeHg once in the intracellular environment (32). Nonetheless, studies employing transporter-deficient mer operons indicate that accumulation of Hg(II) intracellularly propagates elevated cytotoxicity (33). These observations call for a greater understanding of how the interdependence of demethylation and reduction modulates the response to MeHg exposure. Resolving this interdependence will be key in elucidating why merB more commonly occurs in tandem with merA among known MeHg resistance-conferring mer operons and subsequently how MerB might function when not accompanied by MerA.

In the present study, we tested the hypothesis that MeHg exposure would result in a greater toxicity for cells expressing MerB alone than for those expressing MerA and MerB or those that do not express any mer-encoded functionalities. To test this hypothesis, we constructed Escherichia coli strains carrying plasmids encoding MerRETPA and MerB together, MerRETPA only, or MerB only or empty plasmid controls. These transformed strains were then used to characterize MeHg demethylation, Hg species analysis [Hg(II) and MeHg], cytotoxicity, cell lysis, and the cell-associated fate of Hg during growth.

RESULTS

Demethylation of MeHg by MerB can occur independently of MerA.

To assess demethylation and reduction capabilities, E. coli strains expressing various combinations of mer components on plasmids were exposed to MeHg (0.5 μM [0.1 ppm]), and the amount and species of Hg were tracked over time (Fig. 1). In E. coli strains expressing MerA only and strains containing the empty vector, there was no observed change in the amount of MeHg or total mercury [T-Hg; MeHg + Hg(II)] present over the course of 24 h of incubation (Fig. 1A to E). In contrast, all MeHg was depleted following 24 h of incubation in strains expressing both MerA and MerB (Fig. 1A and E). Over this same time frame, a nearly complete loss of T-Hg from the aqueous phase and biomass was observed for MerA- and MerB-containing-strain cultures, presumably due to reduction and volatilization to Hg⁰ into the headspace (Fig. 1C). At the 24-h time point, a 91% loss of T-Hg was seen for the strain carrying MerA plus the MerB from Pseudomonas sp. strain K-62 (K62B) (Fig. 1C and F), while a 61% loss of T-Hg was seen with the strain carrying MerA plus the MerB from Staphylococcus aureus RN23 (RN23B) (Fig. 1C and F). At intermediate time points in these cultures, different amounts of T-Hg were observed: 74% and 14%, respectively, at 10 h and 47% and 11%, respectively, at 18 h, with cultures finally reaching 40% and 9%, respectively, of the initial dose of MeHg remaining at the 24-h time point (Fig. 1C). Together, these results indicate that both MeHg demethylation and Hg(II) reduction occur in cells expressing MerB and MerA, with complete MeHg demethylation but variable amounts of Hg(II) reduction occurring by 24 h after MeHg exposure.

FIG 1.

MeHg demethylation and Hg(II) reduction capabilities of the specified E. coli strains. (A to D). MeHg demethylation (A and B) and Hg(II) reduction (C and D) capability over time was assessed for strains expressing both the MerA and MerB enzymes (A and C) or expressing MerB alone (B and D). MeHg demethylation was assessed by measuring the amount of MeHg in each sample at various time points over 24 h. Demethylation and reduction were assessed by measuring the amount of total Hg (T-Hg) in cultures for each specified E. coli strain at various time points over 24 h of incubation. Data are results from a single representative of three replicates for each construct exposed to 0.5 μM (0.1 ppm) MeHg (0 h). Data are percent MeHg and percent T-Hg remaining of the MeHg concentration that was measured at 0 h in each sample. (E and F). Demethylation (E) and reduction (F) capabilities of all bacterial constructs at 0 h and 24 h after exposure to 0.5 μM MeHg at 0 h. Data are means and SD. Letters indicate pairwise significant differences between samples, where P is <0.05. Two-way ANOVA and Tukey’s HSD were used for all pairwise comparisons. n = 3 to 5.

In strains carrying MerB only, the amount of MeHg in the culture decreased rapidly over time, approaching a complete loss of MeHg at 24 h of incubation (88% loss for K62B, 73% loss for RN23B) (Fig. 1B and E). Interestingly, the kinetics of MeHg degradation in these MerB-only cultures appeared slower than the demethylation seen with the MerA+MerB-expressing strain. Remarkably, no change in the amount of T-Hg was observed even after 24 h of incubation (Fig. 1D and F), indicating that strains expressing only MerB are capable of efficient demethylation of MeHg without the need for subsequent reduction of the generated Hg(II). At intermediate time points, the individual MerB strains (RN23B and K62B) showed different demethylation kinetics (32% and 18% MeHg remaining at 18 h and 27% and 12% MeHg remaining at 24 h, respectively) (Fig. 1B), suggestive of a different enzyme efficiency between the homologs. However, these differences could also reflect variation in plasmid copy number/protein expression levels between the two engineered strains. Nonetheless, taken together, the data illustrate that MerB is capable of degrading MeHg when expressed in the absence of MerA.

MerB-mediated MeHg demethylation inhibits growth and decreases cell viability.

To investigate how MerB-mediated demethylation alone affects cell growth and viability, we first assessed the effects of MeHg on the growth of strains expressing the various Mer combinations (Table 1) using a MIC assay. In the absence of MeHg, all strains grew at the same rate (see Fig. S1B in the supplemental material). In both the empty vector control strain (pET9a) and the MerA-only strain (pHYR1a), the average MICs for MeHg were statistically the same (7.33 μM and 7.17 μM, respectively) (Table 2). The strain expressing both MerA and MerB exhibited the highest MICs (13 to 15 μM) (Table 2), indicative of more MeHg resistance. In contrast, the lowest MIC was observed in the strain expressing MerB alone (4 to 5 μM) (Table 2), indicative of a greater susceptibility to MeHg.

TABLE 1.

Expression constructs used to transform E. coli^a

Construct	mer operon component(s)	Source	Vector backbone	Antibiotic resistance
pET9a			pET9a	Kanamycin
pHYR1a	merRETPA	Bacillus cereus RC607	pHY300PLK	Ampicillin
RN23B	merB	Staphylococcus aureus RN23	pET9a	Kanamycin
K62B	merB1B2	Pseudomonas sp. strain K-62	pET9a	Kanamycin
pHYR1a + RN23B	merRETPA, merB	Bacillus cereus RC607, Staphylococcus aureus RN23	pHY300PLK + pET9a	Ampicillin + kanamycin
pHYR1a + K62B	merRETPA, merB1B2	Bacillus cereus RC607, Pseudomonas sp. strain K-62	pHY300PLK + pET9a	Ampicillin + kanamycin

^aThe host bacterium utilized for all transformations is the chemically competent E. coli strain JM109.

TABLE 2.

MICs for E. coli strains upon exposure to a range of MeHg concentrations^a

Construct	MIC (μM)			ANOVAb
	Avg	Range	SD
pET9a (empty vector)	7.33	7–7.5	0.29	a
pHYR1a (merA)	7.17	6.5–7.5	0.58	a
RN23B (merB)	4.5	4–5	0.5	b
pHYR1a + RN23B (merA + merB)	14.33	13–15	1.15	c

^apET9a was used as an empty vector control and is indicative of bacteria lacking any mercury resistance mechanisms.

^bSignificance of differences in MICs of MeHg, as determined by a one-way ANOVA with Tukey’s HSD for each pairwise comparison. P < 0.05. n = 3.

To further assess the toxicity associated with MeHg for each strain, cell viability was assessed by quantification of CFU. The concentration of MeHg exposure examined (0 or 2 μM [0.4 ppm]) was selected based on prior experiments in which all strains were able to reach a similar optical density at 600 nm (OD₆₀₀) after 48 h of growth in medium containing MeHg in this concentration range (data not shown). Figure 2 shows that all strains displayed a similar viability 48 h after growth in LB medium containing no MeHg (∼1.2 × 10⁸ CFU/mL) (Fig. 2). With 2 μM MeHg exposure, there was a slight, nonsignificant decrease in the number of CFU/mL observed for the strains expressing MerA alone or the empty pET9a vector compared to the unexposed control (29% and 20% decrease in total CFU, respectively). However, a significant decrease in the number of CFU/mL was observed in the strain expressing MerB alone exposed to 2 μM MeHg compared to the 0 μM controls (79% decrease in total CFU). The strain expressing both MerA and MerB showed no decrease in CFU/mL when exposed to 2 μM MeHg. These results indicate that MeHg exposure to cells expressing MerB alone results in impaired cell growth and cell viability compared to strains expressing MerB in combination with MerA.

FIG 2.

Viability of specified E. coli strains after exposure to a sublethal dose of 2 μM MeHg (0.4 ppm), as measured by assessing CFU from each sample following 48 h of incubation. The MIC for the pHYR1a (MerA-only strain) construct was no different from that for a strain carrying an empty vector (data not shown). Data are means and SD. Letters represent significant pairwise differences between samples where P is <0.05. Two-way ANOVA w. Tukey’s HSD were used for all pairwise comparisons. n = 3.

Fate of Hg(II) subsequent to MerB demethylation of MeHg.

We next examined the fate of Hg(II) generated by MeHg demethylation in MerB-expressing cells. We began by assessing the location and species of Hg in each strain following 48 h of growth after exposure to 2 μM MeHg by separating cells from medium by centrifugation, followed by analysis. In all strains, the majority of the T-Hg was cell associated. However, strains that expressed MerB alone had a significantly higher proportion of T-Hg recovered from the extracellular supernatant than all other strains (13 to 31% more in supernatant than the other strains) (Fig. 3A). The MerB-only strain was also the only strain with the majority of T-Hg (80%) as Hg(II) (Fig. 3B). In contrast, more than 99% of the T-Hg in empty-vector controls and MerA-only strains was MeHg (Fig. 3B). The T-Hg recovered from strains expressing both MerA and MerB was much lower than that from the other strains, presumably due to volatilization of Hg⁰. However, the majority of detectable T-Hg (78.8%) was in the form of MeHg at 48 h (Fig. 3B). These results indicate that the formation of Hg(II), without reduction, results in a higher proportion of the T-Hg located in the extracellular environment.

FIG 3.

Location and Hg(II)/MeHg speciation of total mercury (T-Hg) in cultures of specified E. coli strains following 48 h exposure to a sublethal dose of MeHg. (A) Location of T-Hg in cultures of specified E. coli strains after 48 h of growth in 2 μM (0.4 ppm) MeHg. The location of T-Hg was determined by separating cell pellets (cellular) from supernatant (extracellular) and quantifying T-Hg in each fraction. Statistics were used to compare the percentage of T-Hg in the supernatant (extracellular) component between constructs. Letters represent significant differences between samples where P is <0.05. One-way ANOVA with Tukey-Kramer HSD was used for pairwise comparison of means. n = 3 biological replicates. (B) Hg(II)/MeHg speciation of the T-Hg in each sample at the 48-h time point; samples are the same as those used for panel A. n = 3. Data are means and SD.

MerB-mediated formation of Hg(II) from MeHg promotes cell death and lysis.

The increase in toxicity associated with Hg(II) produced from MeHg by MerB, and the accompanying distribution of Hg to the cell medium, indicated that cell lysis may have occurred in strains expressing only MerB. To assess cell lysis, we utilized a LIVE/DEAD staining and imaging method as a probe of membrane integrity/permeability (Fig. 4). Strains were grown for 48 h in the presence (0.4 ppm [2 μM]) or absence of MeHg, at which time they were stained and imaged via epifluorescence microscopy (Fig. 4). Exposure of strains expressing MerB alone to 2 μM MeHg resulted in a significant (P < 0.001 based on one-way analysis of variance [ANOVA]) increase in the percentage of cells with permeable membranes (red cells) compared to 0 μM MeHg controls (an average decrease in permeable cells of 16%). Exposure of strains expressing MerA alone to 2 μM also showed an increase in permeability compared to their 0 μM controls, albeit much less (5%) than the increase seen with MerB alone. In contrast, strains expressing both MerA and MerB showed no effect on membrane permeability following exposure to 2 μM MeHg compared to MerA+MerB strains with no MeHg exposure (Fig. 4G).

FIG 4.

LIVE/DEAD staining and quantification of cell membrane integrity of specified E. coli strains following exposure to MeHg. (A to F) Epifluorescence images of cultures of specified E. coli strains following 48 h exposure to a sublethal dose of MeHg (2 μM [0.4 ppm]) and staining with both SYTO 9 (green [intact membrane]) and propidium iodide (red [permeable membrane]). Cultures were grown in triplicate, with each replicate having three technical replicates for each strain and condition. (G) Percent permeable cells, with standard deviation, for E. coli strains at the indicated MeHg exposures. Letters indicate pairwise significant differences where P is <0.05. Two-way ANOVA with Tukey’s HSD was used for all pairwise comparisons. n = 3.

To further confirm that the permeabilized cells visualized via LIVE/DEAD staining were a result of cell lysis, we performed a Western blot to probe for the presence of aminoglycoside phosphotransferase (NPT). NPT is a cytosolic enzyme that confers kanamycin resistance and is encoded on the pET9a vectors used in the construction of the strains employed in this study. We probed for the presence of NPT protein in cell-associated and supernatant fractions after exposure to 2 μM MeHg (Fig. 5A). Densitometry was utilized to determine the percentage of NPT protein in each fraction analyzed (cell associated and supernatant). Under control conditions (48 h growth in LB with 0 μM MeHg), strains expressing the empty vector (pET9a) and those expressing MerB alone showed almost all (>95%) detectable NPT protein as being cell associated, indicating that these cells remained largely intact. When exposed to 2 μM MeHg, strains expressing the empty vector (pET9a) showed that the majority of detectable NPT (∼90%) was in the cell-associated fractions (Fig. 5A and B). In contrast, 30% of the total NPT was in the supernatant fraction, as assessed by densitometry, in cultures of strains expressing MerB only when exposed to 2 μM MeHg (Table 3). These findings are indicative of NPT release from the cells to the medium as a result of cell lysis, presumably occurring due to Hg(II) accumulation due to degradation of MeHg by MerB.

FIG 5.

Western blot analysis of cellular lysis in specified E. coli strains after exposure to MeHg. (A) Immunoblot of the cytosolic NPT protein from the cell pellets and supernatants of bacterial strains. Samples were grown in medium containing either 0 or 2 μM MeHg for 48 h prior to centrifugation. NPT found in the supernatant fraction indicates cell lysis. (B) Percentage of total protein found in the supernatant fraction of each strain exposed to 0 and 2 μM MeHg for 48 h. Densitometry was performed on each band, and calculations were done using the AUC values from intensity peaks. Mean AUC values and ranges are shown. n = 2.

TABLE 3.

Densitometry analysis of NPT Western blotting for cell and supernatant fractions^a

Fraction	Construct	MeHg concn (μM)	AUC (a.u.)
			Avg	Range
Pellet	pET9a	0	6,162.7	6,137.2–6,188.2
		2	4,860.7	3,621.9–6,099.4
	RN23B	0	3,995.3	3,326.0–4,664.5
		2	3,628.9	3,021.2–4,236.5
Supernatant	pET9a	0	223.6	112.3–334.9
		2	378.4	204.0–552.8
	RN23B	0	50.3	24.1–76.4
		2	1,923.2	1,828.4–2,017.9

^aProtein content for each strain at each MeHg concentration was analyzed via Western blotting and quantified using ImageJ. The average AUC values for all replicates and ranges are reported. n = 2.

DISCUSSION

Here, using a plasmid-based E. coli expression system, we showed that MerB acting independently of MerA and other mer locus gene products can demethylate MeHg to produce Hg(II). This independent MerB activity makes cells more susceptible to MeHg toxicity, as seen by a lowering of the MeHg MIC and reduced CFU after MeHg exposure. Consistent with this reduced viability upon MeHg exposure, we saw evidence of increased MeHg-induced cell lysis in cells expressing MerB alone. As expected with a broad-spectrum mer operon, MerA and MerRPT, together with MerB, confer resistance to MeHg toxicity. Overall, our results support a model whereby MerB-mediated demethylation occurring intracellularly produces Hg(II) that can accumulate to levels that compromise the cell’s metabolism and structural integrity. This activity is averted when MerA activity is also present, presumably due to formation of volatile Hg⁰ that can exit the cell.

Our findings further the understanding of the possible role of MerB in mer operon function and evolution. A prior study using purified MerB and MerA enzymes in vitro showed that these enzymes can function in demethylation of MeHg and reduction of Hg(II), respectively, and independently of one another. However, it was seen that MerB, in the absence of MerA, had a slow turnover and low catalytic efficiency (28). This prior study thus supported the notion that MerB requires MerA to efficiently demethylate MeHg. Here, examining activity in a cellular context, we found that MeHg demethylation can occur efficiently in MerB alone strains. Nonetheless, there appear to be differences between the demethylation rates of the two MerB strains we examined (i.e., K62B versus RN23B). Curiously, this difference resolves to the same overall rate of demethylation when these two MerB strains are each accompanied by MerA. These findings indicate that MerB is catalytically efficient by itself yet is indeed influenced by coexpression with MerA. One caveat to this interpretation is that the apparent kinetics do not account for possible variation in plasmid copy number and protein expression levels in this expression system, where codon optimization was not performed. While a more quantitative kinetic assessment is warranted, it is clear that MerB expressed independently of other mer genes will result in MeHg demethylation.

From our results here, it is reasonable to predict that MerB acting alone could decrease the fitness of cells in the context of MeHg exposure. This is consistent with the conclusions of a recent phylogenetic study of mer homologs among ∼84,000 available archaeal and bacterial genomes that indicated that merB was apparently recruited to the mer operon relatively recently (27). Importantly, this prior study also revealed that as many as 50% of MerB encoding genomes lacked an accompanying MerA gene (27). This evidence of naturally occurring MerB-only organisms suggests that MerA-independent reduction mechanisms, or alternative Hg(II) sequestration mechanisms, must also exist. However, it is also possible that MerB could play an alternative role outside MeHg demethylation that may confer fitness to organisms encoding these systems in an environment free of MeHg. Future studies are warranted to identify and characterize MeHg processing and handling in native organisms that harbor MerB alone. Our results also highlight a previously unappreciated toxicity of Hg(II) relative to MeHg. It is seen that ∼80% of the MeHg added to the cultures is cell associated in empty plasmid control strains and MerA strains. Only when this MeHg is converted intracellularly to Hg(II) is enhanced toxicity seen.

It has long been understood that the toxicities associated with MeHg and Hg(II) differ quite starkly, yet this has often been attributed to the inability of Hg(II) to diffuse across membranes and reach sensitive target sites (34, 35). Our system has bypassed this initial difference in uptake kinetics between Hg species, as the Hg(II) is generated intracellularly from dosed MeHg. Previous work (33) observed an increased toxicity associated with high intracellular Hg(II) levels that were generated by upregulating Hg(II) transporters. In the context of Mer-mediated MeHg resistance, an increased toxicity associated with the formation of Hg(II) suggests that the process of demethylation is an intermediate step which must be coupled with mechanisms to further excrete, sequester, or reduce Hg(II).

We found that, in the MeHg-exposed MerB-only strain, the majority of mercury is in the Hg(II) form and is also found predominantly in the cell-free supernatant. Given the high affinity of Hg(II) for protein thiols in the intracellular space (36, 37), the finding that the majority of Hg(II) is in the extracellular environment suggests that it is likely thiol associated and is indicative of substantial cell lysis. This interpretation is supported by our LIVE/DEAD staining and immunoblotting for the cytosolic NPT protein. However, another possible explanation for this could be that transporters for Hg(II), but not MeHg, are present in our E. coli hosts. While transport was not directly assessed in our current study, previous work suggests that in our experimental design, the loss of Hg(II) to the extracellular environment is unlikely to be due to passive diffusion or active transport of Hg(II) out of the cell (36–38). Multiple pathways can lead to cell lysis (39), and it is plausible that Hg(II) and MeHg can exert differential effects on the physiology of the cells that are in turn required to maintain membrane integrity.

Our study has some limitations with respect to the system we employed; namely, the methods rely on a plasmid-based overexpression system. First, expression of MerA and MerB relies on exogenous promoters and is likely far more efficient at regulating expression and in excess levels than what would be found in naturally occurring mer-harboring organisms. Overexpression may lead to observed enzyme kinetics that are not necessarily what one would observe in wild-type strains. In addition, comparative assessment of the different constructs, such as the variability in rates of demethylation between RN23B and K62B strains, may be subject to variability in copy number of each plasmid, which in turn, could greatly affect protein expression levels. There is also a possibility of nonoptimal codon utilization associated with homologs, relative to the E. coli host, which may result in unequal expression between the two strains of MerB. Additional assays are warranted to assess expression level differences that may exist between these strains containing codon-optimized expression constructs to more rigorously evaluate their kinetic properties. Further research is also needed to evaluate our findings in the context of microorganisms that naturally encode MerB but not MerA, as it is possible that other abiotic/biotic mechanisms for sequestering or reducing Hg(II) may exist in these cells (as outlined by Christakis et al. [27]). For example, thiol-containing reservoirs may serve to complex with Hg(II) in cells that encode MerB but not MerA, and it is possible that divalent transporters, not present in the E. coli strains utilized herein, may exist that aid the rapid excretion of Hg(II) (13).

In conclusion, we demonstrated MerB-mediated MeHg demethylation in the absence of MerA in a cellular context. Without a dedicated mechanism to reduce Hg(II), MeHg exposure was shown to decrease E. coli viability and increase cell lysis, presumably triggered by intracellular production of more toxic Hg(II) compared to MeHg. Our work also adds to previous evidence of different toxicity profiles for Hg(II) and MeHg. Finally, our findings highlight the need for future studies evaluating naturally occurring bacterial strains that harbor MerB alone for the potential existence of a mer-independent Hg(II) reduction/sequestration mechanism.

MATERIALS AND METHODS

Bacterial strains.

Bacterial strains containing selected genes of the mer operon on plasmids were created via manipulation of E. coli-compatible plasmid constructs originally created in the laboratory of Ginro Endo, as reported previously (40). All strains were created by transformation of plasmids into chemically competent JM109 E. coli cells (Promega competent cells). The coding regions from DNA of merRETP and merA from Bacillus cereus RC607 were assembled in a pHY300PLK vector backbone (ampicillin resistance), and the construct was designated pHYR1a. The coding regions of DNA from merB derived from either Pseudomonas sp. strain K-62 or Staphylococcus aureus RN23 were assembled on a pET9a vector backbone (kanamycin resistance) and designated K62B and RN23B, respectively. An empty pET9a plasmid was transformed into E. coli JM109 to be used as an empty vector control. Plasmid compatibility, and expression in E. coli, was confirmed previously by Chien et al. (40). All mer genes on the pHYR1a plasmid were under the control of merR1 from Bacillus cereus RC607, and merB genes on the pET9a plasmid were under the control of a heterologous T7 promoter, inducible by isopropyl-β-d-thiogalactopyranoside (IPTG). The various expression constructs are summarized in Table 1. The mer gene sequences in plasmid DNA from newly derived transformants were sequenced and PCR verified (ACGT, Inc.). Frozen glycerol stocks of each bacterial strain were created and utilized to streak agar plates and obtain single bacterial colonies for experimentation.

Mercury analysis.

Total mercury [T-Hg; MeHg + Hg(II)] was assessed through thermal decomposition and atomic absorption using a DMA-80 direct mercury analyzer (Milestone SRL) and analyzing 100 μL of liquid bacterial cultures. Values were determined on a wet-weight basis and expressed in parts per million (micrograms per gram). Inorganic mercury and methylmercury [Hg(II) and MeHg, respectively] were assessed via analysis of samples using liquid chromatography coupled with inductively coupled plasma mass spectroscopy (LC-ICP-MS). Culture samples of 200 μL were diluted 5-fold in a digestion buffer solution containing 3.25 mM glutathione (GSH), 32.3 mM HCl, and 20 mM KCl and mixed overnight on a platform shaker at room temperature. Digested samples were subsequently filtered through a 0.45-μm cellulose acetate filter, and 50 μL of filtrate was subjected to analysis via LC-ICP-MS using a Perkin Elmer NexION 2000 ICP-MS with a 150-mm Agilent Eclipse XBD C₁₈ high-performance LC (HPLC) column and a pore size of 5 μm. The mobile phase used for all experiments was 0.1% cysteine and 0.1% HCl run isocratically with a flow rate of 1 mL/min. Instrument calibrations were performed prior to analysis using standards of both MeHg and Hg(II) at 500, 250, 125, 62.50, 31.25, 15.62, 7.81, and 3.90 ng/mL. Empower software (Perkin Elmer) was used to generate the LC output, whereby peak volumes generated from all samples were compared against those of the calibration curve to determine Hg concentration. Instrument conditions for all runs were 1,400 W power in standard mode, measuring the Hg²⁰² isotope. The limit of detection (LOD) for both MeHg and Hg(II) in LC ICP-MS experiments was 0.1 ng/mL (ppb), and the limit of quantification (LOQ) is 10× the LOD, 1 ng/mL (ppb).

MeHg demethylation and reduction assays in vitro.

Fresh Difco LB agar (Lennox; BD 240110) plates with the proper antibiotics (abx) for each respective plasmid (100 μg/mL ampicillin, 50 μg/mL kanamycin) were streaked from frozen stocks, and single colonies were chosen for each replicate of this assay. Initially, single colonies were placed in 2 mL of Difco LB broth (Lennox; BD, Franklin Lakes, NJ) + abx and grown overnight in a 14-mL round-bottom polypropylene tube (Falcon) 17 mm wide and 100 mm long. The culture tubes were shaken in a tube rack at an angle to allow increased surface area at 37°C with 200 rpm shaking. Saturated overnight cultures of each bacterial strain were diluted 1:10 into fresh LB broth + abx and 1 mM IPTG to induce the pET9a plasmid constructs. These cultures were grown for 3 h at 37°C and subsequently diluted to an OD₆₀₀ of 0.3. A sample of 270 μL of these diluted cultures was then inoculated into 2.7 mL of fresh LB broth + abx in 14-mL round-bottom polypropylene tubes containing a sublethal dose of MeHg (either 0.1 ppm or 0.2 ppm). Cultures were then incubated at 37°C with 200 rpm shaking for 24 or 48 h. At various time points, subsamples of 300 μL were collected from each liquid culture and assessed for T-Hg, Hg(II), and MeHg as described above. Demethylation was considered to occur if there was no change in T-Hg accompanied by an increase in Hg(II) and a decrease in MeHg over time. Hg(II) reduction was considered to occur if there was a decrease T-Hg over time due to volatilization to Hg⁰. All samples were prepared and grown under the same conditions to ensure that no abiotic or background demethylation occurred due to the experimental setup.

Growth and determination of MIC for MeHg.

The MIC of MeHg was assessed for each of the bacterial strains following a reported method with slight modifications (41). A series of growth assays were run prior to determining MICs to assess growth in the absence of MeHg and to determine the time point at which to assess the MIC. For growth assays, 200 μL of a saturated overnight culture from each strain was inoculated into 2.0 mL fresh LB + abx and a range of MeHg concentrations from 0 to 10 μM. The OD₆₀₀ of these cultures was assessed every 4 h for the first 24 h and then every 8 h until 48 h of total growth time. Under all growth conditions, culture densities reached a maximum value prior to 48 h; therefore, the MICs were determined at the 48-h time point (Fig. S1). From a saturated overnight culture, a new culture was established by diluting to an OD₆₀₀ of 0.20 using fresh LB broth + abx. A subsample of 270 μL of each diluted culture was then inoculated into 2.7 mL of LB broth + abx and supplemented with MeHg at a concentration ranging from 0 to 20 μM (MIC). The OD₆₀₀ of each culture was assessed immediately after inoculation to obtain a baseline cell density, and then cultures were incubated at 37°C with 200 rpm shaking for 48 h. For MIC analysis, a growth/no-growth determination was assessed at 48 h by measuring the OD₆₀₀ of each bacterial strain at each MeHg concentration. Growth at a given concentration of MeHg occurred if the OD₆₀₀ reading at 48 h was above the baseline (0-h) OD₆₀₀. The MIC of MeHg was the lowest concentration of MeHg at which no growth occurred in 48 h.

Cellular viability and lysis.

The viability of bacterial cultures exposed to MeHg was assessed using two methods. First, a CFU assay was utilized to assess the viability of bacterial cells following exposure to a sublethal dose of MeHg, as determined from MIC assays. Samples of 270 μL of overnight cultures prepared as previously described were inoculated into 2.7 mL of LB broth + abx containing 0.4 ppm MeHg. Cultures were allowed to grow for 48 h at 37°C with 200 rpm shaking. After 48 h of exposure, samples were diluted back to an equal OD₆₀₀ of 0.3, and then cultures were further diluted 10⁴ or 10⁵. A sample of 40 μL of each diluted culture was plated onto LB agar plates containing the appropriate abx for each respective plasmid, and plates were incubated for 24 h at 37°C. The number of colonies formed after 24 h of incubation was recorded and compared between the different strains. The number of CFU/mL was determined for each condition as follows: (number of colonies × dilution factor)/volume (ml) of dilute sample plated.

Second, a LIVE/DEAD BacLight bacterial viability kit (Invitrogen, Waltham, MA) was used to assess the membrane integrity of cells after exposure to a sublethal dose of MeHg. Cell cultures were set up in the same manner as was done for the CFU assay; however, after 48 h exposure to MeHg and dilution, subsamples of cultures were stained with both SYTO 9 dye and propidium iodide according to the manufacturers’ protocols. The SYTO 9 dye stains all cells in the culture with a green-fluorescent nucleic acid stain, regardless of membrane integrity. Propidium iodide stains only cells that have permeable membranes, with a red-fluorescent nucleic acid stain; stained cells are considered “dead” cells. Cells were imaged using a Nikon AZ100 wide-field epifluorescent zoom microscope with consistent exposure settings across all samples. For each stain at each MeHg concentration, three biological replicates were imaged, and three separate areas along the microscope slide were analyzed for each biological replicate to obtain counts of on average ∼750 cells. Images were quantified separately, and the averages were used for each replicate. Images of each culture were obtained on both a TRITC (tetramethyl rhodamine isocyanate) and FITC (fluorescein isothiocyanate) channel. Images were then analyzed via ImageJ (NIH) software. Viability was determined as a percentage of the total cell population that stained green (viable cells).

Distribution of mercury isoforms in bacterial cultures.

To determine the location of Hg in the bacterial cultures, a 500-μL subsample of each culture was subjected to centrifugation to separate cell-associated and extracellular (supernatant) content. Briefly, samples were centrifuged in 1.5-mL tubes at 5,000 × g for 15 min at 10°C. Supernatant was carefully removed from the cell pellets and placed in a separate 1.5-mL tube. The remaining cell pellets were resuspended in 500 μL of LB broth. Immediately following separation and resuspension, cell and supernatant fractions were subject to T-Hg analysis as described above. The amount of T-Hg in each fraction is reported in ppm and as a percentage of the combined T-Hg in the cells and supernatant. A subsample of each respective strain that was not centrifuged and separated was analyzed for T-Hg and individual mercury species to control for loss of Hg during the separation process and determine the fraction of the total Hg represented by Hg(II) and MeHg in each sample.

Immunoblotting and protein quantification.

Aminoglycoside phosphotransferase (NPT) is a cytosolic protein that confers kanamycin resistance when expressed via the pET9a vector. Detection of NPT in both cellular and supernatant fractions was therefore utilized as a measure of the release of cytoplasmic content due to membrane lysis in each sample. Cell and supernatant were fractionated by centrifugation described above, and the protein fraction was recovered via shearing samples in lysis buffer using a microcentrifuge tube mortar and pestle (Kontes, Inc.), followed by boiling samples in SDS-PAGE buffer. Immunoblotting for NPT was done using a modified protocol from Sanders and Pavelka (42). Briefly, equal volumes of cell supernatant and cell lysate were separated on a 12% SDS-PAGE gel at 120 V for 90 min. Proteins were subsequently electrotransferred to a polyvinylidene difluoride (PVDF) membrane (Millipore Sigma, Burlington, MA) overnight at 4°C at 36 V. Membranes were probed with rabbit IgG anti-NPT (MyBioSource, Vancouver, BC, Canada) and then incubated with goat anti-rabbit IgG conjugated to horseradish peroxidase. The presence of NPT was detected by chemiluminescence using Clarity Western enhanced chemiluminescence (ECL) assay (Bio-Rad, Hercules, CA) and imaging the gel according to the manufacturer’s recommendations with a ChemiDoc MP imaging system (Bio-Rad, Hercules, CA). Densitometry analysis was done in order to quantify the ratio of total protein found in the cell and supernatant fractions. ImageJ (NIH) software was utilized to quantify the area under the curve (AUC) for each NPT protein band imaged. Total AUC values are reported in Table 3. Using the AUC values for each protein band, a measure of the amount of NPT protein located in the supernatant fraction was calculated by dividing the AUC for just the supernatant band by the total AUC for both bands. This value was then reported as the percentage of NPT located in the supernatant fraction as a proxy measure for the amount of cell lysis in each strain under each condition.

Statistical analysis.

All statistical analyses were completed in JMP Pro version 15.0.0 (SAS Institute, Cary, NC). Comparisons between treatment groups or experimental bacterial strains were evaluated in relation to the relevant untreated group or empty-vector controls, as indicated. For the evaluation of demethylation and reduction, a two-way ANOVA with Tukey’s honestly significant difference (HSD) was used to compare the average concentrations of T-Hg and methylmercury between bacterial strains and over time. To assess differences in MICs between strains, a one-way ANOVA with Tukey’s post hoc test was used to compare the average MIC calculated from three independent biological replicates for each construct. A two-way ANOVA with Tukey’s HSD was used to compare cellular viability by comparing either the average CFU/mL recovered from each culture strain or the average percentage of “live” positive cells in each culture strain at each concentration of MeHg (0 and 2 μM) from three biological replicates. To assess differences in the location of T-Hg between bacterial constructs, a one-way ANOVA was used to compare the mean percentage of T-Hg found in the extracellular environment after exposure to 2 μM MeHg between all groups. Data are presented as means and standard deviations. In all cases, significance was defined as a P value of ≤0.05. All experiments presented herein include data from at least three independent biological replicates.