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Mar 29, 2017 - ... Engineering, Northwestern University, 2145 Sheridan Road, Evanston, Illinois 60208, United ... To gain insight into this Hg(II) bio...
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Cysteine Addition Promotes Sulfide Production and 4‑Fold Hg(II)−S Coordination in Actively Metabolizing Escherichia coli Sara A. Thomas and Jean-François Gaillard* Department of Civil and Environmental Engineering, Northwestern University, 2145 Sheridan Road, Evanston, Illinois 60208, United States S Supporting Information *

ABSTRACT: The bacterial uptake of mercury(II), Hg(II), is believed to be energy-dependent and is enhanced by cysteine in diverse species of bacteria under aerobic and anaerobic conditions. To gain insight into this Hg(II) biouptake pathway, we have employed X-ray absorption spectroscopy (XAS) to investigate the relationship between exogenous cysteine, cellular metabolism, cellular localization, and Hg(II) coordination in aerobically respiring Escherichia coli (E. coli). We show that cells harvested in exponential growth phase consistently display mixtures of 2-fold and 4-fold Hg(II) coordination to sulfur (Hg−S2 and Hg−S4), with added cysteine enhancing Hg−S4 formation. In contrast, cells in stationary growth phase or cells treated with a protonophore causing a decrease in cellular ATP predominantly contain Hg−S2, regardless of cysteine addition. Our XAS results favor metacinnabar (βHgS) as the Hg−S4 species, which we show is associated with both the cell envelope and cytoplasm. Additionally, we observe that added cysteine abiotically oxidizes to cystine and exponentially growing E. coli degrade high cysteine concentrations (100−1000 μM) into sulfide. Thermodynamic calculations confirm that cysteine-induced sulfide biosynthesis can promote the formation of dissolved and particulate Hg(II)-sulfide species. This report reveals new complexities arising in Hg(II) bioassays with cysteine and emphasizes the need for considering changes in chemical speciation as well as growth stage.

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INTRODUCTION Human activities have altered the geochemical cycles of many trace metals, with deleterious consequences for human and ecosystem health. Mercury(II), Hg(II), contamination of natural waters occurs on a global scale, mainly as a result of atmospheric deposition from coal combustion.1 In anoxic sediments1,2 and oxic regions of the water column,2−5 microbes transform Hg(II) into monomethylmercury (MeHg), a potent neurotoxin that biomagnifies in aquatic food webs.6,7 Certain anaerobic bacteria containing the hgcAB gene cluster (e.g., sulfate- and iron-reducers) are responsible for MeHg production,8,9 which is believed to be an intracellular process.9,10 Some bacteria have known Hg(II)-specific transport proteins as part of the mer operon (e.g., MerT and MerC) whose role is to internalize Hg(II) so that MerA, mercuric reductase, will reduce Hg(II) to less toxic Hg(0).11 However, known Hg(II)-methylating bacteria lack the mer operon,10 revealing an unknown Hg(II) uptake pathway that governs Hg(II) methylation. Understanding the factors that influence Hg(II) biouptake in bacteria lacking the mer operon will improve our capacity to predict Hg(II) bioavailability and the potential for MeHg production. The passive diffusion of small, neutrally charged Hg(II) complexes (e.g., Hg(II)-sulfides) was thought to be the universal route of entry through bacteria’s hydrophobic membranes.12−15 However, it was recently reported that © 2017 American Chemical Society

Hg(II) uptake by various aerobic and anaerobic bacterial species is hindered in the presence of carbonylcyanide-mchlorophenylhydrazone (CCCP), a protonophore, suggesting conversely that Hg(II) biouptake is energy-dependent (i.e., potentially occurs by active transport).16,17 The bacterial uptake of Hg(II) can also be repressed in the presence of Zn(II) and Cd(II),17,18 two metals with similar outer electronic configuration as Hg(II). Studies have proposed that Zn(II) and Cd(II) may outcompete Hg(II) for the binding site of an uptake protein, hence inhibiting Hg(II) biouptake, and further supporting the existence of an unknown protein that can transport Hg(II).17,18 Hg(II) has a high affinity for reduced sulfur (e.g., thiols and sulfides), and it is predicted that Hg(II) is predominantly bound to reduced sulfur in the environments where MeHg is produced.15,19,20 Bacterial cell envelopes also contain thiols (∼10% of total functional groups)21 capable of binding Hg(II).22−25 If Hg(II) is indeed internalized by a metal transport protein, then Hg(II) likely interacts with thiol functional groups in the cell envelope prior to uptake. In our previous work, we have determined that when Hg(II) is Received: Revised: Accepted: Published: 4642

December March 15, March 24, March 29,

16, 2016 2017 2017 2017 DOI: 10.1021/acs.est.6b06400 Environ. Sci. Technol. 2017, 51, 4642−4651

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Environmental Science & Technology

(OD600 = 0.2) or 5−10 h after reaching stationary growth phase (OD600 ≈ 0.3; see Figure S1 for growth curve). Cells were washed twice with minimal complexing medium (MCM)the exposure medium for Hg(II) biouptake assaysand resuspended to a density of ∼2 × 108 cells/mL, which is equivalent to an OD600 of 0.2 (Figure S3). MCM (pH = 7.1) consists of 20 mM MOPS buffer, 1 mM Na-β-glycerophosphate, 0.41 mM MgSO4, 12 mM NH4NO3, 0.76 mM isoleucine, 0.76 mM leucine, 3 nM thiamine, 10 mM glucose, and 9.1 mM NaOH.26 To further limit metabolic activity, cells grown to stationary phase were suspended in MCM without the glucose carbon source: MCM (−glucose). Hg(II) and Cysteine Exposure Assays. A 10 mM Hg(NO3)2 stock solution in 1% HNO3 (TMG) was used for all exposure assays and stored at 4 °C. Cysteine stock solutions were prepared in Milli-Q immediately before use. Hg(II) and cysteine were pre-equilibrated at 10 times the final desired concentrations for 1 h in Milli-Q prior to being added to cell suspensions. Hg(II) and cysteine exposure assays were aerobic and conducted in 15 mL loosely capped borosilicate glass vials under dark conditions following previously reported methods.17 Assays were initiated with the addition of 0.7 mL of Hg(II) solution (±cysteine) to 6.3 mL of cell suspension in MCM and mixed for 3 h. Cysteine concentrations of 1, 10, 100, and 1000 μM were tested. To determine the effect of the protonophore CCCP, cells grown to exponential phase in MSM and transferred to MCM were incubated with 50 μM CCCP (from a 50 mM CCCP stock in 99.5% ethanol) for 1 h prior to Hg(II) ± cysteine solution addition. The ethanol alone had no effect on our measurements herein (data not shown). Monitoring of Cysteine Degradation and Oxidation and Biogenic Sulfide Production. During Hg(II) and cysteine exposure assays, we monitored reduced cysteine, oxidized cysteine (cystine), and sulfide in the exposure medium. An adapted version of the Cline method40,41 and the Gaitonde method42 were used to quantify sulfide (detection limit ∼2 μM) and cysteine (detection limit ∼5 μM), respectively. Unfortunately, the presence of CCCP interferes with these two colorimetric methods. Detailed protocols are included in SI Part 2. Quantification of Cellular Hg(II) Localization. After exposure of E. coli cells to Hg(II) ± cysteine for 3 h, a 1 mL aliquot of cell suspension was collected and preserved in ∼1% HCl (TMG) for determination of total recoverable Hg. In addition, a 1 mL aliquot was passed through a 0.2 μm nylon filter (VWR International), and the filtrate was preserved in ∼1% HCl (TMG) for the determination of dissolved Hg. The nylon filters do not bind a significant amount of Hg (data not shown). Around 100−200 μL of acidified solution were added to quartz sample boats for analysis by a Direct Mercury Analyzer (DMA-80, Milestone), which involves thermal decomposition followed by gold amalgamation and detection with atomic absorption spectroscopy.43 To quantify intracellular Hg(II), we adapted a wash method from Schaefer et al.,16,27 which was altered to improve Hg(II) removal from the cell membrane without compromising cytoplasmic membrane integrity. Briefly, after Hg(II) exposure, 4 mL of the cell suspension was centrifuged (7000g for 10 min) and resuspended in 2 mL of 50 mM EDTA and 100 mM oxalate solution (pH = 7.5). After mixing for 10 min, 2 mL of a 10 mM GSH and 3 mM ascorbate solution (pH = 7) were added and mixed for an additional 10 min. Subsequently, a 2 mL aliquot of cells suspended in the EDTA/GSH solution were

quantitatively complexed by EDTA outside the cell, Hg(II) remains bioavailable and undergoes a ligand exchange reaction with thiol moieties in the cell envelope of Escherichia coli (E. coli).25,26 However, it remains unclear if the same would happen when Hg(II) is bound to an extracellular thiolcontaining ligand. Extracellular thiol-containing ligands (e.g., cysteine, glutathione, and dissolved organic matter) will either greatly enhance or hinder the bacterial biouptake of Hg(II) depending on the ligand concentration, cell physiology, and exposure time.16,17,25,27−31 As a result, many recent studies have chosen cysteine as a model ligand in Hg(II) biouptake and methylation assays.18,31−34 Although an essential amino acid, cysteine is strictly regulated in cells due to its toxicity above a certain threshold concentration.35 To modulate cysteine concentrations, bacteria activate cysteine desulfhydrase enzymes which degrade cysteine into sulfide, ammonia, and pyruvate.36−38 However, many Hg(II) biouptake studies involving cysteine do not test for the biodegradation of cysteine to sulfide, which would impact Hg(II) speciation in the exposure medium. Determining how cysteine and its possible degradation products affect the Hg(II) binding environment in bacteria should provide valuable insights into the Hg(II) biouptake mechanism. The objective of this study is to test the hypothesis that the coordination environment of Hg(II) associated with bacterial cells varies with cellular metabolism and Hg(II) localization (i.e., cell envelope or cytoplasm). We exposed E. coli cells with varying levels of metabolic activity to cysteine and low Hg concentrations (50 and 500 nM) under aerobic conditions and characterized cellular Hg(II) localization with a chemical wash method. In parallel, we have determined the coordination environment of Hg(II) in E. coli under the same experimental conditions with Hg LIII-edge X-ray absorption spectroscopy (XAS). A wild-type strain of E. coli without the mer operon or hgcAB gene cluster was chosen as the model organism.39 We also monitored cysteine oxidation and biodegradation to sulfide during these exposure assays to follow changes in extracellular Hg(II) speciation.

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MATERIALS AND METHODS Model Organism. Escherichia coli (E. coli) ATCC 25922 was regenerated from frozen glycerol stock (−80 °C) onto LB agar, Miller (EMD Millipore) plates incubated at 37 °C for 24 h. The plates were stored at 4 °C for no more than 4 weeks. Chemical Reagents and Glassware. The chemical reagents used in this study were ACS grade (≥98%) and obtained from Sigma-Aldrich. The Hg(NO3)2·H2O was trace metal grade (TMG; ≥ 99.99%). L-Cysteine and reduced Lglutathione (GSH) powder were stored at 4 °C and used within 1 year and 6 months of purchase date, respectively. All glassware for Hg(II) biouptake assays was washed with 10% HNO3 and thoroughly rinsed with Milli-Q water. Growth Media and Cell Harvesting. A single colony of E. coli from a refrigerated LB agar plate was inoculated into 50 mL of LB broth, Miller (EMD Millipore) in a sterile, foil-topped 125 mL Erlenmeyer flask and incubated aerobically overnight at 37 °C with medium shaking until exponential phase (OD600 = 0.3−0.4). Subsequently, 20−40 μL of the cell suspension was inoculated into 50 mL of minimal salts medium (MSM; see Table S1 in the Supporting Information, SI) in a sterile, foiltopped 125 mL Erlenmeyer flask and shaken at 37 °C. Cells were either harvested in MSM during exponential growth phase 4643

DOI: 10.1021/acs.est.6b06400 Environ. Sci. Technol. 2017, 51, 4642−4651

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Environmental Science & Technology filtered onto 0.2 μm cellulose nitrate filters, and the whole filters containing cells were analyzed for total Hg content (i.e., intracellular Hg) with a DMA-80. The intracellular Hg(II) concentration was normalized considering the cell density (OD600) of each sample, measured before the wash (in MCM) and after the wash (in EDTA/GSH). The OD600 is linearly related to the cell density (#cells/mL) in all matrices and cell concentrations in this study (Figure S3). We also confirmed that the wash does not compromise cytoplasmic membrane integrity with the LIVE/DEAD BacLight Bacterial Viability Kit (SI Part 4). Hg(II) in the cell envelope is defined as the difference between the total recoverable Hg(II) and the intracellular + dissolved Hg(II). ATP Measurement. Cellular ATP levels before and after a 3-h exposure to Hg(II) and cysteine were measured to assess metabolic activity using the CellTiter-Glo 2.0 Assay (Promega). The ATP assays were conducted in white, polystyrene 96-well plates, where the total assay volume in each well was 100 μL. After mixing cells with Hg(II) ± cysteine for 3 h, 100 μL of room temperature CellTiter-Glo 2.0 reagent was added to each well. After shaking for 15 min, the luminescence intensity in each well was recorded with an Flx 800 Microplate Reader (Biotek). The luminescence value was subtracted from that of the blank (100 μL matrix with no cells and 100 μL CellTiterGlo 2.0 reagent). A standard curve with known concentrations of ATP disodium salt (Sigma-Aldrich) prepared in the same matrix as cell suspensions was concurrently measured on each well plate so that cellular ATP could be quantified. XAS Preparation, Data Collection, and Data Analysis for Hg(II) Standards and Samples. We probed the average coordination environment of Hg(II) associated with whole cells of E. coli with XAS to determine the effect of cell metabolism and cysteine addition. The same approach as described above for exposing cells to Hg(II) and cysteine was followed except that the total assay volume was scaled up to 400 mL to increase the amount of biomass collected. After a 3-h exposure period, cells were washed twice with 0.1 M NaClO4 (7500 g, 10 min), collected as a cell pellet onto 2.5 cm diameter cellulose nitrate filters (0.4 μm pore size), and vacuum filtered to remove excess moisture for 5−10 min. NaClO4 was chosen as the washing agent to remove dissolved Hg(II) that was associated with the cell pellet but not disturb the Hg(II) bound to the cell envelope.22−24 Subsequently, the filters supporting the cells were sandwiched between two pieces of Kapton tape (DuPont). Cells harvested in exponential growth phase were immediately plunged in LN2 to halt cell metabolism while preserving Hg(II) coordination environment and were stored in a −80 °C freezer for no more than 1 week prior to XAS analysis. Since cells in stationary phase were not actively metabolizing, those cell pellets were not flash frozen in LN2 and instead stored at 4 °C for no more than 48 h. We show that the process of flash freezing does not alter Hg(II) coordination in bacteria (Figure S11). Hg LIII-edge XAS spectra were collected on the DuPontNorthwestern-Dow Collaborative Access Team (DND-CAT) bending magnet beamline located at Sector 5 of the Advanced Photon Source. A detailed description of the reference standard preparation, scan parameters, beamline specifications, handling of bacterial samples, and reference standard analysis is included in SI Parts 7, 8, and 9. Principal component analysis (PCA) and target transformation (TT) were performed on the sample spectra to determine the appropriate Hg reference standards to be used in the analysis (SI Part 10). We then determined the

relative fractions of the Hg(II) species associated with the bacterial cells by performing linear combination fits (LCFs) of the extended fine structure (EXAFS) using the reference standards identified by TT (SI Part 11). In addition, we incorporate the near edge structure (XANES) to provide qualitative information on the Hg(II) binding environment (SI Part 11). Thermodynamic Modeling. All Hg(II) speciation calculations were performed with the program ChemEQL.44 The equilibrium constants used in the calculations are reported in Table S4.

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RESULTS Exogenous Cysteine/Cystine Promotes Sulfide Production by Exponential Phase Cells. When E. coli cells in exponential and stationary phase were exposed to 100 and 1000 μM cysteine during aerobic Hg(II) bioassays, the concentration of cysteine outside the cell changed during the 3-h exposure period due to oxidation to cystine, cellular uptake, and degradation (Figure 1). On the addition of cysteine to the

Figure 1. Concentration of reduced and reduced + oxidized cysteine (cystine) detected at time = 0 and 3 h during exposure of exponential and stationary phase E. coli to (A) 100 μM and (B) 1000 μM cysteine. Results of a Tukey’s honest significant differences test are provided separately for subfigure A and B. No reduced cysteine is detected at t = 0 or 3 h when the initial cysteine concentration is 100 μM. Colorimetric interference prevented the measurement of cysteine in CCCP-treated samples. The bars are averages from 3 independent experiments and the error bars are ±1 SD.

cell suspension in MCM (t = 0 h), added cysteine concentrations as high as 100 μM were entirely oxidized (Figure 1A), regardless of growth phase. We determined that cysteine oxidation in MCM occurs in the absence of cells and is not solely caused by dissolved oxygen since 100 μM cysteine remains fully reduced in aerobic Milli-Q (Figure S2). When 1000 μM cysteine is initially added to cell suspensions, around 800 μM of cysteine is reduced at t = 0 h, and this does not significantly change over time for cells in both exponential and stationary growth phase (Figure 1B). We also tested for the degradation of cysteine to sulfide in exposure assays so that Hg(II) speciation could be properly considered. In assays containing exponential phase cells, extracellular sulfide was detected at concentrations of ∼10 μM and ∼30 μM after a 3-h exposure to 100 μM and 1000 μM cysteine, respectively (Figure 2A). Since we measure that 100 μM cysteine is fully oxidized in MCM, it appears that oxidized cysteine (cystine) may be a sulfide source. Extracellular sulfide was not detected for exponential phase cells exposed to 0−10 4644

DOI: 10.1021/acs.est.6b06400 Environ. Sci. Technol. 2017, 51, 4642−4651

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the presence of cysteine or CCCP alone had no effect on the quantification of ATP (data not shown). Potentially, high cysteine concentrations (>100 μM) either rendered CCCP ineffective in disrupting the PMF or promoted an alternative pathway for ATP synthesis. Regardless of the reason, we note that CCCP-treated samples exposed to 100 μM and 1000 μM cysteine were not affected by CCCP treatment as intended. Hg(II) Recovery and Cellular Localization Depend on Cellular Metabolism and Cysteine Addition. When cells are exposed to 50 nM Hg(II) and 0−10 μM cysteine in their exponential growth phase, the cell-associated Hg(II) is predominantly localized in the cell envelope after a 3-h exposure period (Figure 4A). As the cysteine concentration is increased to 100 and 1000 μM, however, the cell-associated Hg(II) becomes predominantly intracellular. Thus, cysteine addition above a threshold concentration promotes Hg(II) transfer through the cytoplasmic membrane, as observed in other bacterial species.16,17,27 Dissolved Hg(II) is very low (∼2% total recoverable Hg(II)) in the exponential growth phase sample with 50 nM Hg(II) and 100 μM added cysteine (Figure 4A). However, a higher added cysteine concentration of 1000 μM in the presence of 50 nM Hg(II) increases the dissolved fraction to 40% of the total recoverable Hg. The dissolved fraction of Hg is even greater for stationary phase cells exposed to 50 nM Hg and 1000 μM cysteine (∼80% of the total recoverable Hg; Figure 4B). Thus, the presence of a high excess of cysteine in its reduced form appears to promote dissolved Hg(II). When exponential phase cells are exposed to 500 nM Hg(II), with and without 1000 μM added cysteine, only ∼30% of the cell-associated Hg(II) is intracellular (Figure 4A), suggesting that Hg(II) internalization is limited at higher Hg(II) concentrations, even in the presence of cysteine. A comparison of the Hg(II) localization results between exponential growth phase cells that were and were not exposed to 50 μM CCCP (Figure 4A and Figure 4C) shows that the presence of CCCP does not affect the concentration of intracellular Hg detected. For the samples exposed to 100− 1000 μM cysteine, this may be related to the inability of CCCP to decrease cellular ATP. However, in stationary phase cells, the concentration of intracellular Hg is significantly lower for samples exposed to 500 nM total Hg (Figure 4B).

Figure 2. Concentration of sulfide in the exposure medium detected in 3-h assays of E. coli exposed to 0−1000 μM cysteine harvested in (A) exponential and (B) stationary growth phase. The presence of 50 nM or 500 nM total Hg(II) had no effect on the production of sulfide (Table S2). Colorimetric interference prevented the measurement of sulfide in CCCP-treated samples. Data points marked with an asterisk are statistically different from the control with no cysteine (p < 0.05). The bars are averages from 3 independent experiments and the error bars are ±1 SD.

μM cysteine or stationary phase cells exposed to 0−1000 μM cysteine (Figure 2). ATP in Exponential Phase, Stationary Phase, and CCCP-Treated Cells. Cells enter stationary growth phase to conserve energy in response to limited nutrient availability.45 We thus observe that cells harvested in stationary growth phase contain around 5 times less ATP per cell than cells harvested in exponential growth phase, regardless of Hg(II) or cysteine presence (Figure 3). Likewise, CCCP can disrupt the proton motive force (PMF), reducing ATP synthesis in aerobically and anaerobically respiring cells.46 Prior to exposure to Hg(II) and cysteine, incubating exponential phase cells with 50 μM CCCP for 1 h decreases the cellular ATP concentration from ∼1.5 to 1.0 × 10−18 mol/cell (compare dotted black lines in Figure 3A,C). A 3-h exposure of the CCCP-treated cells to 50 nM Hg(II) with 0−10 μM cysteine as well as 500 nM Hg(II) decreased ATP further to ∼0.6 × 10−18 mol/cell (Figure 3C). Surprisingly, a 3-h exposure of the CCCP-treated cells to Hg with 100 μM and 1000 μM added cysteine allowed ATP levels to increase back to ∼1.5 × 10−18 mol/cell. We determined that

Figure 3. Amount of ATP per cell measured after exposing E. coli to Hg (50 nM and 500 nM) and cysteine for 3 h. Prior to Hg(II) exposure, cells were harvested in their (A) exponential growth phase and suspended in MCM, (B) stationary growth phase and suspended in MCM with no glucose, and (C) exponential growth phase and suspended in MCM with 50 μM CCCP. Cells were exposed to CCCP for 1 h before addition of Hg(II). The dotted black line represents the ATP concentration in an aliquot from the same sample of cells directly before Hg and cysteine addition. The bars are averages of duplicates from 2 to 3 independent experiments and the error bars are ±1 SD. 4645

DOI: 10.1021/acs.est.6b06400 Environ. Sci. Technol. 2017, 51, 4642−4651

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Figure 4. Fraction of total added Hg (50 nM Hg or 500 nM Hg) that was dissolved, in the cell envelope, and in the cytoplasm (intracellular) after exposing E. coli to Hg and cysteine for 3 h. Prior to Hg(II) exposure, cells were harvested in their (A) exponential growth phase and suspended in MCM, (B) stationary growth phase and suspended in MCM with no glucose, and (C) exponential growth phase and suspended in MCM with 50 μM CCCP. Cells were mixed with CCCP for 1 h before addition of Hg(II). The bars represent averages of at least 3 independent experiments and the error bars are ±1 SD. A statistical analysis of the results is included in Table S2.

Figure 5. Hg LIII-edge k2-weighted EXAFS spectra of Hg(II) in bacterial cells (intracellular + cell envelope) and best-fit curves for 3 models. Prior to Hg(II) and cysteine exposure, cells were harvested in their (A) exponential growth phase and suspended in MCM, (B) stationary growth phase and suspended in MCM with no glucose, and (C) exponential growth phase and suspended in MCM with 50 μM CCCP. Cells were mixed with CCCP for 1 h before addition of Hg(II).

biouptake,27,31 which has been attributed to the formation of a nonbioavailable Hg(cysteine)3 species.27 However, cysteine addition may also induce HgS(s) formation,30 as the Ksp for HgS(s) precipitation is extremely low (log Ksp = −36.8).49 We therefore performed thermodynamic calculations to explore the possibility of HgS(s) precipitation under our experimental conditions in MCM. Prior to addition to cell suspensions, Hg(II) is quantitatively complexed with cysteine (Table S5). However, upon Hg(II) and cysteine addition to cells, Hg(II) speciation changes as cysteine is oxidized and sulfide is released to the exposure medium. In all assays lacking cysteine in its reduced form at t = 0 h, the calculated sulfide threshold concentration to create an oversaturated solution with respect to HgS(s) (∼10−27 M or ∼1 sulfur atom per 1000 L) is well below the sulfide detection limit of 2 μM (Table S6). Thus, it is possible that HgS(s) forms even when sulfide is not detected in the exposure medium. The presence of cysteine in its reduced form greatly increases the solubility of Hg(II) (Figures S7 and S8). We calculated the sulfide threshold concentration for HgS(s) precipitation in 1000 μM cysteine exposure assays at the instant Hg(II) and cysteine

It is clear from Figure 4 that the total recoverable Hg after exposure to Hg(II) for 3 h differs between exponential growth phase cells with and without CCCP treatment (Figure 4A,C). Since it is unlikely that CCCP would alter the amount of Hg that binds to the vial wall in the presence of cells, the Hg loss in the absence of CCCP appears to be due to Hg(II) reduction and volatilization. We do not measure Hg(0) production during bioassays, as this was not anticipated. A BLASTp of the genome of E. coli ATCC 25922 reports no known mercuric reductase genes.39 However, Geobacter species are able to reduce Hg(II) via c-type cytochromes,47,48 and it is possible that E. coli ATCC 25922 also uses this pathway. Cytochromes are components of the electron transport chain, which is coupled to the cell’s PMF. Hence, a PMF inhibitor should effectively prevent Hg(II) reduction. Regardless of the exact cause for Hg loss during the 3-h bioassay, our results inform cellular Hg(II) localization. Statistical comparisons (p < 0.05) of the data presented in Figure 4 are included in Table S3. Thermodynamic Calculations Predict HgS(s) Formation in Hg(II) Bioassays. The addition of high cysteine concentrations (1000 μM) can inhibit bacterial Hg(II) 4646

DOI: 10.1021/acs.est.6b06400 Environ. Sci. Technol. 2017, 51, 4642−4651

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Environmental Science & Technology Table 1. EXAFS Linear Combination Fit Results for Three Models exposure conditions Hg (nM)

Cys (μM)

model 1

model 2

model 3

Hg speciation (%)

Hg speciation (%)

Hg speciation (%)

Hg(Cys)2 (Hg−S2)

exponential phase 50 0 70.7 ± 50 100 22.7 ± 500 0 40.9 ± 500a 1000 25.3 ± stationary phase (− glucose) 50a 0 100 50a 100 100 500a 0 100 500a,b 1000 77.1 ± exponential phase (+50 μM CCCP) 50 100 27.1 ± 500 0 100 500 1000 5.2 ±

Hg(Cys)4 (Hg−S4)

R factor

± ± ± ±

2.6 2.1 1.0 1.1

0.3774 0.3259 0.1557 0.1630

74.1 26.9 43.4 28.1

2.4

22.9 ± 2.4

0.1077 0.1230 0.0948 0.3112

100 100 100 80.5 ± 2.4

3.0

72.9 ± 3.0

1.8

94.8 ± 1.8

0.4835 0.1512 0.2241

31.3 ± 3.0 100 8.4 ± 1.7

2.6 2.1 1.0 1.1

29.3 77.3 59.1 74.7

Hg(Cys)2 (Hg−S2) ± ± ± ±

2.6 2.2 1.1 1.1

β-HgS (Hg−S4)

R factor

± ± ± ±

2.6 2.2 1.1 1.1

0.4093 0.3919 0.1824 0.1671

19.5 ± 2.4

0.1077 0.1230 0.0948 0.3365

25.9 73.1 56.6 71.9

68.7 ± 3.0 91.6 ± 1.7

α-HgS (Hg−S2)

R factor

100

0.1324

0.5324 0.1512 0.2101

These samples were measured at room temperature while all others were measured at −50 °C. bWe note that the EXAFS and XANES closely resemble those of α-HgS (see Figure S19).

a

not improve the fit results to any sample, with the exception of stationary phase cells exposed to 500 nM Hg and 1000 μM cysteine whose XANES and EXAFS spectra are nearly identical to α-HgS (Figures 5 and S19). Because Hg LIII-edge XANES of Hg(II)−S species do not contain many distinguishing features, there is greater uncertainty in XANES analysis compared to EXAFS analysis, from which bond lengths and coordinating atoms are reliably obtained. However, spectral decompositions using the XANES and the EXAFS parts of the spectrum must agree. We have therefore combined a quantitative analysis of Hg(II) coordination from the EXAFS with a qualitative analysis of the XANES derivative (SI Part 11). A table of LCF results as well as the fitted EXAFS spectra are shown in Table 1 and Figure 5, respectively. Since the EXAFS spectra of the Hg(Cys)4 and βHgS standards measured at room temperature are nearly indistinguishable (Figure S17), the fraction of Hg−S2 and Hg− S4 determined from model 1 and model 2 are highly similar. An analysis of the XANES derivative, however, favors model 2 in many samples (Figure S14 and Table S11). Cells exposed to Hg(II) in their exponential growth phase bind Hg(II) differently than those in their stationary growth phase. On the basis of LCF results, all samples of cells in stationary phase have predominantly Hg−S2 coordination. In contrast, cells exposed to Hg(II) in their exponential growth phase bind Hg(II) with varying fractions of Hg−S2 and Hg−S4 coordination, where the fraction of Hg−S4 increases with cysteine addition. Cysteine addition does not influence Hg(II) coordination number when cells are harvested in their stationary growth phase. Cells in stationary growth phase have ATP concentrations ∼5 times lower than cells in exponential growth phase, implying that energy availability, either directly or indirectly related to ATP, is required to coordinate Hg(II) with 4 sulfur atoms. The relationship between cellular ATP concentration and Hg(II) coordination number was further investigated with exponential phase cells treated with 50 μM CCCP without added cysteine, which contain ATP concentrations roughly 3 times less than their untreated counterparts. Again, the cells that experience a significant decrease in cellular ATP due to CCCP treatment coordinate Hg with 2 sulfur atoms. Since the CCCP-treated

are added to cells (t = 0), which is nearly identical for exponential and stationary cells. When cysteine is fixed at ∼850 μM, HgS(s) should form when total sulfide exceeds ∼10 μM and ∼0.5 μM for total Hg(II) concentrations of 50 nM and 500 nM, respectively (Table S6). Since we detect ∼30 μM sulfide in the exposure medium after a 3 h exposure of exponential phase cells to 1000 μM cysteine, regardless of added Hg(II) concentration, HgS(s) should form under these conditions. However, for the stationary phase cells exposed to 1000 μM cysteine, HgS(s) precipitation is only possible when the total Hg(II) concentration is 500 nM, since the sulfide threshold concentration of 0.5 μM is below our detection limit. 4-Fold Hg(II)−S Coordination in Bacteria Depends on Cellular Metabolism and Cysteine Addition. Hg LIII-edge EXAFS spectra of E. coli (containing cell envelope and intracellular fraction of Hg(II)) and Hg(II) references are presented in Figure 5 and Figure S17, respectively. The NaClO4-washed bacterial samples contain approximately 3−30 μg Hg per g of biomass (wet weight), which we calculated from the information provided in Figure 4 and known biomass weight. We demonstrate that the cytoplasmic membrane remained intact after Hg(II) and cysteine exposure (Figure S6). PCA, performed on the EXAFS spectra of 12 Hg(II)-cell samples (k range of 2.5−9.5 Å−1), shows that only 2 components (i.e., 2 Hg binding environments) are required to explain the majority of the variance (Figure S16). Target transforms indicate that 2-fold coordinated Hg(II)−S (Hg−S2) standards and 4-fold coordinated Hg(II)−S (Hg−S4) standards are the spectral components present in the data (Figure S17). We find no evidence for the presence of 3-fold coordinated Hg(II)−S (Hg−S3) in our data set, given that Hg−S2, Hg−S3, and Hg−S4 are 3 independent binding environments.50 A spectral decomposition of the EXAFS by LCFs provides the fraction of Hg−S2 and Hg−S4 in each bacterial sample. We fit our spectra considering 3 models. Model 1 describes Hg(II) as coordinated solely to organic sulfur using Hg(Cys)2 as a reference for Hg−S2 and Hg(Cys)4 as a reference for Hg−S4. Model 2 describes Hg(II) in the cell as a mixture of organic and inorganic species using Hg(Cys)2 as a reference for Hg−S2 and β-HgS (metacinnabar) as a reference for Hg−S4. Model 3 includes α-HgS (cinnabar) as a reference for Hg−S2, which did 4647

DOI: 10.1021/acs.est.6b06400 Environ. Sci. Technol. 2017, 51, 4642−4651

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be a viable candidate for complexing Hg(II) since it is the second most abundant cellular metabolite in glucose-fed E. coli in exponential growth phase.64 However, the 4-coordinate HgGSH complex is not observed at physiological pH.65 Since cysteine promotes the biosynthesis of sulfide and enhances the formation of Hg−S4 in exponential growth phase cells, it is also possible that the Hg−S4 species that we observe is β-HgS. The inclusion of β-HgS in LCFs (model 2) of exponential growth phase cells exposed to 500 nM total Hg(II) improved the quality of the fit results of the XANES derivative (Figure S19), and our thermodynamic models predict HgS(s) precipitation in all assays where sulfide is produced (Table S6). However, we also observe β-HgS-like species in exponential phase cells when we do not detect extracellular sulfide. It is possible that bacteria that are not adapted to sulfidic environments contain low levels of sulfide (below our detection level) capable of binding Hg(II). H2S is involved in metabolic and signaling processes in aerobic organisms, traits that may be widespread in all domains of life.66 While there is evidence for the formation of cell-associated β-HgS in this study, more work is required to eliminate the possibility of the formation of structurally similar Hg−S clusters that would appear like β-HgS with XAS (e.g., Hg in metallothioneins or Hg displacing Fe in Fe−S clusters).67,68 As Hg(II) must pass through the cell envelope during uptake by E. coli, the Hg(II) coordination number in the cell envelope and inside the cytoplasm should provide some insight into the mechanisms involved in Hg(II) biouptake. Comparing cellular Hg(II) localization results with the results for Hg(II) coordination in bacteria (Figure 4 and Table 1), the localization of Hg(II) within the cell seems to have no effect on Hg(II) coordination; Hg−S2 and Hg−S4 both occur in the cytoplasm as well as the cell envelope. In addition, the fraction of Hg−S2 and Hg−S4 has no apparent correlation with the fraction of intracellular Hg(II). For example, when exponential phase cells are exposed to 50 nM Hg and 100 μM cysteine, the majority of cell-bound Hg(II) is intracellular with Hg−S4 as the primary coordination environment. However, exponential phase cells exposed to 500 nM Hg and 1000 μM cysteine also primarily have Hg−S4 coordination, but only around 30% of the cellassociated Hg(II) is intracellular. Thus, the Hg(II) coordination in the cell after exposure to Hg(II) for 3 h must not predict Hg(II) bioavailability alone. For example, it is possible that Hg(II) is internalized by the cell as Hg−S2 and the coordination of Hg(II) later changes to Hg−S4 (due to Hg sequestration or a different chemical composition inside the cell). If the Hg−S4 species that we observe is particulate β-HgS, our results suggest that β-HgS can exist in both the cell envelope and in the cytoplasm and may play a role in Hg(II) uptake. Our finding that E. coli will degrade added cysteine and release sulfide into the exposure medium should compel future Hg(II) biouptake studies involving cysteine to check for cysteine degradation and sulfide biosynthesis. Cysteine desulfhydrase genes are widespread in both prokaryotes and eukaryotes, and it is possible that past studies on Hg(II) biouptake in the presence of cysteine misinterpreted their data due to incomplete information on Hg(II) speciation. Our thermodynamic calculations show that dissolved and particulate Hg(II)-sulfide species can form in the presence of cysteine, obscuring the identity of the Hg(II) species that is actually bioavailable. Additionally, our XAS results provide evidence that Hg(II)-sulfide species (β-HgS) become associated with the

cells with added cysteine do not experience any drop in cellular ATP concentration, we observe the formation of Hg−S4 in the cell. Sulfide production and thermodynamic predictions for HgS(s) formation in exponential phase assays offer an explanation for why XANES analysis favors model 2, which includes β-HgS as the Hg−S4 species.

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DISCUSSION We show that, with and without the addition of cysteine, the binding environment Hg(II) associated with E. coli cells differs with metabolic activity. Cells in exponential growth phase contain Hg−S4 species whereas cells in stationary growth phase contain solely Hg−S2. The treatment of exponential growth phase cells with CCCP, causing a significant decrease in ATP, also represses the formation of Hg−S4 species in the cell. To the best of our knowledge, we are the first to report the formation of Hg−S4 species associated with bacterial cells, likely because previous XAS studies were not performed on cells in exponential growth phase.22−24 Although we are unable to determine the exact nature of the 4-fold Hg(II)−S coordination environment found in our bacterial samples, we propose 3 scenarios: (1) Hg is solely complexed with thiols; (2) Hg exists as a mixture of Hg(SR)2 and β-HgS; and (3) Hg occurs as a mixture of Hg(SR)2, Hg(SR)4, and β-HgS. In biological and environmental samples, Hg(II) is commonly linearly coordinated to 2 organic reduced sulfur atoms.19,51 A 2-fold coordination structure for Hg(II) is considered very stable, the preference being attributed to relativistic effects.52 High-level ab initio calculations suggest that the linear, 2-coordinate Hg(II)-thiolate complex is most stable in solutions at neutral pH, while the trigonal Hg(II)thiolate complex is most stable in biological samples where the thiols are structurally connected.53 A 3-fold Hg(II)−SR complex has been observed in the metalloregulatory protein MerR54 and a quasi 3-fold Hg(II)-thiolate complex (coordinated by 2 sulfurs of cysteine residues and 1 chloride ion) is found in the rabbit liver metallothionein Hg18-MT.55 A recent study on the Hg(II) binding environment in peat discovered metallothionein-like clusters of Hg(II) with 3-fold coordination to organic reduced sulfur.20 Hg complexes with 4-fold Hg(II)− SR coordination are very uncommon in natural samples. However, Hg is 4-fold coordinated to thiols in many crystal structures,56 including the human Hah1 metallochaperone protein where 1 Hg atom is coordinated to 4 cysteine residues (1 at 2.33 Å, 2 at 2.53 Å, and 1 at 2.81 Å).57 A dependence of Hg−S4 coordination on cellular metabolismas reflected here by changes in ATP concentration− implies that either cells expend energy to bind Hg(II) in 4coordinate structures or that the composition of metal-binding sites in the cell differs with the availability of ATP. The synthesis of both amino acids and proteins requires energy in the form of ATP.46 Metallothioneins are small cysteine-rich proteins that sequester metals58 and could be responsible for the Hg−S4 coordination observed in this study.59−61 The bacterial metallothionein SmtA in E. coli coordinates 1 Zn atom with 4 cysteine residues,59 and it is possible SmtA can bind Hg(II) as well. Aside from metallothioneins, other candidates for binding Hg(II) with Hg(SR)2 or Hg(SR)4 coordination include free thiols within the cell. Glutathione (GSH) and cysteine are present in the cytoplasm and periplasm of Gramnegative bacteria and form 2-, 3-, and 4-coordinate Hg(II)−S complexes with Hg(II) in aqueous solution.50,62,63 While free cysteine concentration is strictly regulated in cells,35 GSH may 4648

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(4) Sunderland, E. M.; Krabbenhoft, D. P.; Moreau, J. W.; Strode, S. A.; Landing, W. M. Mercury sources, distribution, and bioavailability in the north pacific ocean: insights from data and models. Global Biogeochem. Cycles 2009, 23, 1−14. (5) Cossa, D.; Averty, B.; Pirrone, N. The origin of methylmercury in open mediterranean waters. Limnol. Oceanogr. 2009, 54 (3), 837−844. (6) Watras, C. J.; Back, R. C.; Halvorsen, S.; Hudson, R. J. M.; Morrison, K. A.; Wente, S. P. Bioaccumulation of mercury in pelagic freshwater food webs. Sci. Total Environ. 1998, 219 (2−3), 183−208. (7) Ruus, A.; Overjordet, I. B.; Braaten, H. F. V.; Evenset, A.; Christensen, G.; Heimstad, E. S.; Gabrielsen, G. W.; Borga, K. Methylmercury biomagnification in an arctic pelagic food web. Environ. Toxicol. Chem. 2015, 34 (11), 2636−2643. (8) Gilmour, C. C.; Podar, M.; Bullock, A. L.; Graham, A. M.; Brown, S. D.; Somenahally, A. C.; Johs, A.; Hurt, R. A.; Bailey, K. L.; Elias, D. A. Mercury methylation by novel microorganisms from new environments. Environ. Sci. Technol. 2013, 47 (20), 11810−11820. (9) Parks, J. M.; Johs, A.; Podar, M.; Bridou, R.; Hurt, R. A.; Smith, S. D.; Tomanicek, S. J.; Qian, Y.; Brown, S. D.; Brandt, C. C.; Palumbo, A. V.; Smith, J. C.; Wall, J. D.; Elias, D. A.; Liang, L. Y. The genetic basis for bacterial mercury methylation. Science 2013, 339 (6125), 1332−1335. (10) Hsu-Kim, H.; Kucharzyk, K. H.; Zhang, T.; Deshusses, M. A. Mechanisms regulating mercury bioavailability for methylating microorganisms in the aquatic environment: a critical review. Environ. Sci. Technol. 2013, 47 (6), 2441−2456. (11) Barkay, T.; Miller, S. M.; Summers, A. O. Bacterial mercury resistance from atoms to ecosystems. FEMS Microbiol. Rev. 2003, 27 (2−3), 355−384. (12) Barkay, T.; Gillman, M.; Turner, R. R. Effects of dissolved organic carbon and salinity on bioavailability of mercury. Appl. Environ. Microbiol. 1997, 63 (11), 4267−4271. (13) Benoit, J. M.; Gilmour, C. C.; Mason, R. P. Aspects of bioavailability of mercury for methylation in pure cultures of Desulfobulbus propionicus (1pr3). Appl. Environ. Microbiol. 2001, 67 (1), 51−58. (14) Benoit, J. M.; Gilmour, C. C.; Mason, R. P. The influence of sulfide on solid phase mercury bioavailability for methylation by pure cultures of Desulfobulbus propionicus (1pr3). Environ. Sci. Technol. 2001, 35 (1), 127−132. (15) Benoit, J. M.; Gilmour, C. C.; Mason, R. P.; Heyes, A. Sulfide controls on mercury speciation and bioavailability to methylating bacteria in sediment pore waters. Environ. Sci. Technol. 1999, 33 (6), 951−957. (16) Schaefer, J. K.; Rocks, S. S.; Zheng, W.; Liang, L. Y.; Gu, B. H.; Morel, F. M. M. Active transport, substrate specificity, and methylation of Hg(II) in anaerobic bacteria. Proc. Natl. Acad. Sci. U. S. A. 2011, 108 (21), 8714−8719. (17) Szczuka, A.; Morel, F. M. M.; Schaefer, J. K. Effect of thiols, zinc, and redox conditions on Hg uptake in Shewanella oneidensis. Environ. Sci. Technol. 2015, 49 (12), 7432−7438. (18) Schaefer, J. K.; Szczuka, A.; Morel, F. M. M. Effect of divalent metals on Hg(II) uptake and methylation by bacteria. Environ. Sci. Technol. 2014, 48 (5), 3007−3013. (19) Skyllberg, U.; Bloom, P. R.; Qian, J.; Lin, C. M.; Bleam, W. F. Complexation of mercury(II) in soil organic matter: EXAFS evidence for linear two-coordination with reduced sulfur groups. Environ. Sci. Technol. 2006, 40 (13), 4174−4180. (20) Nagy, K. L.; Manceau, A.; Gasper, J. D.; Ryan, J. N.; Aiken, G. R. Metallothionein-like multinuclear clusters of mercury(II) and sulfur in peat. Environ. Sci. Technol. 2011, 45 (17), 7298−7306. (21) Yu, Q.; Szymanowski, J.; Myneni, S. C. B.; Fein, J. B. Characterization of sulfhydryl sites within bacterial cell envelopes using selective site-blocking and potentiometric titrations. Chem. Geol. 2014, 373, 50−58. (22) Mishra, B.; O’Loughlin, E. J.; Boyanov, M. I.; Kemner, K. M. Binding of Hg-II to high-affinity sites on bacteria inhibits reduction to Hg-0 by mixed Fe-II/III phases. Environ. Sci. Technol. 2011, 45 (22), 9597−9603.

bacterial cell. In addition to cysteine, other thiol-containing ligands may biodegrade to sulfide. The abiotic transformation of Hg(II) complexes with soil organic matter to β-HgS has already been observed69 and potentially microorganisms can expedite this process.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.est.6b06400. Composition of growth and exposure media, growth curves, methods for sulfide and reduced cysteine detection, cysteine oxidation in absence of bacteria, methods for cell density, methods and results for cell membrane integrity, statistical analysis for cellular Hg(II) localization, Hg(II) speciation calculations, preparation of Hg standards for XAS, XAS data collection, and XAS data analysis (PDF)

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AUTHOR INFORMATION

Corresponding Author

*Phone: (847)-467-1376; e-mail: [email protected] (J.-F.G.). ORCID

Sara A. Thomas: 0000-0003-4970-4431 Jean-François Gaillard: 0000-0002-8276-6418 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We would like to thank Dr. Qing Ma for his beamline assistance at the APS as well as Drs. Kevin Schwartzenberg and Marco Alsina for technical assistance. Dr. Patrice Catty’s assistance with the genomic analysis of E. coli ATCC 25922 is greatly appreciated. Additionally, we are grateful for the suggestions from the anonymous reviewers to improve this manuscript. Portions of this work were performed at the DND-CAT Synchrotron Research Centre located at Sector 5 of the APS. DND-CAT is supported by the E.I. DuPont de Nemours & Co., The Dow Chemical Company, the U.S. National Science Foundation through Grant DMR-9304725, and the State of Illinois through the Department of Commerce and the Board of Higher Education Grant IBHE HECA NWU 96. This work is supported by the National Science Foundation under grant CHE-1308504 and made use of the J.B. Cohen X-ray Diffraction Facility supported by the MRSEC program of the National Science Foundation (DMR-1121262) at the Materials Research Center of Northwestern University.

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REFERENCES

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