Spectroscopic and Microscopic Evidence of Biomediated HgS

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Ecotoxicology and Human Environmental Health

Spectroscopic and microscopic evidence of biomediated HgS species formation from Hg(II)-cysteine complexes: implications for Hg(II) bioavailability Sara Anne Thomas, Kara Rodby, Eric Roth, Jinsong Wu, and Jean-François Gaillard Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.8b01305 • Publication Date (Web): 05 Aug 2018 Downloaded from http://pubs.acs.org on August 6, 2018

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Environmental Science & Technology

Spectroscopic and microscopic evidence of bio-mediated HgS species formation from Hg(II)-cysteine complexes: implications for Hg(II) bioavailability

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Sara A. Thomas,1*+ Kara E. Rodby,1# Eric W. Roth,2 Jinsong Wu,2 and Jean-François Gaillard *

1

Department of Civil and Environmental Engineering, Northwestern University, 2145 Sheridan Road, Evanston, IL, 60208 2

Department of Materials Science and Engineering, NUANCE Center, Northwestern University, Evanston, IL, 60208 +

Present address: Department of Geosciences, Princeton University, Guyot Hall, Princeton, NJ, 08544 #

Present address: Department of Chemical Engineering, MIT, 25 Ames St, Cambridge, MA, 02142

*Corresponding authors: Sara A. Thomas Email: [email protected] Phone: (609)-258-2339 Jean-François Gaillard Email : [email protected] Phone : (847)-467-1376

ACS Paragon Plus Environment


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ABSTRACT

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We investigated the chemistry of Hg(II) during exposure of exponentially-growing bacteria

3

(Escherichia coli, Bacillus subtilis, and Geobacter sulfurreducens) to 50 nM, 500 nM, and 5 µM

4

total Hg(II) with and without added cysteine. With x-ray absorption spectroscopy (XAS), we

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provide direct evidence for the formation of cell-associated HgS for all tested bacteria. The

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addition of cysteine (100 – 1000 µM) promotes HgS formation (> 70% of total cell-associated

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Hg(II)) as a result of the biodegradation of added cysteine to sulfide. Cell-associated HgS species

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are also detected when cysteine is not added as a sulfide source. Two phases of HgS – cinnabar

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(α-HgS) and metacinnabar (β-HgS) – form depending on the total concentration of Hg(II) and

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sulfide in the exposure medium. However, α-HgS exclusively forms in assays that contain an

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excess of cysteine. Scanning transmission electron microscopy (STEM) images reveal that

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nanoparticulate HgS(s) is primarily located at the cell surface/extracellular matrix of gram-

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negative E. coli and G. sulfurreducens and in the cytoplasm/cell membrane of gram-positive B.

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subtilis. Intracellular Hg(II) was detected even when the predominant cell-associated species was

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HgS. This study shows that HgS species can form from exogenous thiol-containing ligands and

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endogenous sulfide in Hg(II) biouptake assays under non-dissimilatory-sulfate reducing

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conditions, providing new considerations for the interpretation of Hg(II) biouptake results.

18 19 20 21 22 23


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INTRODUCTION

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Human activities have greatly enhanced the presence of mercury (Hg) in aquatic

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environments.1 The principal route of human exposure to Hg is the consumption of fish

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contaminated with monomethylmercury (MeHg), a potent neurotoxin. In the environment,

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anaerobic bacteria and archaea containing the hgcAB gene cluster primarily produce MeHg from

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Hg(II),2 and the conversion likely occurs in the cytoplasm.3, 4 Thus, understanding the factors

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that control and promote bacterial Hg(II) uptake is essential to predict the environmental fate of

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Hg.

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Although a pathway for bacterial Hg(II) uptake has yet to be directly confirmed, two

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models currently exist: (1) the passive diffusion of neutral HgS species,5-7 with recent evidence

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indicating small HgS clusters/nano-particulates,8-11 and (2) the energy-dependent uptake of

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Hg(II) by active transport, support for which has been observed in Geobacter sulfurreducens,

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Desulfovibrio desulfuricans, and Shewanella oneidensis.12, 13 The experiments describing Hg(II)

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biouptake by active transport were all performed in the presence of thiol-containing ligands with

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high Hg(II) affinities (e.g., cysteine for G. sulfurreducens and S. oneidensis and glutathione for

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D. desulfuricans), and it is unclear if active uptake would be observed in the absence of these

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ligands.

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The speciation of a metal determines its bioavailability.14 Microbial MeHg production

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predominantly occurs in anoxic environments (e.g., sediments, soils, and marshes),15 in which

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Hg(II) speciation is predicted to be controlled by thiol-containing ligands in natural organic

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matter (NOM) and/or sulfides.16 Micromolar concentrations of some thiol-containing organic

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ligands (e.g., cysteine, glutathione, and dissolved organic matter, DOM) can greatly enhance the

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biouptake and methylation of Hg(II) by various bacterial species for reasons that are not well


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understood.8, 12, 13, 17-21 The influence of thiol-containing ligands is largely attributed to the uptake

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of Hg(II)-thiol complexes12, 22 or thiol-facilitated exchange of Hg(II) with an unknown protein

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able to transport Hg(II).20 However, some of these studies supporting the active transport

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pathway have neglected to consider the rapid degradation of thiol-containing ligands to sulfide

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by bacteria. For example, cysteine degradation to sulfide is known to occur in Escherichia coli23-

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26

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species in the exposure medium, even in the presence of excess cysteine.8, 10, 25, 27 A

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concentration of 100 µM added cysteine can induce the release of sulfide at detectable

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concentrations (> 2 µM) by E. coli within 1 hour,25 which lies within the cysteine concentration

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range commonly tested in Hg(II) biouptake studies (i.e., 1 – 1000 µM). Thus, Hg(II) biouptake

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assays with added thiol-containing ligands, especially cysteine, are likely to contain a mixture of

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Hg(II)-sulfide and Hg(II)-thiol species in the exposure medium.

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and many Desulfovibrio species27 and is predicted to promote the formation of Hg(II)-sulfide

Graham et al. observed that DOM will enhance Hg(II) bioavailability to D. desulfuricans

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ND132 under sulfidic conditions and suggested that the presence of DOM promotes the

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formation of small HgS clusters/nanoparticles that are bioavailable to bacteria.8, 10 Thiol-

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containing ligands (e.g., cysteine, DOM, and NOM) are known to decrease the growth rate of

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HgS(s) particles by stabilizing them against further nucleation or aggregation28-32 as well as

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control the crystallinity of HgS(s),11, 29, 32 which may have led to the increase in HgS(s)

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bioavailability. Interestingly, Graham et al. only observed that DOM will enhance MeHg

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production by D. desulfuricans at low micromolar total sulfide concentrations (< 30 µM), which

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favor HgS nanoparticle/cluster formation.10 At high total sulfide concentrations (> 100 µM),

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which may favor dissolved HgS species, DOM had no effect on MeHg production.10 Regrettably,

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direct evidence for the formation of HgS(s) nanoparticles/clusters within Hg(II) biouptake assays


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is lacking, and the molecular scale interactions of these HgS species with bacterial cells remain

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unknown. We have recently shown using x-ray absorption spectroscopy (XAS) that exponentially

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growing E. coli cells exposed to Hg(II) in the presence and absence of cysteine for 3 hours

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contain β-HgS(s)-like species, associated with both the cell cytoplasm and cell envelope.25

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Cysteine addition promoted the formation of these β-HgS(s)-like species only with exponentially-

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growing cells (i.e, when sulfide from cysteine biodegradation was detected in the exposure

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medium). In the absence of added cysteine, β-HgS(s)-like species likely formed from sulfide

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sources already present in the cells including Fe-S clusters33 and assimilatory sulfate reduction.34

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When cysteine and Hg(II) were added to stationary phase cells under the same conditions,

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cysteine biodegradation to sulfide was undetectable in the exposure medium, and Hg(II) was

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observed as Hg(SR)2 in the bacterial cells. Thus, the β-HgS(s)-like species that we identified in

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the presence of added cysteine are likely particulate β-HgS(s) that form via cysteine

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biodegradation to sulfide. It is even possible that bacterial enzymes can catalyze dealkylation

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reactions to convert Hg(II)-thiolate species to HgS(s), as recently described to occur abiotically.35,

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36

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Hg(II)-S clusters in metallothioneins, for example, would have similar XAS spectra.

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However, we were unable to definitively identify particulate β-HgS(s) in our samples since

The central hypothesis that we are exploring herein is that the addition of thiols

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metabolized by bacteria, cysteine in this case, promotes the formation of mercury sulfide species

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that ultimately control the biouptake of Hg(II). We explore with XAS the effect of cysteine

90

addition and sulfide biosynthesis on the Hg(II) binding environment to three diverse species of

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bacteria in exponential growth phase including 1) the gram negative facultative anaerobe E. coli,

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2) the gram-positive facultative anaerobe Bacillus subtilis, and 3) the gram-negative obligate


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anaerobe G. sulfurreducens. Additionally, we probe samples of these three bacterial species

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exposed to Hg(II) and cysteine with various electron microscopy techniques to observe whether

95

cell-associated particulate HgS(s) forms in the presence of a large excess of thiol-containing

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ligand (cysteine). Lastly, we quantify intracellular Hg(II) in B. subtilis and E. coli exposed to

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Hg(II) and cysteine with an operationally-defined EDTA/GSH wash to determine if Hg(II) is

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bioavailable under conditions where cell-associated HgS(s) is directly observed.

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MATERIALS AND METHODS

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Bacterial species and growth media

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Escherichia coli ATCC 25922, Bacillus subtilis 168, and Geobacter sulfurreducens PCA were

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all obtained from the American Type Culture Collection (ATCC) and stored at -80 oC in glycerol

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stock. B. subtilis and G. sulfurreducens were inoculated directly from frozen stock into their

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respective growth media while E. coli was inoculated from LB agar plates that were stored at 4

105

o

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growth phase in LB broth and subsequently transferred to a transient minimal salts medium

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(MSM; Table S1). After ~ 24 hours of shaking at 150 rpm in MSM, cells reached exponential

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growth phase and were then washed twice in the final exposure medium – a minimally

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complexing medium – and suspended at a cell density of ~2 × 108 cells per mL (OD600 = 0.2).

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MCM (pH = 7.1) consists of 20 mM MOPS buffer, 1 mM Na-β-glycerophosphate, 0.41 mM

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MgSO4, 12 mM NH4NO3, 0.76 mM isoleucine, 0.76 mM leucine, 3 nM thiamine, 10 mM

112

glucose, and 9.1 mM NaOH. B. subtilis was grown aerobically at room temperature to early

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exponential growth phase in nutrient sporulation medium phosphate (NSMP; Table S1) from

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Beveridge and Murray.37 A nonsporulating culture was produced (since the cells never reached

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stationary growth phase), washed twice and then suspended (~5 × 107 cells per mL; OD600 = 0.2)

C for no more than 4 weeks. E. coli was grown aerobically at 37 oC overnight to exponential


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in a modified version of NSMP (pH = 6.8) containing: 20 mM MOPS buffer, 1 mM Na-β-

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glycerophosphate, 0.13 mM methionine, 0.12 mM tryptophan, 10 mM glucose, 0.5% (v/v) metal

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mixture (140 mM CaCl2, 10 mM MnCl2, and 200 mM MgCl2), and 5.0 mM NaOH. G.

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sulfurreducens was grown anaerobically in a glovebox with HEPA filter (MBraun UNIlab) at

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25-30 oC in a defined medium from Schaefer et al. (Table S1) to exponential growth phase.17

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Subsequently, cells were washed once and suspended (1 – 2 × 108 cells per mL) in the assay

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buffer (pH 6.8, N2 atmosphere) containing 10 mM MOPS buffer, 0.1 mM NH4Cl, 1.3 mM KCl,

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1 mM Na-β-glycerophosphate, 0.12 mM MgSO4, 2.5 mM NaOH, 1 mM sodium acetate, and 1

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µg/mL resazurin. Sodium fumarate was added to a final concentration of 1 mM from a filter-

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sterilized stock solution (after autoclaving), and no reducing agent was added to the assay buffer.

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To prepare heat-treated cells, bacteria were grown to exponential phase and then

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incubated for 3 hours at 60 oC in their growth medium. Subsequently, the heat-treated suspension

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was washed in the respective exposure medium as described above.

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Hg(II), cysteine, and sulfide exposure assays

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All assays were conducted in acid-washed (10% HNO3) borosilicate glass under dark conditions.

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The experiments were conducted in loosely-capped 15 mL vials with a total cell suspension

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volume of 7 mL with the exception of XAS experiments which required 500 mL foil-topped

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Erlenmeyer flasks with a total cell suspension volume of 300 – 400 mL. The assays began with

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the addition of the Hg(II) solution (± cysteine) in Milli-Q water to cell suspensions in the

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specified exposure medium. Total Hg concentrations of 50 nM, 500 nM, and 5 µM as well as

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added cysteine concentrations of 100 µM and 1000 µM were employed to make Hg L3-edge

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XAS measurements possible and to maintain Hg:cysteine ratios commonly employed in Hg(II)

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uptake studies. Directly before bacterial exposure, Hg(II) solutions in Milli-Q water were


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prepared in new polypropylene microfuge tubes at 10 times the final desired concentration from

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a 10 mM Hg(NO3)2 stock solution in 1% HNO3 (trace metal grade, TMG). The final NO3-

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concentration in the anaerobic assay medium (G. sulfurreducens) was 1 µM at the highest Hg(II)

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concentration tested (500 nM). When the effect of cysteine was tested, Hg(II) was pre-

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equilibrated with cysteine for 1 hour in Milli-Q water. For anaerobic assays, cysteine and Hg(II)

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solutions were prepared in deoxygenated Milli-Q water in a glovebox (MBraun UNIlab) with N2

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atmosphere and O2 < 0.1 ppm. For assays with added sulfide, a ~50 mM Na2S⋅9H2O stock was

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prepared in deoxygenated Milli-Q water from a single crystal that was washed with Milli-Q

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water and pat dried to remove potential oxidation products. At the start of an assay, small

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volumes (µL) of sulfide stock were added to cell suspensions directly after Hg(II) addition.

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Aerobic assays (E. coli and B. subtilis) were shaken at 150 rpm at room temperature.

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Anaerobic assays (G. sulfurreducens) were incubated statically (mixed for ~5 seconds every

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hour) in a glovebox (MBraun UNIlab) under strict anaerobic conditions. No oxidation was

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noticed during assays as reported by the resazurin indicator.

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XAS sample preparation, measurements, and data analysis.

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To determine the Hg coordination environment in bacteria, cell pellets were collected and

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analyzed with XAS according to our previously reported method.25 Briefly, after a defined

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exposure time to Hg(II), cells were washed twice with 0.1 M NaClO4 (7500 g for 10 min), and

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the cell pellet was spread onto cellulose nitrate filter paper (0.4 µm pore size). We utilized a

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glovebox and a deoxygenated 0.1 M NaClO4 solution to wash the anaerobic samples and

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collected them on filter paper under a stream of N2 gas. Excess moisture was removed from the

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filters with a vacuum pump for 5-10 minutes. We sandwiched the filter and cell pellet between 2

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layers of Kapton tape (DuPont) and immediately plunged the sample in LN2. Samples were


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stored in a -80 oC freezer for no more than 1 week prior to XAS analysis. We previously

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confirmed that the process of plunging the sample in LN2 does not alter Hg coordination

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environment.25

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Hg LIII-edge XANES and EXAFS spectra were collected at the DuPont-Northwestern-

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Dow Collaborative Access Team (DND-CAT) beamline located in Sector 5 of the Advanced

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Photon Source. A description of the scan parameters, beamline specifications, handling of

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bacteria samples, and the preparation of Hg reference spectra is provided in our previous

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publication25 and the Supporting Information (SI), Part 4. We used principal component analysis

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to determine the number of unique Hg coordination environments in our samples, and performed

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linear combination fits of the EXAFS to determine the fraction of various Hg reference standards

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in each sample (SI, Part 5). Spectral analyses were performed with Athena38 and R.

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STEM and TEM sample preparation and imaging

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To prepare bacterial samples for imaging with scanning transmission electron microscopy

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(STEM) and transmission electron microscopy (TEM), cells were mixed with Hg(II) and

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cysteine in their respective exposure media for 3 hours. A 1 – 2 mL aliquot was then collected

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and washed 4 times with 0.1 M NaClO4 by centrifugation (8000 g for 3 min) in a 1.5 – 2 mL

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microfuge tube to remove dissolved and particulate Hg not associated with cells. The collected

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cells were imaged in two ways: (1) as whole cells (E. coli) and (2) as thinly-sliced 200-300 nm

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sections (E. coli, B. subtilis, and G. sulfurreducens).

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resuspended in a final solution of 200 µL filtered Milli-Q water (0.2 µm filter, VWR

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International), and 1 drop (< 5 µL) was immediately placed on a 200 mesh carbon-coated copper

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grid and allowed to air dry for ~10 minutes. The sample was immediately imaged at room

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temperature.

Whole cell E. coli samples were


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For the thin section samples, the cell pellet (washed with 0.1 M NaClO4) was fixed in a

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solution containing 2.5% EM grade glutaraldehyde, 2% paraformaldehyde, and 0.1M phosphate-

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buffered saline (PBS) for 1 hour at room temperature and overnight at 4 ºC. Post fixation

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occurred in 1% osmium tetroxide for 2 hours at room temperature. A graded series of ethanol

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was used for dehydration before infiltration and embedment with EMBed812 epoxy resin. The

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resin blocks were cured at 60 oC for 48 hours.

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To slice bacterial samples into 200 – 300 nm thick sections, ultramicrotomy was done on

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a Leica UC7 ultramicrotome. The sections were collected on 300 mesh thin-bar hex copper grids

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and imaged without staining. STEM images were collected on a Hitachi HD-2300 field emission

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scanning transmission electron microscope (200 kV accelerating voltage) utilizing high-angle

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annular dark field (HAADF), bright field phase contrast, and secondary electron imaging

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modes. Energy dispersive x-ray spectroscopy (EDS) maps, line scans, and point-and-shoot data

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were collected with an ultra-sensitive dual-detector system attachment (Thermo Scientific). TEM

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micrographs and selected area electron diffraction (SAED) patterns were recorded on a Hitachi

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H-8100 transmission electron microscope using an accelerating voltage of 200 kV.

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Quantifying cell-bound and intracellular Hg(II)

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After cell suspensions mixed with Hg(II) and cysteine for the desired amount of time, aliquots

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were collected for the determination of (1) total recoverable Hg (dissolved + cell-bound), (2)

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dissolved Hg (0.2 µm nylon filter, VWR International), and (3) intracellular Hg. Cell-bound Hg

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was calculated as the difference between the total recoverable Hg and the dissolved Hg. Samples

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for determining dissolved and total recoverable Hg were preserved in ~1% HCl (TMG) until the

206

measurement of total Hg with a Direct Mercury Analyzer (DMA-80, Milestone). Fresh 50 nM

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Hg solutions in relevant assay media were measured regularly to ensure the instrument


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calibration was maintained. In addition, the nylon filters do not bind a significant amount of Hg

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(data not shown).

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Intracellular Hg was determined in E. coli and B. subtilis following an EDTA/glutathione

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(GSH) wash procedure described in our previous study.25 Briefly, 4 mL of the cell suspension

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was centrifuged (7000 g for 10 min) and resuspended in 2 mL of 50 mM EDTA and 100 mM

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oxalate solution (pH = 7.5). After mixing for 10 minutes, 2 mL of a 10 mM glutathione (GSH)

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and 3 mM ascorbate solution (pH = 7) were added and mixed for an additional 10 minutes.

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Subsequently, a 2 mL aliquot of cells suspended in the EDTA/GSH solution were filtered onto

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0.2 µm cellulose nitrate filters and the whole filters were analyzed for total Hg content (i.e.,

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intracellular Hg(II)) with a DMA-80.

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Intracellular Hg(II) concentration was normalized considering the OD600 of the sample

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before the wash (in exposure medium) and after the wash (in EDTA/GSH). The density of cells

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(#cells/mL) per OD600 is not significantly different in the exposure medium and the EDTA/GSH

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wash solution for E. coli or B. subtilis (data not shown). We have previously verified that the

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wash does not affect the integrity of the cytoplasmic membrane for E. coli with the

223

LIVE/DEAD® BacLight™ Bacterial Viability Kit.25 However, this same test could not be

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applied to B. subtilis in the adapted NSMP exposure medium (i.e., viable cells with intact

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membranes fluoresced red and did not provide a negative control). Thus, it is unknown whether

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this wash method damages the B. subtilis cytoplasmic membrane.

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Cysteine degradation/oxidation and sulfide production in the exposure medium. At the

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beginning (t = 0 hr) and at various time points during the Hg(II) and cysteine exposure assays, a

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1 mL aliquot was centrifuged (15,000 g for 5 min) for the determination of acid labile sulfide in

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the supernatant by a method adapted from Cline39, 40 as well as cysteine and cystine (oxidized


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cysteine) by a method adapted from Gaitonde.41 Detailed methods are reported in our previous

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publication.25 The detection limit for sulfide and cysteine was 2 µM and 5 µM, respectively.

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Thermodynamic modeling. All Hg(II) speciation calculations were performed with the program

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ChemEQL.42 The equilibrium constants used in the calculations are reported in Table S5.

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RESULTS

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Cysteine degradation/oxidation and sulfide production In agreement with our previous findings, E. coli, G. sulfurreducens, and B. subtilis all

237 238

produce and release sulfide into the exposure medium after exposure to 100 µM and 1000 µM

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added cysteine (Figure 1). Sulfide was below detection limits after cysteine exposure to heat-

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treated cells, suggesting that cysteine degradation to sulfide requires viable cells. Cysteine

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desulfhydrases are the likely candidates for cysteine degradation by the bacteria in this study.23,

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24, 43

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minutes) after exposure of B. subtilis and G. sulfurreducens, respectively, to 1000 µM cysteine.

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Thus, cysteine biodegradation to sulfide in these organisms occurs rapidly.

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Around 6 µM and 4 µM sulfide were released into the exposure medium immediately (0 – 2

In the case of E. coli, cysteine is abiotically oxidized upon contact with the exposure

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medium (Figure S1). The cysteine oxidation mechanism may be transition metal-catalyzed

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autoxidation,44 since we concurrently observe dissolved O2 consumption upon cysteine addition

248

to the E. coli exposure medium (data not shown). However, we do not observe abiotic cysteine

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oxidation in the exposure medium for B. subtilis or in Milli-Q water (Figure S1), which were

250

both aerated, and the potential transition metal catalyst is unclear since transition metals are not

251

intentionally added to the E. coli exposure medium. Consequently, when E. coli is exposed to

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100 µM and 1000 µM cysteine, the added cysteine is immediately oxidized to cystine upon

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contact with the exposure medium, leaving non-detectable and ~834 µM cysteine, respectively,


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at t = 0 h (Table S2). After 3 hours of exposure of E. coli to 1000 µM cysteine, ~774 µM

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cysteine remains in solution, while no cysteine is detected in the 100 µM added cysteine sample.

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In contrast, assays of B. subtilis and G. sulfurreducens exposed to 100 µM and 1000 µM cysteine

257

do not experience any cysteine oxidation at the start of the exposure period. After 3 hours of

258

exposure to 100 µM and 1000 µM cysteine, the B. subtilis assays contain ~56 µM and ~921 µM

259

cysteine, respectively, while the G. sulfurreducens assays contain ~87 µM and ~980 µM

260

cysteine, respectively (Table S2). Cysteine/cystine is also consumed by the organisms, which

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accounts for some loss in addition to cysteine oxidation (Table S2).

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STEM, TEM, and EDS reveal cell-associated HgS(s) nanoparticles

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To determine if HgS(s) will precipitate in assays where the Hg source is pre-equilibrated

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Hg(II)-cysteine complexes, we first probed cells exposed to 500 nM or 5 µM total Hg(II) and

265

1000 µM cysteine with STEM, TEM, and EDS. Precipitates of HgS(s) were observed after a 3-

266

hour exposure period of G. sulfurreducens (Figure 2A), E. coli (Figures 2C and 2D) and B.

267

subtilis (Figures 2F and 2G) to pre-equilibrated 500 nM Hg and 1000 µM cysteine. EDS analysis

268

confirms that the particles are composed of Hg and S (Figures 2B, 2H, S3E, and S3G).

269

Micrographs of 200 – 300 nm sliced cell sections show that the HgS(s) particles associated with

270

G. sulfurreducens and E. coli are located at the cell surface while those associated with B.

271

subtilis are virtually all located within the cell membrane or cytoplasm. The HgS(s) particles were

272

typically concentrated in a few areas for the samples of E. coli and G. sulfurreducens exposed to

273

500 nM total Hg(II) and 1000 µM cysteine and were observed in 10 and 2 individual

274

micrographs for the respective bacterial species. In contrast, the HgS(s) particles were more

275

evenly dispersed in the sample of B. subtilis exposed to 500 nM Hg(II) and 1000 µM cysteine,

276

where many adjacent cells (we observed 21) contained one or two individual HgS(s)


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nanoparticles. The micrograph of HgS(s) particles associated with whole E. coli cells (Figure 2C)

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is likely a more accurate representation of how the nanoparticles interact with cells, as these

279

samples were not fixed or treated with any chemicals prior to imaging.

280

We note that the settling time of HgS(s) particles greater than ~100 nm in diameter,

281

according to Stokes’ law,45 is calculated to be less than the centrifugation time during the

282

collection of bacterial cells (3 min). To further test whether HgS(s) nanoparticles were physically

283

attached to bacterial cells prior to sample collection by centrifugation, we passed aliquots

284

through a 200 nm pore size nylon filter. Hg was not detected in the filtrate of any sample imaged

285

with STEM/TEM (Figure S11), providing further evidence that HgS(s) particles smaller than the

286

200 nm pore size are attached to bacterial cells. The one exception is G. sulfurreducens exposed

287

to 500 nM Hg and 1000 µM cysteine, where ~25% of the total recoverable Hg was present in the

288

filtrate (Figure S11). However, it is very likely that this dissolved Hg is in fact MeHg, which G.

289

sulfurreducens is known to produce and release rapidly into the exposure medium.12

290

Although some of the HgS(s) particles in Figure 2C appear to not be in contact with E.

291

coli, it is possible that they are associated with the cells’ extracellular polymeric substance

292

(EPS).46, 47 An analysis of the same cell group with secondary electron imaging shows a different

293

composition (i.e., potentially EPS) between the region where the HgS(s) particles are

294

concentrated and the background (Figure S3C, blue arrow). In addition, HgS(s) particles trapped

295

in the cells’ fimbriae/pili, the filamentous organelles composed of proteins that encapsulate the

296

cell,48 are visible in the secondary electron image (Figure S3C, red arrow). We tested a higher

297

concentration of Hg(II) (5 µM) with 1000 µM cysteine that was mixed with E. coli for 3 hours

298

and found that the HgS(s) particles that form have a different morphology (Figure 2E); they

299

appear larger and more jagged. Selected area electron diffraction (SAED) patterns of HgS(s)


14

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300

particles in whole cell mounts of E. coli exposed to 1000 µM cysteine and 5 µM total Hg reveal

301

they are polycrystalline α-HgS (Figure S2D). We were unable to observe SAED patterns for

302

HgS(s) particles that formed when E. coli was exposed to 1000 µM cysteine and 500 nM Hg(II);

303

thus, it is possible that particles forming at this lower total Hg(II) concentration are amorphous.

304

Since the HgS(s) particles are forming in a biological system still in the presence of a large excess

305

of cysteine, many factors must control their size, morphology, and phase.

306

Detection of cell–associated Hg(SR)2, Hg-S4, β -HgS, and α-HgS by XAS

307

Hg LIII-edge XANES and EXAFS were used to determine the local coordination environment of

308

Hg(II) associated with E. coli, G. sulfurreducens, and B. subtilis during Hg(II) exposure with and

309

without added cysteine (Figure 3). We additionally assessed the effect of exposure time and

310

added sulfide (without cysteine) on Hg(II) coordination in E. coli (Figure 3A). The total

311

recoverable Hg after the exposure period, cell-bound Hg(II), and dissolved Hg(II) for each of the

312

samples analyzed by XAS is presented in the SI (Figure S11).

313

We performed principal component analysis (PCA) on the EXAFS spectra of 19 cell

314

samples exposed to Hg(II) (k range of 2.5 – 9.5 Å-1) following our previously reported

315

approach.25 PCA shows that only 2 components (i.e., 2 Hg(II) binding environments) are

316

required to explain the Hg(II) coordination environment in cell samples (Figure S6). Two-fold

317

coordinated Hg(II)-S (Hg-S2) and four-fold coordinated Hg(II)-S (Hg-S4) are the spectral

318

components in the data set. However, given the noise in the data, it is difficult to differentiate

319

inorganic and organic Hg(II) species bound to sulfur (i.e., whether the Hg-S2 species is α-HgS or

320

Hg(SR)2 and whether the Hg-S4 species is β-HgS or Hg(SR)4) with just the EXAFS because the

321

spectra of the inorganic and organic reference compounds are nearly identical (Figure S8). It is


15


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322

also possible that some samples contain both the inorganic and organic forms of Hg-S2 and Hg-

323

S4.

324

Spectral decomposition by linear combination fits (LCFs) of the EXAFS provided the

325

fractions of Hg-S2 and Hg-S4 found in bacterial samples, which we identified as Hg(SR)2, α-

326

HgS, or β-HgS with the XANES derivative (Figure 3). The XANES derivative contains a

327

characteristic peak at ~12,305 eV for β-HgS and at ~12,309 eV for α-HgS (Figures S7A and

328

S10). The identity of Hg-S4 in some samples could not be determined from the XANES

329

derivative, and thus, we report the speciation as unidentified 4-coordinate Hg-S in Figure 3.

330

Unidentified Hg-S4 species are prevalent in samples exposed to the lowest total Hg concentration

331

(50 nM) and could consist of β-HgS, Hg(SR)4, Hg-S clusters, or possibly under-coordinated β-

332

HgS29, 32 if the nanoparticles are sufficiently small (i.e., tens of nanometers in diameter).

333

As we observed previously in E. coli,25 the addition of cysteine to G. sulfurreducens and

334

B. subtilis increases the fraction of β-HgS associated with the cell (Figure 3). The presence of

335

particulate HgS(s) in some of those samples (i.e., the 3 bacterial species exposed to 500 nM Hg

336

and 1000 µM cysteine), as observed by STEM, further supports that we correctly identified β-

337

HgS with XAS. Our XAS results show that E. coli and G. sulfurreducens contain Hg-S4 even

338

when cysteine is not added at 500 nM total Hg, which may be due to β-HgS precipitation from

339

endogenous sulfide sources other than cysteine degradation. The fraction of cell-associated Hg-

340

S4 does not change significantly during exposure of E. coli to 500 nM Hg for 30 minutes, 1 hour,

341

or 3 hours (Figure 3A). However, the Hg-S4 species is easily identified as β-HgS only in the

342

sample that mixed for 3 hours. Cell-associated Hg-S4 is present at a fraction of 20-30% in all

343

bacterial cells exposed to 50 nM Hg(II) in the absence of added cysteine. In the presence of 50

344

nM total Hg and 100 µM added cysteine, 70-90% of cell-associated Hg is present as Hg-S4 in all


16

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345

tested bacteria. When E. coli is exposed to 5 µM total Hg (no cysteine) for 3 hours, no HgS

346

species are revealed by XAS, and the XAS spectra resemble the Hg(cysteine)2 reference (i.e.,

347

Hg(SR)2). In the case of B. subtilis, cells exposed to 500 nM total Hg (no cysteine) also contain

348

solely Hg-thiol coordination (as Hg(SR)2).

349

Interestingly, we observe two different phases of HgS (α-HgS and β-HgS) associated

350

with E. coli depending on the total concentration of Hg(II) (500 nM or 5 µM) added with 1000

351

µM cysteine. Our XAS findings show that α-HgS forms in the sample exposed to 5 µM total Hg

352

and 1000 µM cysteine (Figure 3), which agrees with the SAED pattern for the same sample

353

(Figure S2D). Additionally, the XAS results show that the amorphous HgS(s) observed by

354

STEM/TEM to form in E. coli assays with 500 nM Hg and 1000 µM cysteine has local Hg(II)

355

coordination resembling β-HgS. Since E. coli exposed to 1000 µM cysteine produces around 30

356

– 40 µM sulfide regardless of the total concentration of added Hg(II) (Figure 1A), the ratio of

357

Hg(II) to sulfide must influence the phase of HgS(s) that precipitates in the presence of cysteine.

358

However, when we exposed E. coli to 500 nM or 5 µM Hg with 30 µM total sulfide alone (no

359

cysteine), both samples contained predominantly β-HgS after mixing for 3 hours (Figure 3A).

360

Thus, it appears that the formation of α-HgS in our system depends on the presence of excess

361

cysteine in addition to the ratio of total Hg to total sulfide. Samples of E. coli exposed to 500 nM

362

Hg or 5 µM Hg with 1000 µM cysteine display identical XAS results whether the Hg(II)

363

exposure time is 1 hour or 3 hours (Figure 3A), indicating that the HgS species particles form

364

within an hour and do not change phase after an additional 2 hours mixing with E. coli.

365

Hg(II) biouptake and thermodynamic calculations

366

We measured the uptake of Hg(II) by E. coli and B. subtilis under conditions where HgS species

367

will form in the presence of excess cysteine (0 – 500 nM Hg(II) with constant 1000 µM added


17


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368

cysteine; Figure 4A,B). During a 3-hour exposure to Hg(II) and cysteine, the measured

369

intracellular concentration of Hg(II) in E. coli – as determined by the EDTA/GSH wash

370

procedure – does not exceed ~100 nM Hg for all total added Hg concentrations (Figure 4A). We

371

have previously observed that intracellular Hg reaches a similar plateau with increasing total

372

added Hg in the presence of 1000 µM cysteine with a genetically-modified mer-lux E. coli

373

biosensor, an independent method of measuring Hg(II) biouptake.18 A significant fraction of Hg

374

(~200 – 300 nM) is located at the cell envelope at total added Hg(II) concentrations above 250

375

nM (Figure 4A). In contrast, the intracellular concentration of Hg(II) in B. subtilis increases

376

more linearly with total added Hg(II) (Figure 4B). In addition, the concentration of Hg(II)

377

associated with the cell envelope in B. subtilis is less than E. coli, although the measurement

378

error is greater. About half of the total recoverable Hg(II) is dissolved at low total added Hg(II)

379

for E. coli and B. subtilis (see inset in Figures 4A and 4B). At high total added Hg(II), dissolved

380

Hg(II) is not detected in the E. coli exposure assays (i.e., total recoverable Hg is 100% cell-

381

bound), whereas dissolved Hg remains ~15 – 20% of the total recoverable Hg(II) in the B.

382

subtilis assays (Figure 4A,B). The total recoverable Hg for all exposure conditions is less than

383

the total added Hg likely due to Hg(II) reduction to Hg(0) and volatilization, which was observed

384

previously in E. coli.25

385

Thermodynamic calculations were performed to predict the total Hg(II) concentration

386

required to reach saturation in the exposure medium (without bacterial cells) with respect to

387

HgS(s) using the final concentrations of cysteine and sulfide detected after a 3-hour exposure of

388

E. coli and B. subtilis to 1000 µM cysteine (Figures 4C and 4D). As total added Hg(II) is

389

increased, the medium becomes oversaturated with respect to HgS(s) at ~12 nM and ~22 nM total

390

added Hg(II) for E. coli and B. subtilis, respectively. Therefore, above ~12 nM and ~22 nM total


18

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391

added Hg(II), HgS(s) formation is thermodynamically favorable for the experimental data shown

392

in Figures 4A and 4B, respectively.

393

DISCUSSION

394

We report direct evidence of HgS formation in experiments performed with three diverse

395

species of bacteria – E. coli, B. subtilis, and G. sulfurreducens – exposed to Hg(II) with and

396

without cysteine under non-dissimilatory-sulfate reducing conditions. Cysteine addition induces

397

the formation of HgS as a result of cysteine biodegradation to sulfide, even when Hg(II) is

398

introduced to cells as a pre-equilibrated Hg(II)-cysteine complex and while the remaining

399

cysteine concentration is in excess of Hg(II) and sulfide. Since no sulfide was detected in the

400

exposure medium for heat treated cells under the same conditions, cysteine degradation is

401

biologically mediated. In addition to the microbial species in this study, other bacteria that are

402

known to cleave cysteine and form sulfide include many Hg(II)-methylating and non-

403

methylating Desulfovibrio species.27, 49

404

The STEM/TEM observations provide some insights into the association between cells

405

and HgS(s) particles that can in turn inform about Hg(II) bioavailability. Recent studies have

406

proposed that the Hg(II) in HgS(s) nanoparticles or Hg-S clusters is bioavailable to bacteria.8, 9

407

Our STEM and TEM images of whole cells and thin sections of bacteria exposed to 500 nM

408

Hg(II) and 1000 µM cysteine directly show HgS(s) particles associated with the cell

409

envelope/EPS for the two gram-negative species (E. coli and G. sulfurreducens). The HgS(s)

410

particles are in direct contact with the lipophilic cell envelope or EPS material likely because

411

they are hydrophobic.28 The entrapment of HgS(s) particles in the extracellular matrix (e.g., EPS,

412

fimbriae, or pili) could explain why we observe that intracellular Hg(II) in E. coli reaches a

413

plateau with increasing total added Hg(II) and 1000 µM added cysteine. In addition, HgS(s)


19


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414

entrapment in the cell envelope could explain previous results showing limited Hg methylation

415

or uptake at higher total added Hg(II) concentrations.18, 27, 50, 51 Similar trapping of silver

416

nanoparticles by EPS has been shown previously in E. coli.52 However, it is difficult to know

417

whether the HgS(s) particles form in the exposure medium and then attach to cellular material or

418

nucleate at the surface of the cell/EPS for these gram-negative bacteria. EPS may play a

419

significant role in regulating the bioavailability of HgS(s) nanoparticles in Hg-methylating

420

bacteria and should be explored further. Although the HgS(s) particles visible with STEM are

421

located outside the cell and at the cell surface in E. coli, our biouptake studies suggest that ~100

422

nM of 500 nM total added Hg(II) is still being internalized. While the form of Hg that passes

423

through the membrane layers is still unknown, we show that Hg(II) remains bioavailable under

424

conditions with direct spectroscopic and microscopic evidence of HgS(s) formation.

425

In contrast, HgS(s) particles that precipitate in the B. subtilis assays are all observed

426

within the cell membrane or cytoplasm. At the start of the exposure of B. subtilis to 500 nM

427

Hg(II) and 1000 µM cysteine, thermodynamic calculations predict that the solution will become

428

oversaturated with respect to HgS(s) at a total sulfide concentration of greater than ~0.5 µM.

429

Since sulfide release from cysteine degradation by B. subtilis occurs rapidly, producing ~5 µM

430

sulfide in the exposure medium within the first few minutes of a 1000 µM cysteine addition, it is

431

conceivable that HgS(s) nucleation begins outside of the cell (at least outside of the cytoplasm).

432

The fact that B. subtilis is gram-positive and thus has a different membrane structure than the

433

two gram-negative bacteria in this study may explain the observed differences between HgS(s)

434

localization within the cells. It is possible that small HgS species (e.g., dissolved HgS or HgS

435

clusters) can passively diffuse into the cytoplasm of B. subtilis, where they may develop into

436

HgS(s) nanoparticles. Or, it is possible that B. subtilis can internalize HgS(s) nanoparticles. The


20

Page 21 of 32


437

uptake of nano-TiO2 by Salmonella typhimurium is reported in the literature,53 although the

438

uptake mechanism is unclear. Potentially, the presence of EPS plays a role in determining the

439

bioavailability of HgS(s) particles by acting as a barrier. E. coli K-12 contains five sets of genes

440

that are known to be involved in EPS synthesis,54 which also appear to be present in E. coli

441

ATCC 25922 from this study,55 whereas B. subtilis 168 is a strain that is reportedly defective in

442

producing EPS.56, 57

443

It is also important to note that we only observe particulate HgS(s) with STEM/TEM at

444

total Hg(II) concentrations of 500 nM and above. Particulate HgS(s) was not observed by STEM

445

for experiments performed using 50 nM total Hg(II). However, this does not mean that particles

446

or clusters do not form at 50 nM total Hg(II) since our XAS data imply that β-HgS-like species

447

are associated with all the bacterial species tested in presence or absence of 100 µM cysteine. It

448

is possible that we were unable to locate them during our observation time with the electron

449

microscope or that they were too small to be detected. Interestingly, nearly all of the Hg(II)

450

associated with E. coli and B. subtilis is intracellular (no cell envelope fraction, as determined

451

from the EDTA/GSH wash) when the cells are exposed to less than 75 nM total Hg(II) and 1000

452

µM cysteine for 3 hours (Figure 4A and 4B). Under these conditions, the exposure medium is

453

oversaturated with respect to HgS(s) according to thermodynamic calculations (Figure 4C and

454

4D). The relatively low Hg(II) concentration, combined with the coexistence of cysteine and

455

sulfide in the exposure medium, may then lead to the formation of smaller, cysteine-

456

functionalized HgS(s) particles that can diffuse into the cytoplasm, as Graham et al. suggest.8 We

457

note that some prior studies documenting cysteine’s effect on Hg(II) biouptake employed lower

458

total added Hg (1 – 40 nM) and cysteine (1 – 50 µM). We therefore performed a simple

459

calculation using the thermodynamic model in Table S5 to predict how our findings could be


21


Page 22 of 32

460

applied to these previous studies. At a total Hg concentration as low as 0.5 nM, HgS(s) formation

461

is predicted to occur when cysteine is 1 µM and sulfide is 0.1 µM (i.e., the Hg:cysteine:sulfide

462

ratio in this study), suggesting that our results could be applicable to experiments with lower

463

total Hg and cysteine.

464

We also observe the formation of β-HgS and α-HgS from Hg(II)-cysteine complexes in

465

relatively short time scales (as quick as 1 hour). Manceau et al. recently described that β-HgS(s)

466

abiotically forms from Hg(II)-(cysteine ethyl ester)2 and Hg(II)-thiol complexes of natural

467

organic matter (NOM) under aerobic conditions within days.36 The proposed mechanism for β-

468

HgS(s) formation from Hg(SR)2, which is supported theoretically,35 starts with the cleavage of the

469

S-C bond in one thiolate ligand (RS) to create RS-(HgS)n-R chains. The chains may converge to

470

initially form disordered β-HgS and then crystalline β-HgS over time, or the chains may initially

471

align in parallel to yield α-HgS. Similar mechanisms may have been in play in our systems,

472

which may have been biologically catalyzed by cysteine desulfhydrase enzymes known to cleave

473

the S-C bond in cysteine.23 However, our conditions slightly differ due to presence of

474

biosynthesized free sulfide also in the assay medium.

475

Our results highlight the importance of endogenous sulfide and the sulfide from

476

exogenous cysteine biodegradation in controlling Hg(II) speciation in diverse species of bacteria.

477

These pathways for sulfide formation have rarely been considered in previous Hg methylation

478

studies. As we show here, cell-associated HgS is directly observed in Hg(II) biouptake assays

479

with E. coli, B. subtilis, and G. sulfurreducens, even under conditions that are aerobic (E. coli

480

and B. subtilis) and non-dissimilatory-sulfate reducing. Hg(II) speciation in biouptake assays that

481

contain cysteine is clearly dynamic. We demonstrate that Hg(II) remains bioavailable even when

482

HgS is a dominant cell-associated species (according to XAS measurements), further supporting


22

Page 23 of 32


483

that bacterial Hg(II) biouptake can occur under conditions that favor HgS formation. However,

484

additional studies are necessary to pinpoint the mechanism for Hg(II) uptake in complex

485

systems, where Hg(II) speciation changes from Hg(II)-thiol to HgS over time.

486 487

Acknowledgements

488

We are grateful to Qing Ma for his beamline assistance at the APS. We thank Dr. Isabelle

489

Michaud-Soret for helpful discussions regarding this study. This work is supported by the

490

National Science Foundation under grant CHE-1308504 and a grant to K.E.R. from the

491

Undergraduate Research Grant Program administered by Northwestern University's Office of the

492

Provost. Portions of this work were performed at the DND-CAT Synchrotron Research Center

493

located at Sector 5 of the APS. DND-CAT is supported by the E.I. DuPont de Nemours & Co.,

494

The Dow Chemical Company, the U.S. National Science Foundation through Grant DMR-

495

9304725, and the State of Illinois through the Department of Commerce and the Board of Higher

496

Education Grant IBHE HECA NWU 96. The STEM and TEM work made use of the BioCryo

497

and EPIC facility of Northwestern University’s NUANCE Center, which has received support

498

from the Soft and Hybrid Nanotechnology Experimental (SHyNE) Resource (NSF ECCS-

499

1542205); the MRSEC program (NSF DMR-1121262) at the Materials Research Center; the

500

International Institute for Nanotechnology (IIN); the Keck Foundation; and the State of Illinois,

501

through the IIN. Finally, we thank 4 anonymous reviewers for their helpful suggestions.

502

Supporting Information

503

Composition of growth media, cysteine/cystine concentration in exposure media, additional

504

STEM/TEM/EDS/SAED images, XAS data collection, XAS data analysis, Hg-cell sorption

505

results, equilibrium constants, Hg speciation calculations.


23


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Page 24 of 32

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25. Thomas, S. A.; Gaillard, J.-F., Cysteine addition promotes sulfide production and 4-fold Hg(II)–S coordination in actively metabolizing Escherichia coli. Environ. Sci. Technol. 2017, 51, (8), 4642-4651. 26. Mironov, A.; Seregina, T.; Nagornykh, M.; Luhachack, L. G.; Korolkova, N.; Lopes, L. E.; Kotova, V.; Zavilgelsky, G.; Shakulov, R.; Shatalin, K.; Nudler, E., Mechanism of H2Smediated protection against oxidative stress in Escherichia coli. Proc. Natl. Acad. Sci. U.S.A. 2017, 114, (23), 6022-6027. 27. Graham, A. M.; Bullock, A. L.; Maizel, A. C.; Elias, D. A.; Gilmour, C. C., Detailed assessment of the kinetics of Hg-cell association, Hg methylation, and methylmercury degradation in several Desulfovibrio species. Appl. Environ. Microbiol. 2012, 78, (20), 73377346. 28. Deonarine, A.; Hsu-Kim, H., Precipitation of mercuric sulfide nanoparticles in NOMcontaining water: Implications for the natural environment. Environ. Sci. Technol. 2009, 43, (7), 2368-2373. 29. Gerbig, C. A.; Kim, C. S.; Stegemeier, J. P.; Ryan, J. N.; Aiken, G. R., Formation of nanocolloidal metacinnabar in mercury-DOM-sulfide systems. Environ. Sci. Technol. 2011, 45, (21), 9180-9187. 30. Ravichandran, M.; Aiken, G. R.; Ryan, J. N.; Reddy, M. M., Inhibition of precipitation and aggregation of metacinnabar (mercuric sulfide) by dissolved organic matter isolated from the Florida Everglades. Environ. Sci. Technol. 1999, 33, (9), 1418-1423. 31. Slowey, A. J., Rate of formation and dissolution of mercury sulfide nanoparticles: The dual role of natural organic matter. Geochim. Cosmochim. Acta 2010, 74, (16), 4693-4708. 32. Poulin, B. A.; Gerbig, C. A.; Kim, C. S.; Stegemeier, J. P.; Ryan, J. N.; Aiken, G. R., Effects of sulfide concentration and dissolved organic matter characteristics on the structure of nanocolloidal metacinnabar. Environ. Sci. Technol. 2017, 51, (22), 13133-13142. 33. Blanc, B.; Gerez, C.; de Choudens, S. A., Assembly of Fe/S proteins in bacterial systems: Biochemistry of the bacterial ISC system. Biochim Biophys Acta, Mol. Cell. Res. 2015, 1853, (6), 1436-1447. 34.

Kredich, N. M., Biosynthesis of cysteine. EcoSal Plus 2008, 3, (1).

35. Enescu, M.; Nagy, K. L.; Manceau, A., Nucleation of mercury sulfide by dealkylation. Sci. Rep. 2016, 6. 36. Manceau, A.; Lemouchi, C.; Enescu, M.; Gaillot, A. C.; Lanson, M.; Magnin, V.; Glatzel, P.; Poulin, B. A.; Ryan, J. N.; Aiken, G. R.; Gautier-Luneau, I.; Nagy, K. L., Formation


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of mercury sulfide from Hg(II)-thiolate complexes in natural organic matter. Environ. Sci. Technol. 2015, 49, (16), 9787-9796. 37. Beveridge, T. J.; Murray, R. G. E., Uptake and retention of metals by cell-walls of Bacillus subtilis. J. Bacteriol. 1976, 127, (3), 1502-1518. 38. Ravel, B.; Newville, M., Athena, Artemis, Hephaestus: Data analysis for x-ray absorption spectroscopy using IFEFFIT. J. Synchrotron Radiat. 2005, 12, 537-541. 39. Cline, J. D., Spectrophotometric determination of hydrogen sulfide in natural waters. Limnol. Oceanogr. 1969, 14, (3), 454-&. 40. Nagy, P.; Palinkas, Z.; Nagy, A.; Budai, B.; Toth, I.; Vasas, A., Chemical aspects of hydrogen sulfide measurements in physiological samples. Biochim. Biophys. Acta, Gen. Subj. 2014, 1840, (2), 876-891. 41. Gaitonde, M. K., A spectrophotometric method for direct determination of cysteine in presence of other naturally occurring amino acids. Biochem. J. 1967, 104, (2), 627-&. 42. http://www.eawag.ch/en/department/surf/projects/chemeql/ (accessed November 12, 2016). 43. Oguri, T.; Schneider, B.; Reitzer, L., Cysteine catabolism and cysteine desulfhydrase (CdsH/STM0458) in Salmonella enterica Serovar Typhimurium. J. Bacteriol. 2012, 194, (16), 4366-4376. 44. Kachur, A. V.; Koch, C. J.; Biaglow, J. E., Mechanism of copper-catalyzed autoxidation of cysteine. Free Radical Res. 1999, 31, (1), 23-34. 45. Laidlaw, I.; Steinmetz, M., Introduction to differential sedimentation. In Analytical ultracentrifugation, Scott, D. J.; Harding, S. E.; Rowe, A. J., Eds. 2005; pp 270-290. 46. Li, W. W.; Yu, H. Q., Insight into the roles of microbial extracellular polymer substances in metal biosorption. Bioresour. Technol. 2014, 160, 15-23. 47. Tourney, J.; Ngwenya, B. T., The role of bacterial extracellular polymeric substances in geomicrobiology. Chem. Geol. 2014, 386, 115-132. 48. Hahn, E.; Wild, P.; Hermanns, U.; Sebbel, P.; Glockshuber, R.; Haner, M.; Taschner, N.; Burkhard, P.; Aebi, U.; Muller, S. A., Exploring the 3D molecular architecture of Escherichia coli type 1 pili. J. Mol. Biol. 2002, 323, (5), 845-857. 49. Forsberg, C. W., Sulfide production from cysteine by Desulfovibrio-desulfuricans. Appl. Environ. Microbiol. 1980, 39, (2), 453-455.


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50. Gilmour, C. C.; Elias, D. A.; Kucken, A. M.; Brown, S. D.; Palumbo, A. V.; Schadt, C. W.; Wall, J. D., Sulfate-reducing bacterium Desulfovibrio desulfuricans ND132 as a model for understanding bacterial mercury methylation. Appl. Environ. Microbiol. 2011, 77, (12), 39383951. 51. Lin, H.; Morrell-Falvey, J. L.; Rao, B.; Liang, L. Y.; Gu, B. H., Coupled mercury-cell sorption, reduction, and oxidation on methylmercury production by Geobacter sulfurreducens PCA. Environ. Sci. Technol. 2014, 48, (20), 11969-11976. 52. Joshi, N.; Ngwenya, B. T.; French, C. E., Enhanced resistance to nanoparticle toxicity is conferred by overproduction of extracellular polymeric substances. J. Hazard. Mater. 2012, 241, 363-370. 53. Kumar, A.; Pandey, A. K.; Singh, S. S.; Shanker, R.; Dhawan, A., Cellular uptake and mutagenic potential of metal oxide nanoparticles in bacterial cells. Chemosphere 2011, 83, (8), 1124-1132. 54. Ionescu, M.; Belkin, S., Overproduction of exopolysaccharides by an Escherichia coli K12 rpoS mutant in response to osmotic stress. Appl. Environ. Microbiol. 2009, 75, (2), 483-492. 55. Minogue, T. D.; Daligault, H. A.; Davenport, K. W.; Bishop-Lilly, K. A.; Broomall, S. M.; Bruce, D. C.; Chain, P. S.; Chertkov, O.; Coyne, S. R.; Freitas, T.; Frey, K. G.; Gibbons, H. S.; Jaissle, J.; Redden, C. L.; Rosenzweig, C. N.; Xu, Y.; Johnson, S. L., Complete genome assembly of Escherichia coli ATCC 25922, a serotype O6 reference strain. Genome Announcements 2014, 2, (5), e00969-14. 56. Marvasi, M.; Visscher, P. T.; Martinez, L. C., Exopolymeric substances (EPS) from Bacillus subtilis: Polymers and genes encoding their synthesis. FEMS Microbiol. Lett. 2010, 313, (1), 1-9. 57. Aguilar, C.; Vlamakis, H.; Losick, R.; Kolter, R., Thinking about Bacillus subtilis as a multicellular organism. Curr. Opin. Microbiol. 2007, 10, (6), 638-643.


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40 30

C

B. subtilis

a

20

40

20

20

30

G. sulfurreducens

a

b

0 nM Hg 0 µM Cys

c c

c

c c

c

c

0

c c c c

e e e e e

50 nM Hg 500 nM Hg 5 µM Hg 100 µM Cys 1000 µM Cys 1000 µM Cys

0 hr

heat-treated 0 hr

0 nM Hg 0 µM Cys

0.5 hr

de de

cd

cd e

bc de

de

0

10

10

b

10

b

0

737 738 739 740

B

50

a

a

40

E. coli

50

A

30

50

Figures

Sulfide (µM)

734 735 736


50 nM Hg 100 µM Cys

500 nM Hg 1000 µM Cys

3 hr

bc d d d d

cd d

0 nM Hg 0 µM Cys

50 nM Hg 500 nM Hg 100 µM Cys 1000 µM Cys

d

d

d

heat-treated 3 hr

Figure 1: The concentration of sulfide measured in the exposure medium after exposure of (A) E. coli, (B) G. sulfurreducens, and (C) B. subtilis to various concentrations of total Hg(II) and cysteine for different exposure times. Cells in exponential growth phase with and without heat-treatment (3 hours incubation at 60 oC) to halt metabolic activity were tested. The letters in the plots are results of Tukey’s honest significant difference test (p < 0.05) and can be compared among results within each bacterial species. The time point of 0.5 hr was only tested for G. sulfurreducens.

741 742

743 744 745 746 747 748

Figure 2: High-angle annular dark field STEM images (A,C,D,E,F,G) and EDS maps (B,H) of bacterial cells and HgS(s) nanoparticles (bright white spots and areas) in (A,B) 200 – 300 nm sliced sections of G. sulfurreducens, (C) whole cells of E. coli, (D,E) 200 – 300 nm sliced sections of E. coli, and (F,G,H) 200 – 300 nm sliced sections of B. subtilis. All samples imaged were initially exposed to 500 nM Hg with 1000 µM cysteine except (E) where cells were exposed to 5 µM Hg with 1000 µM cysteine.


29

B

C

α-HgS

2-coordinate Hg-S

C ys

(3

hr )

hr )

hr )

µM 0

10 0 +

H g

g 50 0

nM

H nM 50

(3

nM

H g

C ys µM

0 +

50 0

H g nM 50

0 10 0 +

H g nM 50 0

(3

(3

hr )

hr )

hr )

0

20

40

60

80

100

B. subtilis

(3

(3

C ys

(3 nM

H g

C ys µM

50 0

H g nM 10

0

50

+ g H nM 50

β-HgS

µM

hr ) (3

hr )

C ys µM

0

hr )

0

20

40

60

80

100

G. sulfurreducens

(3

(1 h

(3

ys C µM 10 0 +

µM

H g

H g µM 5

r)

hr )

hr ) H g +

10 00

5

µM

su lfi d

30 + H g

+ H g

µM 5

Hg(SR)2

5

µM

µM

su lfi d

e

e

(3

(3

(3

hr )

hr )

hr ) 30

10 0 nM 50 0

*5 00

nM

H g

H g

+

+

10 0

0

0

µM

µM

C ys

C ys

(1

(3

(1

H g nM

*5 00 50 0

nM

H nM 0

hr )

hr )

hr ) H g

nM

(0 .5 50 0

nM 50 0

µM 0 10

g

+

H g

C ys

H g nM *5 0

(3

(3

hr )

hr )

0

20

40

60

80

100

E. coli

Page 30 of 32

10

A

*5

749 750 751 752 753 754 755 756 757 758 759 760 761 762 763 764 765 766 767 768 769 770 771 772 773 774 775 776 777 778 779 780 781 782 783 784 785 786 787 788 789 790 791 792 793 794

Cell-associated Hg species (%)


Unidentified

4-coordinate Hg-S

Figure 3: Hg(II) speciation determined from LCFs of the Hg LIII-edge EXAFS in whole cells of (A) E. coli, (B) G. sulfurreducens, and (C) B. subtilis exposed to varying concentrations of total Hg(II), cysteine, and sulfide for different exposure times. The Hg species consist of Hg bound to 2 sulfurs (2-coordinate Hg-S) and Hg bound to 4 sulfurs (4-coordinate Hg-S), which are conclusively identified as β-HgS, α-HgS, or Hg(SR)2 in some samples. In other samples, the 4-coordinate Hg-S species were not able to be identified and could be β-HgS, Hg(SR)4, Hg-S clusters, or a combination of all three. Samples marked with asterisks (*) were reported in our previous study (Thomas & Gaillard, 2017).


30

Page 31 of 32

40

Measured Hg (nM)

400

B

60

20

300

0 0

25

50

75

200

100

(As portion of added Hg)

Measured Hg (nM)

(As portion of added Hg)

A 500

E. coli

0 0

100

200

300

400

500

400

60 40 20

300

0 0

B. subtilis

0 0

cell-bound

20

HgS(s) precipitation 10

E. coli 0

10

20

30

40

50

D Total dissolved Hg (nM)

[sulfide]T = 43.3 µM

0

75

100

200

300

400

500

Added Hg (nM) with 1000 µM Cys

[cysteine]Red = 750 µM

30

50

100

500

total

C

25

200

Added Hg (nM) with 1000 µM Cys

Total dissolved Hg (nM)

795 796 797 798 799 800 801 802 803 804 805 806 807 808 809 810 811 812 813 814 815 816 817 818 819 820 821 822 823 824 825 826 827 828 829 830 831 832 833 834 835 836 837 838 839 840


intracellular

[cysteine]Red = 920 µM

30

[sulfide]T = 20.6 µM

20

HgS(s) precipitation

10

B. subtilis

0 0

Total Hg (nM)

10

20

30

40

50

Total Hg (nM)

Figure 4: The total recoverable Hg(II) (dissolved + cell envelope + intracellular), cell-bound Hg(II) (cell envelope + intracellular), and intracellular Hg(II) – as determined by an EDTA/GSH wash – detected after a 3-hour exposure of (A) E. coli and (B) B. subtilis to varying total Hg(II) in the presence of 1000 µM cysteine. The cell density of E. coli and B. subtilis was ~2 × 108 cells per mL and ~5 × 107 cells per mL, respectively. The calculated total dissolved Hg(II) as a function of total added Hg(II) in the exposure medium containing the cysteine and sulfide concentrations detected after a 3-hour exposure of (C) E. coli and (D) B. subtilis to 1000 µM cysteine. The calculations were performed without considering the potential for Hg binding to the bacterial surface.


31


841

Page 32 of 32

TOC Non-sulfate reducing conditions HgS(s)

Bio-mediated formation

XANES Derivative

Hg(cysteine)2(aq)

β-HgS Bacteria pellet

2 µm 12275

12300

12325

12350

Energy (eV)

842

HgS(s) interaction with bacterial cells


32

Spectroscopic and Microscopic Evidence of Biomediated HgS

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