Microbial generation of elemental mercury from dissolved methylmercury in seawater

Abstract

Elemental mercury (Hg⁰) formation from other mercury species in seawater results from photoreduction and microbial activity, leading to possible evasion from seawater to overlying air. Microbial conversion of monomethylmercury (MeHg) to Hg0 in seawater remains unquantified. A rapid radioassay method was developed using gamma-emitting ²⁰³Hg as a tracer to evaluate Hg⁰ production from Hg(II) and MeHg in the low pM range. Bacterioplankton assemblages in Atlantic surface seawater and Long Island Sound water were found to rapidly produce Hg⁰, with production rate constants being directly related to bacterial biomass and independent of dissolved Hg(II) and MeHg concentrations. About 32% of Hg(II) and 19% of MeHg were converted to Hg⁰ in 4 d in Atlantic surface seawater containing low bacterial biomass, and in Long Island Sound water with higher bacterial biomass, 54% of Hg(II) and 8% of MeHg were transformed to Hg⁰. Decreasing temperatures from 24°C to 4°C reduced Hg⁰ production rates cell⁻¹ from Hg(II) 3.3 times as much as from a MeHg source. Because Hg⁰ production rates were linearly related to microbial biomass and temperature, and microbial mercuric reductase was detected in our field samples, we inferred that microbial metabolic activities and enzymatic reactions primarily govern Hg⁰ formation in subsurface waters where light penetration is diminished.

Introduction

Elemental Hg (Hg⁰) can be found at all depths in the ocean and is usually supersaturated in surface waters (Mason and Fitzgerald 1993), resulting in its subsequent evasion from the sea surface to the overlying air. Several studies have demonstrated the significance of Hg⁰ evasion from the surface ocean (Fitzgerald et al. 1984; Kim and Fitzgerald 1986; Mason et al. 1994; Mason and Sheu 2002; Rolfhus and Fitzgerald 2001). The sea-air exchange of Hg⁰ is rapid and is associated directly with Hg⁰ concentrations in surface waters. Therefore, the transformation of other Hg species to Hg⁰ is an important process in marine Hg cycling. In the mixed layer, Hg⁰ can be produced by biological and/or photochemical reduction of reactive Hg(II) (Amyot et al. 1997; Amyot et al. 1994; Rolfhus and Fitzgerald 2004), leading to decreased Hg(II) concentrations in the water column and therefore avoiding further methylation of Hg(II). Moreover, monomethylmercury (MeHg) can also be converted to Hg⁰ through biotic and/or abiotic demethylation processes in soils and lakes (Oremland et al. 1991; Sellers et al. 1996). Both Hg(II) and MeHg are toxic, and the latter is bioaccumulative in aquatic organisms and can build up in aquatic food chains (Lawson and Mason 1998; Mason et al. 1996). Hence, there is interest in processes that influence mercury speciation in natural waters.

Hg⁰ formation in the surface mixed layer can involve a variety of complex abiotic and biotic processes. Abiotic processes such as photochemical reduction of Hg(II) has been shown to produce Hg⁰, where solar radiation (visible and ultraviolet), dissolved organic matter, and inorganic free radicals are considered key factors involved in the photochemical formation of Hg⁰ from Hg(II) (Amyot et al. 1997; Costa and Liss 1999; Costa and Liss 2000; Nriagu 1994; Vost et al. 2012). In addition, photodegradation of MeHg can also produce Hg⁰ (Sellers et al. 1996; Suda et al. 1993).

Biologically mediated processes like microbial Hg(II) reduction and MeHg demethylation have also been shown to form Hg⁰ (Barkay et al. 2003; Barkay et al. 1991; Robinson and Tuovinen 1984). These processes can be attributable to prokaryotes possessing the Hg-resistant mer operon at a relatively high Hg exposure condition (Morel et al. 1998; Yu et al. 1996). For example, the microbial mercuric reductase enzyme, merA governs the conversion of dissolved Hg(II) to volatile Hg⁰. Microorganisms with enzymes of Hg-resistance are widespread from diverse environments (Barkay et al. 1989; Gionfriddo et al. 2016; Osborn et al. 1997; Vetriani et al. 2005). Whether the transcription of the mer operon can be induced at natural Hg concentrations (pico- or femtomolar range) is not fully understood. Poulain et al. (2007) reported that merA genes were present and expressed by microbes from remote polar waters. However, some studies suggested Hg⁰ formation can occur by other uncharacterized microbial processes (Barkay et al. 2003; Kuss et al. 2015; Oremland et al. 1991). Recent studies further suggest that methanotrophs may play a role in controlling Hg transformation such as MeHg degradation (Lu et al. 2017; Vorobev et al. 2013). Hg(II) reduction can also be carried out by eukaryotic phytoplankton (Ben-Bassat and Mayer 1977; Mason et al. 1995), but the mechanism of this pathway is not well-understood or is often overlooked. Recent studies showed that phototrophic microorganisms (bacteria and algae) can use Hg(II) as an electron sink to maintain intracellular redox homeostasis, producing Hg⁰ and therefore modifying the availability of Hg to methylation sites (Grégoire and Poulain 2016; Grégoire and Poulain 2014). Also, intracellular photochemical degradation and volatilization of Hg(II) and MeHg in a marine microalga was recently reported (Kritee et al. 2017). Overall, microorganisms, especially bacteria, play a fundamental role in Hg(II) reduction and MeHg degradation to form Hg⁰.

In the surface mixed layer, the relative importance of abiotic (photochemical) and biotic (microbial) processes on Hg transformation remains uncertain. Over the past decade, mass-independent fractionation (MIF) and mass-dependent fractionation (MDF) of mercury isotopes have been developed to investigate photochemical reactions of Hg species in various environments (Bergquist and Blum 2007; Bergquist and Blum 2009). The distinct MIF signature caused by photochemical processes allows determination of the degree of photoreduction of Hg(II) (Zheng and Hintelmann 2009) and photodegradation of MeHg in natural waters (Point et al. 2011; Tsui et al. 2013). Although microbial transformation of Hg (mer mediated) exhibits only the MDF signature, the Hg isotope ratios can still serve as a tool in determining the role of microbial Hg(II) reduction (Kritee et al. 2008; Kritee et al. 2007) and MeHg degradation (Kritee et al. 2009) in the environment.

To further explore the roles that microbial activity play in Hg⁰ product, here we demonstrate how marine bacterioplankton convert MeHg to Hg⁰ at prevailing ocean temperatures. We focused on Hg⁰ formation mediated by natural assemblages of marine microorganisms from Atlantic Ocean surface seawater collected off Southampton, NY (SSW) and Long Island Sound surface water (LIW) collected off Stony Brook, NY exposed to picomolar Hg concentrations. Studies quantifying the kinetics of Hg⁰ transformation from MeHg are few, and our study investigates the kinetics of Hg⁰ formation from MeHg as well as Hg(II) mediated by natural occurring marine bacterioplankton. We developed a device to determine the kinetics of Hg⁰ formation from either dissolved Hg(II) or MeHg and used ²⁰³Hg as a gamma-emitting radiotracer of Hg⁰ gas formation. We further investigated how temperature and microbial abundance and metabolic activity influence Hg⁰ production in seawater.

Materials and methods

²⁰³Hg preparation and Me²⁰³Hg synthesis

The gamma-emitting radioisotope, ²⁰³Hg (half-life = 46.6 d) was used in this study to trace the transformation of Hg species between dissolved and gaseous phases. Previously, this isotope was used in experiments to quantify bioaccumulation and trophic transfer of mercury in marine plankton assemblages exposed to environmentally realistic mercury concentrations (Lee and Fisher 2016; 2017). A solution of ²⁰³Hg(II) in 1 M HCl was obtained from Eckert and Ziegler Isotope Products (Valencia, California) with a specific activity of 185 GBq g⁻¹ and was further converted to methyl-²⁰³Hg (Me²⁰³Hg) in the lab following established methods (Lee and Fisher 2016; Rouleau and Block 1997). In brief, ²⁰³Hg(II) solution was mixed with methylcobalamin (C₆₃H₉₁CoN₁₃O₁₄P) and acetate buffer at pH 5. Allowing the reaction to proceed in the dark for 18 to 24 h, Me²⁰³Hg was formed spontaneously. Following extraction by dichloromethane (CH₂Cl₂) and purification processes, Me²⁰³Hg was re-dissolved in Milli-Q^® water and was ready to use. The conversion yield (fraction of ²⁰³Hg recovered as Me²⁰³Hg) was 98 ± 2% (n = 3). Activities of ²⁰³Hg and Me²⁰³Hg used in this study ranged from 0.019 kBq L⁻¹ to 2.23 kBq L⁻¹, corresponding to concentrations of 1 to 100 pM of total (stable plus radioactive) Hg. Radiotracer ²⁰³Hg can provide precise, rapid and direct measurement of Hg⁰ formation in seawater, avoiding lengthy analytical procedures and potential contamination.

Natural bacterioplankton assemblages

The natural microbial assemblages used in this study were collected from Southampton, NY (8 km offshore, Lat. 40.77°N, Long. 72.43°W, 35 psu) and Stony Brook, NY (Lat. 40.92°N, Long. 73.15°W, 30 psu). Southampton seawater (SSW, referring to Atlantic surface water) was collected from the upper 10 m and was coarsely filtered (~100 μm) 6 years previously, stored at 4°C in the dark, and brought to room temperature (22°C) 2 d before experiments. Long Island Sound surface water (LIW) was freshly collected during high tide one day prior to experiments. Background concentrations of Hg(II) were ~5 pM, and MeHg was close to the detection limit (~5 fM) in these two waters, as determined using Tekran 2600 and 2700 mercury analyzers, respectively (Munson et al. 2014). Seawater used for experiments was filtered through sterile 1 μm polycarbonate membranes to remove all large particles including phytoplankton and zooplankton, leaving only bacterioplankton and similarly sized (or smaller) particles in the seawater. In general, SSW and LIW contain microbial densities of about 0.2–0.4×10⁶ cells mL⁻¹ and 4–5×10⁶ cells mL⁻¹, respectively. We used SSW_{1 μm} and LIW_{1 μm} to indicate 1 μm-filtered seawater with bacteria, and SSW_ctrl and LIW_ctrl to represent experimental controls, sterile-filtered (0.2 μm) or autoclaved seawater (see below).

General approach

Experimental setup

Figure 1 shows a schematic depiction of the experimental apparatus used in this study. All the experiments were performed in 250 mL Pyrex^® Erlenmeyer flasks with Teflon-lined screw caps (containing two holes drilled through the caps for an air inlet and outlet). 200 mL of seawater with its naturally occurring bacteria was injected with microliter quantities of ²⁰³Hg or Me²⁰³Hg solutions (dissolved in 1 M HCl and Milli-Q^® water, respectively) with no equilibration period, yielding concentrations ranging from 1 to 100 pM. Flasks were incubated at a constant temperature (24 or 4°C) under a 14 h:10 h light/dark cycle provided by cool white fluorescent lamps. Seawater was continuously bubbled gently with particle-free and Hg-free air and the produced gaseous ²⁰³Hg species was then collected in a Hg trap through the air outlet. The Hg trap was made of a quartz tube, glass wool and gold-coated beads purchased from Brooks Rand Instruments (Seattle, Washington) (Fig. 1). Gold traps have been used previously to capture volatile mercury species (Fitzgerald and Gill 1979; Fitzgerald et al. 2007); the quantity of Hg released from these cultures was far less than the binding capacity of the gold traps. Radioactivity of ²⁰³Hg in the Hg traps from each culture was directly determined with an LKB Wallac 1282 Compugamma NaI(Tl) gamma detector at 279 keV. All samples were counted with standards and decay-corrected. Propagated counting errors were <5%. From measured activities on traps, the percentage of Hg species transformed to Hg⁰ gas were calculated as:

$$\%{\text{Hg}}^0\text{\hspace{0.17em}evolved=}\frac{{\left[{\text{Bq}}^{\text{203}}\text{Hg}\right]}_{\text{trap}}}{{\left[{\text{Bq}}^{\text{203}}\text{Hg}\right]}_{\text{added}}}$$

[²⁰³Hg]_added was measured at the beginning of the experiment (t = 0) to determine variation in added amounts. Radioactivity was converted to molar concentrations of Hg using the specific activity of the isotope at the time of measurement. The rate of Hg⁰ formation attributed to microbial activity were calculated as:

$$\frac{{\left[{\text{Hg}}^0\text{evolved}\right]}_{\text{bacteria}}-{\left[{\text{Hg}}^0\text{evolved}\right]}_{\text{control}}}{{\left[\text{Hg}\left(\text{II}\right)\text{\hspace{0.17em}or\hspace{0.17em}MeHg}\right]}_{\text{dissolved}}\times \text{Time}}$$

Assuming microbial Hg(II) reduction and MeHg demethylation as irreversible pseudo-first order kinetics, the following equations apply:

$$\frac{d{\left[\text{Hg}\left(\text{II}\right)\text{\hspace{0.17em}or\hspace{0.17em}MeHg}\right]}_{\text{dissolved}}}{dt}=-k_r{\left[\text{Hg}\left(\text{II}\right)\text{\hspace{0.17em}or\hspace{0.17em}MeHg}\right]}_{\text{dissolved}}=\frac{d{\left[{\text{Hg}}^0\right]}_{\text{evolved}}}{dt}$$

where k_r is the pseudo-first order rate constant for gross reactions under the experimental conditions. Reversible reactions such as Hg⁰ oxidation and Hg(II) methylation were neglected because with the continuous purging used in our experiments the Hg⁰ concentrations would be negligible and these reactions would be unlikely to occur.

Figure 1.

Schematic layout of the incubation set-up and the Hg trap.

Of the various Hg species, the Henry’s law coefficients of Hg⁰ and dimethylmercury (DMHg) indicate they are highly volatile and could volatilize from the dissolved phase at ambient temperatures. Recent studies have shown the potential abiotic pathways of DMHg formation from MeHg mediated by reduced sulfur groups on mineral or organic surfaces (Jonsson et al. 2016), and in sulfidic aqueous solutions (Kanzler et al. 2018). To verify whether DMHg was produced in our experiments as well as distinguish Hg⁰ from DMHg in the gas traps, we used another Hg trap, Bond Elut ENV (Agilent Tech., Santa Clara, California), which contains styrene-divinylbenzene polymeric resins and effectively collects volatile organomercury compounds (Baya et al. 2013). In preliminary experiments, produced gaseous Hg was first passed through a Bond Elut trap followed by a gold trap without exceeding the reported breakthrough volume of the Bond Elut trap. We found very little Hg retained in the Bond Elut trap and >95% of the Hg was collected in the gold trap. The Bond Elut trap can also collect a small fraction of Hg⁰ (Baya et al. 2013). Thus, we concluded that Hg⁰ was the dominant gaseous species produced in our study and Bond Elut traps were used in selected treatments to confirm no DMHg formation.

Short-term incubation experiments

We first performed a short term (1 d) incubation experiment to test the influence of Hg concentrations and microbial densities on Hg gas production. For experiments assessing different Hg concentrations, triplicate samples of 200 mL SSW with the same initial microbial density were injected with ²⁰³Hg(II) or Me²⁰³Hg at 1, 10, or 100 pM and then incubated for 24 h at 24°C under a 14:10 light/dark cycle. For experiments assessing the influence of initial microbial density, duplicate samples of 200 mL SSW with varying initial microbial densities (by mixing with 0.2 μm filtered SSW) were prepared, with the addition of 10 pM of ²⁰³Hg(II) or Me²⁰³Hg in each treatment and incubated for 24 h at 24°C. After 24 h with continuous purging, Hg traps and 1 mL of unfiltered seawater were collected for radioanalysis, and the microbial densities were measured.

Long-term incubation experiments

A series of long-term (4 d) incubation experiments were carried out in order to evaluate the rates at which Hg gas evolved from each flask to overlying air. First, we determined the Hg gas production from three different types of controls, 0.2 μm sterile-filtered SSW (removing bacteria detectable with DAPI staining), autoclaved SSW (120°C for 30 min), and SSW with antibiotics (1% GE HyClone™ Penicillin-Streptomycin). Second, we examined how temperature (24°C vs. 4°C) and light (light vs. dark) influenced the Hg gas evolved from SSW. Third, we examined the Hg gas production in freshly collected LIW, as well as the difference between unfiltered LIW and 1 μm-filtered LIW.

Experimental procedures for long-term incubations were similar to those described in short-term experiments. Briefly, 200 mL seawater was injected with ²⁰³Hg(II) or Me²⁰³Hg at concentrations from 40 to 80 pM and incubated at either 4°C or 24°C with or without light. The Hg trap was collected and replaced with a new one at each sample time (4, 8, 12, 24, 36, 48, 72, and 96 h). In addition, 1 mL of unfiltered seawater was collected for microbial density determination at each sample time. Control treatments (seawater without bacteria) containing equivalent amounts of ²⁰³Hg were performed simultaneously. To calculate the Hg mass balance ([Hg]_added = [Hg]_suspension + [Hg]_trap + [Hg]_{flask wall}, suspension = particulate + dissolved) in each flask, sorption of Hg(II) or MeHg onto culture flasks was examined by acid washing (1 M HCl) the flask walls at the end of the experiment. Across all experiments, the mass balance of Hg(II) and MeHg treatments were 94 ± 10% and 97 ± 2%, respectively.

Microbial density and microbial production

A modified DAPI (4’,6-diamidino-2-phenylindole) staining method (Sherr et al. 2001) was used to count microbial densities. Microbial cell samples were fixed with 2% glutaraldehyde solution and were stored at 4°C. Prior to cell counts, cell samples were mixed with DAPI (1 μg mL⁻¹) and stained cells were then filtered onto a 0.2 μm black polycarbonate membrane at a vacuum pressure <300 mm Hg. Cell counts were obtained with the use of epifluorescence on an inverted microscope under ultraviolet light excitation. For each sample, the number of luminant dots (bacteria) in a whole grid for 10 randomly chosen fields over the membrane was counted and averaged. The average microbial density in each samples (cells mL⁻¹) were calculated from the equation:

$$\left[\frac{\left(\text{average\hspace{0.17em}cells\hspace{0.17em}per\hspace{0.17em}grid\hspace{0.17em}}-\text{\hspace{0.17em}background\hspace{0.17em}cells\hspace{0.17em}per\hspace{0.17em}grid}\right)\times \text{grids\hspace{0.17em}per\hspace{0.17em}filter}}{\text{volume\hspace{0.17em}of\hspace{0.17em}sample}}\right]$$

where background refers to 0.2 μm-filtered seawater subjected to the same DAPI-staining procedure. Microbial production measurements were performed as a parallel experiment to the Hg⁰ formation experiments, using leucine incorporation method (Kirchman 1993). Radiolabeled L-[4,5-³H]-leucine was purchased from PerkinElmer, Inc. In brief, 20 mL of unfiltered seawater was collected in an incubation vial at each time point (6, 12, 24, 48, 72, and 96 h), injected with ³H-leucine to yield a concentration of 20 nM (corresponding to 24.8 kBq per vial) and incubated for 6 h. Incubation was terminated by adding trichloroacetic acid (TCA) at a concentration of 5%. After heating for 20 min at 80°C, microbial cells were collected onto a 0.22 μm cellulose acetate filter at a vacuum pressure <300 mm Hg. The filter with cells was dissolved by adding ethyl acetate in a scintillation vial, mixed with Ultima Gold™ XR liquid scintillation cocktail. Beta radioactivity was determined using a PerkinElmer Tri-Carb^® 2810 TR liquid scintillation analyzer. Because preliminary experiments indicated negligible microbial growth at 4°C, no microbial production measurements were conducted at this temperature.

Conventional PCR analysis for mercuric reductase gene merA

Total DNA was extracted from seawater, using a PowerWater^® Sterivex™ DNA isolation kit (MO BIO, Carlsbad, CA). About 10 L of SSW and 2 L of LIW were filtered to collect enough DNA concentrations for analysis and merA was determined following the procedure of Poulain et al. (2015). PCR was performed using a Biometra^® T3000 Thermocycler. PCR conditions consisted of a 10-min incubation at 95 °C, followed by 35 cycles of 95°C for 30 s, 63°C for 15 s, 72°C for 30 s, and a final extension step at 72°C for 5 min. Final concentrations of the PCR mixture were 1×PCR buffer, 2.5 mM MgCl₂, 0.2 μM forward/reverse primers (merA2 F: CCTGCGTCAACGTCGGCTG, merA2 R: GCGATCAGGCAGCGGTCGAA), 0.2 mM dNTPs, and 1 U Taq Polymerase. Samples were analyzed with a positive control (Pseudomonas aeruginosa containing the Hg resistance plasmid pVS1) (Lloyd et al. 2016), and a negative control (reagents only). Amplicons were separated by electrophoresis and visualized under ultraviolet light. Amplification of PCR product at the expected amplicon size (~300 bp) was used to determine the presence of merA in our samples.

Results

Effects of Hg concentrations and microbial biomass

The total percentage of Hg⁰ gas evolved after 24 h as a function of dissolved Hg species concentrations and average microbial density is shown in Fig. 2. The %Hg⁰ evolved from each culture stayed almost constant across three different Hg concentrations (1 to 100 pM) at ~20% for Hg(II) treatments and ~5% for MeHg treatments (Fig. 2a). In these cultures, the initial microbial densities were 0.34–0.40×10⁶ cells mL⁻¹, and the microbial densities at 24 h were 0.81–1.2×10⁶ cells mL⁻¹. In separate experiments which used cultures with varying microbial cell densities and 10 pM of Hg(II) or MeHg, the %Hg⁰ that evolved as a gas ranged from 6.2% to 31% in Hg(II) treatments and from 2.1% to 13% in MeHg treatments depending on the microbial abundance (Fig. 2b); Hg⁰ released was positively correlated with microbial density. We performed each experiment twice and had consistent results. At the same microbial density, Hg⁰ production was always higher from Hg(II) treatments than from MeHg treatments.

Figure 2.

Total percentage of ²⁰³Hg⁰ evolved from ²⁰³Hg(II)- and Me²⁰³Hg-treated SSW (Southampton seawater, 1 μm-filtered) after 1-day incubation as a function of (a) dissolved Hg(II) or MeHg concentrations and (b) mean microbial cell density. Data points are the means from two or three replicate cultures with error bars of one standard deviation. Added concentrations of Hg(II) and MeHg were 10 pM in (b). Note that background concentrations of Hg(II) and MeHg in SSW were ~5 pM and ~5 fM, respectively.

Long term incubations

Background Hg⁰ production in SSW

The Hg⁰ formation kinetics in three different control treatments are shown in Fig. 3. Generally, in Hg(II) treatments (Fig. 3a), 0.2 μm sterile-filtered SSW and autoclaved SSW had very similar patterns, rapid Hg⁰ formation in the first 12 h followed by very slow formation, with an overall 5–9% of Hg⁰ evolved. Controls with antibiotics showed almost no Hg⁰ production in the first 24 h, but increased Hg⁰ formation was observed after 36 h (Fig. 3a). In MeHg treatments (Fig. 3b), all three types of controls showed very low Hg⁰ formation over 72 h (overall < 1%), suggesting MeHg was very stable in seawater without bacteria. For all other experiments, we used 0.2 μm sterile-filtered seawater (SSW or LIW) for controls.

Figure 3.

Total percentage of Hg⁰ evolved over time at 24°C from different controls (Southampton seawater without microbes) treated with (a) Hg(II) or (b) MeHg. Data points are the means from three replicate cultures with error bars of one standard deviation; error bars are often too small to see. Added concentrations of Hg(II) were 54 pM and of MeHg were 56 pM.

Effects of temperature and light

Figure 4 shows the Hg⁰ evolved from (a) Hg(II) and (b) MeHg treatments in SSW over 96 h at two different temperatures, and (c) their corresponding microbial cell densities. Rapid formation of Hg⁰ in Hg(II)-treated SSW with bacteria within 24 h was observed (Fig. 4a). Total %Hg⁰ evolved was 32.6 ± 1.6% at 24°C after 96 h, but at 4°C it was only 8.0 ± 2.3%; this compares with total %Hg gas produced in controls of 7.6 ± 3.5% at 24°C and 5.6 ± 1.4% at 4°C. Thus, net production in microbial communities over 96 h was about 25% at 24°C and 2.5% at 4°C. Total Hg⁰ evolved from MeHg-treated SSW with bacteria after 4 d was 18.9 ± 1.7% at 24°C and 10.2 ± 0.4% at 4°C, and both showed the same linear increasing patterns over time (Fig. 4b). Controls at two temperatures both showed very low Hg⁰ production after 96 h (< 0.4%) and no microbial growth. Microbial growth was more pronounced at 24°C (more than one doubling after 96 h) than at 4°C, where scant growth occurred (Fig. 4c). The results of microbial production at 24°C are shown in Fig. 4d. The microbial production, as inferred from leucine uptake, increased over time, ranging from 0.09 to 0.35 μg-C L⁻¹ h⁻¹. Normalized to microbial density, the estimated production rate ranged from 0.21 to 0.44 fg C cell⁻¹ h⁻¹; normalized to bacterial carbon (20 fg C cell⁻¹) (Lee and Fuhrman 1987) the microbial production increased at a rate of 1.1 to 2.2% h⁻¹. As the lower estimated value, which is the rate reflective of the initial production rate, closely matches the observed growth rate of the bacteria (increase in cell number day⁻¹) (Fig. 2c), it appears from the leucine incorporation rates that all the cells observed by DAPI were metabolically active.

Figure 4.

Total percentage of Hg⁰ evolved from (a) Hg(II)-treated SSW and (b) MeHg-treated SSW at two different temperatures, (c) microbial cell density, and (d) microbial production (24°C only) over time. Treatments with bacteria were 1 μm-filtered and controls were 0.2 μm sterile-filtered. Data points are the means from three replicate cultures with error bars of one standard deviation. Added concentrations of Hg(II) were 56 pM and of MeHg were 55 pM.

For both Hg(II) and MeHg treatments, there was no significant difference in Hg⁰ production between light and dark conditions (Fig. 5). Neither Hg⁰ formation in controls nor microbial growth were affected by light (data not shown).

Figure 5.

Total percentage of Hg⁰ evolved from Hg(II)-treated SSW and MeHg-treated SSW under light and dark conditions. Controls were sterile-filtered (0.2 μm) SSW. Data points are the means from two replicate cultures with error bars of one standard deviation. Added concentrations of Hg(II) were 45 pM and of MeHg were 70 pM.

Hg⁰ production in freshly collected Long Island Sound Seawater

The results of Hg⁰ evolved from freshly collected LIW are shown in Figure 6 (note that presented data contain unfiltered and 1 μm-filtered LIW). For Hg(II)-treated LIW, 70.5 ± 5.4%, 53.4 ± 3.1%, and 11.2 ± 1.3% of Hg⁰ evolved from LIW_unfiltered, LIW_{1 μm}, and LIW_ctrl after 96 h, respectively (Fig. 6a). For MeHg-treated LIW, 18.8 ± 0.6%, 7.9 ± 0.2%, and 0.4 ± 0.1% of Hg⁰ evolved from LIW_unfiltered, LIW_{1 μm}, and LIW_ctrl after 96 h, respectively (Fig. 6b). Microbial densities in LIW ranged from 4.3 to 7.1×10⁶ cells mL⁻¹ (Fig. 6c).

Figure 6.

Total percentage of Hg⁰ evolved from (a) Hg(II)-treated LIW and (b) MeHg-treated LIW, and (c) microbial cell density at 24°C over time. Data points are the means from two replicate cultures with error bars of one standard deviation; error bars are often too small to see. Added concentrations of Hg(II) were 74 pM and of MeHg were 67 pM. Controls were sterile-filtered (0.2 μm) LIW.

Hg⁰ production rate

After correcting for abiotic Hg⁰ production in controls, average Hg⁰ formation rates by bacteria in different treatments over 4 d of incubation are presented in Table 1. The Hg⁰ production rates by bacteria at different time intervals were also calculated (Fig. 7a–b, data from LIW_unfiltered not shown). For Hg(II)-treated seawater, the production rate was high at the first day (0–24 h) and then declined afterward (Fig. 7a). Notably, a significant decline after 1 day can be found in SSW (both 24°C and 4°C treatments). In contrast, a significant decline can be found after 3 days in LIW. For MeHg-treated seawater, the production rate gradually increased over time (Fig. 7b). The calculated Hg⁰ production rate in Fig. 7a–b could have been biased by differences in the microbial biomass among treatments. Therefore, we further normalized the Hg⁰ production rate to microbial density (Fig. 7c–d).

Table 1.

Production of Hg⁰ by marine bacteria in SSW and LIW in 4-d incubation experiments. All production rates from treatments with bacteria are corrected for abiotic production (controls, 0.2 μm sterile-filtered). Mean bacterial cell density represents the average bacterial density of all time points. In controls, bacterial cell densities were below detection limit (BDL).

Seawater type	Incubation temp. (°C)	Added Hg species	Total Hg0 formation after 4 d (%)	Hg0 formation (% d−1)	Mean bacterial density (☓106 cells mL−1)	Hg0 formation (% d−1 ☓106 cells−1)	Hg0 formation (amol-Hg d−1 cells−1)	Rate constant (☓10−6 s−1)
SSW1μm	24	Hg(II)	25.1 ± 1.2	6.3 ± 0.3	0.86 ± 0.10	7.3 ± 0.9	0.82 ± 0.10	1.1
SSW1μm	24	MeHg	18.5 ± 1.7	4.6 ± 0.4	0.74 ± 0.09	6.2 ± 0.9	0.69 ± 0.11	0.61
SSWctrl	24	Hg(II)	7.6 ± 3.5	1.9 ± 0.9	BDL	-	-	0.14
SSWctrl	24	MeHg	0.5 ± 0.4	0.1 ± 0.1	BDL	-	-	<0.01
SSW1μm	4	Hg(II)	2.5 ± 2.7	0.6 ± 0.7	0.47 ± 0.07	1.3 ± 1.5	0.20 ±0.22	0.06
SSW1μm	4	MeHg	9.9 ± 0.4	2.5 ± 0.1	0.52 ± 0.11	4.8 ± 1.0	0.56 ± 0.12	0.30
SSWctrl	4	Hg(II)	5.6 ± 1.4	1.4 ± 0.4	BDL	-	-	0.17
SSWctrl	4	MeHg	0.2 ± 0.0	0.05 ± 0.01	BDL	-	-	<0.01
LIW1μm	24	Hg(II)	42.2 ± 3.1	10.5 ± 0.8	5.02 ± 0.58	2.1 ± 0.3	0.31 ± 0.04	1.3
LIW1μm	24	MeHg	7.6 ± 0.2	1.9 ± 0.1	5.26 ± 0.83	0.4 ± 0.1	0.05 ± 0.01	0.17
LIWunfiltered	24	Hg(II)	59.3 ± 5.4	14.8 ± 1.3	5.54 ± 0.39	2.7 ± 0.3	0.39 ± 0.05	2.2
LIWunfiltered	24	MeHg	18.4 ± 0.6	4.6 ± 0.2	5.44 ± 0.38	0.8 ± 0.1	0.12 ± 0.01	0.47
LIWctrl	24	Hg(II)	11.2 ± 1.3	2.8 ± 0.3	BDL	-	-	0.27
LIWctrl	24	MeHg	0.4 ± 0.1	0.09 ± 0.03	BDL	-	-	<0.01

Figure 7.

Hg⁰ production rate (corrected for controls) at different periods of time from (a) Hg(II)-treated seawater and (b) MeHg-treated seawater, and Hg⁰ production rate (corrected for controls) normalized to microbial cell density from (c) Hg(II)-treated seawater and (d) MeHg-treated seawater. All data shown here were for 1 μm-filtered seawater and bars represent means of two or three replicate cultures with error bars of one standard deviation.

Discussion

Effects of Hg concentration and microbial densities

Regardless of the concentrations of dissolved Hg(II) or MeHg, the percentage of the total that resulted in Hg⁰ formation in SSW_1μm stayed constant over a 1-day incubation, suggesting that the fraction of total Hg that was bound to microbial cells remained constant and reached equilibrium rapidly across three concentrations. Limited by the specific activity of ²⁰³Hg, the lowest Hg concentration used in this study was 1 pM, which was still higher than typical concentrations (fM to pM range) in nearly all marine ecosystems (especially for MeHg). Because there was no significant difference in Hg behavior across three concentrations (over 2 orders of magnitude), extrapolation of the experimental results to lower Hg concentrations may be valid.

Hg⁰ production was positively correlated with microbial density, consistent with observations of Rolfhus and Fitzgerald (2004) in coastal seawater. Other studies demonstrated that Hg⁰ production correlated with enzymatic activity (Barkay et al. 1991; Siciliano et al. 2002) and the biomass of planktonic organisms (Barkay et al. 1989; Mason et al. 1995), which also support the idea that biological activity contributes to Hg⁰ production. The y-intercepts of the regression lines observed in Fig. 2b represent the percentage of abiotic Hg⁰ formation in each treatment, ~9% for Hg(II) and ~1% for MeHg. These values are comparable to the results of background Hg gas formation described above in Results.

Abiotic Hg⁰ production from control treatments

The Hg⁰ formation in SSW controls that were 0.2 μm sterile-filtered or autoclaved was consistent with findings that reactive Hg(II) might be reduced abiotically in a very short time in the presence of reductants in seawater or on flask walls (Mason et al. 1995). For controls of 0.2 μm-filtered SSW with antibiotics, no rapid Hg(II) reduction was observed in the first 24 h, which might be attributable to complexation of Hg(II) by the antibiotics, thereby inhibiting abiotic Hg(II) reduction. However, the effectiveness of antibiotics lasted only one day as Hg⁰ formation increased up to 18% by 48 h, possibly due to degradation of antibiotics over time. No significant Hg⁰ production was found among all MeHg-treated controls of SSW or LIW, suggesting that MeHg was very stable in seawater with respect to reduction without no bacteria and/or particles larger than 0.2 μm.

Effects of temperature and light

Nearly the same amount of Hg⁰ was collected in controls between 24°C and 4°C, suggesting that continuous bubbling of air can sufficiently remove all produced Hg⁰ from the dissolved phase and therefore the influence of changes in Hg⁰ solubility due to varying temperatures was negligible. Note that this is within the normal annual range of water temperatures in Long Island waters. It was clear that more Hg⁰ was produced at 24°C than at 4°C in either Hg(II)- or MeHg-treated SSW_1μm. For Hg(II) treatments, total %Hg⁰ production in SSW_1μm was about four times higher at 24°C than at 4°C and the Q₁₀ (calculated over the period 0–96 hours) for the gas production rate process was 2.03 for Hg(II), consistent with a biologically mediated process. It is unlikely that abiotic reduction of Hg(II) occurring on microbial cell surfaces also contributed to the Hg gas production because Hg⁰ formation in SSW_ctrl at 4°C was close to SSW_1μm at 4°C where bacteria cells were present (Fig. 4a), suggesting that surface reduction only accounted for a small part of Hg⁰ formation. In contrast, for MeHg-exposed cells, the Q₁₀ of the Hg⁰ production calculated over the first 96 hours was only 1.11. The Q₁₀ values calculated over the first 24 hours for Hg(II) and MeHg were 2.09 and 1.38, respectively. There was a much sharper decline in Hg formation rates for Hg(II) than for MeHg after the first 24 hours of exposure, expressed either on a whole culture basis or normalized to a microbial cell basis (Table 1).

The likeliest mechanism of Hg⁰ formation from MeHg involves cleavage of the carbon-mercury bond by the organomercurial lyase, producing Hg(II), followed by the reduction of Hg(II) to Hg⁰ by mercuric reductase (Ullrich et al. 2001). The Hg⁰ production in MeHg-treated cultures at 4°C was similar to that of gross but greater than net production rates in Hg(II)-treated cultures at 4°C. Because gas formation in Hg(II)-treated seawater was much more influenced by temperature than in MeHg-treated seawater, it appears that the demethylation process might not be as sensitive to temperature as reduction of Hg(II) or, the gross demethylation process might be partially driven by an uncharacterized abiotic reaction which is not affected appreciably by temperature. Alternatively, abiotic MeHg degradation with thiols (Jonsson et al. 2016) and sulfides (Kanzler et al. 2018) to form DMHg has been shown to occur spontaneously. No sulfide was added in our treatments, but DMHg formation might be achieved if MeHg reacts with reduced sulfur groups on microbial cell surfaces. However, the tandem trap (Bond Elut ENV + gold beads) test showed that Hg⁰ was the dominant species and the DMHg formation, if any, was negligible. The higher microbial production at 24°C than at 4°C, as inferred from leucine uptake, coincided with increased microbial cell densities over time. The measured values of microbial production were comparable to reports for natural assemblages of bacteria in field studies (Anderson and Taylor 2001; Shiah and Ducklow 1994).

Photochemical reactions through radiation of ultraviolet and visible light are known to reduce Hg(II) (Amyot et al. 1994; Costa and Liss 2000; O’Driscoll et al. 2007) and degrade MeHg (DiMento and Mason 2017; Hammerschmidt and Fitzgerald 2006; Lehnherr and St. Louis 2009; Sellers et al. 1996; Suda et al. 1993), resulting in Hg⁰ formation. However, we did not observe significant differences in Hg⁰ evolution between light and dark treatments. The reason was probably due to our experimental settings where cool-white fluorescent light with very low ultraviolet radiation (Kritee et al. 2017) was used in our study. Moreover, its light intensity was less than one-twentieth of incident irradiation at full sunlight. Regardless of any light effects that may occur in nature, it was clear from the laboratory experiments that biologically mediated processes significantly enhanced the evasion of Hg from these cultures.

Unfiltered LIW vs. 1 μm-filtered LIW

Hg⁰ production in Hg(II)-treated seawater was higher in LIW_unfiltered than in LIW_1μm. Microbes smaller than 1 μm accounted for 71% of Hg(II) reduction in LIW, suggesting that these organisms are the primary Hg⁰ producers, consistent with the conclusion of Mason et al. (1995). Hg⁰ was also produced in MeHg-treated LIW_unfiltered to a greater extent than in LIW_1μm, but microbes smaller than 1 μm only accounted for 41% of MeHg degradation. Whether this discrepancy is attributable to differential distribution of organomercurial lyase (MerB), the enzyme that is responsible for MeHg degradation, in different sized organisms is not known. In contrast, there appears to be broad distribution of the gene (merA) coding for mercuric reductase (MerA) which is responsible for Hg(II) reduction to gaseous Hg⁰ and is the core enzyme of bacterial mercury resistance (Barkay et al. 2003).

Comparison of Hg⁰ formation between SSW and LIW

The greater production of Hg⁰ in Hg(II)-treated LIW_1μm than in Hg(II)-treated SSW_1μm was probably attributable to the greater microbial biomass in LIW. However, less Hg⁰ production was found in MeHg-treated LIW_1μm than in MeHg-treated SSW_1μm which cannot be explained by differences in microbial biomass or other factors like salinity, dissolved organic matter and nutrient concentrations. Further, the Hg⁰ production in LIW was significantly lower than from SSW when normalized to microbial biomass. Assuming Hg⁰ formation was mainly driven by microbial enzymatic processes, the discrepancy between SSW and LIW may have been a consequence of different microbial communities in these two waters, which in turn may have been partially attributable to the long storage of the SSW (6 years) compared with the freshly collected LIW. Regardless, the rapid production of Hg⁰ in SSW within 2 days of raising the temperature of this water from 4°C to 24°C was noteworthy.

The Hg-resistance gene, merA, is considered broadly distributed among bacteria and archaea probably due to horizontal gene transfer in evolution (Boyd and Barkay 2012; Osborn et al. 1997), but the prevalence of Hg-resistance among bacteria can vary from one location to another (Barkay et al. 2011). Further investigations using metagenomic analysis could elucidate the microbial community composition in SSW and LIW.

Bacterial mercuric reductase gene merA

There are two known MeHg degradation processes mediated by microbes, reductive demethylation by merAB yielding CH_4, and oxidative demethylation related to C1 metabolism in anaerobic prokaryotes yielding CO₂ and CH₄ (Barkay et al. 2011; Barkay et al. 2003). Studies suggest that the reductive pathway is dependent on intracellular Hg concentrations (typically found in more heavily impacted environments) in more aerobic conditions, while the oxidative demethylation is more prevalent in anaerobic environments (Barkay et al. 2003; Grégoire and Poulain 2014; Hintelmann 2010). In addition, recent studies show a novel biological pathway of MeHg demethylation by methanotrophs but no production of gaseous Hg⁰ is observed (Lu et al. 2017; Vorobev et al. 2013). Reductive demethylation is the likeliest process to have occurred in our experiments (assuming the demethylation was mediated by microbes), because the other processes are typically observed under low redox conditions which did not occur in our experiments. As noted above, a new pathway of Hg(II) reduction has been observed for phototrophs that use Hg(II) as an electron sink to maintain intracellular redox homeostasis and generate Hg⁰ (Grégoire and Poulain 2016), although this pathway has not yet been demonstrated for MeHg or specifically in heterotrophic bacterioplankton.

Following the procedure described by Poulain et al. (2015), merA genes in our positive control were successfully amplified at 300 bp (Fig. S1 in Supplementary). Moreover, we found amplification of PCR product at ~300 bp (Fig. S1), which was used to determine presence of merA in our samples. The results suggest that the Hg⁰ formation in our study may have been driven by bacterial mercuric reductase. It has been suggested that merA-mediated reduction is induced at high ambient Hg(II) concentrations of >50 pM (Morel et al. 1998); such a “threshold” concentration of Hg(II) is much higher than in most marine environments. Several field studies indeed showed detectable transcription of mer genes in highly contaminated environments (Nazaret et al. 1994; Schaefer et al. 2004). However, Poulain et al. (2007) reported detectable merA transcription in microbes in the Canadian Arctic where the ambient total Hg concentrations were about 10 pM, implying that bacteria may have continuous activity of Hg reductase even at natural, low Hg concentrations. Furthermore, they concluded that merA-mediated activity could account for up to 90% of Hg⁰ production at a greater depth in the water column. In fact, mer induction should not only depend on absolute Hg concentrations but also other factors that affect actual Hg bioavailability to bacteria (Barkay et al. 2011). Our results showed that bacteria still exhibited significant capacity to produce Hg⁰ at ambient concentrations as low as ~5 pM. However, we did not determine if merA transcription was induced in our samples during the incubation. Further work is needed to elucidate whether threshold concentrations of Hg are applicable to bacteria.

Production kinetics and rates

Mason et al. (1995) examined Hg⁰ formation in Hg(II)-treated seawater in diverse cultures of marine microorganisms (mostly phytoplankton, including cyanobacteria), and reported Hg⁰ production rates of 0.4–9.7% d⁻¹ (corresponding to 0.027–0.48 amol-Hg cell⁻¹ d⁻¹), which were comparable with our results (Table 1). However, it should be noted that their added Hg(II) concentrations were 0.5 nM which was about 10 times higher than ours. Monperrus et al. (2007) reported that microbial and/or phytoplanktonic reduction of Hg(II) in surface waters of the Mediterranean Sea ranged from 2.2–12.3% d⁻¹, which were also comparable to our study (Table 1).

The patterns of Hg⁰ evolved from Hg(II)-treated seawater (initially rapid formation rates followed by much slower formation rates) were similar to the results from Barkay et al. (1991), where Hg(II) volatilization by an indigenous freshwater microbial community was measured. The decline in production rate in Hg(II) treatments was probably because of a decline in bioavailable Hg(II) for the bacteria and/or reactive Hg(II) was converted to a non-reducible form by complexation with dissolved organic matter. The Hg⁰ production rates normalized to microbial abundance were significantly higher in SSW than in LIW, particularly within the first 48 h, possibly because the bacteria in SSW had a higher capacity to reduce Hg(II). The reaction of Hg(II) reduction from SSW_1μm over 96 h seemed to be a mixed order reaction. But treated as a first order reaction, the calculated rate constant was 1.1×10⁻⁶ s⁻¹. The reaction for LIW can perfectly fit the model of a first order reaction and the rate constants were 1.3×10⁻⁶ s⁻¹ for LIW_1μm and 2.2 × 10⁻⁶ s⁻¹ for LIW_unfiltered. These gross values are comparable to dark (or low light) gross values from incubations for the Baltic Sea (1.5 ± 0.5×10⁻⁶ s⁻¹) (Kuss et al. 2015) and Chesapeake Bay and coastal shelf waters (0.67–1.8×10⁻⁶ s⁻¹) (Whalin et al. 2007), but rate constants of microbial production can be one to two orders of magnitude lower than those attributable to photochemical transformation at the ocean surface (O’Driscoll et al. 2006; Qureshi et al. 2009; Vost et al. 2012; Whalin et al. 2007).

Very few studies reported microbial MeHg demethylation rates and kinetics in seawater. Our calculated Hg⁰ formation rates in MeHg-treated seawater (1.9–4.6% d⁻¹, Table 1) were comparable to Hg⁰ production rates (2.8–10.9% d⁻¹) reported by Monperrus et al. (2007) in the Mediterranean. The Hg⁰ evolved from MeHg-treated seawater was linear over time, implying a zero-order reaction with an independence of substrate (MeHg) concentrations. Unlike the Hg(II)-treated seawater, the Hg⁰ formation rates in MeHg treatments did not change much over time. The modest increase over time is probably due to microbial growth over time as indicated by the low variability in Hg⁰ production rate normalized to microbial densities. As noted earlier in the text, the formation of Hg⁰ from MeHg is a likely two-step reaction involving the demethylation of MeHg to Hg(II), and the subsequent reduction to Hg⁰. The demethylating process might be the rate limiting step, resulting in a linear increase of Hg⁰ formation which was very different from the Hg(II)-treated seawater. In order to compare with Hg(II) treatments and previous literature, here we also used first order kinetics to calculate the rate constants for Hg⁰ formation in MeHg-treated seawater. The calculated values were 0.61×10⁻⁶ s⁻¹ for SSW_1μm, 0.17×10⁻⁶ s⁻¹ for LIW_1μm, and 0.47×10⁻⁶ s⁻¹ for LIW_unfiltered. Whalin et al. (2007) reported that the rate constants of demethylation in the water column of Chesapeake Bay were <1–5×10⁻⁶ s⁻¹ where the demethylation was driven by both photochemical and microbial processes. In a lab study using coastal seawater, rate constants attributable solely to photochemistry were reported to range from 10 to 19×10⁻⁶ s⁻¹ (DiMento and Mason 2017). DiMento and Mason (2017) also demonstrated that photodegradation of MeHg decreases exponentially with depth, with degradation rates being <10% of those in surface waters at depths <5 m in estuaries, and 60 m in open ocean waters. Thus, biologically mediated conversion of MeHg would be expected to be greater than photochemical degradation of MeHg in subsurface waters, the absolute depth depending on light penetration in different water columns.

The distribution of Hg species in microbial cells may influence the kinetics of Hg(II) reduction and MeHg degradation. It has been shown that more MeHg can penetrate into the cytoplasm of phytoplankton cells than Hg(II) (Lee and Fisher 2016; Mason et al. 1996; Pickhardt and Fisher 2007). If this is also applicable to bacteria, rapid accumulation of MeHg into bacterial cytoplasm may therefore provide enough substrate (MeHg) to induce demethylation by the organomercurial lyase, resulting in continuous Hg⁰ formation. In contrast, only part of the Hg(II) (free ion or neutral species) may be allowed to enter bacterial cells in the beginning of incubations and be reduced by mercuric reductase. Hu et al. (2013) reported that adsorption of Hg(II) by bacterial cells (Geobacter sulfurreducens PCA) would compete with, and therefore inhibit Hg(II) reduction. They suggested that the inhibition can be attributed to bonding between Hg(II) and the thiol functional groups on cells. Bioavailability of Hg(II) to microorganisms may be lowered because of further complexation of Hg(II) with other ligands (Barkay et al. 1997; Hsu-Kim et al. 2013; Schartup et al. 2015; Ullrich et al. 2001), explaining why less Hg⁰ production was observed at a later stage of incubation (Fig. 7). Although MeHg can also form strong complexes with ligands which decrease its bioavailability to microorganisms (Ndu et al. 2016), Hg(II) is more prone to form metal complexes than MeHg because of its greater formation constant with ligands. Similar cases can be seen in previous studies regarding MeHg production by Hg-methylating bacteria, where the highest methylation rate can be found in the first few days (Compeau and Bartha 1984; Hamdy and Noyes 1975; Ullrich et al. 2001).

In natural waters, Hg⁰ formation is a combination of photochemical and microbial processes. The importance and relative contribution of each process on total Hg⁰ formation may vary from one place to another. Some studies concluded that photochemical reduction of Hg(II) was the dominant process; for instance, earlier studies suggested that sunlight induced gaseous Hg⁰ formation was the most important route in lake water (Amyot et al. 1994) and coastal seawater (Amyot et al. 1997). Rolfhus and Fitzgerald (2004) estimated that approximately 70% of bulk reduction was driven photochemically and 20% was by microbial mediation in coastal surface seawater from Long Island Sound. Whalin et al. (2007) also suggested that biotic processes were relatively unimportant to Hg(II) reduction in surface waters of the Chesapeake. Tsui et al. (2013) and Point et al. (2011) used stable Hg isotopes as a proxy and concluded that photodegradation of MeHg is predominant in stream and Arctic marine ecosystems, respectively; they did not determine Hg⁰ production. Although the photochemical production of Hg⁰ can be very efficient, it is limited to the daytime and at the ocean’s surface. Rapid water column attenuation of sunlight with depth can diminish the importance of the photochemical process, especially in high latitude regions. For example, Poulain et al. (2007) reported that at a depth of 10 m below the surface of the Arctic Ocean, up to 90% of Hg⁰ was produced by microbial reduction of Hg(II) rather than photochemical reduction. Kuss et al. (2015) reported that microbial production of Hg⁰ under low-light conditions in the Baltic accounted for ~40% of total Hg⁰ production. If the microbial transformation of Hg⁰ can occur at any depth, the biological reduction of Hg(II) should become a significant source to the total pool of Hg⁰ in the water column. Our findings indicate that microbial activity can convert MeHg as well as Hg(II) to Hg⁰ and under light conditions typically prevalent at water depths of >10 m, can contribute significantly to Hg⁰ production. The Hg⁰ gas produced has the potential to be released to the atmosphere, as shown here, but the extent to which this occurs under prevailing oceanographic conditions remains to be determined.