Mercury methylation and hydrogen sulfide production among unexpected strains isolated from periphyton of two macrophytes of the Amazon


  • Dario Achá,

    Corresponding author
    1. Instituto de Ecología, Carrera de Biologia, Universidad Mayor de San Andrés, La Paz, Bolivia
    • Environmental and Life Sciences Graduate Program, Trent University, Peterborough, ON, Canada
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  • Cecilia A. Pabón,

    1. Carrera de Biología, Universidad Mayor de San Andrés, La Paz, Bolivia
    2. Department of Chemistry, Trent University, Peterborough, ON, Canada
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  • Holger Hintelmann

    1. Department of Chemistry, Trent University, Peterborough, ON, Canada
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Correspondence: Dario Achá, Campus Universitario de Cota Cota Calle 27. Instituto de Ecologia. Casilla 680 Correo Central La Paz, Bolivia. Tel.: +59 122 792582; fax: +59 122 772522; e-mail:


The periphyton of macrophytes had previously been identified as important spots for mercury methylation in the Amazon basin, but the microorganisms that facilitate methylation in such compartment are still to be identified. Here, bacteria were isolated from periphyton associated with Eichhornia crassipes and Polygonum densiflorum in Widdel and Pfennig medium and tested for mercury methylation with a stable isotope tracer technique using 198HgCl, hydrogen sulfide production and molybdate inhibition. Three Pleomorphomona spp., one unidentified Deltaproteobacteria, two Klebsiella spp., and one Tolumonas sp. were isolated. All except Tolumonas sp. were able to methylate mercury (up to 5% of the 198HgCl added) and produce up to 4 mM of H2S, while the Deltaproteobacteria was also able to demethylate methylmercury. Although these bacteria may not be as strong mercury methylators as sulfate-reducing bacteria, they have the potential to contribute to methylmercury accumulation in the system.


Mercury methylation is the first and crucial step for the introduction of mercury to the trophic chain. Methylmercury bio-accumulates and biomagnifies increasing by orders of magnitude the concentrations in predatory fish, which are the main source of human exposure to methylmercury (For a review Risher et al., 2002). Mercury methylation can be abiotic or biologically mediated, but the last produces most of the methylmercury in aquatic environments (Berman & Bartha, 1986; Coelho-Souza et al., 2006). In marine sediments, methylation is facilitated mainly by sulfate-reducing bacteria (SRB) (Compeau & Bartha, 1985; King et al., 2000). However, mercury methylation takes place not only in sediments, but also in the water column (Eckley & Hintelmann, 2006) and periphyton (Guimaraes et al., 2006) of fresh water environments where other bacteria are more likely to play a major role in carbon mineralization.

SRB link to mercury methylation was first established with molybdate inhibition experiments (Compeau & Bartha, 1985). However, in some compartments mercury methylation does not get completely inhibited by molybdate (Acha et al., 2005, 2011; Fleming et al., 2006). This shows that other groups of bacteria may be involved. So far, iron-reducing bacteria have been identified as potential mercury methylators (Fleming et al., 2006; Kerin et al., 2006) as well as methanogenic bacteria (Hamelin et al., 2011).

Mercury methylation in periphyton associated with macrophyte roots is the main site for mercury methylation in Amazon lakes (Guimarães et al., 2000). We found that mercury methylation in Eichhornia crassipes and Polygonum densiflorum root-periphyton samples amended with molybdate was inhibited by up to 39% and 51%, respectively, which suggests that SRB are not the only important mercury methylators in these environments (Acha et al., 2011). This is in agreement with other studies demonstrating that molybdate does not completely inhibit mercury methylation in the periphyton of Amazonian macrophytes (Acha et al., 2005), tropical lake sediments (Guimarães et al., 1998), boreal epilithon (Desrosiers et al., 2006) and lake sediments (Fleming et al., 2006). To explore which bacteria could be involved in mercury methylation in this compartment, we attempted to isolate bacteria with similar growing requirements as SRB from the periphyton associated with roots of E. crassipes and P. densiflorum. Then, we tested their ability to methylate mercury under similar conditions to those in which SRB were tested. We also evaluated the effect of different culture medium and conditions for mercury methylation.

Materials and methods

Sample collection and culture conditions

Roots plus associated periphyton from E. crassipes (Water Hyacinth) and P. densiflorum (Denseflower Knotweed) were manually collected at lake La Granja, which is an oxbow lake further described elsewhere (Acha et al., 2005, 2011). Around the macrophyte periphyton, the temperature was 26–31 °C and pH 6.9–7.5. A portion was saved for an in situ mercury methylation experiment, and ~ 2 cm3 of sample was transferred to a serum bottle containing 18 mL of anaerobic water and sealed again to prevent further oxygenation. Then, the samples were manually mixed for a few minutes to separate the periphyton from the roots. Subsamples were taken, using the Hungate technique to keep anaerobic conditions, and then inoculated into Widdel and Pfenning medium with lactate, acetate, or propionate as electron donor (Widdel & Bak, 1992) in 10-fold serial dilutions and three replicates for most probable counts and isolation of individual strains.

Periphyton was diluted and cultured in the fresh water version of Widdel and Pfennig medium (Widdel & Bak, 1992) reduced with hydrogen sulfide (1.5 mM), thioglycolate (88 μM) or titanium-NTA solution (0.1 mM). The electron donor in the medium was either lactate (20 mM), acetate (10 mM), or propionate (1 mM). The medium contained a nonchelated trace element mixture (1 mL L−1). Cultures from rhizosphere samples were initially kept in the dark at room temperature (25–30 °C). Subsequently, bacterial cultures were held at approximately 22 °C in the dark and were manipulated by standard Hungate technique.

Strain isolation and hydrogen sulfide production

Initially, the samples were diluted in culture medium by a factor of 10−4, and then the enriched cultures were again diluted several times until isolated colonies were distinguishable in the solid Widdel and Pfennig medium (~ 1 g agar L−1). The isolated colonies were extracted with a syringe and inoculated into a fresh medium.

The production of H2S by the isolated strains was first determined visually by precipitation of black iron sulfide (Widdel & Bak, 1992). Iron was added to the bacteria saturated medium as few drops of iron sulfate (FeSO4). Immediate formation of a black precipitate was assumed to be the result of H2S reaction with iron. A quantitative experiment was also conducted for some of the strains. Hydrogen sulfide concentration was determined by formation of methylene blue, separated by high-performance liquid chromatography (HPLC) coupled to UV–visible spectrometer as described by Small (2005). Sulfate reduction rates were calculated during the exponential phase of sulfide production.

To further evaluate the ability of bacteria to produce H2S, the presence of the dissimilatory sulfite reductase (dsr) gene was tested by PCR with DSR1F (5′-AC[C/G] CAC TGG AAG CAC G-3′) and DSR4R (5′-GTG TAG CAG TTA CCG CA-3′) primers for alpha and beta subunits of the dsr gene (Wagner et al., 1998). The amplification was considered positive only when the product matched the expected size (~ 1900 bp), as determined by agarose gel electrophoresis. The identity of the PCR product was verified by sequencing. We also tested molybdate (5 mM final concentration) inhibition.

DNA extraction and 16S rRNA gene sequencing

Genomic DNA was extracted with the UltraClean Microbial DNA kit (MoBio Inc.) following the recommendations of the manufacturer. Almost the entire 16S rRNA gene sequence was amplified by polymerase chain reaction (PCR) with fD1 (5′-AGA GTT TGA TCC TGG CTC AG-3′) and rP2 (5′-ACG GCT ACC TTG TTA CGA CTT-3′) (Weisburg et al., 1991) using Taq polymerase and dNTPs mixture from Promega at recommended concentrations. Each reaction comprised preheating at 95 °C for 2 min, 30 cycles of denaturation at 95 °C for 1 min followed by annealing at 55 °C for 1 min and extension at 72 °C for 1 min, and finally 72 °C for 5 min. PCR product size was verified with electrophoresis in 1.5% agarose gel prestained with SYBR green in 0.5× Tris-borate-EDTA buffer (45 mM Tris, 45 mM boric acid, 1.0 mM EDTA) and visualized by UV illumination. PCR products were purified with Wizard® SV Gel and PCR Clean up system (Promega) as recommended by the manufacturer and sent to be sequenced in the DNA laboratory sequencing facility at Trent University with 8f (5′-AGT TTG ATC CTG GCT CAG-3′), 1492r (5′-GGT TAC CTT GTT ACG ACT T-3′), 800f (5′-ATT AGA TAC CCT GGT AG-3′), 800r (5′-CTA CCA GGG TAT CTA AT-3′), 1050f (5′-TGT CGT CAG CTC GTG-3′), and 1050r (5′-CAC GAG CTG CGA CA-3′) primers.

A second partial sequencing was used to verify culture purity after experiments were concluded. Any strain that fail to pass this test was not included in this study, under the suspicion of culture contamination.

16S rRNA gene sequence analysis and strain identification

The 16S rRNA gene partial sequences obtained were manually corrected and assembled using BioEdit (version 7.0) and TraceEdit. Once assembled, the sequences were preliminarily identified using the Classifier tool of the Ribosomal Database Project (RDP) (Wang et al., 2007). Using the tool Sequence Match of the RDP (Cole et al., 2007), most similar sequences were downloaded to construct a phylogenetic tree using Mega version 4 (Tamura et al., 2007) and to confirm the classification.

Sequences of 400 to 1370 bp from the 16S rRNA were obtained for identifying the strains. Sequencing errors and other artifacts were corrected through proof reading of each fragment. It was verified that the sequences obtained had no anomalies by analysis with Pintail software (Ashelford et al., 2005) and Bellerophon online analysis (( (Huber et al., 2004). The sequences were identified with the Classifier, a tool of the Ribosomal Database Project II release 9.51 (RDP) ( (Cole et al., 2007).

Methylmercury analysis

Methylmercury analysis was conducted as previously described (Hintelmann et al., 1995; Hintelmann & Evans, 1997). Briefly, mercury species were extracted by distillation from approximately 2 mL of sample, and CH3201HgCl2 (50 pg) was added as a yield tracer to estimate the procedural recovery. The distillation product was transferred to a reaction vessel, where the different mercury species were ethylated with NaBEt4. The volatile reaction products were purged onto a tenax trap, which were then heated to thermodesorb the mercury species onto a gas chromatography (GC) column. Mercury species were separated by isothermal gas chromatography and quantified by ICP-MS (Hintelmann & Evans, 1997). Concentrations of newly methylated mercury and demethylated methylmercury were calculated using spread sheets as described in Hintelmann & Ogrinc (2003).

Mercury methylation experiments

Strains were initially incubated for 16–40 h until growth could be evidenced by the turbidity in the medium. Initial density measured as absorbance at 680 nm was between 0.01 and 0.02. Then the culture medium was spiked with 198HgCl2 to obtain final concentrations between 0.01 and 2000 ng mL−1 to test the influence of various amendments and spike concentration on the degree of mercury methylation. Subsamples for methylmercury determination and bacterial growth quantification by absorbance at 680nm were collected at different times depending on the rate of the bacterial growth. Microbial mercury methylation in the remaining sample was stopped by acidifying the sample with HCl to a final concentration of 0.5% and stored at 4 °C until methylmercury analysis.

The degree of mercury methylation was calculated using the change in CH3198Hg+ concentration during the exponential phase of growth. A net mercury methylation rate was calculated as a function of time during the linear increase in methylmercury period identified by plotting methylmercury production against time or as the maximum degree of methylmercury production at any given time interval. For some samples, mercury methylation was also expressed as a function of bacterial growth or sulfide production.

Three mercury methylation experiments were conducted. Firstly, the degree of mercury methylation among samples and controls was determined for each strain in triplicate. Second, the effect of different spike concentrations was evaluated with some strains. Third, the ability of bacteria to simultaneously methylate and demethylate methylmercury was tested by adding CH3202Hg+ after approximately 52 h of incubation, which is between late exponential and stationary phases. Positive and blank control samples included Desulfovibrio sp. 12ML1 (FJ865472.1) (Achá et al., 2012) and Desulfovibrio desulfuricans ssp. desulfuricans 27774. Autoclaved and uninoculated medium served as negative controls.

We used mostly regression analysis to evaluate the relation between Hg methylation, bacterial growth, and H2S production.

Results and discussion

Six of the seven strains isolated and identified from E. crassipes and P. densiflorum root-periphyton assemblages were able to methylate added 198Hg (II) (Table 1). Three of the strains belong to the class Alphaproteobacteria genus Pleomorphomonas, two to Gammaproteobacteria genus Klebsiella, and the last is an unclassified Proteobacteria (Fig. 1). None of these groups were previously described as potential mercury methylators. The strong significant (< 0.05) positive log linear relation between bacterial growth and methylmercury production (Fig. 2) suggests that methylation is dependent on the bacterial cell density, confirming their potential to produce methylmercury. Abiotic methylation would show a simple linear increase with no relation with the exponential increase in cell density. Only Tolumonas sp. E1P1 cultures did not show any significant (> 0.05) relation between mercury methylation and bacterial growth.

Table 1. Comparison of mercury methylation (ng L−1 day−1 ± standard error or% day−1) determined using different Hg(II) spikes and culture temperatures
Strain  CH3Hg+References
Hg(II) (μg L−1)Temp °C(ng L−1 day−1)(% day−1)
Pleomorphomonas sp. E2A10.02220.038 ± 0.0090.16This study
Klebsiella sp. E2A20.81300.136 ± 0.0350.07This study
0.01220.002 ± 0.0020.06
Pleomorphomonas sp. E2A30.01220.002 ± 0.0110.03This study
Pleomorphomonas sp. E2A40.81301.010 ± 0.0530.09This study
0.01220.021 ± 0.0530.21
1.05224.120 ± 5.2220.37
Unclassified Proteobacteria P5P30.81301.253 ± 0.0520.12This study
0.02220.057 ± 0.0020.23
Klebsiella sp. E1L10.01220.034 ± 0.0240.34This study
Desulfovibrio desulfuricans desulfuricans 277740.06220.0780.13This study
Desulfovibrio africanus7.3937547.44 ± 60.005.47Ranchou-Peyruse et al. (2009)
Desulfovibrio africanus1.0028333.27 ± 145.4224.55Ekstrom et al. (2003)
Desulfovibrio desulfuricans LS7388.492735.000.35Choi & Bartha (1993)
Desulfovibrio desulfuricans61.7828107.52 ± 283.200.13King et al. (2000)
Enteroboacter aerogenes0.92373.1360.04Hamdy & Noyes (1975)
Clostridium cochlearium370783716.67< 0.01Pan-Hou & Imura (1982)
Desulfococcus multivorans61.7828110.88 ± 513.600.13King et al. (2000)
Desulfococcus multivorans 1be11.0028230.29 ± 87.4616.96Ekstrom et al. (2003)
Desulfobacter sp. BG-861.782837.20 ± 511.200.04King et al. (2000)
Desulfobacterium sp. BG-3361.7828180.72 ± 237.600.22King et al. (2000)
Desulfobulbus propionicus61.782825.20 ± 729.600.03King et al. (2000)
Desulfobulbus propionicus 1pr31.0028274.79 ± 37.7820.24Ekstrom et al. (2003)
Desulfobulbus propionicus 1pr37.39372064.96 ± 113.2820.65Ranchou-Peyruse et al. (2009)
D. propionicus MUD1.0028729.67 ± 161.4653.75Ekstrom et al. (2003)
Figure 1.

Neighbor-Joining phylogenetic tree constructed from the 16S rRNA gene sequences of the isolated strains (black circles) and the closest sequences found with the Ribosomal database project seqmatch tool. The phylogeny was evaluated by Boostrap with 1000 replicates (number next to the node).

Figure 2.

Relationship between logarithm base 10 of the bacterial growth of Pleomorphomonas sp. E2A4 expressed as absorbance at 680 nm and 198Hg(II) methylation.

To further verify the ability of the isolated strains to methylate mercury, we compared methylmercury production in the strains with nonspiked and autoclaved controls. Methylmercury production of the nonmethylating SRB control (Desulfovibrio desulfuricans ssp. desulfuricans 27774) was one order of magnitude lower to that of Pleomorphomonas sp. E2A4, unclassified P5P3 or Klebsiella sp. E2A2 (Table 1).

Even if the strains described here are not as strong mercury methylators as many SRB and IRB (Fleming et al., 2006; Kerin et al., 2006; Ranchou-Peyruse et al., 2009), they may account for part of the methylation observed in molybdate amended samples (Acha et al., 2011) because they do not get fully inhibited by molybdate (Table 2). In a previous work, we found a strong relation between mercury methylation and Desulfobacteriaceae as well as mercury methylation stimulated by acetate (Acha et al., 2011). Our results are not in contradiction with such a hypothesis, because the strongest methylator we found (Pleomorphomonas E2A4) was also isolated in acetate as well as other three of six Hg methylating isolates.

Table 2. Strain metabolic characterization in Widdel and Pfennig medium
StrainAcetateEthanolLactatePropionateNo reducing agentNo sulfateOxygenMolybdate
  1. Nd, not determined; +, full growth evidenced by saturation of the media; (+), limited bacterial growth with some turbidity observed but no saturation of the media; −, no growth was observed (OD680 nm not different from blank).

Pleomorphomonas sp. E2A4++++++(+)
Pleomorphomonas sp. E2A3++Nd(+)+++(+)
Pleomorphomonas sp. E2A1+++(+)+++Nd
Klebsiella sp. E2A2++++++++
Klebsiella sp. E1L1++++++NdNd
Tolumonas sp. E1P1++++(+)NdNd

16S rRNA gene-based identification

The bacteria isolated from the rhizospheres were diverse and corresponded to three different classes of the phylum Proteobacteria. At least three of them were Alphaproteobacteria (E2A1[FJ750583], E2A3[FJ750587], and E2A4 [FJ750582]), three Gammaproteobacteria (E1L1[FJ750584], E1P1[FJ750585], and E2A2[FJ750586]) (Fig. 1), and one unclassified Proteobacteria (P5P3). Alphaproteobacteria were highly similar to Pleomorphomonas oryzae (97%, 95%, and 99% respectively by NCBI blastn). Although this does not mean that they are the same species, it suggests that they are likely to belong to the same genus. E1L1 and E2A2 were up to 99% similar to Klebsiella pneumoniae strains, while E1P1 strain had a similarity of 97% with Tolumonas auensis according to NCBI blastn.

Most probable number (MPN)

Most probable number counts in the rhizosphere from the Amazonian macrophytes showed higher abundance of acetate-utilizing bacteria in rhizospheres from both E. crassipes and P. densiflorum. The MPN of acetate-utilizing sulfide-producing bacteria was more than 106 bacteria per cm3 of root-associated periphyton, because bacterial growth and H2S were observed in the lowest dilution. There was no significant difference (P > 0.05) between the MPN of lactate and propionate-utilizing bacteria in samples from either rhizosphere (Supporting Information, Fig. S1). However, in samples from E. crassipes rhizosphere, there was a slightly higher MPN of lactate and propionate-utilizing bacteria (1.15 × 105 and 9.33 × 104 respectively) than in samples from P. densiflorum (7.41 × 103 and 2.76 × 104 respectively).

Sulfate reduction

Control medium without bacterial inoculation did not show detectable sulfide (< 0.05 μM). Two bacteria testing negative for sulfide production upon iron sulfate addition contained only 2.43–9 μM of hydrogen sulfide in solution. The bacteria that tested positive produced between 0.124 and 2.797 mM of H2S day−1 (Table 3). Such amounts are comparable to the sulfide production by SRB (King et al., 2000) (Table 3) and in the case of Klebsiella strain are consistent with previous studies that show hydrogen sulfide production under anaerobic conditions (Holmes et al., 1997; Sharma et al., 2000). The sulfide production follows the same exponential trend as the bacterial growth and methylmercury production until 40h of incubation (Fig. 3), so does not appear to be the result of bacterial decomposition. The inverse relation with methylmercury production after 40h may be attributed to demethylation processes. It could be argued that part of the measured sulfide may originate from the thioglycolate added as reducing agent, but its concentration was only about 0.9 mM and some strains produced up to 4 mM of sulfide.

Figure 3.

Hydrogen sulfide and CH3198Hg+ production over time in a culture of Pleomorphomonas sp. E2A3.

Table 3. Degree of sulfide and methylmercury production normalized to sulfide. The uncertainty is expressed as standard error for data from this study and as standard deviation for the rest
StrainH2S mM day−1198Hg methylated/H2S ng L−1 mM−1
  1. a

    Cultured using titanium-NTA (1 mM) as reducing agent.

  2. b

    Cultured using thioglycolate as reducing agent.

  3. c

    Obtained from King et al. (2000).

Unclassified Proteobacteria P5P3a0.2661 ± 0.10430.299 ± 0.096
Unclassified Proteobacteria P5P3b0.1243 ± 0.0352ND
Pleomorphomonas sp. E2A40.5087 ± 0.41160.727 ± 0.040
Klebsiella sp. E2A22.7970 ± 0.09060.211 ± 0.074
Desulfovibrio desulfuricans ATCCc0.3655 ± 0.1824 
Desulfobulbus propionicus ATCCc0.4075 ± 0.3096 
Desulfococcus multivorans ATCCc0.0996 ± 0.4800 
Desulfobacter sp. BG-8c0.0473 ± 0.2928 
Desulfobacterium sp. BG-33c0.0326 ± 0.1992 

The possibility that the bacteria here isolated are dissimilatory SRB under certain conditions is intriguing. No SRB have been reported among Alphaproteobacteria or Gammaproteobacteria, but as SRB are phylogenetically widely distributed (Castro et al., 2000) and horizontal gene transfer of sulfate reduction genes was demonstrated (Klein et al., 2001; Friedrich, 2002; Zverlov et al., 2005), the observation is not entirely surprising. However, it is not clear whether some of these strains are producing hydrogen sulfide through a dissimilatory sulfate reduction pathway. We were not able to sequence the dsrAB gene with standard primers, which may be attributed to inadequate primers, but suggests that they are not typical SRB.

Mercury methylation potentials

Autoclaved and blank controls showed almost no detectable amounts of MMHg (< 0.05 and 0.2 pg mL−1 respectively), which tend to decline over time. The potential of some of the strains to produce methylmercury is comparable to that of SRB reported by King et al. (King et al., 2000) and Choi and Bartha (Choi & Bartha, 1993), but up to two orders of magnitude lower than the rates reported elsewhere (Ekstrom et al., 2003; Ranchou-Peyruse et al., 2009) (Table 1). However, the differences in the magnitude of methylmercury produced can be attributed to factors that are not necessarily related to the significance of these bacteria for Hg methylation in the environment. The large variations in the degree of mercury methylation found in the literature, even among experiments with same strain, are likely a product of different culture conditions, concentration of the Hg (II) spike and of how well adapted the strain is to a particular medium.

We tested the effect of initial Hg(II) concentration and different temperatures in some of the samples (Table 1). The amount of Hg(II) spike had a significant effect on the percentage methylated by each strain. Generally, the degree of methylation increased with the increasing spike concentrations (Table 1). This is in agreement with Gilmour et al. (2011) results, but is in contradiction to Choi & Bartha's (1993) findings. However, the concentrations of added Hg(II) used in the second study were three orders of magnitude higher than those used in the present study (Table 1). We also attempted to test the effect of temperature and different reducing agents, but we encountered adaptation problems (data not shown).

Culture conditions can also affect methylation (Choi & Bartha, 1993; Gilmour et al., 2011). All strains cultured here had the ability to produce sulfide and grow under sulfate-reducing conditions so they were tested for methylation under such conditions. This is important because it helps in comparison with methylation potentials of SRB, and because it has been suggested that a neutral HgS0 compound is the most bioavailable form for methylation (Benoit et al., 1999, 2001). The fact that the isolated bacteria can grow using sulfate and the same electron donors as many SRB could mean that they compete for such resources and both contribute to methylmercury accumulation. However, their ability to methylate mercury may vary considerably depending on the conditions in their natural environment, which may differ significantly from culture conditions.

It must be acknowledge that the ability of a strain to methylate mercury in culture conditions does not imply that such a strain is an important methylmercury producer in situ. An instructive example are methanogens, which were early on suggested to be potential mercury methylators (Wood et al., 1968), but later in situ examination has repeatedly found that they are not important in most compartments (Compeau & Bartha, 1985, 1987; Gllmour et al., 1992; Pak & Bartha, 1998). It was just recently demonstrated that methanogens could be important methylators in periphyton of temperate fluvial lakes (Hamelin et al., 2011) Also, it appears that mercury methylation is species or even strain specific (Ranchou-Peyruse et al., 2009; Gilmour et al., 2011) and therefore the isolation of a single strain able to methylate mercury is not necessarily a good indicator for methylmercury production potentials of the family or class. However, as the large diversity of bacteria present in most natural systems cannot be tested simultaneously at the family or genus level for its role in mercury methylation in situ, culture studies remain highly relevant to identify groups to be further studied in situ.

Methylmercury demethylation

After noticing that in several experiments the concentration of methylmercury declines at the end of the incubation (Fig. 3), the possibility of demethylation occurring during such process was tested. Of the three strains tested (E2A4, E2A2, and P5P3), only P5P3 demethylated methylmercury. The net demethylation was 11.13 ng L−1 day−1 or 27% of the added CH3202Hg+, but demethylated instantaneously (at time 0 after CH3202Hg+ addition) about 55% of the CH3202Hg+ added. Such demethylation is not observed with the newly produced CH3198Hg+. This may be explained by the sequestration of methylmercury and by a protective effect of thioglycolate toward methylmercury. Probably, methylmercury produced in the cell remains associated with the bacteria during its growth, but when the cells start to decay, methylmercury becomes available for demethylation, while the added methylmercury may be susceptible to fast demethylation. This is in agreement with our observations of in situ behavior of methylmercury formation and demethylation (unpublished data). However, as the actual demethylation of added methylmercury was only observed in one of the strains and in a particular medium, the causes for such behavior are unclear and should be further investigated.


This work was supported by an NSERC Strategic Research Grant to H.H. and an NSERC Discovery Grant to Janet Yee. We thank Mark Dzurko for providing the control and C. Gilmoure for providing the original strains. We also thank Joy Zhu, Brian Dimock, Dephine Foucher, and Olivier Clarisse for guidance in mercury analysis and general laboratory assistance.