Volatilization and Precipitation of Tellurium by Aerobic, Tellurite-Resistant Marine Microbes

Media and reagents.Chemicals and reagents were purchased from Sigma-Aldrich (St. Louis, MO), Fisher Scientific (Pittsburgh, PA), or VWR Scientific (West Chester, PA) and were of the highest grade available. All growth media were based on Luria-Bertani (LB) broth which contains the following, per liter: 10 g tryptone, 5 g yeast extract, and 5 g NaCl. A previously described trace element solution (49) was added to the medium at 1 ml liter⁻¹. The medium was solidified with 1.5% (wt/vol) agar (biotech grade; Fisher Scientific) for colony isolation and streaking. All growth on plates was carried out at room temperature, while liquid cultures were grown at 30°C with shaking at 250 rpm.

Isolation and growth of strains.An optimized medium, called LB-marine, contained the following, per liter: 2.0 g tryptone, 1.0 g yeast extract, 12.5 g sodium chloride, and 1 ml of trace element solution. This mixture was adjusted to pH 8.1, 1.5% (wt/vol) agar was added for plates when desired, and then the mixture was autoclaved for 15 min at 121°C. After cooling, 20 ml of sterile 1 M magnesium sulfate was added per liter of medium prior to pouring plates or inoculating liquid cultures. A 0.22-μm-filter-sterilized 10 mg ml⁻¹ stock of sodium tellurite was added to the media to achieve a final concentration of 150 μg ml⁻¹ (0.7 mM) for the selection of resistant strains.

Mud samples, from the upper 2 cm of sediment, were collected from fringing salt marsh bordering the Indian River inlet in Rehoboth Beach, DE, in May of 2004. The mud samples were slurried 1:10 (vol/vol) in 0.45-μm-filter-sterilized water collected at the sampling site, transported to the laboratory, and stored under air at room temperature without shaking. Enrichment for tellurite-resistant organisms was performed by amending slurries with 150 μg Na₂TeO₃ ml⁻¹.

Strains were isolated from primary dilutions of the slurries on LB-marine plates both in the presence and absence of 150 μg Na₂TeO₃ ml⁻¹. Single tellurite-resistant colonies were purified by restreaking until a single colony morphology was consistently obtained. Purified strains were grown in liquid medium, and frozen glycerol stocks were prepared for long-term storage (5) at −70°C. Strains were revived from glycerol stocks by streaking onto LB-marine agar with tellurite.

Tellurite resistance determination.Tellurite resistance was assessed by growing each strain in LB-marine medium in the absence of tellurite and by quantifying their ability to grow on LB-marine agar containing tellurite. Cell concentrations in liquid cultures were determined by direct counting using a Hausser counting chamber (Fisher Scientific, Pittsburgh, PA). Cultures were diluted to a concentration of ∼2 × 10³ cells ml⁻¹ in LB-marine medium, and 100 μl of this suspension (∼200 CFU) was spread onto LB-marine agar without amendment (control) or containing concentrations of Na₂TeO₃ ranging from 75 to 1,200 μg ml⁻¹ (0 to 5.4 mM) and was incubated for at least 2 weeks to allow for the observation of slow-growing colonies.

Gram staining.Gram stains of culture or colony smears were performed with commercial reagents (Protocol Gram stain set; VWR, West Chester, PA) according to manufacturer instructions. Stained samples were observed on an Olympus (Central Valley, PA) BX61 microscope equipped with a UApo/340 40× objective.

rRNA gene amplification and sequence analysis.Genes encoding 16S/18S rRNA from individual isolates were amplified by PCR using the universal primer pair 519F-1406R (46). Amplified products were cloned into the pCR2.1 plasmid vector by using a commercial kit (Invitrogen, Carlsbad, CA) and were used to transform Escherichia coli TOP10 (Invitrogen, Carlsbad, CA) by electroporation. White colonies on LB plates containing 50 μg kanamycin ml⁻¹ were screened for inserts by PCR with M13-Reverse and T7-Promoter primers (Invitrogen, Carlsbad, CA). Two clones for each strain were sequenced by standard protocols, using the same primers, at the University of Delaware Sequencing and Genotyping Center. Sequences were compared with the Vector NTI program suite (Invitrogen, Carlsbad, CA) to generate a consensus contig using the Contig Express module.

Sequence alignments were carried out in MEGA3 (31) using the Clustal W algorithm. The phylogenetic trees produced from the resulting distance matrices were analyzed by neighbor joining with bootstrapping (1,000 replicates) using the Kimura 2-parameter correction method.

Quantification of solid and dissolved tellurium species.Tellurium content in liquid and solid phases of samples was determined using a Perkin-Elmer (Waltham, MA) model 3300 atomic absorption spectrometer equipped with a graphite furnace accessory HGA-600 (graphite furnace-atomic absorption spectrometry) and an autosampler. A hollow cathode lamp was employed as the emission source at 214.3 nm with a slit width of 2 nm and 30 mA lamp current. Measurements were performed in peak area (integrated absorbance) mode. Tubes with pyrolytic graphite coating were used throughout the experiments. High-purity argon was used as the internal gas. The temperature-time program was performed according to Shiue et al. (45). The formation of tellurium oxides TeO (g) during pyrolysis can lead to analyte losses (40). To overcome this issue, a 20-μl aliquot of the sample (i.e., 0.5 to 2 ng Te) was injected into the furnace, followed by 20 μl of palladium (30 μg ml⁻¹, i.e., 0.6 μg Pd) mixed with magnesium (200 μg ml⁻¹; i.e., 4 μg Mg) matrix modifier. With these techniques, a linear range was found between 0 and 16 μM Te (0 to 2 μg ml⁻¹) by using a commercially prepared standard solution (Aldrich Chemical, Milwaukee, WI) in 5% (vol/vol) HNO₃.

For each sample, culture was centrifuged (9,000 × g, 25 min) and supernatant transferred to a Teflon beaker, where it was evaporated to dryness at 90 to 95°C. The residue was dissolved with high-purity, sub-boiled HNO₃, dried at 90 to 95°C, and redissolved in 5% (vol/vol) HNO₃. The cell pellet from the centrifugation described above was resuspended with Te-free medium and collected again by centrifugation. When this wash supernatant was analyzed as described above, the tellurium was less than 1% of that measured in the original supernatant. Pellets were dissolved with sub-boiled HNO₃, dried as described above, and redissolved in 5% (vol/vol) HNO₃. Samples from the supernatant or pellet were diluted by factors ranging from 5- to 210-fold in order to obtain tellurium concentrations between 25 and 100 ppb. Samples containing no added tellurium were analyzed with each batch of samples to estimate the level of potential contamination introduced by lab operations. In all cases, these were indistinguishable from the background level.

Detection of volatile tellurium species by gas chromatography-mass spectrometry (GC-MS).The headspace above cultures was sampled with a manual solid-phase microextraction (SPME) holder using a 75-μm carboxen-polydimethylsiloxane fiber (Supelco, Bellefonte, PA). Cultures (2 ml) amended with tellurite were grown in sealed 18-ml headspace vials (Supelco, Bellefonte, PA) under room air for sampling. The fiber was exposed to the headspace over the liquid cultures amended with tellurite for 48 h. The fiber was then retracted and removed from the vial and immediately inserted into the injection port of a Hewlett-Packard (Palo Alto, CA) 6850 gas chromatograph coupled with a Hewlett-Packard 5973 mass selective detector (electron impact). A 0.75-mm inner-diameter liner and predrilled thermogreen septa for SPME were used. Ultra-high-purity helium was the carrier gas with a flow rate of 1 ml min⁻¹. An Agilent HP-5ms (5% phenylmethylpolysiloxane; Agilent Technologies, Palo Alto, CA) column was used. All samples were analyzed under splitless injection with a desorption time of 10 s at an injector temperature of 280°C. Reinserting the SPME fiber after the run did not show any carryover. The GC oven was thus kept at 25°C (room temperature) for 3 min after desorption and then heated to 200°C with a ramp of 5°C/min. The final temperature was held for 2 min. The interface and the ion source temperatures were 280°C.

Compounds were identified by matching their mass spectra to the NIST and Wiley spectral libraries, requiring a resemblance percentage above 85% for identification. Commercial standards of dimethyl telluride (DMTe) and dimethyl ditelluride (DMDTe) are unavailable. However, the natural isotopic abundance of tellurium makes Te-containing fragments clearly identifiable in MS. The peaks of DMTe and DMDTe appeared after a retention time of approximately 2.5 to 3.7 and 16.6 to 19.3 min, respectively. To validate the elution time of DMTe, a >98% pure standard was synthesized from trimethyl telluronium iodide after the work by Kuhn et al. (30).

Localization of precipitated tellurium.Culture samples were observed by phase-contrast microscopy on an Olympus (Central Valley, PA) BX61 microscope equipped with a UPlan Fl 40× Ph2 objective and phase condenser. Images were acquired with a Retiga EXi charge-coupled-device camera (QImaging, Surrey, BC, Canada) and stored as TIFF files.

Cells from cultures were harvested by centrifugation and fixed with 2% glutaraldehyde and 2% paraformaldehyde in 0.1 M Na cacodylate (primary fixative) and 1% OsO₄ (secondary fixative). Resin infiltration was carried out with Embed-812 (Electron Microscopy Sciences, Hatfield, PA). Blocks were sectioned on a Reichert-Jung ultra-cut E microtome (Leica Microsystems, Bannockburn, IL) with a diamond knife. Thin sections were approximately 60 to 70 nm (silver interference color) and were collected on copper grids (Electron Microscopy Sciences, Hatfield, PA). The sections were poststained with uranyl acetate and methanol as well as Reynolds' lead citrate (44). The samples were viewed using a Zeiss (Goettingen, Germany) CEM 902 transmission electron microscope at 80 kV, and images taken with a Soft Imaging system mega view II camera (Olympus Soft Imaging, Lakewood, CO).

Nucleotide sequence accession numbers.The rRNA gene sequences reported here have been deposited in GenBank under accession numbers EU405881 to EU405886.

Isolation of marine tellurite-resistant bacteria.The effect of medium composition on the recovery of culturable microbes from collected salt marsh sediments was assessed by systematically altering the basic recipe for LB broth to optimize it for use with salt marsh sediments. The total amount of tryptone plus yeast extract, in a 2:1 (wt/wt) ratio, varied between 1.5 g liter⁻¹ and 30 g liter⁻¹, with the peak number of strains recovered at 3.0 g liter⁻¹. Sodium chloride varied between 85 mM and 428 mM, with consistently improved culturability observed above 171 mM. Magnesium sulfate was added to the medium up to 80 mM, with the optimum at 20 mM. The initial pH of the medium varied between 7.2 and 8.7, with the optimum at 8.1. The addition of 1 ml liter⁻¹ of a standard trace element solution afforded a 1.2-fold increase in strain recovery over basal LB broth. In general, optimizing each parameter provided a 1.2- to 2.0-fold improvement over basal LB broth. Optimized LB-marine contained, per liter, 2.0 g tryptone, 1.0 g yeast extract, 214 mM sodium chloride, 20 mM MgSO₄, and 1 ml liter⁻¹ of trace element solution at a pH of 8.1.

The total culturable population of aerobic microbes recovered on LB-marine medium in the absence of tellurite, selection averaged 1.2 × 10⁴ CFU ml⁻¹ in 1:10 sediment slurries indicating a culturable population of 1.2 × 10⁵ CFU per milliliter of original sediment. The total number of tellurite (0.70 mM)-resistant organisms recovered was 9.0 × 10² CFU ml⁻¹ in sediment slurries, indicating an initial population size of 9.0 × 10³ CFU ml⁻¹ tellurite-resistant strains in the original sediment. Thus, ∼8% of the total culturable population was found to be tellurite resistant. Enrichment with 0.70 mM tellurite in sediment slurries for periods of up to 2 weeks increased the proportion of tellurite-resistant strains twofold, to ∼15% of the total culturable microbial population (data not shown).

When isolated strains from LB-marine medium without tellurite were inoculated on plates containing 150 μg Na₂TeO₃ ml⁻¹ (0.70 mM), 8% of these strains were found to be tellurite resistant, duplicating the original fraction of tellurite resistance observed in the initial isolation experiment. All tellurite-resistant strains from the original isolation grew in the absence of tellurite and maintained their tellurite resistance. A total of 30 strains were purified by repeated streaking of isolated colonies on LB-marine medium plus tellurite and carried forward for characterization.

Characterization of tellurite-resistant strains.Tellurite-resistant isolates were grouped initially on the basis of colony morphology and subsequently characterized for their tellurite resistance range on LB-marine plates (Table 1). Based on these two criteria, the 30 isolates could be divided into three clusters. Six representative model strains from these clusters were carried forward to further examine their properties (Table 1).

Cluster 1 is composed of highly tellurite-resistant isolates (Fig. 1A) that form compact, nonspreading rose-pink colonies. On plates containing tellurite, these colonies are dark black in color. Cluster 2 is composed of isolates that display moderate tellurite resistance (Fig. 1B). These organisms have variable colony morphology. One of the model strains for this cluster, strain 6A, forms moderately sized, white colonies with a fungal appearance. In contrast, strain 28A forms large, shiny, pale-orange colonies in the absence of tellurite and gray to black minute colonies in the presence of tellurite. Cluster 3 is composed of isolates that display relatively weak tellurite resistance (Fig. 1C). Even though strain 14B, the cluster 3 model strain, was isolated in the presence of 0.70 mM Na₂TeO₃, it grows poorly at this concentration both on plates and in liquid cultures. Therefore, this strain was routinely propagated in the presence of 0.17 mM Na₂TeO₃. Colonies grown without tellurite are buff colored or white. In the presence of tellurite, colony sizes are greatly diminished, and colonies were colored slightly gray. Liquid cultures of this strain tend to grow as gelatinous aggregates, rather than as dispersed cultures typical of the other isolates.

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FIG. 1.

Resistance of model strains cultured in the absence of tellurite to various concentrations of sodium tellurite on LB-marine plates. Approximately 200 CFU were seeded onto each plate, and the viable population observed on LB-marine plates without tellurite was defined as 100% viability. (A) Cluster 1 strains 1A, 13B, and 30B. (B) Cluster 2 strains 6A and 28A. (C) Cluster 3 strain 14B. Symbols for each strain are noted in the figure. Data points are the average of the results for two independent experiments for each strain.

All tellurite-resistant strains isolated to date in this study stained gram positive. Isolated colonies recovered on LB-marine medium in the absence of tellurite selection contained both gram-positive and gram-negative organisms with nearly equal frequencies (data not shown). Thus, it appears that a specific subset of gram-positive organisms was identified by tellurite selection and that the gram-negative organisms in the upper sediment layers sampled were not tellurite resistant under the conditions tested. Strains in all clusters were also resistant to 0.70 mM tellurate, selenate, selenite, arsenate, and arsenite under aerobic growth conditions (data not shown).

Taxonomic assignment of isolates.A ca.-900-bp fragment of rRNA genes was PCR amplified from each of the six model strains in Table 1, cloned, and sequenced. Comparison of cluster 1 rRNA gene sequences to those in known databases indicated that these strains are all eukaryotes similar (98.8 to 99.8% identity) to the yeast Rhodotorula mucilaginosa, strains of which are frequently isolated from marine and estuarine sediments (22, 34) (Fig. 2A). Comparison of rRNA gene sequences from the isolates within clusters 2 and 3 unambiguously identified them as members of the family Bacillaceae, order Bacillales of the class Bacilli within the phylum Firmicutes of gram-positive bacteria (Fig. 2B). Cluster 2 strains were most similar (99.6 to 99.7% identity) to various uncharacterized marine Bacillus isolates (23). The cluster 3 strain 14B was most similar (99.2% identity) to strains of Bacillus halodenitrificans (syn. Virgibacillus halodenitrificans [53] and Oceanobacillus iheyensis [37]). None of the strains closely related to those identified here have been reported as being resistant to tellurium, selenium, or arsenic oxyanions.

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FIG. 2.

Taxonomic affiliations of tellurite-resistant strains characterized in this study. Isolates from this study are indicated by their cluster and strain number in boldface text. Individual sequences are noted by their GenBank identifier, and strains isolated from marine environments are noted in italics. Major clades aside from those containing tellurite-resistant strains have been collapsed for clarity. Bootstrap support from 1,000 replicates is indicated at each node. Scale bars indicate the numbers of substitutions per site. (A) Combined 16S/18S rRNA gene sequence tree including isolates from cluster 1. (B) Bacterial 16S rRNA gene sequence tree including isolates from clusters 2 and 3.

Volatilization of tellurium by isolated strains.The hypothesis that tellurite resistance is linked with tellurite volatilization was tested in the model strains by GC-MS analysis of headspace gas samples (Fig. 3). DMTe was the most-abundant volatile tellurium species detected, followed by DMDTe and the mixed species dimethyltellurenyl sulfide (DMTeS) (Fig. 3A). The identity of all putative Te compounds was verified by detailed mass-spectral analysis to ensure that they displayed previously observed fragmentation patterns characteristic of organotellurium compounds (Fig. 3B). GC-MS analysis of the headspace above sterile controls did not detect any volatile tellurium species (see Fig. S1 in the supplemental material).

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FIG. 3.

Representative SPME-GC-MS analysis of the headspace above cluster 1 strain 13B cultured for 48 h. The total ion count trace (A) indicates the relative proportions of each species detected. The detailed mass spectrum for DMTe (B) displays the characteristic signature expected based on Te isotope natural abundance and fractionation patterns resulting from the loss of methyl groups.

The ability of strains to produce a suite of volatile compounds was assayed at time points ranging from 2 days to 5 weeks of time in culture. All strains tested produced volatile sulfur compounds, while only strains in clusters 1 and 2 produced volatile tellurium compounds (Table 2). Dimethyldisulfide was detected in nearly every sample analyzed between 48 h and 5 weeks of growth. Dimethylsulfide (DMS) and dimethyltrisulfide were also detected with somewhat lower frequency. In cluster 1 and 2 strains, DMTe was more abundant than any of the volatile sulfur compounds detected (Fig. 3A). Generally, cluster 1 strains appeared to be most proficient at both S and Te volatilizations, as judged by the variety of compounds produced and the frequency of detection of these compounds over 5 weeks of culture.

Comparing clusters 1 and 2, two of three cluster 1 strains produced DMDTe, while this compound was not observed in headspace samples above cultures of cluster 2 isolates. The mixed species DMTeS was also observed frequently in cluster 1 strains but less frequently in cluster 2 strains. This is likely due to the timing of Te volatilization and relative activities of cluster 1 versus those of cluster 2. As shown in Fig. 4 for DMTe, volatile Te species were observed at all times in cluster 1 strains (Fig. 4A), with a peak abundance at 2 weeks. In contrast, volatile tellurium species were only observed after 5 weeks of culture in cluster 2 strains (Fig. 4B). Cluster 1 strains also produced between 5- and 20-fold greater amounts of volatile Te compounds than did cluster 2 strains, as can be seen by comparing the axis limits in Fig. 4. While the SPME-GC-MS analysis was not quantitative in absolute terms, relative abundances can be determined as all samples were treated identically.

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FIG. 4.

Time course of DMTe production by strains from clusters 1 (A) and 2 (B). Relative amounts of DMTe in each headspace sample were determined by SPME-GC-MS analysis after 48 h, 2 weeks, or 5 weeks of culture. Note that the scale of the y axis in panel B is 100-fold lower than that of panel A. Data points are the average of the results for two independent cultures per strain for each time point.

Within cluster 1, strain 30B behaved somewhat differently from strains 1A and 13B, as it consistently failed to produce DMDTe. DMS, dimethyltrisulfide, and DMTeS were less frequently observed in this strain (Table 2). In addition, while the temporal pattern of DMTe levels peaked after 2 weeks of culture in all cluster 1 strains, strain 30B produced three- to fivefold smaller amounts of volatile Te compounds than did the other two strains in this cluster.

The facts that no volatile Te compounds were detected in the headspace above the cluster 3 strain 14B and that DMDTe was observed only with cluster 1 strains indicate that the production of DMTe and DMDTe is biologically mediated. If abiotic chemical reactions were responsible, similar types and amounts of compounds should have been observed in all cultures. Furthermore, no volatile Te compounds were observed in the headspace above sterile medium incubated under identical conditions (See Fig. S1 in the supplemental material).

Cell-associated, particulate Te is the major product of Te metabolism by the isolates.As SPME-GC-MS analysis did not allow a direct determination of the amount of Te volatilized by each strain, we sought to determine the maximum possible amount of Te in volatile species by measuring Te in the soluble and particulate fractions after various times in culture (Fig. 5). The amount of volatilized Te was then obtained by subtraction from the amount of Te initially added into each culture. In each case, the soluble and particulate Te was found to account for nearly all the added Te. Total recovery of Te in the soluble and particulate fractions ranged from 80 to 110% of the amended Te in any given measurement, with a mean of 95% ± 6%. Cluster 1 strains appear to be more efficient at precipitating Te, converting ∼98% of added Te to a particulate form (Fig. 5A), while cluster 2 and 3 strains converted only 30 to 40% of added Te to a particulate form (Fig. 5B and C) over 5 weeks of culture. While it is possible that some of the soluble Te observed was a dissolved alkyl species, the total amount of alkylated Te species is likely much less than 5% of the total tellurite converted by these strains.

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FIG. 5.

Recoveries of soluble and particulate Te in model strains for each cluster. The concentration of Te in each fraction was determined by graphite furnace-atomic absorption spectrometry. A total of 0.65 mM TeO₃²⁻ was added to cultures of strains 13B (cluster 1) (A) and 28A (cluster 2) (B), while the culture of strain 14B received 0.16 mM TeO₃²⁻ (cluster 3) (C), denoted by the dashed lines in each panel. Dark bars denote tellurium recovered as precipitates; light bars represent tellurium recovered in the liquid and are the mean of four measurements (± the standard deviation).

Cellular localization of particulate Te.To determine the localization of Te particulates, cultures were examined directly by phase-contrast microscopy to determine if they were producing extracellular crystalline materials, but no significant amounts were observed in any of the strains (data not shown). Thin sections of fixed cells were examined by transmission electron microscopy (TEM), and strains in all clusters were found to contain electron-dense bodies that were present only when strains were cultured in the presence of tellurite (Fig. 6). Generally, these electron-dense bodies were found evenly distributed throughout the cell sections. This lack of distinct localization indicates that the Te is precipitated intracellularly without any obvious membrane association. Strain 28A in cluster 2 was the exception to this rule, as it tended to form precipitates in regions close to the cell periphery, suggesting that the tellurium-precipitating activity in this strain is membrane associated.

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FIG. 6.

Localization of precipitated Te in model strains from each cluster. Representative TEM images are displayed for model strains grown in the presence (+) or absence (−) of tellurite. Images are arranged in rows, with the strain indicated at the left edge of each row and the growth condition noted at the top of each column. A sevenfold-magnified subsection (indicated by a white box) of the +TeO₃²⁻ image for each strain is shown to highlight the different types of precipitates produced by each strain.

Strains appeared to produce different shapes and sizes of precipitates. Cluster 1 strains generally formed clusters of short needles of <100 nm in length, though individual cells sometimes contained clusters over 300 nm in length. As an aside, the images of strain 13 support the eukaryotic affiliation of cluster 1 strains inferred from 18S rRNA gene sequencing, as nuclei and mitochondria were clearly distinguishable in most sections. Strains in cluster 2 displayed more variability in the Te precipitate structure. Strain 6A formed spheres and amorphous aggregates that ranged from <10 to >50 nm in diameter. In contrast, strain 28A precipitates were primarily observed as aggregates of needles at the cell periphery a few hundred nanometers in length. Precipitates produced by the cluster 3 strain 14B were less electron dense and less compact than those of other strains. This may be due to the relatively low tellurite levels (0.18 mM) required for the growth of strain 14B in liquid culture and the tendency of this strain to aggregate in culture. Aggregation may protect cells by exclusion of tellurite leading to lower intracellular concentrations for precipitation.