Environ. Sci. Technol. 1991, 25,302-305
New Method for Detecting Methylmercury by Its Enzymatic Conversion to Methane Franco Bald1*!t and Marco Filippelli~
Dipartimento di Biologia Ambientale, Universits di Siena, via P.A. Mattioli, 4, 1-53100 Siena, Italy, and Laboratorio Chimico d’Igiene e Profilassi, 1-19100 La Spezia, Italy
rn A new method is suggested for determining methylmercury in biological samples. The determination is based on the use of whole cells of a “broad-spectrum” mercuryresistant bacterium, Pseudomonas p u t i d a , strain FB1, which is induced to produce the enzymes organomercurial lyase and mercuric reductase. These biocatalysts convert organomercurials to elemental mercury and their derivative hydrocarbons. The extraction procedure for methylmercury in biological samples follows the conventional toluene extraction method, which is retained until methylmercury is concentrated in an aqueous phase containing 0.01 M Na2S20,. This solution is mixed with bacterial cells in exponential growth phase and then incubated in microreaction vessels. A complete biological degradation of methylmercury to methane occurs. The efficiency of derivatization depends on the organomercurial concentration, the incubation time, and the cell density. The detection limit is 15 ng of methylmercury in 1 g of biological tissue and the coefficient of variation is 1.9% in 10 replicate samples with 100 ng/mL. Introduction The investigation of methylmercury in the environment is still of great interest, especially the natural alkylation of Hg(I1). So far, all procedures to detect methylmercury are mainly based on Westoo’s method ( I ) or on Cappon’s method (2). The extraction of methylmercury from samples generally consists of acidic hydrolysis of the organic matter, followed by sequential extractions with solvent and aqueous solutions. The organomercurial can be determined with different techniques, the most common being the detection of the chlorine moiety of methylmercury chloride in the solvent phase by gas chromatography with a 63Nielectron capture detector (ECD). Other methods to determine this organomercurial consist in detecting total mercury in the reextracted aqueous layer ( 3 , 4 )by flameless atomic absorption spectrophotometry (AAS). Still other methods are based on gas-liquid chromatography ( 5 , 6 ) , and on spectrometric determination ( 7 ) . Recently, Filippelli (8) suggested a more effective method to determine methylmercury in biological samples by comparing results obtained by both AAS equipped with a graphite furnace (GF-AAS) and GC-ECD techniques. For a more accurate speciation of organomercurials, sophisticated methods are also used, such as gas chromatography in line with mass spectrometry (GC/MS), or a “purge and trap” device connected with a GC in line with AAS (9). It is possible to use the specificity of microbial enzymes to detect methylmercury in environmental samples. For example, bacteria are resistant to mercury by virtue of two different enzymes: mercuric reductase ( I O ) , which reduces Hg(I1) to Hg(O),and organomercurial lyase ( I I , 1 2 ) ,which splits C-Hg bonds (C = methyl, ethyl, and phenyl), releasing the hydrocarbon and inorganic mercury (13-15). These enzymes are usually codified by genes often hart Universith
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bored in plasmids (16). The two enzymes also have been demonstrated to be inducible and specific for mercury compounds only, and no other organometallic compounds can be transformed by the organomercurial lyase ( I 7 ) . In a previous paper (18),we demonstrated that 98% of methylmercury was degraded in 130 min a t 37 “C to methane and elemental mercury by a culture of Pseudomonas p u t i d a , strain FB1. The two byproducts were detected, respectively by gas chromatograph/Fourier transform infrared spectroscopy (GC-FTIR) and by atomic absorption spectrophotometry. We also obtained a “cured” bacterial strain, FB4, by chemically depriving strain FB1. Thus, strain FB4 had lost its transforming activity for organomercurials. Then FB1, and FB4 strains were used to establish the microbial origin of methane from methylmercury degradation. The aim of this research is to suggest an alternative method for detecting methylmercury, based on its enzymatic derivatization to methane.
Experimental Section Apparatus. Methane analysis was carried out by a gas chromatograph (Hewlett-Packard Model 5890A) equipped with a flame ionization detector (FID). A 30 m X 0.530 mm GS-Q gas-solid open tubular megabore column (J & W Scientific) was operated isothermally a t 70 “C with a helium flow rate of 30 mL/min. The injection port temperature was held a t 200 “C, and the detector a t 250 “C. The integration area peaks were recorded by an integrator (Hewlett-Packard 3396 A). Total organic mercury analysis was made by a PerkinElmer Model 372 atomic absorption apparatus, equipped with a deuterium background corrector, an HGA 500 graphite furnace, and a mercury hollow-cathode lamp operated a t 6 mA (spectral band pass 0.2 nm) and a t a resonance wave length of 253.6 nm. A Sargent Model SRG recorder was used for mercury measurements. The experimental furnace conditions were as follows: drying, 110 “C for 20 s, ramp time 5 s; ashing, 200 “C for 10 s, ramp time 5 s; atomization, 1000 “C for 5 s, ramp time 1 s; cleaning, 2700 “C for 2 s, ramp time 1 s. The nitrogen internal gas flow (40 mL) was stopped in the atomization step. The organic mercury speciation was performed with a gas chromatograph (Carlo-Erba HRGC 5300 Mega series), equipped with a 63Nielectron capture detector. A CP-Si1 8 column (Chromopack wide-bore fused-silica 50 X 0.53 mm i.d., film thickness 2 pm) was held at 180 “C isothermally. The injector temperature was 250 “C and the detector temperature was maintained a t 250 “C. The nitrogen carrier gas flow rate was 10 mL/min. The bacterial density was measured by means of its absorbance ( A ) with a UV-visible spectrophotometer (UV-160 Shimadzu) at 600 nm and then compared with the dry weight (dw) of bacteria collected on nitrocellulose membranes (0.22 pm). The absorbance of the liquid culture was directly correlated to the dry weight of cells as follows:
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0 1991 American Chemical Society
y ( A ) = 0.976(mg of cells dw)
n = 10 r2 = 0.98 Bioderivatization Enzymatic Analysis. The cells (50 mL) were routinely grown a t 37 "C in Erlenmeyer flasks (250 mL) and agitated a t 200 rpm on an orbital shaker (New Brunswick Scientific Model G75). Cells were pelleted at 5000g with a centrifuge (ALC Model 4227 R) and rinsed twice with PIPES [piperazine-N,N'-bis(2-ethanesulfonic acid)] buffer (pH 7.4). For this analysis, 1 ml of sample containing methylmercury was extracted with toluene and then with thiosulfate (0.01 N) aqueous solution. The thiosulfate solution was mixed with 1mL of FB1 cells in liquid culture in microreaction vessels (2 mL; Supelco Inc., Bellafonte, PA), sealed with Mininert valve caps (Supelco). The mixture in the vials was stirred with a triangular magnetic bar (Supelco), a t room temperature. For headspace sampling, a 0.1-mL syringe (Dynatech Precision) was used. Cell-free extracts were obtained by 1-min sonication a t four intervals of 15 s each with a sonicator (New Brunswick Model MSE). The supernatant was recovered after 5 min a t 12000g in a microcentrifuge (Eppendorf Model 5415). The enzymes were activated by a complete assay mixture (0.75 mL) added to the cell free extract (0.25 mL), and they were mixed together with 1 mL of an extracted aqueous layer containing methylmercury (final volume 2 mL). Reagents. A 0.01 M solution of sodium thiosulfate (Na2S2O3.5H2O) was prepared by dissolving 0.2482 g in 100 mL of double distilled water (DDW). Iron peptone broth was prepared by dissolving in 1L of DDW 5.0 g of Universalpeptone M 66 (Merck), 0.5 g of glucose, 0.1 g of (NH4)2S04,and 0.1 g of FeS04.7H20and then autoclaving a t 120 "C for 20 min. The inducer of the FB1 cells was a solution of 0.1 M dimercury dibromofluorescein (merbromin, Merck), which was prepared by dissolving 0.7507 g in 10 mL of DDW. A buffering solution a t pH 7.4 for washing cells was obtained by dissolving 3.024 g/L PIPES (0.01 M). The enzymatic activity was measured by a complete assay mixture (CAM) containing a final concentration in PIPES (40 mL) of 0.5 mM EDTA, 0.6 mM MgC12, 1mM 2-mercaptoethanol (Bio-rad), 0.2 mg of bovine serum albumin (Sigma)/mL, 5 mg of chloramphenicol. The final NADPH (Fluka) concentration ranged from 0.3 to 0.9 mM (19). Standards. A total of 0.1251 g of methylmercury(I1) chloride (BDH Chemicals Ltd.) was dissolved in 1 L of DDW and stored a t 4 "C. Methane (1%) in nitrogen (Scotty I1 Supelchem) was used. A certified sample of homogenated mussel (MA-M-2/TM) from the International Atomic Energy Agency (IAEA) (20) was used for method intercalibration study. Procedures. Biological sample (1.0 g) plus 1.0 mL of concentrated HC1 were placed in 20-mL glass vials equipped with Teflon septa screwcaps. The vials were heated in a water bath at 100 "C for 5 min. The samples were cooled and made to 10 mL with DDW, and 1.0 mL of toluene was added. Vials were stoppered and shaken horizontally for 30 min and then centrifuged a t 6000g for 5 min. Thereafter, the toluene layer were transferred to 10-mL vials. The operation was repeated three times. Finally, the three toluene layers were combined, 0.5 mL of 0.01 M Na2S20, aqueous solution was added, and the resultant mixture was vortexed for 30 s (8). The toluene was carefully discarded to avoid inhibition of FB1 strain metabolism. One milliliter of the aqueous layer was added to 1 mL of an active culture of FB1 strain, previously induced by 0.1 mM merbromin solution (21). The 2-mL
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reaction vessels were sealed with Mininert valves and stirred with magnetic triangular bars for 4 or 18 h, depending on the amount of methylmercury in the sample. A volume of 0.1 mL from the headspace (0.7 mL) was injected in a gas chromatograph equipped with a flame ionization detector for determining methane produced by the biological derivatization of methylmercury. This process was carried out both with cell free extract and with actively growing cells. The cell-free extract was obtained by sonication of 20 mL of FB1 cells; a 1:lO dilution gave an absorbance (600 nm) of 0.61, for 1-min sonication with intervals of 15 s each under ice to avoid enzyme denaturation by heating. The cell-free extract was added to the assay mixture (CAM) plus the sample containing methylmercury. Results and Discussion The degradation of organomercurials and the determination of their byproducts is a well-documented process. P. putida strain FB1, a broad-spectrum mercury-resistant strain, is able to enzymatically convert methylmercury, ethylmercury, and phenylmercury to Hg(0) and the methane, ethane, and benzene (18) either in whole cells or in cell-free extracts. The cured strain, FB4, lost its ability to degrade organomercurials (10, 22). The conversion rate of organomercurials to the respective hydrocarbon is a fast and complete reaction (10). Therefore, this enzymatic feature can be utilized for determining organomercurials, particularly methylmercury, in biological samples. Two different conversion rates of MeHgCl to methane are obtained (Figure 1) for the in vivo and in vitro experiments. Although both correlations are significant, the in vitro experiment (cell-free extract) shows a conversion rate 3 times lower than the in vivo experiment (whole cells). The different results between the two systems might be attributed to lower efficiency of the complete assay mixture, and likely to a reduced activity of the organomercurial lyase. In addition, the aliquot of transformed methylmercury in cell-free extract depends on the concentration of NADPH added in the CAM (Figure 2). In whole cells, the enzyme and its cofactors instead are produced continuously until methylmercury undergoes a Environ. Sci. Technol., Vol. 25, No. 2, 1991 303
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complete degradation. Consequently, the in vivo system was preferred to the cell-free extract in further experiments. The in vivo degradation system is affected by the following parameters: (1)Time of incubation. The degradation of low concentrations of methylmercury (1pg) is completed in 2 h of incubation (Figure 3). However, if the content of MeHgCl in the microreaction vessels is above 10 pg, the enzymes are inhibited (Figure 4). If the incubation is longer than 18 h, the degradation is also completed at high concentrations and is linear over a large range (from 0.015 to 20 pg) (Figure 1). (2) Number of actively growing cells. The conversion of methylmercury (1 pg/mL) to methane depends on the number of bacteria present in the microreaction vessel (Figure 5 ) . T o achieve the total conversion of methyl304
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Flgure 5 . Enzymatic determination of methylmercury (1 pg/mL) correlated to different concentrations of biomass.
mercury to methane, the weight of bacteria (dw) or the absorbance (A600)should be, respectively, above 1mg/mL or 1.0 absorbance unit. Accuracy of the Method. To verify the efficiency of methylmercury conversion to methane in matrices from environmental samples, we spiked different concentrations of methylmercury into extracts of liver tissue from a dolphin containing 1.38 pg/g methylmercury (Figure 6) and also in other biological tissue (data not reported). The addition curve showed no matrix interferences with the enzymatic activity of whole cells of FB1. To further investigate the reliability of the method, an intercalibration exercise was performed with an IAEA mussel homogenate sample. We determined methylmercury by enzymatic (GC-FID), GF-AAS, and GC-ECD methods. The enzymatic method gave the same value as the other methods (Table I), which agreed well with the
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results from the interlaboratory study. The sensitivity of the method was 0.158; the detection limit based on a Student's t test (95%) was 15 ng of methylmercury extracted from 1 g of biological tissue, considering that the injected volume into the GC-FID was 100 pL and the headspace of 2-mL microreaction vessels was 0.7 mL. The coefficient of variation (CV), calculated with 10 replicates of the same sample containing an average value of 0.095 pg of methylmercury in 1 mL of thiosulfate solution, was 1.9%.
Conclusion The bioderivatization of methylmercury to methane is efficient not only in standard solutions, but also in biological extracts. The tissue sample matrix and 0.01 M thiosulfate solution does not affect the enzymatic activity of the whole cells of P. putida strain FB1. The importance and the applicability of the method is summarized as follows: (1) The use of microbial cells, as a source of biocatalysts for analytical determination, is innovative.
(2) This method is an alternative biological technique for methylmercury or other organomercurials analyses. (3) The GC-ECD is a sensitive but not specific method. The GC/MS is very specific, but expensive and not as sensitive a technique as the enzymatic method, which is also specific. (4) The chemical interferences are negligible in the enzymatic determination of methylmercury. (5) The organomercurial lyase reacts only with organomercurials and not with other organometallic compounds (18). The only exception is a report by Walts and Walsh (23),who demonstrated that a rare environmental stannous compound, triethylvinyltin, undergoes turnover by the organomercurial lyase to yield ethylene and ethane. The major difficulty in applying the enzymatic method is that this procedure might be difficult for laboratories without microbiological facilities. The method in its current form does not provide a time advantage over conventional methods. However, it maybe possible to improve the method by using immobilizing cells or distributing cells of FB1 strain in sealed microreaction vessels for storage and subsequent use. However, the specificity of this determination places the method among the most reliable ones. Registry No. CH,, 74-82-8.
Literature Cited (1) Westoo, G. Acta Chem. Scand. 1968, 22, 2277. (2) Cappon, C. J.; Smith, J. C. Anal. Chem. 1977, 49, 365. (3) Gage, J. C. Analyst (London) 1961,86, 457. (4) Davies, I. M. Anal. Chim. Acta 1978, 102, 189. (5) Fishbein, L. Chromatogr. Reu. 1970, 13, 83. (6) Mushak, P. Enuiron. Health Perspect. 1973, 4, 55. (7) Jones, P.; Nickless, G Analyst (London) 1978, 103, 1121. (8) Filippelli, M. Anal. Chem. 1987, 59, 116. (9) Blair, W. R.; Iverson, W. P.; Brinckman, F. E. Chemosphere 1974, 3, 167. (10) Fox, B.; Walsh, C. T. J. Biol. Chem. 1982, 257, 2498. (11) Begley, T. P.; Walts, A. E.; Walsh, C. T. Biochemistry 1986, 25, 7186. (12) Begley, T. P.; Walts, A. E.; Walsh, C. T. Biochemistry 1986, 25, 7192. (13) Furukawa, K.; Susuki, T.; Tonomura, K. Agric. Biol. Chem. 1969, 23, 128. (14) Tezuka, T.; Tonomura, K. J . Biochem. 1976,80, 79. (15) Robinson, J. B.; Tuovinen, 0. H. Microbiol. Reu. 1984,48, 95. (16) Brown, N.; Minsra, L.; Winnie, T. K.; Schmidt, J. N.; Seiff, A. M.; Silver, S. MGG, Mol. Gen. Genet. 1986, 202, 143. (17) Baldi, F.; Filippelli, M.; Olson, G. J. Microbiol. Ecol. 1989, 17, 263. (18) Baldi, F.; Cozzani, E.; Filippelli, M. Enuiron. Sci. Technol. 1988, 22, 836. (19) Shottel, J. L. J . Biol. Chem. 1978, 12, 4341. (20) Report no. 26. Atomic Agency Laboratory of Marine Radioactivity, Oceanographic Museum, Monaco, October 1985. (21) Weiss, A. A.; Murphy, S. D.; Silver, S. J . Racteriol. 1977, 132, 197. (22) Baldi, F.; Coratza, G.; Manganelli, R.; Pozzi, G. Microbios 1988, 54, 7. (23) Walts, A. E.; Walsh, C. T. J . Am. Chem. SOC.1988, 110, 1950.
Received for review June 8, 1990. Revised manuscript received September 20,1990. Accepted September 21,1990. We gratefully acknowledge financial support by the Commission of the European Communities, Research Project EV4V-0136-I.
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