Vol. 55, No. 5APPLIED AND ENVIRONMENTAL MICROBIOLOGY, May 1989, p. 1196-12020099-2240/89/051196-07$02.00/0Copyright © 1989, American Society for Microbiology

Environmental Significance of the Potential for mer(Tn2l)-MediatedReduction of Hg2+ to Hgo in Natural Waterst

TAMAR BARKAY,1* CYNTHIA LIEBERT,2 AND MARK GILLMAN2Microbial Ecology and Biotechnology Branch,' and Technical Resources Inc.,2 Environmental Research Laboratory,

U.S. Environmental Protection Agency, Gulf Breeze, Florida 32561

Received 7 November 1988/Accepted 8 February 1989

The role of mer(Tn2l) in the adaptation of aquatic microbial communities to Hg2+ was investigated.Elemental mercury was the sole product of Hg2+ volatilization by freshwater and saline water microbialcommunities. Bacterial activity was responsible for biotransformation because most microeucaryotes did notsurvive the exposure conditions, and removal of larger microbes (>1 ,Lm) from adapted communities did notsignificantly (P > 0.01) reduce Hg2+ volatilization rates. DNA sequences homologous to mer(Tn2l) were foundin 50% of Hg2+-resistant bacterial strains representing two freshwater communities, but in only 12% of strainsrepresenting two saline communities (the difference was highly significant; P < 0.001). Thus, mer(Tn2l) playeda signfficant role in Hg2' resistance among strains isolated from fresh waters, in which microbial activity hada limited role in Hg2+ volatilization. In saline water environments in which microbially mediated volatilizationwas the major mechanism of Hg2e loss, other bacterial genes coded for this biotransformation.

Adaptation of microbial communities to pollutants resultsin the enhancement of biodegradation and the developmentof resistance to these noxious substances (9). Hypotheses onthe nature of the mechanisms and microbial interactions thatlead to adaptation have been proposed and investigated (31).These hypotheses include (i) the derepression of preexistingenzymatic activities and selection of subpopulations of theactive organisms (42); (ii) the emergence of novel metabolicpathways for degradation of the pollutant by genetic ex-change among organisms of contaminated environments(23); and (iii) the selection and enrichment of populationsthat degrade the pollutant or that can survive in its presence(7, 32). Selection plays a role in enrichment of populationscreated by the former two mechanisms. Our approach forimproving the understanding of mechanisms underlying mi-crobial adaptation has been to relate changes in the genepool of the adapting community with metabolic processesand physiological responses that occur during adaptation.The bacterial operon coding for resistance and reduction

of Hg2+ was selected to study the role of specific genes inadaptation to an environmental pollutant. Aquatic microbialcommunities adapt to Hg2+ by rapidly volatilizing (and thuseliminating) it from their immediate environment (7). Be-cause Hg2+ reduction to volatile elemental mercury (Hgo) isa well-established resistance mechanism in pure bacterialcultures (35), its role in adaptation and volatilization by theaquatic communities was investigated by DNA-DNA colonyhybridization with a DNA probe constructed of a gram-negative mer operon, mer(Tn2J) (8). Although resistancewas widely distributed among aerobic .ulturable hetero-trophs of adapted communities, DNA sequences homolo-gous to mer were rarely (if at all) detected (7).Three hypotheses were considered to explain this discrep-

ancy. (i) Hg2+ was not transformed to Hg° but to one ormore alternative volatile forms by processes not genetically

* Corresponding author.t Contribution no. 660 of the U.S. Environmental Research

Laboratory, Gulf Breeze, FL 32561.

encoded by the mer gene. Ethyl and methyl mercury arevolatile compounds (24). Although little is known about theirformation, microbial methylation of Hg2+ may proceedunder aerobic conditions (28, 37). Methyl mercury formationhas been documented in the water column of marine envi-ronments (38) and freshwater lakes (43).

(ii) Volatilization of Hg2+ was mediated by microbes otherthan bacteria. Although the mer-mediated reduction of Hg2+is the only biological process known to have specificallyevolved to detoxify this heavy metal, microbial metabolismmay indirectly result in Hg2+ reduction. For example, Hg2+volatilization induced by the alga Chlorella pyrenoidosa hasbeen described previously (11). This process was mediatedby organic acids released from algal cells, was light depen-dent, and was inhibited by specific inhibitors of photosyn-thesis (12). A general mechanism for such indirect Hg2+volatilization may be its reduction to Hg+ and the sponta-neous disproportionation of Hg22 to Hg0 and Hg24 (6).Very low concentrations of reductants, such as those thatare present in distilled water, are sufficient to drive Hg2 outof solution by this mechanism (39).

(iii) Mercuric mercury resistance and volatilization wasmediated by bacterial genes that were not homologous to thecharacterized mer operon. The DNA sequences of meroperons originating in three gram-negative bacteria havebeen shown to have a high degree of homology (30), and thishomology was sufficient to result in cross-hybridization withmer(Tn2l) (8). However, Hg2+-resistant strains that do nothybridize with this probe have been isolated from differentenvironments (2, 10; N. Hamlett, personal communication),and NADPH-dependent mercuric reductase activity hasbeen found in three mer(Tn2l)-negative marine caulobacters(22).

In the experiments described here we examined the threehypotheses. We demonstrated that Hg2+ is biotransformedto Hg0 by freshwater and estuarine microbial communities,that this activity is solely mediated by the bacterial compo-nent of both communities, and that strains with alternativemer genes play an important role in the ecology of Hg2+resistance and volatilization in aquatic environments.

1196

on January 3, 2020 by guesthttp://aem

.asm.org/

Dow

nloaded from

mer(Tn2l)-MEDIATED Hg2+ REDUCTION IN NATURAL WATERS

MATERIALS AND METHODS

Sampling sites and sample collection and processing.

Coastal marine samples were collected from a pier on the

Gulf of Mexico at Pensacola Beach, Fla. Estuarine samples

were obtained from nearby Santa Rosa Sound, and freshwa-

ter samples were collected at Thompson's Bayou, Fla. A salt

marsh sample was collected at Range Point, Fla. These sites,

the sampling procedures, and the methods used during

physicochemical measurements have been described previ-

ously (7). An additional freshwater sample was collected 90

miles (145 km) east of Pensacola, Fla., at Vortex Spring,

which is a part of the Floridian limestone aquifer system with

a high mineral content and slightly alkaline pH (29).

Determination of volatile mercurial species. To obtain mi-

crobial communities with full Hg2+ volatilization activity,

water samples were exposed in microcosms to 250 jig of

Hg2+ per liter for 2 days at 29°C. It was previously shown (7)

that volatilization by microbial communities of saline waters

commenced only after a lag period that lasted 6 to 12 h. A

total of 50 ml of Hg2+-adapted cell suspensions was placed in

100-ml serum bottles (Wheaton Industries, Millville, N.J.)

that were plugged with butyl rubber stoppers and crimp

caps. 203HgC12 (Amersham Corp. Arlington Heights, Ill.)

was added to a final concentration of 250 pug of Hg2+ per liter

(specific activity is indicated in footnote a of Table 1).

Water-saturated air, which was sterilized by passage through

a 0.3-p.m-pore-size hydrophobic glass fiber filter (Gelman

Sciences, Inc., Ann Arbor, Mich.), was delivered by stain-

less steel needles to the bottom of the bottles. Air leaving

these systems passed through a 25-ml trapping solution (10%

KBr, 1.5% HgBr2, 0.1% CUSO4) in test tubes (150 by 18 mm;

Bellco Glass, Inc., Vineland, N.J.). After 24 h of incubation

at 28°C, the experimental vessels were disassembled, and

1-ml-samples from bottles and traps were removed to scin-

tillation vials for determination of radioactivity. The control

consisted of filter-sterilized (pore size, 0.22 ,um) sample

water, as described by Barkay (7). Recoveries (see footnote

b of Table 1) ranged between 83 and 95 percent.

The method adapted by Compeau and Bartha (15) from

that of Longbottom et al. (27) was followed to determine the

mercury species in trapping solutions. A differential extrac-

tion procedure was used to distinguish whether mercury was

trapped as Hgo or as methyl mercury. Organomercurial

compounds were extracted from solutions by partition into

toluene, and under the same conditions (27) inorganic mer-

cury remained in the aqueous phase. The 23 ml of trapping

solution was acidified to a pH of <0.5 by the addition of 0.35

ml of concentrated H2SO4 and extracted with three fractions

(3.7 ml each) of toluene. The aqueous and toluene phases

were separated, and the toluene fractions were pooled (total

volume, 11.1 ml). Samples (1 ml) of the aqueous and organic

phases were removed to scintillation vials. Counting of

radioactivity and data analysis were performed as described

previously (7).The capacity of the trapping solution that was used to

absorb both Hgo and methyl mercury and the effectiveness

of toluene as an extractant of methyl mercury were tested.

Elemental mercury was produced by the chemical reduction

of 203HgC12 dissolved in distilled water by using a 10%solution of SnCl2 (41). A total of 95+ 17% of the trapped Hgowas partitioned with the aqueous phase; radioactivity in the

organic phase was below the level of detection. Methyl

mercury was biologically produced by the activity of an

estuarine isolate of Desulfovibrio desulfuricans (16). The

organism was grown in a high-concentration chloride me-

dium in a 100-ml serum bottle (Wheaton Industries), asdescribed by Compeau and Bartha (16), in the presence of 15,ug of 203Hg2" per ml (as HgCl2; specific activity, 1.2 ,uCi permg of Hg2+) for 48 h at 25°C. At the end of the incubationperiod, 12.5 ml of fivefold-concentrated trapping solutionfollowed by 0.9 ml of concentrated H2SO4 were injectedthrough the septum into the bottle, producing a single-strength trapping solution at a pH of <0.5. The solution wasextracted with three fractions (12.5 ml each) of toluene, and1-ml samples of the aqueous and organic phases wereremoved for counting. Radioactivity partitioned with theorganic phase amounted to 13.00 ± 0.93 ng/ml. This corre-sponded to the yield of methyl mercury produced by the teststrain (15 ng/ml) after 48 h of incubation, as reported byCompeau and Bartha (16).

Experiments to determine the role of microeucaryotes inHg2+ volatilization. (i) Preparation of Hg2+-adapted microbialcommunities. An estuarine sample (pH, 8.16; temperature,25°C; salinity, 26%o) was obtained on 31 May 1988, and afreshwater sample (pH, 6.70; temperature, 27°C; salinity,below the level of detection) was collected from Thompson'sBayou on 5 July 1988. Four replicate adapted communitieswere prepared as described by Barkay (7). After 2 days ofpreexposure, two replicate communities were passedthrough a 1-ji.m-pore-size polycarbonate filter (NucleporeCorp., Pleasanton, Calif.) by using a vacuum pump operatedat a pressure of 30 to 40 mm of Hg to remove the largermicrobes. These communities were called filtered, adaptedcommunities, and the remaining two replicate communitieswere called unfiltered, adapted communities. Cells werespun down and suspended in filter-sterilized sample water.

(ii) Hg2+ analysis. Hg2+ volatilization was monitored asdescribed previously (7), except that 1-ml samples wereremoved to scintillation vials, acidified by the addition ofHNO3 (70.0 to 71.0%; Hg content, <0.005 ppm[,ug/ml]; J. T.Baker Chemical Co., Phillipsburg, N.J.) to a final concen-tration of 0.5 N, and stored frozen until they were analyzedfor total mercury by a modification of the cold vapor atomicabsorption procedure (40). Diluted samples (8 ml) wereplaced in 30-ml graduated glass impingers (Wheaton Indus-tries); 0.5 ml of concentrated H2SO4 (95.0 to 98.0%; Hgcontent, S ppb [ug/jip]; Fisher Scientific Co., Fair Lawn,N.J.) was added, followed by the addition of 0.25 ml ofconcentrated HNO3. Samples were incubated for 15 min atroom temperature after the addition of 1.5 ml of 5% KMnO4(Fisher Scientific Co.), 0.8 ml of 5% K2S208 (J. T. BakerChemical Co.) and 0.6 ml of a solution consisting of 12%NaCl-12% (NH20H)2. H2SO4 (Fisher Scientific Co.) werethen added sequentially. A suspension of 10% SnC12 in 0.5 NH2SO4 (0.5 ml) was added, the aeration device of a glassimpinger connected to a mercury analyzer system (modelMAS-SOB; The Perkin-Elmer Corp., Norwalk, Conn.) wasimmediately inserted, and light absorption was measured at453.7 nm. The log of the ratio between light absorbed, asdetermined from the peak heights produced on the chart ofan Omniscribe recorder (model B521-75; Houston Instru-ment, Austin, Tex.), and light emitted was used to determinetotal mercury in the sample with a standard curve. The limitof the detection procedure was 50 ng per diluted sample.

(iii) Bacterial and microflagellate counts. Heterotrophicmicroflagellates and bacterial cells were counted at the timeof sampling, after 2 days of preexposure, after suspension infilter-sterilized water, and at termination of the volatilizationexperiment. Samples were fixed by the addition of formal-dehyde (final concentration, 3.7%) and stored at4°C untilbacterial and microflagellate counts were performed by the

VOL. 55, 1989 1197

on January 3, 2020 by guesthttp://aem

.asm.org/

Dow

nloaded from

APPL. ENVIRON. MICROBIOL.

TABLE 1. Determination of the chemical species of volatile mercurial compound produced by aquatic microbial communities

Mercury in fraction:Hg (,ug) produced bya: Remaining in Trapping Recovery Aqueous Organic phase

bottles solution (%)b phase (Hgo) [(CH3)2HgI

Coastal marine communityLive 6.57 ± 0.09 4.46 ± 0.30 88 4.13 ± 0.01 <0.11CSterile 10.02 ± 0.04 0.37 ± 0.05 83 0.38 ± 0.02 <0.11

Freshwater communityLive 7.23 ± 0.03 4.50 ± 0.53 94 4.66 ± 0.08 <0.08Sterile 11.49 ± 1.54 0.38 ± 0.02 95 1.71 ± 0.01 <0.08" Total mercury (as Hg2+) was calculated by considering the final volume of each sample (see text) in the indicated fraction. Calculations were based on specific

activities of 4.56 ,Ci of Hg2+ per mg for coastal marine samples and 5.88 ,iCi of Hg2+ per mg for freshwater samples.b Percent recovered calculated as: [(Hg2+ remaining in bottle + mercury in trapping solution)/total Hg2+ added (12.5 Rg)] x 100.' Below the level of detection, assuming 100 dpm/ml as the background level.

procedures of Hobbie et al. (20) and Caron (13), respec-tively.

(iv) Primary production measurements. The amount of "'Cincorporated into cellular material was determined by themethod of Strickland and Parsons (34) at the time of sam-pling and after 2 days of preexposure. An estuarine samplewas collected on 21 March 1988, and primary productionwas determined by the addition of 2 ~iCi of NaH14CO3(specific activity, 7.0 mCi/mmol; DuPont, NEN ResearchProducts, Boston, Mass.) to three 65-ml fractions. Onesample was immediately filtered through a GC/F filter (What-man, Inc., Clifton, N.J.) for time zero determination, and theremaining two samples were incubated at 30°C for 5 h beforefiltration. Filters were immersed in PCS scintillation cocktail(Amersham Corp.) and counted as described above for203Hg2 . Primary production by a freshwater sample (col-lected on 23 August 1988 from Thompson's Bayou) wasdetermined similarly, except that NaH"4CO3 (DuPont, NENResearch Products) with a specific activity of 8.4 mCi/mmolwas used.

Isolation of Hg2+-resistant bacterial strains and colonyhybridization. Diluted water samples were plated onto petridishes containing growth medium supplemented with 10 ,ugof Hg2+ per ml (as HgCl2) and onto medium containing 50 ,ugof Hg2+ per ml, as described previously (7). By using arandom-number table, 60 bacterial colonies were selected torepresent each community at each Hg2+ concentration.Isolated colonies were streaked onto fresh medium (supple-mented with appropriate concentrations of Hg2+) and Gramstained; gram-negative cultures were stored at -70°C in 50%(vol/vol) glycerol. Colonies of these strains were hybridizedto the mer(Tn2l) probe as described by Barkay (7).

Statistical analysis. Regression analysis and comparisonbetween decay curves of Hg2+ was performed by using thestatistical analysis system (SAS Institute, Cary, N.C.) asdescribed previously (7). A P value of <0.01 was used todetermine the significance of differences observed betweendecay rates, following the rationale presented by Cripe et al.(17). Chi-square analysis of colony hybridization results wasperformed as described by Armitage (3).

RESULTSChemical species of volatile mercurial products. The vola-

tile mercurial products of coastal marine and freshwatercommunities were examined because these communities haddistinct patterns of Hg2+ volatilization. Volatilization re-quired a lag period with saline water communities, butproceeded immediately on exposure of freshwater commu-nities (7). Elemental mercury was identified as the only

major product of volatilization by both coastal marine andfreshwater communities (Table 1). Dimethylmercury did notaccount for more than approximately 2% of the volatilemercurial compounds, considering the limit of detection.Thus, the hypothesis that volatile dimethylmercury is amajor product of microbial biotransformation of Hg2 + innatural waters was rejected.

Role of microeucaryotes in Hg2+ volatilization. A role inHg2+ volatilization could be attributed to microeucaryotes inadapted microbial communities if they survived the 2 days ofpreexposure to Hg2' and if their elimination caused adecrease in Hg2+ volatilization rates. The susceptibility ofphotosynthetic and heterotrophic microeucaryotes to Hg2+was investigated by primary productivity measurements andcell counts of heterotrophic microflagellates, respectively.Photosynthetic microflagellates were less abundant thanheterotrophic microflagellates in the estuarine and freshwa-ter samples (R. Coffin, personal communication). The pos-sible indirect role of photosynthesis in Hg2+ reduction (11,12) was ruled out by demonstrating that primary productionwas inhibited by Hg2+ (Table 2). In estuarine samples, only1 to 2% of the original activity remained after 2 days ofpreexposure. Photosynthetic microeucaryotes in the fresh-water sample were more tolerant to Hg2+, with 13% of theinitial activity remaining after 2 days of preexposure.

Estuarine heterotrophic microflagellates were also highlysensitive to Hg2+, with 95% mortality during preexposure.Only 10% of surviving microflagellates passed the filtrationstep and potentially could contribute to Hg2+ volatilizationby filtered communities. Freshwater heterotrophic mi-croflagellates were more tolerant to Hg2+, with 41% surviv-ing preexposure. Filtration removed most of these cells, asindicated by a 91% decrease in counts in the filtered, adaptedcommunity (Table 2). Preexposure and filtration had noeffect on bacterial counts in estuarine samples, but filtrationresulted in a substantial decrease in bacterial counts infreshwater samples. This was due to removal of bacteria thatwere attached to particulate matter. Microscopic observa-tions of freshwater samples, prior to filtration, indicated thatapproximately 40% of bacterial cells were attached to parti-cles.

Statistical analysis of first-order decay curves indicatedthat larger eucaryotic microbes did not play a significant rolein Hg2+ volatilization in estuarine and freshwater microbialcommunities (Fig. 1). In the estuarine samples (Fig. 1A),biological activity was responsible for volatilization (P <0.0001 for the difference between filter-sterilized control andbiologically active samples), and the differences betweenloss mediated by filtered, adapted and unfiltered, adapted

1198 BARKAY ET AL.

on January 3, 2020 by guesthttp://aem

.asm.org/

Dow

nloaded from

mer(Tn2l)-MEDIATED Hg2+ REDUCTION IN NATURAL WATERS

TABLE 2. Bacterial and heterotrophic microflagellate counts in, and primary production by, Hg2+-exposedaquatic microbial communitiesa

Estuarine Freshwater

Sample Bacteria Microflagellates Primary Bacteria Microflagellates Primary

(cells/ml) (cells/ml) production (cells/ml) (cells/ml) (dpm/65-ml sample)

Before preexposure 2.02 x 106 2,577 51,895 ± 6,700 1.39 x 107 2,702 13,868 ± 554

After 2 days of exposure 2.12 x 106 167 721 ± 42 1.44 X 107 1,102 1,864 ± 285

After suspension in filtered sample waterAdapted community 8.68 x 105 469 NDb 1.48 x 107 837 NDFiltered, adapted community 8.06 x 105 34 ND 8.90 x 106 100 ND

End of volatilization periodAdapted community 3.13 x 106 3 ND ND ND NDFiltered, adapted community 2.13 x 106 34 ND 4.34 x 106 ND ND

a Microbial communities were exposed to 250 ,ug of Hg2+ per liter.bND, Not determined.

communities was not significant (P = 0.2138). Volatilizationof Hg2+ from freshwater samples (Fig. 1B) was largely dueto nonbiological processes, with only a small (yet significant;P = 0.0092) difference between decay curves for the filter-sterilized control and the unfiltered community. The ob-served difference between the two adapted communities (notsignificant; P = 0.4388) was probably due to the removal ofbacterial cells that were attached to particles in this sample(see above). Thus, microeucaryotes (heterotrophic and pho-tosynthetic) in adapted microbial communities could nothave played a significant role in the volatilization of Hg2+.The presence of mer(Tn2l) in the genomes of Hg2+-resistant

strains. The hypothesis that DNA sequences coding forHg2+ resistance and volatilization in saline water and fresh-water communities had no homology to the characterizedmer operon was tested by hybridization of randomly se-lected representative strains with a mer(Tn2l) DNA probe.Gram-positive strains were excluded from this analysisbecause their Hg2+ resistance genes bear little homologywith mer(Tn21) (30, 35) and homology was not sufficient for

hybrid formation under the stringency conditions used in thisstudy (8). However, only a minority of the resistant strainswere gram positive (ranging from 7.5% in the Vortex Springcommunity to 1.9% in the coastal marine community). Atotal of 60 strains were isolated (final samples of 50 strainsper community per Hg2+ concentration were expected). Adecrease in the number of strains were anticipated duringhandling because of the loss of viability, loss of the resis-tance phenotype, and exclusion from analysis of gram-positive strains. This was the case with one freshwater andtwo saline water communities, whereas strains isolated fromVortex Spring rapidly lost their resistance phenotype. Thereason for this loss is not clear, but it resulted in a smallersample representing the Vortex Spring community (Table 3).Two phenomena distinguished Hg2'-resistant strains iso-

lated in freshwater environments from those isolated insaline environments. The first was resistance to a higherHg2+ concentration (50 ,ug/ml) (Table 3). Bacteria fromsaline water samples did not grow on plates containing 50 ,ugof Hg2+ per ml, even when cells were concentrated from 1

IA. ESTUARINE 100-

80-

60-

40-

20-

24 3 7 12TIME (HOURS)

FIG. 1. Volatilization of Hg2+ mediated by filtered (0) and unfiltered (-) microbial communities of estuarine (A) and freshwater (B)environments. Filter-sterilized sample water (A) was used as a biologically inactive control.

100

800zz< 60w

+ 40

aR

IB. FRESHWATER

24

VOL. 55, 1989 1199

on January 3, 2020 by guesthttp://aem

.asm.org/

Dow

nloaded from

1200 BARKAY ET AL.

TABLE 3. Presence of mer(Tn2l) in the genomes ofHg2+-resistant bacterial strains isolated from saline water

and freshwater samples

No. of mer(Tn2J)-positive bacteriaSample resistant to Hg2+ at":

10 jig/ml 50 jig/ml

FreshwaterVortex Spring 11 (27) 16 (31)Thompson's Bayou 26 (50) 27 (52)

Saline waterCoastal marine 13 (52) NonebSalt marsh 0 (53) None

a The concentration of Hg2+ in the growth medium used for selection ofresistant strains is given. Numbers in parentheses indicate the total number ofstrains tested.

b No colonies grew after 7 days of incubation.

liter of water. The number of highly resistant strains reached31.7 ± 7.1 CFU/ml in the Thompson's Bayou sample and0.031 ± 0.017 CFU/ml in the Vortex Spring sample. Therewas no difference in the frequency of mer(Tn2J) homologoussequences among strains resistant to 50 p.g of Hg2+ per ml,as compared with those isolated on 10 jig of Hg2+ per ml(Table 3).The second phenomena was that mer(Tn2J) was signifi-

cantly more abundant. A total of 50% of freshwater strainscarried mer(Tn2J)-homologous DNA sequences, whereasonly 12% of strains from saline environments hybridizedwith the probe. Chi-square analysis indicated that this dif-ference between organisms from fresh and saline waters washighly significant (P < 0.001). However, a highly significantdifference (P < 0.001) was also found between the two salinecommunities. Hybridization with mer(Tn2l) occurred withone-quarter of the coastal marine strains but with no saltmarsh strains (Table 3). Thus, half of the resistant freshwaterstrains and the majority of the saline water strains did nothybridize with a gene probe constructed of the mer operon ofgram-negative bacteria. Hypothesis iii in the Introduction,that genes with no homology to mer(Tn2J) coded for resis-tance to, and volatilization of, Hg2+, was therefore ac-cepted.

DISCUSSIONRecent developments in the application of molecular ap-

proaches and biotechnological methods to microbial ecology(21) have created experimental tools that allow researchersto analyze molecular events that take place during microbialadaptation to pollutant stress (9). One approach to this studyrelates the distribution of biodegradative genes (detected bynucleic acid gene probes) in the active microbial communitywith adaptation to pollutants. However, a myriad of factors,in addition to the genetic potential of the indigenous micro-bial community, determines the fate and effects of pollutantsin a given environment. Among them are physical chemicalprocesses (5) and alternative mechanisms of biotransforma-tion mediated by any of the populations making up themicrobial community. In addition, the distribution of genescould be estimated erroneously if alternative genes code forthe same reactions. Thus, the validity of our approach to thestudy of the mer(Tn2l) role in adaptation to Hg2+ (7) wasbased on the assumption that Hgo was the volatile product ofHg2+ volatilization, that the bacterial component of aquaticmicrobial communities mediated this activity, and that asignificant number of representative Hg2+-resistant strainscarried mer(Tn2l)-homologous genes.

Elemental mercury was the major volatile mercurial com-pound formed by adapted aquatic communities. Thus, thesame reaction, the reduction of Hg2" to Hg0, was employedby whole microbial communities and by a mer(Tn21)-car-rying bacterial strain (36). However, our data do not rejectthe possibility that the aquatic communities that we studiedalso methylated Hg2". Considering the limit of detection (2%of added 203Hg2+) of the analysis of volatile mercurialcompounds, methylation would have been detected only ifmore than 5 jig of dimethylmercury was produced per liter.Reported values for methyl mercury production in naturalwaters are in the nanogram-per-liter range, and methylmercury usually accounts for 1 to 0.1% of added Hg2+. Xunet al. (43) observed conversion of approximately 0.1% ofadded 203Hg2+ to CH3Hg+ in epilimnitic water, and similarvalues were reported by Furutani and Rudd (18) for methyl-ation in unstratified lake waters. Methylation in sediments(14, 33) and by mercury-methylating bacterial strains (16;C. C. Gilmour, E. A. Henry, and R. Mitchell, Abstr. Annu.Meet. Am. Soc. Microbiol. 1988, Q143, p. 306) also resultedin very low concentrations of methyl mercury. Thus, even ifdimethylmercury was produced by the communities that westudied, we would not have detected it.

Bacteria were implicated as the active component in bothfreshwater and estuarine microbial communities. Appar-ently, microeucaryotes were more susceptible to Hg2+ thanwere the bacteria (Table 2). Their elimination in filteredcommunities did not significantly affect Hg2+ loss (Fig. 1).Interestingly, tolerance of primary productivity and hetero-trophic microflagellates to Hg2+ was higher in the freshwatersample than in the saline sample. Additional experiments areneeded to understand this difference in susceptibilities.The prominent role of bacteria in the volatilization of

Hg2+ suggests that bacterial genes in the community genepool coded for Hg2+ reduction. Because the mer operonmediated this reaction in many species of bacteria, andbecause alternative Hg2+ resistance mechanisms have beenrarely reported and not fully investigated (30), it was pre-sumed that mer genes in the aquatic communities mediatedHg2+ volatilization. DNA-DNA hybridization with gram-negative strains representing Hg2+-resistant bacteria in thecommunities that we studied indicated that mer genes withno homology to the mer(Tn2I) gene are largely responsiblefor Hg2+ resistance and volatilization. Further work hasdemonstrated that some of these mer(Tn2J)-negative strainsvolatilized Hg2+ (T. Barkay, manuscript in preparation).

In comparison with saltwater strains, freshwater strainsmore frequently had mer(Tn2l)-homologous DNA se-quences, and some were resistant to a higher Hg2+ concen-tration (Table 3). The latter observation could be explained ifresistance to higher concentrations of Hg2+ had not evolvedin estuarine and marine bacteria, because salt concentra-tions, such as those found in marine and estuarine environ-ments, reduce the toxicity of Hg2+ (4).

Others have reported the absence of DNA sequences withhomology to mer(Tn2I) in resistant bacteria from salineenvironments (2; N. Hamlett, personal communication).Possibly, resistance to Hg2+ in saline environments neces-sitates additional or different gene products than resistancein freshwater environments does. Inorganic mercury in theformer exists mostly as HgCl3-/HgC142, whereas in thelater the hydrated form [Hg(OH)2] dominates (25). Theseforms of mercury may be different substrates for Hg2+-resistant organisms. Transport of mercury is the likely stepat which marine bacteria are different from freshwaterstrains, because osmoregulating mechanisms maintain a

APPL. ENVIRON. MICROBIOL.

on January 3, 2020 by guesthttp://aem

.asm.org/

Dow

nloaded from

mer(Tn21)-MEDIATED Hg2+ REDUCTION IN NATURAL WATERS

stable environment in the cell cytoplasm (26) and the simi-larity between mercuric reductases of gram-negative bacte-ria and those of marine caulobacters has been demonstrated(22). Neither form of mercury, Hg(OH)2 and HgCl3-/HgC142-, diffuses through lipid bilayer membranes (19); andboth require an active transport through the bacterial cellwall.Other environmental factors besides salinity may affect

the distribution of Hg2+ resistance genes, because a signifi-cant difference was also observed between the two salinecommunities (Table 3). More research is needed to deter-mine how environmental factors affect the distribution ofalternative genes that code for biotransformations of pollu-tants.The data presented in Fig. 1 demonstrate that the signifi-

cance of microbial reduction of Hg2+ in natural watersdepends on its relation to physical chemical processes thatinduce volatilization of Hg2+. A rapid loss of Hg2+ wasobserved in filtered freshwater samples. This could be due toreduction and disproportionation (6) or the presence of freeradicals associated with humic acids (1). In some freshwatersamples, nonbiological Hg2+ loss is so rapid that microbialactivity cannot be distinguished (T. Barkay, unpublisheddata). The bacterial role in Hg2+ volatilization is clearer insaline water environments. The loss of Hg2+ in filtered waterwas negligible compared with the loss from biologicallyactive samples. One reason for this difference may be thepresence of Cl- in marine and estuarine waters. Chlorideions inhibit disproportionation when they react with Hg22+to form Hg2Cl2 (calomel) (6).

In conclusion, mer(Tn2J) coded for Hg2+ resistance inapproximately 50% of resistant freshwater strains, but bio-logically induced volatilization had only a marginal ecologi-cal role in this environment because of a strong nonbiologi-cal loss of Hg2+. In contrast, bacterial activity was the majorcause of Hg2+ volatilization in saline water environments inwhich genes homologous to mer(Tn2J) had a limited distri-bution among Hg2+-resistant organisms. Thus, the ecologi-cal significance of mer(Tn21)-mediated reduction of Hg2+ toHgo, in saline water and freshwater environments, dependson interactions of the pollutant with indigenous bacteria andwith environmental factors.

ACKNOWLEDGMENTS

Gratitude is extended to Hap Pritchard, Peter Chapman, RickCoffin, and Ralph Turner for stimulating discussions; Richard Bar-tha and Michael Berman for sending bacterial strains; RichardDevereux and Rick Coffin for help with the growth of D. desulfuri-cans and the microeucaryote experiments, respectively; ChristineDeans and Rick Cripe for help with statistic analysis; and Steve Fossand Yvonne Ford for the illustration.

LITERATURE CITED1. Alberts, J. J., J. E. Schindler, R. W. Miller, and D. E. Nutter, Jr.

1974. Elemental mercury evolution mediated by humic acid.Science 184:895-897.

2. Anast, N., and J. Smit. 1988. Isolation and characterization ofmarine caulobacters and assessment of their potential for ge-netic experimentation. Appl. Environ. Microbiol. 54:809-817.

3. Armitage, P. 1971. Statistical methods in medical research.Blackwell Scientific Publications, Ltd., Oxford.

4. Babich, H., and G. Stotzky. 1979. Differential toxicities ofmercury to bacteria and bacteriophages in sea and lake waters.Can. J. Microbiol. 25:1252-1257.

5. Babich, H., and G. Stotzky. 1986. Environmental factors af-fecting the utility of microbial assays for the toxicity andmutagenicity of chemical pollutants, p. 9-42. In B. J. Dutka and

G. Bitton (ed.), Toxicity testing using microorganisms, vol. II.CRC Press, Inc., Boca Raton, Fla.

6. Baltisberger, R. J., D. A. Hildebrand, D. Grieble, and T. A.Ballintine. 1979. A study of the disproportionation of mercury(I) induced by gas sparging in acidic aqueous solutions forcold-vapor atomic absorption spectrometry. Anal. Chim. Acta111:111-122.

7. Barkay, T. 1987. Adaptation of aquation microbial communitiesto Hg2+ stress. Appl. Environ. Microbial. 53:2725-2732.

8. Barkay, T., D. L. Fouts, and B. H. Olson. 1985. Preparation ofa DNA gene probe for detection of mercury resistance genes ingram negative bacterial communities. Appl. Environ. Micro-biol. 49:686-692.

9. Barkay, T., and P. H. Pritchard. 1988. Adaptation of aquaticmicrobial communities to pollutant stress. Microbiol. Sci. 5:165-169.

10. Barkay, T., and G. S. Sayler. 1988. Gene probes as a toll for thedetection of specific genomes in the environment, p. 29-36. InW. J. Adams, G. A. Chapman, and W. G. Landis (ed.), Aquatictoxicology and hazard assessment, 10th vol. ASTM STP 971.American Society for Testing and Materials, Philadelphia.

11. Ben-Bassat, D., and A. M. Mayer. 1977. Reduction of mercurychloride by Chlorella: evidence for a reducing factor. Physiol.Plant 40:157-162.

12. Ben-Bassat, D., and A. M. Mayer. 1978. Light-induced Hgvolatilization and 02 evolution in Chlorella and the effect ofDCMU and methylamine. Physiol. Plant. 42:33-38.

13. Caron, D. A. 1983. Technique for enumeration of heterotrophicand phototrophic nanoplankton, using epifluorescence micros-copy, and comparison with other procedures. Appl. Environ.Microbiol. 46:491-498.

14. Compeau, G., and R. Bartha. 1984. Methylation and demethyl-ation of mercury under controlled redox, pH, and salinityconditions. Appl. Environ. Microbiol. 48:1203-1207.

15. Compeau, G., and R. Bartha. 1985. Sulfate-reducing bacteria:principal methylators of mercury in anoxic estuarine sediment.Appl. Environ. Microbiol. 50:498-502.

16. Compeau, G., and R. Bartha. 1987. Effect of salinity on mercu-ry-methylating activity of sulfate-reducing bacteria in estuarinesediments. Appl. Environ. Microbiol. 53:261-265.

17. Cripe, C. R., W. W. Walker, P. H. Pritchard, and A. W.Bourquin. 1987. A shake-flask test for estimation of biodegrad-ability of toxic organic substances in the aquatic environment.Ecotoxicol. Environ. Safety 14:239-251.

18. Furutani, A., and J. W. M. Rudd. 1980. Measurement ofmercury methylation in lake water and sediment samples. Appl.Environ. Microbiol. 40:770-776.

19. Gutknecht, J. 1981. Inorganic mercury (Hg2+) transport throughlipid bilayer membranes. J. Membr. Biol. 61:61-66.

20. Hobbie, J. E., R. J. Daley, and S. Jasper. 1977. Use ofNuclepore filters for counting bacteria by fluorescence micros-copy. Appl. Environ. Microbiol. 33:1225-1228.

21. Jain, R. K., R. S. Burlage, and G. S. Sayler. 1988. Methods fordetecting recombinant DNA in the environment. Crit. Rev.Biotechnol. 8:33-84.

22. Ji, G., S. P. Salzberg, and S. Silver. 1989. Cell-free mercuryvolatilization activity from three marine Caulobacter strains.Appl. Environ. Microbiol. 55:523-525.

23. Kellogg, S. T., D. K. Chatterjee, and A. M. Chakrabarty. 1981.Plasmid-assisted molecular breeding: new technique for en-hanced biodegradation of persistent toxic chemicals. Science214:1133-1135.

24. Kimura, Y., and V. L. Miller. 1960. Vapor phase separation ofmethyl- or ethylmercury compounds and metallic mercury.Anal. Chem. 32:420-424.

25. Krenkel, P. A. 1974. Mercury: environmental considerations,part II. Crit. Rev. Environ. Control 4:251-339.

26. Le Rudulier, D., A. R. Strom, A. M. Dandekar, L. T. Smith, andR. C. Valentine. 1984. Molecular biology of osmoregulation.Science 224:1064-1068.

27. Longbottom, J. E., R. C. Dressman, and J. J. Lichtenberg. 1973.Metals and other elements: gas chromatographic determinationof methyl mercury in fish, sediment, and water. J. Assoc. Off.

VOL. 55, 1989 1201

on January 3, 2020 by guesthttp://aem

.asm.org/

Dow

nloaded from

APPL. ENVIRON. MICROBIOL.

Agric. Chem. 56:1297-1303.28. Olson, B. H., and R. C. Copper. 1976. Comparison of aerobic

and anaerobic methylation of mercuric chloride by San Fran-cisco Bay sediments. Water Res. 10:113-116.

29. Rosenau, J. C., G. L. Faulkner, C. W. Hendry, Jr., and R. W.Hull. 1977. Springs of Florida. Florida Department of NaturalResources and Bureau of Water Resources Management, Flor-ida Department of Environmental Regulation, Tallahassee.

30. Silver, S., and T. Misra. 1988. Plasmid-mediated heavy metalresistances. Annu. Rev. Microbiol. 42:717-743.

31. Spain, J. C., P. H. Pritchard, and A. W. Bourquin. 1980. Effectsof adaptation on biodegradation rates in sediment/water coresfrom estuarine and freshwater environments. Appl. Environ.Microbiol. 40:726-734.

32. Spain, J. C., P. A. Van Veld, C. A. Monti, P. H. Pritchard, andC. R. Cripe. 1984. Comparison of p-nitrophenol biodegradationin field and laboratory test systems. Appl. Environ. Microbiol.48:944-950.

33. Steffan, R. J., E. T. Korthals, and M. R. Winfrey. 1988. Effectsof acidification on mercury methylation, demethylation, andvolatilization in sediments from an acid-susceptible lake. Appl.Environ. Microbiol. 54:2003-2009.

34. Strickland, J. D. H., and T. R. Parsons. 1972. A practicalhandbook of seawater analysis. Fisheries Research Board ofCanada, Ottawa, Ontario.

35. Summers, A. 0. 1986. Organization, expression, and evolutionof genes for mercury resistance. Annu. Rev. Microbiol. 40:607-634.

36. Summers, A. O., and S. Silver. 1972. Mercury resistance in aplasmid-bearing strain of Escherichia coli. J. Bacteriol. 112:1228-1236.

37. Summers, A. O., and S. Silver. 1978. Microbial transformationsof metals. Annu. Rev. Microbiol. 32:637-672.

38. Topping, G., and I. M. Davies. 1981. Methylmercury productionin the marine water column. Nature (London) 290:243-244.

39. Toribara, T. Y., C. P. Shields, and L. Koval. 1970. Behavior ofdilute solutions of mercury. Talanta 17:1025-1028.

40. U.S. Environmental Protection Agency. 1974. Methods for chem-ical analyses of water and wastes, p. 118-137. U.S. Environ-mental Protection Agency, Washington, D.C.

41. Weiss, A. A., S. D. Murphy, and S. Silver. 1977. Mercury andorganomercurial resistance determined by plasmids in Staphy-lococcuis aiureus. J. Bacteriol. 132:197-208.

42. Wyndham, R. C. 1986. Evolved aniline catabolism in Acineto-baicter calcoaceticus during continuous culture of river water.Appl. Environ. Microbiol. 51:781-789.

43. Xun, L., N. E. R. Campell, and J. W. M. Rudd. 1987. Measure-ments of specific rates of net methyl mercury production in thewater column and surface sediments of acidified and circumneu-tral lakes. Can. J. Fish. Aquat. Sci. 44:750-757.

1202 BARKAY ET AL.

on January 3, 2020 by guesthttp://aem

.asm.org/

Dow

nloaded from