APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Dec. 1987, p. 2725-27320099-2240/87/122725-08$02.00/0Copyright © 1987, American Society for Microbiology

Vol. 53, No. 12

Adaptation of Aquatic Microbial Communities to Hg2" StresstTAMAR BARKAY

Microbial Ecology and Biotechnology Branch, Environmental Research Laboratory, U.S. Environmental ProtectionAgency, Gulf Breeze, Florida 32561

Received 26 May 1987/Accepted 28 August 1987

The mechanism of adaptation to Hg2+ in four aquatic habitats was studied by correlating microbiallymediated Hg2+ volatilization with the adaptive state of the exposed communities. Community diversity,heterotrophic activity, and Hg2+ resistance measurements indicated that adaptation of all four communitieswas stimulated by preexposure to Hg2+. In saline water communities, adaptation was associated with rapidvolatilization after an initial lag period. This mechanism, however, did not promote adaptation in a freshwatersample, in which Hg2+ was volatilized slowly, regardless of the resistance level of the microbial community.Distribution of the mer operon among representative colonies of the communities was not related to adaptationto Hg2+. Thus, although volatilization enabled some microbial communities to sustain their functions inHg2+-stressed environments, it was not mediated by the genes that serve as a model system in molecular studiesof bacterial resistance to mercurials.

Adaptation governs the activity of natural microbial com-munities in human-affected environments. Such environ-ments are subjected to a myriad of toxic xenobiotic com-pounds that challenge the integrity of the indigenous flora.Typically, the community responds to this challenge by theevolution of resistant populations (7, 14) that sustain micro-bial processes (13, 19) in the presence of the inhibitorytoxicants. Further, adaptation processes may relieve stressby biodegradation of toxicants (29, 34). Evolution of resis-tant populations and biodegradation have been studied inde-pendently, leaving unanswered the important question ofhow elimination of inhibitors promotes continuation of es-sential microbial functions. This relationship may play a keyrole in adaptation to toxic biotransformable pollutants, suchas some heavy metals (13, 36) and pesticides (16).Mechanisms of biotransformation have been investigated

with pure cultures isolated from impacted environments. Forexample, biodegradation of 3-chlorobenzoate was studiedwith Pseudomonas cepacia isolated from a sewage sample(10), and bacterial reduction of Hg2+ to volatile Hgo wasstudied with organisms exposed to mercurial disinfectants inthe hospital environment (32). The significance of specificbiotransformation mechanisms in the response of naturalmicrobial communities to stress has been examined to alimited extent. This is an important issue because studieswith pure cultures may lead to erroneous conclusions. Forexample, based on theoretical considerations and pure cul-ture studies with cell extracts of a methanogen (41), it haslong been assumed that methylation of Hg2+ in anaerobicenvironments was due to activity of methanogenic orga-nisms (36). However, when methyl mercury formation wasstudied with whole sediment communities, sulfate reducerswere found to carry out this activity and, contrary toexpectations, specific inhibition of methanogens stimulatedmethylation (11). Utilizing information obtained with purecultures to explore ecological phenomena could confer un-derstanding of how molecular mechanisms operate in situ atthe community level. This is especially important for theprospect of using genetically engineered organisms as ameans of environmental management (16).

t Contribution no. 608 of the U.S. Environmental ResearchLaboratory, Gulf Breeze, Florida.

Mercury is an excellent model stressor for study of howbiotransformation of a toxicant can affect the adaptiveresponse of exposed communities. Mercuric ion (Hg2+)alters microbial community structure (33) and function (19),and it can be biotransformed by microorganisms to a lesstoxic form, Hg°. The molecular details of this biotransforma-tion are well understood (35). Some mercury-resistant orga-nisms possess a cytoplasmic NADPH-dependent mercuricreductase that transfers two electrons to Hg2+. The resultingHg0 is lost by volatilization owing to its high vapor pressure.Therefore, Hg2+ reduction is a detoxification mechanism.The gene coding for mercuric reductase is a part of the meroperon together with genes coding for transport of Hg2+ intothe cell and regulation of the expression of the operon (35).This mechanism has been shown to confer resistance uponorganisms isolated from natural waters (25, 26), soils (20),and the clinical environment (32). Biologically mediatedmercury evolution has been observed in soils (30) andnatural waters (25) spiked with mercurials. Informationregarding the kinetics of mercury evolution by environmen-tal samples, or the nature of the mechanisms which mediateit, is not available.A DNA gene probe consisting of the widely studied mer

operon was constructed to study distribution of this genesystem in environmental microbial communities (6). It wassubsequently shown that mer genes are more widely dis-tributed in microbial communities of mercury-laden sedi-ments than in communities of unimpacted sediments (5). Thepresent investigation reveals relationships among volatiliza-tion of Hg2+, adaptive response of aquatic communities toHg2+ stress, and distribution of the mer gene that codes forboth volatilization and resistance.

MATERIALS AND METHODS

Sampling locations and procedures. Sampling sites in thevicinity of Pensacola, Fla., represented four types of aquaticenvironment (Fig. 1). Santa Rosa Sound is an estuarylocated between the Florida mainland and Santa Rosa Is-land, a barrier island. It is subjected to active small-boattraffic and to runoff from the nearby municipality, but nomajor industrial or domestic wastes enter this system. TheRange Point site is a small enclosed salt-marsh pond located

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FIG. 1. Area map of sampling locations in the vicinity of Pensacola, Fla. (A) Estuary, Santa Rosa Sound; (B) salt marsh, Range PointPond; (C) fresh water, Thompson's Bayou; (D) coastal marine, Gulf of Mexico.

on Santa Rosa Island. The source of water in the pond isrunoff and seepage waters from Santa Rosa Sound. The pondis not subjected to pollution and supports abundant aquaticvegetation. The third sampling site, Thompson's Bayou, is afreshwater tributary of the Escambia River. It is part of anature sanctuary located on the University of West Floridacampus and is protected from exogenous effects. Watersamples representative of a coastal marine environmentwere collected in the Gulf of Mexico at Pensacola Beach.Water samples were brought in sterile flasks to the labo-

ratory for processing. Water temperature was obtained witha thermometer at the site during sampling. Water pH wasdetermined in the laboratory with a pH meter (model 044;Beckman Instruments, Inc., Fullerton, Calif.). Salinity wasmeasured with a salinometer (model TS; Reichert ScientificInstruments, Buffalo, N.Y.), and samples for total organiccarbon determination were stored in acid-washed (1 N HCl)glass tubes at -20°C until analyzed with a model 0524C TotalCarbon System (Oceanograph International Corp., CollegeStation, Tex.) as recommended by the manufacturer.Sample incubation. Samples were divided into four

subsamples (200 ml each) and dispensed into 250-ml gradu-ated glass cylinders (Kimax; Kimble Products, Toledo,Ohio) whose spouts had been removed (remaining cylinderdepth was 27 to 29 cm). The remainder of each sample wasstored at 4°C for future use. Each cylinder was plugged withan 8817-A Neoprene stopper (size 8; Arthur H. Thomas Co.,Philadelphia, Pa.) through which a stainless-steel needle wasinserted and adjusted to reach the bottom of the cylinder.Aeration of the samples was by bubbling water-saturatedand filter-sterilized compressed air through Tygon tubingconnected to the needle by a one-way syringe stopcock. Asampling port was constructed by inserting a glass tube (3 cm

long, 9 mm in diameter) into the stoppers. This port wasplugged with a smaller (size 00) Neoprene stopper that hadan air outlet. Air leaving the samples was passed throughTygon tubing to a trap containing 3.6 N H2SO4 and 0.28 mMKMnO4 for removal of volatile mercury.

Two subsamples were dosed with Hg2+ by the addition of5 p.l of HgCl2 stock solution (10 mg/ml as Hg2+) to a finalconcentration of 250 p.g/liter. The microbial communities ofthese samples were called preexposed communities. NoHg2+ was added to the two remaining subsamples used ascontrol communities (called control communities). All cyl-inders were incubated in a water bath at 29°C with a slowbubbling of air. After 40 to 48 h of incubation, samples werecentrifuged at 5,858 x g in 250-ml bottles at 4°C in a SorvallRC-2B Superspeed centrifuge (Dupont Instruments, Wilming-ton, Del.), and pellets were rinsed once with 10 ml of cold0.85% NaCl solution and suspended in 200 ml of filter-sterilized (0.22-p.m pore size) sample water that had beenstored at 4°C during the preexposure period. Samples werekept on ice during this procedure, which yielded 100%recovery of CFU. The following analyses were performedwith the cell suspensions.

(i) Fate of Hg2+. Samples (25 ml) of cell suspension weredispensed into 30-ml graduated glass impingers (WheatonIndustries, Millville, N.J.). Water-saturated sterile air (pre-pared as described above) was bubbled through the samples,and the air leaving the systems was passed through rubbertubing to a trap containing 10 ml of freshly prepared trappingsolution (see above) to remove volatile mercurial com-pounds. 203Hg2+ stock solution was prepared by mixing203HgC12 (Amersham Corp., Arlington Heights, Ill.) withcold HgCl2 (10 mg/ml as Hg2+) to a specific activity of 100pCi/mg of Hg2+ and a concentration of 250 pg of Hg2+ perml. This solution was added to the 25-ml samples to yield afinal Hg2+ concentration of 250 p.g/liter. Samples (1 ml) wereremoved immediately for time zero measurements, and theremainder was incubated at 29°C. Samples for Hg2+ remain-ing were taken periodically (times indicated in Fig. 2), mixedwith 10 ml of PSC scintillation cocktail (Amersham), andcounted with a Beckman LS250 liquid scintillation counterprogrammed for quench correction with standards providedby the manufacturer. Traps were replaced after each sam-pling, and 1-ml samples were removed and counted to

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TABLE 1. Physicochemical parameters of the studied aquatic environments

Dateof Salinity ~~~~~~~~~~~~~~~~~~~TotalorganicEcosystem Date of pH Temp ('C) Salinity carbon (,ag/sampling ~~~~~~~~~~~~~~~~~~~~ml,± SD)

Estuarine 5/31/86 8.10 29 26 8.7 + 2.2Salt marsh 7/14/86 8.81 30.5 27.5 7.3 + 1.8Freshwater 8/04/86 6.39 25 0 4.35 + 0.5Coastal marine 9/09/86 8.24 29.5 34.5 ND"

" ND, Not determined.

account for evolved mercurials. A filter-sterilized (0.22-p.mpore size) water sample was used as a control for the fate ofHg2+ in biologically inactive samples. Removal of mercuryfrom solution by adhesion to glass walls was examined byrinsing impingers with concentrated nitric acid as suggestedby Jenne and Avotins (18). Radioactivity removed by thistreatment regularly amounted to less than 1% of the amountadded at the beginning of the experiment.

(ii) Total and Hg2+-resistant aerobic heterotrophic counts.Cell suspensions were diluted serially in 0.85% NaCl andspread on solid media for total colony counts and on similarmedia containing HgCl2 (10 p.g/ml as Hg2+) for enumerationof resistant colonies. Media were selected according to theorigin of each sample. Culturable aerobic heterotrophicbacteria in estuarine and salt-marsh samples were enumer-ated on plate count agar (Difco Laboratories, Detroit, Mich.)modified with an estuarine salt solution (26). Freshwater andcoastal marine bacteria were enumerated on plate count agarand Marine Agar 2216 (Difco), respectively. Plates wereincubated for 48 to 72 h at room temperature before enumer-ation.

(iii) Direct bacterial counts. Samples (2 ml) of cell suspen-sions were fixed, stained with acridine orange solution,filtered through black polycarbonate membrane filters(0.2-p.m pore size; Nuclepore Corp., Pleasanton, Calif.), andcounted with a Nikon Optiphot microscope equipped with anEF-D fluorescence attachment (Nippon Kogakuk K.K.,Tokyo, Japan) as described by Hobbie et al. (17).

(iv) Incorporation of [3H]TdR into cellular material and itsinhibition by Hg2'. The test procedure of Jonas et al. (19)was modified as follows. [methyl-3H]thymidine ([3H]TdR;specific activity, 78.3 Ci/mmol; New England NuclearCorp., Boston, Mass.) was added to 5 ml of cell suspensionto a final concentration of 3.2 pmollml. Incorporation wasfollowed in duplicates at room temperature. A 2% formalde-hyde-treated control was included to determine nonbiologi-cal uptake. Incubation continued for periods which did notexceed the linear stage of the incorporation reaction (0.5 to3 h) as determined by preliminary experiments. Reactionswere terminated, and samples were prepared for counting asdescribed by Jonas et al. (19). Filters were transferred toscintillation vials, dissolved in 1 ml of ethyl acetate, andcounted in 10 ml of PCS cocktail as described above for203Hg. The Hg2" concentration which inhibited the incorpo-ration rate of [3H]TdR by 50% (50% inhibitory dose [ID50])was determined by adding 5 p.l of HgCl2 stock solution to cellsuspensions before the addition of [3H]TdR. The final Hg2+concentrations were 0, 0.1, 1.0, 10, and 100 p.g/liter forcontrol communities and 0, 100, 1,000, and 5,000 p.g/liter forpreexposed communities. ID50 values were calculated asdescribed by Jonas et al. (19).

(v) Community structure analysis. Shannon-Weaver diver-sity indices were calculated by using the natural logarithm asdescribed by Barkay et al. (7), except that the morphology ofbacterial colonies grown on solid medium was used to

characterize individual strain types as described by Maki etal. (23).

(vi) Distribution of mercury resistance gene system (mer).Colonies were lifted on nitrocellulose filters (BA85, 82 mm;Schleicher & Schuell, Inc., Keene, N.H.) by placing thefilters on the agar of petri dishes containing 30 to 300colonies. The filters remained in contact with the surface fora few minutes until they were wetted completely. Duplicatefilters were obtained from each plate. Bacteria in the colo-nies were lysed immediately, and their DNA was fixed to thesupport filters as described by Berent et al. (8). One filterwas hybridized with the mer probe (6) and the other withplasmid pERC-1, which contained the rrnH operon of Esch-erichia coli (24). Procedures and materials for nick transla-tion of the probes, hybridization reactions, treatment offilters after hybridization, and exposure to X-ray films wereas described previously (6). Autoradiograms of duplicatefilters reacted with the two probes were compared, and onlycolonies that hybridized with the rRNA gene probe but notwith the mer probe were recorded as mer negative. Coloniesthat did not hybridize with the rRNA gene probe wereconsidered to have escaped lysis and were excluded from theanalysis.Data analysis. Means and standard deviations for the

results obtained with all the tests for duplicate subsamples ofpreexposed and control communities are reported. Ratesand statistical significance of 203Hg2" volatilization wereobtained by an SAS (statistical analysis system; SAS Insti-tute, Cary, N.C.) regression analysis. Half-life (t1/2) wascalculated as t12 = -0.693/-K1, where K1 is the first-orderrate constant obtained from the slopes of the lines describingthe loss of Hg2+ over time. A general linear model procedure(SAS) was used to compare slopes of two or more Hg2+ losslines.

RESULTS

Adaptation experiments were performed at least twice foreach community. The results presented are for each com-munity experiment, for which a more complete set of datawas obtained.

Physicochemical characterization of samples. The fourphysicochemical parameters measured are known to affectthe activity of aquatic microbes as well as the chemical formand thus bioavailability of mercury (21). Redox potential,which affects the chemical form of mercurial compounds,was not measured because incubation of the samples in thelaboratory was under fully aerated conditions. The estuarineand salt-marsh samples were very similar to each other, withpH values around 8.0, temperature of approximately 30°C,salinity of 26 to 27%o, and total organic carbon of 7 to 9 p.g/ml(Table 1). The freshwater sample had a slightly acidic pHand was at 25°C at the time of sampling; its salt content wasbelow the level of detection, and it contained approximatelyhalf as much organic carbon (4.35 ± 0.5 p.g/ml) as the saline

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CY 201 20-

O12 24 6 2 24

S D. COASTAL MRNE

< )00 ICFRS6R1224 0 6 122

w

standarddevation (bars)

60- ~~~~~~~60-

036 12 ~24 12 24

TIME (HOURS)

FIG. 2. Loss of Hg2l from water samples spiked with 250 p.g of Hg2+ per liter. Symbols: loss mediated by preexposed communities;

0, loss mediated by control communities; A, loss mediated by filter-sterilized (0.2-~~.m pore size) water samples. Error corresponds to±

standard deviation (bars).

water samples. The physicochemical parameters of thecoastal marine sample were similar to those of the othersaltwater samples, except that salinity was higher (34.5%o).

Volatilization of 203Hg2' by aquatic microbial communities.The effect of preexposure on the volatilization of Hg2" was

tested by following the loss of 203Hg2+ mediated bypreexposed and control communities (Fig. 2). The majorityof added Hg2+ was volatilized during 24 h of exposure.

Mercury loss curves followed first-order kinetics (Fig. 2),and half-life values obtained by regression analysis are

presented in Table 2. Volatilized mercury was accumulatedin traps containing KMnO4 and sulfuric acid, but no corre-

lation between amounts of Hg2+ lost and mercury evolvedcould be obtained. Since the mass balance of mercury couldnot be calculated, quantitative data describing mercuryevolved are not presented.Two patterns of loss were observed. (i) Rapid loss was

mediated by the preexposed community, with a delayedreaction mediated by the control community after an initiallag phase. This pattern, demonstrated by the estuarine andcoastal marine communities (Fig. 2A and D, respectively),

indicates adaptation. The preexposed estuarine communityvolatilized Hg2" with a t112 of 4.8 + 0.45 h. The controlcommunity, after a lag of 6 to 12 h, reacted at similar rate,with a t1/2 of 5.0 ± 0.56 h. Initially, Hg2+ was lost from thecontrol sample at a rate (t1/2 of 30.1 ± 9.16 h) approximatelythat of the sterile control (52.1 ± 7.44 h), suggesting that lossduring the lag period was due largely to nonbiologicalprocesses. A similar pattern was observed with the coastalmarine sample: rapid volatilization mediated by the pre-exposed community (41/2 of 5.5 ± 0.7 h) with a delayed (lagperiod of 12 to 24 h) response of the control community (t1/2of 8.0 ± 0.18 h). Hg2+ loss occurred slowly in the controlsample during the lag stage (41/2 of 86.6 ± 10.83 h) and in thesterile sample during the entire experiment (4112 of 247 ±

114.9 h). Adaptation was also apparent with the salt-marshcommunity, as preexposure resulted in rapid volatilization(4112 of 8.6 ± 1.8 h; Fig. 2B). However, Hg2+ loss by thecontrol community did not have two clear phases. This maybe due to substantial loss which occurred in the absence ofbiological activity, as is evident by the sterile control. Lossfrom this sample (4112 of 23.9 ± 2.97 h) was only slightly

TABLE 2. Half-lives of Hg2" from samples containing preexposed and control communities and from sterile water samples

tl/2 (h)"Sample

Preexposed Control' Sterile

Estuarine 4.8 ± 0.45 30.1 ± 9.16 (early) 52.1 ± 7.445.0 ± 0.56 (late)

Salt marsh 8.6 ± 1.80 16.1 ± 0.82 23.9 ± 2.97Freshwater 15.8 ± 9.34 19.8 ± 2.52 34.7 ± 3.12Coastal marine 5.5 ± 0.70 86.6 ± 10.83 (early) 247.0 ± 114.90

8.0 ± 0.18 (late)a 1/2 calculated by regression analysis from the plot of In Hg2 concentration over time (see Materials and Methods).bt1/2 for Hg2+ loss mediated by the estuarine and coastal marine control communities were calculated for the two stages of the reaction: early, the lag phase;

late, the active phase.

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TABLE 3. Structural and functional parameters describing the adaptive state of preexposed and control communities

13HITdRCommunity TVC (CFU/ml)" TVC-Hg' (CFU/ml)5 Direct counts incorporattion IDa, Shannon-Weaver

(cells/ml)" i-ate (pmol/h per (,ug of Hg/liter) diversity index'liter)

EstuarinePreexposed (2.2 ± 1.1) X 105 (1.5 ± 1.1) x 105 (68) NA' 113.2 ± 96.1 1.800 ± 141 1.013 (721)Control (8.2 ± 8.2) x 104 (2.6 ± 1.0) X 103 (3) NA 83.9 ± 9.1 5.9 ± 1.0 2.010 (721)

Salt marshPreexposed (4.7 ± 1.3) x 105 (4.1 ± 2.1) x 105 (87) (2.2 ± 0.6) x 106 18.4 + 7.9 36.5 ± 33.2 0.5355 + 0.55 (61)Control (1.4 ± 0.8) X 103 (1.3 ± 0.6) x 102 (9) (2.1 ± 0.7) x 106 NA NA 1.2280 t 0.49 (61)

FreshwaterPreexposed (8.3 ± 6.0) X 104 (6.0 ± 4.8) X 104 (72) (1.5 ± 0.4) x 106 NA NA 0' (100)Control (4.4 ± 1.5) x 103 (1.4 ± 0.2) x 102 (3) (9.5 + 5.0) x 105 NA NA 1.805 ± 0.57 (100)

Coastal marinePreexposed (2.1 ± 2.0) x 105 (2.6 ± 3.1) X i0s (124) (4.0 ± 1.0) x i05 149.9 ± 192.0 897.5 + 434.9 0.6811 ± 0.16 (100)Control (2.0 ± 0.1) x 104 (4.5 ± 2.4) x 103(23) (4.4 + 0.2) x 105 13.48 0.17 2.2760 (100)

a Total viable counts (CFU) of aerobic heterotrophic culturable bacteria.b Total viable counts resistant to Hg2. Numbers in parentheses denote percent TVC.' Acridine orange direct counts.d Numbers in parentheses indicate size of analyzed community.e NA, Not available (see discussion).f Community composed of a single strain type.

slower than that from the control community (t112 of 16.1

0.82 h). They differed significantly at the 0.0042 level, andafter 48 h of incubation, 17.0 + 5.2 and 29.9 ± 2.1% of theadded Hg2+ remained in control and sterile samples, respec-

tively. The substantial nonbiological Hg2+ loss from thesalt-marsh sample may have been due to humic acids. Freeradical electrons of these compounds reduce Hg2+ to Hgo(1), and the dark color of the salt-marsh waters suggestedtheir abundance.

(ii) The second pattern of volatilization is represented bythe freshwater community (Fig. 2C). Preexposed and controlcommunities volatilized Hg2+ at the same rate (differencenot significant at the 0.2363 level) with tl2s of 15.8 ± 9.34and 19.8 ± 2.52 h for the preexposed and control communi-ties, respectively (Table 2). These rates were lower thanthose observed for saline preexposed communities. Loss ofHg2+ owing to nonbiological processes (i.e., sterile sample)was at a rate significantly slower (at the 0.001 level) than thatof the preexposed and control communities (t1,2 of 34.7 ±

3.12 h). This suggests that although biological volatilizationof Hg2+ contributed significantly to cleansing of the studiedfreshwater environment, adaptation was not involved in thisprocess.

Adaptive state of preexposed and control communities. Thecapacity of aquatic communities to maintain structure andfunction in the presence of Hg2+ was defined by comparingselective characteristics of preexposed and control commu-nities (Table 3). Since the four communities respondedsimilarly, the following is a general description of observedresults. Examples of specific communities are given whenpertinent. Preexposure resulted in the following.

(i) Increased potential for heterotrophic activity. All butthe estuarine preexposed communities contained at least 1order of magnitude more heterotrophic aerobic culturableorganisms as compared with control communities. Thisincrease was at the expense of nonculturable organisms inthe community; total biomass as indicated by direct micro-scopic counts was not affected by preexposure. For exam-

ple, biomass of the salt-marsh community was approxi-

mately 2.1 x 106 cells per ml. In the preexposed adaptedcommunity about 20% (4.7 x 105 CFU/ml) were aerobicheterotrophs, whereas the corresponding value for the con-trol community was only 0.1% (1.4 x 103 CFU/ml). Thisincrease in CFU may correspond to increased heterotrophicactivity as suggested by higher incorporation rates of[3H]TdR by preexposed communities (Table 3).

(ii) Increased tolerance to Hg2+ as shown by a highernumber of Hg2'-resistant heterotrophic aerobic organismsas well as by increased ID50s for preexposed communities.Between 68% (for estuarine) and 124% (for coastal marine)of the CFU of preexposed communities were resistant,whereas only 3% (for estuarine) to 23% (for coastal marine)of CFU of control communities grew on Hg2'-containingmedia (Table 3). This observation corresponds to an in-creased tolerance as indicated by the effect of Hg2+ onincorporation of [3H]TdR.

(iii) Decreased diversity of preexposed communities asdemonstrated by smaller Shannon-Weaver indices, indicat-ing that these communities developed in the presence of astressor. An extreme example of this was provided by thefreshwater community with an index of 0 (i.e., one straintype) for the preexposed adapted community and an index of

TABLE 4. Distribution of mer in preexposed and control aquaticcommunities

Community Date of (iner/lmll)Community smlnsampling Preexposed Control

Estuarine 5/31/86 1.7 x 104 <104Salt marsh 6/20/86 <i0lo <102

7/14/86 < lo,3 <lo,Freshwater 8/04/86 <103 2.5 x 102

8/17/86 (8.5 + 9) x 102 (2.9 + 5) x 102Coastal marine 8/25/86 <102 <101

9/09/86 <103 <10'

Numbers are of colonies grown on plate count agar which were hybridizedto the ,ner probe (Materials and Methods).

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1.805 ± 0.57 (accounting for 13 to 16 strain types) for thecontrol community (Table 3).

Distribution of mer gene systems. If the mer operon codedfor Hg2" resistance and volatilization, its distribution inpreexposed communities would be increased. Results ofduplicate experiments are presented in Table 4. Distributionof mer in one of the estuarine experiments was detected byrandom transfer of colonies to nitrocellulose filters, and only2 of 260 (0.8%) and 12 of 536 (2.2%) of preexposed andcontrol colonies, respectively, were mer positive. Coloniescontaining DNA sequences homologous to mer (as indicatedby positive hybridization signals) were infrequently ob-served in estuarine and freshwater communities. Distribu-tion of mer did not correlate with the number of Hg2+-resistant aerobic heterotrophs in these communities (Table3). For example, mer in the preexposed estuarine commu-nity (1.7 x 104/ml) could account for only 10% of theresistant heterotrophs (1.5 x 105/ml). These results suggestthat as far as sensitivity of the method used allows (seebelow), mer, even in communities in which it is present,contributes very little to community response to Hg2".

DISCUSSION

The adaptive response of four aquatic microbial commu-nities to Hg2+ stress was described by following bio-transformation of Hg2+ to a volatile form, by its effect onstructure and function of the community, and by distributionof the gene system which codes for the specific detoxifica-tion process. It is hoped that this approach will lead tounderstanding of how a specific molecular mechanism me-diates a process of ecological significance to sustain micro-bial activities in a stressed environment.The concentration of Hg2+ (250 jLg/liter) used in this study

was selected because preliminary experiments in whichcommunities were exposed to gradually increased concen-trations indicated that their response to this dose clearlydistinguished between adapted and unadapted communities.Although this concentration is higher than that reported forcontaminated natural waters (15, 25), it is likely that com-munities of such environments are exposed occasionally tohigher concentrations because inorganic mercury is rapidlyremoved from solution by adsorption to particulates (21, 31).The techniques of Spain et al. (34) to show adaptation of

estuarine communities to p-nitrophenol were used to studyadaptation to Hg2+. Two patterns of Hg2+ volatilizationwere observed. The first, common to all saline communities,consisted of a period of adaptation followed by loss owing torapid volatilization. In the second, no adaptation period wasrequired for onset of this process by a freshwater community(Fig. 2; Table 2). These responses of the microbial flora maybe explained by physicochemical parameters of the aquaticenvironment (Table 1) which determine chemical form andthus toxicity. Speciation of mercury as a function of pH andsalinity (21) indicated that under conditions which existed inthe freshwater samples (little salt and pH below 7.0), thehydrated form, Hg[OH12, was dominant. On the other hand,at salinities above 20%c and at pH 8.0 or above (as observedfor the three saline samples), HgCI3- and HgC142- were mostabundant. Babich and Stotzky (4) demonstrated that toxicityof Hg2+ to pure bacterial cultures was lower in brothcontaining salt or in seawater, as compared with that in brothwithout salt or in lake water. Accordingly, it is suggestedthat toxicity of Hg2+ in the freshwater sample was higherthan that in the saline samples. Increased toxicity may haveelicited an immediate, albeit slow, volatilization of the toxic

element (Fig. 2). Alternatively, other factors which variedamong the studied water samples, such as microenviron-ments, particulate matter, and presence of toxicants, couldhave affected the activity of the microbial communities andthe chemical form of mercury.Response of aquatic communities to Hg2+ stress was

defined by changes in structure and function and by anincrease in the tolerance level to Hg2+ of preexposedsubsamples. Increase in tolerance of all four test communi-ties resulted from exposure to Hg2+. This result was ex-pected because of observations indicating enrichment ofresistant populations in communities in metal-contaminatedenvironments (12-14, 25, 27) and evolution of resistance inmicrocosm systems (38). This study goes a step further byconsidering the nature of the resistance mechanism. Corre-lation between developed resistance and rapid inducible lossof Hg2+ in saline communities suggests that tolerance wasinduced by mechanisms that increased the rate of moleculardisassociation followed by volatilization. If so, this demon-strates that a specific molecular mechanism is responsiblefor adaptation of natural communities to Hg2+. However,this discussion remains speculative because appearance ofresistance to Hg2+, and its volatilization, could be indepen-dent events both triggered by preexposure. Volatilizationclearly was not the resistance mechanism in the freshwatercommunity, because both preexposed and control sub-samples volatilized Hg2+ at the same rate (Fig. 2C; Table 2)despite the lower tolerance of the latter (Table 3). Alterna-tive resistance mechanisms, such as accumulation of mer-cury by resistant cells (40) or reduced permeability to Hg2+owing to the synthesis of new membrane proteins (28), couldbe active in aerobic aquatic environments. Resistance in thefreshwater community may be due to either of those mech-anisms or additional, as yet unexplored, mechanisms.

Diversity indices may serve two purposes: indication ofstress and insight into the functional status of the community(3). In the present study, preexposed communities hadsmaller diversity indices, suggesting that they developedunder the influence of a stressor. This decline in diversityhad no inhibitory effect on at least one essential microbialfunction, heterotrophic activity (Table 3). On the contrary,this activity was stimulated by preexposure: preexposedcommunities had a higher rate of [3H]TdR incorporation anda greater number of heterotrophic aerobic culturable orga-nisms as compared with the control communities. Similarstimulation of microbial activities by Hg2+ was reported byCapone et al. (9), who observed an increase in methan-ogenesis and CO2 evolution in salt-marsh sediments supple-mented with 1,000 ppm (1,000 ,ug/g) HgCl2. An explanationfor this phenomenon was suggested by Vaccaro et al. (39),who observed stimulation of the rate of ['4C]glucose assim-ilation by Cu2+-stressed aquatic microbial populations in amesocosm system. A decline in chlorophyll a and increasedexcretion of 14C-labeled organic carbon were noted in thesame system (37), suggesting that growth-supporting hetero-trophic substrates were released by Cu2+-sensitive phyto-plankton. Similarly, increased heterotrophic activities of theaquatic communities could be due to stimulation ofheterotrophs nourished by growth substrates released fromHg2+-susceptible flora. This hypothesis is supported bydirect microscopic observations of cell lysis in water sam-ples exposed to Hg2+ (C. Liebert and T. Barkay, unpub-lished observation). The generality of this phenomenon isnot clear because evidence for inhibition of microbial pro-cesses by metals is also available (2, 9). It seems that theresponse of metal-stressed aquatic communities involves an

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equilibrium between several processes which are triggeredby the toxicants.An attempt was made in this study to select a set of

measurements which together will define the state of thecommunity under stress. [3H]TdR incorporation rate wasselected to measure heterotrophic activity because Jonas etal. (19) found it most sensitive to mercury toxicity andbecause it can be used to calculate in situ reproduction rates.However, the [3H]TdR incorporation rate was found todecline rapidly during 48 h of preincubation, often beyondthe level of detection, resulting in the loss of data forsalt-marsh and freshwater communities (Table 3). In addi-tion, a high variability between duplicate subsamples (largestandard deviations) rendered the differences betweenpreexposed and control communities insignificant. The datawere included in Table 3 as a record of the first use of thisapproach to study microbial adaptation. The utility of alter-native tests systems is currently under study.

Distribution of mer was determined by the colony hybrid-ization procedure in which colonies are grown on mediumdevoid of Hg2". Under these conditions, the limit of detec-tion was low (1 of 103 colonies). However, the absence ofmer+ colonies in preexposed salt-marsh and coastal marinecommunities and the low occurrence of mer in preexposedfreshwater and estuarine communities strongly suggest thatmer has no role in adaptation to Hg2+ in these communities.There are at least two possible hypotheses to account for thisobservation. (i) At least one additional Hg2+ resistancegenetic determinant exists in nature; resistant organismswhich do not hybridize with the characterized TnSOJ andTn2J are abundant (22; T. Barkay and G. Sayler, in 10thASTM Aquatic Toxicology Symposium, in press; N.Hamlett, personal communication). Thus, resistance or vol-atilization or both by the aquatic communities could havebeen mediated by an alternative determinant(s). If thishypothesis is true, mer would not be detected in Hg2+-resistant organisms isolated from the studied communitiesreported here.

(ii) In addition to Hg°, dimethylmercury is a highly volatilecompound (36). If aquatic communities biotransform Hg2+to dimethylmercury rather than reduce it, a different mech-anism (and thus gene system) mediates resistance and vola-tilization. Although aerobic Hg2+ methylation has beensuggested, little is known about its mechanism or the orga-nisms which carry it out (36). This hypothesis can be testedby identifying the chemical form of the volatilized mercurial.Experiments to test these two hypotheses are in progress.

ACKNOWLEDGMENTS

Gratitude is extended to Al Bourquin, Hap Pritchard, and PeterChapman for advice and support during this study, Jon Tuttle andBob Jonas for instructions in the [3H]TdR incorporation method asa toxicity measurement, Rick Cripe for help with statistical analy-ses, and Steve Foss for the figures.

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