INTRODUCTION

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Heavy metals are an important class of pollutants and derive from both point sources (e.g., sludge dumping, industrial effluents, mine tailings) and diffuse sources (e.g., highway runoff [10]). Metals such as cadmium, chromium, copper, lead, mercury, uranium, and zinc have been shown to exist at significantly elevated levels in ground waters and in soil and sediments at U.S. Department of Energy facilities and in other sediments (23, 40, 55). Great interest in metal-microbe interactions has arisen in recent years as scientists and engineers try to remove, recover, or stabilize heavy metals in soils. Metal toxicity towards microorganisms is of environmental concern because of possible inhibition of essential microbe-assisted processes, such as the degradation of organic matter (3, 28, 41) and the transfer of accumulated metals to higher organisms in the food chain (21, 26). Biological treatment of toxic heavy metals, efficient management of bacterial processes in ex situ-engineered treatments, and effective manipulation of indigenous bacterial communities to stimulate in situ activity all require knowledge of the toxic effects of various heavy metals on bacterial populations.

Both in laboratory studies and in field studies, it has been shown that the toxicity of a given metal depends on species and chemical properties as well as environmental factors (e.g., adsorption to solid surfaces, complexation, or precipitation) (22, 26, 34, 43, 57). Heavy metal toxicity is also known to interfere with important microbial processes including aerobic and anaerobic degradation of organic matter (3, 28, 41). Toxic effects include ion displacement and/or substitution of essential ions from cellular sites and blocking of functional groups of important molecules, e.g., enzymes, polynucleotides, and essential nutrient transport systems (35). This results in denaturation and inactivation of enzymes and disruption of cell organelle membrane integrity (21, 36). Microorganisms require some metals like Cu(II), Zn(II), Co(II), and Ni(II) at low concentrations as essential micronutrients for vital cofactors for metalloproteins and certain enzymes (21, 35). However, at higher concentrations, it has been reported that these metals interact with nucleic acids and enzyme active sites and in Saccharomyces cerevisiae can also lead to a rapid decline in membrane integrity, which is generally manifested as leakage of mobile cellular solutes (e.g., K⁺) and cell death (11, 25, 37, 49).

Sulfate reduction may be the predominating pathway of terminal carbon oxidation in anoxic sediments since sulfate-reducing bacteria (SRB) often effectively outcompete methanogens for common substrates (1, 31, 56) and have higher theoretical growth yields (51). SRB are well known to utilize various electron donors (organic acids such as lactate, acetate, pyruvate, and formate) and hydrogen, electron acceptors including sulfate, sulfite, thiosulfate, elemental sulfur, fumarate, nitrate, nitrite, and heavy metals like Fe(III), Cr(VI), U(VI), Mn(IV), and Tc(VII) (14). In addition, SRB are present in many contaminated subsurface sites (5). SRB carry out dissimilatory reduction of sulfate to sulfide. The resulting HS is very reactive and forms insoluble precipitates with heavy metals like Pb(II), Cd(II), Cu(II), Zn(II), Ni(II), and Hg(II) (7, 39, 54). Further, the formation of metal sulfides in situ can help maintain a low redox potential barrier that hinders the reoxidation of heavy metal precipitates. Finally, SRB are also known to enzymatically reduce Cr(VI) and U(VI) to form insoluble mineral phases (32, 33) and have been shown to reduce soluble Pd(II) to zerovalent Pd (29).

While these organisms can catalyze a variety of heavy metal transformations, it has been demonstrated that heavy metals at toxic levels may inhibit or prevent SRB growth, despite their release of sulfide (39, 43). Reports regarding the toxicity of heavy metals to SRB (8, 10, 24, 27, 30, 39, 42, 46, 50) have generally been qualitative in nature and have used microbial media designed to optimize growth rather than to examine metal toxicity. Often in these studies the authors reported abiotic formation of metal precipitates and/or significant metal complexation that prevented meaningful quantitative assessment of metal toxicity. Of particular note, however, Poulson et al. (39) quantified the toxicity of nickel and zinc to Desulfovibrio desulfuricans in a chemically defined medium and calculated metal activities using the geochemical species identification program MINTEQA2 (2).

In a previous report (43), Pb(II) toxicity to D. desulfuricans G20 was shown to be strongly influenced by the physicochemical properties of the SRB ambient environment (e.g., presence of chelators, buffers, and reductants). These observations led to the development of an SRB metal toxicity medium (MTM), in which no abiotic precipitation of Cu(II), Pb(II), or Zn(II) was observed. MTM was specifically designed to determine the effects of metal toxicity on SRB and has been used to generate a baseline for the studies presented here. The present investigation examines the effect of copper (a well-known toxic metal) on the growth of the sulfate-reducing bacterium D. desulfuricans G20 in MTM. In this study, the degree of toxicity was quantified in terms of (i) inhibition of total cell protein, (ii) inhibition of number of cells with intact membranes, (iii) reduction in maximum specific growth rates, (iv) longer lag times and in some cases no measurable growth of D. desulfuricans. The effects of Pb(II) and Zn(II) on the growth of D. desulfuricans were also investigated and compared to that of Cu(II).

	MATERIALS AND METHODS

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Microorganism, medium, and cultivation conditions. The D. desulfuricans strain used in the study was G20, which was a gift of J. Wall, University of Missouri. D. desulfuricans G20 was maintained in MTM (43), which contained (in grams per liter) sodium lactate (4.6), sodium sulfate (2.23), calcium chloride dihydrated (0.06), ammonium chloride (1.0), magnesium sulfate (1.0), yeast extract (0.05), tryptone (0.5), and PIPES [piperazine-N,N'-bis(2-ethanesulfonic acid)] disodium salt monohydrate (10.93). The pH was adjusted to 7.2 with 1 N HCl. The medium components and heavy metal standard solutions were of analytic grade and were purchased from Fisher Scientific (Pittsburgh, Pa.) with the exception of the following: yeast extract and tryptone were obtained from Difco Chemical Company, and PIPES buffer, resazurin, and sodium sulfate were obtained from Aldrich Chemical Company. The preparation of serum bottles and cultivating conditions were the same as described previously (43). In brief, the serum bottles containing media were autoclaved and were put immediately in an anaerobic chamber under vacuum (10 in. Hg) to remove headspace oxygen. The serum bottles were sealed with butyl rubber septa, capped, and crimped with an aluminum seal and pressurized with ultrapure nitrogen at 10 lb/in² above atmospheric pressure. D. desulfuricans G20 contained the green fluorescent protein (GFP) reporter gene construct that was unused in the present study, but to maintain the GFP plasmid, 20 µg of chloramphenicol per ml was added to the medium prior to inoculation.

Preparation of cells for inoculation and experimentation. Hydrogen sulfide initially present in an active inoculum was removed by flushing with ultrapure nitrogen for 1 h. The cells were centrifuged in the presence of ultrapure nitrogen at 10,000 × g for 10 min. The supernatant was discarded, and the cell pellets were suspended in 0.89% NaCl. This process was repeated twice, and washed cells were used as inoculum. Prior to inoculation of the serum bottles, the inoculum was examined under epifluorescence microscope with an excitation filter of 480 nm and an emission filter of 520 nm. The cells were found to be motile and fluorescent. The apparent absence of elongated cells indicated that anaerobic conditions were maintained during growth since it has been reported that Desulfovibrio strains develop typically elongated cells when growing in the presence of oxygen (45). Stock solutions of CuCl₂, ZnCl₂, and PbCl₂ were added to the serum bottles to give the desired metal concentrations. To examine the inhibitory effects of Cu(II) on D. desulfuricans, a 16 µM concentration of Cu(II) was selected by preliminary screening tests, since at this concentration, the inhibition in total cell protein and maximum specific growth rate was approximately 50%. For comparison, 16 µM concentrations of Zn(II) and Pb(II) were also used. However, at 16 µM Pb(II), no growth of D. desulfuricans was observed even after prolonged incubation of 3 weeks (unpublished data), such that for Pb(II), 5 µM was used for comparison under otherwise identical conditions. Pb(II) concentrations of 10 µM were examined, but the lag time at this concentration was >200 h, such that it did not compare well with the Zn(II) data.

Determination of total cell protein, maximum specific growth rate, and cell numbers. Total cell protein was determined using a quantitative colorimetric Coomassie assay method (Pierce, Rockford, Ill.) described previously (43). For determination of maximum specific growth rates, growth curve data were evaluated by polynomial curve fitting in Microsoft Excel. The polynomial was used to calculate the derivative, dx/dt, which was then used to calculate the specific growth rates using the formula µ = (1/x) (dx/dt) (44). Fit curves had an R² of >0.95 in all cases and did not differ by more than 10% from any experimental data point.

Determination of organic and inorganic anions and metal ions. Samples for lactate, sulfate, and acetate were filtered (Gelman Acrodisc; pore diameter, 0.2 µm), and concentrations were determined using a Dionex Ion Chromatograph (DX-500 equipped with conductivity detector-20) with an IonPac AS11-HC4-mm column. Elution was carried out using a sodium hydroxide gradient (1 to 100 mM). Aqueous sulfide concentration was determined spectrophotometrically using the methylene blue method applied to liquid samples that were added to 0.5 ml of a 10% (wt/vol in water) zinc acetate solution (19, 20). Sulfide in the gaseous compartment was not measured.

	RESULTS

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Effect of copper on the growth of D. desulfuricans G20. In this study, assessment of metal toxicity was quantified as the inhibition of D. desulfuricans G20 growth in MTM based on total cell protein, the number of cells with intact membranes as indicated by live/dead staining, and the reduction in maximum specific growth rates. Growth profiles of D. desulfuricans in MTM at Cu(II) concentrations of 0 to 30 µM are shown in Fig. 1. It can be seen from Fig. 1 that the toxicity of Cu(II) to D. desulfuricans was dependent on Cu(II) concentration and that Cu(II) caused inhibition in the final cell protein yield, longer lag times, and lower growth rates or no measurable growth. As is shown in Fig. 1, at 6 and 12 µM Cu(II), cell protein concentration increased with time, and lag times were approximately the same as that of the copper-free control. However, growth rates in the presence of even 6 µM Cu(II) were lower than in the copper-free control. Also in the presence of Cu(II), the final cell protein concentration was lower than that of the copper-free control by 18 and 35% for 6 and 12 µM Cu(II), respectively. At 18 µM Cu(II), the growth rate decreased further; moreover, lag time increased to 24 h, and the final cell protein concentration was reduced by 59% to that of the copper-free control. It can be seen from Fig. 1 that under the same conditions, Cu(II) completely inhibited growth at 30 µM, since no measurable growth was observed for up to 384 h. From the data presented in Fig. 1, the values of t_1/2 (the time at which the total cell protein is half of the maximum total cell protein) were also evaluated. At 6 µM Cu(II), the value of t_1/2 was approximately the same (21 h) as that of the copper-free control. However, at 12 and 18 µM Cu(II), the values of t_1/2 increased to 28 and 46 h, respectively.

View larger version (25K): [in this window] [in a new window]   FIG. 1.   Effect of Cu(II) on the growth of D. desulfuricans G20 as measured by total cell protein. The points are the averages of duplicates, and error bars indicate ± standard deviations of the means (n = 2). Error bars smaller than the symbols are not shown.

View larger version (11K): [in this window] [in a new window]   FIG. 2.   (A) Inhibition of maximum cell protein of D. desulfuricans G20 as a function of Cu(II) concentration. (B) Inhibition of maximum specific growth rate of D. desulfuricans G20 as a function of Cu(II) concentration.

Effects of copper, zinc, and lead on the growth of D. desulfuricans G20. To examine the inhibitory effects of Cu(II) on D. desulfuricans, a 16 µM concentration of Cu(II) was selected by preliminary screening tests, since at this concentration, the inhibition in total cell protein and maximum specific growth rate was approximately 50%. For comparison, 16 µM concentrations of Zn(II) and Pb(II) were also used. However, at 16 µM Pb(II), no growth of D. desulfuricans was observed even after prolonged incubation of 3 weeks (unpublished data), such that for Pb(II), a 5 µM concentration was used for comparison under otherwise identical conditions. The effects of Cu(II), Zn(II), and Pb(II) on the growth of D. desulfuricans are shown in Fig. 3. All metals were tested individually. It can be seen from Fig. 3 that growth rates with Pb(II) and Zn(II) were similar to that of the metal-free control. However, with Cu(II), growth rates were reduced significantly. Lag times were also increased by the addition of heavy metals. The lag times with Zn(II), Pb(II), and Cu(II) were observed to be 48, 72, and 120 h, respectively, compared to 24 h for the metal-free control.

View larger version (22K): [in this window] [in a new window]   FIG. 3.   The effects of Cu(II), Zn(II), and Pb(II) on the growth of D. desulfuricans G20 as measured by total cell protein. The points are the averages of duplicates, and error bars indicate ± standard deviations of the means (n = 2). Error bars smaller than the symbols are not shown.

1

1

View larger version (19K): [in this window] [in a new window]   FIG. 4.   Direct count of live cells during the growth of D. desulfuricans G20 in the presence of Cu(II), Zn(II), and Pb(II). The points are the averages of duplicates, and error bars indicate ± standard deviations of the means (n = 2). Error bars smaller than the symbols are not shown.

(1)

View larger version (25K): [in this window] [in a new window]   FIG. 5.   Aqueous lactate (A), sulfate (B), and acetate (C) concentrations during the growth of D. desulfuricans G20 in the presence of Cu(II), Zn(II), and Pb(II). The points are the averages of duplicates, and error bars indicate ± standard deviations of the means (n = 2). Error bars smaller than the symbols are not shown.

View larger version (24K): [in this window] [in a new window]   FIG. 6.   Aqueous metal ion concentrations during the growth of D. desulfuricans G20 in the presence of Pb(II), Zn(II), and Cu(II). The points are the averages of duplicates and error bars indicate ± standard deviations of the means (n = 2). Error bars smaller than the symbols are not shown.

	DISCUSSION

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In this study, D. desulfuricans was found to be very susceptible to Cu(II) at very low concentration (6 µM) in MTM. In the presence of even very low Cu(II) concentrations, the culture required more time than in the copper-free control to lower the redox potential (E_h), based on visual observation of the resazurin indicator. At higher Cu(II) concentrations (47 µM), D. desulfuricans was unable to reduce the E_h even after a prolonged incubation of four months. At the same time, however, live/dead staining indicated that most of the cells had intact membranes (i.e., were alive), indicating that cells were inhibited by Cu(II) but had not lysed. It has been reported that Cu(II) can act as an inhibitor of periplasmic hydrogenase (17, 18, 29). It has also been reported that in D. desulfuricans and D. vulgaris, the periplasmic proteins, which contain hydrogenase and cytochrome c₃, were found to catalyze oxygen reduction with high rates (13, 14). In our results, the presence of heavy metals significantly inhibited the rate at which D. desulfuricans lowered the E_h of the medium; thus, lag times were increased compared to metal-free control. Thus, while they do not constitute a conclusive proof, our results support previous work that suggests that Cu(II) likely inhibits periplasmic hydrogenase, which would in turn impair the organism's ability to scavenge trace oxygen, thus lengthening the time required to lower the medium E_h.

From our data, the IC₅₀ of Cu(II) on D. desulfuricans was calculated to be 16 µM. This value is significantly lower than those previously reported in the literature. Song et al. (46) reported a sulfate removal IC₅₀ for Cu(II) (concentration causing 50% inhibition of SRB sulfate removal efficiency) of 1.57 mM Cu(II). Their value is 100 times higher than that obtained in this study. They observed no inhibition in sulfate reduction at concentrations up to 79 µM Cu(II). It should be noted that they used a mixed SRB culture and an anaerobic growth medium containing constituents that have been shown to be responsible for significant metal complexation and/or precipitation (43). Many previous studies have used growth media that resulted in significant abiotic precipitation and/or complexation of toxic metal ions. For example, Jalali and Baldwin (27) observed no significant inhibition in sulfate reduction at initial concentrations of up to 787 µM Cu(II); however, they also observed up to 78% abiotic copper removal by precipitation, indicating that only 173 µM Cu(II) could have remained in solution. In addition, they used Postgate's medium C (38), which contains metal complexants, reductants, and chelators that can significantly reduce metal bioavailability and thus significantly alter observed metal toxicity (43).

Staining of D. desulfuricans using the LIVE/DEAD Baclight bacterial viability kit indicated that in the presence of Cu(II) most cells showed green fluorescence, indicating that they had intact membranes. It was also observed, however, that after exposure to Cu(II), cells had much less fluorescence than did those in the presence of Pb(II) or Zn(II) or the metal-free control. It may be possible that exposure of D. desulfuricans to Cu(II) reduced the efficiency of uptake of SYTO 9 green fluorescent nucleic acid stain. Ohsumi et al. (37) observed that under the copper-stressed conditions, S. cerevisiae releases nucleic acids through the membrane. In our results, it is possible that a similar effect may have occurred in D. desulfuricans under Cu(II) stress, and since SYTO 9 dye stains the nucleic DNA, this may result in the low fluorescence observed with Cu(II) compared to those of Pb(II), Zn(II), and the metal-free control.

The results presented here show that Cu(II) toxicity to D. desulfuricans can be quantified in terms of inhibition in total cell protein and cell number, longer lag times, and t_1/2 values, lower maximum specific growth rates, and at concentrations of 30 µM or higher no measurable growth. In the presence of Pb(II) and Zn(II) at 5 and 16 µM, respectively, once growth had started, even after long lag times, for D. desulfuricans in MTM, we observed no inhibition in the final cell protein. Interestingly, this was not the case for Cu(II), where final cell protein concentration was linearly reduced with increasing Cu(II) concentrations up to 30 µM. In contrast, Zn(II) and Pb(II) exerted no permanent effect on cell growth, which may indicate that no lasting structural damage occurred in the cells of D. desulfuricans organisms. As soon as D. desulfuricans detoxified and removed Zn(II) or Pb(II) from the aqueous phase, it grew in a manner similar to that of the metal-free control. In contrast to Pb(II) and Zn(II), Cu(II) appeared to effect some permanent structural alteration in D. desulfuricans cells. Even after Cu(II) concentrations were below the detection limit, growth did not resume to match the metal-free control, such that exposure to Cu(II) resulted in continuing low rates of growth and low final cell protein concentrations. These data indicate that there is a significant residual effect of Cu(II) on the exposed organisms and that perhaps Cu(II) exerts a toxicity mechanism different from those of Pb(II) and Zn(II). Similar effects have been reported where Cu(II) altered the physical properties of cell membranes and/or inhibited critical functional enzymes of algae (Nitzschia closterium and Chlorella pyrenoidosa) and yeast (S. cerevisiae) and was responsible for reduced growth (4, 47). Our results suggest that the mechanism of metal toxicity to SRB may not be a fortuitous feature of sulfide production but is apparently more complex and differs with different metals.

In this study, we observed decreases in aqueous metal concentrations, which may have resulted from the following processes: (i) biosorption to cell surfaces (12), (ii) release of extracellular polymeric substances that can complex and detoxify Cu(II) (6), (iii) complexation and precipitation of Cu(II) as CuS (7, 39), and (iv) intracellular penetration and accumulation (16). In our system, all four mechanisms may have been responsible for the measured decrease in aqueous metal concentrations. However, for the residual inhibition of growth observed in the case of Cu(II), it appears that processes i through iii are not likely to be responsible and that residual inhibition is probably the result of Cu(II) penetrating into the periplasm or cytoplasm to react with intracellular components.

Mechanisms of metal toxicity and inhibition in microbiological systems, especially with SRB, are not well understood. To have a physiological or toxic effect, most heavy metal cations have to enter the cell (35). Cu(II) at toxic concentrations is known to bind to free thiols (e.g., glutathione) and other functional groups (e.g., -SH) of enzymes and may also replace metals that are constituents and the active centers of enzymes, cofactors, or other biomolecules. This results in denaturation and inactivation of enzymes and disruption of cell organelle membrane integrity and cell division (21, 36, 48, 53). Further research is under way to better understand how Cu(II) inhibits the growth of D. desulfuricans and what factors are responsible for reduction in aqueous phase Cu(II) concentrations.

The results of this study clearly show that heavy metal toxicity to the sulfate-reducing bacterium D. desulfuricans G20 was demonstrated by inhibition in total cell protein, longer lag times, lower maximum specific growth rates, and in some cases no measurable growth. When the toxicities of Pb(II) and Zn(II) were studied, however, no inhibition in the final cell protein concentration was observed. It can be concluded that in the absence of strong chelators and reductants, Cu(II) concentrations of 6 µM are toxic to D. desulfuricans at pH 7.2 and at 30 µM no growth will occur. Under the same conditions, at 47 µM Cu(II), D. desulfuricans could not even lower the redox potential. Live/dead staining showed that Cu(II), Pb(II), and Zn(II) at the concentrations tested did not kill the cells of D. desulfuricans even when, in some cases, growth was utterly inhibited. It was observed that even after a long lag phase [more than 96 h with 16 µM Cu(II)], more than 99% of the cells gave indications that cell membranes were intact. Comparison of our metal toxicity results with literature values indicates that in some conditions heavy metals may be much more toxic than previously thought, since earlier studies used media that contained significant amounts of metal complexing and/or precipitating agents (e.g., Postgate's C medium).

For SRB, our measured IC₅₀ for Cu(II) of 16 µM is approximately 100 times smaller than the previously reported IC₅₀ of 1.57 mM (46). This indicates that MTM may be a good medium for measuring metal toxicity to SRB and could provide a baseline for comparison of natural and industrial waters. In addition, we envision that in future studies MTM could be supplemented with specific complexing agents to help better understand the interactive effects of aqueous system components and toxic heavy metals on SRB. The results indicate that D. desulfuricans clearly showed different growth patterns and significantly reduced maximum specific growth rate, indicating that Cu(II) toxicity may have proceeded by a mechanism different from that used by Pb(II) or Zn(II) toxicity. While the use of MTM and a pure culture of D. desulfuricans may overestimate metal toxicity in the natural environment (e.g., where chemical complexants and other microorganisms are present), the results presented here have fundamental relevance to SRB found in natural systems that contain heavy metals and also to efforts to use SRB to remediate heavy metal contamination.