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. 2001 Oct;67(10):4765–4772. doi: 10.1128/AEM.67.10.4765-4772.2001

Copper-Induced Inhibition of Growth of Desulfovibrio desulfuricans G20: Assessment of Its Toxicity and Correlation with Those of Zinc and Lead

Rajesh Kumar Sani 1, Brent M Peyton 1,*, Laura T Brown 1
PMCID: PMC93230  PMID: 11571183

Abstract

The toxicity of copper [Cu(II)] to sulfate-reducing bacteria (SRB) was studied by using Desulfovibrio desulfuricans G20 in a medium (MTM) developed specifically to test metal toxicity to SRB (R. K. Sani, G. Geesey, and B. M. Peyton, Adv. Environ. Res. 5:269–276, 2001). The effects of Cu(II) toxicity were observed in terms of inhibition in total cell protein, longer lag times, lower specific growth rates, and in some cases no measurable growth. At only 6 μM, Cu(II) reduced the maximum specific growth rate by 25% and the final cell protein concentration by 18% compared to the copper-free control. Inhibition by Cu(II) of cell yield and maximum specific growth rate increased with increasing concentrations. The Cu(II) concentration causing 50% inhibition in final cell protein was evaluated to be 16 μM. A Cu(II) concentration of 13.3 μM showed 50% inhibition in maximum specific growth rate. These results clearly show significant Cu(II) toxicity to SRB at concentrations that are 100 times lower than previously reported. No measurable growth was observed at 30 μM Cu(II) even after a prolonged incubation of 384 h. In contrast, Zn(II) and Pb(II), at 16 and 5 μM, increased lag times by 48 and 72 h, respectively, but yielded final cell protein concentrations equivalent to those of the zinc- and lead-free controls. Live/dead staining, based on membrane integrity, indicated that while Cu(II), Zn(II), and Pb(II) inhibited growth, these metals did not cause a loss of D. desulfuricans membrane integrity. The results show that D. desulfuricans in the presence of Cu(II) follows a growth pattern clearly different from the pattern followed in the presence of Zn(II) or Pb(II). It is therefore likely that Cu(II) toxicity proceeds by a mechanism different from that of Zn(II) or Pb(II) toxicity.

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

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.

After the addition of metal ions and cells, serum bottles were sampled for cell growth as total cell protein, number of cells with intact and damaged membranes, and aqueous concentrations of Cu(II), Pb(II), Zn(II), lactate, acetate, sulfate, and sulfide. During the incubation of D. desulfuricans in the presence or absence of metal, the redox potential (Eh) was monitored visually by resazurin. Resazurin is colorless at 0.5 mg/liter in a medium at pH 7 that has a value for Eh of ≤−100 mV (52). Resazurin was added to the medium prior to autoclaving. It was also shown in a previous study (43) that in MTM D. desulfuricans must first reduce the redox potential before growth can occur. Each experiment was carried out in duplicate and repeated for each set of conditions.

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.

The respective numbers of cells with intact and damaged membranes were measured by using a LIVE/DEAD Baclight bacterial viability kit (Molecular Probes, Eugene, Oreg.), a Petroff-Hausser counting chamber, and an epifluorescence microscope (Leica DMLB; Leica Microsystems Inc., Deerfield, Ill.) with an excitation filter of 480 nm and an emission filter of 520 nm. The LIVE/DEAD Baclight bacterial viability kit includes mixtures of the green fluorescent nucleic acid stain SYTO 9 and the red fluorescent nucleic acid stain propidium iodide. The SYTO 9 stain generally labels all bacteria in any population, both those with intact membranes and those with damaged membranes, i.e., both live and dead bacteria (9, 15). In contrast, propidium iodide penetrates only bacteria with damaged membranes, causing a reduction in the SYTO 9 stain fluorescence when both dyes are present. Damaged cells were counted by their red color and were scored as having damaged membranes (dead).

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.

Samples for aqueous Cu(II) concentration were prepared by filtering through a 0.2-μm-pore-size membrane filter, and concentrations of Cu(II) (0 to 3 μM) were determined using a quantitative colorimetric porphyrin method (Hach Company, Loveland, Colo.). Sample size and analytical reagents were reduced proportionately to allow use of a smaller sample volume. The absorbance was measured at 425 nm and compared to a standard curve generated for known concentrations of cupric chloride. The detection limit for Cu(II) was estimated to be 0.2 μM. Samples for aqueous zinc concentration were prepared by filtering through a 0.2-μm-pore-size membrane filter, and concentrations of Zn(II) (0 to 30 μM) were determined using a quantitative colorimetric Zincon method (Hach Company) modified to reduce the required sample volume. The absorbance was measured at 620 nm and compared to a standard curve generated for known concentrations of ZnCl2. This method gave a detection limit of 0.6 μM. Samples for aqueous Pb(II) concentration were filtered through a 0.2-μm-pore-size membrane filter, diluted with 3% HNO3 prepared with nanopure water, and measured on an Agilent 4500 inductively coupled plasma mass spectrometer (ICP-MS). Calibration of this spectrometer was done using standard Pb(II) solutions of 0, 0.25, 0.5, and 1 μM, to give a Pb(II) detection limit of 0.05 μM.

RESULTS

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 t1/2 (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 t1/2 was approximately the same (21 h) as that of the copper-free control. However, at 12 and 18 μM Cu(II), the values of t1/2 increased to 28 and 46 h, respectively.

FIG. 1.

FIG. 1

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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.

To quantify Cu(II) toxicity in MTM, the percent inhibition in maximum cell protein at different concentrations of Cu(II) was calculated and plotted as shown in Fig. 2A. A least-squares line was obtained (R2 = 0.997) and was used to calculate the Cu(II) concentration estimated to cause a 50% inhibition in total cell protein (IC50) of D. desulfuricans. An IC50 of 16 μM Cu(II) was determined, which is significantly lower than that reported in the literature (46). In addition to the IC50, the percent reduction in maximum specific growth rates was calculated and plotted versus Cu(II) concentrations (Fig. 2B). It can be seen from Fig. 2B that maximum specific growth rates decreased with increasing Cu(II) concentration. As in Fig. 2A, a least-squares line (R2 = 0.972) was obtained relating percent inhibition in maximum specific growth rate to Cu(II) concentrations. It can be seen that 13.3 μM Cu(II) caused 50% inhibition in specific growth rates, which was similar to the IC50 for Cu(II) obtained for the data in Fig. 2A.

FIG. 2.

FIG. 2

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(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.

FIG. 3.

FIG. 3

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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.

Lag times also correlated with observations on the change in color of resazurin. The metal-free serum bottle contents went from blue to pink to colorless within 24 h, indicating that Eh was ≤−100 mV, while with Zn(II) and Pb(II), the medium became colorless after 36 h. However, with Cu(II) the serum bottle contents required 72 h to become colorless. Thus, under the same conditions, with Cu(II) the bacteria required more time to reduce the redox potential than with Zn(II) or Pb(II). In the presence of 47 μM Cu(II), D. desulfuricans was not able to lower the Eh to ≤−100 mV and no growth was observed (data not shown). It is interesting that with Zn(II) and Pb(II), growth started after a longer lag time than the metal-free control yet ultimately attained the same total cell protein as the metal-free control. However, with Cu(II), the growth rates decreased drastically and the final cell protein concentration was reduced by 68% compared to the copper-free control.

The values of t1/2 were also calculated from Fig. 3 and are given in Table 1. It has been observed that the values of t1/2 increased with the addition of heavy metals. The values of t1/2 with Zn(II), Pb(II), and Cu(II) were found to be 93, 116, and 163 h, respectively, compared to 85 h for the metal-free control. In addition to the values of t1/2, the maximum specific growth rates in the presence or absence of heavy metals were also calculated and listed in Table 1. The maximum specific growth rates with Zn(II), Pb(II), and Cu(II) were found to be 0.27, 0.26, and 0.06 h−1, respectively, compared to 0.24 h−1 for metal-free control. It can be seen from Table 1 that for Zn(II) and metal-free control, the values of t1/2 and the maximum specific growth rate are not statistically significantly different, indicating that Zn(II) at 16 μM showed no significant toxicity.

TABLE 1.

Influences of different metal ions on maximum cell protein, t1/2, maximum specific growth rate, final sulfide, and pH of D. desulfuricansa

Metal ion and concn (μM) Maximum cell protein concn (mg/liter) t1/2 (h)b Final aqueous sulfide concn (mM) Final pH Maximum specific growth rate, μc (h−1)
None (uninoculated control) NAd NA 0 7.21 ± 0.01 NA
None (metal-free control) 37.8 ± 0.5 85 ± 6.2 11.8 ± 1.3 7.46 ± 0.01 0.24 ± 0.02
Cu, 16 11.9 ± 0.5 163 ± 10.1 4.0 ± 1.0 7.36 ± 0.05 0.06 ± 0.01
Zn, 16 40.0 ± 0.9 93 ± 6.1 13.3 ± 1.1 7.46 ± 0.15 0.27 ± 0.02
Pb, 5 39.3 ± 1.2 116 ± 8.1 13.03 ± 1.7 7.42 ± 0.11 0.26 ± 0.02
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a

Values are reported as averages ± standard deviations. 

b

t1/2, time at which the total cell protein is half of the maximum total cell protein. 

c

Specific growth rate (μ) = dx/(xdt), as described in Materials and Methods. 

d

NA, not applicable. 

It was hypothesized that exposure of D. desulfuricans to Cu(II), Zn(II), and Pb(II) could lead to a decline in membrane integrity. This has been observed for S. cerevisiae, where in the presence of Cu(II), disruption of plasma membrane integrity was measured by release of K+, amino acids, and nucleic acids (3, 37). Extensive metal-induced disruption of membrane integrity might be responsible for the loss of cell viability, longer lag times, and/or reduced growth. To check this hypothesis, the numbers of live cells (with intact membranes) were determined using epifluorescence microscopy and the LIVE/DEAD Baclight bacterial viability kit. It can be seen from Fig. 4 that the profiles of cell numbers were similar to the profiles of cell protein. During the growth of D. desulfuricans, in all experiments with and without heavy metals, the numbers of cells with damaged membranes accounted for less than 1% of the total cell count (data not shown). It was observed, however, that in the presence of Cu(II), cells had much less fluorescence than did those in the presence of Pb(II) or Zn(II) or the metal-free control.

FIG. 4.

FIG. 4

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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.

In addition to cell protein and cell number, the aqueous concentrations of lactate, sulfate, and acetate in the serum bottles were measured over time; results are shown in Fig. 5A, B, and C, respectively. It can be seen from Fig. 5A that in the presence of Zn(II) or Pb(II) (after a short lag phase) and in the metal-free control, the rates of lactate biotransformation were approximately the same. However, in the case of Cu(II), lactate biotransformation rates were appreciably reduced. With Zn(II), Pb(II), and the metal-free control, D. desulfuricans utilized 90% of the available lactate as an electron donor compared to only 38% in the presence of Cu(II) (Fig. 5A). Similar results were observed in the utilization of sulfate as an electron acceptor and formation of acetate (Fig. 5B and C). Dissimilatory bacterial sulfate reduction, using lactate as an electron donor, is described in equation 1 (51):

graphic file with name M1.gif 1

It can be seen from Fig. 5A, B, and C that the consumption in lactate and sulfate was significantly slower in the presence of Cu(II) and that the cultures containing Pb(II) and Zn(II) behaved in a manner similar to that of the control. The lactate-to-sulfate utilization ratios were 1.95, 2.28, 2.03, and 1.95 for the metal-free control, Cu(II), Zn(II), and Pb(II), respectively, which correspond well to the theoretical value of 2 shown in equation 1. These data indicate that although significant inhibition by Cu(II) was observed, only a slight difference in the expected electron donor and acceptor utilization ratio was observed for the Cu(II)-containing cultures. Correspondingly, the ratios of lactate consumption to acetate production were 0.97, 1.04, 0.95, and 0.90 for the metal-free control, Cu(II), Zn(II), and Pb(II), respectively, which also matched well with a theoretical value of 1 (equation 1). Final aqueous phase concentrations of sulfide with metal-free control, Cu(II), Zn(II), and Pb(II) were found to be 11.81, 4.03, 13.34, and 13.03 mM, respectively (Table 1). It can also be seen from Table 1 that the pH remained essentially constant throughout the experiments, which is important since metal ion species often changes with pH.

FIG. 5.

FIG. 5

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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.

Aqueous concentrations of Cu(II), Zn(II), and Pb(II) for the cultures are indicated in Fig. 6, which shows that for up to 24 h, there was no significant reduction in aqueous-phase Cu(II), Pb(II), or Zn(II) concentrations. The aqueous Pb(II) and Zn(II) concentrations decreased rapidly after 24 and 48 h, respectively, and were both below the detection limit after 90 h. However, in the case of Cu(II), the reduction was slower than Zn(II) and Pb(II) and the Cu(II) concentration did not reach the detection limit until 120 h. No precipitation or removal of Cu(II), Zn(II), or Pb(II) was observed in abiotic controls (Fig. 6). Comparison of Fig. 3, 4, and 5 with Fig. 6 indicates that very little microbial activity was observed until concentrations of Cu(II), Zn(II), and Pb(II) were less than 0.2, 3.4, and 0.05 μM, respectively.

FIG. 6.

FIG. 6

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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

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 (Eh), based on visual observation of the resazurin indicator. At higher Cu(II) concentrations (47 μM), D. desulfuricans was unable to reduce the Eh 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 Eh 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 Eh.

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 t1/2 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.

ACKNOWLEDGMENTS

We gratefully acknowledge the financial support provided by the Natural and Accelerated Bioremediation Research program (NABIR), Office of Biological and Environmental Research, U.S. Department of Energy (grant DE-FG03-98ER62630/A001). The support of the Center for Multiphase Environmental Research and the Department of Chemical Engineering also contributed significantly to this research.

We thank Gill Geesey for input.

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