Reductive Dechlorination of cis-1,2-Dichloroethene and Vinyl Chloride

cis-Dichloroethene (DCE) and vinyl chloride (VC) often accumulate in contaminated aquifers in which tetrachloroethene (PCE) or trichloroethene (TCE) u...
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Environ. Sci. Technol. 2001, 35, 516-521

Reductive Dechlorination of cis-1,2-Dichloroethene and Vinyl Chloride by “Dehalococcoides ethenogenes” XAVIER MAYMO Ä -GATELL,‡ IVONNE NIJENHUIS, AND STEPHEN H. ZINDER* Department of Microbiology, Wing Hall, Cornell University, Ithaca, New York 14853

cis-Dichloroethene (DCE) and vinyl chloride (VC) often accumulate in contaminated aquifers in which tetrachloroethene (PCE) or trichloroethene (TCE) undergo reductive dechlorination. “Dehalococcoides ethenogenes” strain 195 is the first isolate capable of dechlorinating chloroethenes past cis-DCE. Strain 195 could utilize commercially synthesized cis-DCE as an electron acceptor, but doses greater than 0.2 mmol/L were inhibitory, especially to PCE utilization. To test whether the cis-DCE itself was toxic, or whether the toxicity was due to impurities in the commercial preparation (97% nominal purity), we produced cis-DCE biologically from PCE using a Desulfitobacterium sp. culture. The biogenic cis-DCE was readily utilized at high concentrations by strain 195 indicating that cis-DCE was not intrinsically inhibitory. Analysis of the commercially synthesized cis-DCE by GC/mass spectrometry indicated the presence of approximately 0.4% mol/mol chloroform. Chloroform was found to be inhibitory to chloroethene utilization by strain 195 and at least partially accounts for the inhibitory activity of the synthetic cis-DCE. VC, a human carcinogen that accumulates to a large extent in cultures of strain 195, was not utilized as a growth substrate, and cultures inoculated into medium with VC required a growth substrate, such as PCE, for substantial VC dechlorination. However, high concentrations of PCE or TCE inhibited VC dechlorination. Use of a hexadecane phase to keep the aqueous PCE concentration low in cultures allowed simultaneous utilization of PCE and VC. At contaminated sites in which “D. ethenogenes” or similar organisms are present, biogenic cis-DCE should be readily dechlorinated, chloroform as a co-contaminant may be inhibitory, and concentrations of PCE and TCE, except perhaps those near the source zone, should allow substantial VC dechlorination.

Introduction The solvents tetrachloroethene (PCE) and trichloroethene (TCE) are major groundwater pollutants (1). At anaerobic sites, these solvents can undergo reductive dechlorination to less-chlorinated ethenes. At some sites reductive dechlorination proceeds completely to ethene, while at others * Corresponding author phone: (607)255-2415; fax: (607)255-3904; e-mail: [email protected]. ‡ Present address: McKinsey & Company, Barcelona, Spain. 516

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dechlorination is incomplete (2). Large accumulations of cisdichloroethene (DCE) are often found at contaminated sites undergoing reductive dechlorination, and at some sites there is little or no dechlorination past cis-DCE. The reasons for cis-DCE accumulation are partially understood. It appears that organisms that can dechlorinate PCE to cis-DCE are diverse and cosmopolitan. Several different organisms have been isolated from diverse sites that reductively dechlorinate PCE and TCE no further than cis-DCE (3-9). Organisms that dechlorinate past cis-DCE are not always present at contaminated sites and seem to be more nutritionally fastidious and slower growing than the above organisms (10-14). Presently, the only isolated organism known to reductively dechlorinate chloroethenes further than DCE is provisionally named “Dehalococcoides ethenogenes”. Growth of this organism on cis-DCE was considerably slower than with the other electron acceptors (PCE, TCE, 1,1-DCE, and 1,2-dichloroethane (DCA)). In this communication, we examine more extensively the utilization of this environmentally significant intermediate and demonstrate that slower growth on cis-DCE is due to contaminants in the commercial preparation. When dechlorination proceeds past cis-DCE at a site, accumulations of vinyl chloride (VC) are also undesirable since VC is considered a human carcinogen (15). VC can be utilized aerobically as a growth substrate by various microorganisms (16-18), and evidence for its oxidation under anaerobic conditions in the presence of Fe3+ (19) or humic acids (20) has been obtained. VC can be reduced to ethene and ethane under anaerobic conditions by a variety of mixed cultures (14, 21-25) and in microcosms and field sites (10, 26, 27). When chloroethenes are utilized by strain 195 or cultures containing strain 195, VC conversion to ethene (ETH) was rate limiting and showed first-order kinetics (13, 28). Moreover, its reductive dechlorination did not support growth of strain 195 (29) so that VC utilization can be considered cometabolic. In this publication, we describe the utilization of these two important substrates by “D. ethenogenes” strain 195 in greater detail. We demonstrate that difficulties utilizing cisDCE by strain 195 were due to inhibitory impurities in the commercially synthesized product, one of which is chloroform. We also demonstrate that VC utilization by strain 195 requires another chloroethene to serve as an electron acceptor for growth; however, high concentrations of these chloroethenes also inhibit VC utilization during the period of their utilization.

Materials and Methods Chemicals and Analyses of Chloroethenes. Chloroethenes and other chemicals were purchased as described previously (24). The commercially produced cis-DCE, with a nominal purity of 97%, was purchased from Aldrich Chemical Co., Milwaukee, WI. We were unable to find cis-DCE from another supplier. For quantitative analysis of chloroethenes and ETH, headspace gas samples (100 µl) were analyzed using a temperature programmed Perkin-Elmer 8500 gas chromatograph (GC) equipped with a flame ionization detector (FID). The GC contained a 60m × 0.53 mm RTX-502.2 capillary column operated in splitless injection mode (3 µm film thickness) (Restek Corp., Bellefonte, PA). The carrier gas utilized was helium at a flow of 10 mL/min. Peak areas were calculated using the software supplied with the GC and were compared to standard curves. In some experiments, chloroethenes were quantified using a Perkin-Elmer Autosystem 10.1021/es001285i CCC: $20.00

 2001 American Chemical Society Published on Web 12/23/2000

XL FID gas chromatograph fitted with a 2 m × 3 mm stainless steel column packed with 60/80 mesh Carbopak B/1% SP1000 (Supelco, Bellefonte, PA) and operated isothermally at 200 °C as described previously (13). For analyses of contaminants in cis-DCE, the column was maintained at 45 °C for 8 min, increased at 8 °C per minute to 220 °C, and then held at that temperature for 15 min. For gas chromatographic/mass spectrometric analyses of contaminants in cis-DCE, a Hewlett-Packard Model 5890 Series II gas chromatograph equipped with a 30 m × 0.25 mm × 0.23 µm film thickness, HP-5 (5%phenylmethyl silicone, Hewlett-Packard) fused silica capillary column connected to a Hewlett-Packard 5971 quadrupole mass selective detector was operated under the conditions previously described (30). The column oven was maintained at 40 °C for 8 min after injection of a 0.1 mL headspace sample, then increased by 8 °C per minute to 220 °C, at which it was held for 15 min. Under these conditions, both chloroform and cis-DCE had a retention time of 2.7 min. Growth Medium and Culture Conditions. Cultures of strain 195 were grown as previously described (29). Unless otherwise stated, inoculum sizes were 2% v/v, all incubations were done in duplicate, and each experiment presented was performed at least twice with similar results. Solutions of PCE in hexadecane (HD) were prepared as follows. HD was dispensed into 18 × 150 mm crimp-top culture tubes (Bellco Glass, Vineland, NJ) and was purged with a mixture of 70% N2/30% CO2 for 15 min. The tubes were then sealed with Teflon-coated butyl rubber stoppers (Wheaton, Millville, NJ) and aluminum crimps and were autoclaved at 121 °C for 30 min. PCE was purged with 70% N2/30% CO2 for 5 min and was then added by filter sterilization through a sterile Swinnex filter assembly fitted with a 0.2 µm cellulose acetate filter (Millipore Inc., Woburn, MA) as described by Holliger et al. (31). Cultures in 27 mL tubes containing 9.25 mL of medium received 0.75 mL of either a 2% v/v (called “low”) or 5% v/v (called “high”) PCE dissolved in HD, equivalent to 14.7 or 37.5 mmol added per liter of medium, respectively. Our initial studies indicated that upon equilibration the aqueous concentration of PCE was the equivalent of ca. 0.3 and 0.65 mmol PCE added per liter and that VC was only sparingly soluble and ETH practically insoluble in the HD phase. To accommodate the greatly increased amounts of PCE in cultures containing HD, increased amounts of H2 and HCO3- were added to respectively serve as electron donor and increase buffer capacity. Cultures were shaken at 150 rpm on a gyrotory shaker to ensure adequate transfer of H2 and PCE between the three phases. The amounts of VC and ETH in these cultures were corrected for the absorption in the HD phase through use of calibration curves prepared with the appropriate amounts of HD and medium present. Production of Biogenic cis-DCE. Biogenic cis-DCE was prepared using a culture isolated in this laboratory, called Desulfitobacterium strain DCE, which was isolated from a contaminated site at Plattsburgh Air Force Base, NY. This culture reduced PCE to cis-DCE (>99% of the products detected). We determined ca. 1000 bases of its 16S rDNA sequence (data not presented) as previously described (13), and sequence analysis showed that it is a member of the genus Desulfitobacterium. Its cultural characteristics resemble those of the recently described Desulfitobacterium frappieri sp. strain PCE (7). Cultures of Desulfitobacterium DCE were grown in 120 mL serum vials containing 60 mL of basal mineral medium used for strain 195 without any of the other nutrient supplements and were provided with 7 g/L sodium lactate as the electron donor and 1 mL 10% v/v PCE dissolved in hexadecane as previously described (31). After consumption of the PCE by the culture (lactate was present in excess), cis-DCE, which is considerably less soluble in

FIGURE 1. Effects of different dosing schedules on product formation from cis-DCE by strain 195. The doses of cis-DCE added to the cultures were (a) 0.3, 0.5, and 0.7 mmol added per liter of culture medium and (b) 0.15, 0.25, 0.35, and 0.50 mmol added per liter. hexadecane than is PCE, was quantified in the culture headspaces. Samples of the headspace were removed with a sterile N2-flushed syringe and were added to cultures of strain 195 in 27 mL tubes containing 10 mL of medium to achieve appropriate cis-DCE concentrations. Tubes not inoculated with strain 195 and receiving PCE showed no signs of dechlorination.

Results and Discussion Utilization of Chemically Synthesized cis-DCE by Strain 195. Cultures of strain 195, when transferred from medium containing PCE as electron acceptor to medium containing cis-DCE obtained from a commercial manufacturer, initially utilized cis-DCE with increasing rates, indicative of growth, but at much slower rates than other chloroethene electron acceptors such as PCE, TCE, 1,1-DCE, or 1,2-dichloroethane (29). When the dosing schedule typically used for PCE (24) of ca. 0.3, 0.5, and 0.7 mmol cis-DCE added per liter of culture medium was used (Figure 1a), the final dose was not consumed, even after incubation for 8.5 days. When the doses of cis-DCE added were reduced by 50% to ca. 0.15, 0.25, 0.35, and 0.50 mmol added per liter (Figure 1b), the initial performance of the culture was better. After 8.5 days from the start of the experiments, the culture in Figure 1b produced a total of 0.92 mmol VC per liter, nearly double the 0.52 mmol VC per liter produced by the culture in Figure 1a. Using the dosing schedule in Figure 1b, cultures could be successfully transferred in medium with cis-DCE at least three times (data not presented). Utilization of PCE and Other Chloroethenes by Cultures Grown on Chemically Synthetic cis-DCE. We have previously shown that cultures of strain 195 grown on TCE, 1,1-DCE, and 1,2-dichloroethane are capable of immediately using PCE (29), indicating that PCE dechlorination is constitutive when grown on those substrates. We examined the ability of cis-DCE-grown cells of strain 195 to dechlorinate chloroVOL. 35, NO. 3, 2001 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

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FIGURE 2. Utilization of PCE by cells grown on cis-DCE. ethenes containing an equal or higher number of chlorines after having consumed several doses of cis-DCE. Strain 195 was able to dechlorinate cis-DCE, 1,1-DCE, and TCE without any lag as has been previously demonstrated for other substrates (29). PCE, in contrast to the other chloroethenes tested, was not dechlorinated, even 32 days after it was added to the culture (Figure 2). Cultures grown on cis-DCE would not utilize a dose of PCE in several repetitions of this experiment, nor could they be transferred to fresh growth medium containing PCE as the electron acceptor. However, cultures transferred one time to a medium containing either 1,1-DCE or TCE regained the ability to utilize PCE after receiving two doses of the alternate substrate. Magnuson et al. (32) have shown that mixed cultures containing strain 195 contain a PCE reductive dehalogenase which dechlorinates PCE to TCE, and a TCE reductive dehalogenase, which dechlorinates TCE, cis-DCE, 1,1-DCE to VC, and VC slowly to ETH. Preliminary results indicate that these two enzyme are present in strain 195. These results are consistent with greater inhibition of the PCE dehalogenase by the inhibitory factor(s) in cis-DCE than TCE dehalogenase. Utilization of Biogenic cis-DCE by Strain 195. The slow utilization of commercially prepared cis-DCE, especially at high concentrations, indicated toxicity of this preparation to strain 195. However, it was not clear whether it was the cisDCE itself or an impurity in the preparation we used that was responsible for the toxicity. Initial studies indicated that gaseous cis-DCE derived from the headspace of a vial containing the neat liquid was as inhibitory to growth as was the liquid itself (data not presented), indicating that the toxic agent was either cis-DCE or a volatile contaminant. We decided to examine the toxicity of cis-DCE further by producing cis-DCE biologically using a culture, called Desulfitobacterium DCE, which reductively dechlorinated PCE stoichiometrically to cis-DCE (see Materials and Methods). As shown in Figure 3, a culture of strain 195 transferred to medium containing 0.14 or 0.65 mmol/L biogenic cis-DCE converted it nearly stoichiometrically to VC within a few days, whereas cultures supplied with synthetic cis-DCE did not convert it in that time. Cultures grown on biogenic cis-DCE retained the ability to utilize PCE (data not presented). Thus, cis-DCE is not intrinsically inhibitory, since biogenic cis-DCE was readily utilized by strain 195. There is other evidence of toxicity of synthetic cis-DCE to chloroethenedechlorinating cultures. There is evidence for cis-DCE toxicity to an anaerobic enrichment culture that reductively dechlorinated cis-DCE and VC (14), as demonstrated lack of toxicity of a biogenic cis-DCE preparation (P. L. McCarty, personal communication). This culture had properties different from those of strain 195 in that it could not use PCE or TCE and used VC more rapidly than does strain 195 (14). Neumann et al. (33) noted a 50% inhibition of a PCE reductive dehalogenase from Dehalospirillum multivorans by 14 mM cis-DCE. Since cis-DCE has not found widespread industrial 518

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FIGURE 3. Utilization of synthetic or biogenic cis-DCE by strain 195. Vinyl chloride was the primary product of reductive dechlorination in these cultures. Data points represent the average of duplicate tubes.

FIGURE 4. Analyses of synthetic and biogenic cis-DCE. Gas chromatograms of biogenic cis-DCE (a), synthetic cis-DCE (b), and chloroform (c). Analyses were performed as described in the Materials and Methods. Mass spectral analysis (d) of the gas chromatographic peak eluting at 2.7 min (see Materials and Methods) obtained from a sample containing synthetic cis-DCE. Both cis-DCE and chloroform elute at this time. use, it is generally considered to be biogenic rather than anthropogenic at contaminated sites. Therefore, it is unlikely that this inhibition phenomenon is relevant to those sites. Analysis of Synthetic and Biogenic cis-DCE and Chloroform Inhibition of Dechlorination. Figure 4a-b shows gas chromatograms obtained for synthetic cis-DCE and biogenic cis-DCE at low attenuation so that the cis-DCE peak eluting near 13.5 min was offscale and small peaks representing impurities are emphasized. The chromatogram for biogenic cis-DCE (Figure 4a) showed peaks that comigrated with 1,1DCE and with TCE, the former being a minor sideproduct the latter an intermediate in PCE dechlorination to cis-DCE by strain DCE. The chromatogram for synthetic cis-DCE (Figure 4b) showed several small peaks, including ones comigrating with 1,1-DCE and TCE, and more prominent peaks eluting immediately before cis-DCE and after cis-DCE as a shoulder. The peak eluting before cis-DCE as well as the other small peaks did not comigrate with any compound tested, including dichloromethane, 1,1-dichloroethane, 1,1,1-

trichloroethane, or carbon tetrachloride. The peak that migrated as a shoulder on the cis-DCE comigrated with chloroform (Figure 4c). We were unable to obtain better resolution between cisDCE and chloroform using a RTX-502.2 capillary column (see Materials and Methods) which is designed for use with chlorinated solvents and provides excellent resolution between the three DCE isomers. A perusal of sample chromatograms for various GC columns designed to separate aliphatic chlorinated organic compounds did not reveal any columns in which chloroform did not elute almost immediately after cis-DCE, thereby making it making it difficult to quantify chloroform in the presence of large amounts of cis-DCE. The boiling points of cis-DCE and chloroform are similar (60.7 °C and 61.7 °C, respectively (34)) and their Henry’s Law constants are 0.167 and 0.150 at 24.8 °C (35). Thus, their similar boiling points and hydrophobicity may partially explain the difficulty in resolving them. Analysis of the synthetic cis-DCE preparation indicated that if the peak was indeed chloroform, it would represent ca. 0.4% of the amount of cis-DCE in the sample. We also examined the synthetic cis-DCE preparation by gas-chromatography/mass spectroscopy. Chloroform and cis-DCE coeluted at 2.7 min from the column available for use with this instrument (see Materials and Methods), but analysis of the peak at 2.7 min from synthetic cis-DCE showed that ions with m/z values of 83, 85, and 87 were present (Figure 4d) which were not found in biogenic cis-DCE (data not presented) or the database mass spectrum for cis-DCE. These three mass values represent the fragment produced by removal of a chlorine atom from chloroform to form CHCl2+. This fragment forms a triplet in mass spectrograms because it can contain either 35Cl or 37Cl (natural ratio ≈ 3:1 (34)) and therefore each fragment can consist of CH35Cl2+, CH35Cl37Cl+, or CH37Cl2+ in order of abundance. The ratio between these peaks was 8.9/5.9/1.0 in the synthetic cisDCE preparation, whereas their ratios were 9.5/6.1/1.0 and 9.0/5.9/1.0 in mass spectra from authentic chloroform and the database values for chloroform, respectively. A set of small peaks from chloroform centered near m/z 118 representing the fragment CCl3+ were also visible in the spectrum from the synthetic cis-DCE (Figure 4d) but not in the biogenic material, while peaks from chloroform near m/z 47 were obscured by the cis-DCE peaks in that region. Although this analysis was not quantitative, it did verify the presence of a component in the synthetic cis-DCE with a mass spectrum essentially identical with chloroform. The fragment CHCl2+ could also be derived from dichloromethane, but previous gas chromatographic analysis (Figure 4b) demonstrated that this compound was not present. We examined whether adding chloroform to biogenic cisDCE in amounts comparable to the amount found in the synthetic preparation was toxic to growth of strain 195. We found that 1.6 µM, about double the amount of chloroform as that estimated a completely inhibitory dose of synthetic cis-DCE, was required for complete inhibition of cis-DCE dechlorination (data not presented). Gas chromatographic analysis (Figure 4b) of the synthetic cis-DCE preparation indicates the presence of other compounds which could also be toxic. Also unclear was the fate of small amounts of chloroform in the cultures. Preliminary studies (not presented) indicate little or no chloroform absorption into the stopper, but it is not clear whether chloroform is metabolized by strain 195. No dichloromethane was found in gas chromatographic traces of cultures exposed to chloroform. Whether or not chloroform is completely responsible for the inhibition of reductive dechlorination in strain 195 by cis-DCE, it is a potent inhibitor. There is other evidence for chloroform inhibition of reductive dechlorination of chlo-

FIGURE 5. VC dechlorination to ETH by strain 195 in the absence (a) and presence (b) of PCE at the time of inoculation. After 30 days of incubation, three consecutive increasing doses of PCE (0.3, 0.5, and 0.7 mmol/L) were added to the culture in (a). roethenes. Studies in J. M. Gossett’s laboratory (36) on a mixed PCE-dechlorinating anaerobic culture showed that ca. 1 mg/L aqueous concentration (ca. 8.4 µM) of chloroform led to complete inhibition of PCE dechlorination within a day, an inhibition similar to that which we detected in our culture (Figure 4). This similar inhibition is not surprising since the culture studied was derived from the one from which strain 195 was isolated. Chlorinated methanes were inhibitory to a PCE reductive dehalogenase from D. multivorans, with chloroform (50% inhibition at 0.25 µM) and chloromethane (50% inhibition by 0.8 µM) being the most inhibitory (33). This same enzyme was also inhibited by high concentrations of cis-DCE, which was presumably commercially synthesized. Chloroform is known to be a potent inhibitor of methanogenesis (37, 38) and is believed to interfere with corrinoid function. The presence of corrinoids has been demonstrated or implicated in chloroethene reductive dehalogenases (32, 39, 40). Thus, the inhibition of reductive dechlorination of chloroethenes by chloroform may be a general phenomenon that may be relevant to sites at which chloroform is a cocontaminant with chloroethenes. These problems may be obviated by the breakdown of chloroform at those sites. At this time, the only process known for anaerobic breakdown of chloroform is a slow cometabolic dehalogenation by anaerobic respirers such as methanogens, acetogens, and sulfate reducers (37, 41, 42). Indeed recent studies on a highrate PCE-dechlorinating enrichment culture (43) in which molecular biological techniques indicated the presence of an organism closely related to strain 195 showed that carbon tetrachloride was reductively dechlorinated to chloroform, which was subsequently transformed to dichloromethane. There was considerable inhibition of reductive dechlorination of chloroethenes in this culture. ETH Production from VC in Inoculated Cultures. As shown in Figure 5a, during 30 days of incubation with ca. 0.4 VOL. 35, NO. 3, 2001 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

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FIGURE 6. ETH formation in the presence or absence of PCE during growth. Cultures were either fed a new dose of PCE before the previous one had been consumed (a) or were allowed 2 days to dechlorinate VC after each PCE dose had been consumed (b). mmol VC added per liter of growth medium, there was a slow conversion of VC to ETH. The slightly more rapid VC disappearance versus ETH appearance was attributed to absorption into the stopper as indicated by uninoculated controls (data not presented). Figure 5c shows this ETH appearance with greater sensitivity and demonstrates that there was a gradual decrease in the rate of ETH appearance during the first 30 days. A dose of 0.3 mmol/l PCE added at Day 30 was utilized in 14 days, but with only a slight accumulation of dechlorination products (mainly VC), with the decrease in PCE concentration attributed mainly to its absorption into the stopper. Two subsequent PCE doses were consumed at increasing rates, following which there was significant ETH accumulation. As shown in Figure 5b, when a dose of PCE was added together with VC at the time of inoculation, PCE was metabolized to VC by Day 4, and, and once the PCE was depleted, there was a much more rapid accumulation of ETH than when VC was added in the absence of PCE. Thus VC dechlorination by strain 195 does not support growth, follows first-order kinetics (13, 28), and requires the addition of a metabolically useful substrate, such as PCE, for sufficient growth to occur to allow significant VC dechlorination. These are hallmarks of a cometabolic process. It is not clear why strain 195 cannot conserve energy from VC dechlorination, since it is as thermodynamically favorable as dechlorination of the other chloroethenes (44). There is evidence that other organisms are capable of conserving energy from reductive dechlorination of VC to ETH (14, 21), but the microorganisms in these mixed cultures are as yet poorly characterized. Inhibition of VC dechlorination to ETH by PCE. Figure 6a demonstrates that in a culture in which four doses of PCE were added in rapid succession without allowing PCE to be depleted to concentrations below 1 mmol per liter, ETH only accumulated slightly until the final dose of PCE was consumed, after which there was significant ETH accumulation. In cultures in which an additional dose of PCE was added 2 days after the previous dose was consumed (Figure 520

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FIGURE 7. Total ETH (a) and VC (b) produced and aqueous levels of PCE (a) by strain 195 grown in media with a hexadecane (HD) phase. PCE was added differently to cultures in a HD phase either at “low” (total of 14.7 mmol/L PCE added) or “high” (total of 36.7 mmol/L PCE added) concentrations (see Materials and Methods). Also included are cultures without HD (4 doses; total of 2.4 mmol/L PCE added). 6b), ETH was produced almost exclusively during the periods when PCE was absent from the medium. Cultures in which 4 days were allowed for VC dechlorination after PCE was consumed only led to about 20% more ETH produced from VC than those shown in Figure 6b (data not presented). The other chloroethenes that strain 195 could use as electron acceptors, TCE, cis-DCE, and 1,1-DCE, had similar inhibitory effects on VC dechlorination to ETH (data not presented). These results are similar to those for the mixed culture containing strain 195 (28) in which all other chloroethenes except trans-DCE were inhibitory to VC dechlorination. It cannot be determined whether this inhibition is direct, i.e., binding to the same site on an enzyme or cofactor, or perhaps a competition for electrons derived from H2. Recent evidence in our laboratory (Nijenhuis and Zinder, unpublished) has shown that dechlorination rates in whole cells were much higher when the artificial electron donor reduced methyl viologen was used as than when H2 was utilized, suggesting that electron flow from H2 was rate limiting in these cells. VC Dechlorination to ETH in Cultures Grown with PCE Dissolved in Hexadecane. To examine further the interaction between the presence of PCE and VC dechlorination to ETH, we introduced a hexadecane (HD) phase into our cultures. PCE is highly soluble in hexadecane, and such a method is a useful way to add large amounts of PCE while maintaining a low aqueous concentration (31). Preliminary studies showed that similar to results with Dehalobacter restrictus (31), strain 195 grew well when PCE was dissolved in a HD phase. Figure 7 shows a comparison for the total amounts of VC and ETH produced by cultures to which two different concentrations of PCE dissolved in HD had been added (called “low” and “high” see Materials and Methods) as well as a control culture to which 2.4 mmol/L of PCE was added in four doses, not allowing the PCE concentration to reach

zero, as in Figure 6a. Cultures receiving the “low” PCE/HD solution produced the same amount of VC as controls without HD during the first 7 days and somewhat more VC than cultures amended with the “high” PCE/HD solution. Significantly, cultures receiving the “low” PCE/HD solution, in which the PCE concentration was maintained below 0.3 mmol/L, produced about 4-fold more ETH than the cultures amended with the “high” PCE/HD solution. ETH began accumulating by Day 6 in the “low” PCE cultures, long before the concentration of the PCE decreased significantly. It is unlikely that the actual aqueous concentrations were much lower than those measured in the culture headspaces at this time since the cultures were vigorously agitated and the rate of PCE utilization was still slow at this time. By Day 20, product formation stopped in all cultures, presumably due to nutrient depletion or possibly buildup of toxic products. Thus, the interaction between VC utilization and that of other chloroethenes used for dehalorespiration by strain 195 is complex. Since strain 195 cannot grow using VC, it needs an electron acceptor it can use for dehalorespiration and growth such as PCE to reach high enough cell densities to cause appreciable VC utilization. However, at the same time, high concentrations of these chloroethenes inhibit VC conversion to ETH although the mechanism is unclear. At chloroethene-contaminated sites at which reductive dechlorination is occurring, VC is produced biogenically from more highly chlorinated chloroethenes, so that there should be sufficient growth of D. ethenogenes or similar organisms. It is also unlikely that the concentrations of these chloroethenes would exceed 600 µM or higher needed to fully inhibit VC dechlorination (Figure 6), except perhaps immediately at the source zone. Moreover, there may be organisms present more proficient at using VC than D. ethenogenes (14, 21) so that the prospects for complete dechlorination of chloroethenes at most contaminated sites with appropriate electron donors and organisms would be good.

Acknowledgments Supported by the U. S. Air Force Armstrong Laboratory, Environmental Quality Directorate, Tyndall Air Force Base, Florida and the “La Caixa” Foundation, Catalonia, Spain (X. Maymo´-Gatell). We thank James Gossett and Donna Fennell for advice and providing butyrate-grown enrichment culture used as a nutrient supplement in these studies, Tim Anguish for help with culture studies, Eugene Madsen for use of his gas chromatograph/mass spectrometer, and Amy Carroll for determining part of the 16S rDNA sequence of Desulfitobacterium strain DCE. We also thank Wil de Bruijn for useful early conversations about the toxicity of cis-dichloroethene.

Literature Cited (1) Westrick, J. J.; Mello, J. W.; Thomas, R. F. J. Am. Water Works Assoc. 1984, 76 (5), 52-59. (2) Hinchee, R. E.; Leeson, A.; Semprini, L. Bioremediation of Chlorinated Solvents; Battelle Press: Columbus, OH, 1995; p 338. (3) Holliger, C.; Hahn, D.; Harmsen, H.; Ludwig, W.; Schumacher, W.; Tindall, B.; Vazquez, F.; Weiss, N.; Zehnder, A. J. B. Arch. Microbiol. 1998, 169, 313-321. (4) Gerritse, J.; Renard, V.; Pedro-Gomes, T. M.; Lawson, P. A.; Collins, M. D.; Gottschal, J. C. Arch. Microbiol. 1996, 165, 132140. (5) Krumholz, L. R. Int. J. Syst. Bacteriol. 1997, 47, 1262-1263. (6) Miller, E.; Wohlfarth, G.; Diekert, G. Arch. Microbiol 1997, 166, 379-87. (7) Miller, E.; Wohlfarth, G.; Diekert, G. Arch. Microbiol. 1997, 168, 513-519. (8) Sharma, P. K.; McCarty, P. L. Appl. Environ. Microbiol. 1996, 62, 761-765. (9) Neumann, A.; Scholz-Muramatsu, H.; Diekert, G. Arch. Microbiol 1994, 162, 295-301.

(10) Harkness, M. R.; Bracco, A. A.; M. J. Brennan, J.; Deweerd, K. A.; Spivak, J. L. Environ. Sci. Technol. 1999, 33, 1100-1109. (11) Holliger, C.; Wohlfarth, G.; Diekert, G. FEMS Microbiol. Rev. 1998, 22, 383-398. (12) Lo¨ffler, F. E.; Flynn, S.; Tiedje, J.; Fathepure, B.; Shulz, N.; Adriaens, P. Abstracts of the General Meeting of the American Society for Microbiology; 1998, 98, 429. (13) Maymo´-Gatell, X.; Chien, Y. T.; Gossett, J. M.; Zinder, S. H. Science 1997, 276, 1568-1571. (14) Rosner, B. M.; McCarty, P. L.; Spormann, A. M. Appl. Environ. Microbiol. 1997, 63, 4139-4144. (15) Federal Register 1989, 54, 22062-22160. (16) Hartmans, S.; de Bont, J.; Tramper, J.; Luyben, K. Biotechnol. Lett. 1985, 7, 383-388. (17) Davis, J. W.; Carpenter, C. Appl. Environ. Microbiol. 1990, 56, 3878-3880. (18) Bradley, P. M.; Chapelle, F. H. Environ. Sci. Technol. 1998, 32, 553-557. (19) Bradley, P. M.; Chapelle, F. H. Environ. Sci. Technol. 1996, 30, 2084-2086. (20) Bradley, P. M.; Chapelle, F. H.; Lovley, D. R. Appl. Environ. Microbiol. 1998, 64, 3102-5. (21) Lo¨ffler, F. E.; Ritalahti, K. M.; Tiedje, J. M. Appl. Environ. Microbiol. 1997, 63, 4982-4985. (22) DiStefano, T. D.; Gossett, J. M.; Zinder, S. H. Appl. Environ. Microbiol. 1991, 57, 2287-2292. (23) Freedman, D. L.; Gossett, J. M. Appl. Environ. Microbiol. 1989, 55, 2144-2151. (24) Maymo´-Gatell, X.; Tandoi, V.; Gossett, J. M.; Zinder, S. H. Appl. Environ. Microbiol. 1995, 61, 3928-3933. (25) de Bruin, W. P.; Kotterman, M. J. J.; Posthumus, M. A.; Schraa, G.; Zehnder, A. J. B. Appl. Environ. Microbiol. 1992, 58, 19962000. (26) Beeman, R. E.; Howel, J. E.; Shoemaker, S. H.; Salazar, E. A.; Buttram., J. R. A field evaluation of in situ microbial reductive dehalogenation by the biotransformation of chlorinated ethenes; Hinchee, R. E., Leeson, A., Semprini, L., Ong, S. K., Eds.; Lewis Publishers: Boca Raton, Florida, 1993; pp 14-27. (27) Major, D. W.; Hodgins, E. W.; Butler, B. J. Field and laboratory evidence of in situ biotransformation of tetrachloroethene to ethene and ethane at a chemical transfer facility in north Toronto; Hinchee, R. E., Olfenbuttel, R. F., Eds.; Butterworth-Heinemann: Boston, 1991; pp 113-133. (28) Tandoi, V.; DiStefano, T. D.; Bowser, P. A.; Gossett, J. M.; Zinder, S. H. Environ. Sci. Technol. 1994, 28, 973-979. (29) Maymo´-Gatell, X.; Anguish, T.; Zinder, S. H. Appl. Environ. Microbiol. 1999, 65, 3108-3113. (30) Wilson, M.; Madsen, E. L. Environ. Sci. Technol. 1996, 20992103. (31) Holliger, C.; Schraa, G.; Stams, A. J. M.; Zehnder, A. J. B. Appl. Environ. Microbiol. 1993, 59, 2991-2997. (32) Magnuson, J. K.; Stern, R. V.; Gossett, J. M.; Zinder, S. H.; Burriss, D. R. Appl. Environ. Microbiol. 1998, 64, 1270-1275. (33) Neumann, A.; Wohlfarth, G.; Diekert, G. J. Biol. Chem. 1996, 271, 16515-16519. (34) Lange’s Handbook of Chemistry, 13th ed.; Dean, J. A., Ed.; McGraw-Hill: New York, 1985. (35) Gossett, J. M. Environ. Sci. Technol. 1987, 21, 202-208. (36) Carney, A. P. M.S. Thesis, Cornell University, 1995. (37) Bagley, D. M.; Gossett, J. M. Appl. Environ. Microbiol. 1995, 61, 3195-201. (38) Bauchop, T. J. Bacteriol. 1967, 94, 171-175. (39) Schumacher, W.; Holliger, C.; Zehnder, A. J.; Hagen, W. R. FEBS Lett. 1997, 409, 421-5. (40) Neumann, A.; Wohlfarth, G.; Diekert, G. J. Bacteriol. 1998, 180, 4140-4145. (41) Becker, J. G.; Freedman, D. L. Environ. Sci. Technol. 1994, 28, 1942-1949. (42) Mikesell, M. D.; Boyd, S. A. Appl. Environ. Microbiol. 1990, 56, 1198-1201. (43) Adamson, D. T.; Parkin, G. F. Environ. Sci. Technol. 2000, 34, 1959-1965. (44) Vogel, T. M.; Criddle, C. S.; McCarty, P. L. Environ. Sci. Technol. 1987, 21, 722-736.

Received for review May 22, 2000. Revised manuscript received October 17, 2000. Accepted November 8, 2000. ES001285I

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