Environ. Sci. Technol. 1991, 25, 982-984
COMMUNICATIONS Stimulation of the Reductive Dechlorination of Tetrachloroethene in Anaerobic Aquifer Microcosms by the Addition of Toluene G. W. Sewell*+tand S. A. Gibson2 US. Environmental Protection Agency and NSI Technology Services Corporation, Robert S. Kerr Environmental Research Laboratory, Ada, Oklahoma 74820
Introduction Chlorinated solvents are among the most common industrial contaminants of groundwater (1). Tetrachloroethene (PCE) is resistant to aerobic biodegradation; however, it is occasionally reductively dechlorinated in anoxic contaminated aquifers (2, 3). This is particularly true if the subsurface also contains other organic compounds that can serve as electron donors and whose utilization by subsurface bacteria will deplete the available oxygen. Chlorinated solvents are followed closely in importance as groundwater contaminants by alkylbenzenes. Alkylbenzenes most commonly enter the subsurface in the form of fuel spills and are degraded under both aerobic and anaerobic conditions by subsurface microorganisms. Freedman and Gossett ( 4 ) have shown that a number of low molecular weight organic compounds can serve as electron donors for reductive dechlorination, and recently, Scholz-Muramatsu et al. ( 5 ) demonstrated that benzoate can also serve as a source of reducing equivalents for reductive dechlorination of PCE. Grbic-Galic and Vogel (6) have shown that benzoate is an intermediate of anaerobic toluene degradation. Overall these findings suggest that certain alkylbenzenes, such as toluene, could potentially serve as electron donors for the reductive dechlorination of PCE. Field studies conducted by this laboratory have suggested that this may indeed be occurring a t field sites exposed to both alkylbenzenes and chlorinated solvents (7). In this study, the biologically mediated interactions of toluene and PCE under anaerobic conditions were investigated by using microcosms constructed with aquifer solids from an area that was exposed to both alkylbenzenes and chlorinated ethenes at the U S . Coast Guard Air Station, in Traverse City, MI. Materials and Methods Construction, Incubation, and Sampling of Microcosms. Aquifer solids were collected by using the aseptic/anaerobic coring procedure as described by Leach et al. (8) from an anaerobic subsurface zone thought to contain microorganisms capable of metabolizing dissolved fuel components or partially degraded fuel components. The core material was a medium-grain beach sand, light gray to dark gray in color, with a total organic carbon content ranging from 0.040 to 0.011% (w/w). Sealed, collected cores were placed in an anaerobic chamber where 50 g of mixed, saturated core material was added aseptically to sterile 160-mL serum bottles, which were then completely filled with sterile spring water (Byrd's Mill Spring-Municipal water supply for the city of Ada, OK, pH 7.4, total US. Environmental Protection Agency.
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alkalinity 326 CaC03 equiv) amended with ammonium phosphate (10 mM ammonium and 5 mM phosphate final concentration). Reducing conditions were maintained by the addition of sodium sulfide (- 1 mM final concentration), and resazurin (0.0001% w/v) was added as a redox indicator. Microcosms were spiked with PCE (6 ppm), or PCE and toluene (10 ppm), or were unamended. Bottles were sealed with Teflon-faced butyl rubber stoppers and aluminum crimp seals. Microcosms were incubated in an anaerobic chamber a t 20 "C and protected from light. Experiments were performed with three to four active experimental units and two to four autoclaved controls for each condition. Autoclaved controls were constructed as above with the exception that the serum bottles were autoclaved for 1h on two successive days after core addition and before being filled and spiked. Microcosms were repetitively sampled by introduction of a syringe needle down the side of a partially opened stopper and removal of a 2-mL aqueous subsample. Zero headspace was maintained by the addition of 2 cm3 of clean, sterile 6-mm Kimex glass beads immediately after subsample removal. All replicates were analyzed a t each time point and reported concentrations represent mean values. Reflected-Light Fluorescence Microscopy. The presence of metabolically active (fluorescent) methanogenic bacteria in aquifer microcosms was qualitatively determined with a Zeiss Axioplan microscope in the epi-illumination mode: exciter filter 395-440, chromatic beam splitter F T 460, barrier filter L P 470. HPLC Analysis. Metabolic acids (acetate, lactate, propionate, and butyrate) were determined by HPLC as described Chamkasem et al. (9). The lower limit for acetate quantification was 1.3 ppm. GC Analysis. Chloroethenes were determined by using a cryogenically equipped headspace GC system with a flame ionization detector. The detector temperature was set a t 200 "C and the gas flow rates were set at 40 mL/min for H,, 400 mL/min for air, and 15 mL/min for the carrier gas (high-purity helium). The analysis was done using a 15-m DB-5 capillary column (0.517-mm i.d., and a 1.5-pm film]. The initial temperature was -50 "C for 1 min, followed by a temperature program to a final temperature of 90 "C. The septum purge was a t 2 mL/min. The headspace sampler bath temperature was 80 "C, valve and loop temperature was 75 "C, carrier pressure was 0.6 bar, carrier flow rate was 1 2 mL/min, and auxiliary pressure was 0.4 bar. Samples were allowed to equilibrate at 80 "C for 1h before injection. The lower limit quantification for chloroethenes and toluene was 200 ppb. Results and Discussion After 85 days of incubation toluene levels were lower in active vs autoclaved microcosms (Figure 1). Toluene levels
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were approximately 1/3 of the original concentration after 100 days of incubation in active toluene/PCE-spiked microcosms while autoclaved controls showed no significant loss. After 130 days of incubation, analysis of toluene/PCEspiked, active microcosms indicated a loss of PCE, which was not seen in autoclaved microcosms or microcosms spiked with PCE alone (Figure 2). Acetate was detected in toluene/PCE-spiked microcosms after 100 days of incubation, reaching a maximum concentration of approximately 180 pM (11 ppm) by 184 days of incubation (Figure 3). Of the four metabolic acids analyzed for by HPLC (acetate, propionate, lactate, and butyrate), only acetate was detected in above background levels. No accumulation of acetate was detected in autoclaved, unspiked, or PCE-spiked microcosms. In PCE/toluene-spiked microcosms, trichloroethene (TCE) and dichloroethene (DCE) were detected after 120 and 140 days, respectively. In the absence of an added oxidizable substrate (toluene), no daughter products were detected in PCE-spiked microcosms over the entire duration of the experiment (232 days). The initiation of reductive dechlorination activity varied within the set of toluene/PCE microcosms and the results in Figure 3 represent the mean values; however, within individual microcosms PCE and DCE(s) were never detected together a t the same sampling time point. DCE was present as a mixture of cis- and trans-DCE a t a ratio of approximately
Figure 3. Toluene, PCE, and metabolic products in active microcosms spiked with toluene and PCE.
12, respectively, which was fairly constant through out the experiment. No 1,l-DCE was detected a t any time. DCE levels reached a maximum of slightly less than 3 ppm after 175 days of incubation and did not significantly decrease after 230 days. The final concentration of DCE(s) was approximately 20 pM, which would indicate that little or none of the original PCE was transformed past the level of DCE, This would explain our failure to detect vinyl chloride or ethene by GC/mass spectroscopy. The reason for the “stalling” of the reductive dechlorinating activity a t the level of DCE is unclear. We have in the past been able to drive the dechlorination of PCE by the addition of oxidizable substrate to the level of vinyl chloride (data not shown), as others have reported ( 4 ) . The reductive dechlorination of chloroethenes requires the gain of 2 mol of electrons/mol of chloride ion removed (10). Assuming complete oxidation of toluene to C 0 2 by the following equation, 14H20+ C7H8 7 c 0 2 + 36H’ + 36e-, then 18 reducing equivalents (18 electron pairs) would theoretically be available for reductive dechlorination reactions. Apparently most of the reducing equivalents had an alternate fate, with only -3% of the theoretical total being used for reductive dechlorination. Even assuming the conversion of toluene to acetate, C7H8 7H20 3.5C2H4O2 22H+ 22e-, over 90% of the potential reducing equivalents are unaccounted for. One possible fate for the unaccounted for reducing potential is in the formation of methane. Although methane was detected in toluenespiked microcosms it was not at significantly greater levels than that detected in unamended or autoclaved microcosms. However, metabolically active, coccoid-shaped, methanogenic bacteria were repeatedly visualized in samples from the active toluene and toluene/PCE-spiked microcosms and were not seen in unamended microcosms. The onset of reductive dechlorinating activity does not closely parallel the loss of toluene (Figure 3). Chronologically, the formation of acetate, which also lags behind toluene removal, appears more related to the onset of reductive dechlorinating activity. This would be consistent with an intermediate of toluene degradation, such as benzoate, acting as the immediate electron donor for PCE reduction in a manner similar to that reported by Scholz-Muramatsu et al. (5). It is also possible that acetate could be acting as an electron donor ( 4 ) . Overall, the results of this study indicate that toluene can act as an initial source of reducing potential for the reductive dechlorination of chloroethenes under anaerobic conditions. Also, the cultivation of this consortium from a site impacted by alkylbenzenes and chloroethenes is an indication that this process may occur in contaminated aquifers.
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Acknowledgments
We note the important contributions of NSI Technological Services, analytical group, Ada, OK, in this work. Literature Cited (1) Westrick, J. J.; Mello, J. W.; Thomas, R. F. J.-Am. Water Works Assoc. 1984, 76, 52. (2) Roberts, P. V.; Schreiner, J.; Hopkins, G. D. W a t e r Res. 1982, 16, 1025. (3) Wilson, B. H. In U.S. Geological Survey Toxic Substances Hydro 1ogy Progra m-Proce ed ings o f t h e T e chnica 1 Meeting, Phoenix, AZ, September 26-30, 1988; WaterResour. Znuest. Rep. (U.S. Geol. Surv.) 1989, No. 88-4220. (4) Freedman, D. L.; Gossett, J. M. Appl. Environ. Microbiol. 1989, 55, 2144. (5) Scholz-Muramatsu, H.; Szewzyk, R.; Szewzyk, U.; Gaiser, S. FEMS Microbiol. L e t t . 1990, 66, 81. (6) Grbic-Galic, D.; Vogel, T. M. Appl. Environ. Microbiol. 1987, 53, 254.
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(7) Sewell, G. W.; Wilson, B. H.; Wilson, J. T.; Kampbell, D. H.; Gibson, S.A. In Chemical and Biochemical Detoxification o f Hazardous Waste II; Glaser, J. A., Ed.; Lewis Publishers: Chelsea MI, in press. (8) Leach, L. E.; Beck, F. P.; Wilson, J. T.; Kampbell, D. H. In Proceedings of the Second National Outdoor Action
Conference on Aquifer Restoration, Ground Water Monitoring and Geophysical Methods, Las Vegas, NV; National Water Well Association: Dublin, OH, 1988; p 31. (9) Chamkasem, N.; Hill, K. D.; Sewell, G. W. J . Chromatogr. 1991,536, 193. (10) Vogel, T. M.; Criddle, C. S.; McMarty, P. L. Enuiron. Sci. Technol. 1987, 21, 722. Received f o r review December 3, 1990. Revised manuscript received January 15,1991. Accepted January 22,1991. Although the research described i n this paper is supported by the U.S. Environmental Protection Agency through a n in-house research program, it has not been subjected to Agency review and therefore does not necessarily reflect the views of t h e Agency, and no official endorsement should be inferred.
