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Methods
Moravek Biochemicals, Inc., Brea, CA) as described previously (2, 3). The radiochemical purity of the [1,2-14C]DCE (0.6 µCi/ µmol specific activity) was confirmed in our lab by radiometric detection gas chromatography and liquid scintillation counting to be g99.9%. Study Site. [1,2-14C]DCE mineralization studies were initiated using bed sediments from the Naval Air Station (NAS) Cecil Field, Jacksonville, FL. The NAS Cecil Field bed sediments, which were described in detail previously (2), were collected from a shallow, freshwater stream that receives groundwater contaminated with low concentrations (e0.2 µM or 20 µg/L) of cis-DCE. Microcosm Studies. Aerobic microcosms were prepared as described previously (12). In brief, 20-mL serum vials were created with 15 g of saturated bed sediment, sealed with Teflon-lined butyl rubber stopper/base trap assemblies (12), and flushed with an excess (1000 mL) of zero air. Experimental treatments were prepared in triplicate. Duplicate killed control microcosms were prepared as described and autoclaved twice for 1 h at 15 psi and 121 °C. Sediment microcosms were preincubated for 3 days and then amended with approximately 0.01 µCi of [1,2-14C]DCE. Initial dissolved concentrations in equilibrium with the headspace were estimated based on experimentally determined adsorption (2) and Henry’s law coefficients (2) to be about 5 µM. 14CO2 was collected in 3 M KOH and quantified by liquid scintillation counting (12). Recovery as 14CO2 was confirmed in select microcosms as described previously (12, 13). [1,2-14C]DCE mineralization was also examined in liquid cultures with [1,2-14C]DCE as the sole electron donor and an air atmosphere. Following the microcosm experiment, the experimental microcosm with the highest rate of [1,2-14C]DCE mineralization was shaken and allowed to settle. A total of 0.1 mL of clear supernatant (dissolved organic content, DOC e 100 mg/L) was then transferred to 25-mL serum tubes containing 9.9 mL of sterile, minimal media. The minimal media consisted of 98% phosphate buffer (8.3 mM, pH ) 7.2), 1% Wolfes mineral solution (14), 1% Wolfes vitamin solution (14), and 0.5 µCi of [1,2-14C]DCE. Initial dissolved concentrations in equilibrium with the headspace were estimated based on experimentally determined adsorption (2) and Henry’s law coefficients (2) to be about 50 µM. Experimental treatments were prepared in triplicate. Duplicate killed control microcosms were prepared in the same manner and autoclaved twice for 1 h at 15 psi and 121 °C. Duplicate cell-free controls were prepared as described without the addition of bed sediment innoculum. [1,2-14C]DCE mineralization was monitored as described for the microcosm study. Following each incubation, an aliquot of culture media was removed from each tube and used as the innoculum for the subsequent incubation. A total of three transfers were completed for a 107 final dilution of the original bed sediment slurry innoculum. On the basis of a measured DOC of e100 mg/L in the initial bed sediment slurry innoculum, the final concentration of bed sediment DOC in transfer 3 was less than 10 ng/L. For the final transfer, the potential for microbial growth under these culture conditions was assessed by monitoring absorbance (450 and 660 nm) throughout the incubation and initial and final direct counts using the acridine orange method.
Chemicals. Aerobic oxidation of 1,2-DCE was evaluated using a neat mixture of [1,2-14C]DCE (29% trans, 71% cis isomers,
Results and Discussion
Aerobic Microbial Mineralization of Dichloroethene as Sole Carbon Substrate PAUL M. BRADLEY* AND FRANCIS H. CHAPELLE U.S. Geological Survey, 720 Gracern Road, Suite 129, Columbia, South Carolina 29210-7651
Microorganisms indigenous to the bed sediments of a black-water stream utilized 1,2-dichloroethene (1,2-DCE) as a sole carbon substrate for aerobic metabolism. Although no evidence of growth was observed in the minimal salts culture media used in this study, efficient aerobic microbial mineralization of 1,2-DCE as sole carbon substrate was maintained through three sequential transfers (107 final dilution) of the original environmental innoculum. These results indicate that 1,2-DCE can be utilized as a primary substrate to support microbial metabolism under aerobic conditions.
Introduction Dichloroethene (DCE) isomers are U.S. EPA priority groundwater pollutants due to their innate toxicity and their tendency to be reduced to vinyl chloride (VC, a known human carcinogen) under anaerobic conditions. Thus, identifying the biodegradation mechanisms that affect the fate and transport of these compounds is a significant environmental concern. A number of studies have demonstrated that microorganisms can degrade DCE under aerobic conditions (1-11). Cometabolic oxidation of DCE to CO2 under aerobic conditions has been demonstrated for methanotrophs (810), phenol oxidizers (1, 5, 11), toluene oxidizers (5), and propane oxidizers (7). For each of these processes, oxidation of DCE was apparently a fortuitous occurrence with no clear benefit to the responsible organisms and, consequently, required a primary substrate in sufficient concentration to support microbial growth and energy production (7-10). In contrast, several recent investigations indicate that microorganisms can oxidize DCE to CO2 in the absence of an apparent alternative substrate (2-4, 6). Significant aerobic oxidation of cis-1,2-DCE was demonstrated for an organicrich stream bed sediment (2, 3), organic-rich surface soils (6), and organic-poor aquifer sediments (3, 4, 6). However, even though no alternative substrates were apparent in these studies, each was conducted with a natural sediment and the complexity of such matrixes makes mechanistic conclusions equivocal (3, 4, 6). The purpose of this paper is to present evidence that microorganisms collected from a stream bed sediment and maintained on minimal salts media were able to oxidize 1,2-DCE as a sole carbon substrate under aerobic conditions.
* Corresponding author telephone: (803) 750-6125; fax: (803) 7506181; e-mail:
[email protected]. 10.1021/es990785c Not subject to U.S. Copyright. Publ. 2000 Am. Chem. Soc. Published on Web 11/24/1999
Rapid aerobic oxidation of [1,2-14C]DCE to 14CO2 was observed in bed sediment microcosms under aerobic conditions (Figure 1). Approximately 100% of the initial [1,2-14C]DCE VOL. 34, NO. 1, 2000 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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FIGURE 1. Aerobic mineralization of [1,2-14C]DCE to 14CO2 in microcosms containing stream bed sediment from NAS Cecil Field. Data are means ( SD for triplicate experimental and duplicate killed control microcosms. Differences between treatment means with different superscripts are statistically significant according to the Kruskal-Wallis one-way analysis of variance on ranks (P < 0.05).
FIGURE 2. Aerobic mineralization of [1,2-14C]DCE to 14CO2 in liquid cultures containing [1,2-14C]DCE as the sole carbon substrate. Data are shown for three consecutive transfers of the original stream bed sediment innoculum. Data are means ( SD for triplicate experimental, duplicate sterile control, and duplicate cell-free control microcosms. Differences between treatment means with different superscripts are statistically significant according to the Kruskal-Wallis one-way analysis of variance on ranks (P < 0.05). radiolabel was recovered as 14CO2 within 8 days. [1,2-14C]DCE mineralization was attributable to biological activity because the recovery of 14CO2 in autoclaved control microcosms was only 3% (Figure 1). These results are consistent with previous reports, at this (2, 3) and other sites (4, 6), that microbial oxidation of DCE in the absence of an apparent alternative substrate can be significant in environmental samples under aerobic conditions. Because the stream bed sediments at NAS Cecil Field are a complex environment, however, a conclusive identification of the underlying mechanism was not possible under these culture conditions. Efficient mineralization of [1,2-14C]DCE was also observed in all liquid culture incubations conducted in this study (Figure 2). In general, the rate and extent of [1,2-14C]DCE mineralization decreased with each dilution. However, 46 ( 5% recovery as 14CO2 was observed even in the final culture transfer (107 dilution of the original bed sediment slurry). Consistent with previous investigations of DCE mineralization by these bed sediment microorganisms (2, 3), final headspace analyses indicated that both cis and trans isomers of DCE 222
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were degraded in approximately equimolar amounts in liquid cultures (data not shown). For all dilutions, [1,2-14C]DCE mineralization was attributable to biological activity, because the recovery of 14CO2 in autoclaved and cell-free control microcosms was e7% (Figure 2). These results demonstrate that microorganisms indigenous to the stream bed sediments at NAS Cecil Field were able to use [1,2-14C]DCE as the sole carbon substrate to support metabolism under aerobic conditions. This in turn suggests that utilization of 1,2-DCE as a primary substrate is a significant mechanism contributing to the rapid degradation of 1,2-DCE observed in these stream bed sediments under aerobic conditions. No evidence of microbial growth was observed in liquid cultures containing [1,2-14C]DCE as sole carbon substrate (data not shown). Although metabolism of 1,2-DCE as the sole carbon substrate was the focus of this study, the potential for microbial growth on [1,2-14C]DCE was assessed in Transfer-3 culture tubes (107 dilution). No significant change in absorbance (450 and 660 nm) or acridine orange direct counts (104-105 cells/mL throughout the incubation) was observed. Although the lack of microbial growth under these culture conditions may reflect an inability by the indigenous microorganisms to grow on 1,2-DCE as sole carbon substrate, growth also may have been limited by the relatively low substrate concentration. For this study, an initial 1,2-DCE concentration of about 50 µM was selected specifically to assess substrate utilization at environmentally relevant 1,2DCE concentrations. Successful growth of microorganisms, however, generally requires millimolar or greater carbon substrate concentrations (15). For example, microbial growth on the only chloroethene compound presently known to serve as a primary carbon substrate for microbial growth was demonstrated on 5% (v/v) vinyl chloride under aerobic conditions (16, 17). Alternatively, growth in this study may have been limited by the absence of a required growth factor. Specific growth factor requirements have been described previously for chloroethene-degrading microorganisms (18). Further investigation is required to assess the ability of microorganisms to grow on 1,2-DCE as sole carbon substrate. The results of this study have important implications for the natural attenuation of DCE in contaminated environments. Although aerobic cometabolism of DCE is wellestablished (1, 5, 7-11), this process is probably restricted to the fringe of contaminant plumes (19, 20) because readily oxidizable substrates and oxygen rarely co-occur within the core of mature contaminant plumes. However, the results of this study demonstrate that microorganisms can also utilize DCE as a sole carbon substrate to support metabolism. This suggests that DCE can be degraded as a primary substrate in microbial metabolism and that this process may contribute to the natural attenuation of DCE even under circumstances where aerobic cometabolism is not favored.
Acknowledgments We thank Mike Maughon of the Naval Facilities Engineering Command for assistance in collecting sediments and providing background information for the site. This research was supported by the U.S. Geological Survey Toxic Substances Hydrology Program and the Southern Division Naval Facilities Engineering Command.
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(5) Hopkins, G. D.; McCarty, P. L. Environ. Sci. Technol. 1995, 29, 1628-1637. (6) Klier, N. J.; West, R. J.; Donberg, P. A. Chemosphere 1999, 38, 1175-1188. (7) Malachowsky, K. J.; Phelps, T. J.; Teboli, A. B.; Minnikin, D. E.; White, D. C. Appl. Environ. Microbiol. 1994, 60, 542-548. (8) Moore, A. T.; Vira, A.; Fogel, S. Environ. Sci. Technol. 1989, 23, 403-406. (9) Semprini, L.; Roberts, P. V.; Hopkins, G. D.; McCarty, P. L. Ground Water 1990, 28, 715-727. (10) Semprini, L.; McCarty, P. L. Ground Water 1991, 29, 365-374. (11) Semprini, L. Environ. Health Perspect. 1995, 103, 101-105. (12) Bradley, P. M.; Chapelle, F. H. Environ. Sci. Technol. 1996, 30, 2084-2086. (13) Davis, J. W.; Carpenter, C. L. Appl. Environ. Microbiol. 1990, 56, 3878-3880. (14) Atlas, R. M. Handbook of Microbiological Media; CRC Press: Boca Raton, FL, 1993; p 49.
(15) Tanner, R. S. Manual of Environmental Microbiology; ASM Press: Washington, DC, 1997; pp 52-60. (16) Hartmans, S.; de Bont, J. A. M.; Tramper, J.; Luyben, K. C. A. M. Biotechnol. Lett. 1985, 7, 383-388. (17) Hartmans, S.; de Bont, J. A. M. Appl. Environ. Microbiol. 1992, 58, 1220-1226. (18) Maymo-Gatell, X.; Tandoi, V.; Gossett, J. M.; Zinder, S. H. Appl. Environ. Microbiol. 1995, 61, 3928-3933. (19) Anderson, J. E.; McCarty, P. L. Appl. Environ. Microbiol. 1997, 63, 687-693. (20) Dolan, M. E.; McCarty, P. L. Environ. Sci. Technol. 1995, 29, 1892-1897.
Received for review July 14, 1999. Revised manuscript received November 1, 1999. Accepted November 1, 1999. ES990785C
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