Environ. Sci. Technol. 1996, 30, 2412-2416
In Situ Dynamics of Aromatic Hydrocarbons and Bacteria Capable of AH Metabolism in a Coal Tar Waste-Contaminated Field Site E. L. MADSEN,* C. T. THOMAS, M. S. WILSON, R. L. SANDOLI, AND S. E. BILOTTA Section of Microbiology, Wing Hall, Cornell University, Ithaca, New York 14853-8101
Field experiments utilizing randomized block designs were implemented to assess the mobility of both coal tar-derived aromatic hydrocarbons and bacteria capable of metabolizing these substances at a contaminated field site. Arrays of sorbent materials wrapped in fiberglass mesh fabric were inserted into organic matter-rich freshwater sediments in order to intercept mobile chemicals and bacteria carried by the prevailing hydraulic gradient. Polyurethane foam plugs served as a sorbent for aqueous-phase coal tar components while sterile sand from the site served as a substrate for colonization by bacteria. Replicate sorbents were removed from the sediments at varying intervals and assessed for organic compounds (via gas chromatography/mass spectrometry) and for numbers of aromatic hydrocarbon-degrading bacteria (via viable plate counts). Organic contaminants including naphthalene, methyl naphthalene, indenes, and substituted benzenes were detected in the foam sorbents. Contaminant concentrations reached a maximum after 15 days before diminishing. Both naphthalene- and phenanthrene-utilizing bacteria were mobile and reached peak titers of 104 and 103.3, respectively, within 11 days. Thus, comigration of both contaminants and microorganisms occurred at the study site. Furthermore, the in situ abundances of contaminants and microorganisms reflect a dynamic balance between processes causing accrual and elimination.
Introduction The impact of organic environmental contaminant compounds upon potential biological receptors is strongly influenced by contaminant mobility. Contaminant mobility, in turn, is governed by a variety of reaction processes * To whom all correspondence should be addressed: telephone: 607-255-3086; fax: 607-255-3904; e-mail address: ELM3@ CORNELL.EDU.
2412
9
ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 30, NO. 7, 1996
(1, 2) including those that enhance mobility (e.g., dissolution, volatilization, desorption, entrainment in flowing water, colloid-mediated transport) and those that diminish mobility (e.g., precipitation, condensation, sorption, deposition, uptake by biota). Biodegradation of organic compounds is often exceptional among naturally occurring processes that occur in field sites because biodegradation can break intramolecular carbon-carbon bonds, thereby altering the molecular structure and often completely mineralizing organic contaminants to carbon dioxide (3). Contaminant migration is determined by the balance between the variety of simultaneous processes that act in field sites. When biodegradation is a dominant process, contaminant plumes in subsurface habitats have been shown to contract (4-6). However, many studies have shown that the mobility of microorganisms in soils and sedimentary environments is quite limited (7-9). This casts doubts on the probability that microorganisms capable of metabolizing organic contaminants will also migrate. Clearly, comigration of contaminants and contaminantutilizing microorganisms would increase the duration of interactions between cells and contaminants, thereby increasing the likelihood of successful biodegradation. Conversely, successful contaminant biodegradation may be precluded in certain contexts if mobile contaminants migrate more rapidly than the microorganisms responsible for biodegradation. As part of an ongoing series of studies designed to assess the fate of coal tar-derived organic compounds at a field site in New York state (10-13), we have begun to investigate the degree to which organic contaminants and bacteria capable of contaminant metabolism are mobile in the surface sediments of the study site. To pursue these objectives, sorbents were inserted into the organic matterrich sediments of the study site and periodically removed for both chemical and microbiological analyses.
Experimental Section Field Study Site. The coal tar waste-contaminated field site, located in South Glens Falls, NY, has been extensively characterized, hydrologically, chemically, and microbiologically (10). A schematic cross-section of the site is presented in Figure 1. In this investigation, all field procedures and measurements were carried out in the seep area, where contaminated water emerges from an aquifer and enters organic matter-rich sediments at the base of a hill. Groundwater flow velocity has been estimated by hydrologists to be tens of meters per year. Overview of Experimental Design. The sorbents utilized were either clean sterilized polyurethane foam plugs (for contaminant sorption assays) or 2-g portions of sterilized site-derived sand (for microorganism sorption assays). Both sorbent types were enclosed in fiberglass mesh packets with ends secured with copper wire, one end of which carried a flag for retrieval purposes. Sixteen foam sorbents and 20 sand sorbent packets were placed into the sediments by briefly opening and then closing a cavity in the moist mud with a small stainless steel spatula. The sorbents were inserted at a location where water exits the seep area of the study site (Figures 1 and 2) and flows into the nearby surface stream and toward a distant river. Later, the sorbents were periodically retrieved and analyzed.
S0013-936X(96)00035-1 CCC: $12.00
1996 American Chemical Society
recovered from the sediments. To assure accurate contaminant recovery, immediately after removal from sediments the foam sorbents were unwrapped, freed of excess water (by squeezing), and placed into Teflon-sealed 25-mL vials (Ichem; Newcastle, DE), and 6 mL of extractant (1:1, acetone:hexanes) was added. Sorbents were handled with sterile forceps. Following transport back to the laboratory, gas chromatography/mass spectroscopic (GC/MS) analysis was completed (see below).
FIGURE 1. Diagram of study site in vertical cross-section showing source of coal tar contaminants, direction of groundwater flow, and relationship between upland and seep areas of the site. Neither horizontal nor vertical dimensions are to scale.
FIGURE 2. Detailed plan view of seep area of the study site. Bold lines represent free-flowing streamwater. Bold arrows indicate direction of water flow. Lines flanking the stream flow delineate water-saturated organic matter-rich sediments contaminated with coal tar constituents.
Sorbent Preparation. For contaminant mobility, polyurethane foam plugs (2.0 cm in diameter × 3.5 cm in length; Gaymar Industries Inc., Buffalo, NY) were wrapped in fiberglass mesh fabric (used widely in the repair of fiberglass boats) and then autoclaved for 20 min. For colonization of sand by microorganisms, 2 g of site-derived sand was placed and secured in 7.5 × 7.5 cm squares of fiberglass mesh fabric prior to being autoclaved three times for 45 min. Sorbent Deployment, Sampling, and Microbiological Analyses. The area of the seep chosen for sorbent deployment is depicted in Figure 2. Placement positions in 4 × 4 or 4 × 5 arrays were selected in a level portion of the organic matter-rich sediments. Each sorbent was inserted to a depth of 2 cm. Periodically, during the autumn of 1993, three or four replicates of the sorbents were removed from the sediment using a randomized design (14) and subjected to extraction and chemical analysis (foam sorbent) or microbiological analysis (sand sorbent). At some sampling times, only three of the four replicates were
After removal from the field sediment, the sand sorbents were immediately transferred using an aseptic technique to sterile plastic bags and then placed on ice. At the time of sorbent removal, a companion sediment sample from adjacent sediments was also aseptically collected. Dilution and plating of the microorganisms colonizing the sand occurred within 24 h after transport back to the laboratory, using a general heterotrophic medium (5% PTYG; 15) and contaminant-specific media [mineral salts (16) or mineral salts plus naphthalene vapor or overlaid with ether-applied phenanthrene (17)]. For each dilution series prepared from each sample, organisms on duplicate plates were counted. Incubations were aerobic at 22 °C in the dark. In enumerating microorganisms able to grow on aromatic compounds, only large colonies (after 2-3 weeks incubation) occurring in excess of those found in the medium containing mineral salts, alone, were counted. Chemical Analyses. After return from the field site, vials containing extractant and foam were stored at 4 °C until injection of 1 µL of the solvent phase into a Hewlett-Packard 5890 Series II gas chromatograph fitted with a Nukol glass capillary column (30 m × 0.25 mm i.d., 0.25 µm film thickness; Supelco, Bellefonte, PA). The unit was equipped with a 5971A mass selective detector operated in scanning mode at a voltage of 2047, temperature at 300 °C, scanning mass range of 10-180 m/z, an electron volt energy of 70 eV, and an ion source pressure maintained at 1.0 × 10-5 Torr. The injector temperature was 250 °C, and the oven temperature program was 60 °C, 1 min; ramp at 10 °C/min to 200 °C; hold at 200 °C for 10 min. Identification of coal tar-derived analytes was accomplished by injecting authentic standards (1-methylnaphthalene, naphthalene, indene, and 1,2,3-trimethylnaphthalene, obtained from Sigma Chemical Co., St. Louis, MO) whose GC retention times and mass fragmentation patterns matched the corresponding site-derived analytes. Quantification was achieved by preparing triplicate linear calibration curves for each authentic standard and expressing concentrations as ppm in each fully hydrated foam sorbent plug. Authentic standards were not available for all compounds whose fragmentation patterns were matched by the HewlettPackard spectral comparison software to entries in the National Bureau of Standards spectral library. When the fragmentation pattern of coal tar components matched a library entry with a score of 95 or higher, such compounds were tentatively identified. However, without authentic standards, rigorous identification and quantification could not be carried out, and the abundances of these compounds could only be reported in relative terms, as integration units per foam plug. Removal of [14C]Naphthalene from Aqueous Solution. To test the efficacy of aqueous contaminant sorption by polyurethane foam plugs, triplicate sterile 40-mL vials containing 38 mL of sterile naphthalene-saturated water
VOL. 30, NO. 7, 1996 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
9
2413
FIGURE 3. Removal of aqueous [14C]naphthalene from aseptically prepared solutions by polyurethane foam. Vertical axis depicts dpm remaining in an initially saturated (30 ppm) naphthalene solution. Treatments were fiberglass mesh wrapping with and without foam plugs. Each point is the average of three replicates. The range of standard deviations for the various data points was from 0.8 to 10% of each value shown; the average standard deviation was 4.3%. were prepared. To each flask was added 4.5 × 103 dpm of aqueous phase [1-14C]naphthalene (10.1 mCi/mmol, >98% radiopurity; Sigma Radiochemicals) and a single sterile foam plug (wrapped in fiberglass mesh, as described above). Vials omitting the foam plugs but including the fiberglass mesh and copper wire wrapping were also prepared in triplicate. Periodically, 0.5 mL of fluid was removed from each flask and mixed with 4.0 mL of Ecoscint scintillation cocktail (National Diagnostics, Manville, NJ), and 14C activity was quantified in a scintillation counter (Model 5000CE, Beckman Instruments, Inc., Fullerton, CA). Depletion of 14C from the aqueous phase in excess of the controls was interpreted as naphthalene sorption by the foam plugs.
Results Prior to deploying the clean, sterile foam sorbents in the field site sediments, we tested the affinity of the polyurethane foam sorbent for naphthalene, a contaminant previously found at the site. When suspended in aqueous-phase naphthalene, the foam sorbent effectively reduced the concentration from 30 to less than 1 ppm within approximately 100 h (Figure 3). Note that the naphthalene concentration diminished somewhat even in the treatment to which no sorbent was added (Figure 3). This was probably due to volatilization and/or sorption to the fiberglass mesh. However, the difference in the rate and extent of [14C]naphthalene recovery from the two treatments shown in Figure 3 can be attributed to uptake by the foam sorbent. A typical chromatogram of site contaminants recovered from the foam sorbents is depicted in Figure 4A. Data shown in this figure demonstrate the complexity of the chemical components present in the site. From the numerous compounds detected by the GC/MS, we focused on seven compounds for inclusion in this study because of their relatively high abundances and consistent occurrence in the chromatograms. Four of these (1-methylnaphthalene, 1,2,3-trimethylbenzene, indene, and naphthalene) were quantified because authentic standards were available. An example of how the unknown contaminant, 1,2,3-trimethylbenzene, was identified by matching its fragmentation pattern to an authentic standard is shown in Figure 4B,C. After GC retention times were also found to match, standard curves for quantification were prepared (data not shown). Authentic standards for three other contaminants (1-methyl-2-cyclopropen-l-ylbenzene, 1-ethyl3-methylbenzene, and 2,3-dihydro-4-methyl-1H-indene) were not available from chemical manufacturers; however,
2414
9
ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 30, NO. 7, 1996
FIGURE 4. Chromatogram of contaminants from the field site (A) and an example of the mass fragmentation pattern of a site contaminant (B), later identified as 1,2,3-trimethylbenzene after matching an authentic standard (C).
these compounds appeared consistently throughout our sampling period and received a very high score (>95) in the mass spectrum comparison program that matched mass fragmentation patterns between unknown compounds and those in the library database. Figure 5 displays the results of chemical analyses performed on the sediment-deployed foam sorbents. The initially clean polyurethane foam plugs were immersed in saturated sediments. Then after periods ranging from 5 min to 23 days, sorbents were removed from the site and extracted, and GC/MS analyses were performed. Processes reflected in the contaminant retrieval data in Figure 5 include sorption/desorption reactions, aqueous-phase transport, filtration of colloids into the foam sorbent, and microbial metabolism of the compounds occurring in the foam. All compounds were initially at very low concentrations. After initial insertion into the site sediments, the seven compounds did not simply accrue in the foam sorbents. Instead, a maximum concentration was found on day 15 followed by a slow decline in concentration by day 23. The reason for the decline in concentration was uncertain. However, a shift in the relative rates of loss versus accrual mechanisms must have been responsible and may have involved leaching of the materials after rainfall events and/or microbial metabolism. Inspection of local weather records revealed that 7.5 cm of rain fell during the sampling period.
FIGURE 5. Concentrations of coal tar-derived compounds retrieved from polyurethane foam sorbents inserted into the seep study area. Compounds were analyzed by GC/MS. Authentic standards were available for the identified and quantified compounds shown in panel A. Authentic standards were unavailable for the compounds shown in panel B; hence, identification is tentative, and absolute concentrations could not be ascertained. Each point represents the average of three or four replicate samples. The range of standard deviations for the various data points was from 1.2 to 98% of each value; the average standard deviation was 19%.
The results of the bacterial mobility experiments are presented in Figure 6. This displays bacterial numbers recovered from the previously sterilized sorbent sand as well as bacterial abundances in sediment adjacent to the sorbent array and collected simultaneously. To assess colonization on day 0, initially sterile sand was immersed in the water-saturated sediment for a 1-min period. As shown in Figure 6A,B, the number of naphthalene and phenanthrene-utilizing bacteria were initially low and reached peak titers of 104 and 103.3 g sediment-1, respectively, within 11 days of sorbent insertion into the sediment. General heterotrophic bacteria (grown on 5% PTYG medium; Figure 6C) recovered from the sand sorbent also seemed to increase with time. As expected, general heterotrophs were several orders of magnitude more abundant than microorganisms able to grow on aromatic compounds. The microbial abundance data revealed that heterotrophic microorganisms, including contaminantmetabolizing bacteria, were present and mobile in the water and sediment at the field site. The microbial colonization experiment was designed to assess dynamic changes in components of the microbial community. These changes occurred via rapid equilibration with the sediment, by the transport of microorganisms into the sand sorbent, and/or by the subsequent growth or release of the microorganisms. The sorbent deployment methodology alone could not easily distinguish these mechanisms of change from one another. However, the degree to which population shifts on the initially sterile sorbent in Figure 6A-C were caused simply by instantaneous equilibration with the surrounding sediment versus growth and accrual can be assessed by comparing the
FIGURE 6. Numbers of naphthalene- (A), phenanthrene- (B) , and PTYG- (C) grown bacteria retrieved at various times from initially sterile sand sorbents inserted into the seep study area. Each group of bacteria was enumerated after aseptic serial dilution and plating onto agar media containing each respective carbon source. Each data point represents bacterial numbers recovered from the packets of sand sorbent (three or four replicate samples) and on bulk sediment (three replicates on day 0, single unreplicated samples for remaining days). The range of standard deviations for the replicated samples was from 16 to 150% of each value; the average standard deviation was 60%. number of bacteria retrieved from the sterile sorbent to the number of bacteria retrieved from the independently gathered adjacent field sediments at each sampling time. Panels A and C of Figure 6 show that the initial numbers of bacteria in field sediment samples that grew on naphthalene and PTYG media were either 2.5 or 1.6 log units higher than numbers retrieved from the briefly immersed sand sorbents. As determined by Student’s t-test, these differences were significant at P values of 0.012 and 0.23, respectively; then, over the next two sampling periods, the numbers increased asymptotically in the sand sorbents toward those in the surrounding sediment. Thus, for naphthalene-metabolizing (Figure 6A) and general heterotrophic bacteria (Figure 6C), the data suggest that gradual colonization and/or bacterial growth occurred on the initially sterile sorbents. By contrast, for phenanthrenemetabolizing bacteria (whose numbers were 1 and 3 orders of magnitude lower than those found on naphthalene and PTYG media, respectively), there was no distinguishable difference between bacterial numbers in field sediment versus those retrieved from the sand sorbent (Figure 6B). Thus, the data shown in Figure 6B do not argue for dynamic successional population shifts of phenanthrene-degrading bacteria within the initially sterile sorbents. Instead of accrual and growth of phenanthrene-utilizing bacteria on the sorbent, the population shifts appear to simply reflect an instantaneous equilibration with the ambient water and sediment.
Discussion Passive sampling strategies have been long established in microbial ecology in efforts to assess in situ colonization
VOL. 30, NO. 7, 1996 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
9
2415
of microorganisms for a variety of habitats. For instance, the Rossi slide technique [described by Parkinson et al. (18)] involves insertion of an initially microorganism-free glass slide into soil. Subsequent removal and microscopic examination of the glass slide can provide insights into the propensity of soil microorganisms to adhere and grow under in situ conditions. More recently, the same principles have been applied to investigating groundwater microorganisms (19). Hirsch and Rades-Rohkohl (19) inserted an initially sterile column of aquifer sand into well water and subsequently characterized microorganisms that attached and grew on the sand over a 12-week period. Implicit in studies investigating microbial colonization is the choice of a sterile substrate, whose surface properties will inevitably influence which microorganisms are retrieved. Rather than selecting planar glass surfaces (i.e., slides) or some other arbitrary substrate for deployment and retrieval from the seep area of our study site, we used the sandy subsoil native to the site because of its ecological relevance. Nonetheless, it must be recognized that this sand almost certainly had its surface properties altered by the sterilization process (autoclaving). Thus, devising colonization substrates that avoid sampling biases remains a challenge. Biases must also be recognized when devising passive in situ sampling schemes for organic contaminant compounds. In the present study, polyurethane foam plugs were selected because of their low cost and convenient size and because precedent had been set for their use in environmental sampling (20). Furthermore, we verified that the foam plugs effectively scavenged a model analyte (naphthalene) from aqueous solution (Figure 3). However, the scavenging selectivity, efficiency, and capacity of the foam plugs were not fully characterized for all analytes described in this study. When Prest et al. (21) compared a suite of organic contaminants sequestered from San Joaquin River water by both semipermeable membrane devices and clams (the latter used as a sentinel species), the contaminants retrieved matched qualitatively but not quantitatively. Despite inevitable biases implicit in the design of sampling devices (e.g., materials, geometry, equilibration times), deployment of passive samplers thought to integrate contaminant fluxes over time (21-23) are gaining increased attention and use. The present study was designed to build upon existing passive sampling procedures in two ways. First, both organic contaminants and microorganisms capable of metabolizing related contaminants were measured. Second, a randomized block design allowed the same sampling locations to be repeatedly assayed so as to document the dynamic changes in contaminant and microbial abundances. Resulting data showed that (i) aromatic contaminants were consistently retrieved by foam sorbents inserted into the seep area of a coal tar-contaminated field study site; (ii) concentrations of aromatic contaminants increased during a 15-day period and then gradually declined; and (iii) contaminant-metabolizing heterotrophic microorganisms indigenous to the field study site were mobile, capable of colonizing initially sterile sand, and increased in abundance over time. The reasons for dynamic shifts in the abundances of both contaminants and microorganisms are uncertain but necessarily reflect a balance of mechanisms that add (e.g., via transport, sorption, cell growth) and remove (e.g., via desorption, leaching, biodegradation, cell death) the chemicals and microorganisms. The impetus for inserting sorbents for organic contaminants and mi-
2416
9
ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 30, NO. 7, 1996
crobial cells into the field site sediments was to ascertain how dynamic these two key components were at the study site. The fact that both contaminants and contaminantmetabolizing microorganisms were mobile strongly suggest that co-transport is occurring at the site. This bodes well for continued attenuation of contaminants, should they be mobilized toward potential off-site receptors.
Acknowledgments This research was supported by a grant from the Air Force Office of Scientific Research (Grants AOFSR-91-0436, 93NL-073, and F49620-95-1-0346). Expert manuscript preparation by P. Lisk is gratefully acknowledged. The authors also thank Niagara Mohawk Power Corporation and E. Neuhauser for access to the study site.
Literature Cited (1) Hemond, H. F.; Fechner, E. J. Chemical Fate and Transport in the Environment; Academic Press: San Diego, CA, 1994. (2) Thibodeaux, L. J. Environmental Chemodynamics, 2nd ed.; John Wiley & Sons: New York, 1995. (3) Madsen, E. L. Environ. Sci. Technol. 1991, 25, 1662-1673. (4) Chiang, C. Y.; Salanitro, J. P.; Chai, E. Y.; Colthart, J. D.; Klein, C. L. Ground Water 1989, 27, 823-834. (5) Hinchee, R. E., Wilson, J. T., Downey, D. C., Ed. Intrinsic Bioremediation; Battelle Press: Columbus, OH, 1995. (6) Wilson, B. H.; Wilson, J. T.; Kampbell, D. H.; Bledsoe, B. E.; Armstrong, J. M. Geomicrobiol. J. 1991, 8, 225-240. (7) Gannon, J.; Tan, Y.; Baveye, P.; Alexander, M. Appl. Environ. Microbiol. 1991, 57, 2497-2501. (8) Harvey, R. W.; Kinner, N. E.; Bunn, A.; MacDonald, D.; Metge, D. Appl. Environ. Microbiol. 1995, 61, 209-217. (9) Scholl, M. A.; Harvey, R. W. Environ. Sci. Technol 1992, 26, 14101416. (10) Madsen, E. L.; Sinclair, J. L.; Ghiorse, W. C. Science 1991, 252, 830-833. (11) Herrick, J. B.; Madsen, E. L.; Batt, C. A.; Ghiorse, W. C. Appl. Environ. Microbiol. 1993, 59, 687-694. (12) Wilson, M. S.; Madsen, E. L. Environ. Sci. Technol. 1996, 30, 2099-2103. (13) Madsen, E. L.; Bilotta, S. E. Environ. Toxicol. Chem., in press. (14) Sokal, R. R.; Rohlf, F. J. Biometry, 2nd ed.; W. H. Freeman and Co.: New York, 1981. (15) Balkwill, D. L.; Ghiorse, W. C. Appl. Environ. Microbiol. 1985, 50, 580-588. (16) Stanier, R. Y.; Palleroni, N. J; Doudoroff, M. J. Gen. Microbiol. 1966, 43, 159-271. (17) Kiyohara, H.; Nagao, K.; Yana, K. Appl. Environ. Microbiol. 1982, 43, 454-457. (18) Parkinson, D.; Gray, T. R. G.; Williams, S. T. Methods for Studying the Ecology of Soil Micro-organisms; Blackwell Scientific Publishers: Oxford, England, 1971. (19) Hirsch, P.; Rades-Rohkohl, E. Appl. Environ. Microbiol. 1990, 56, 2963-2966. (20) Rappe, C.; Kjeller, L.-O.; Andersson, R. Chemosphere 1989, 19, 13-20. (21) Prest, H. F.; Jarman, W. M.; Burns, S. A.; Weismu ¨ ller, T.; Martin, M.; Huckins, J. N. Chemosphere 1992, 25, 1811-1823. (22) Huckins, J. N.; Manuweera, G. K.; Petty, J. D.; Mackay, D.; Lebo, J. A. Environ. Sci. Technol. 1993, 27, 2489-2496. (23) Lebo, J. A.; Gale, R. W.; Petty, J. D.; Tillitt, D. E.; Huckins, J. N.; Meadows, J. C.; Orazio, C. E.; Echols, K. R.; Schroeder, D. J.; Inmon. L. E. Environ. Sci. Technol. 1995, 29, 2886-2892.
Received for review January 17, 1996. Revised manuscript received March 14, 1996. Accepted March 21, 1996.X ES960035S X
Abstract published in Advance ACS Abstracts, May 15, 1996.
