Environ. Sci. Technol. 1999, 33, 4346-4351
Chemical-Extraction Methods To Estimate Bioavailability of DDT, DDE, and DDD in Soil JIXIN TANG, BOAKAI K. ROBERTSON, AND MARTIN ALEXANDER* Institute for Comparative and Environmental Toxicology and Department of Soil, Crop, and Atmospheric Sciences, Cornell University, Ithaca, New York 14853
A study was conducted to find a method to determine the bioavailability to earthworms of DDT, DDE, and DDD in soils. Measurements were made of the uptake by Eisenia foetida of the three compounds in seven soils in which they were freshly added or had persisted in the field for up to 49 yr. The worms assimilated 3.15-66.2% of the compounds in the test samples. Different amounts of the three compounds were sorbed by C18 membrane disks placed in suspensions of these soils, and different amounts also were extracted by solutions of tetrahydrofuran in water. The amounts of the compounds taken up by the C18 membrane disks and extracted by the tetrahydrofuran-water mixture correlated well with uptake by E. foetida from 10 samples of these soils. The correlation coefficients (r) were 0.921 or higher for the C18 membrane disk assay and 0.8310.948 for the tetrahydrofuran-extraction assay in correlations of the chemical and biological assays. These methods thus offer promise as chemical assays to predict bioavailability of DDT and related compounds.
Introduction Assessment of exposure of living organisms to toxic chemicals in soil requires information on the concentration that is available to those species. The current approach to exposure assessment commonly relies not on the level that is biologically available, however, but rather on the total concentration as determined by vigorous extraction. Nevertheless, considerable evidence exists that the amount that is available to mammals (1, 2), invertebrates (3, 4), plants (5, 6), and microorganisms (7, 8) is less, sometimes appreciably less, than that anticipated by analytical procedures based on vigorous extractions. Assessing exposure based on such procedures is made even more difficult because seemingly small differences in soil properties may markedly change the availability of the same quantity of an organic compound for acute toxicity to animals (3) and plants (5) and for genotoxicity (9). In addition, even at a constant total concentration, the level that is biologically available may decline with time because the compound becomes sequestered as it ages in soil (10, 11). As a result, attempts have been made to devise chemical assays that reflect bioavailability. For example, analysis of the water in soil pores has been proposed as a means of predicting the bioavailability of pesticides (12, 13), and * Corresponding author phone: (607)255-1717; fax: (607)255-2644; e-mail:
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equilibrium partitioning has been suggested as a way of measuring the bioaccumulation of polycyclic aromatic hydrocarbons by earthworms (14). An approach based on nonexhaustive extraction with organic solvents showed that the decline in extractability of phenanthrene and atrazine was parallel to the decline in their availability to earthworms and bacteria (15). Other methods have been proposed for organic compounds, but usually the results have not been correlated with the data obtained by bioassays, and such correlations are essential for any procedure to be ultimately accepted for exposure assessments. We have recently found that the quantities of three aged and unaged polycyclic aromatic hydrocarbons taken up by earthworms were correlated with the amounts recovered from soil by a mild extraction with organic solvents (16). The study reported here was designed to assess the feasibility of two procedures for determining the bioavailability of organic compounds in soil, namely, solid-phase extraction with C18 membrane disks and liquid-phase extraction with an aqueous solution containing tetrahydrofuran. A parallel study was designed to evaluate an approach using Tenax TA beads as a solid-phase extractant (17). The compounds chosen were DDT [1,1,1-trichloro-2,2-bis(pchlorophenyl)ethane], DDE [1,1-dichloro-2,2-bis(p-chlorophenyl)ethane], and DDD [1,1-dichloro-2,2-bis(p-chlorophenyl)ethylene].
Materials and Methods Materials. DDT, DDE, and DDD (all of 99.5% purity) were purchased from ChemService (West Chester, PA). Tetrahydrofuran was obtained from VWR Scientific Products (Bridgeport, NJ). Novo-Clean 25-mm C18 membrane [poly(tetrafluoroethylene) impregnated with a bonded octyl silica sorbent] was purchased from Alltech Associates (Deerfield, IL). All solvents were HPLC grade. Redworms (Eisenia foetida), which were obtained from Carolina Biological Supply (Burlington, NC), were maintained on a commercial worm bedding that was kept moist with deionized water. The worms were fed with a commercial mixture of crude protein and carbohydrate (Magic Products, Amherst Junction, WI). The worms that were used were of almost identical sizes (ca. 0.3 g wet weight), they were mature as determined by the presence of a clitellum, and they were active at the time they were introduced into the soil. Soil samples were collected from experimental plots containing Sassafras silt loam (pH 6.5, 4.4% organic matter) and Chester loam (pH 5.8, 6.5% organic matter) located at Beltsville Agricultural Research Center, U.S. Department of Agriculture, Beltsville, MD. The plots were treated with DDT in 1949 (18). Samples of a sandy loam (pH 6.6, 6.0% organic matter) contaminated approximately 30 yr ago with DDT were obtained from a remediation site at the U.S. Navy Surface Weapons Testing Center in Dahlgren, VA. Soils not previously treated with insecticides [Kendaia loam (pH 6.6, 12.6% organic matter), Collamer silt loam (pH 6.8, 3.1% organic matter), Hudson silt loam (pH 6.8, 1.6% organic matter), Lima loam no. 1 (pH 7.9, 2.6% organic matter), and Lima loam no. 2 from a forested location (pH 7.1, 11.4% organic matter)] were from New York. All soils were passed through a 2-mm mesh sieve and mixed well. The soil samples from New York were sterilized with 2.5 Mrad of γ-irradiation from a 60Co source. No growth was observed when samples of the irradiated soils were placed on Trypticase-soy agar. Laboratory Aging. Various concentrations of DDT, DDE, and DDD dissolved in 15 mL of dichloromethane were added to the sterile soils. The soil was mixed thoroughly, the solvent 10.1021/es990581w CCC: $18.00
1999 American Chemical Society Published on Web 10/12/1999
TABLE 1. Concentration of DDT, DDD, and DDE in Test Soils concentration (µg/g)
a
soil
aging time
DDTa
DDEa
DDDa
Sassafras silt loam Chester loam Dahlgren sandy loam Collamer silt loam Lima loam no. 1 Lima loam no. 2 Hudson silt loam Hudson silt loam Kendaia loam Kendaia loam
49 yr 49 yr ca. 30 yr 924 day 186 day 0 day 924 day 0 day 186 day 0 day
3.12 ( 0.21 12.2 ( 1.6 58.7 ( 4.7 194 ( 13 22.9 ( 3.2 209 ( 17 78.5 ( 6.3 155 ( 13 197 ( 17 248 ( 17
3.09 ( 0.14 4.54 ( 0.32 49.3 ( 4.0 6.07 ( 0.45 0.58 ( 0.04 1.96 ( 0.11 3.12 ( 0.20 1.50 ( 0.08 4.03 ( 0.28 2.12 ( 0.13
1.14 ( 0.11 3.89 ( 0.23 60.3 ( 5.5 3.80 ( 0.32 0.65 ( 0.08 3.30 ( 0.35 1.12 ( 0.16 2.40 ( 0.18 19.7 ( 2.3 3.58 ( 0.31
Mean ( standard error.
was allowed to volatilize for 48 h, and autoclaved water was added to bring the moisture level of the soil to 80% of field capacity at one-third bar. The soil was mixed with a sterilized spatula and transferred to sterile 50-mL test tubes or 125-mL flasks sealed with sterile screw caps lined with silicone-backed Teflon liners. The tubes or flasks were incubated in the dark (22 ( 2 °C) for varying periods of time. Bioavailability. To measure bioavailability, triplicate groups of eight worms were placed in 50-mL test tubes having soil (at a moisture content of 80% of field capacity) containing DDT, DDE, and DDD. The dry weight of the soil was 10 g. The bottoms of the tubes were wrapped with aluminum foil, and the tubes were kept at 22 ( 2 °C in the dark. Ten days later, the worms were removed from the tubes, rinsed with deionized water, and placed on moist filter paper for 24 h to allow for depuration. The worms were washed with water, dried with paper towels, weighed, and then frozen at -10 °C. The worms were ground with 25-30 g of anhydrous Na2SO4 with a mortar and pestle prior to Soxhlet extraction. Soxhlet Extraction. Triplicate soil samples (1.0 g dry weight) or ground worm tissues were transferred to cellulose extraction thimbles, and the materials were subjected to Soxhlet extraction for 10-12 h with 120 mL of hexane at a rate of 5-6 min/cycle. The extract was concentrated to near dryness under vacuum, the resulting material was dissolved in 5 or 10 mL of hexane, and the solution was passed through a 0.22-µm Teflon filter prior to gas chromatographic analysis. To measure recoveries, 10 µL of hexane was added to 10 g of Kendaia loam. The recoveries of DDT, DDE, and DDD added to soil were 95.8 ( 8.5, 93.6 ( 7.7, and 99.2 ( 7.5%, respectively. Solid-Phase Extraction. A 1-g (dry weight) soil sample was added with 50 or 100 mL of deionized water to a 125or 250-mL flask containing a C18 membrane disk, which had been previously conditioned by rinsing once with 20 mL of methanol for 5 min and twice with 400 mL of deionized water for 2 min. To prevent biodegradation, 1.0 mL of a 0.2% aqueous HgCl2 solution was added to each flask. The flasks were shaken for different time periods, and the disks were rinsed twice by shaking them with 400 mL of water to dislodge soil particles. The disks were dried on paper towels and transferred to 20-mL vials. Hexane (2 or 5 mL) was added to elute DDT, DDE, and DDD from the membrane. To determine the recovery of DDT from water by C18 membrane disks, 20 ng of DDT in 20 µL of hexane was added to 250-mL flasks, and the solvent was allowed to evaporate in the hood. Deionized water (100 mL) and a conditioned C18 membrane disk were added to the flask, which was then placed on a rotary shaker operating at 150 rpm for 24 or 96 h. The DDT sorbed to the membrane was eluted and analyzed. The recovery of 20 ng of DDT in aqueous solution by the C18 membrane was 90.3-98.5%.
Extraction with Cosolvents. Triplicate 1-g (dry weight) soil samples were placed in 50-mL Teflon centrifuge tubes, and 20 mL of 25% tetrahydrofuran in water was added to each tube. The tubes were shaken for 24 h on a reciprocal shaker. The slurry was centrifuged at 15000g for 20 min. The supernatant was added to a 50-mL glass test tube, and the compounds were recovered with a Novo-Clean C18 extraction membrane by a modification of EPA Method 525.2 (19). The C18 membrane was placed in a 25-mm Kontes filtration apparatus (VWR Scientific Products, Bridgeport, CN), and 5 mL of methanol was added to the solvent reservoir of the filtration apparatus. After 3 min, a vacuum was applied to allow most of the solvent to pass through the membrane, leaving about 5-mm depth of solvent above the membrane, and 5 mL of methyl tert-butyl ether (MTBE) was added to the solvent reservoir. The liquid was passed through the membrane, which was then dried under vacuum. The membrane was conditioned by first adding 10 mL of methanol, which was then pulled through by use of a vacuum, and the procedure was repeated with 15 mL of deionized water. The supernatant from the extract was added and then pulled through the membrane by vacuum in 5-10 min until the membrane became dry. During the conditioning and extraction, the disk was not allowed to dry out until the sample had been drawn through. The sorbed chemicals were eluted from the membrane by adding 5 mL of methanol, which was allowed to soak the membrane for 3 min, and it was then drawn through. Then, 5 mL of MTBE was allowed to soak into the membrane for 3 min and drawn through, followed by 10 mL of hexane. The methanol, MTBE, and hexane were combined and concentrated by evaporation to near dryness, and 5 or 10 mL of hexane was added prior to analysis. Chromatographic Analysis. The concentrations of DDT, DDE, and DDD were determined with a gas chromatograph (model 5890, Hewlett-Packard, Rochester, NY) equipped with an electron-capture detector. The analysis was performed essentially by EPA Method 8080 (20) with an HP-608 column (30 m, 0.53-mm i.d., 0.5 µm film thickness); a flow rate of N2 of 10.0 mL/min; and oven, injector, and detector temperatures of 225, 275, and 275 °C, respectively. The retention times of DDT, DDE, and DDD were 4.7, 2.8, and 4.0 min, respectively. Data Analysis. The relationship between assimilation by the worms and chemical extraction was determined by linearregression analysis. An analysis of variance of each regression was used to determine the statistical significance of the correlations.
Results Analyses were conducted to determine the concentrations of the compounds of interest in the test soils. The concentrations of DDT, DDE, and DDD were 3.12-209, 0.58-49.3, VOL. 33, NO. 23, 1999 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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FIGURE 1. Effect of time of exposure on the uptake from soil of DDT, DDD, and DDE by earthworms. and 0.65-60.3 µg/g, respectively (Table 1). These values are for soils in which the compounds had not been aged or had been aged for up to 49 yr. Measurements were made of the effect of time of exposure of the redworms to soil containing DDT, DDE, and DDD on the uptake of these compounds. For this purpose, the worms were introduced into Kendaia loam in which the compounds (200 µg/g) had been aged for 186 days. For the first few days, the amounts of the three compounds that were assimilated increased (Figure 1). After 14 days, the concentration of DDT in the worms fell and the level of DDD increased, suggesting that the DDT was being transformed to DDD. Because the uptake of the total of the three compounds reached a maximum at about 10 days, that time period was selected for use in subsequent bioavailability measurements. Determinations were made of the amounts of the three compounds assimilated by the worms using the same assay conditions for each of the soils. The concentrations varied widely among the 10 soil samples (Table 2). This is not unexpected in view of the dissimilar concentrations in the soil, the differing periods of aging of the compounds, and the different soil properties. More DDT, DDE, and DDD were assimilated by the worms from soils containing the unaged than the aged compounds, from soils with lower contents of organic matter, and from soils containing higher chemical concentrations. DDT, DDE, and DDD were not detected in control worms. The percentage of the compounds assimilated by the worms from the soils varied greatly. Because the aim of this investigation was to develop a chemical assay, no attempt was made to relate the amounts or uptake percentages to concentrations in the soil, soil properties, or aging time. We have previously investigated the effects of soil properties and aging time on uptake of two compounds by earthworms (11) and of aging time on DDT effects on insects (10). Several experimental variables were evaluated to develop the assay. To test the effect of water:soil ratio on DDT sorption by the C18 membrane, 25, 50, 100, 150, or 200 mLof deionized water was added to a 250-mL flask containing 1.0 g of Kendaia soil in which DDT (200 µg/g) had aged for 178 days. Each flask contained a C18 membrane disk. The sorption of DDT by the C18 membrane decreased linearly based on sorbed amount and percentage when the water:soil ratio increased from 25:1 to 200:1 (v/w). Water:soil ratios of 50:1 and 100:1 were chosen for solid-phase extraction. The rate of sorption by the C18 membrane disks and the rate of extraction of DDT from soil were determined. A 1-g sample of Kendaia soil in which DDT had aged for 186 days was added to 18 250-mL flasks each of which contained 100 4348
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mL of deionized water and a conditioned C18 membrane. The flasks were shaken for different periods of time, and the C18 membrane was removed at 6, 12, 24, 48, 120, and 216 h. The disks were rinsed with water and dried, and the amount of DDT was determined. The amount of DDT sorbed by the membrane increased linearly from 1.82 to 40.5 µg/membrane as the time increased from 6 to 216 h. After 216 h, 32.3% of DDT in the soil was sorbed by the membrane. The quantities of the three compounds extracted in 24 h from the 10 soils by use of the C18 membrane disks and 50 mL of water were measured. From 0.54 to 15.1% of the DDT, 2.59-26.4% of the DDE, and 2.18-28.0% of the DDD were extracted from the 10 soils under identical conditions. When the contact time was increased to 72 h and 100 mL of water was used, the trends among the soils were similar to that observed in 24 h with 50 mL of water; 1.12-24.0% of the DDT, 5.36-38.1% of the DDE, and 4.45-40.6% of the DDD were extracted. The effectiveness of 20% aqueous solutions of several organic solvents in extracting DDT from soil was assessed. For this purpose, 0.5 g of Kendaia loam containing DDT aged for 163 days was added to 50-mL Teflon centrifuge tubes containing 20 mL of 20% aqueous solutions of acetone, ethanol, methanol, propanol, or tetrahydrofuran (THF). The tubes were placed on a reciprocal shaker for 24 h and then centrifuged at 15000g, and DDT in the supernatant was removed by C18 membranes. The percentages of the DDT extracted by 20% aqueous solutions of acetone, ethanol, methanol, propanol, and THF were 3.47 ( 0.31, 3.04 ( 0.36, 0.93 ( 0.15, 1.43 ( 0.33, and 21.5 ( 2.4%, respectively. Because of the higher recoveries, THF was selected for further testing. The recoveries of DDT by the C18 membrane were determined using different percentages of THF in aqueous solution. For this purpose, a solution (0.3 mL) of DDT in hexane (10 µg/mL) was added to 50-mL Teflon centrifuge tubes, the hexane was allowed to evaporate, and then 20 mL of aqueous solution containing different amounts of THF was added. The tubes were shaken for 10 s on a vortex mixer. The DDT in solution was removed by use of C18 membranes. The recoveries of DDT were 10.1, 44.6, 70.4, 91.3, 84.8, and 25.5% at 5, 10, 20, 25, 30, and 40% THF, respectively. Therefore, a 25% THF aqueous solution was chosen for further investigation. The percentages of the three compounds in the 10 soils that were removed by extraction with a THF-water solution (25:75; v/v) differed markedly among the soils. Using this extraction procedure, 7.31-42.8% of the DDT, 16.2-52.7% of the DDE, and 12.3-51.6% of the DDD were brought into solution. In view of the greatly different amounts and percentages of the three compounds assimilated by the earthworms from these soils and the markedly different amounts extracted from them by the C18 membrane and THF-water solution, it was concluded that the soil samples were adequate to test the possible utility of the chemical procedures for predicting bioavailability. The correlations between the chemical assays and earthworm uptake were assessed by linear-regression analysis. The quantity of the three compounds and the sum of the three amounts extracted in 24 h from 1.0 g of soil by the C18 membrane with 50 mL of water were well-correlated with earthworm uptake (Figure 2). The correlation coefficients (r) were 0.967, 0.984, 0.940, and 0.947 for DDT, DDE, DDD, and total of the three, respectively. The corresponding equations for the regression lines are y ) 10.69x - 1.01, y ) 6.54x + 1.59, y ) 10.81x + 1.13, and y ) 10.83x - 0.63. In instances in which 10 points are not evident in a panel in the figure, the size of the symbol obscured separate points. When the C18 membrane was used with 50 mL of water to extract 1.0 g of soil in 120 h, good correlations were also
TABLE 2. Uptake of DDT, DDD, and DDE by Eisenia foetida from Different Soils uptake (%)a
uptake (µg/g) soil
DDT
DDE
DDD
DDT
DDE
DDD
Sassafras silt loam Chester loam Dahlgren sandy loam Collamer silt loam Lima loam no. 1 Lima loam no. 2 Hudson silt loam Hudson silt loam Kendaia loam Kendaia loam
3.36 ( 0.42 1.42 ( 0.08 7.81 ( 1.02 73.7 ( 6.5 19.3 ( 1.4 140 ( 15 42.5 ( 3.5 111 ( 9 67.6 ( 5.4 99.6 ( 7.6
2.94 ( 0.21 2.89 ( 0.32 12.2 ( 1.4 6.21 ( 0.72 1.49 ( 0.09 3.25 ( 0.28 3.04 ( 0.25 3.22 ( 0.24 2.62 ( 0.18 2.88 ( 0.22
1.38 ( 0.12 0.90 ( 0.07 12.5 ( 1.6 4.95 ( 0.39 1.87 ( 0.13 7.13 ( 1.01 1.34 ( 0.09 4.99 ( 0.36 11.0 ( 1.1 5.57 ( 0.48
27.8 ( 3.4 3.53 ( 0.26 3.15 ( 0.34 9.47 ( 1.28 19.4 ( 1.6 17.7 ( 1.8 13.5 ( 1.5 17.6 ( 1.6 10.8 ( 1.1 13.6 ( 1.4
24.6 ( 2.7 18.4 ( 1.8 5.06 ( 0.61 25.4 ( 2.2 60.3 ( 3.4 43.8 ( 5.1 24.3 ( 2.1 54.8 ( 4.8 20.5 ( 2.1 45.8 ( 3.4
30.4 ( 3.1 6.68 ( 0.48 4.82 ( 0.36 32.3 ( 3.0 66.2 ( 5.5 56.7 ( 6.2 27.7 ( 2.4 53.6 ( 5.1 17.4 ( 1.4 53.0 ( 4.5
a
Percentage of the compound in soil that was assimilated by the earthworms.
corresponding equations for the regression lines are y ) 6.46x - 1.23, y ) 4.20x + 1.62, y ) 4.40x + 1.94, and y ) 6.82x 8.55. When the assay was based on extraction of 1.0 g of soil with THF-water (25:75, v/v), the chemical assay was wellcorrelated with uptake by the earthworms (Figure 3). The r values were 0.918, 0.973, 0.831, and 0.880 for DDT, DDE, DDD, and total of the three, respectively. The corresponding equations for the regression lines are y ) 2.77x + 0.16, y ) 2.20x + 1.72, y ) 2.11x + 2.42, and y ) 2.82x + 1.24.
Discussion
FIGURE 2. Correlation between uptake by earthworms of DDT, DDE, DDD, and total of the three compounds with sorption by C18 membrane. The extraction time was 24 h, and 1.0 g of soil and 50 mL of water were used. obtained. The r values were 0.968, 0.984, 0.921, and 0.931 for DDT, DDE, DDD, and total of the three, respectively. The
The two procedures described here and the method reported in a parallel study from this laboratory (17) offer considerable promise as means for determining the bioavailability of DDT and related compounds. The results of each of the chemical assays were highly correlated with earthworm uptake of the three test compounds present at different concentrations in dissimilar soils and following different periods of aging. Nevertheless, additional study is required before a final approach is established, particularly in instances where earthworm uptake may not parallel bioavailability because of the effects of other toxicants, poor soil structure, or low organic matter levels on their behavior. Moreover, the choice of earthworms as an appropriate indicator for bioavailability to humans or other ecological receptors needs to be clarified further. The procedures commonly used for the analysis of toxicants in soils and sediments not only are designed to give data on the total concentration but also entail the use of reasonably large volumes of organic solvents. In contrast, the methods described here only use small quantities of organic solvents. Solid-phase extractants have been used previously for analysis of organic compounds in soils, but those studies have focused on the total quantity rather than the bioavailable fraction (21-23). Although the amounts of the compounds extracted by the two procedures were correlated with the quantities assimilated by the earthworms, additional investigation is warranted to fully delineate an acceptable procedure. Information is needed on the number of worms and their period of exposure, the optimal soil:water ratio for solidphase extractants, the length of time for the membrane assay, and the best solvent system and rigor of extraction to be used. Of particular importance are the influence of other compounds and the presence of nonaqueous-phase liquids on both the biological and chemical assays. A variety of other procedures have been suggested to estimate bioavailability by chemical and physical methods; e.g., extraction with alcohols or alcohol-water mixtures (11, 15) and by organic acids (24, 25). Approaches based on analysis of the water in soil pores (12, 13) or making use of equilibrium partitioning (14, 26) offer promise for certain VOL. 33, NO. 23, 1999 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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to be done with awareness of the difficulties. Correlations of the assay with the availability to one species, as shown here, may fail when other species are deemed to be the receptors of concern. The concentration-response curve in the chemical or physical assay may not parallel the dose-biological response curve because of a no-effect threshold in the organisms or lack of a further biological response but not a chemical or physical response at high toxicant concentrations. Other toxicants at the site or unfavorable conditions in the soil may suppress the organisms, or antagonistic or synergistic effects among the toxicants may not be evident in the surrogate test. However, the need for finding a rapid, inexpensive, and precise means for determining bioavailability in soil is great, and additional research may lead to findings that will obviate or minimize some of these confounding factors. Hopefully, the approach described herein represents one step in that direction.
Acknowledgments This research was supported by funds provided by Research Grant ES05950 from the National Institutes of Environmental Health Sciences with funding provided by the U.S. Environmental Protection Agency.
Literature Cited
FIGURE 3. Correlation between uptake by earthworms of DDT, DDE, DDD, and total of the three compounds with the amount extracted by a 25% tetrahydrofuran-water solution. compounds. Indeed, equilibrium partitioning has been used for the prediction of the biological effects of DDT, DDE, and DDD in sediments (27, 28). Work in this laboratory has shown the utility of Tenax TA beads for predicting the bioavailability of DDT and related compounds in soils (17), and supercritical fluid extraction has been suggested for determination of the availability of PAHs for biodegradation (29). In addition, C18coated silicate particles have been proposed as a means for assessing uptake of hydrophobic compounds from sediments (30). Such methods should be applicable for determining the bioavailability of other classes of compounds, although the procedures would need to be modified based on the properties of those chemicals. Preliminary studies in this laboratory suggest their usefulness for determining the bioavailable fraction of polycyclic aromatic hydrocarbons in soil. In view of the array of techniques available, it is likely that nonbiological procedures will be validated to determine or predict bioavailability. On the other hand, extrapolation from the results of a chemical or physical assay to bioavailability in the field needs 4350
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Received for review May 24, 1999. Revised manuscript received September 13, 1999. Accepted September 14, 1999. ES990581W
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