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a CH2M Hill, 4350 West Cypress Street, Tampa, FL 33607
b Department of Agronomy, Purdue University, West Lafayette, IN 47907
* Corresponding author (kbanks{at}ecn.purdue.edu)
Received for publication October 27, 2004.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSREFERENCES |
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Abbreviations: MGP, manufactured gas plant • PAH, polycyclic aromatic hydrocarbon
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSREFERENCES |
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The impact of PAHs on ecosystems is significant. Several lower molecular weight PAHs are toxic to green algae and significantly reduce photosynthesis (Kusk, 1981). These compounds also may cause digestive cell death in marine mussels (Pipe and Moore, 1986). Phenanthrene decreases reproduction rates in young rainbow trout (Oncorhynchus mykiss), and increases mortality in crustaceans (Savino and Tanabe, 1989). Passino-Reader et al. (1995) found that phenanthrene significantly increased rainbow trout fry mortality with an LC50 of 0.2 mg L–1 (where LC50 is the concentration that kills 50% of the samples in a given time). A number of PAHs have been reported to be carcinogenic in humans (Phillips, 1985; Williams and Weisburger, 1986). Consequently, the USEPA has prioritized 16 PAHs for environmental regulation (Sayles et al., 1999).
Even though regulatory standards are based primarily on concentration limits, the use of bioassays for assessment of contaminant toxicity in soil is gaining in popularity (Saterbak et al., 2000; Van Gestel et al., 1992). However, only limited ecotoxicity research has been reported for PAHs in terrestrial systems, although other hazardous organic soil pollutants have been shown to significantly affect the activity of plants and soil invertebrates (Rust et al., 2004; Alexander et al., 2002; Tang and Alexander, 1999). Several bioassays have proven effective in assessing the bioavailability of PAHs (Thorsen et al., 2004). In fact, strong correlations were recently reported between the assimilation of PAHs in earthworms and bioavailable extractable concentrations (Tang et al., 2002).
Plants can be used to increase the rates of PAH degradation due to rhizosphere abiotic and biotic processes. The rhizosphere is a zone of increased microbial activity at the root–soil interface. Soil planted with perennial ryegrass resulted in significantly lower PAH and pentachlorophenol concentrations when compared to unplanted soil (Ferro et al., 1999), although Lalande et al. (2003) found that easily degradable organic matter in the rhizosphere slowed pyrene degradation. Reilley et al. (1996) reported that tall fescue, sudangrass, and switchgrass independently resulted in 30 to 40% more degradation of pyrene and anthracene than in soil without plants, while Liste and Alexander (2000) demonstrated enhanced degradation of pyrene by nine different plant species.
In addition, degradation of higher molecular weight PAHs has been observed using soil microorganisms. For example, Heitkamp et al. (1988) observed the utilization of pyrene by Mycobacterium sp. as a sole source of carbon and energy. Also, a Mycobacterium sp., isolated from a former MGP plant, was found to metabolize fluoranthene at relatively high degradation rates (Rehmann et al., 2001). However, information is limited on the biodegradation of five- and six-ring PAHs (Kanaly et al., 2001) in natural soil environments.
As mentioned previously, evidence of PAH dissipation in rhizosphere soils exists for a small number of plant species. However, there is only limited published research focusing on the degradation of PAHs in the rhizosphere of trees. The objective of this study was to assess the potential use of trees to enhance degradation of PAHs located in MGP-impacted soils.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSREFERENCES |
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Experimental Design
The greenhouse study was designed to assess the degradation of recalcitrant PAHs in the rhizosphere of several tree species. Polyvinyl chloride (PVC) pipes with a length of 2.13 m were driven to a depth of 1.83 m into wetland soil and sediments adjacent to the MGP site. The depth to ground water at the time of sampling was approximately 1 m. The sampling location was chosen based on previous identification of the area as highly contaminated (Battelle, 1999). Initial samples were collected and analyzed to confirm prior identification of impacted soil zones. Within the zone of highest contamination, intact soil cores were removed from the wetland using 15-cm-diameter PVC pipe. Immediately adjacent to every soil core taken, a time zero core also was taken. The time zero cores were sealed and transferred to the laboratory for temporary storage at 4°C until analysis. The treatment cores were capped and sealed, then transferred immediately to the greenhouse for plant establishment. Ground water was collected at the site from an adjacent monitoring well for use in the greenhouse study. This water was stored at 4°C until use. Additional ground water was collected as needed throughout the study period.
Preparation of each core for plant establishment occurred in the greenhouse. The soil core was placed on a support stand and the bottom was leveled by removing excess PVC material. A 20-cm square piece of protective fiberglass screen material was affixed to the bottom of the core to prevent the loss of contaminated material during the column placement process. Approximately 50 small holes (0.3 cm in diameter) were placed on the influent and effluent sides of the lower 0.9 m of the core (Fig. 1) . Fiberglass screen material was secured over the holes to prevent loss of soil. Two 1.2-m-long by 1.9-cm aluminum U-channels were screwed onto opposing sides of the core flush with the core base. The channels securely held the baffle boards. A watertight containment vessel was created for each core and was comprised of a 30.5-cm PVC pipe with a cap placed on one end (bottom) to create a watertight container. Each soil core was placed upright inside the cap. Then, the outer containment pipe was placed over the core and secured to the cap. The cap was sealed with 100% silicone sealant to prevent leakage. Between the inner and outer cores, wooden baffles were wedged on opposite sides of the inner core to create a barrier between the two sections and driven tightly against the cap. This barrier allowed temporary storage of water within one half of the inner space at a higher elevation than the opposite cell's outlet, forcing the passage of water through the inner core. Ground water collected from the MGP site monitoring well was used in this study. Water levels were raised and lowered periodically as needed for replenishment of water lost through transpiration and/or evaporation. All water was recycled through each column and was not allowed to leave the system except through evaporation or transpiration processes.
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Sample Handling and Processing
At each column takedown event, the soil core was removed from the water-tight containment vessel and placed horizontally on a support stand. Days before each takedown, standing water was allowed to dissipate, either by evaporation or transpiration, to ensure that there was no PAH loss through the aqueous phase. Soil cores were opened lengthwise by use of a circular saw to expose the entire soil profile. Bulk soil samples were removed in 30.5-cm increments from the open PVC core and placed into amber glass jars. Jars were sealed and immediately transferred to the laboratory for analysis.
Microbial Analyses
Anthracene (99% purity; Aldrich Chemical, Milwaukee, WI), naphthalene (99% purity; ACROS Organics, New York, NY), phenanthrene (98% purity; Aldrich Chemical), and pyrene (>97% purity; Fluka Chemika, Buchs, Switzerland) were used as the primary PAH-degrader selective growth substrates. The PAHs were dissolved in high purity n-pentane (ACROS Organics, Newark, NJ). The PAH substrate mixture was prepared in pentane by adding 2.0 g L–1 of phenanthrene, 0.2 g L–1 of anthracene, 0.2 g L–1 of napthalene, and 0.2 g L–1 of pyrene. All PAHs were completely dissolved and filter sterilized (0.22-µm filter) before addition to the microtiter plates.
All plate preparations were conducted within a sterile hood. The PAH degraders were enumerated using sterile 96-well microtiter plates (CoStar 3361 EIA/RIA Plates; Corning, Inc., Corning, NY), each with 8 rows and 12 columns. Plates were prepared by adding 50 µL well–1 of the PAH growth substrate solution. The pentane evaporated rapidly, and PAHs remained on the interior surfaces of each well. Bushnell-Hass mineral medium (DIFCO Laboratories, Detroit, MI) was used as the growth medium for the procedure. The medium was prepared, sterilized by autoclaving, and 180 µL well–1 was added.
Following preparation of the MPN plates, 5 g of test soil was added to an autoclaved glass bottle containing 45 mL of a sterilized, saline buffer solution that contained 1 g L–1 of sodium pyrophosphate (pH 7.5) and 85 g L–1 of sodium chloride. Samples were placed on a reciprocal shaker for 30 min and were set aside for 10 min to allow the solids to settle out of the solution. Ten-fold serial dilutions were performed and 20 µL of each dilution was added to each well in specified columns of the microtiter plate, with the rows used to differentiate the dilution series. The plate cover was replaced and plates then covered in plastic wrap. Samples were incubated at 25°C in the dark for 3 wk.
After 3 wk of incubation, positive results were indicated by a yellow color in the wells. Positive wells became yellow or brown due to the accumulation of the partial oxidation products of aromatics (Wrenn and Venosa, 1996). Positive wells were counted for each dilution level and recorded. An enumeration program provided by the USEPA (MPN Calculator, Version 4.04; USEPA, Cincinnati, OH) was used to analyze the raw data set.
Contaminant Analysis
The PAH concentrations in the soil were quantified using a shaking extraction method (Schwab et al., 1999) followed by gas chromotography. Contaminated soil was first sieved to 2 mm. Approximately 2 g of soil was added to a 45-mL centrifuge tube. One hundred microliters of 1000 mg L–1 tetracosane (matrix spike) was added to the soil. Twenty grams of dichloromethane was transferred to the tube. The tube was securely capped and shaken for 30 min on a reciprocating, platform shaker at 120 cycles min–1. Shaking cycles ranged between 30 and 45 min. The moisture content of a subsample sample was determined gravimetrically. Initial soil sample mass, soil moisture content, and final extract mass data were recorded. Extracts were centrifuged for 10 min at 1350 x g within a high-speed centrifuge (Sorvall RC-5B; Kendro Laboratory Products, New Haven, CT). The supernatant was decanted and stored. The extraction process was repeated once with 10 g of clean solvent added to the extracted soil to begin a new cycle. Extracts from all cycles for a given sample were combined and weighed. Extracts were placed in a sealed container and stored at 4°C until analysis. Recovery of the matrix spike was greater than 95% in all cases.
Total PAHs were quantified by gas chromatography. A 1.5-mL aliquot of the extract was transferred to a gas chromatography vial and spiked with 5 µL of 1000 mg L–1 5
-androstane (Accustandard, New Haven, CT) as an internal standard. An ampule containing a premixed certified standard of 16 priority PAHs was used to prepare standard concentration curves (ULTRA Standard PAH Mixture PM-810; ULTRA Scientific, North Kingston, RI). A minimum of five nonzero concentrations were used to construct the standard curve. The PAHs were identified by developing standardized curves for each PAH compound at known concentrations. Five increasing concentrations of PAHs within the range of interest included 10, 100, 250, 500, and 1000 mg kg–1 in dichloromethane.
The PAHs were analyzed by injecting the supernatant into a Hewlett-Packard (Palo Alto, CA) Model 6890 gas chromatograph–flame ionized detector equipped with a Hewlett-Packard 7673A autosampler. A HP-5PMS fused silica column with dimensions of 30-m x 0.25-mm nominal diameter (0.25-µm film) was used for analyte detection. Instrument settings included a splitless injection, initial temperature of 40°C, after which the temperature increased at a rate of 12.0°C min–1 to a final temperature of 320°C. The initial total gas flow was 7.7 mL min–1 with an initial pressure of 171 KPa (24.85 psi), and an average velocity of 85 cm s–1 using helium as a carrier gas.
Lettuce Toxicity Assay
Contaminated soil was first sieved to 2 mm. Each soil sample was completely homogenized. A small subsample was removed from each sample to determine moisture content. After the moisture content was determined, distilled deionized water was added to each soil sample to ensure 85% of the soil water-holding capacity was achieved before commencement of the bioassay.
Approximately 125 g of moisture adjusted, sieved soil was transferred to a 150.0- x 15.0-mm Petri dish. The soil within the dish was lightly packed to a uniform depth. Lettuce (Lactuca sativa L.) seeds (Midwest Seed Growers, Lenexa, KS) were sieved and sorted into fractions of 40 seeds each. Exactly 40 seeds were scattered across the soil of one Petri dish to within 1.2 cm of the edge. A spatula was used to press the seeds into the soil surface. A thin coat of 16 mesh silica sand was placed on top of the soil surface. The lid of the Petri dish was replaced and sealed with a 2.5-cm strip of parafilm. A set of four control dishes were prepared in the same manner except clean, fine mesh silica sand was used in place of contaminated soil. Control dishes were clearly labeled and randomly placed in the test group. The test containers were placed in a growth chamber at 22 ± 0.2°C and left in darkness for 48 h. After the initial 48-h time period, the containers were subjected to 16-h light and 8-h dark cycles for another 72 h.
Upon completion of the test period, all germinated seeds within each dish were counted. The number of seedlings in all control soil dishes was recorded. Final seedling counts for experimental samples were divided by the mean control counts so germination was represented as percent germination over control.
Earthworm Toxicity Assay
Soil was prepared as described in the lettuce germination assay. An artificial soil was prepared based on a modified method (Lanno and McCarty, 1997). This clean, artificial soil was composed of (by dry weight) 10% sphagnum peat, 20% colloidal kaolinite clay, and 70% grade-70 silica sand (Best Sand Corp., Cleveland, OH). Soil moisture was adjusted to 75% field capacity.
Earthworms (Eisenia foetida) are considered representative soil macroinvertebrates for examining effects of soil contamination (Lanno and McCarty, 1997) and were used to evaluate the changes in toxicity in this soil. Approximately 8 h before the start of the experiment, worms were hydrated and allowed to void the gut contents. Worms, in groups of 12, were removed from their container, cleaned of soil debris, and placed into beakers containing 250 mL of distilled water at 20 ± 2°C. Beakers were securely covered with paper towels and placed in a dark environmental chamber at 20 ± 2°C for 8 h. A pair of adult earthworms were prepared for the test using a hydration process where the worms are externally cleaned of soil, placed in a covered beaker with approximately 100 mL of distilled water, and incubated overnight at 20 ± 2°C.
Approximately 50 g of soil was placed in small plastic bags that were uniformly punctured 10 times just below the seal to permit air circulation. Next, 2 g of plant protein–based worm supplement, Magic Worm Food (Magic Products, Amherst Junction, WI), was placed on the soil surface within the bag. Two adult earthworms were selected together, placed briefly on a piece of filter paper to remove excess moisture, then weighed as a pair with a Mettler Model AE163 balance to ±0.1 mg (Mettler-Toledo, Hightstown, NJ). Each pair was added to the previously prepared soil and incubated at 20 ± 2°C on a 12-h day and 12-h night light cycle in a Model E-36L environmental chamber (Percival Scientific, Boone, IA). There were four replicates for each soil sample.
After incubating for 14 d, the soil from each bag was emptied and the adult worms were removed, noting mortality. Surviving adults were hydrated using the identical procedure as previously described by placing each pair in separate bottles filled with approximately 50 mL of distilled water. Bottles were loosely capped and incubated in darkness at 20 ± 2°C for another 8 h allow for purging of gut contents. Each pair was then placed on a labeled aluminum weighing dish, covered, and dried at 100 ± 5°C for 48 h. The final dry mass of each worm pair was determined using a Mettler Model AE163 balance to ±0.1 mg.
Since the initial dry mass of each pair was not measurable, it was estimated by multiplying the hydrated mass by a 0.14 coefficient determined by linear regression. This coefficient was based on the linear regression (forced through zero) of hydrated mass for actual dry mass for a sample set of 15 worms treated using the same hydration procedure described previously. Each sample set was dried in covered aluminum weighing dishes at 100 ± 5°C for >48 h and the final dry mass of each set of worms was determined. A regression coefficient of 0.1403 was based on the regression of dry mass to wet mass (r2 = 0.9846).
Statistical Analyses
All data were subjected to analysis of variance. The expected mean squares correspond to a linear model for a four-factor experiment arranged in a split plot–split block design in which blocks are random and factors treatment, time, and depth are fixed. Fisher's least significant difference (LSD) was used to compare means. The LSD was more conservative than the t test when evaluating more than two groups. For all variables, the comparisons of interest were the interaction means where all means at a given depth, vegetative treatment, and time could be compared. When indicated in the text, measurements at a given time were normalized to values before treatment (i.e., time zero). Analyses for transformations by log (as indicated) or arsin (used only for Most Probable Number for PAH degraders), were performed by the Box–Cox regression method.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSREFERENCES |
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Trees from the 18-mo study period set also produced significant shoot growth. Ash had an average growth of 73.5 cm, with poplar growth averaging 65.7 cm, and willow producing an average growth of 63 cm. Over the same period, nearly all of the trees produced prolific root growth throughout the core length and most roots had penetrated into the highly contaminated zone of 1.5 m depth.
Due to the complexity of the statistical analyses, data will be presented primarily in a table format to simplify presentation. Data were transformed as necessary to perform analysis of variance. To allow for quick assessment of sample variability, standard deviation of the original data set is shown in parentheses in all tables.
Lettuce germination assays were used in this study to evaluate the relative toxicity of the soil before and after treatment with trees. A nonvegetated soil core was incorporated into the study to serve as a control treatment. Germination data in all treatments after 18 mo were stable while the soil control showed improvement (Table 1). Although roots were noted in the contaminant zone, their presence did not significantly improve germination.
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Earthworm assays did not demonstrate a strong treatment response over the initial 9-mo study period when comparing vegetated to the unvegetated soil (Table 2). However, earthworm biomass did increase significantly (p < 0.01, n = 48) when evaluated with respect to time over the 9-mo study period. Average earthworm biomass increased from 43 mg at time zero to 136 mg after 9 mo when examined at the 1.2- to 1.5-m depth (Table 2). Soil in the poplar rhizosphere specifically had large increases in worm biomass over the 18-mo study period, but nearly equal increases were noted in the controls over the same period. Therefore, no relationship could be established based on a treatment effect, although characteristics of the soil horizon within the core had changed to allow these organisms to thrive.
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Average earthworm biomass increased from 94 mg initially to 138 mg after 18 mo of plant growth when averaged across all depths and treatments. Although no treatment effect was predominant within this bioassay as a whole, a significant decline in toxicity response was observed throughout the entire soil profile, including the depth with high concentrations of coal tar residuals. Reductions in bioassay toxicity at depths greater than 1 m are encouraging.
Bacterial enumeration by the MPN method was highly significant (p < 0.01, n = 4) at several interaction levels. At the three-way interaction level (treatment x time x depth), the greatest significant increase was seen within the willow treatment at the 1.2- to 1.5-m level (log10 4.15 cfu g–1 dry soil initially to log10 6.26 cfu g–1 dry soil after 9 mo; cfu = colony forming unit), with values normalized to time zero values. Roots found at this depth penetrated the highly contaminated zone more abundantly than all other vegetated treatments during the 9-mo study (Table 3). The presence of significant root mass in the willow cores may have contributed to the increase in soil microbial counts. Data from the 18-mo study period also show a considerable increase in PAH degraders compared to the unvegetated soil controls.
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Although microbial counts in the poplar rhizosphere soil declined statistically during the 9-mo study period (specifically at the depth of greatest contamination), poplar rhizosphere soil had the highest overall increase in microbial numbers compared to the no plant columns, with data normalized to time zero values. Early reduction in microbial numbers in the ash and poplar treatments in the 9-mo study may be attributable to the lack of the establishment of significant root systems at depth. Later in the 18-mo study period, extensive rooting in the willow and poplar treatments was shown to correlate with higher microbial populations. This interaction was significant in this study because our data provide strong empirical evidence supporting the hypothesis that the presence of deep-rooting vegetation (i.e., trees) can stimulate PAH-degrading microbial populations within the soil profile, thus contributing to the degradation of PAHs.
Concentrations of all 15 contaminants of interest decreased significantly (p < 0.05, n = 16) over time at the 1.2- to 1.5-m level. Losses of PAHs occur for many reasons, including volatilization, photodegradation, microbial oxidation, soil matrix sorption, and various chemical reactions. Lower molecular weight PAHs, naphthalene and phenanthrene (e.g., with two or three benzene rings), may have rapidly biodegraded in the soil. Two-ring and three-ring PAHs showed greater decreases at the 1.2- to 1.5-m level than the higher molecular weight contaminants assessed at the same depth.
Significant decline in contaminant concentration was observed for five of the contaminants when concentrations were normalized to time zero values. Acenaphthene, anthracene, fluoranthene, naphthalene, and phenanthrene had significant decreases (p < 0.05, n = 24) in the ash and poplar treatments when compared to the unvegetated control. The decrease is noticeably lower for naphthalene and phenanthrene after 18 mo at a depth of 1.2 to 1.5 m in the presence of vegetation (Tables 4 and 5).
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Although PAH compounds with increasing number of rings may be microbially degraded, limitations exist which may hinder bioremediation efficiency. High molecular weight PAHs (such as benzo[a]pyrene or benzo[ghi]perylene) have been reported to be degraded by soil microorganisms, but data supporting degradation in the presence of a mixture of contaminants are limited. Results from this study indicate that significant decreases in the higher molecular weight PAHs in soil are possible. Reduction trends for four-ring and five-ring PAHs were noted over the 18-mo study period, although none were at significant levels when compared to unvegetated controls. Decreases in the concentrations of chrysene and pyrene are noticeably smaller after 18 mo at a depth of 1.2 to 1.5 m in the presence of poplar and willow vegetation (Tables 6 and 7). Trends are less apparent for benzo[a]pyrene (Table 8).
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Rates of decrease in PAH concentrations were calculated for the interval between the 9- and 18-mo experimental takedowns. The rate of dissipation of the PAHs was greatest for willow, followed by poplar, ash, and the unvegetated soil controls (Table 9). These data support the hypothesis that trees can enhance the degradation of target PAHs in soil. However, it is important to note that these degradation rates were measured under greenhouse conditions. Rates of dissipation under field conditions could vary significantly from these results.
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ACKNOWLEDGMENTS |
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