Through mechanical weathering processes, these clay fragments become incorporated into the soil and may provide a long-term source of PAH contamination in the environment . Polycyclic aromatic hydrocarbons are ubiquitous environmental contaminants and 16 PAHs are considered priority pollutants by the U.S. .There have been a few ecotoxicological evaluations concerning the large PAH concentrations from clay target fragments, but these studies have reported that the PAHs elicit low toxicity in aquatic organisms . This low toxicity was determined to be primarily due to the low bio-availability of PAHs resulting from the process of making clay targets in which the PAHs in the binding agent are bound under heat and pressure with dolomitic limestone . In addition, due to their aromatic nature and hydrophobicity, PAHs typically bind to nonpolar soil domains such as organic matter, further decreasing their bio-availability . However, a recent clay target ecotoxicity study using Eisenia andrei showed that the content of clay fragments in soils was correlated with PAH bio-accumulation in the terrestrial soil organism, suggesting that direct ingestion can be a more important route of exposure and potentially explain the lack of toxicity in exposed aquatic organisms . Clay-target contaminated site evaluations have also concluded that the elevated PAH concentrations in the soil from the clay target fragments pose an unacceptable level of risk to future potential residents and current site workers.
There are several PAH remediation strategies involving physical, chemical, biological,stackable flower pots and thermal technologies; however, conventional PAH removal methods such as incineration, excavation, and land filling are expensive and inefficient . Because of these issues, biological remediation practices such as bio-augmentation and phytoremediation have become preferred in situ treatment technologies as they are considered to be cost-effective and more environmentally friendly for the cleanup of PAH-contaminated soils . However, biological remediation operations can also be ineffective due to the limited PAH soil bio-availability that is a consequence of the clay target manufacturing process and the physicochemical properties of these compounds, which can be further exacerbated by the aging effect in field-contaminated soils . These PAH bio-availability limitations can be overcome through the use of surfactants that increase the desorption of PAHs from the soil to the aqueous phase, thus increasing their bio-availability to the degrading soil microbes . Bio-surfactants such as rhamnolipids or glycolipids offer an environmentally-friendly alternative to synthetic surfactants and are becoming more economically-feasible through the use of low-cost substrates and offer distinct advantages to synthetic surfactants such as reduced toxicity, high biodegradability, and greater stability under different temperature, pH, or salinity conditions . In practical surfactant-enhanced PAH contaminated soil remediation applications, mixtures of surfactants are commonly used to take advantage of the potential synergistic effects that can result in increased solubilization at a reduced effective surfactant concentrations . The bioaugmentation of biosurfactant-producing soil microbes has also been shown to be an effective strategy for the remediation of PAH-contaminated soils. For example, M. vanbaalenii PYR-1, a glycolipid-producing microorganism isolated from an oilcontaminated estuary near the Gulf of Mexico, has been shown to enhance PAHsolubility and degradation in PAH-contaminated soils .
Another in situ biological remediation treatment commonly used to increase PAH bio-availability is phytoremediation, or the use of plants and the associated rhizosphere to restore contaminated sites . Phytoremediation is considered to be an effective, low-cost alternative to cleanup large contaminated sites . The PAH bio-availability is enhanced in the plant rhizosphere, as plant roots secrete root exudates that promote PAH desorption from the soil matrix . In addition, plant roots may release enzymes that play a key role in the degradation of PAHs including oxygenases, dehydrogenases, phosphatases, and lignolytic enzymes . Finally, plant roots also provide easily degradable carbon sources and other nutrients that increases microbial biomass, diversity, and activity, contributing to enhanced PAH degradation through direct metabolism or co-metabolism . Because of the numerous benefits provided by the rhizosphere, grass species are recommended for phytoremediation treatments due to their extensive fibrous root systems and large root surface area, and hence more extensive interactions between PAHs and the rhizosphere microbial community . Typically, the primary contaminants of concern during the remediation of outdoor shooting range soils are heavy metals from ammunition; however, large concentrations of PAHs from the clay target fragments remain in the contaminated soil and could possibly become more bioavailable during the remediation of the metals.Brij-35 nonionic surfactant and sodium dodecyl sulfate anionic surfactant were purchased from Sigma-Aldrich. Rhamnolipid biosurfactant isolated from P. aeruginosa NY3 was purchased from AGAE Technologies .Diatomaceous earth, Ottawa sand, and all GC-MS grade solvents used in this study were purchased from Thermo Fisher Scientific . All substrates utilized for soil enzymatic analyses were purchased from Tokyo Chemical Industry Co., .
Bermudagrass, switch grass, and lettuce [Lactuca sativa] seeds were purchased from Lowe’s.A Vista coarse sandy loam was collected manually using a shovel from the 0-15 cm soil depth of an abandoned shooting range located near Lake Elsinore, California that was littered with clay target fragments with no prior soil remediation or waste removal from the site. The collected soil was air-dried for 5 d at approximately 23 °C and sieved through a 2-mm stainless-steel mesh screen.The soil pH and electrical conductivity were determined potentiometrically in a 1:2 soil-to-water suspension . Total metal analysis was carried out using an Optima 7300 DV inductively coupled, argon-plasma optical emission spectrometer following U.S. EPA Method 3050B after a 6-h digestion in a mixture of nitric acid, hydrogen peroxide, and hydrochloric acid at 95 °C .Mycobacterium vanbaalenii PYR-1 was stored at -80 °C in a 30% glycerol stock and the inoculum was prepared according to a previous method in MBS solution amended with pyrene as a carbon source . The CMC of Brij-35 and rhamnolipid biosurfactant was determined previously . The CMC of the Brij-35/SDS surfactant mixture was determined by measuring the surface tension of surfactant solutions over a concentration range using a Du Noüy ring-tensiometer and using the inflection in the plot of surface tension against surfactant concentration. The CMC was determined to be 0.099 mM at 0.5/0.5 molar fraction, which was similar to a previous study .After the soil was thoroughly mixed, 1 kg soil was placed in a stainless-steel bowl and 150 mL of distilled water was added and mixed to achieve a soil water potential of approximately -33 kPa determined by a soil tensiometer. For the M. vanbaalenii PYR-1 bioaugmented treatments, 15 mL M. vanbaalenii PYR-1-MBS solution was added to yield approximately 106 CFU/g soil and thoroughly mixed . The same procedure using only the MBS solution was added to the non-inoculated, or native, soil as the control. The PYR-1-MBS solution was reapplied every 2 months by adding the inoculum solution into the soil rhizosphere 5 cm below the soil surface . Once the soil treatments were prepared, the soil was added to the phytoremediation sample containers,flower pots for sale which consisted of 800-mL glass jars that were first painted on the outside with black paint, followed by aluminum enamel to prevent exposure to light . The pots contained approximately 50 g of 2-cm diameter gravel at the bottom to allow for accumulation of any excess soil water . Bermudagrass and switch grass seeds were surface-sterilized by three sequential washings in 0.1% sodium hypochlorite, followed by two rinses with sterile distilled water . Bermudagrass and switch grass seeds were planted at a rate of 20 seeds/pot and sealed with plastic wrap for 1 week for optimal seedling emergence conditions. After 2 weeks, plants were thinned to 8 plants/pot and amended with a commercial fertilizer for bermuda grass establishment. Treatments were then fertilized monthly with 100 mg/kg-N as urea, and 12.5 mg/kg-P as monobasic potassium phosphate . Due to the potential toxicity of surfactants to emerging plant seedling , surfactant addition at 50 mg/kg was initiated 1 week after plant thinning and initial fertilizer application. Since rhamnolipid biosurfactants have been previously shown to be degraded by the soil microbial community and are considered more biodegradable than the synthetic surfactants used in this study, surfactants at the initial rate were reapplied to the soil surface every 40 d .
Each pot was placed randomly in one of four blocks, each containing one replication of all treatment combinations in a climate-controlled growth chamber . The PAH phytoremediation experiment was continued for 8 months in the growth chamber under a 12/12 hour day/night period at 23±1/19±1 °C and 40% relative humidity. The average light intensity was obtained through fluorescent and incandescent lighting in the growth camber . Each pot was weighed daily for 8 months and the soil moisture was gravimetrically adjusted to 20% by application of distilled water . The quantity of distilled water added to the soil to achieve proper soil moisture was not adjusted for vegetation biomass produced during the study. Plant shoots were trimmed to a height of 5 cm every 3 months in order to stimulate continuous plant growth . At the end of the 8-month phytoremediation experiment, plant shoots and roots were separated from the soil as described in section 2.7. Once the vegetation was removed, the soil was sieved to pass through a 2-mm sieve and separated into two subsamples. The first soil subsample was air-dried for 7 d at approximately 23 °C in the dark and used for PAH analysis and toxicity assay . The second soil subsample was used for soil enzyme analysis and kept at fieldmoist conditions and analyzed within 1 week after the termination of the experiment. set at 60 °C and then raised at 5°C/min to 280°C . Quantification of PAHs was performed using an internal standard-normalized calibration curve and coefficients of determination for all calibration curves fulfilled the requirement of R2 ≥ 0.99. Soil dehydrogenase soil activity was analyzed by the use of 2–3- -5-phenyl tetrazoliumchloride as a substrate . A 1.0-g soil aliquot was mixed with Tris buffer and INT substrate in a stoppered 100-mL Erlenmeyer flask, and the mixture was incubated for 2 h at 40 °C in the dark. After incubation, the mixture was extracted using 10 mL N,Ndimethylformamide:ethanol mixture for 1 h at 23 °C in the dark and shaken every 20 min. Immediately after filtration, iodonitrotetrazolium formazan formation was measured colorimetrically at 464 nm against the reagent blank using a UV-Visible spectrophotometer . Soil dehydrogenase activity was expressed as µg INTF produced/g dry soil 2h. Soil polyphenol oxidase activity was measured by the utilization of pyrogallic acid as a substrate to form purpurogallin . Ten mL of 1.0% pyrogallol was added to 1.0 g soil sample and incubated at 30 °C for 2 h at 200 rpm. Afterwards, 5 mL of citrate-phosphate buffer was added to the treatment to stop the reaction, followed by the addition of 35 mL ether and shaking for 30 min at 200 rpm. The colored ether with dissolved purple gallic prime was measured colorimetrically at 430 nm on a UV-Visible spectrophotometer.The polyphenol oxidase activity was expressed as mg purpurogallin produced/g dry soil 2h. Control assays for each soil enzyme activity included autoclaved soil treatments, assays without soil, and assays without substrate addition during incubation . Results of soil enzyme activities are reported on an oven-dry-weight basis.At the end of the phytoremediation experiment, plant shoots were cut at the soil surface and rinsed with distilled water to remove any adhering soil. Approximately 4 g shoot subsample was taken and freeze-dried for PAH extraction and the remaining shoots were dried to a constant weight at 55 °C and weighed to determine total shoot biomass. The freeze-dried plant shoots were ground to pass a 2-mm, stainless-steel mesh screen using a Wiley Mill Grinder and 2 g was used to determine PAH shoot concentrations using procedures similar to those for soil PAH extraction . Plant roots were manually collected from the soil using forceps, placed on a 500-µm stainless-steel sieve, and thoroughly rinsed with distilled water to remove any adhering soil particles. Approximately 2 g root subsample was taken and freeze-dried and 1 g was used to determine PAH root concentrations similar to shoot analysis. The remaining plant roots were dried to a constant weight at 55 °C and weighed to determine total root biomass. The lettuce seed toxicity assay was performed to evaluate changes in phytotoxicity before and after remediation treatments by following a method in Cofield et al. . Briefly, 100 g soil at 85% water-holding capacity was placed in a 150 mm ´ 15 mm Petri dish and 40 lettuce seeds were evenly distributed and pressed into the soil.