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Microbial community and ecotoxicity analysis of bioremediated, weathered hydrocarbon-contaminated soil.

Introduction

Contamination by hydrocarbons such as polycyclic aromatic hydrocarbons (PAHs), has attracted considerable public attention due to their toxic, mutagenic, and carcinogenic potentials (Husaini et al. 2008; Peng et al. 2009). The frequent transportation of hydrocarbons as a result of their widespread use as an energy source has led to increased risks of environmental pollution of vital soil and water resources. These increased risks of pollution have raised legitimate concerns about public health safety and environmental conservation (Philip et al. 2005; Peng et al. 2009).

Hydrocarbons are diverse (Lindley 1992) and susceptible to microbial degradation processes, which can remove considerable quantities of hydrocarbons from the environment (Genovese et al. 2008). Bacterial groups such as Pseudomonas, Acinetobacter, and Rhodococcus (and to a lesser extent fungi) are thought to be important hydrocarbon degraders and consequently have been well studied (Boonchan et al. 2000; Hamamura et al. 2008; Wu et al. 2008). These microbial groups are thought to mediate the process of natural attenuation of hydrocarbon contaminants in polluted soils (Bento et al. 2005).

The disposal of soils contaminated by hydrocarbon is usually carried out after either physicochcmical or biological treatments have substantially reduced the levels of total petroleum hydrocarbon (TPH), metals/metalloids, organics, and other toxic substances to specific concentrations deemed to be environmentally safe. These levels or guidelines, which are usually set by legislation, vary from one country to another and in Australia are set by the National Environment Protection Measure (NEPM) (NEPC 1999). The NEPM guidelines are partly derived from the 'Australian and New Zealand Guidelines for the Assessment and Management of Contaminated Sites', prepared by the Australian and New Zealand Environment and Conservation Council and National Health and Medical Research Council and based on standards from Europe and North America and studies carried out in Australia (ANZECC/NHMRC 1992). These measures are designed to provide adequate protection of human health and the environment through the development of an efficient and national approach. The guidelines are considered for both in situ and ex situ treatment of polluted soil and disposal of affected soil after successful treatment.

These guidelines have some practical limitations which may affect their use for the determination of the environmentally safe levels of toxic substances. For example, the assessment of environmental hazards of remediated soil is limited to specific contaminants of concern, such as benzo(a)pyrene (Plaza et al. 2005), and a universal safe TPH value (NEPC 1999; EPA 2010) is usually set for remediated hydrocarbon-contaminated soils. These values may not adequately recognise the effects of factors such as soil type (sand or clay), oil type (weathered or fresh), and chemical characteristics (Dorn and Salanitro 2000) on hydrocarbon toxicity. Soils with high TPH values (unsafe) may be easily remediated because they contain easily degradable alkanes. In contrast, another soil may have a legally safe TPH value but may contain recalcitrant compounds which may not be bioavailable. However, such soils should not be assumed to be safe without suitable microbiological and ecotoxicity tests to ensure that the petroleum compounds and their metabolites are not harmful to soil flora and fauna (Wang et al. 2010).

The impact of hydrocarbon pollution on microbial community can be assessed using a variety of molecular techniques, such as denaturant gradient gel electrophoresis (Muyzer et al. 1993) and real rime polymerase chain reaction (PCR). Such investigations can show whether bioremediation processes have caused any change in the soil microbial communities of such polluted soils (Evans et al. 2004; Kao et al. 2010). Soil organisms such as earthworms are also important biological indicators of toxic compounds and ecotoxicity tests are needed to verify that the treated soil is also safe for these macroorganisms. Although various ecotoxicity assays exist for assessing remediated soils, the correlation between the results of these assays and the mandated chemical analysis for determining the safety of treated soils is limited (Mao et al. 2009). Ecotoxicity testing could be useful as a supplementary tool and an additional safety measure for supporting management decisions for remediation strategies (Salanitro et al. 1997; Plaza et al. 2005) and the disposal of remediated soils.

In Australia, there are presently no legal requirements to carry out ecotoxicity assessments on remediated soils before use for residential purposes or disposal as wastes. Consequently, remediated soils are being used for a variety of purposes without propedy assessing their impacts on soil biota. Therefore, the aims of this study were to (i) assess the effectiveness of bioremediation treatments on the breakdown of weathered hydrocarbon in contaminated soils, and (ii) carry out a safety appraisal via ecotoxicity tests on these remediated soils that have met the legislated safe hydrocarbon levels.

Material and methods

Soil characteristics and metal and pesticide analyses

The original soil contamination was caused by an accidental spillage of transformer oil at an industrial site, leading to high levels of soil TPH (~18 000 kg/mg, Table 1). The soil was excavated after this pollution event and stored as a pile at a landfill site in Adelaide, South Australia, for approximately 3 years. During this time, the pile was exposed to natural elements (weathered) and subject to monitored natural attenuation. The soil samples used for this study were obtained from this 3-year-old, weathered pile. Stones were removed manually, and the sample was sieved (10mm) to remove large bulking agents and finally fine-sieved (<2 mm). Organic carbon, nitrogen, moisture content, organic matter, and pH of the soil were determined using standard methods (Mishra et al. 2001). Table 2 summarises the main characteristics of the treated, contaminated soil. The soil samples were also analysed for metal and pesticide contents using a combination of USEPA methodologies: Microwave Digestion USEPA 3051A (www.epa.gov/osw/hazard/testmethods/sw846/pdfs/3051a.pdf, accessed 4 June 2010); and ICP/MS determination, USEPA (www.epa.gov/osw/hazard/testmethods/sw846/pdfs/6020a.pdf, accessed 4 June 2010). The results are summarised in Table 3.

Experimental design and soil treatments

The soil samples were subject to different bioremediation treatment methods in order to reduce the soil TPH levels to legally safe levels for landfill disposal and domestic (residential) use. This was carried out in laboratory-based microcosms which were performed in triplicate, with each microcosm containing 200 g of soil sample. The experiment consisted of four treatment methods: control, with no additional treatment (natural attenuation, NA); addition of nutrients (biostimulation, BS); addition of known hydrocarbon degraders (bioaugmentation, BA); and a combination of biostimulation and bioaugmentation (BABS). The BS microcosms had the addition of a modified nutrient formulation optimised for fungal inoculum growth based on Bushnell Haas medium at 0.06mL/g soil (Eriksson et al. 2000). The BA microcosms had the addition of hydrocarbonoclastic fungal mycelia (Scedosporium apiospermum) (0.5mg/g soil) (Martin-Gill et al. 2008) which had been successfully used to treat other hydrocarbon-contaminated soils (Makadia et al. 2011). The BABS treatment consisted of the nutrient formulation (0.06 mL/g soil) and fungus (0.5 mg/g soil). The microcosms were incubated at 30[degrees]C and adjusted to 50% of soil water holding capacity for 12 weeks. Aeration and moisture content of microcosms were maintained on a weekly basis.

Determination of total petroleum hydrocarbon degradation analysis

The TPH analysis was carried out on soil samples from the different microcosms at selected time intervals (Day 0 and Weeks 2, 4, 6, 8, 10, and 12) and procedural blanks. Hydrocarbons in replicate soil samples were extracted using a modified standard protocol of determining hydrocarbon content in soil according to Intemational Standard Organisation (ISO 2004), ISO/DIS 16703 GC method. Five mL of acetone was used to extract oil from 1 g of homogenised, contaminated soil in a glass tube. A 2-mL aliquot containing a mixture of standard solutions (retention rime window, RTW; consisting of n-decane and n-tetracontane at 20 and 30 [micro]g/mL, respectively, in heptane) was then added to the contaminated soil, after which the rest of the protocol as described in the 1SO/DIS 16703 GC-method was followed. A standard calibration curve was constructed from hydrocarbon mixture (RTW solution) dilutions, and the equation from the standard calibration curve was used in conjunction with the area under each chromatogram (area between [C.sub.6] and [C.sub.36]) to determine TPH concentrations. The residual hydrocarbons in soil samples were analysed on an 8200 Autosampler Gas Chromatograph equipped with a Varian 8200 Autosampler and Flame Ionization Detector. A programmed temperature splitless injection was used. The capillary column used was an Alltech EC 5 (30m by 0.25 mm with 0.25 [micro]m film thickness), with helium as a carrier gas flowing at a rate of 2 mL/min in a constant flow mode.

DNA extraction and polymerase chain reaction

Genomic DNA was extracted from the soil samples using a DNA Isolation Kit (MoBio PowerSoil, Carlsbad, CA, USA) as per the manufacturer's protocol. Community DNA extractions were performed in replicates at selected rime points (Weeks 0, 4, 8, and 12) to analyse the changes in the microbial (bacteria and fungi) community profiles over the course of the 12-week experiment.

Bacterial 165 rDNA amplification

The PCR amplification of bacterial 16S rDNA (rDNA) was performed in a 50-[micro]L PCR reaction mixture containing 2 [micro]L of forward primer 341FGC (10pmol/[micro]L), 2 [micro]L of reverse primer 518R (10 pmol/[micro]L), 5 [micro]L of magnesium chloride (25 mM), 1 [micro]L of deoxynucleoside triphosphate (dNTP) mixture (10mM), 10 [micro]L of GoTaq Flexi buffer (5x), 0.25 [micro]L of Taq polymerase enzyme (5 U/[micro]L), 1 [micro]L of bovine serum albumin, and 26.75 [micro]L of sterile nuclease-free water per PCR reaction. For each sample, 2 [micro]L of purified template DNA extract was added to 48 [micro]L of PCR master mix. The PCR was conducted with universal bacterial primers 341FGC as the forward and 518R as the reverse primer (Muyzer et al. 1993). The thermocycling program used consisted of one cycle of 5 min at 95[degrees]C; 33 cycles of 30 s at 94[degrees]C, 30 s at 55[degrees]C, 1 min at 72[degrees]C; and a final extension at 72[degrees]C for 10 min.

Fungal DNA amplification

Internal Transcribed Spacer regions (ITS) of fungi were targeted using a two-step PCR process. The first round of PCR was with the primers ITS1F and ITS4 (Anderson et al.

2003), while the second round of amplification was with the primers ITS1FGC and ITS2 (Anderson and Parkin 2007). The master mixes of both reactions comprised 2 [micro]L of forward primer (10pmol/[micro]L), 2 [micro]L of reverse primer (10pmol/[micro]L), 5 [micro]L of magnesium chloride (25 mM), 1 [micro]L of dNTP mixture (10mM), 101[micro]L of GoTaq Flexi buffer (5x), 0.251[micro]L of Taq polymerase enzyme (5 U/[micro]L), and 27.75 [micro]L of sterile nuclease-free water. For each sample, 2 [micro]L of purified template DNA extract was added to 48 [micro]L of PCR master mix with amplicons from the first reaction being used as template DNA for the second reaction. The thermocycling program used consisted of 1 cycle of 5 min at 95[degrees]C; 35 cycles of 45 s at 94[degrees]C, 45 s at 58[degrees]C, and 45 s at 72[degrees]C and a final extension at 72[degrees]C for 10 min.

AlkB functional group based PCR

A two-step PCR employing a first PCR round using forward (F485) and reverse (R851) primers without a GC clamp was used to facilitate PCR amplification. This was followed by a second-round PCR with forward primer F485 and reverse primer R851 GC with a GC clamp (Hamamura et al. 2008). The PCR cycle consisted of 1 cycle of 10 min at 94[degrees]C; 30 cycles of 45 s at 95[degrees]C, 45 s at 64[degrees]C, and 1 min 30 s at 72[degrees]C; and a final extension at 72[degrees]C for 7 min. The same two-step procedure was used to assay for alkB genes using group-specific Pseudomonas and Acinetobacter primers as described by Hamamura et al. (2008).

Denaturing gel gradient electrophoresis (DGGE)

The changes in microbial community composition during bioremediation were monitored with DGGE. The PCR amplicons were analysed on a DCode apparatus (BioRad) on an 9% polyacrylamide gel (acrylamide-N,N'-methylenebisacrylamide ratio 37 : 1) in 1 x TAE. Electrophoresis was carried out for 18 h at 600C at a constant voltage of 60 V with a linear denaturant gradient of 47--60% (universal bacterial community), 45-62% (alkB community), and 40-50% for fungal community. After electrophoresis, the gels were silver-stained (McCaig et al. 2001), scanned, and saved as TIFF with an Epson V700 scanner. Digitised DGGE gel images were processed using TotalLab analysis package, (Version TL3; Nonlinear Dynamics, USA). The banding patterns were analysed with TotalLab to generate similarity profiles using the unweighted pair group method with mathematical averages (UPGMA) and overall community diversity using the Shannon Weaver Diversity Index (H').

Ecotoxicity analysis

This was carried out to assess the safety of bioremediated soils to soil biota. As the monitored NA soils had the lowest TPH content (legally safe levels for in situ bioremediation and landfill disposal as low level contaminated waste) after 12 weeks of incubation, these samples were used for ecotoxicity assays. The ecotoxicity tests were conducted by the ASTM guideline (ASTM 2004) to determine the safety of the treated soil (ar legally safe levels) to earthworms and radish seeds (soil flora and fauna).

Earthworm (Eisena fetida) assay

Twenty-four hours before the ecotoxicity assay, earthworms (Eisena fetida) were placed into the reference soil, a premium grade potting mix (Amgrow Organix, NSW), to ensure that the potting mix was non-lethal. A range of contaminated soil concentrations utilising only NA samples from Week 12 (in replicate) was prepared by mixing the premium-grade potting mix with various concentrations (w/v) of contaminated soils: 15% (~825 mg/kg), i.e. legally safe level for ex situ bioremediated soil use for domestic and residential purposes; 25%, 50%, and 75%, concentrations legally safe for landfill disposal; and 100%, i.e. only contaminated soil. Controls were set up with 100% potting mix. An appropriate soil volume (200 mL) was added to a clean glass beaker. Adult earthworms (10 per sample, washed and weighed) were placed on the surface of the soil and allowed to burrow. Each container was covered with perforated cling wrap (to allow aeration) and incubated ar room temperature (21[degrees]C) for a natural day/night period of 14 days. At appropriate time intervals (Days 1, 2, and 14) living earthworms were removed, counted, washed, and weighed before being placed back into the containers.

Radish seed germination assay

A slightly modified method from Banks et al. (2003) was used with the previously described ranges of contaminated soil concentrations. Seven radish (Raphanus sativus) seeds (Mr Forthergill, NSW) were planted in Petri dishes containing 50 g of NA soil sample. The Petri dishes were covered and incubated in the dark for 48h at 21[degrees]C. After 48h, the samples were randomly placed in a growth chamber, lids removed, with a light cycle of 16 h/8 h light/dark for 5 days. The number of germinations were calculated, with each seed being considered germinated if it sprouted above the cover soil (Banks et al. 2003).

Statistical analyses

Statistical significance of the data was evaluated by repeated-measure analysis of variance (ANOVA) and Tukey's multiple comparison tests. The tests were used to determine the statistical difference of total petroleum hydrocarbon degradation between treatments and time periods. Data were considered to be significantly different among values if P < 0.05. Version 2.0 of SigmaStat (Systat Software, CA) was used for all statistical analyses.

Results

Biodegradation of TPH

At time zero, the TPH ([C.sub.6]-[C.sub.36]) concentrations of the soil mesocosm samples were ~10 133mg/kg for NA, 9666mg/kg for BS, 9800mg/kg for BA, and 10 933mg/kg for BABS (Table 1). The TPH levels in both the treated and NA microcosms thereafter fluctuated until Week 12 with final concentrations of 5733 mg/kg for NA, 6500mg/kg for BS, 6666mg/kg for BA, and 6866mg/kg for BABS for C6-36 fractions. The data showed that there was a significant difference between the initial and final TPH concentrations (P<0.05), although the gas chromatograms showed only slight changes in the hydrocarbon profile of treated and control samples over the 12-week period. These chromatograms showed the presence of high-end hydrocarbons [C.sub.16]-[C.sub.35] (96%) in the weathered, oil-contaminated soils (data not shown). As the NEPM guidelines require only the [C.sub.16]-[C.sub.35] fractions (NEPC 1999), adjusting final TPH values of the microcosm to reflect the 96% composition of [C.sub.16]-[C.sub.35] resulted in (mg/kg) 5503 for NA, 6240 for BS, 6399 for BA, and 6591 for BABS (Table 1). Therefore, the NA values were lower than the safety threshold of NEPM guidelines for in situ soil bioremediation (Table 3).

Effect of treatments on bacterial and fungal community profile and diversity values

Analysis of the DGGE-based soil microbial profiles of the different microcosms showed that both bacterial and fungal community UPGMA clusters were largely influenced by sampling rime rather than by treatment. This indicated that treatment had very little effect on the microbial community banding patterns at Weeks 4, 8, and 12 (data not shown). The assessment of the alkB bacterial community (Fig. 1) was only successfully carried out with Rhodococcus group specific primers, with few or no amplicons being obtained from Pseudomonas and Acinetobacter PCR despite different optimisations. However, the alkB rhodococcal community also showed clustering largely by time rather than by treatment, although to a lesser extent than as observed in the universal bacterial community. The Shannon Diversity values (data not shown) of the bacterial (universal and alkB) and fungal communities in both treated and control samples were not significantly different (P>0.05) from one another throughout the 12-week incubation period. This suggested that the bacterial and fungal communities were largely stable in the different microcosms.

[FIGURE 1 OMITTED]

Ecotoxicity analysis

Earthworms

The control samples containing 100% premium-grade potting mix were found to be non lethal to the earthworms, with all worms surviving the 14-day period. The earthworms successfully burrowed into the potting mix without showing any behavioural signs of distress (Fig. 2). However, the earthworm survival rates were different in potting mixes containing different percentages of NA soils from Week 12 with increasing concentrations of contaminated soils being associated with an increase in earthworm mortality. Ali earthworms in the 100% (5503 mg/kg) contaminated soil died within 24h, with a 60% survival rate in 75% (4127mg/kg) contaminated soils and 100% survival in 15-50% (825-2751 mg/kg) contaminated soils within the same time frame (24h). By Day 2 (48 h) the survival rate dropped to 0% in 50% and 75% contaminated soils. At the end of the experiment, only the control and 15% contaminated soils (100% survival rate) and 25% (1375mg/kg) contaminated soils (90% survival rate) still had living earthworms (Fig. 2).

Radish seeds

Soil toxicity was also assessed through the use of plant germination index and emergence using radish seeds. In Fig. 3, the success of germination by the emergence of the radish through the soil from the control (premium potting mix) soil indicated that it was not toxic to the seeds. When treated contaminated soil was mixed ata concentration of [greater than or equal to] 50% with potting mix soil, no germination was observed (Fig. 3). In addition, the percentage reduction in growth compared with the control showed that contaminated samples at concentrations of 10% (550mg/kg, legally safe ex situ bioremediated soil TPH levels for domestic use) and 25% (1375mg/kg) had a germination rate that was half of the control. The figure also illustrated high percentage reduction (>70%) in root elongation in most contaminated soil-potting mix samples.

[FIGURE 2 OMITTED]

Discussion

Analysis of TPH and microbial response

The soil used in the study was contaminated with hydrocarbons in 2007 (18000mg/kg) and naturally attenuated until 2009 (final TPH of 10 000 mg/kg) (Table 1). As the levels of soil TPH (10 000 mg/kg) and metals were within acceptable guidelines of NEPM (NEPC 1999) and Environmental Protection Authority (EPA 2010) with PAH not being detected (Table 3), the soil samples were legally suitable for discharge into selected landfills as low level contaminant wastes (Table 3). However, the gas chromatography profile showed that a substantial percentage (at least 96%) of the TPH comprised [C.sub.16]-[C.sub.35] fractions, which were probably rapidly degraded by microorganisms. Consequently, this contaminated soil was then subject to a variety of bioremediation strategies (biostimulation and/or bioaugmentation) to degrade these long-chain hydrocarbon fractions.

Analysis of the final TPH values showed substantial reduction by Week 12, which ranged from 31% in BA to 43% in NA. This result indicated that these bioremediation strategies conferred no additional enhancing effect of hydrocarbon degradation compared with natural attenuation. While there are reports on the beneficial effects of these bioremediation strategies (Margesin et al. 2007; Manceralopez et al. 2008), other researchers have shown that biotreatment (just as observed in this study) may not always result in enhanced hydrocarbon degradation (Bento et al. 2005; Vinas et al. 2005). This may be due to available (enhanced) soil microbial hydrocarbon catabolic potential, soil ion-binding properties, the type of contaminating hydrocarbons (Bento et al. 2005), or soil physico-chemical processes such as volatilisation. While processes such as volatilisation tend to occur with lower end hydrocarbons and at elevated temperatures, microbial degradation can occur at different temperatures. Microbial degradation of alkanes (natural attenuation) involves an initial oxidation of n-alkanes to their corresponding primary alcohol by enzymes such as monoxygenases, dioxygenases, and hydroxylases (Wentzel et al. 2007). The rest of the process and hydrocarbon degradation pathways are well described in literature (Wentzel et al. 2007; Rojo 2009).

[FIGURE 3 OMITTED]

Assessment of the soil microbial community profile using DGGE analysis revealed strong and diverse bacterial and fungal communities which were not substantially altered by biotreatment. This lack of change in the microbial community might have been occurred because the communities had already adapted to the presence of hydrocarbons. Nevertheless, the detection of alkB genes confirmed the presence of hydrocarbon-degrading communities (capacity), which may also partly explain why there was very little change in the microbial community composition during treatment. Therefore, the addition of extra nutrients did not increase the rate of hydrocarbon degradation. As very few changes were observed in the gas chromatography and microbial profiles of the biotreated soils, ecotoxicity analyses were carried out to assess the safety of the soils (at legally safe TPH levels) to other soil biota after mixing them with potting mix.

Ecotoxicity analysis and disposal levels

Ecotoxicity tests are usually carried out with earthworms (Eisena fetida) and radish seeds (Raphanus sativus). The earthworm assay is a sensitive test, and several effect-based end-point measures including mortality, growth, cocoon output, juvenile production, and avoidance behavioural response can account for the bioavailability of petroleum products in soil and therefore highlight toxicological impacts of exposure (Wang et al. 2010). The radish seed germination test is designed to highlight any phytotoxic issues with the remediated soils. Seeds have a seed coat, which is a primary line of defence between the embryo and its immediate environment (Adam and Duncan 2002; Hubalek et al. 2007). However, this protection is limited and the inhibitory effect of oil on seed germination may be attributed to physical constraints rather than biological damage to the seed, especially if the seed is coated with contaminating oil which may prevent water from getting to the seed.

Currently in Australia, there are well-defined guidelines and different disposal levels (based on whether the soil was remediated ex situ or in situ) for remediated soils. If a soil is treated ex situ, then a TPH level of 1000mg/kg (health investigation levels) (>C9 TPH) is legally acceptable for domestic and residential use of the soil, including gardening (Table 3). In this study the TPH level of the naturally attenuated soil sample was sufficiently low to be legally disposed of as wastes or when mixed in potting mix fit for use for domestic and residential purposes. Yet the treated soils at 10% (~550mg/kg) concentrations, which were within the acceptable health investigations level (Table 3), were still toxic to radish seed germination. Interestingly, no ecotoxicity investigations are presently legally mandatory and no guide ecological investigation levels are available before soil use for residential purposes in the current NEPM guideline (NEPC 1999). The results from this study, however, indicate that these soils, which potentially could have been used for gardening, were still toxic and probably not appropriate for domestic and residential use despite meeting the recommended chemical (safe) guidelines. Therefore, the fact that no ecotoxicity tests are legally required makes ir difficult to accurately assess the safety of any remediated soil to soil biota under present Australian guidelines. Although the reason for this toxicity was not investigated in this study, it was possible that the observed soil toxicity was related to the contaminating hydrocarbon type (no heavy metals, pesticides, aromatics detected in soil) (Table 3).

According to the 'Australian Guidelines on Investigating Levels of Soil and Groundwater' (NEPM guideline, NEPC 1999), once the [C.sub.16]-[C.sub.35] aliphatic soil concentration falls below 5600mg/kg, the soil being remediated in situ should be safe (health investigation levels) for residential use (Table 3). Although the 5503mg/kg ([C.sub.16]-[C.sub.35]) TPH concentrations recorded in naturally attenuated soils in this study were off-site (and technically an ex situ process), the fact that they were obtained in naturally attenuated soils indicated that these reductions could have potentially happened on-site. Yet, these soils at this concentration (5503mg/kg, 100% contaminated soil) caused 100% earthworm mortality (Fig. 2) and completely inhibited seed growth (Fig. 3), with the danger to other soil macro-organisms (which are important for soil processes and fertility) remaining unquantified. Even at lower naturally attenuated soil concentrations 50-75% (~2751-4127 mg/kg) for earthworm and 10-75% (~550-4127 mg/kg) for seed germination), the soil was still toxic to both earthworms and radish seeds. This indicated that even when in situ remediation contaminant levels are acceptable, they may still be potentially unsafe for soil biota. Although it was possible for macro-organisms such as earthworms and plants to adapt to the presence of contaminants over rime, leading to an increase their survival or growth rates, this soil could still be unsafe especially if used for gardening (possible bioaccumulation of toxicants in plants that may end up in the food chain). Therefore, it may be beneficial to the environment and human health if ecotoxicity tests are carried out in order to validate their safety. We recommend that ecotoxicity tests form part of the legal requirements for assessment of contaminated soils.

Conclusion

The study has shown that monitored natural attenuation could be as efficient as other bio-treatments and the need to conduct ecotoxicology analysis of remediated contaminated soil before disposal. Ecotoxicity assays are already being applied for assessing the success of soil bioremediation methods (Schultz et al. 2004; Molina-Barahona et al. 2005; Plaza et al. 2005; Parrish et al. 2006; Hubalek et al. 2007) and it would be beneficial to the environment if ecotoxicity assays are part of the mandatory tests required for assessing the safety of treated soils. There is also a need for the development of different safety guidelines for different soil types and different thresholds for safe hydrocarbon levels (as obtained in other countries) which take into account the type and nature (fresh or weathered) of the hydrocarbon contaminant. Currently there are guidelines for different hydrocarbon compounds and fractions, but there are other important factors which affect biodegradation ability such as age, oil characteristics, bioavailability and soil type which are not adequately accounted for in the present regulation.

10.1071/SR10159

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Manuscript received 2 August 2010, accepted 11 November 2010

Petra J. Sheppard (A,B), Eric M. Adetutu (A), Tanvi H. Makadia (A), and Andrew S. Bali (A)

(A) School of Biological Sciences, Flinders University of South Australia, GPO Box 2100, Adelaide, SA 5001, Australia.

(B) Corresponding author. Email: shep0131@flinders.edu.au
Table 1. Soil sample total petroleum hydrocarbon ([C.sub.6]-
[C.sub.36] and [C.sub.16]-[C.sub.35], mg-Icg) history analysed by
gas chromatography-mass spectrometry Total petroleum hydrocarbons
removal by the bioremediation treatment: natural attenuation
(NA), biostimulation (BS), bioaugmentation (BA), and combined
bioaugmentation-biostimulation (BABS). Data are means of three
replicates f standard deviation

                2007                     2008

                                 [C.sub.6]-[C.sub.36]

       NA 18 000 [+ or -] 3000   17 333 [+ or -] 1527
BS               --                       --
BA               --                       --
BABS             --                       --

               2009

       10 133 [+ or -] 757
BS       9666 [+ or -] 230
BA       9800 [+ or -] 529
BABS   10 933 [+ or -] 1171

          2009 (Week 0)         2009 (Week 12)

                 [C.sub.16]-[C.sub.35]

         9727 [+ or -] 726    5503 [+ or -] 775
BS       9279 [+ or -] 220    6204 [+ or -] 1221
BA       9408 [+ or -] 507    6399 [+ or -] 110
BABS   10 495 [+ or -] 1124   6591 [+ or -] 442

Table 2. Soil characteristics measured

pH                                     8.3
Nitrogen (%)                           0.34
Carbon (%)                             9.6
Moisture (%)                          16.4
Organic matter (%)                     3.43
Nutrient content (mg/kg)
  Phosphate (P[O.sub.4.sup.3]          1.23
  Nitrite (N[O.sub.2]                  0.35
  Nitrate (N[O.sub.3]                  6.67
  Ammonium ([N[H.sub.4.sup.+]          0.97

Table 3. Recommended guidelines on investigation levels for soil
and groundwater (mg/kg) and comparison to soil sample results

NEPM, National Environmental Protection Measure; EPA,
Environmental Protection Authority; PHC, petroleum hydrocarbon
components; NA, natural attenuation. Guidelines A: in situ
bioremediation standard residential with garden/accessible soil
(this category includes children's day care centres,
kindergartens, preschools, and primary schools). Ex situ
bioremediation guidelines: Interim urban, interim ecological
investigation level for urban settings, based on consideration of
phototoxicity; Waste-fill soils, can be used for residential
purposes; Low level waste soils, can only be disposed of in
approved landfills. -, No data available

Guidelines                             NEPM investigation levels

                                     Health:        Ecological:
                                     Guidelines A   Interim urban

Metals/metalloids
  Arsenic                                 100             20
  Cadmium                                  20              3
  Copper                                 1000            100
  Lead                                    300            600
  Manganese                              1500            500
  Nickel                                  600             60
Organics
  DDT+DDD+DDE                             200             -
  Benzo(a)pyrene                            1             -
PHC (constituents)
  >[C.sub.16]-[C.sub.35] aromatics         90             -
  >[C.sub.9] PAH                           20             -
  >[C.sub.16]-[C.sub.35] aliphatic       5600             -
  >[C.sub.9] TPH                           -              -

Guidelines                                     EPA
                                            Waste soil

                                     Waste-fill   Low level
                                                  waste soil

Metals/metalloids
  Arsenic                                 20          750
  Cadmium                                  3           60
  Copper                                  60         7500
  Lead                                   -            -
  Manganese                              -            -
  Nickel                                 -            -
Organics
  DDT+DDD+DDE                              2           50
  Benzo(a)pyrene                           1            5
PHC (constituents)
  >[C.sub.16]-[C.sub.35] aromatics       -            -
  >[C.sub.9] PAH                           5          200
  >[C.sub.16]-[C.sub.35] aliphatic       -            -
  >[C.sub.9] TPH                        1000        10000

Guidelines
                                     This study
                                     Soil sample
                                     NA

Metals/metalloids
  Arsenic                                  0
  Cadmium                                 0.6
  Copper                                  35
  Lead                                    40
  Manganese                              200
  Nickel                                   7
Organics
  DDT+DDD+DDE                              0
  Benzo(a)pyrene                           0
PHC (constituents)
  >[C.sub.16]-[C.sub.35] aromatics
  >[C.sub.9] PAH                           0
  >[C.sub.16]-[C.sub.35] aliphatic      5503
  >[C.sub.9] TPH                        5804
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Author:Sheppard, Petra J.; Adetutu, Eric M.; Makadia, Tanvi H.; Ball, Andrew S.
Publication:Soil Research
Article Type:Report
Geographic Code:8AUST
Date:May 1, 2011
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