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College of Science and Management, University of Northern British Columbia, 3333 University Way, Prince George, BC, Canada V2N 4Z9
* Corresponding author (rutherfm{at}unbc.ca)
Received for publication June 23, 2005.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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Abbreviations: CCME, Canadian Council of Ministers of the Environment • F2, nC10–nC16 petroleum hydrocarbon fraction • F3, nC16–nC34 petroleum hydrocarbon fraction • PHC, petroleum hydrocarbon
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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The establishment and growth of plants in PHC-contaminated soils can be challenging for a number of reasons. Petroleum hydrocarbons and associated contaminants (e.g., salts or metals) may be toxic to plants, reducing plant germination, emergence, or vigor (Adam and Duncan, 2002; Bizecki-Robson et al., 2004; Rutherford et al., 2005). In addition to direct toxicity, the presence of PHCs may disrupt soil biological and biogeochemical properties and may contribute to soil structural degradation and increased hydrophobic tendencies (Kuperman et al., 2002; McGill et al., 1981; Roy et al., 1999; Visser et al., 2003). Nutrient limitations can result from microbial biomass production through decomposition of PHC compounds. In some cases, relatively high concentrations of available C in PHC-contaminated soils may lead to net N immobilization as soil organisms utilize contaminant C and mineral N for growth and metabolism (Xu et al., 1995; Xu and Johnson, 1997). Phosphorus may also be limiting in PHC-contaminated soils. Addition of inorganic fertilizers and/or organic amendments can been used to help overcome nutrient deficiencies in PHC-contaminated soils (Brook et al., 2001; Juteau et al., 2003; Pichtel and Liskanen, 2001; Sadowsky and Turco, 1999; Schwab and Banks, 1999; Williams et al., 1999).
Despite many reports of the benefits of biosolids addition to non-contaminated soils, few reports exist on the utilization of biosolids during phytoremediation of PHC-contaminated soils. It is expected that biosolids addition would prove beneficial for several reasons. In addition to being a readily available resource, biosolids have relatively high N and P content, low C to N ratio, and exhibit slow nutrient release properties (e.g., due to net N mineralization) (Barbarick and Ippolito, 2000; Pierzynski et al., 2000). Biosolids can also produce other improvements in soil chemical and physical properties such as decreased bulk density and increased cation exchange capacity, porosity, and water-holding capacity (Aggelides and Londra, 2000; Lindsay and Logan, 1998; Wong and Ho, 1991). Organic amendments have been found to reduce the toxicity of PHCs to grasses grown in diesel-contaminated soil (Vouillamoz and Milke, 2001). Some authors have reported on the influence of biosolids addition on PHC removal during bioremediation of non-vegetated soils. For example, Juteau et al. (2003) found that biosolids addition (activated sludge) resulted in greater removal of alkanes from non-vegetated soil contaminated with oily sludge as compared to soil treatments amended with ammonium nitrate or sterile sludge filtrate. Rivera-Espinoza and Dendooven (2004) found that biosolids addition accelerated the decomposition rate of diesel and total petroleum hydrocarbons in non-vegetated, laboratory contaminated soil, but did not increase the extent of degradation.
Depending on the source, biosolids may contain elevated concentrations of pathogens, trace elements, and/or trace organic contaminants (O'Connor et al., 2005). Consequently, land application of biosolids in the United States is regulated by federal and state government agencies (Sims and Pierzynski, 2000). In Canada, biosolids quality standards, and the agricultural use of biosolids, are regulated by the federal Fertilizers Act, and by provincial government agencies.
This 32-wk greenhouse study was initiated to investigate the influence of biosolids addition and smooth brome establishment on the rate and extent of PHC degradation in diesel-contaminated soil. It was hypothesized that the addition of biosolids would stimulate plant growth and microbial activity such that degradation of diesel PHCs would be enhanced over unamended controls. Another goal of the research was to investigate the influence of biosolids addition on plant N uptake and mineral N dynamics within the contaminated soil.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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Anaerobically digested sewage sludge (biosolids) from the Prince George (British Columbia, Canada) Wastewater Treatment Centre was collected on 29 Aug. 2003 after belt-pressing to remove excess water. The biosolids were sampled at a depth of approximately 50 to 100 cm from a large holding pile and were stored in sealed plastic buckets at 4°C until needed, at which time they were homogenized by hand mixing before addition to the soil. Preliminary work showed that total elemental concentrations of Ca, Mg, K, Na, and P were 32.4, 4.1, 0.86, 0.54, and 17.1 g kg–1 oven-dry biosolids, respectively. Available (Bray) P was 1.0 g kg–1 oven-dry biosolids. Trace element concentrations were 4.9, 4, 3, 36, 2760, 68, 5.9, 16, 25, 10, and 839 mg kg–1 oven-dry biosolids for As, Cd, Co, Cr, Cu, Pb, Hg, Mo, Ni, Se, and Zn, respectively.
Experimental Setup and Greenhouse Conditions
Smooth brome (cv. Carlton) seeds were obtained from Fosters Seeds in Beaverlodge, Alberta, Canada. The experimental design consisted of six treatments with two factors: vegetation (planted and non-planted) and biosolids addition at rates of zero (0), 13.34 g oven-dry biosolids kg–1 oven-dry soil (low), and 26.68 g oven-dry biosolids kg–1 oven-dry soil (high). Each of these treatments was replicated four times. Sampling dates corresponded to 8, 16, 24, and 32 wk after the onset of the experiment with an initial analysis done on all applicable measurements at time zero. A total of 96 experimental units were placed in a completely randomized design in the greenhouse. Day temperature was kept at 25°C with a 16-h light period (400-W high pressure sodium supplemental lighting used); night temperature was 15°C.
For each experimental unit, a total of 2.72 kg fresh soil, equivalent to 2.50 kg oven-dry soil, was added to plastic bags along with the appropriate amount of moist biosolids. The biosolids were then thoroughly mixed with the soil in each bag by hand. Unamended soils were mixed in the same way as biosolids-amended soils to assure similar abiotic losses of volatiles (e.g., PHCs and NH3) in amended and unamended treatments. Soil or soil-plus-biosolids were added to 3.7-L plastic pots (18-cm height by 17-cm diameter). Twenty-five seeds were placed at a 1-cm depth in the appropriate treatment pots to give approximately 1 seed per 7 cm2. Water content for each pot was brought up to approximately 80% water-holding capacity by daily waterings (with tap water).
The biosolids application rates corresponded to N additions of 726 mg total N kg–1 oven-dry soil and 1452 mg of total N kg–1 oven-dry soil for the low- and high-amendment treatments, respectively. These rates were estimated to meet (low-amendment) or exceed (high-amendment) the 80 kg N ha–1 plant requirements for the species and region (Zebarth et al., 2000) as well as to achieve a 25:1 contaminant C to N amendment ratio (to meet microbial requirements), based on the initial mineral N content of the biosolids (before addition to soil) and an estimated first year net N mineralization rate of 20% of organic N added by biosolids. Contaminant C content was estimated by the difference in total C content in non-solvent extracted versus solvent (1:1 acetone to hexane) extracted soils. It was assumed that net N mineralization from the native soil organic matter was negligible due to the low total N content of the soil (0.41 g N kg–1 oven-dry soil).
Sample Processing
At each sampling date, four randomly selected pots from each treatment were destructively sampled for selected analyses. The soil in each pot was divided in half using shoot placement as a guide to ensure that an equal number of plants occurred on both halves. One half of the total soil was only used for root biomass determination via root washing, while the other half was homogenized and subsampled for selected soil and root (e.g., N content) analyses. Non-vegetated pots were processed and handled in the same way as vegetated treatments to minimize differences in potential abiotic PHC losses between treatments. Soil samples were passed through a 2-mm sieve (10 mesh) before most chemical analyses (except PHC determination). Roots were handpicked from soil subsamples that were used for soil analyses (e.g., PHC extraction); these soil samples had <1% root biomass (by mass) before use for various soil analyses. Roots that were removed during sieving (10 mesh), and manually through handpicking, were gently washed with deionized water (to remove adhering soil) and were air-dried before N analysis. The PHC analysis on root tissue showed negligible concentrations of extractable PHCs.
Plant Biomass
Shoot and root biomass were determined at each destructive sampling date (8, 16, 24, and 32 wk). Due to the rapid growth of the smooth brome in the biosolids-amended treatments, intermediate, nondestructive harvests of shoots were also performed; this biomass was added to the total shoot biomass for that sampling period. To ensure uniformity during intermediate harvests, shoots were trimmed to the same height in both the low- and high-amendment treatments; that is, approximately to the height of the non-amendment plants (not harvested intermediately). At specified sampling dates, shoots were trimmed at the soil surface and placed into pre-weighed paper bags. The bags were then placed in a drying oven at 70°C (to constant weight) before weighing for biomass determination. Root biomass (only) was determined from half of the total soil in each pot using a root washing procedure. The root-soil mass was soaked overnight in dilute soapy water (10 mL phosphate-free dish soap per 1 L tap water) before the resulting mixture was run through a 500-µm sieve; roots were further cleaned by hand, rinsed with tap water, and placed in pre-weighed paper bags. The root biomass for half of the pot was obtained as described for the shoot biomass above; this value was then multiplied by 2 to give the total root biomass reported for the whole pot.
Analysis of Soil, Biosolids, and Plant Tissue
Soil water-holding capacity was determined by measuring the gravimetric moisture content of a column (15-cm length x 4.5-cm i.d.) of <5-mm soil (or soil-plus-biosolids mixtures) after saturating with water (overnight) and allowing to drain freely for 24 h under conditions that minimized evaporative moisture losses. Gravimetric moisture content was determined by drying soil overnight at 105°C. The following soil analyses were conducted on either <2-mm sieved soil samples (or soil plus biosolids mixtures) or on non-sieved biosolids samples. Soil pH was measured in a 1:4 soil to deionized H2O suspension (Hendershot et al., 1993a). Electrical conductivity of soil was determined in saturated paste extracts using a Model 3100 conductivity meter (YSI, Yellow Springs, OH) (Janzen, 1993). Effective cation exchange capacity was determined by the summation method in 0.1 M BaCl2 extracts, using inductively coupled plasma–atomic emission spectroscopy (ICP–AES) (ARL 3560; Thermo Electron, Waltham, MA) (Hendershot et al., 1993b). Available N (NH4+–N and NO3––N) was determined by extraction with 0.5 M K2SO4 (1:5 soil to solution ratio), followed by colorimetric N determination using an Alpkem Flow System IV Auto Analyzer (OI Analytical, College Station, TX).
Total C and N in soil and biosolids, and total N of plant tissue (shoots or roots), were determined on <100-mesh samples (air-dried, then ground in a Model MM200 ball mill; Retsch, Haan, Germany) by dry combustion using a Model 1500 NC Elemental Analyzer (Fisons, Milan, Italy).
Petroleum Hydrocarbon Analysis
The Canadian Council of Ministers of the Environment (CCME) has developed Canada-Wide Standards for PHCs in soils. These Tier 1 standards are grouped according to the equivalent normal straight-chain hydrocarbon (nC) boiling point ranges and can be defined by four fractions (Canadian Council of Ministers of the Environment, 2001a, 2001b). Fraction 1 (F1) ranges from nC6 to nC10, Fraction 2 (F2) from nC10 to nC16, Fraction 3 (F3) from nC16 to nC34, and Fraction 4 (F4) from nC34 to nC50. Fractionation in such a fashion is useful because complex mixtures of PHCs may contain hundreds to thousands of individual compounds; compounds within CCME fractions generally exhibit similar chemical, physical, and toxicological behaviors (Canadian Council of Ministers of the Environment, 2000).
Petroleum hydrocarbons for fractions F2 (nC10–nC16) and F3 (nC16–nC34) were determined according to the CCME's Reference Method for the Canada-wide Standard for Petroleum Hydrocarbons in Soil—Tier 1 Method (Canadian Council of Ministers of the Environment, 2001b). Preliminary work showed that F1 and F4 fractions were negligible and could be ignored in this study.
Briefly, PHCs were extracted from <5-mm soil using a Soxhlet extraction method with a 1:1 hexane to acetone (by volume) mixture (Canadian Council of Ministers of the Environment, 2001b). Twenty-five grams of soil (wet weight) was mixed with 2 g of diatomaceous earth and extracted for 23 h at four cycles per hour. The resulting extractant was made up to 250 mL with the hexane to acetone solvent. A column cleanup procedure was applied to a 50-mL subsample of the resulting extractant to remove polar organic compounds (Canadian Council of Ministers of the Environment, 2001b). The extract was concentrated into 1 mL toluene; this mixture was quantitatively transferred to a gas chromatography vial and made up to 2.00 mL with cyclohexane. The gas chromatography vial was capped with a Teflon-lined lid and stored at 4°C until analysis.
Solvent extracts were analyzed on a Model CP 3800 gas chromatograph equipped with a flame ionization detector (Varian, Palo Alto, CA). The column used was a 30-m MXT-1 with a 0.53-mm internal diameter and a 0.25-µm film thickness (Restek, Bellefonte, PA). A 1.0-µL volume was injected via a Varian CP 8400 Auto Sampler into a split/splitless injection port (Model 1079 PTV) held at 120°C for 2 min. The injector port was then increased at 200°C min–1 to 350°C, held for 2 min and was then decreased to 250°C and held for 5 min before cooling to the start temperature. The split ratio (5:1), initially off, was activated after 1 min. A pressure pulse injection technique was used to increase the amount of sample on the column. Pulse pressure was about 69 kPa (10.0 psi) (0.25-min duration). The column was initially held at 35°C for 4 min and was then increased at 15.0°C min–1 to 330°C, and held for 10 min; total run time was 34 min per sample. Helium carrier gas flow was 7.5 mL min–1. The flame ionization detector was kept at 340°C.
Peak retention times (i.e., peak maximums) of the external standards decane (nC10), hexadecane (nC16), and tetratriacontane (nC34) dissolved in 50:50 cyclohexane to toluene (final concentrations of 62.5, 125, and 250 µg mL–1) were used to identify CCME F2 and F3 regions on sample gas chromatography–flame ionization detector chromatograms; samples were run concurrently with the standards (Canadian Council of Ministers of the Environment, 2001b). Peak areas were integrated within the F2 and F3 regions and concentrations were calculated using the average response factor obtained from the following external standards (Sigma-Aldrich, St. Louis, MO) each dissolved in the above solvents and run at three concentrations (62.5, 125, and 250 µg mL–1): hexadecane (nC16), nonadecane (nC19), eicosane (nC20), dotriacontane (nC32), and tetratriacontane (nC34). Decane was not included in the calculation of the average response factors because this standard eluted several minutes before the first PHCs eluted from sample extracts.
Statistical Analysis
Statistical analysis was performed using Statistica Version 6.1 software (StatSoft, 2003) with reference to Siegal (1956) and Sokal and Rohlf (1995). After initial exploratory data analysis, it was determined that nonparametric statistics were required since homogeneity of variance and normality of data were not met. All significant results were found at the 
0.05 level. Plant data (biomass and plant N within each sampling date) and initial soil properties data were analyzed using the nonparametric Kruskal–Wallis one-way ANOVA (Siegal, 1956). Within subsequent sampling dates, soil properties data and % PHC remaining were analyzed using the two-way ANOVA (Scheirer–Ray–Hare extension of the Kruskal–Wallis test) on ranked data (Sokal and Rohlf, 1995). Interaction effects were also tested. Where statistical (p 0.05) significance was found, a Mann–Whitney U test (Siegal, 1956) was performed to test pair-wise comparisons of ranks.
A modified first-order decay equation of Nocentini et al. (2000) was used to fit degradation rates of PHC fractions:
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Nonlinear regression was used to fit the model to the PHC data. The coefficient of multiple determination (R2) for each treatment was calculated according to Bailey and McGill (2001) by the equation:
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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0.05) increased the total C, total N, NH4+–N, pH, electrical conductivity, and water-holding capacity of the soil, but did not significantly change soil NO3––N concentrations (Table 1). The soil showed hydrophobic tendencies and was quite slow to wet at the onset of the experiment. The addition of biosolids at both rates appeared to reduce this tendency and increased the apparent wettability (i.e., ease of wetting) of the soil as compared to the unamended treatment. Cation exchange capacity at 32 wk (data not shown) was significantly (p 0.05) greater in amended soils (average of low- and high-amendment treatments: 11.9 cmolc kg–1 soil) than unamended soils (averaged 10.2 cmolc kg–1 soil).
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0.05) increased shoot and root biomass at every sampling date; total biomass was greatest in the high-amendment treatment (Table 2). Plant N in shoots and roots was significantly (p 0.05) increased by biosolids addition as compared to the unamended treatment. By 32 wk, total plant N (shoot N + root N) for low- and high-amendment treatments was 75-fold greater and 135-fold greater than the unamended treatment, respectively. Maximum % recovery of biosolids-derived N into plants occurred at 32 wk, with significantly greater recovery occurring in the low-amendment rate (26.4%) as compared to the high-amendment rate (23.9%) (Table 2).
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0.05) interaction effect between the vegetation and amendment treatments was found for NH4+–N at 8 wk. Vegetated treatments had a greater initial decrease and had significantly (p 0.05) less total mineral N and NH4+–N as compared to non-vegetated pots after 8 wk. Nitrate N in the non-vegetated, biosolids-amended soils increased between 0 and 8 wk and remained significantly (p 0.05) greater in the non-vegetated treatments for the remainder of the 32-wk study.
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0.05) greater total soil N as compared to unamended treatments and total soil N in all treatments remained relatively constant throughout the 32-wk study (Table 3). Although total soil N did not show a significant vegetation effect, there was a tendency for the vegetated treatments to have lower concentrations than non-vegetated treatments.
Petroleum Hydrocarbon Degradation
Initial PHC concentrations were 1406, 2086, and 3492 mg kg–1 oven-dry soil for F2, F3, and total PHC (= F2 + F3), respectively. The greatest decline in PHC concentrations occurred during the first 8 wk of the experiment, where the vegetated, low-amendment treatment showed the greatest decrease (86% for F2; 50% for F3) (Table 4). By 32 wk, 6 to 20% of F2 remained, and there was no significant effect due to amendment. At this time, 30 to 49% of F3 remained, and significantly (p 0.05) less F3 remained in the unamended treatment as compared to the high-amendment treatment (Table 4). There was no significant difference in F3 remaining between the unamended and low-amendment treatments, or between the low- and high-amendment treatments. Within biosolids-amended treatments, vegetated treatments had significantly (p 0.05) lower F2, F3, and total PHC remaining at 32 wk as compared to nonvegetated treatments.
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0.05) greater than rate constants for unamended treatments. The low-amendment treatments tended to have a greater F3 rate constant than both the unamended and high-amendment treatments, but these trends were not significant. Likewise, the residual fraction (y0) tended to be lower in planted treatments as compared to non-planted treatments, but these trends were not significant.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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The addition of biosolids-derived mineral N, and potentially mineralizable N, likely played an important role in stimulating plant growth and plant N uptake. Petroleum hydrocarbon contaminated soils often are low in plant available N due to net immobilization of mineral N as PHC degrading organisms utilize contaminant C and soil mineral N for growth and metabolism (Xu et al., 1995; Xu and Johnson, 1997). In this study, low concentrations of mineral N in the amended, vegetated treatments suggests that mineralizable N from the biosolids was being used by plants as quickly as it was becoming available. Although total mineral N concentrations remained below 1 mg kg–1 soil in the unamended treatments, total mineral N concentrations in the amended, non-vegetated treatments exceeded 40 mg kg–1 soil throughout the entire study. This suggests that the supply of mineral N in the amended, non-planted treatments exceeded microbial demand. The fact that NO3––N increased with time in amended, non-vegetated treatments shows that nitrifying bacteria were not inhibited by the presence of PHCs in this soil. Others have indicated that nitrifiers may be sensitive to PHCs in contaminated soils (Xu et al., 1995).
The recoveries of biosolids-derived N by smooth brome in the low- and high-amendment treatments of this study (23.9–26.4% at 32 wk) are in the range typically reported for non-contaminated soils suggesting that plant N uptake was unaffected by PHC contamination. Nitrogen recovery by forage grasses from biosolids amendments in non-contaminated field soils typically ranges from 14 to 40%, depending on soil properties, field conditions, and types of biosolids used (Cogger et al., 1999; Kiemnec et al., 1987; Smith and Tibbett, 2004; Sullivan et al., 1997; Warman, 1986). It should be noted that the recoveries of biosolids-derived N in this study are based on the initial N content of biosolids that were mixed into the soil. Recoveries account for losses of volatile N (e.g., NH3) that may have occurred during addition of biosolids to soil.
Modifying the simple first-order decay equation to include the parameter y0, the residual PHC fraction present in the soil (Nocentini et al., 2000), yielded a good fit to the PHC data (Table 5).
Examination of the degradation rate constants suggests that F2 hydrocarbons were degraded faster than F3 hydrocarbons (see Table 5). In general, rate constants for F2 (0.151–0.322 wk–1) were two to three times greater than those for F3 (0.073–0.219 wk–1). The lone exception occurred in the non-vegetated, low-amendment treatment where the degradation rate for F2 (0.274 wk–1) was slightly greater than that for F3 (0.219 wk–1). These trends generally agree with the work of Namkoong et al. (2002) who in a 30-d laboratory study found greater degradation rates for a PHC fraction composed of short-chain normal alkanes (0.37–0.54 d–1 for nC10–nC15) as compared to a PHC fraction containing longer-chain normal alkanes (0.18–0.24 d–1 for nC16–nC20).
The greater degradation rate constant for F2 versus F3 in the current study is consistent with reports of crude oil or refinery sludge biodegradation in Canadian soils (Huang et al., 2005; Visser et al., 2003). A lesser degradation rate for the F3 fraction is expected, given the larger average size of the component molecules as compared to the F2 fraction. Also, the F3 fraction has a greater carbon-normalized sorption coefficient, Koc, than the F2 fraction (Canadian Council of Ministers of the Environment, 2000), which may contribute to stronger sorption (or partitioning) to native- or amendment-derived soil organic carbon. Strong sorption processes may limit PHC bioavailability to hydrocarbon degraders in soil (Alexander, 2000; Rogers, 1996; Semple et al., 2003).
The low-amendment treatment significantly increased the degradation rate of the total PHC in this study by approximately twofold over the untreated control (Table 5). A laboratory study reported by Namkoong et al. (2002) observed that the addition of sewage sludge to a spiked, diesel-contaminated soil at a ratio of 1:0.1 (wet-weight) soil to sewage sludge (approximately equivalent to high biosolids rate in this study) also increased the degradation rate of PHCs almost twofold over the non-treated control. Whereas Namkoong et al. (2002) observed a degradation rate of 0.505 wk–1 we observed a degradation rate of 0.142 wk–1 in the non-vegetated high-amendment treatment of this study. Differences in experimental conditions and starting materials are most likely responsible for the different degradation rates. That is, degradation rates in recently spiked soils, such as that of Namkoong et al. (2002), are expected to be higher than in aged or weathered soils due to greater bioavailability of compounds (Alexander, 2000; Loehr and Webster, 1996). Brook et al. (2001) examined the bioremediation of PHCs in a weathered, diesel-contaminated soil (2.2% PHC by dry weight) and found degradation rates closer to those found in this study. When inorganic N was added to the soil to achieve a C to N ratio of 20:1, PHC degradation rate values corresponding to 0.027 to 0.224 wk–1 were observed over 60 d depending on the N source applied. Degradation rate values were 1.5 to 12 times greater than those for the untreated control.
Although the low-amendment rate resulted in a greater degradation rate of total PHCs, the extent of PHC removal by 32 wk was not increased over that of the unamended treatments (Tables 4 and 5). These findings are similar to Juteau et al. (2003) and Rivera-Espinoza and Dendooven (2004), who reported that biosolids addition increased the rate, but not the extent, of PHC removal in non-vegetated, diesel-contaminated soils.
Though there was no significant effect of vegetation on the degradation rate constant, k, the y0 values in Table 5 would suggest that vegetated treatments did decrease the amount of residual F2, F3, and total PHC remaining at 32 wk. This is supported by data in Table 4, where vegetated treatments in the biosolids-amended soils had significantly (p 0.05) lower concentrations of PHC fractions than non-vegetated treatments at the end of the study. Several factors likely contributed to these trends. First, the presence of plant roots may have stimulated rhizodegradation of diesel-derived PHCs in biosolid-amended soils, resulting in more extensive removal (as compared to non-vegetated treatment) by 32 wk. A rhizosphere effect may have included processes that stimulated the degradative activity of PHC degrading organisms and/or processes that increased the bioavailability (e.g., facilitated desorption) of diesel PHCs to degrading organisms (Siciliano and Germida, 1998; Singer et al., 2003).
A second factor that likely contributed to lower residual PHC concentrations in vegetated versus non-vegetated soils receiving biosolids is a biosolids-derived interference in the F3 fraction. The chromatogram for the biosolids-amended soil at time 0 (Fig. 1A) shows a small peak in the F3 window that was not observable in unamended soils (chromatogram not shown). Initial concentrations of F3 (and F2) in amended soils were not significantly greater than unamended soils. By the end of the study, the biosolid-derived contribution was not very visible in vegetated soils (Fig. 1B), but was still present in non-planted soils (Fig. 1C). Close examination of all chromatograms obtained for 32 wk samples showed that the presence of plants tended to result in reduced diesel- and biosolid-derived components, relative to non-planted soils. The reduction in biosolid-derived components was often more apparent than that for the diesel-derived components. Perhaps root-induced microbial activity stimulated the decomposition or transformation of biosolids components, thereby reducing the solvent-extractable component. The biosolids amendment rates in this study were low (1–3% of soil, dry mass basis). The challenges of quantifying contaminants in soil or other media containing high organic matter content are problematic and are discussed in Canadian Council of Ministers of the Environment (2001b) and Rogers (1996).
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CONCLUSIONS |
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ACKNOWLEDGMENTS |
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