Article pubs.acs.org/JAFC
Novel Biochar-Plant Tandem Approach for Remediating Hexachlorobenzene Contaminated Soils: Proof-of-Concept and New Insight into the Rhizosphere Yang Song,*,† Yang Li,† Wei Zhang,‡,§ Fang Wang,*,†,‡ Yongrong Bian,† Lisa A. Boughner,‡ and Xin Jiang*,† †
State Key Laboratory of Soil and Sustainable Agriculture, Institute of Soil Science, Chinese Academy of Sciences, 71 East Beijing Road, Nanjing 210008, PR China ‡ Department of Plant, Soil and Microbial Sciences, and §Environmental Science and Policy Program, Michigan State University, East Lansing, Michigan 48824, United States S Supporting Information *
ABSTRACT: Volatilization of semi/volatile persistent organic pollutants (POPs) from soils is a major source of global POPs emission. This proof-of-concept study investigated a novel biochar-plant tandem approach to effectively immobilize and then degrade POPs in soils using hexachlorobenzene (HCB) as a model POP and ryegrass (Lolium perenne L.) as a model plant growing in soils amended with wheat straw biochar. HCB dissipation was significantly enhanced in the rhizosphere and near rhizosphere soils, with the greatest dissipation in the 2 mm near rhizosphere. This enhanced HCB dissipation likely resulted from (i) increased bioavailability of immobilized HCB and (ii) enhanced microbial activities, both of which were induced by ryegrass root exudates. As a major component of ryegrass root exudates, oxalic acid suppressed HCB sorption to biochar and stimulated HCB desorption from biochar and biochar-amended soils, thus increasing the bioavailability of HCB. High-throughput sequencing results revealed that the 2 mm near rhizosphere soil showed the lowest bacterial diversity due to the increased abundance of some genera (e.g., Azohydromonas, Pseudomonas, Fluviicola, and Sporocytophaga). These bacteria were likely responsible for the enhanced degradation of HCB as their abundance was exponentially correlated with HCB dissipation. The results from this study suggest that the biochar-plant tandem approach could be an effective strategy for remediating soils contaminated with semi/volatile organic contaminants. KEYWORDS: persistent organic pollutants, bioavailability, microbial community structure, soil rhizosphere, biochar
■
INTRODUCTION Persistent organic pollutants (POPs) include a broad range of toxic, persistent, and bioaccumulative contaminants capable of being transported over a long-range.1 One important group of POPs are organochlorinated pesticides, such as hexachlorobenzene (HCB) and 2,2-bis(4-Chlorophenyl)-1,1,1-trichloroethane (DDT), that were primarily used on agricultural soils of the world before their use was banned.2 Because of the volatile properties of these contaminants (e.g., log octanol/air partition coefficient of 7.2 and Henry’s law constant of 30 Pa m3 mol−1 for HCB), it is generally believed that POPs were transported over a long distance by atmospheric circulation from low to high latitudinal areas.3 In fact, many POPs including HCB have been detected in soils and animals from the most remote Arctic and Antarctic regions.4,5 The atmospheric HCB is thought to come predominantly through volatilization from contaminated soils.3,6 A conventional remediation technique for volatile organic contaminants is air stripping, which consists of pumping them out of the contaminanted soils.7 This strategy is costly and could also increase the release of volatile POPs to the atomosphere. Once these contaminants become airborne, they tend to migrate over large geographic regions and pose direct exposure risks to humans and other organisms.3 Since the rate and efficiency of HCB degradation is far greater in soils than the atmosphere,3 a novel alternative is to immobilize these © 2016 American Chemical Society
contaminants in soils, reducing their release to the atmosphere, and then recruit soil microorganisms to degrade them in situ. Recently, biochar has been increasingly considered a costeffective soil amendment for agronomic and environmental applications.8 Because of biochar’s large surface area and high microporosity, it typically has strong sorption capacities for many organic contaminants, including polycyclic aromatic hydrocarbons (PAHs) and pesticides.9,10 Amendment of contaminated soil with biochar has been reported to reduce the uptake and accumulation of POPs by both plants and earthworms.11,12 Similarly, our previous studies also demonstrated that wheat straw biochar could efficiently immobilize chlorobenzenes in soils, thus reducing their volatilization and uptake by earthworms.13,14 However, due to the strong sorption of POPs to biochar, their bioavailability to microorganisms may be decreased, reducing their degradation rates.15 Many perceive that this obstacle may be overcome with phytoremediation. This hypothesis is well supported by previous reports that planting either ryegrass, alfalfa, or celery enhanced the degradation of Received: Revised: Accepted: Published: 5464
March 3, 2016 May 17, 2016 June 21, 2016 June 21, 2016 DOI: 10.1021/acs.jafc.6b01035 J. Agric. Food Chem. 2016, 64, 5464−5471
Article
Journal of Agricultural and Food Chemistry
area (BET) of 4.8 m2 g−1. Detailed soil and biochar properties are provided in a previous publication.13 Rhizodegradation of HCB in Biochar-Amended Soil. To immobilize HCB in soil with biochar, multiple 5-kg soil batches were spiked with HCB and amended with 1% or 2% biochar by weight, respectively.13 The soil without biochar addition was used as the control. Hence, there were three treatments, i.e., control (0% biochar), 1% biochar, and 2% biochar, all in triplicate. The treated soils just described were incubated at 25 °C for 3 months in the dark, the aged soils were then air-dried, ground, and sieved through a 0.84 mm sieve before being used for planting experiments (Figure S1). A rhizobox as described in previous studies25 was used for planting ryegrass (Lolium perenne L.) in the HCB aged soils with or without biochar amendment. Briefly, as shown in Figure S1, the central zone (20 mm width) of the rectangular rhizobox was used to grow the ryegrass. On either side of the rhizosphere (R), the soils were separated into 1−5 mm near rhizosphere (NR) and >5 mm far rhizosphere (FR) zones with a nylon mesh of 5 mm FR soils) were analyzed by high-throughput sequencing as described later. Sorption and Desorption of HCB in Relation to Biochar As Affected by Root Exudates. A hydroponic experiment was first conducted to examine the main components of ryegrass root exudates in response to the HCB exposure. Briefly, the pregerminated ryegrass was exposured to HCB with a concentration of 0, 0.1, 0.5, 1, 2, or 5 mg L−1 in Hoagland solution. After 8 d, the ryegrass roots were cleaned and immersed for 2 h into 5 mL Milli-Q water in test tubes. Then, the root exudates containing solutions were collected and filtered with a 0.45-μm Millipore Millex (PES) filter and topped-off with Milli-Q water to a constant-volume of 5 mL. Total sugar and LMWOAs in root exudates were measured by spectrophotometer and high performace liquid chromatography, respectively. The detailed experimental procedure and detection conditions are provided in the Supporting Information (SI). Generally, with increasing HCB concentrations, the total amount of LMWOAs released from the ryegrass roots increased (Table S1). In all cases, the concentration of oxalic acid was significantly greater than that of the other LMWOAs (Table S1). There was no significant difference among total sugar concentrations in the ryegrass root exudates, with or without HCB. Batch sorption experiments were then performed in triplicate to elucidate the effect of root exudates on the sorption of HCB to biochar. Since oxalic acid was detected as the main component of ryegrass root exudates (Table S1), an oxalic acid solution of 0, 1, 10, 20, or 40 mg L−1 was used as the background solution. According to
POPs in biochar-free soils.16,17 The increased degradation of POPs largely resulted from their increased bioavailability and the enhanced microbial activities in the rhizosphere, both of which were induced by plant root exudates such as low molecular weight organic acids (LMWOAs).18,19 For instance, LMWOAs could enhance the release of dissolved organic matter (DOM) from soil particles to the solution phase, and then binding of PAHs with DOM would promote desorption of PAHs from soils.20 Increased desorption of POPs from soils could, in principle, increase their bioavailability. However, it remains unclear whether plant root exudates enhance the bioavailability of POPs in biochar-amended soils. Previously, it was reported that plant roots could grow into and break the biochar pore structure.21 Mineralization of biochar in soils was also accelerated by root exudates.22 Consequently, it is very likely that in biochar-amended soils, plant roots may facilitate the release of sorbed POPs into soil water, subsequently becoming accessible to POPs-degrading microbes. Moreover, plant root exudates could influence the microbial community by providing readily available carbon sources.19,23 Thus, this plant root−microbe interaction may play an important role in the degradation of POPs in the rhizosphere. However, in biochar-amended soils, considering the increased macro- and micronutrients (e.g., calcium, phosphorus, and potassium),8 the strong affinity of some nutrients to biochar, and the recalcitrant nature of biochar carbon, it is not clear how the presence of biochar in soils may modulate changes in the microbial community structure within the rhizosphere and nonrhizosphere soils.24 Therefore, the objectives of this study were to (i) investigate whether plants could enhance the dissipation of POPs after being immobilized by biochar in soils and (ii) elucidate the effects of biochar addition on the dissipation of POPs and the changes in the microbial community structures within the rhizosphere, near rhizosphere, and far rhizosphere soils. In this proof-of-concept study, we used the rhizobox experimental design with ryegrass as a model plant and HCB as a model POP. The bacterial community structure was analyzed by highthroughput sequencing. Investigations into the effects of ryegrass root exudates on the sorption of HCB by biochar and desorption of HCB from biochar-amended soil were conducted to better understand the influence of ryegrass on the dissipation of HCB in biochar-amended soil.
■
MATERIALS AND METHODS
Chemicals. The HCB standard (>99.5% purity) was purchased from Dr. Ehrenstorfer (Augsburg, Germany). Tenax TA (60−80 mesh), used for desorption experiments, was purchased from Kanglin Science & Technology Co. Ltd. (Beijing, China). Before use, the Tenax beads were rinsed with hexane/acetone (1/1 by volume) 3 times and then dried overnight at 75 °C. The solvents and all other chemical reagents, purchased from Nanjing Chemical Factory (Nanjing, China), were of analytical grade. Anhydrous sodium sulfate was oven-dried at 400 °C for 4 h. Soil Sampling and Biochar Preparation. An agricultural soil (classified as Ferri-Udic Argosols, according to Chinese Soil Taxonomy) was collected from the top 20 cm in a vegetable field, air-dried, gently ground, and then passed through a 2 mm sieve prior to being used in this study. The soil had a pH of 7.6, total carbon content of 3.1%, with a composition of 13.6% clay, 63.1% silt, and 23.3% sand. A wheat straw biochar was produced under oxygenlimited conditions at 500 °C as previously described.13 The produced biochar had a pH of 10.5; total carbon content of 48.5%; dissolved organic carbon (DOC) content of 646 mg kg−1; and a specific surface 5465
DOI: 10.1021/acs.jafc.6b01035 J. Agric. Food Chem. 2016, 64, 5464−5471
Article
Journal of Agricultural and Food Chemistry our previously developed method,13 HCB sorption isotherms were measured in oxalic acid background solutions at the initial HCB concentrations of 0.1−2 mg L−1. The detailed experimental procedure is provided in the SI. The measured HCB sorption isotherms were then fitted with the linear Henry model as Q e = Kd × Ce
MOTHUR v.1.30.1 (http://www.mothur.org/). The principal coordinates analysis (PCoA) was performed according to Bray−Curtis dissimilarity metrics. Heatmap analysis, based on vegdist and hclust, was conducted in R v.3.2.1 with the vegan package.29 Quality Control and Statistical Analysis. To estimate the HCB loss during sorption experiments, sorbent-free blank samples were included in the sorption experiments. The average recovery of HCB from blank samples was 96.2 ± 2.1%. The recoveries of HCB from spiked soil and ryegrass samples were determined to be 91.6 ± 4.3% and 92.4 ± 3.7%, respectively (SI). Data were statistically analyzed using analysis of variance (ANOVA) and the least significant difference (LSD) posthoc comparison tests with SPSS V17.0 at a p < 0.05 significance level.
(1)
where Qe (mg kg−1) is the sorbed HCB concentration on biochar, Ce (mg L−1) is the HCB equilibrium concentration in solution, and Kd (L kg−1) is the distribution coefficient. Another parallel sorption experiment was conducted to detect the DOC concentrations in the equilibrium solutions. The effect of oxalic acid on the desorption of HCB from biochar was determined after HCB had been sorbed to biochar without oxalic acid. The detailed experimental procedure is provided in the SI. Desorption of HCB in Biochar-Amended Soil with Tenax Extraction. To elucidate the effect of oxalic acid on the bioavailability of HCB in biochar-amended soil, Tenax extraction based desorption experiments were conducted in triplicate according to the previously described method.26 Desorption of HCB from the soils was measured after their exposure to oxalic acid solutions of 0, 1, 10, 20, or 40 mg L−1 for 6-h or 400-h, as described in detail in the SI. The Tenax 6-h and 400-h extractions were used to measure both the rapid and slow desorption fractions of soil contaminants, respectively.26 Bacterial Community Structure in Soils. The rhizosphere, 2 mm near rhizosphere and >5 mm far rhizosphere soils were subjected to microbial analyses at the end of the rhizobox experiment (Figure S1). DNA was extracted from soil samples using the E.Z.N.A. Soil DNA Kit (OMEGA, USA) following the protocols recommended by the manufacturer. DNA concentration was measured with a Nanodrop (Thermo Scientific), and quality was assessed by agarose gel electrophoresis. The V4−V5 bacterial 16S rRNA gene regions were amplified by PCR (95 °C for 2 min, followed by 25 cycles at 95 °C for 30 s, 55 °C for 30 s, and 72 °C for 30 s, and a final extension at 72 °C for 5 min) using primers 338F 5′-barcode-ACTCCTACGGGAGGCAGCA-3′ and 806R 5′- GGACTACHVGGGTWTCTAAT-3′, where “barcode” is an eight-base sequence unique to each sample.27 PCR reactions were performed in triplicate in 20 μL mixtures containing 4 μL of 5 × FastPfu Buffer (Sangon Biotech, Shanghai, China), 2 μL of 2.5 mM dNTPs, 0.8 μL of each primer (5 μM), 0.4 μL of FastPfu polymerase (2.5 U/μL, TransGen Biotech Co., Ltd., Beijing, China), and 10 ng of template DNA. Amplicons were purified using the AxyPrep DNA Gel Extraction Kit (Axygen Biosciences, Union City, CA, USA) following the manufacturer’s instructions and quantified using Quanti Fluor-ST (Promega, USA). The purified amplicons were then combined in equimolar concentrations, and Illumina adapters were added by ligation (TruSeq DNA LT Sample Prep Kit). The adapter-ligated fragments were further amplified with 10 cycles to obtain a sufficient yield for sequencing and paired-end sequenced (2 × 250) on an Illumina MiSeq platform by Majorbio Bio-Pharm Technology Co., Ltd. (Shanghai, China). The raw reads were deposited into the NCBI Sequence Read Archive (SRA) database (Accession Number: SRP065866). Raw fastq files were demultiplexed, quality-filtered using QIIME (version 1.17) with the following criteria: (i) The 250 bp reads were truncated at any site receiving an average quality score 0.05). The HCB concentration in the ryegrass root was significantly higher than that in the ryegrass shoot, regardless of treatment (Figure 2). The ryegrass root-to-
Figure 2. HCB concentrations in the shoots and roots of ryegrass grown in soils with and without biochar amendment. Different lowercase (shoot) and capital letters (root) indicate the significant differences among the treatments at p < 0.05.
shoot translocation factor for the control, 1% biochar, and 2% biochar treatments was 0.008, 0.077, and 0.079, respectively. Therefore, HCB was preferentially bioaccumulated in the ryegrass root resulting in low root-to-shoot translocation factors. Moreover, the presence of biochar significantly reduced the uptake of HCB by ryegrass root (Figure 2). On the basis of mass balance calculations (Table S2), after a 6-month growing period, the dissipation percentages of HCB in R soils were 56.11%, 46.56%, and 37.18% for the control, 1% biochar, and 2% biochar treatments, respectively. Whereas the dissipation percentages of HCB in FR soils were 24.48%, 16.59%, and 8.10% for the control, 1% biochar, and 2% biochar treatments, respectively. Therefore, compared to FR soils, the dissipation percentage of HCB in R soils was increased by 31.63%, 29.96%, and 29.08% for the control, 1% biochar, and 2% biochar treatments, respectively. These observations indicate that the rhizodegradation of HCB could be efficient in biochar-amended soils. Because the amount of HCB in the shoot and root was less than 1.1% of the initial HCB amount (Table S2), the plant uptake was not a dominant factor for the removal of the HCB in the R soils. Intriguingly, the lowest concentration of HCB was not measured in R soil, but in the NR 2 mm away from the ryegrass root in all treatments, indicating that the fastest dissipation of 5467
DOI: 10.1021/acs.jafc.6b01035 J. Agric. Food Chem. 2016, 64, 5464−5471
Article
Journal of Agricultural and Food Chemistry
oxalic acid significantly enhanced the 6-h desorption of HCB (p < 0.05), but no significant desorption enhancement was observed for oxalic acid concentrations beyond 10 mg L−1 (p > 0.05). Increased desorption of HCB by oxalic acid was in agreement with the enhancing effect of oxalic acid on HCB desorption from pure biochar as discussed above (Figure 3), again likely resulting from the enhanced release of DOC from biochar-amended soils. Similarly enhanced desorption of some POPs, such as DDTs36 and PAHs,20 by LMWOAs from biochar-free soils has already been reported. Of notable interest, oxalic acid did not enhance the 400-h desorption of HCB, implying that root exudates, such as LMWOAs, could only desorb the bioavailable fraction of sorbed HCB. Since the average oxalic acid concentrations in rhizosphere soil water are in the range of 0.1−10 mg L−1,37 our results indicate that root exudates could increase the bioavailability of POPs under relevant field conditions. Bacterial Community Structure in Soil. Since bacteria have already been reported as the predominant degraders of POPs in soil,38 the sequencing of the rhizobox soil was conducted to determine the bacterial community structure. The amendment of soil with biochar increased the bacterial biodiversity as expressed by the Shannon index (Figure 5).
Figure 3. Sorption isotherms of HCB to biochars (a) and the desorption percentage of HCB from biochars (b), in the presence of varying concentrations of oxalic acid. In part a, the lines were fitted with the linear Henry’s model. In part b, the 1773 and 3073 mg kg−1 were the amounts of HCB that sorbed to biochar without the influence of oxalic acid. Different letters in b indicate the significant difference among treatments at p < 0.05.
Figure 5. Bacterial diversity expressed as Shannon index in the rhizosphere (R), near rhizosphere (NR), and far rhizosphere (FR) soils with and without biochar amendment. Different letters indicate the significant difference among treatments at p < 0.05.
The bacterial biodiversity was lower in the R soil than in the FR soil, regardless of treatments (Figure 5). Interestingly, the 2 mm NR soil showed the lowest bacterial diversity, where the fastest dissipation of HCB was observed (Figure 1b). The PCoA analysis showed that the bacterial communities in the R, NR, and FR soil samples were individually clustered together. The first two PCs explained a total of 69.1% variance of bacterial communities, with PC1 explaining 52.4% of the variance (Figure S3). A heatmap coupled with cluster analysis (Figure 6) showed that the bacterial community structure similarity among the R (Cluster 3) and NR (Cluster 2) was higher than that of the FR (Cluster 1). This observation indicates that planting ryegrass, rather than biochar addition, was the main factor controlling the similarity of bacterial community structure in soils. The release of root exudates may result in conditions optimal for certain kinds of bacteria that are able to establish ecological niches that are ideal for the dissipation of HCB in soils. At the genus-level, the relative abundances of Comamonadaceae-unclassif ied, Azohydromonas, and Pseudomonas were significantly higher in Cluster 2 and Cluster 3 (blue region) than those in Cluster 1, suggesting that
Figure 4. Desorption percentages of HCB from biochar-free and biochar-amended soils at varying oxalic acid concentrations. Different lowercase (6-h desorption) and capital letters (400-h desorption) indicate the significant difference among treatments at p < 0.05.
400-h desorption percentages in 1% and 2% biochar treatments were lower than those in the control treatment (p < 0.05). In all tested soils, the presence of 1 mg L−1 oxalic acid did not significantly affect the HCB 6-h desorption, relative to the oxalic acid-free treatment. However, the presence of 10 mg L−1 5468
DOI: 10.1021/acs.jafc.6b01035 J. Agric. Food Chem. 2016, 64, 5464−5471
Article
Journal of Agricultural and Food Chemistry
Figure 6. Heatmap of bacterial community structure in the rhizosphere (R; Cluster 3), near rhizosphere (NR; Cluster 2), and far rhizosphere (FR; Cluster 1) soils with and without biochar amendment. The boxed regions indicate certain genera with high relative abundance.
Betaproteobacteria and Arthrobacter may be the most important bacteria for the dissipation of HCB in these soils. The abundance of Betaproteobacteria in the NR under 1% biochar treatment was significantly higher than that in other treatments. Previously, it was reported that Betaproteobacteria dominate the bacterial community of PAH-contaminated soil.45 Also, Arthrobacter was highly competitive in the presence of root exudates46 and has the potential for degrading biphenyl and other hydrocarbons.47 It should be noted that the cooperative effort of a microbial community is usually more efficient than a single organism in biodegradation of soil contaminants. However, elucidating the community of microbes responsible for the dissipation of HCB needs further investigation, especially with the help of stable isotope probing technology. Environmental Implications. The findings of this study provide several interesting implications for remediating soils contaminated with semivolatile or volatile POPs such as HCB. In the biochar-plant tandem remediation scheme, biochar soil amendment could substantially immobilize HCB in soils and consequently reducing ryegrass root uptake of HCB. Considering that plant uptake and volatilization were minor in biochar-amended soils, microbial degradation was likely the main route of HCB dissipation. Ryegrass root exudates not only increased the bioavailability of sorbed HCB in biocharamended soils but also enriched it with potential HCBdegrading bacteria. Moreover, root exudates could increase the soil biota activity as reported elsewhere.18 Consequently, by
the root and/or root exudates stimulated the growth of these bacteria. Among these bacteria, Pseudomonas are considered the predominant group of soil microorganisms that biodegrade complex organic compounds, 23 such as phenol, PAH, tetrabromobisphenol A, etc.39−41 They could also use both root exudates and PAH as a carbon source.42 In this study, there was also a linear relationship between the percent loss of HCB and the log relative abundance of Pseudomonas in soils, with a correlation coefficient (r) of 0.79 (Figure S4 and Table S3). Therefore, Pseudomonas could be partly responsible for the dissipation of HCB in the R and NR soils. These significant linear relationships were also observed for Comamonadaceaeunclassif ied and Azohydromonas (Figure S4 and Table S3). As shown in Figure 6, the genera Ohtaekwangia, Lactococcus, Cellvibrio, Solibacillus, Fluviicola, Betaproteobacteria_unclassified, Sporocytophaga, possible_genus_04, Bacillus, Chryseolinea, and Arthrobacter from Cluster 2 (red region) were higher in their abundance than in Cluster 3 and Cluster 1, indicating that in the 2 mm NR soil these genera showed the highest relative abundances. Moreover, these same genera all showed linear relationships between the dissipation percentages of HCB and the log relative abundances for these bacteria, with an r value of 0.37−0.89 (Figure S4 and Table S3). Ohtaekwangia are copiotrophic and fast-growing bacteria, which mainly use the labile organic substance in the rhizosphere.43 Lactococcus could produce lactic acid as a byproduct of glucose fermentation and is able to degrade dinitrotoluene.44 Among these genera, 5469
DOI: 10.1021/acs.jafc.6b01035 J. Agric. Food Chem. 2016, 64, 5464−5471
Article
Journal of Agricultural and Food Chemistry
contaminant management in soil and water: A review. Chemosphere 2014, 99, 19−33. (10) Uchimiya, M.; Wartelle, L. H.; Boddu, V. M. Sorption of triazine and organophosphorus pesticides on soil and biochar. J. Agric. Food Chem. 2012, 60, 2989−2997. (11) Gomez-Eyles, J. L.; Sizmur, T.; Collins, C. D.; Hodson, M. E. Effects of biochar and the earthworm Eisenia fetida on the bioavailability of polycyclic aromatic hydrocarbons and potentially toxic elements. Environ. Pollut. 2011, 159, 616−622. (12) Yang, X. B.; Ying, G. G.; Peng, P. A.; Wang, L.; Zhao, J. L.; Zhang, L. J.; Yuan, P.; He, H. P. Influence of biochars on plant uptake and dissipation of two pesticides in an agricultural soil. J. Agric. Food Chem. 2010, 58, 7915−7921. (13) Song, Y.; Wang, F.; Bian, Y.; Kengara, F. O.; Jia, M.; Xie, Z.; Jiang, X. Bioavailability assessment of hexachlorobenzene in soil as affected by wheat straw biochar. J. Hazard. Mater. 2012, 217−218, 391−397. (14) Song, Y.; Wang, F.; Kengara, F. O.; Yang, X.; Gu, C.; Jiang, X. Immobilization of chlorobenzenes in soil using wheat straw biochar. J. Agric. Food Chem. 2013, 61, 4210−4217. (15) Jones, D. L.; Edwards-Jones, G.; Murphy, D. V. Biochar mediated alterations in herbicide breakdown and leaching in soil. Soil Biol. Biochem. 2011, 43, 804−813. (16) Wang, M. C.; Chen, Y. T.; Chen, S. H.; Chien, S. W. C.; Sunkara, S. V. Phytoremediation of pyrene contaminated soils amended with compost and planted with ryegrass and alfalfa. Chemosphere 2012, 87, 217−225. (17) Wenzel, W. W. Rhizosphere processes and management in plant-assisted bioremediation (phytoremediation) of soils. Plant Soil 2009, 321, 385−408. (18) Chen, K. J.; Zheng, Y. Q.; Kong, C. H.; Zhang, S. Z.; Li, J.; Liu, X. G. 2,4-dihydroxy-7-methoxy-1,4-benzoxazin-3-one (DIMBOA) and 6-methoxy-benzoxazolin-2-one (MBOA) levels in the wheat rhizosphere and their effect on the soil microbial community structure. J. Agric. Food Chem. 2010, 58, 12710−12716. (19) Lu, Y. C.; Zhang, S.; Miao, S. S.; Jiang, C.; Huang, M. T.; Liu, Y.; Yang, H. Enhanced degradation of herbicide isoproturon in wheat rhizosphere by salicylic acid. J. Agric. Food Chem. 2015, 63, 92−103. (20) Gao, Y.; Ren, L.; Ling, W.; Kang, F.; Zhu, X.; Sun, B. Effects of low-molecular-weight organic acids on sorption-desorption of phenanthrene in soils. Soil Sci. Soc. Am. J. 2010, 74, 51−59. (21) Joseph, S. D.; Camps-Arbestain, M.; Lin, Y.; Munroe, P.; Chia, C. H.; Hook, J.; van Zwieten, L.; Kimber, S.; Cowie, A.; Singh, B. P.; Lehmann, J.; Foidl, N.; Smernik, R. J.; Amonette, J. E. An investigation into the reactions of biochar in soil. Aust. J. Soil Res. 2010, 48, 501− 515. (22) Hamer, U.; Marschner, B.; Brodowski, S.; Amelung, W. Interactive priming of black carbon and glucose mineralisation. Org. Geochem. 2004, 35, 823−830. (23) Glick, B. R. Using soil bacteria to facilitate phytoremediation. Biotechnol. Adv. 2010, 28, 367−374. (24) Lehmann, J.; Rillig, M. C.; Thies, J.; Masiello, C. A.; Hockaday, W. C.; Crowley, D. Biochar effects on soil biota - A review. Soil Biol. Biochem. 2011, 43, 1812−1836. (25) He, Y.; Xu, J.; Lv, X.; Ma, Z.; Wu, J.; Shi, J. Does the depletion of pentachlorophenol in root-soil interface follow a simple linear dependence on the distance to root surfaces? Soil Biol. Biochem. 2009, 41, 1807−1813. (26) Song, Y.; Wang, F.; Yang, X.; Liu, C.; Kengara, F. O.; Jin, X.; Jiang, X. Chemical extraction to assess the bioavailability of chlorobenzenes in soil with different aging periods. J. Soils Sediments 2011, 11, 1345−1354. (27) Srinivasan, S.; Hoffman, N. G.; Morgan, M. T.; Matsen, F. A.; Fiedler, T. L.; Hall, R. W.; Ross, F. J.; McCoy, C. O.; Bumgarner, R.; Marrazzo, J. M.; Fredricks, D. N. Bacterial communities in women with bacterial vaginosis: High resolution phylogenetic analyses reveal relationships of microbiota to clinical criteria. PLoS One 2012, 7, e37818.
planting ryegrass the dissipation of HCB in the biocharamended soils increased. Therefore, this study demonstrates the proof-of-concept that biochar−plant tandem application could effectively control and eliminate contaminants in the soil. Future studies are needed to investigate whether this new remediation scheme could be used for other agriculturally important field crops and/or other types of POPs.
■
ASSOCIATED CONTENT
S Supporting Information *
The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jafc.6b01035. Diagram of experimental design, supplmental methods and supplmental results including root exudate compositions, HCB mass balance in rhizobox experiments, dissolved organic carbon in the equilibrium solution, PCoA analysis of bacterial communities, and correlations of bacterial abundance and HCB dissipation percentages (PDF)
■
AUTHOR INFORMATION
Corresponding Authors
*(Y.S.) Tel: +86 25 86881195/86881193. Fax: +86 25 86881000. E-mail:
[email protected]. *(F.W.) E-mail:
[email protected] *(X.J.) E-mail:
[email protected] Funding
This study was financially supported by the National Key Basic Research Program of China (2014CB441105), the National Natural Science Foundation of China (41301240 and 21277148), the Outstanding Youth Fund of Natural Science Foundation of Jiangsu, China (BK20150050), and the Natural Science Foundation of Jiangsu, China (BK20131049). Notes
The authors declare no competing financial interest.
■
REFERENCES
(1) Zhu, Y. F.; Liu, H.; Xi, Z. Q.; Cheng, H. X.; Xu, X. B. Organochlorine pesticides (DDTs and HCHs) in soils from the outskirts of Beijing, China. Chemosphere 2005, 60, 770−778. (2) Wei, D. B.; Kameya, T.; Urano, K. Environmental management of pesticidal POPs in China: Past, present and future. Environ. Int. 2007, 33, 894−902. (3) Barber, J. L.; Sweetman, A. J.; van Wijk, D.; Jones, K. C. Hexachlorobenzene in the global environment: Emissions, levels, distribution, trends and processes. Sci. Total Environ. 2005, 349, 1−44. (4) Andersen, M. S.; Fuglie, E.; Konig, M.; Lipasti, I.; Pedersen, A. O.; Polder, A.; Yoccoz, N. G.; Routti, H. Levels and temporal trends of persistent organic pollutants (POPs) in arctic foxes (Vulpes lagopus) from Svalbard in relation to dietary habits and food availability. Sci. Total Environ. 2015, 511, 112−122. (5) Corsolini, S. Industrial contaminants in Antarctic biota. J. Chromatogr. A 2009, 1216, 598−612. (6) Bailey, R. E. Global hexachlorobenzene emissions. Chemosphere 2001, 43, 167−182. (7) Lee, W. J.; Shih, S. I.; Chang, C. Y.; Lai, Y. C.; Wang, L. C.; Chang-Chien, G. P. Thermal treatment of polychlorinated dibenzo-pdioxins and dibenzofurans from contaminated soils. J. Hazard. Mater. 2008, 160, 220−227. (8) Sohi, S. P.; Krull, E.; Lopez-Capel, E.; Bol, R. A review of biochar and its use and function in soil. Adv. Agron. 2010, 105, 47−82. (9) Ahmad, M.; Rajapaksha, A. U.; Lim, J. E.; Zhang, M.; Bolan, N.; Mohan, D.; Vithanage, M.; Lee, S. S.; Ok, Y. S. Biochar as a sorbent for 5470
DOI: 10.1021/acs.jafc.6b01035 J. Agric. Food Chem. 2016, 64, 5464−5471
Article
Journal of Agricultural and Food Chemistry (28) Amato, K. R.; Yeoman, C. J.; Kent, A.; Righini, N.; Carbonero, F.; Estrada, A.; Gaskins, H. R.; Stumpf, R. M.; Yildirim, S.; Torralba, M.; Gillis, M.; Wilson, B. A.; Nelson, K. E.; White, B. A.; Leigh, S. R. Habitat degradation impacts black howler monkey (Alouatta pigra) gastrointestinal microbiomes. ISME J. 2013, 7, 1344−1353. (29) Dixon, P. VEGAN, a package of R functions for community ecology. J. Veg. Sci. 2003, 14, 927−930. (30) Takagi, K.; Iwasaki, A.; Kamei, I.; Satsuma, K.; Yoshioka, Y.; Harada, N. Aerobic mineralization of hexachlorobenzene by newly isolated pentachloronitrobenzene-degrading Nocardioides sp Strain PD653. Appl. Environ. Microbio. 2009, 75, 4452−4458. (31) Yan, D. Z.; Mao, L. Q.; Li, C. Z.; Liu, J. Biodegradation of hexachlorobenzene by a constructed microbial consortium. World J. Microbiol. Biotechnol. 2015, 31, 371−377. (32) Beesley, L.; Moreno-Jimenez, E.; Gomez-Eyles, J. L. Effects of biochar and greenwaste compost amendments on mobility, bioavailability and toxicity of inorganic and organic contaminants in a multielement polluted soil. Environ. Pollut. 2010, 158, 2282−2287. (33) Zhang, P.; Sheng, G. Y.; Feng, Y. C.; Miller, D. M. Role of wheat-residue-derived char in the biodegradation of benzonitrile in soil: Nutritional stimulation versus adsorptive inhibition. Environ. Sci. Technol. 2005, 39, 5442−5448. (34) Ding, N.; Hayat, T.; Wang, J.; Wang, H.; Liu, X.; Xu, J. Responses of microbial community in rhizosphere soils when ryegrass was subjected to stress from PCBs. J. Soils Sediments 2011, 11, 1355− 1362. (35) Chiou, C. T.; Kile, D. E.; Brinton, T. I.; Malcolm, R. L.; Leenheer, J. A.; Maccarthy, P. A comparison of water solubility enhancements of organic solutes by aquatic humic materials and commercial humic acids. Environ. Sci. Technol. 1987, 21, 1231−1234. (36) Luo, L.; Zhang, S.; Shan, X. Q.; Zhu, Y. G. Oxalate and root exudates enhance the desorption of p,p′-DDT from soils. Chemosphere 2006, 63, 1273−1279. (37) Xiao, M.; Wu, F. A review of environmental characteristics and effects of low-molecular weight organic acids in the surface ecosystem. J. Environ. Sci. 2014, 26, 935−954. (38) Xie, X. M.; Liao, M.; Yang, J.; Chai, J. J.; Fang, S.; Wang, R. H. Influence of root-exudates concentration on pyrene degradation and soil microbial characteristics in pyrene contaminated soil. Chemosphere 2012, 88, 1190−1195. (39) Hasan, S. A.; Jabeen, S. Degradation kinetics and pathway of phenol by Pseudomonas and Bacillus species. Biotechnol. Biotechnol. Equip. 2015, 29, 45−53. (40) Peng, X.; Qu, X.; Luo, W.; Jia, X. Co-metabolic degradation of tetrabromobisphenol A by novel strains of Pseudomonas sp. and Streptococcus sp. Bioresour. Technol. 2014, 169, 271−276. (41) Sun, K.; Liu, J.; Gao, Y.; Jin, L.; Gu, Y.; Wang, W. Isolation, plant colonization potential, and phenanthrene degradation performance of the endophytic bacterium Pseudomonas sp Ph6-gf p. Sci. Rep. 2014, 4, 1−8. (42) Rentz, J. A.; Alvarez, P. J. J.; Schnoor, J. L. Repression of Pseudomonas putida phenanthrene-degrading activity by plant root extracts and exudates. Environ. Microbiol. 2004, 6, 574−583. (43) Li, X.; Rui, J.; Mao, Y.; Yannarell, A.; Mackie, R. Dynamics of the bacterial community structure in the rhizosphere of a maize cultivar. Soil Biol. Biochem. 2014, 68, 392−401. (44) Shin, K. H.; Lim, Y.; Ahn, J. H.; Khil, J.; Cha, C. J.; Hur, H. G. Anaerobic biotransformation of dinitrotoluene isomers by Lactococcus lactis subsp lactis strain 27 isolated from earthworm intestine. Chemosphere 2005, 61, 30−39. (45) Martin, F.; Torelli, S.; Le Paslier, D.; Barbance, A.; MartinLaurent, F.; Bru, D.; Geremia, R.; Blake, G.; Jouanneau, Y. Betaproteobacteria dominance and diversity shifts in the bacterial community of a PAH-contaminated soil exposed to phenanthrene. Environ. Pollut. 2012, 162, 345−353. (46) Haichar, F. e. Z.; Marol, C.; Berge, O.; Rangel-Castro, J. I.; Prosser, J. I.; Balesdent, J.; Heulin, T.; Achouak, W. Plant host habitat and root exudates shape soil bacterial community structure. ISME J. 2008, 2, 1221−1230.
(47) Leigh, M. B.; Pellizari, V. H.; Uhlik, O.; Sutka, R.; Rodrigues, J.; Ostrom, N. E.; Zhou, J.; Tiedje, J. M. Biphenyl-utilizing bacteria and their functional genes in a pine root zone contaminated with polychlorinated biphenyls (PCBs). ISME J. 2007, 1, 134−148.
5471
DOI: 10.1021/acs.jafc.6b01035 J. Agric. Food Chem. 2016, 64, 5464−5471
