Biochar reduces the bioaccumulation of PAHs from soil to carrot (Daucus carota L.) in the rhizosphere: A mechanism study

Science of the Total Environment 601–602 (2017) 1015–1023

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Science of the Total Environment journal homepage: www.elsevier.com/locate/scitotenv

Biochar reduces the bioaccumulation of PAHs from soil to carrot (Daucus carota L.) in the rhizosphere: A mechanism study Ni Ni a,b, Yang Song a, Renyong Shi a,b, Zongtang Liu c, Yongrong Bian a, Fang Wang a, Xinglun Yang a, Chenggang Gu a, Xin Jiang a,⁎ a b c

Key Laboratory of Soil Environment and Pollution Remediation, Institute of Soil Science, Chinese Academy of Sciences, Nanjing 210008, PR China University of the Chinese Academy of Sciences, Beijing 100049, PR China Jiangsu Key Laboratory for Bioresources of Saline Soils, Yancheng Teachers University, Yancheng, Jiangsu 224051, PR China

H I G H L I G H T S

G R A P H I C A L

A B S T R A C T

• Corn straw and bamboo-derived biochars reduced PAH bioaccumulation in carrot roots. • BB pyrolyzed at 700 °C reduced the PAH bioavailability primarily via immobilization. • CB at 300 °C reduced the PAH bioavailability primarily via promoting biodegradation. • Dominant retention mechanism depends on the type of biochar in the rhizosphere.

a r t i c l e

i n f o

Article history: Received 2 April 2017 Received in revised form 28 May 2017 Accepted 28 May 2017 Available online xxxx Editor: Jay Gan Keywords: Biochar Polycyclic aromatic hydrocarbons Bioavailability Microbial community structure Metagenome prediction

a b s t r a c t The aim of this study was to reveal the mechanisms on how biochar reduces bioaccumulation of polycyclic aromatic hydrocarbons (PAHs) in tuberous vegetables. Corn straw-derived biochar pyrolyzed at 300 °C (CB300) or bamboo-derived biochar pyrolyzed at 700 °C (BB700) was amended into PAH-contaminated soil planted with carrot (Daucus carota L.). After 150 days, 2% CB300 or 2% BB700 amendments significantly reduced the bioaccumulation of PAHs in carrot root (p b 0.05), especially for high-molecular-weight PAHs. In the non-rhizosphere, either CB300 or BB700 suppressed PAH dissipation and decreased the bioavailability via adsorption processes. Compared to the control, the total concentration of PAHs in the rhizosphere was higher in the 2% BB700 treatment but the bioavailable concentration was lower. This indicates that BB700 decreased the bioavailability of PAHs primarily via immobilization (adsorption processes). By contrast, the total and bioavailable PAH concentrations were both lower in the 2% CB300 treatment than those in the control. The abundance of bacteria such as Arthrobacter and Flavobacterium and the total number of genes playing important roles in microbial PAH degradation processes increased significantly (p b 0.05), which were likely responsible for the rapid dissipation of PAHs in the 2% CB300 treatment in the rhizosphere. These results indicate that CB300 decreased the PAH bioavailability primarily via increasing degradation of PAHs by indigenous microorganisms. The two biochars both showed better effectiveness at reducing the bioavailability of high-molecular-weight PAHs than the low-molecular-weight PAHs in the rhizosphere. Therefore, the mechanisms on how biochar reduces the PAH uptake into carrot are dependent on the type of biochar (e.g., pyrolysis temperature and feedstock) and root presence. © 2017 Elsevier B.V. All rights reserved.

⁎ Corresponding author at: 71 East Beijing Road, Nanjing 210008, PR China. E-mail address: [email protected] (X. Jiang).

http://dx.doi.org/10.1016/j.scitotenv.2017.05.256 0048-9697/© 2017 Elsevier B.V. All rights reserved.

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1. Introduction Polycyclic aromatic hydrocarbons (PAHs) are recognized as toxic, persistent and bioaccumulative contaminants (Man et al., 2013; Ruby et al., 2016; Gong et al., 2017). PAHs enter the atmosphere, aquatic and soil environment primarily as a result of anthropogenic activities, e.g., transportation emissions, sludge application and wastewater irrigation, in addition to the incomplete combustion of fossil fuel (Usman et al., 2016). In farmland soils, PAHs persist for a long time because of the large quantities discharged as well as their semi-volatility, low water solubility, and resistance to biodegradation (Wang et al., 2011). In a recent investigation, the total PAH concentrations varied widely, from 8.8 to 3880 μg/kg, in a collection of surface soil samples at 109 sites in agricultural soils in eastern China (Sun et al., 2017). The available PAHs in soil which are freely available to cross an organism's cellular membrane (Barnier et al., 2014) can find their way into the food chain after their uptake into crops, eventually posing a threat to human health (Usman et al., 2016). PAHs enter plants from contaminated soils primarily via two pathways: root uptake from soil solution (Su and Zhu, 2008; Namiki et al., 2013) and particle-phase deposition onto the waxy cuticle of plant leaves (Collins et al., 2006; Odabasi et al., 2015) after volatilizing into the air from soil. Therefore, reducing the volatilization and bioavailability of PAHs is the critical step to reduce the migration from soils into plants. Biochar is widely used as a cost-effective soil amendment in environmental remediation (Meyer et al., 2011). Amendment with biochar in the soil could reduce the uptake of PAHs into some plants, which is mainly affected by the physicochemical properties of the biochar (e.g., polarity, functional groups, pore structure, feedstock, elemental composition) and the chemical structures and the PAH concentrations as well as the plant species (Ahmad et al., 2014; Anyika et al., 2015). For example, uptake of PAHs into maize was significantly decreased in soil amended with pine woodchip-derived biochar pyrolyzed at 450 °C (Brennan et al., 2014). Sewage sludge biochar pyrolyzed at 550 °C, 500 °C, 550 °C could reduce the mobility of PAHs in the soils and therefore reduce the bioaccumulation of PAHs in Cucumis sativa L., Lactuca sativa L. and Solanum lycopersicum, respectively (Waqas et al., 2014; Khan et al., 2013; Waqas et al., 2015). Furthermore, compared with leafy crops, tuberous vegetables (e.g., carrot, radish and turnip) contain much higher PAH concentrations because the accumulation of PAHs in roots is higher than that in other parts of the plant (Florence et al., 2015). However, studies focusing on the effect of biochar on the bioavailability of PAHs to tuberous vegetables are limited (Khan et al., 2015). Whether and how biochar can reduce the accumulation of PAHs in tuberous vegetables need to be further elucidated. Generally, the reduced bioavailability of PAHs in biochar-amended soils is primarily attributed to the sorption ability of biochar (Wang et al., 2011). Khan et al. (2015) reported that treating soil with soybean straw biochar, rice straw biochar or peanut shell biochar (PNBC), all pyrolyzed at 500 °C, significantly decreased the available concentrations of PAHs for turnips (Brassica rapa L.), and the decreases in availability varied with the type of biochar used. In this set of biochar, PNBC has the largest surface area and lowest polarity and thus the highest sorption capacity for PAHs. Biochar produced under high pyrolytic temperature is highly efficient in improving the sorption capacity of PAHcontaminated soil (Chen and Yuan, 2011). Due to the recalcitrance, pore structures and nutrient properties of biochar, microbial degradation of PAHs in biochar-amended soils could also be influenced simultaneously (Anyika et al., 2015). Biochar affected the microbial activity and community structures through its physicochemical properties, such as nutrients, pH and energy (Ennis et al., 2012). The biomass, activity and diversity of PAH-metabolizing microorganisms as well as gene expression are observed to be enhanced after biochar application (Liu et al., 2015; Anyika et al., 2015; Tang et al., 2013). Moreover, the rhizosphere (R) is the gateway for pollutants to transfer from soil into the food chain (D'Orazio et al., 2013; Storey et al., 2014), and reducing

PAH bioavailability in the rhizosphere is the critical step for reducing the bioaccumulation in crops (Liu et al., 2013). Plant root-microbe interactions play an important role in the degradation of PAHs in R soils (Cébron et al., 2011; Storey et al., 2014; Thomas and Cébron, 2016). However, less information is available regarding the effect of biochar on the microbial community structure in relation to PAH dissipation in soils with an active R region. Whether the sorption or promotion of degradation of PAHs occurs in the R soil of tuberous vegetables in biochar-amended soil also remains unclear. Therefore, the objective of this study was to investigate whether biochar could reduce the accumulation of PAHs in tuberous vegetables, and if so, to determine the mechanisms for the actions of different biochars. Corn straw-derived biochar (CB) and bamboo-derived biochar (BB) were selected and amended into moderate PAH-contaminated farmland soil planted with a root vegetable, carrot (Daucus carota L.). Phospholipid fatty acid (PLFA) profiles were examined as reliable tracers of the effects of environmental stress on soil microbial highlevel taxonomic groups, and high-throughput sequencing for 16S rDNA genes was used to shed light on the detailed structure of the soil bacterial community. The software PICRUSt (Phylogenetic Investigation of Communities by Reconstruction of Unobserved States) was used to predict metagenome functional content from marker gene surveys and full genomes. 2. Materials and methods 2.1. Soil sampling and characterization An agricultural soil contaminated with PAHs for N 40 years was sampled from a depth of 0–20 cm in a region near a steel factory in a suburb of Nanjing, Jiangsu Province (31°89′75″N, 118°61′30″E). The soil was air-dried, sieved through a 2 mm mesh and manually homogenized for the pot experiment. The soil was classified as silty loam and the physicochemical properties are shown in Table S1. The sum and individual concentrations of 16 PAHs listed by the United States Environmental Protection Agency (US EPA) as priority pollutants in the soil sample are shown in Table S2. The concentrations of 11 PAHs in the soil exceed the environmental quality standards for farmland soils in China (GB 15618– 2008) by two orders of magnitude, respectively (Table S2). 2.2. Biochar preparation and characterization The CB, produced under anoxic conditions at 300 °C (CB300) using a patented biochar reactor (NO. ZL2009 2 0232191.9) (Jia et al., 2013), has shown great sorption efficiency for PAHs in a previous study (Ni et al., 2017). The BB, produced at 700 °C (BB700) and purchased from Shanghai Hainuo Charcoal Co., Ltd., was also a commercial efficient sorbent for organic contaminants (Denyes et al., 2012). The surface area, pore volume and pore size of the materials were determined via the BrunauerEmmett-Teller (BET) method using a V-Sorb 2800P analyzer (Gold APP Instruments Corporation, Beijing, China). The elemental compositions, including carbon (C), nitrogen (N), sulfur (S), hydrogen (H) and oxygen (O), were measured with an elemental analyzer (ANA1500, Carlo Erba, Milano, Italy) (Yuan et al., 2011). The details of biochar properties are presented in Table S3. 2.3. Experimental design Briefly, 3 kg of the soil were manually mixed thoroughly with 15 g CB300 (0.5%) in a storage box. The amended soil was subsequently transferred into cylindrical polyvinyl chloride pots (20 cm in height and 20 cm in bottom diameter) and then compacted to soil bulk density of 1.3 g/cm3. Treatments with 2% CB300, 0.5% BB700 and 2% BB700 were applied using the same procedure. Soil without biochar addition was used as the control. Hence, the five treatments were control, 0.5% CB300, 2% CB300, 0.5% BB700 and 2% BB700, each in triplicate. The

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soils were irrigated with deionized water and then kept for 1 week before carrot cultivation. The carrot seeds were sterilized in 30% H2O2 for 10 min, washed, and then immersed in deionized water for one week on moist filter paper in culture dishes. Ten uniformly germinated seeds were sown in each pot. The plants were grown in a greenhouse under natural diurnal light conditions from May to October in 2015. Every 30 days, 10 g of the non-rhizosphere (NR) soil from each pot was sampled using a soil borer to measure the total and bioavailable concentrations of PAHs. Carrots were sampled on days 90, 120 and 150 during the growth period, and two carrots were sampled at each time point. The collected shoots and roots were thoroughly washed with deionized water and freeze-dried. At the end of incubation, a brushing method was used to separate the R soil from the root surface (Zhang et al., 2017). Five grams of the NR and R soil was collected separately and stored at − 80 °C until further analysis for PLFA profiles and high-throughput sequencing. 2.4. PAHs extraction and quantification in soil and plant Total PAH concentrations in soils/carrots were extracted by accelerated solvent extraction (ASE 200, Dionex, Sunnyvale, CA, USA) (Zhang et al., 2013a). Briefly, 1 g soil or 0.5 g carrot sample was homogenized with 5 g of diatomaceous earth and extracted by hexane/acetone (4:1, V/V) at 100 °C with a pressure of 1500 psi. The extracts were rotary evaporated at 50 °C to 1 mL. The concentrated samples were successively washed with silica gel/anhydrous sodium sulfate column for soil and sulfonated silica/anhydrous sodium sulfate column for carrot and then eluted with 15 mL of hexane/dichloromethane (9:1, V/V). The eluate was concentrated to 1 mL for detection of PAHs by gas chromatography-mass spectrometer analysis (GC–MS, Agilent 7890A/ 5975C, Santa Clara, CA). Available PAH concentrations in soil were determined by extracting PAHs from soil with hydroxypropyl-β-cyclodextrin (HPCD), which is an effective method for evaluating the bioavailability of PAHs (Oleszczuk et al., 2016). Briefly, 1 g of freeze-dried soil was extracted with 20 mL of HPCD (50 mmol/L) in a glass centrifuge tube by shaking on an orbital shaker at 200 r/min for 20 h followed by centrifugation for 30 min. The supernatant was discarded, and the residue soil was shaken with deionized water for 10 min and then centrifuged again. The supernatant was also discarded. Then, the total PAH concentrations in the residue soil were measured using the method described above. The bioavailable PAH concentration in the soil was calculated by subtracting the total PAH concentration in residue soil after HPCD extraction from the total PAH concentration in the soil before HPCD extraction. 2.5. Microbial PLFA analysis PLFA assay was used to analyze the soil microbial community structure (Lazcano et al., 2013). Briefly, 2 g of freeze-dried soil was extracted with 15 mL of chloroform-methanol-citrate buffer solution (10:20:8, V/ V/V). Polar lipids (including phospholipids) were subsequently separated from neutral lipids and glycolipids using Sep-Pak solid-phase extraction columns (Waters, Milford, MA, USA) by eluting with 1 mL of chloroform, 5 mL of acetone, and 6 mL of methanol. Fatty acid methyl esters (FAMEs) were extracted successively by saponification and methylation and then quantified by GC (Agilent 6890 Agilent Technologies, Little Falls, DE, USA). FAMEs were identified and quantified using MIDI software with MIDI microbial calibration standards (MIDI, Inc., Newark, DE, USA). The PLFAs were categorized into four groups of gram-positive bacteria, gram-negative bacteria, fungi and actinomycetes, following the reported rules (Huygens et al., 2011). 2.6. High-throughput sequencing The R and NR soils collected at harvest were subjected to highthroughput sequencing analyses. Genomic DNA of the samples was

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extracted using the PowerSoil DNA Isolation kit (MoBio Laboratories Inc., Carlsbad, USA). DNA concentration was measured with a NanoDrop 2000C ultramicrospectrophotometer (Thermo Scientific, Waltham, USA) and quality was checked using 0.8% agarose gel electrophoresis. The V3–V4 region of bacterial 16S rRNA was amplified by PCR for high-throughput pyrosequencing. The 16S rRNA gene V3-V4 region of bacteria was amplified using the universal primers 338F (5′-ACTCCT ACGGGAGGCAGCA-3′) and 806R (5′-GGACTACHVGGGTWTCTAAT-3′). The PCR program was as follows: 95 °C for 3 min, 27 cycles of 95 °C for 30s, 55 °C for 30s, and 72 °C for 45 s, with a final extension of 72 °C for 10 min. The amplicons were determined using the Illumina MiSeq PE300 sequencing platform (Illumina, Inc., CA, USA) according to the manufacturer's recommendations. The raw sequencing data were deposited in the SRA database of NCBI under the study accession number SRP100491. The barcoded 16S RNA gene sequences were trimmed into libraries using Quantitative Insights into Microbial Ecology (QIIME) program version 1.8.0 (Caporaso et al., 2010). To retain only high-quality sequences for the downstream analysis, sequences that were b100 bp in length after quality trimming, contained one or more ambiguous base-calls (N), or had b90% quality scores N Q20 were eliminated. High-quality sequences were clustered into operational taxonomical units (OTUs) at a 97% similarity level using UPARSE version 7.1 (Edgar, 2013). The phylogenetic affiliation of each 16S rRNA gene sequence was analyzed with RDP Classifier (http://rdp.cme.msu.edu/) against the silva (SSU117) 16S rRNA database using a confidence threshold of 70% (Amato et al., 2013). The Shannon index representing α-diversity was calculated in MOTHUR v.1.30.1 (Schloss et al., 2009). 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 (Dixon, 2003). PICRUSt used evolutionary modeling to predict metagenomes from 16S data compared with a reference genomedatabase (Langille et al., 2013). The metagenomes were collapsed into the Kyoto Encyclopedia of Genes and Genomes (KEGG, http://www.kegg.jp/) Orthology (KO). Briefly, the OTUs of 16S rRNA sequences were normalized in PICRUSt, after which, the KO profiles were calculated with the PICRUSt algorithm. Genes related to PAH degradation among different treatments were screened out. 2.7. Quality control and statistical analysis The recovery of 16 PAHs was measured for quality control. Briefly, 10 g of soil were spiked with the mixed standard samples of 16 PAHs (500 μg/kg). Extraction and purification of the soil samples were performed using the procedure described above. To estimate the recoveries for PAH residues in carrots, 0.5 g of freeze-dried carrot samples were spiked with the mixed standard samples of 16 PAHs (500 μg/kg). The sample extraction and analyses were performed using the same procedures described above. The average recoveries of triplicate samples were 96% ± 7% in the soil sample and 89% ± 5% in the carrot sample. Data were statistically analyzed using analysis of variance (ANOVA) and the least significant difference (LSD) post-hoc comparison tests with SPSS V14.0 (International Business Machines Corporation, New York, USA) at a p b 0.05 significance level. 3. Results and discussion 3.1. Carrot biomass and uptake kinetics of PAHs by carrot The effects of the biochars on carrot biomass production after 150 days of growth are shown in Fig. S1. No significant differences in carrot biomass were observed among the Control, 0.5% CB300, 0.5% BB700 and 2% BB700 treatments. However, the carrot root weight was increased up to 1.83-fold in the 2% CB300 treatment compared with

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Fig. 1. Concentrations of ∑16 PAHs in carrot shoots (A) and roots (B) grown in soils amended with/without corn straw-derived biochar (CB) and bamboo-derived biochar (BB). Control: no biochar addition; 0.5% CB300: 0.5% corn straw-derived biochar addition; 2% CB300: 2% corn straw-derived biochar addition; 0.5% BB700: 0.5% bamboo-derived biochar addition; 2% BB700: 2% bamboo-derived biochar addition. Different letters indicate significant differences among the treatments by LSD post-hoc comparison tests at p b 0.05. Error bars indicate the standard deviation (N = 3).

the control. The soils amended with 2% CB300 contained the highest levels of organic matter, dissolved organic matter and nitric nitrogen (Table S4), which may explain the increase in yield. Biochars that are not thoroughly carbonized may lead to increased plant growth through improving plant nutrient uptake and the availability of phosphorus (P), cadmium, and potassium in soil (Schmidt et al., 2014). As shown in Fig. 1, the concentration of ∑16 PAHs in the carrot root was significantly higher than that in the carrot shoot. The carrot root-toshoot translocation factor was b0.15 (Table S5), verifying that PAHs were preferentially bioaccumulated in the carrot root (Zohair et al.,

2006). No significant differences in the concentration of ∑ 16 PAHs were detected in carrot shoots among all treatments (Fig. 1A), and no differences in ∑16 PAH concentrations in carrot roots were observed among the Control, 0.5% CB300 and 0.5% BB700 treatments (p b 0.05, Fig. 1B). However, compared with the control, the amendments with 2% CB300 and 2% BB700 effectively decreased the PAH accumulation in carrot roots after 150 days, with decreases of 51% ± 3% and 30% ± 2%, respectively (Fig. 1B). The bio-concentration factors of PAHs in carrot roots amended with 2% CB300 and 2% BB700 were also significantly lower than that in the control (Table S5). These results suggest that the

Fig. 2. Time course of total concentrations of ∑16 PAHs (A), 2(+3)-ring (B), 4-ring (C), and 5(+6)-ring (D) PAHs in non-rhizosphere soils amended with/without corn straw-derived biochar (CB) and bamboo-derived biochar (BB). Control: no biochar addition; 0.5% CB300: 0.5% corn straw-derived biochar addition; 2% CB300: 2% corn straw-derived biochar addition; 0.5% BB700: 0.5% bamboo-derived biochar addition; 2% BB700: 2% bamboo-derived biochar addition. Different letters indicate significant differences among the treatments by LSD posthoc comparison tests at p b 0.05. Error bars indicate the standard deviation (N = 3).

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retention effect of the biochars used was shown in carrot roots, where PAHs preferentially bio-accumulated, and that the amendment with 2% CB300 was the most effective in reducing the uptake of PAHs into carrot. The decrease of the PAH concentrations in both shoots and roots could be attributed partly to the growth dilution, and partly to metabolism or the formation of non-extractable bound residues in carrots (Salam et al., 2017). After 150 days, compared to the control, the reductions in 2(+ 3)-, 4-, and 5(+ 6)-ring PAHs bioaccumulated in carrot roots in the 2% CB300 and 2% BB700 treatment varied greatly, by 42% ± 2%, 58% ± 3%, 63% ± 2% and 21% ± 1%, 29% ± 1%, 55% ± 3%, respectively (Fig. S2), suggesting that the bioaccumulation of highmolecular-weight (HMW, containing four to six benzene rings) PAHs in carrot was more substantially reduced by the addition of CB300 or BB700 than the bioaccumulation of low-molecular-weight (LMW, containing two to three benzene rings) PAHs. This result may be because that HMW PAHs sorb more strongly to soils and biochar-amended soils than LMW PAHs (Khan et al., 2015; Brennan et al., 2014). 3.2. Dynamic changes in concentrations of total and bioavailable PAHs in NR soils Although CB300 and BB700 contained a certain level of PAH, the amendment of soil with CB300 or BB700 did not significantly enhance the initial PAH concentrations in the soils (contributing b1%). The dissipation of PAHs in the NR soils amended with/without biochars is presented in Fig. 2. After 150 days, the total concentrations of ∑16 PAHs, 2(+3)-, 4-, and 5(+6)-ring PAHs in the control were significantly decreased, while no significant differences were observed between the Control and 0.5% CB300 treatment. After 60 days, the total concentrations of ∑ 16 PAHs, 2(+ 3)-, 4-, and 5(+ 6)-ring PAHs in the 2% CB300, 0.5% BB700, and 2% BB700 treatments were higher than in the

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control (p b 0.05), indicating that CB300 or BB700, at appropriate rates, effectively suppressed the dissipation of either LMW or HMW PAHs in the NR soil. After 150 days, the residue level of ∑ 16 PAHs, LMW or HMW PAHs, in soil was highest in the 2% BB700 treatment, followed by 0.5% BB700 and 2% CB300. The percentages of HPCD-extracted concentrations to the total concentrations were calculated to express the bioavailability of PAHs in soils (Fig. 3). In the control, the percentages of ∑16 PAHs, 2(+3)-, 4-, and 5(+ 6)-ring PAHs did not decrease significantly throughout the incubation period. After 150 days, the percentages of ∑16 PAHs, 2(+ 3)-, 4-, and 5(+ 6)-ring PAHs were significantly lower in the 2% CB300, 0.5% BB700 and 2% BB700 treatments than in the control, which may be due to the higher residues (Fig. 2) and the lower HPCDextractable concentrations of PAHs (Fig. S3) in the biochar-amended soils than in the control. These results indicate that the biochars, used at appropriate rates, could reduce the bioavailability of either LMW or HMW PAHs in the NR soil via adsorption processes. No significant differences in the percentages of HPCD-extracted concentrations to the total concentrations of ∑16 PAHs, 2(+3)-, 4-, and 5(+6)-ring PAHs were observed between the Control and 0.5% CB300 treatments, indicating the absence of the immobilization effect with the 0.5% CB300 amendment. At the end of the incubation, the HPCD extractions of ∑ 16 PAHs, 2(+ 3)-, 4-, and 5(+ 6)-ring PAHs, were lower in the BB700amended soils than those in the CB300-amended soils, indicating that BB700 showed a stronger ability to reduce the bioavailability of either LMW or HMW PAHs than CB300 in the NR soil. This result is consistent with a previous study in which BB700 had a higher sorption capacity for phenanthrene (Phe)/pyrene (Pyr) in an aqueous system than CB300 (Ni et al., 2017). Sorption of PAHs by biochar is mainly affected by the production temperature of biochar and PAH chemical structures (Anyika et al., 2015). Aromatic hydrocarbons are slightly polar and can

Fig. 3. Time course of the percentage of HPCD-extracted concentrations to the total concentrations of ∑16 PAHs (A), 2(+3)-ring (B), 4-ring (C), and 5(+6)-ring (D) PAHs in nonrhizosphere soils amended with/without corn straw-derived biochar (CB) and bamboo-derived biochar (BB). Control: no biochar addition; 0.5% CB300: 0.5% corn straw-derived biochar addition; 2% CB300: 2% corn straw-derived biochar addition; 0.5% BB700: 0.5% bamboo-derived biochar addition; 2% BB700: 2% bamboo-derived biochar addition. Different letters indicate significant differences among the treatments by LSD post-hoc comparison tests at p b 0.05. Error bars indicate the standard deviation (N = 3).

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be covalently bonded to polar surfaces of biochar. As a result, sorption of PAHs into biochar is related to the aromaticity of the biochar (Wang et al., 2010). In this study, BB700 has larger surface area, higher aromaticity (defined by a H/C ratio of 0.08) and lower polarity (defined by an O/C ratio of 0.16) than CB300 (Table S3), which facilitates the sorption process (Khan et al., 2015). 3.3. PAH concentrations in R soils amended with or without biochar As shown in Fig. S4, the bioavailable concentrations of ∑16 PAHs, 2(+ 3)-, 4-, and 5(+ 6)-ring PAHs in the R soils were well correlated with the PAH concentrations in carrot roots (R2 = 0.81–0.89, p b 0.05), indicating that the amendments with biochars reduced PAH bioaccumulation in carrot roots by decreasing the bioavailability of PAHs in the R soils. The concentrations of ∑16 PAHs, 2(+3)-, 4-, and 5(+ 6)-ring PAHs were higher in the R soil amended with 2% BB700 than those in the control (Fig. 4), while the percentages of HPCDextracted concentrations to the total concentrations were lower in the 2% BB700 treatment than those in the control (Fig. 5). This result was consistent with that in the NR soils, suggesting that BB700 decreased the bioavailability of either LMW or HMW PAHs in the R soil primarily via immobilization. By contrast, both of the total concentrations and percentages of HPCD-extracted concentrations to the total concentrations of ∑16 PAHs, 2(+3)-, 4-, and 5(+ 6)-ring PAHs were lower in the R soil amended with 2% CB300 than those in the control (Fig. 4 and Fig. 5). This result reveals that the effect of CB300 on the dissipation and bioavailability of PAHs in the R soils differed from that in the NR soils and from the effect of BB700. Because the contribution of PAHs uptake into carrot to the total PAH dissipation in the soil was b 0.02% (Table S6), the decreased total concentrations of either LMW or HMW PAHs in the 2% CB300 treatment may be mainly attributed to the increased biodegradation of PAHs caused by 2% CB300. Poultry manurederived biochar containing high N and P was observed to have greater impact on microbial degradation. It could be an excellent candidate to provide simultaneous impacts on sorption and biodegradation (Cimò et al., 2014). In this study, compared to BB700, CB300 contained more abundant nutrients (Table S3) that were released into the soil (Table S4), likely modulating changes in the microbial community structure and increasing in the biodegradation of PAHs. Therefore, in addition to immobilization, the 2% CB300 amendment also likely reduced

Fig. 4. Total concentrations of ∑16 PAHs, 2(+3)-ring, 4-ring, and 5(+6)-ring PAHs in rhizosphere soils amended with/without corn straw-derived biochar (CB) and bambooderived biochar (BB) after 150 days of carrot growth. Control: no biochar addition; 0.5% CB300: 0.5% corn straw-derived biochar addition; 2% CB300: 2% corn straw-derived biochar addition; 0.5% BB700: 0.5% bamboo-derived biochar addition; 2% BB700: 2% bamboo-derived biochar addition. Different capital (low-molecular-weight PAHs) and lowercase letters (∑ 16 and high-molecular-weight PAHs) indicate significant differences among the treatments by LSD post-hoc comparison tests at p b 0.05. Error bars indicate the standard deviation (N = 3).

Fig. 5. Percentage of HPCD-extracted concentrations to the total concentrations of ∑16 PAHs, 2(+3)-ring, 4-ring, and 5(+ 6)-ring PAHs in rhizosphere soils amended with/ without corn straw-derived biochar (CB) and bamboo-derived biochar (BB) after 150 days of carrot growth. Control: no biochar addition; 0.5% CB300: 0.5% corn strawderived biochar addition; 2% CB300: 2% corn straw-derived biochar addition; 0.5% BB700: 0.5% bamboo-derived biochar addition; 2% BB700: 2% bamboo-derived biochar addition. Different letters indicate significant differences among the treatments by LSD post-hoc comparison tests at p b 0.05. Error bars indicate the standard deviation (N = 3).

the bioavailability of PAHs in the R soil by enhancing the biodegradation of available PAHs. Additionally, the ability of either BB700 or CB300 to decrease the bioavailability of HMW PAHs was more effective than that toward LMW PAHs (p b 0.05) in the R soils, resulting in their better performance at reducing the bioaccumulation of HMW PAHs in carrot roots than that of LMW PAHs (Fig. S2). 3.4. Bacterial community structure To further explore the possible reasons for the rapid dissipation of PAHs in the R soils, particularly in the 2% CB300 treatment, the microbial PLFAs extracted from the R and NR soils were identified (Table 1). After 150 days, no significant differences in total PLFAs between the control and the 2% BB700 treatment in the R or NR soils were observed. Compared with the control, the amendment of 2% CB300 remarkably increased the total PLFAs in the R and NR soils by 31% ± 1% and 22% ± 1%, respectively (p b 0.05). The addition of the biochars did not affect the abundance of gram-positive bacteria, actinomycetes or fungi. However, the biomass of gram-negative bacteria in 2% CB300 treatments was increased by 53% ± 6% and 37% ± 2% in the R and NR soils, respectively, compared with the control (p b 0.05). Gomez et al. (2014) found a similar shift in microbial community structures caused by the addition of oak-derived biochar pyrolyzed at 550 °C. Moreover, the increase in bacterial PLFAs was best correlated with the PAH dissipation in the R soils (R2 = 0.87–0.96, p = 0.008) (Table S7). Most bacteria that degrade persistent organic pollutants (e.g., Pseudomonas, Bacillus and Arthrobacter) are gram-negative (Liu et al., 2015; Song et al., 2016). Therefore, bacteria dominated the shift in the soil microbial community and the dissipation of PAHs in the R soils, particularly in the 2% CB300 treatment. To further evaluate bacterial community structures in the R soils amended with biochar, high-throughput sequencing of the soils from the Control, 2% CB300, and 2% BB700 treatments was conducted. No significant difference was detected in the bacterial diversity (as expressed by the Shannon index) between the control and 2% BB700 R treatments, while bacterial diversity in the 2% CB R treatment was lowest in the treatments with fastest PAH dissipation (Table S8). In the R soil, bacterial community structures were similar among the Control, 2% CB300 and 2% BB700 treatments but different from the bacterial community structures in the NR soil (Fig. S5A), indicating that planting carrots,

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Table 1 Microbial biomass indicated by phospholipid fatty acid (PLFA) concentrations in all treatments after 150 days of growth. Microbial biomass indicated by PLFA concentrations (nmol/g)

Rhizosphere

Non-rhizosphere

Control 0.5% CB300 2% CB300 0.5% BB700 2% BB700 Control 0.5% CB300 2% CB300 0.5% BB700 2% BB700

Gram-positive bacteria

Gram-negative bacteria

Actinobacteria

Fungi

5.03 5.24 5.19 4.79 4.92 2.83 3.24 3.15 3.11 2.87

5.89 7.04 8.99 5.58 6.05 4.21 4.39 5.77 4.22 4.32

2.45 2.01 2.21 2.19 2.37 1.85 1.81 1.94 1.76 1.80

1.16 1.09 1.51 0.94 1.13 0.75 0.82 0.92 0.85 0.89

± ± ± ± ± ± ± ± ± ±

0.21a 0.13a 0.09a 0.31a 0.15a 0.23b 0.16b 0.09b 0.13b 0.12b

± ± ± ± ± ± ± ± ± ±

0.23b 0.08b 0.05a 0.05b 0.11b 0.08c 0.12c 0.21b 0.05c 0.13c

± ± ± ± ± ± ± ± ± ±

0.07a 0.16abc 0.20abc 0.11abc 0.10ab 0.12c 0.09c 0.14bc 0.23c 0.08c

± ± ± ± ± ± ± ± ± ±

Total PLFAs 0.24ab 0.12bc 0.10a 0.06bc 0.04bc 0.11c 0.07bc 0.15bc 0.06bc 0.08bc

15.09 ± 1.14bc 15.55 ± 1.25b 19.72 ± 0.96a 14.68 ± 1.13c 15.11 ± 0.28bc 9.72 ± 0.15e 10.27 ± 0.04e 11.82 ± 0.08d 10.18 ± 0.09e 9.95 ± 0.11e

Control: no biochar addition; 0.5% CB300: 0.5% corn straw-derived biochar addition; 2% CB300: 2% corn straw-derived biochar addition; 0.5% BB700: 0.5% bamboo-derived biochar addition; 2% BB700: 2% bamboo-derived biochar addition. Mean values with the same letter in a column are not significantly different among treatments by LSD post-hoc comparison tests at the 5% level. Means ± standard deviation (N = 3).

rather than biochar addition, was the primary factor controlling the similarity of the bacterial community structures in the soils. Moreover, the microbial community composition in the 2% BB700 R treatment was similar to that in the control. However, the addition of 2% CB300 significantly changed the microbial community composition in the R soil (Fig. S5B). The rapid dissipation of PAHs in the R soil indicated that the carrot root likely create optimal conditions for certain types of bacteria to

establish ecological niches, which might be ideal for PAH dissipation. At the genus level, the relative abundances of Arthrobacter, Rhizobium, Altererythrobacter, Terrimonas, Flavihumibacter, Sphingobium, Ohtaekwangia, Flavobacterium, Chryseolinea, Thermomonas, Arenimonas, Lysobacter, Pseudoxanthomonas were higher among the Control, 2% CB300 and 2% BB700 treatments in the R soils than those in the NR soils (Fig. 6). Sphingobium was reported to increase in the presence of biochar and degrade recalcitrant compounds (Anderson et al., 2011).

Fig. 6. Heatmap of bacterial community structure in the non-rhizosphere (NR; Cluster 1) and rhizosphere (R; Cluster 2) soils with/without biochar amendment. Control: no biochar addition; 2% CB300: 2% corn straw-derived biochar addition; 2% BB700: 2% bamboo-derived biochar addition. The red-boxed regions indicate certain genera with higher relative abundance in the R than in the NR soils. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

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Ohtaekwangia and Chryseolinea were highly competitive in the presence of root exudates and primarily utilize the labile organic substances in R soils (Li et al., 2014; Kim et al., 2013). Among these genera, the relative abundances of Pseudoxanthomonas, Lysobacter, Arenimonas, Thermomonas, Altererythrobacter, Rhizobium, Arthrobacter, and Flavobacterium showed linear relationships with the dissipation of ∑16 PAHs, LMW or HMW PAHs among the treatments in the R soils, respectively with R2 values of 0.59–0.85, 0.63–0.86, 0.61–0.91 (Table S9). Particularly, Arthrobacter and Flavobacterium might be partly responsible for the fastest dissipation of PAHs in the R soil of the 2% CB300 treatment. Arthrobacter has the potential to degrade biphenyl and other hydrocarbons (Haichar et al., 2008), and biochar-amended soil contains more members of Flavobacterium, which are known to be degraders of PAHs (Gregory et al., 2015). 3.5. Bacterial functional genes composition Different functional genes are involved in PAH-degradation processes. Therefore, a prediction of gene function related to PAH degradation was conducted to further investigate the changes in bacterial communities in soils. After 150 days, the predictive functional genes among the Control, 2% CB300 and 2% BB700 treatments in the R soils were significantly different from those in the NR soils (Fig. S6A), and the 2% CB300 amendment notably changed the functional genes composition in the R region (Fig. S6B). This result was consistent with the bacterial community composition in the soils (Fig. S4). Genes encoding approximately 10 enzymes related to PAH degradation, i.e. 1, 2–dihydroxynaphthalene dioxygenase, 3–hydroxyanthranilate 3,4–dioxygenase, 4–hydroxyphenylpyruvate dioxygenase, 4–hydroxyacetophenone monooxygenase, 1, 4– dihydroxy–2–naphthoyl–CoA hydrolase, nicotinamide adenine dinucleotide phosphate (NADP)–dependent aldehyde dehydrogenase, cis–1, 2– dihydro–1, 2–dihydroxynaphthalene, 1, 4–dihydroxy–6–naphthoate synthase, PAH dioxygenase large subunit, and 1–hydroxy–2–naphthoate dioxygenase (Muangchinda et al., 2015; Zhang et al., 2013b), were detected in the soils (Table S10), verifying the intrinsic PAH degradation capacity of native microflora. Consistent with the shifts in the bacterial community structure, the mean proportions of 3–hydroxyanthranilate

3, 4–dioxygenase, 4–hydroxyphenylpyruvate dioxygenase and NADPdependent aldehyde dehydrogenase were significantly higher in the R soil of 2% CB300 treatment than those of the Control or 2% BB700 treatments (Fig. 7). These genes with increased abundance mainly encoded the bacterial dioxygenase and dehydrogenase activities involved in PAH degradation (Wang et al., 2016; Fuchs et al., 2011), therefore likely increased the dissipation of PAHs in the R soil amended with 2% CB300. The results suggested that CB300 may decrease the bioavailability of PAHs primarily by inducing more bacteria and functional genes involved in PAH degradation in the R soil, while the influence of BB700 on the bacterial community structure and PAH-degradation genes was relatively small.

4. Conclusion In the rhizosphere, the amendment of CB300 or BB700 at appropriate rates could reduce the migration of PAHs from soil into tuberous vegetables (i.e., carrot), and show a greater efficiency for HMW than LMW PAHs. The mechanisms for the actions of these two biochars were different. BB700 suppressed PAHs dissipation and decreased the bioavailability mainly via immobilization, while CB300 enhanced PAHs dissipation and decreased the bioavailability primarily by promoting bacterial degradation. However, in the non-rhizosphere, both CB300 and BB700 decreased the bioavailability of PAHs mainly via immobilization. Therefore, the mechanisms on how biochar reduced the bioaccumulation of PAHs in the tuberous vegetables (i.e., carrot) depend on the biochar type (e.g., pyrolysis temperature and feedstock) and root presence.

Funding This study was supported by the National Key Basic Research Program of China (2014CB441105); the National Natural Science Foundation of China (41671236); the “135” Plan and Frontiers Program of the Institute of Soil Science, Chinese Academy of Sciences (ISSASIP1614); and the Outstanding Youth Fund of Natural Science Foundation of Jiangsu Province, China (BK20150050). Appendix A. Supplementary data Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.scitotenv.2017.05.256.

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Fig. 7. Mean proportions of 3-hydroxyanthranilate 3,4-dioxygenase, 4hydroxyphenylpyruvate dioxygenase and NADP-dependent aldehyde dehydrogenase in the rhizosphere (R) and non-rhizosphere (NR) soils amended with/without corn strawderived biochar (CB) and bamboo-derived biochar (BB) after 150 days of carrot growth. Control: no biochar addition; 2% CB300: 2% corn straw-derived biochar addition; 2% BB700: 2% bamboo-derived biochar addition. Different letters indicate significant differences among the treatments by LSD post-hoc comparison tests at p b 0.05. Error bars indicate the standard deviation (N = 3).

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