Effects of di-n-butyl phthalate on rhizosphere and non-rhizosphere soil microbial communities at different growing stages of wheat

Ecotoxicology and Environmental Safety 174 (2019) 658–666

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Effects of di-n-butyl phthalate on rhizosphere and non-rhizosphere soil microbial communities at different growing stages of wheat Minling Gaoa,b, Ze Zhanga, Zhengguo Songc,

T

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a

School of Environmental Science and Engineering, Tianjin Polytechnic University, No. 399 Binshui West Road, Xiqing District, Tianjin 300387, China State Key Laboratory of Separation Membranes and Membrane Processes, Tianjin Polytechnic University, No. 399 Binshui West Road, Xiqing District, Tianjin 300387, China c Agro-Environmental Protection Institute, Tianjin 300191, China b

A R T I C LE I N FO

A B S T R A C T

Keywords: DBP Microbial community Rhizosphere Non-rhizosphere Enzyme activities Microbial functional diversity

The potential effects of dibutyl phthalate (DBP) on soil ecosystems and biological processes have recently aroused great concern because of the ubiquitous nature of this pollutant. However, the effects of DBP-associated disturbance on rhizosphere and non-rhizosphere soil microbial communities remain poorly understood. In the present study, we investigated the effects of DBP contamination on microbial function and soil enzyme activities in rhizosphere and non-rhizosphere soils throughout the growing season of wheat. We conducted pot experiments under glasshouse conditions and used different concentrations of DBP: 10, 20, and 40 mg kg–1. We found that the average well color development value and McIntosh index in rhizosphere and non-rhizosphere soils increased in the 10 and 20 mg kg–1 DBP treatments, but declined in the 40 mg kg–1 DBP treatment at the seedling and tillering stages, particularly, in the non-rhizosphere soil. DBP addition enhanced the Shannon-Wiener and Simpson indexes in rhizosphere and non-rhizosphere soils throughout the growing period of wheat. A principal component analysis clearly differentiated the treatments from the control, indicating that DBP led to different patterns of potential carbon utilization in rhizosphere and non-rhizosphere soils. The microbial use of amino acids was significantly increased in rhizosphere and non-rhizosphere soils after DBP addition, while the use of carbohydrates was significantly declined (p < 0.05). The dehydrogenase, urease, and acid phosphatase activities were significantly stimulated (p < 0.05) at the seedling stage, while the phenol oxidase and β-glucosidase activities were inhibited. The 40 mg kg–1 DBP treatment significantly decreased the phenol oxidase and β-glucosidase activities in rhizosphere and non-rhizosphere soils at the seedling stage, particularly in non-rhizosphere soil (p < 0.05). The microbial function and soil enzymatic activities were gradually restored following the wheat growing stage. These results offer a better understanding of the effects of DBP on the activities and functional diversity of microbial communities in farmland soils.

1. Introduction Phthalate esters (PAEs) are industrial chemicals that are widely used as plasticizers and additives. To date, global plastic production has reached a level of 150 million tons, and 6–8 million tons of PAEs are used by humans each year (Net et al., 2015). Numerous studies have demonstrated that some PAEs are endocrine-disrupting compounds that cause estrogenic effects in animals and humans (Sun et al., 2015; Gao et al., 2016). Soil is the principal receptor and an environmental reservoir of semivolatile organic contaminants (Wang et al., 2016). The concentrations of PAEs have reached milligrams per kilogram in most agricultural soils across China (Sun et al., 2016). Yan et al. (2010) reported that the volume of soil residue was 26–50 kg ha–1 in arable areas

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that were covered with plastic film mulch over a period of 10 years. Yang et al. (2013) also observed high concentrations (19.50 μg g−1) of PAEs in areas with increased anthropogenic activities resulting from urbanization and industrialization, and in agricultural districts. The content of DBP in fluvo-aquic soils of Handan is 3.18–29.37 mg kg−1 and 9.02–13.0 mg kg−1 in agricultural soils of Qingdao (Lü et al., 2018). Commonly, the PAE levels in agricultural soils have increased drastically because of the use of irrigation and application of plastic film, pesticides, and sewage sludge in agricultural fields (Zhao et al., 2015). PAEs can accumulate in crops and throughout the food chain because of their low water solubility and high octanol/water partition coefficients (Yuan et al., 2010). Therefore, the PAE contamination of agricultural soils poses a threat to environmental and human health.

Corresponding author. E-mail address: [email protected] (Z. Song).

https://doi.org/10.1016/j.ecoenv.2019.01.125 Received 30 July 2018; Received in revised form 19 January 2019; Accepted 22 January 2019 0147-6513/ © 2019 Elsevier Inc. All rights reserved.

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2. Materials and methods

The soil microbial community plays an important role in many ecosystem processes, such as the biogeochemical cycling of nutrients, soil structural and hydrological properties, and energy flow (Cao et al., 2017), and is an essential indicator of soil health. PAEs and their residues might persist in the soil for long periods because of their hydrophobic nature and consequently strong adsorption on soil particles (Wang, 2014). The PAE contamination can change microbial community composition, and might, therefore, affect the microbial functions and soil enzyme activities (Yuan et al., 2010; Zhang et al., 2015; Wang et al., 2016). Blondel et al. (2017) reported dynamic changes in soil functionality under lindane and chlordecone exposure in areas with or without maize plants and found that soil ammonium concentration, potential nitrogen mineralization, and microbial biomass decreased after exposure to organochlorine pesticides. Wang et al. (2016) investigated the effects of plastic film residues on the occurrence of soil PAEs and microbial activities using a batch pot experiment. These authors found that the PAE concentrations increased with increasing plastic film residues and that the soil microbial carbon and nitrogen contents, enzyme activities, and microbial diversity decreased significantly as a result. Zhou et al. (2005) found that in the constructed wetlands, dibutyl phthalate (DBP) enhanced the activities of dehydrogenase, catalase, protease, and phosphatase, but inhibited the activities of urease, cellulase, and b-glucosidase when the influent DBP concentration was 9.84 mg L–1. These studies indicate that PAEs pose a potential threat to soil health; however, additional studies are needed to assess the impact of PAEs on the rhizosphere soil microbial communities. Numerous microbial activities take place in the rhizosphere because the secretions derived from roots serve as the primary food sources for microbes and, hence, drive their population density and activeness (Li et al., 2016). Within the rhizosphere, microorganisms offer a range of ecosystem services to the plant, including nutrient acquisition, longterm soil sustainability, and resistance to perturbations (Van der Voort et al., 2016). Microbes that can degrade organic pollutants might use organic compounds released by plants as carbon and nitrogen sources for their growth and long-term survival (Wei et al., 2017). In addition, plants might regulate the microbiome in their rhizospheres to enhance their tolerance of abiotic stresses (Li et al., 2014). Plants secrete 10–20% of their photosynthate in root exudates, which support the growth and metabolic activities of diverse fungal and bacterial communities in the rhizosphere (Lefevre et al., 2013). Guo et al. (2017) investigated the effect of the ryegrass rhizosphere on the degradation of polycyclic aromatic hydrocarbons in an aged contaminated agricultural soil and found that the presence of ryegrass promoted the dissipation of polycyclic aromatic hydrocarbons and changed the structures of active bacterial communities in the soil. Zhang et al. (2015) investigated the effect of DBP on the root physiology and rhizosphere microbial community of cucumber seedlings (1–21 days) and found a consequent decrease in the root protein content and root activity. These authors also found that the abundance, structure, and composition of rhizosphere bacteria significantly changed when the concentration of DBP was higher than 50 mg L−1 (Guo et al., 2017; Zhang et al., 2015). However, the effect of DBP on microbial communities in the rhizosphere soil is not clearly understood, and few studies have focused on the relationships among phthalate occurrence, wheat growing stage, and rhizosphere soil microbial activities. The main objectives of this study were: 1) to investigate the impact of DBP on soil microbial functional diversity (using the BIOLOG system), 2) to examine the effect of DBP on soil enzyme activities, 3) to explore different responses between rhizosphere and non-rhizosphere soils under DBP stress, 4) the growing period-dependent changes in the DBP-induced effects on soil microbial community function and soil enzyme activities.

2.1. Chemicals DBP (99% purity) was purchased from J&K (Beijing, China). Methanol (98% purity) and H2O2 were obtained from Jinke Chemical Reagent Co., Ltd. (Tianjin, China). Other chemicals (analytical purity) were acquired from Fengchuan Chemical Reagent Co., Ltd. (Tianjin, China). The DBP stock solution (concentration: 1000 mg L–1) was prepared by dissolving DBP (99% purity) in methanol and subsequently stored at 4.0 °C. For conducting tests, the working concentrations of DBP were prepared at 150, 300, and 600 mg L–1. 2.2. Sterilization of wheat seeds The wheat seeds (Jinqiang 8) were provided by the AgroEnvironmental Protection Institute, Ministry of Agriculture, Tianjin, China. The seeds were sterilized using 30% (v/v) H2O2 for 30 min and then washed several times in distilled water. 2.3. Soil preparation and treatments The soil used was cinnamon soil, which was collected from Shanxi Province (China). The organic matter content of the soil was 1.50%. The soil pH was 7.92, and the total nitrogen, phosphorus, and potassium concentrations were 0.73, 1.03, and 24.3 g kg−1, respectively. The soil was air-dried and sieved through a 2-mm nylon sieve before used. The different DBP methanol solutions (0.70 L) were mixed uniformly with 1.50 kg oven-dry soil. The spiked soils were thereafter vented for 24 h to allow the methanol to vaporize. The soil treatments were then mixed thoroughly with 13.5 kg of uncontaminated soil. Deionized water was added to the contaminated soils in order to maintain a moisture level of approximately 60% of their water-holding capacity (WHC). The contaminated soils were equilibrated in a greenhouse for 5 days. The original, untreated soil was used as the control (CK). A methanol CK group was also prepared. Three replicates were conducted for each CK and treatment (n = 24 pots in total). 2.4. Greenhouse rhizo-box experiment A rhizo-box experiment was conducted in the greenhouses of the Agro-Environmental Protection Institute (Tianjin, China). The rhizobox used in this study was designed according to He et al. (2005) (Fig. S1). The rhizobox was made from polyvinylchloride (polypropylene PP), with the dimensions of 180 × 160 × 220 mm (length × width × height). The box was divided by a nylon mesh (300 mesh, < 53 µm pore size) into three compartments: a root compartment or a rhizosphere zone in the middle (50 mm in width) and nonrhizosphere zones on the left and right sides (each 65 cm in width). The design successfully constrained root hairs from entering the adjacent non-rhizosphere soil and separated the soil zones. Nitrogen and phosphorus, obtained from urea 0.32 g kg–1 soil and KH2PO4 0.15 g kg–1 soil, respectively, were added to each pot. Soil moisture was adjusted to 60% (v/w) of the WHC before use. The rhizo-boxes were randomly arranged in the greenhouse and watered with 150 mL deionized water daily to maintain moderate soil moisture. The rhizosphere and non-rhizosphere soils of the different treatments were sampled during the seedling, tillering, flowering, and ripening stages. The collected soils were ground and homogenized by sieving through a 1-mm stainless steel sieve after removing the stones and residual roots, and then stored in aluminum foil bags at − 80 °C. Precautions were taken during sampling and sample processing to avoid PAE contamination. 659

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The Shannon-Wiener index (H′):

Table 1 Substrates in the EcoPlateTM (Biolog, Inc.) assay categorized. Substrate class

Substrate name

Amines

Phenylethylamine Putrescine Glycyl-L-glutamic acid L-Arginine L-Phenylalanine L-Threonin L-Asparagine L-Serine α-D-Lactose β-Methyl-D-glucoside D-Mannitol i-Erythritol D-Cellobiose D-Xylose N-Acetyl-D-glucosamine γ-Hydroxy butyric acid α-Keto butyric acid 4-Hydroxy benzoic acid 2-Hydroxy benzoic acid D-Galactonic acid γ-lactone D-Galacturonic acid D-Glucosaminic acid D-Malic acid Itaconic acid D,L-Glycerol Glucose-1-phosphate Pyruvic acid methyl ester α-Cyclodextrin Glycogen Tween 80 Tween 40

Amino acids

Carbohydrates

Carboxylic acids

Miscellaneous

Polymers

H′ = − ∑ (Pi × LogPi ) The Simpson index (D):

D=1

− ∑ (Pi )2

McIntosh index (U):

U=

ni 2

where Pi is the proportion of the absorbance of each well to the sum of the absorbance of all wells, and ni is the absorbance of the ith well. 2.6. Soil enzymes The activities of protease, β-glucosidase, dehydrogenase, urease, polyphenol oxidase, and acid phosphatase for each soil sample were determined according to the methods of Serra-Wittling et al. (1995) and Saiya-Cork et al. (2002). 2.7. Statistical analyses Statistical analyses were performed using SPSS 20.0 (IBM Corporation, New York, USA). The data were analyzed for significant differences between the CK treatment or between treatments using oneway analysis of variance. Comparisons of means were conducted using the Tukey's test. Differences were considered significant at p < 0.05. All data are expressed as means ± standard deviations. 3. Results

Note: Substrate classes were seen Zak et al. (1994).

3.1. Effect of DBP on microbial functional diversity indexes 2.5. Biolog EcoPlate analysis The dynamic changes in the AWCD values are shown in Fig. 1a. At the seedling stage, the AWCD values increased by 15.3% and 8.24% in the rhizosphere soil and by 14.3% and 10.0% in the non-rhizosphere soil, compared to the CK soil when the concentrations of DBP were 10 and 20 mg kg–1, respectively, whereas the AWCD value decreased by 8.24% in the rhizosphere soil and by 17.1% in the non-rhizosphere soil upon treatment with 40 mg kg–1 DBP (p < 0.05). At the tillering stage, the AWCD values increased by 8.79% and 5.49% and by 5.48% and 2.74% in the in the rhizosphere and non-rhizosphere soils exposed to the DBP concentrations of 10 and 20 mg kg–1, respectively, compared with the CK soil, and decreased by 4.40% and 5.48% in the rhizosphere and non-rhizosphere soils treated with 40 mg kg–1 of DBP. At the flowering and ripening stages, no significant change in the AWCD value was observed in rhizosphere and non-rhizosphere soils of wheat compared to the CK (p > 0.05). Moreover, the AWCD value in the rhizosphere soils was significantly greater than that of the non-rhizosphere soils (p < 0.05) at all growth stages, except for the ripening stage. Exposure to DBP caused a general increase in the Shannon-Wiener index in both the rhizosphere and non-rhizosphere soils in response to the DBP concentration used throughout the growing period of wheat, as shown in Fig. 1b. The Shannon-Wiener index significantly increased in the DBP treatments at the seedling and tillering stages in both rhizosphere and non-rhizosphere soils (p < 0.05). At the flowering and ripening stages, the Shannon-Wiener index increased slightly, but not statistically significantly, in response to the presence of DBP (p > 0.05). The maximum Shannon-Wiener index was observed during the seedling stage in the 20 mg kg–1 DBP exposure treatment and was 1.03 times and 1.04 times that of the rhizosphere and non-rhizosphere CK, respectively. Compared to the CK, the Simpson index of the rhizosphere and nonrhizosphere soils after DBP treatments notably increased in the seedling stage (p < 0.05) (Fig. 1c). However, no significant difference was observed in the Simpson index of the DBP-treated rhizosphere and non-

Biolog EcoPlates (BIOLOG, Hayward, USA) were used to study the metabolic profiles of the microbial community (Wang et al., 2016). Each 96-well plate consisted of 31 sole carbon sources and water as the CK, with three replicates of each substrate. According to the methodology of Zak et al. (1994), we grouped the 31 substrates into substrate classes: amines, amino acids, carbohydrates, carboxylic acids, miscellaneous, and polymers (Table 1). A total of 1 g of fresh soil samples (with the soil moisture content tested before weighing) were suspended in triangular flasks filled with 100 mL of sterile buffer solution (0.85% NaCl) in a biological safety cabinet. The flasks were then sealed and incubated in a shaker for 30 min. After settling for 15 min, the soil suspensions were obtained and diluted (1:1000) with a sterile inoculating solution (0.85% NaCl). Subsequently, 0.1 mL soil suspension was transferred into 96 wells on the Biolog EcoPlates. The plates were incubated in the dark at 28 °C, and the absorbance was determined using a micro-plate reader (PERLONG, China) at 595 nm after 96 h. Each reading of the Biolog EcoPlates was corrected by subtracting the value of CK and standardized by dividing the values of the average well color development (AWCD) of the replicates (Insam et al., 1996). The AWCD value reflected the microbial activity and was calculated using the following formula (Garland, 1997):

AWCDECO =

∑ (C − R)/n

where, C is the value of absorbance of each well with a carbon source, R is the value of absorbance of the blank well, and n is the number carbon sources (i.e., n = 31). The functional diversity of the soil microbial community can be determined by the Shannon-Wiener index, Simpson index, and McIntosh index based on the absorbance at 96 h. The equation of each index is given below: 660

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Fig. 1. The dynamic changes of microbial functions from the five DBP treatments (CK, Methanol, DBP10, DBP20, DBP40) in rhizosphere and non-rhizosphere soils during the whole growing period of wheat (values are means of three replicates ± standard error. The same lowercase letter on bars means no significant differences among treatments (p < 0.05). (a) Average well color development (AWCD); (b) H′ index (Shannon index); (c) D index (Simpson index); (d) U index (McIntosh index)).

rhizosphere soils after the seedling stage, compared to the CK (p > 0.05). The McIntosh indexes in the rhizosphere and non-rhizosphere soils are shown in Fig. 1d, and they exhibited a similar trend as seen with the AWCD values throughout the wheat growing period. At the seedling stage, the McIntosh indexes in the 10 mg kg–1 DBP-treated rhizosphere and non-rhizosphere soils increased by 9.86% and 7.25%, respectively, compared to the CK. However, when the soils were treated with 20 and 40 mg kg–1 DBP, the McIntosh indexes decreased by 0.37% and 13.9% and by 1.48% and 21.5% in the rhizosphere and non-rhizosphere soils, respectively. The McIntosh index did not significantly change after the tillering stage, compared to the CK (p > 0.05). However, the McIntosh index in the rhizosphere soil was significantly higher than that of the non-rhizosphere soil throughout the growing period (p < 0.05), except during the ripening stage. 3.2. Effect of DBP on microbial community carbon source utilization Fig. 2 describes the influence of DBP treatments on six groups of carbon sources (amino acids, polymers, carbohydrates, carboxylic acids, miscellaneous, and amines). Overall, the microbial community used carboxylic acids, amino acids, polymers, and carbohydrates more than amines and miscellaneous carbon sources in both rhizosphere and non-rhizosphere soils. The amino acids showed the highest microbial utilization in rhizosphere and non-rhizosphere soils in the presence of DBP throughout the growing period of wheat (Fig. 2a). A peak in the amino acid usage emerged in the tillering stage, and increased with increasing DBP concentrations by 43.7%, 53.6%, and 29.5% in the rhizosphere soil and by 44.8%, 70.8%, and 46.0% in the non-rhizosphere soil (Fig. 2a). Although the utilization of polymers was promoted by the presence of DBP in the rhizosphere and non-rhizosphere soils (Fig. 2b), the degree of promotion was gradually weakened. At the seedling stage, the utilization of polymers at the DBP concentrations of 10, 20, and 40 mg kg–1 DBP increased by 32.6%, 16.71%, and 5.82% in the rhizosphere soil and by 36.1%, 21.5%, and − 6.03% in the nonrhizosphere soil, respectively, compared to the CK. However, carbohydrates in the CK were metabolized more than the other carbon sources in the rhizosphere and non-rhizosphere soils (Fig. 2c). The utilization of carbohydrates in the rhizosphere and non-rhizosphere soils of wheat was significantly inhibited by the increase in DBP concentration throughout the growing period (p < 0.05). For example, the utilization of carbohydrates decreased by 4.86%, 24.8%, and 34.3% in the rhizosphere soil and by 13.6%, 29.8%, and 41.7% in the non-rhizosphere soil under the conditions of 10, 20, and 40 mg kg–1 DBP treatments at the seedling stage, respectively. No significant difference in the utilization of carboxylic acid, amine, and miscellaneous carbon sources was observed between the CK and rhizosphere and non-rhizosphere soils (p > 0.05) throughout the growing period, except in the seedling stage (Fig. 2d–f). In addition, the utilization of the six groups of carbon sources in the rhizosphere soil was significantly higher than that in the non-rhizosphere soil for four growing seasons (p < 0.05). 3.3. Principal component analysis (PCA) A PCA analysis of the substrate utilization patterns is shown in Fig. 3. The first and second principal components (PC1 and PC2) explained 32.7% and 20.9% of the rhizosphere soil (Fig. 3a), and 29.6% 661

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Fig. 3. Principal component analysis (PCA) of the utilization pattern of substrate carbon sources in rhizosphere and non-rhizosphere soils.

and 15.9% of the non-rhizosphere soil (Fig. 3b), respectively. It was obvious that the rhizosphere soil of wheat treated with DBP was clearly different from the soil in the CK treatment throughout the growing season. Similar observations were obtained in the non-rhizosphere soil (Fig. 3b). Therefore, the PCA showed that DBP application could lead to different patterns of carbon source utilization and different microbial communities both in rhizosphere and non-rhizosphere soils. In addition, a clear discrimination was observed in sampling at the seedling, tillering, flowering, and ripening stages, indicating the effects of wheat growth stages on the carbon substrate utilization pattern within a microbial community in rhizosphere and non-rhizosphere soils. The analysis of the loadings of carbon sources on PC1 and PC2 showed that both DBP addition and growth stages appeared to be the factors that could influence the catabolic diversity of microbial communities under our experimental conditions. Tables 2, 3 present the loadings of BIOLOG substrates in the rhizosphere and non-rhizosphere soils, respectively. The largest loadings on PC1 in the rhizosphere soil were from L-arginine, L-aspartic acid, Tween 40, oxybenzoic acid, L-phenylalanine, Tween 80, γ-hydroxybutyrate, L-threonine, itaconic acid, glycine-L-glutamic acid, phenethylamine, and DL glycerol. The largest loadings on PC2 in the rhizosphere soil were from D-galactonic acid-γ-lactone, D-xylose, Dgalacturonic acid, N-acetyl-D-glucosamine, D-glucosamine, D-cellobiose, and glucose-1-phosphate (Table 2). Table 3 shows that in the non-rhizosphere soil, major contributions to PC1 were made by hydroxybenzoic acid, L-phenylalanine, Tween 80, L-serine, γ-hydroxybutyric acid, L-threonine, itaconic acid, glycine, and L-glutamic acid and to PC2 by N-acetyl-D-glucose and glycogen.

Fig. 2. Utilization of different carbon sources by the soil microbial community after 96 h incubation (values are means of three replicates ± standard error. The same lowercase letter on bars means no significant differences among treatments (p < 0.05)).

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Table 2 Loadings of BIOLOG substrates to the two principal components (PC1–PC2) in rhizosphere soil. Substrate

PC1

PC2

β-Methyl-D-glucoside D-Galactonic acid γ-lactone L-Arginine Pyruvic acid methyl ester D-Xylose D-Galacturonic acid L-Asparagine Tween 40 i-Erythritol 4-Hydroxy benzoic acid L-Phenylalanine Tween 80 D-Mannitol 2-Hydroxy benzoic acid L-Serine α-Cyclodextrin N-Acetyl-D-glucosamine γ-Hydroxy butyric acid L-Threonin Glycogen D-Glucosaminic acid Itaconic acid Glycyl-L-glutamic acid D-Cellobiose Glucose-1-phosphate α-Keto butyric acid Phenylethylamine α-D-Lactose D,L-Glycerol D-Malic acid Putrescine

− 0.131 − 0.0360 0.627 0.452 − 0.385 − 0.211 0.718 0.669 0.561 0.908 0.886 0.918 0.309 0.530 0.538 0.468 − 0.254 0.715 0.746 0.581 0.0770 0.893 0.871 − 0.075 − 0.251 0.433 0.637 0.437 0.697 0.539 0.251

0.487 0.726 − 0.114 − 0.244 0.862 0.679 0.195 0.123 0.542 − 0.286 − 0.0660 − 0.004 0.501 0.359 − 0.322 0.380 0.870 − 0.0970 0.0230 0.410 0.764 − 0.133 − 0.154 0.679 0.788 0.409 0.170 0.423 0.290 − 0.223 0.472

Table 3 Loadings of BIOLOG substrates to the two principal components (PC1–PC2) in non-rhizosphere soil. Substrate

PC1

PC2

β-Methyl-D-glucoside D-Galactonic acid γ-lactone L-Arginine Pyruvic acid methyl ester D-Xylose D-Galacturonic acid L-Asparagine Tween 40 i-Erythritol 4-Hydroxy benzoic acid L-Phenylalanine Tween 80 D-Mannitol 2-Hydroxy benzoic acid L-Serine α-Cyclodextrin N-Acetyl-D-glucosamine γ-Hydroxy butyric acid L-Threonin Glycogen D-Glucosaminic acid Itaconic acid Glycyl-L-glutamic acid D-Cellobiose Glucose-1-phosphate α-Keto butyric acid Phenylethylamine α-D-Lactose D,L-Glycerol D-Malic acid Putrescine

− 0.410 − 0.410 0.470 0.460 − 0.800 − 0.490 0.510 0.490 0.210 0.940 0.840 0.710 0.000 0.270 0.610 0.170 − 0.660 0.660 0.660 0.200 − 0.460 0.840 0.860 − 0.400 − 0.620 0.180 0.500 0.170 0.500 0.550 − 0.140

0.0200 0.550 0.210 − 0.0400 0.430 0.450 0.550 0.250 0.550 − 0.0700 0.170 0.500 0.450 0.390 − 0.0100 0.510 0.610 0.040 0.280 0.710 0.560 0.330 − 0.100 0.550 0.520 0.470 0.170 0.380 0.330 − 0.190 0.400

(caption on next page)

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Fig. 4. Effects of different DBP concentrations on the enzyme activities in rhizosphere and non-rhizosphere soils for different wheat growth stages (values are means of three replicates ± standard error. The same lowercase letter on bars means no significant differences among treatments (p < 0.05). (a) urease; (b) dehydrogenases; (c) protease; (d)acid phosphatase; (e) polyphenol oxidase; (f) b-glucosidase.).

activity in the rhizosphere soil was 119% of that in the non-rhizosphere soil. 3.4.5. Polyphenol oxidase activity Fig. 4e describes the impact of DBP on the polyphenol oxidase activity in the rhizosphere and non-rhizosphere soils. The DBP treatment showed no significant inhibition of the polyphenol oxidase activity in the rhizosphere soil, except for a significant decrease in the 40 mg kg–1 DBP treatment at the seedling and tillering stages. A general decrease in the polyphenol oxidase activity was observed in the non-rhizosphere soil in response to the increase in DBP concentration at the seedling and tillering stages. At the seedling stage, the polyphenol oxidase activity in the non-rhizosphere soil was reduced by 0.48%, 11.5%, and 24.9% with an exposure to the DBP concentrations of 10, 20, and 40 mg kg–1, respectively. The DBP-mediated inhibition of polyphenol oxidase activity gradually weakened in both rhizosphere and non-rhizosphere soils with wheat development.

3.4. Effect of DBP on soil enzyme activities The effects of DBP on enzyme activities in the rhizosphere and nonrhizosphere soils are shown in Fig. 4a–f. Different enzymes showed different response patterns to different DBP concentrations and wheat growth stages. 3.4.1. Urease activity Fig. 4a shows that in the DBP-treated rhizosphere and non-rhizosphere soils, the urease activity increased significantly with increasing DBP concentration at the seedling and tillering stages (p < 0.05), but not in the flowering and ripening stages (p > 0.05), when compared to the CK soil. In the tillering stage with 40 mg kg−1 DBP, the maximum urease activity in the rhizosphere and non-rhizosphere soils was 108% and 127% of that in the CK soil, respectively.

3.4.6. β-glucosidase activity The changes in the β-glucosidase activity in response to DBP exposure are shown in Fig. 4f. At the seedling and tillering stages, the βglucosidase activity decreased significantly with increasing DBP levels in both rhizosphere and non-rhizosphere soils, compared to the CK soil (p < 0.05). The activity of β-glucosidase remained constant in the rhizosphere and non-rhizosphere soils at the flowering and ripening stages.

3.4.2. Dehydrogenase activity The dehydrogenase activities in the DBP-treated rhizosphere and non-rhizosphere soils are shown in Fig. 4b. Compared to the CK soil, the dehydrogenase activity in the rhizosphere soil was not significantly different (p > 0.05) within the same growing stages, except for a significant increase (2.66%) in the 40 mg kg–1 DBP treatment at the seedling stage (p < 0.05). The dehydrogenase activity in the non-rhizosphere soil exhibited a steady increase with increasing DBP concentration at the seedling and tillering stages. In the non-rhizosphere soil treated with DBP concentrations of 10, 20, and 40 mg kg–1, the dehydrogenase activity increased by 1.79%, 6.23%, and 10.0% at the seedling stage and by 0.12%, 4.00%, and 6.75% at the tillering stage, respectively. Compared to the CK, the activity of dehydrogenase increased slightly, but not statistically significantly, in the rhizosphere and non-rhizosphere soils from the tillering to the ripening stage.

4. Discussion The production and utilization of agricultural plastic film have rapidly increased in recent years and are the potential cause of the increasing soil PAE pollution in China. Over the last decade, PAEs have frequently been identified in farmland soils from different regions of China (Sun et al., 2018). Previous studies have found that environmental pollution might affect the metabolic activities of soil microbes and consequently change the soil microbial community composition and diversity (Akmal et al., 2005; Pringault et al., 2015). The AWCD value is an important index of microbial functional diversity and is used to assess the total metabolic activity of soil microbes. It provides a general indication on the reactive capacity of the microbial metabolic diversity with respect to 31 different carbon sources (Wang et al., 2015). The functional diversity indexes H′, D, and U of soil microbes can be used to estimate the microbial diversity, dominant microbial diversity, and microbial distribution uniformity, respectively (Cao et al., 2017). In this study, the AWCD values and McIntosh indexes decreased in the seedling stage in the soils treated with high concentrations of DBP. This might be attributed to a decline in the metabolic activity as a result of cell membrane fluidity being destroyed or the abundance of some cultivable microbial community (Wang et al., 2016). In addition, studies have shown that the Simpson and ShannonWiener indexes in the rhizosphere and non-rhizosphere soils were enhanced by DBP in all contamination treatments (10, 20, and 40 mg kg–1) throughout the growing period of wheat. It is probable that pollutants in soils could enrich tolerant species via the environmental filtering process, which, in turn, might affect the overall ecosystem functions and natural balance of soil microorganism communities (Sun et al., 2018). Wang et al. (2015) reported the impacts of dimethyl phthalate on microbial community function in black soils. Similar to our findings from the AWCD and biodiversity assessments, the changes in the utilization of different carbon sources indicated that the metabolic functions of the soil microbial community were sensitive to DBP contamination, and the metabolic features of soil microbes were altered. The PCA was able to distinguish between the CK soil and the rhizosphere and non-rhizosphere soils treated with DBP. The difference in the pattern of carbon substrate use between the DBP treatments and CK

3.4.3. Protease activity The protease activity in the rhizosphere and non-rhizosphere soils is shown in Fig. 4c. At the seedling stage, the protease activity in the rhizosphere and non-rhizosphere soils treated with 10 and 20 mg kg–1 DBP increased by 3.79% and 0.57% and by 22.1% and 12.5%, respectively, compared to that in the CK soil. However, the protease activity decreased by 15.0% and 16.2% in the rhizosphere and non-rhizosphere soils treated with 40 mg kg–1 DBP, respectively. At the tillering stage, when the soil was treated with 40 mg kg–1 of DBP, the protease activity increased significantly in the rhizosphere soil (10.3%), but decreased in the non-rhizosphere soil (22.8%) (p < 0.05). At the flowering and ripening stages, the protease activity in the rhizosphere and non-rhizosphere soils was not markedly altered with an increase in DBP concentration (p > 0.05). 3.4.4. Acid phosphatase activity Fig. 4d shows that the acid phosphatase activities in both rhizosphere and non-rhizosphere soils increased significantly with an increase in DBP concentration at the seedling stage (p < 0.05). No significant increase in acid phosphatase activity was observed in the rhizosphere and non-rhizosphere soils after the seedling stage, while the acid phosphatase activity in the rhizosphere soil slightly increased with the increase in DBP concentration (p > 0.05). An obvious difference was observed in the acid phosphatase activity between the rhizosphere and the non-rhizosphere soils (p < 0.05). At the tillering stage in the 40 mg kg−1 DBP treatment, the maximum acid phosphatase 664

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soil is not (or is only slightly) affected by plants roots and root exudates, with a consequently lower level of microbial activity and soil fertility. Guo et al. (2017) found that the ryegrass root enhanced the functional bacterial diversity in the early stages of growth (0–10 days); these findings coincided with our results. Simultaneously, the higher enzyme activities in the rhizosphere soil could be linked to the larger microbial populations and higher enzyme activities, since microbial populations contribute to the release of these enzymes. Furthermore, our results showed that the inhibition of protease enzyme activity caused by 40 mg kg–1 DBP in the rhizosphere soil was not significant, except for a significant increase at the tillering stage (p > 0.05). A comparatively greater reduction in the AWCD value in the rhizosphere soil treated with 40 mg kg–1 DBP than in the similarly treated non-rhizosphere soil was seen in the seedling stage. Enhanced enzymatic activity might improve plant growth through rhizosphere soil nutrient enrichment and increase in resistance against disturbance, although enzyme synthesis and secretion might decline in response to the DBP treatment (Wei et al., 2017). Thus, the resistance of soil microorganisms to high concentrations of DBP might be enhanced by plant growth. This result is comparable with the study conducted by Ma et al. (2016), who showed that the soil microbial activity was significantly affected by the addition of OTC and might be ameliorated by planting alfalfa or Sedum plumbizincicola. A possible cause might be that the soil supplied with sufficient nutrients and an abundance of microorganism populations might easily adapt to the presence of this organic pollutant. Another possible reason is that plant roots might influence the bioavailability and decontamination of soil PAEs by altering water flux and oxygenation, increasing soil porosity, and changing microbial communities through the release of exudates (Xie et al., 2012). Obviously, soil enzymes were insensitive to DBP after the tillering stage, and the differences in soil enzyme activities between the CK and DBP treatments faded gradually over time. It was possible that DBP was dissipated and adsorbed in soil. Thus, soil microbial community function could recover or improve because of DBP dissipation in soil. The dissipation of PAEs in rhizosphere soil can be a result of abiotic dissipation (such as hydrolysis, photolysis, evaporation, etc.), biodegradation, and plant uptake and accumulation. The uptake and accumulation by plants is a significant pathway in the migration and transformation of PAEs in the environment (Lin et al., 2017). Zhao et al. (2015) found that DBP and di-(2-ethylhexyl) phthalate could be taken up simultaneously by various cultivars of Chinese flowering cabbage. Sun et al. (2015) reported that plants (lettuce, strawberry, and carrot) could take up and accumulate PAEs from the soil and transform them in both whole plants and plant cells. However, Tang et al. (2015) evaluated the influence of various factors (flow types, substrates, plants, hydraulic loading rates) on PAE removal and concluded that the uptake of plants appeared to be of minor importance compared to the microbial degradation processes.

might be attributed to the tolerance discrepancy of different microorganism to pollutants and to the utilization of DBP as a source of carbon and energy (Zhang et al., 2014). Alternatively, plants might play a key role in selecting efficient microbial metabolic capacities and communities in the use of rhizodeposits as substrates, and they might also affect microbial community resistance to DBP (Deng et al., 2015). Nevertheless, the pattern of carbon substrate use was similar in both rhizosphere and non-rhizosphere soils. It is likely that the root exudates of low-molecular-weight compounds could disperse from the rhizosphere to more distant soils because of their diffusion in soil (Blondel et al., 2017). Falchini et al. (2003) reported the soil diffusion gradient of carbohydrates, organic acids, and amino acids by monitoring the concentrations of C-14-labeled glucose, oxalic acid, or glutamic acid. Owing to their high specificity and catalytic efficiency, soil enzymes secreted by plants, soil microorganisms, and animals play important roles in biological metabolic processes (Lebrun et al., 2012). Soil enzymes are the catalysts of important metabolic processes, including the detoxification of xenobiotics and heavy metals (Thavamani et al., 2012). The activities of urease and phosphatase might reflect potential alterations in the soil contents of N and P (Lebrun et al., 2012). Our study showed that the urease and acid phosphatase activities in rhizosphere and non-rhizosphere soils were significantly increased after DBP application at the seedling stage (p < 0.05). The increased levels of urease and acid phosphatase activities might be explained by enzyme activation or induction (Shackle et al., 2000). Lyubun et al. (2013) reported that the arsenic pollution stimulated the soil phosphatase activity, and such stimulation might be explained by the presence of Psolubilizing bacteria, which increase the presence water-soluble phosphorus in soil. In addition, this observation suggests that DBP influences the nitrogen and phosphorus cycle in soil. The activities of β-glucosidase and polyphenol oxidase are involved in the biochemical transformation rate of soil organic carbon (Cao et al., 2017). Our results showed that the polyphenol oxidase and β-glucosidase activities in both rhizosphere and non-rhizosphere soils were significantly inhibited when the soils were contaminated with a DBP concentration of 40 mg kg–1 during the seedling stage of wheat (p < 0.05). The negative effect of DBP on the activities of phenol oxidase and β-glucosidase might be attributed to the DBP-mediated dephosphorylation of regulatory proteins in physiological processes; this, in turn, might denature and degrade the activities of phenol oxidase and β-glucosidase (Shackle et al., 2000). Zhang et al. (2014) concluded that organic chemicals could alter enzyme activities by interacting with the enzyme-substrate complex, by denaturing the enzyme protein, or by interacting with the protein-active groups. Furthermore, changes in the community structure might indirectly modify enzyme activities. Our findings showed that the protease activity was significantly increased in the 10 and 20 mg kg–1 DBP treatments, but significantly decreased in the 40 mg kg–1 DBP treatment in the non-rhizosphere soil (p < 0.05); these findings were similar to those reported by Wang et al. (2015). Dehydrogenase has been used as an important index for determining the influence of contaminants and is related to overall microbial activity in soil (Zhou et al., 2005). In the present study, the dehydrogenase activity in both rhizosphere and non-rhizosphere soils was promoted by DBP. It is probable that DBP acted as a substrate and induced microbial biomass, which, in turn, promoted the activities of these enzymes. Cycoń et al. (2006) reported that fungicides and insecticides could act as carbon sources and nutrients for microbial growth, leading to an increase in bacterial biomass. Furthermore, our results clearly indicated that the microbial communities of the rhizosphere soil exhibited greater activity and functional diversity, likely resulting from the release of root components. Plant roots release organic compounds, such as sugars, polysaccharides, amino acids, fatty acids, sterols, proteins, and secondary metabolites in their exudates (Kumar et al., 2016). Root components might enhance the biodegradation of bioaccessible fractions and influence the rate of microbial growth (Wei et al., 2017). Conversely, the non-rhizosphere

5. Conclusion In this study, different microbial indicators were used to assess the impact of soil microbial community structure and activities. In the biolog assay, the wheat rhizosphere functionality did not seem to be altered by low- and medium-concentration DBP exposure, while the non-rhizosphere soil functioning showed significant modifications caused by DBP, especially at the seeding and tillering stages. The patterns of potential carbon utilization were changed upon treatment with different DBP concentrations, especially in wheat soil. In addition, DBP enhanced the activities of soil enzymes associated with nitrogen and phosphorus metabolism and inhibited the activities of soil enzymes related to carbon utilization in both rhizosphere and non-rhizosphere soils. The differences between the DBP treatments and CK weakened in the later growth stages of the plant. Moreover, the enzyme activities and microbial functional diversity in the rhizosphere were generally higher than those in the non-rhizosphere soil throughout the growing period of wheat, and plants enhanced the resistance of DBP. It is vital 665

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that further studies are conducted to elucidate the molecular mechanisms of changes in the soil microbial community structure due to plastic film residues so that appropriate measures could be taken to reduce plastic film residues and protect soil quality.

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