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Biodegradability and ecological safety assessment of Stenotrophomonas sp. DDT-1 in the DDT-contaminated soil
Ecotoxicology and Environmental Safety 158 (2018) 145–153
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Ecotoxicology and Environmental Safety journal homepage: www.elsevier.com/locate/ecoenv
Biodegradability and ecological safety assessment of Stenotrophomonas sp. DDT-1 in the DDT-contaminated soil
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Hua Fang, Yanfei Deng, Qiqing Ge, Jiajia Mei, Houpu Zhang, Huifang Wang, Yunlong Yu Institute of Pesticide and Environmental Toxicology, College of Agriculture & Biotechnology, Zhejiang University, Hangzhou 310058, China
A R T I C LE I N FO
A B S T R A C T
Keywords: DDT Stenotrophomonas Soil Bioremediation Ecological safety assessment
The biodegradability and ecological safety assessment of the previously isolated DDT-degrading bacterial strain Stenotrophomonas sp. DDT-1 were investigated in the DDT-contaminated soil under laboratory and field conditions. Under laboratory conditions, the degradation rates of fresh p,p′-DDT in soil were enhanced by 2.0–3.0-fold with the introduction of the strain DDT-1 compared to those of the control treatments. A similar enhancement in the dissipation of DDTs (p,p′-DDT, p,p′-DDE, p,p′-DDD, and o,p′-DDT) in the aged DDT-contaminated field plot soils resulted from the inoculation with this strain. Meanwhile, the degradation rates of DDTs increased by 2.9–5.5- and 2.8–7.6-fold in the inoculated greenhouse and open field soils, respectively, after field demonstration application of strain DDT-1 preparation. Moreover, no significant differences in the soil enzyme activity, microbial functional diversity, and bacterial community structure were observed between the inoculated and uninoculated field soils, but several soil microbial genera exhibited some fluctuations in abundance. It is concluded that strain DDT-1 could accelerate the removal of DDTs residues in field soils, and furthermore, its inoculation was ecologically safe.
1. Introduction 1,1,1-Trichloro-2,2-bis(4-chlorophenyl) ethane (DDT) has been extensively applied since the 1940s as an insecticidal agent against agricultural pests and arthropod diseases (Ren et al., 2016). DDT can be transformed into some major metabolites, including 1-dichloro-2,2-bis (4-chlorophenyl)ethylene (DDE) and 1,1-dichloro-2,2-bis-(4-chlorophenyl ethane (DDD), which may be more recalcitrant and ecologically toxic (Yu et al., 2011; Chen et al., 2013). Although the use of DDT has been prohibited in most countries since the early 1970s and in China since 1983, DDT residues and its metabolites have still been widely detected in soils, sediments, surface water, and groundwater (Fang et al., 2016). Exposure to DDT residues can cause a wide range of symptoms such as nausea, headache, vomiting, tremors, and confusion (Purnomo et al., 2011; Singh et al., 2016). Meanwhile, the high lipophilicity of DDT allows it to accumulate in food sources and fatty tissues of organisms, disrupting the central neural system and causing reproductive disorders (Mcglynn et al., 2008). Considering these negative effects, it is urgent to seek an effective remediation method to remove residual DDT. Some researchers have reported that soil microorganisms play a key role in the detoxification metabolism of organochlorine compounds (Tong et al., 2014; Chen et al., 2016a, 2016b), and thus bioremediation
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has been recognized as a useful, cost-effective, and safe approach to remove or detoxify DDTs (p,p′-DDT, p,p′-DDE, p,p′-DDD, and o,p′-DDT) residues from contaminated soils (Fang et al., 2010;). The biodegradation of DDTs residues by DDT-degrading strains under laboratory conditions has been well documented (Fang et al., 2016; Pan et al., 2017). Bajaj et al. (2014) reported that a glycolipid-producing Rhodococcus sp. strain IITR03 could degrade 282 µM DDT under aqueous culture conditions. Xiao et al. (2011) found that the white rot fungi Phlebia lindtneri and Phlebia brevispora could remove 70% and 30% of DDT, respectively, in a pure culture of low-nitrogen medium after incubation for 21 d, and further degrade the metabolites DBP and DBH via hydroxylation of the aromatic ring. Ziya and Teresa (2016) demonstrated the ability of the aerobic bacterium Alcaligenes eutrophus A5 to degrade DDT in both aqueous and soil phases. Grewal et al. (2016) observed that approximately 42% of the initial 50 mg/l of DDT disappeared from a culture system after 10-d inoculation with the bacterial strain Serratia marcescens NCIM 2919. However, little information is available on the in situ bioremediation potential and ecological safety using DDT-degrading strains in contaminated field soils. In our previous study, a DDT-degrading bacterial strain DDT-1 was isolated and identified as a member of Stenotrophomonas sp., and this strain could convert DDT into DDE/DDD, DDMU, DDOH, and DDA via dechlorination, hydroxylation, and carboxylation, and ultimately
Corresponding author. E-mail address: [email protected] (Y. Yu).
https://doi.org/10.1016/j.ecoenv.2018.04.026 Received 13 December 2017; Received in revised form 26 March 2018; Accepted 12 April 2018 Available online 24 April 2018 0147-6513/ © 2018 Elsevier Inc. All rights reserved.
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(CFUs)/g and thoroughly intermixed by stirring and sieving to ensure the uniform distribution of strain DDT-1. The soil water content was adjusted to 60% of the maximum water holding capacity and maintained by periodic distilled water additions. Finally, soil samples were transferred into 500-ml plastic cups and immediately covered with aluminum foil with 5 small holes (1 mm diameter). Soil samples amended with the same volume of sterilized water were used as the controls. All plastic cups were placed in an artificial climate chamber at 25 °C in the dark. Each treatment was performed in triplicate, and 25 g of soil samples were randomly collected from each plastic cup using a plastic spoon at 0, 7, 14, 21, and 28 d, respectively, after the inoculation of strain DDT-1 for the determination of p,p′-DDT residues.
mineralized to carbon dioxide (Fang et al., 2016). The objectives of this study were 1) to examine the degradation ability and adaptability of strain DDT-1 to fresh p,p′-DDT residues in different types of soils under laboratory conditions; 2) to investigate the feasibility of using strain DDT-1 to remove or detoxify aged DDTs residues in the field plot soils; 3) to conduct a field demonstration of DDTs biodegradation by strain DDT-1 in the greenhouse and open field soils under different planting and farming conditions; and 4) to assess the ecological safety of in situ bioremediation of the DDT-contaminated soils by strain DDT-1. This study will develop a promising and safe technology for in situ bioremediation of DDT-contaminated soils. 2. Materials and methods
2.5. DDTs residues in the field soil 2.1. Chemicals One hundred forty seven soil samples were collected by a five-point sampling method from a vegetable base located in Cixi, Zhejiang, China. The soils were passed through a 2-mm sieve to remove plant materials and debris after air-drying at room temperature, and then stored at − 20 °C until the determination of the DDTs residues.
Standard samples of p,p′-DDT (purity ≥ 99.5%), p,p′-DDE (purity≥99.5%), p,p′-DDD (purity ≥ 99.5%), and o,p′-DDT (purity ≥ 99.5%) were provided by Dr. Ehrenstorfer (Augsburg, Germany). Anhydrous sodium sulfate of analytical grade was provided by Shuanglin Chemical Co. (Hangzhou, China). Analytical grade of acetone, n-hexane, and sodium chloride were provided by China National Chemical Reagent Corporation (Jiangsu, China). All other chemical reagents used in the experiment were analytical reagent grade.
2.6. Field plot trials To evaluate the feasibility of strain DDT-1 to remove or detoxify DDTs residues in soil, a bioremediation study with a completely randomized block design was performed in the vegetable field located in Cixi, Zhejiang, China. Each block covered an area of 6 m2 (2 m × 3 m) and was separated with PVC boards. A suspension of strain DDT-1 amended with 2% yeast powder was sprayed evenly with water at an inoculation level of approximately 1.0 × l08 CFUs/g using an electric sprayer. Meanwhile, the surface soils (0–10 cm) were tilled to ensure the homogenous distribution of the inocula. The control experiment without inoculation was performed under the same conditions. All plots were planted with corn using the same field management measures. Four successive inoculation treatments were performed every 30 d to ensure the survival and colonization of strain DDT-1 in the field soil. Each plot was performed in triplicate. Each soil subsample consisted of five soil cores, and six surface soil subsamples (0–10 cm, 50 g for each) were randomly collected from each plot using a stainless-steel manual auger. All subsamples from each plot were combined into a single soil sample. Soil samples were collected from all plots at 0, 15, 45, 90, 120, 150, and 210 d, respectively. Five soil subsamples from each plot were thoroughly mixed to obtain a composite sample. Subsequently, all soil samples were sieved (2 mm) to remove plant materials and debris, and then stored at − 20 °C until the determination of the DDTs residues.
2.2. Soils The fresh surface soil samples (0–10 cm) used in this study were collected from two paddy fields located at Xiaoshan (XS) and Jinhua (JH) and a mulberry field located on the Huajiachi (HJC) campus of Zhejiang University, Huangzhou, China. All soil samples were sieved (2 mm) to remove stones and debris after air-drying at room temperature. The residues of DDTs were below limit of detection (LOD) in all soil samples. The physicochemical properties of the soil samples were determined by the Zhejiang Academy of Agricultural Sciences, Hangzhou, China, and are summarized in Table S1. 2.3. Inocula preparation Stenotrophomonas sp. DDT-1 could utilize DDT as its sole carbon and energy source (Fang et al., 2016). Strain DDT-1 was repetitively cultured in the Luria-Bertoni (LB) agar (yeast extract, 5 g; peptone, 5 g; NaCl, 5 g; agar, 20 g; distilled water, 1000 ml; pH 7.2) plates, and a single colony was placed into 100 ml of sterilized LB liquid media and incubated for 24 h on a rotary shaker at 25 °C and 150 r/min in the dark. Then, the colony was transferred into a centrifuge tube and centrifuged for 10 min at 7000 r/min. The supernatant was removed and the remaining was immediately washed 3 times with 40 ml of NaH2PO4-Na2HPO4 buffer (0.1 mol/l, pH 7.0), and ultimately suspended in the same phosphate buffer as the inocula.
2.7. Field demonstration application The field demonstration areas for the strain DDT-1 preparation were divided into 12 experimental zones and designated as B1-B12, including greenhouse and open field soils, and their related information is summarized in Table S2. Each experimental zone included strain DDT-1 inoculation treatment and the corresponding un-inoculated control treatment, and each treatment was replicated at least six times. The suspension of strain DDT-1 supplemented with 2% yeast powder was evenly sprayed with water using an electric sprayer at a dose of 800 L/ ha, and the inoculation level of strain DDT-1 reached 2.0 × 106 CFUs/g dry soil in the greenhouse and open field soils. The addition of yeast powder provided a nitrogen source and energy source for the initial survival and propagation of strain DDT-1 in the inoculated greenhouse and open field soils. Four successive inoculation treatments were performed every 30 d, and mechanical tillage was carried out on the demonstration areas of B7 and B11 during the inoculation of the strain DDT-1 preparation. The un-inoculated areas receiving the same amount of distilled water were used as the controls. Each soil subsample consisted of five soil cores, and six surface soil subsamples (0–10 cm, 100 g
2.4. Biodegradation of DDTs in laboratory soils Soil samples (0.4 kg, dry weight equivalent) were divided into two equal parts. The first part was transferred into a 5-L plastic basin, and the standard solution of p,p′-DDT in acetone at a concentration of 400 mg/l was sprayed on the soil using a small sprinkling can in combination with an appropriate amount of sterile distilled water. Subsequently, the second part was also transferred into this plastic basin. All soil samples were thoroughly stirred using a plastic spoon and passed through a 2-mm sieve to ensure uniform distribution of the added p,p′-DDT at an initial concentration of 1 mg/kg dry weight soil, and were then kept for 1 h on a laminar flow bench to evaporate the solvent. Meanwhile, the strain DDT-1 preparation was inoculated into the soils at an inoculation level of 6.0 × 1010 colony-forming units 146
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2.11. DNA extraction and MiSeq sequencing
for each) were randomly collected from each plot using a stainless-steel manual auger. All subsamples from each experimental zone were combined into a soil sample. Soil samples were collected from all plots at 0, 30, 60, 90, 120, 180, 240, 270, 360, and 400 d, respectively. Subsequently, each sample was sieved (2 mm) to remove plant materials and debris after air-drying at ambient temperature and stored at − 20 °C until the determination of the DDTs residues.
The total DNA was extracted from 0.5 g of soil sample using a nucleic acid extractor and FastDNA SPIN Kit for soil (Qbiogene, Carlsbad, CA, USA) according to the manufacturer's protocol. The purity and concentration of the extracted DNA were determined using a spectrophotometer (NanoDrop ND-1000, Wilmington, DE, USA). The extracted DNA was further analyzed by 0.8% agarose gel electrophoresis (with TAE buffer containing 0.8 µg/ml of ethidium bromide) at a constant voltage of 120 V for 20 min. The 16 S rDNA V4 region was amplified using the universal forward primer 520F (5′-AYTGGGYDTAAAGNG-3′) and the reverse primer 802R (5′-TACNVGGGTATCTAATCC-3′), and a barcode unique to each sample was added to the reverse primer to allow multiplex sequencing. The PCR reaction system contained 25.0 μl, which included 1.0 μl of the 20 ng/μl template, 2.5 μl of buffer (10 ×), 2.0 μl of 2.5 mM dNTP, 1.0 μl of each primer (10 μM), 0.125 μl of Taq DNA polymerase (5 U/μl), and 17.375 μl of ultra-pure water. The PCR running conditions consisted of an initial denaturation at 94 °C for 4 min followed by 30 cycles of 94 °C for 30 s, 50 °C for 30 s, 72 °C for 30 s and a final extension at 72 °C for 5 min. Subsequently, the bacterial 16S amplicon sequencing was performed under the Illumina MiSeq platform (2 × 250 bp, San Diego, CA, USA) by the Personal Biotechnology Co. (Shanghai, China).
2.8. Extraction and determination of DDTs residues in soils The residues of DDTs in the soils were extracted following the method described by Fang et al. (2010). The concentrations of DDTs were determined by an Agilent 6890 N gas chromatography (Agilent Technologies, USA) equipped with an electron capture detector (µ-ECD) and a DB-1701 silica capillary column (30 m × 0.32 mm × 0.25 µm, Agilent Technologies, USA). The oven temperature was initially 160 °C and increased to 220 °C at a rate of 10 °C/min and then rose to 240 °C at a rate of 5 °C/min with maintenance of the final temperature for 2 min. The injector and detector temperatures were set at 230 °C and 280 °C, respectively. The injection mode was splitless and the injection volume was 1.0 μl. The flow rate of the helium carrier gas was set at 2 ml/min. 2.9. Soil enzyme activity assays
2.12. Recovery study
The catalase activity was evaluated based on the recovery rate of hydrogen peroxide. Soil samples (2.0 g dry soil weight) were mixed with 40 ml of distilled water and 5 ml of 0.3% H2O2 in a 100-ml volumetric flask, and then shaken for 30 min at 150 rpm and 30 °C in the dark. After shaking, 5 ml of H2SO4 (1.5 mol/l) was added to the flask to stabilize the un-decomposed hydrogen peroxide, and the sample was filtered through 7-cm quantitative dense filter paper. Subsequently, 25 ml of the filtrates were titrated with 0.02 mol/l KMnO4 and the results are expressed as ml KMnO4/(h·g dry soil). Meanwhile, control experiments without soil samples were conducted using the same procedure. The neutral phosphatase activity was measured referred to the method described by Rahmatpour et al. (2017). Soil samples (1.0 g, dry soil weight), 0.2 ml of toluene, 4 ml of sodium hydrogen phosphatecitric acid buffer (pH 7.0), and 1 ml of p-nitrophenyl sodium hydrogen phosphate solution (0.025 mol/l) were mixed in a 100-ml volumetric flask. The mixture was incubated for 1 h at 35 °C in the dark and the reaction was stopped by adding 1 ml of CaCl2 solution (0.5 mol/l) and 4 ml of NaOH solution (0.5 mol/l). The suspension was shaken for a few seconds and filtered, and the color intensity at 420 nm was determined using a spectrophotometer. The controls were unamended with p-nitrophenyl sodium hydrogen phosphate solution and were processed using the same procedure. The results were modified by the calibration plot of the p-nitrophenol (PNP) standard, and the neutral phosphatase activity is expressed as μg PNP/(h·g dry soil)
To evaluate the validity of the extraction method for DDTs residues, a recovery experiment was performed at spiking levels of 0.1, 1, and 10 mg/kg dry weight soil. The same extraction and detection methods for DDTs residues were used as described above. 2.13. Statistical analysis Analysis of variance (ANOVA) of the degradation half-lives and rates between the inoculated soils and un-inoculated controls was performed using SPSS 19.0 (SPSS, Chicago, IL, USA). The average values and standard deviations of the data were calculated using Microsoft Excel 2007 (Microsoft Corporation, Redmond, WA, USA). The average well color development (AWCD), Shannon index (H′), McIntosh index (U), and Simpson index (1/D) were calculated according to the methods described by Fang et al. (2014). To distinguish the differences in the diversity and abundance of the soil microbial communities after inoculation of strain DDT-1, heat maps of the top 30 soil bacterial genera were visualized using Matlab 7.0 (Mathworks, Natick, MA, USA). 3. Results and discussion 3.1. Evaluation of the recovery The recoveries of DDTs at spiking concentrations (0.1, 1.0 and 10.0 mg/kg) ranged from 91.9% to 100.7% with relative standard deviation (RSD) ≤ 4.4% in the indoor soils, and 87.1–100.7% with RSD ≤ 0.7% in the field soils. The LOD for p,p′-DDT, p,p′-DDE, p,p′-DDD, and o,p′-DDT residues in the soil was 0.005 mg/kg, 0.005 mg/kg, 0.005 mg/kg, and 0.005 mg/kg, respectively. The corresponding limit of quantitation (LOQ) was 0.001 mg/kg, 0.001 mg/kg, 0.001 mg/kg, and 0.001 mg/kg, respectively. These data indicate that the extraction method was satisfactory for the analysis of DDTs residues.
2.10. BIOLOG assay Five grams of the soil samples (dry-weight equivalent) were mixed with 45 ml of physiological saline (0.85%, w/v) in a 100-ml triangular flask and shaken for 1 h on an orbital shaker at 25 °C and 150 r/min in the dark. Subsequently, the mixtures were allowed to settle for 30 min, and a portion of the supernatant was serially diluted using sterile physiological saline in a sterile tube to obtain a 10−3 dilution. Each micropore in the BIOLOG ECO microplate™ (BIOLOG Inc., Hayward, USA) was inoculated with 150 μl of the 10−3 dilution, and all microplates were placed in a thermostat incubator for 168 h at 25 ± 1 °C. Color development in the wells was measured at 4, 24, 48, 72, 96, 120, 144, and 168 h using the BIOLOG reader (BIO-TEK, Winooski, VT, USA) at wavelengths of 750 nm and 590 nm. The output data were recorded by the automatic threshold option through BIOLOG software.
3.2. DDTs residues in the field soils The residual levels of p,p′-DDT, p,p′-DDE, p,p′-DDD, o,p′-DDT and DDTs in the field soils are shown in Figs. S1-S5. p,p′-DDT, p,p′-DDE, p,p′DDD and o,p′-DDT were detected in almost all field soils, and the residual concentration of p,p′-DDE was much higher than other three homologues in concentration, indicating that p,p′-DDT was mainly degraded to p,p′147
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DDE in the field soils under aerobic conditions (Huang et al., 2018; Zhang et al., 2018). Furthermore, the ratio of p,p′-DDT/p,p′-DDE in the field soils was less than 1, with a range of 0.01–0.67, suggesting that DDTs residues may be attributed to its substantial use in cotton fields prior to banning. As shown in Fig. S1, the residual levels of DDTs in the field soils were 0.06–1.02 mg/kg, and the mean residual concentration was 0.31 mg/kg, which showed that this investigation area has been seriously polluted by DDTs. The residues of DDTs in soils have been reported extensively. Fang et al. (2010) reported that the residual level of DDTs was 0.14 mg/kg in the pumpkin-growing field soils located in Cixi, China, and these residues of DDTs may have resulted from extensive use of DDT during the 1960s and 1970s. Zhu et al. (2005) reported that the mean level of DDTs residues was 0.14 mg/kg in the shallow subsurface soils in the outskirts of Beijing. Neitsch et al. (2016) also reported that the total concentration of p,p′-DDT, p,p′-DDE, and p,p′-DDD was 0.12 mg/kg in a conventionally farmed land located in Germany.
Fig. 2. Degradation of DDTs residues in the field plot soils after inoculation of strain DDT-1. Data followed by a different letter in the same treatment time are significantly different (p ≤ 0.05).
3.3. Biodegradation of DDT by strain DDT-1 in different soils under laboratory conditions
degradation percentages of p,p′-DDT by strain DDT-1 were 24.5%, 21.0%, and 20.9% in HJC, XS, and JH soils, respectively, and the corresponding percentages were only 9.5%, 8.9% and 8.4% in the un-inoculated control samples, which showed the much higher degradation percentages of p,p′-DDT in inoculated soils compared with those in the un-inoculated control soils. As shown in Table S3, the degradation rates of p,p′-DDT in strain DDT-1 inoculated HJC, XS, and JH soils were 2.7-, 2.0-, and 3.0-fold higher, respectively, than those in the un-inoculated controls. The degradation half-lives of p,p′-DDT in strain DDT-1 inoculated HJC, XS, and JH soils were 2.4-, 1.5-, and 3.8-fold lower than those in the un-inoculated controls. The significantly higher degradation rate in the inoculation treatment than in the un-inoculated controls indicated that strain DDT-1 could efficiently degrade p,p′-DDT residues in soils, and furthermore, this bacterium could be widely adapted to different types of soil. Similarly, Gao et al. (2015) reported that approximately 60.1%, 61.6%, 63.5%, and 65.0% of the initial p,p′-DDT was degraded by the plasmid donor bacteria E. coli TGI(pDOD-gfp) in JH, JX, XS and HZ soils, respectively. Purnomo et al. (2011) demonstrated the ability of brown-rot fungi G. trabeum to degrade DDT in historically contaminated soils under laboratory conditions. Additionally, the biodegradation of p,p′DDT in the HJC, XS, and JH soils was correlated to the soil physicochemical properties. As shown in Table S1 and Fig. 1, a significant positive correlation (R = 0.997) between the biodegradation rate of p,p′-DDT and soil organic matter content was found in all soils. Han et al. (2016) also found a similar positive correlation between the degradation half-life of p,p′-DDT and soil organic matter content as well as soil clay mineral content.
The degradation of p,p′-DDT by strain DDT-1 in different soils under laboratory conditions is shown in Fig. 1. After inoculation for 28 d, the
3.4. Biodegradation of DDTs residues by strain DDT-1 in field plot soils The biodegradation of DDTs residues in the field plot soils after inoculation of strain DDT-1 is shown in Fig. 2. After inoculation for 210 d, the degradation percentage of DDTs was 38.0%, but the corresponding value was only 11.1% in the un-inoculated controls. As shown in Fig. 2 and Fig. S6, the degradation percentage of DDTs in the inoculated soils was 1.5–3.6-fold higher than that in the un-inoculated control soils. As shown in Table S4, the degradation half-lives of p,p′DDT, p,p′-DDE, p,p′-DDD, o,p′-DDT, and DDTs in the inoculated soils were 47.4%, 18.3%, 33.3%, 68.0%, and 34.0%, respectively, of those in the corresponding un-inoculated soils. The corresponding degradation rates in the inoculated soils were 1.5-, 3.9-, 2.3-, 1.3-, and 2.5-fold higher than those in the un-inoculated soils, which indicated more rapid degradation in the inoculated soils than in the un-inoculated soils. The incorporation of strain DDT-1 into field plot soils could efficiently remove DDTs residues. Several similar studies on the
Fig. 1. Degradation of p,p′-DDT in different types of soils inoculated with strain DDT-1 under laboratory conditions. Data followed by a different letter in the same treatment time from each subfigure are significantly different (p ≤ 0.05). 148
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biodegradation of DDTs in field soils have been conducted. Qu et al. (2015) reported that 80.4% of DDT was removed by strain Chryseobacterium sp. PYR2 in a pilot-scale field soils after inoculation for 45 d. Fang et al. (2010) reported that 56.05% of DDTs were degraded by strain Sphingobacterium sp. D6 in the pumpkin-grown field soils. Wang et al. (2010) also reported that the inoculation of the DDT-degrading strain Pseudoxanthomonas sp. Wax into fine sandy-loam soils at an inoculation level of 108 CFUs/g could thoroughly eliminate 20 mg/kg of DDTs after treatment for 20 d. Some researchers reported that the survival and propagation of the inocula determined the success of DDTs bioremediation. However, the biodegradation activity of the inocula after inoculation in field soils depended largely on some factors, including competition and antagonism by indigenous microorganisms, predation by protozoa, soil physicochemical properties (moisture, pH, aeration, clay content, organic matter content, salinity etc.), environmental conditions (temperature, humidity, rainfall, solar irradiation etc.), and electron acceptors as well as the bioavailability of DDTs (Baczynski et al., 2010; Fang et al., 2010; Purnomo et al., 2011). Additionally, the similar high biodegradation ability of strain DDT-1 for DDTs in the field plot soils and laboratory soils indicated that strain DDT-1 could be applied to the field for the removal of DDTs residues.
Fig. 3 and Figs. S7 and S8. The residual concentration of DDTs did not decrease significantly in any of the un-inoculated controls. After inoculation for 420 d, the mean degradation percentages of p,p′-DDT, p,p′-DDE, and DDTs were 84.9%, 36.7%, and 42.1% in the open field soils, and 76.1%, 38.0%, and 38.7% in the greenhouse soils, respectively. The mean degradation rates of p,p′-DDT, p,p′-DDE, and DDTs in the inoculated greenhouse soils were 1.08-, 1.04-, and 1.08-fold higher than those in the inoculated open field soils. As shown in Tables S5-S7, the degradation half-lives of p,p′-DDT, p,p′-DDE, and DDTs were 15.3–77.0%, 12.6–27.7%, and 10.9–32.5% (open field soils) and 30.8–39.4%, 16.7–28.8%, and 20.6–30.8% (greenhouse soils) shorter in the inoculation treatments than those in the un-inoculated controls. The degradation rates of p,p′-DDT, p,p′-DDE, and DDTs were 1.1–2.4, 2.7–6.3, and 2.8–7.6 (open field soils) and 1.5–1.8, 3.2–5.2, and 2.9–5.5 (greenhouse soils) fold higher in the inoculation treatments than those in the un-inoculated controls. The application of strain DDT-1 preparation could accelerate the degradation of DDTs residues in the greenhouse and open field soils. A similar result has been reported that soil inoculation with strain DDT-1 could decrease the DDTs concentration from 0.489 mg/kg to 0.265 mg/ kg within 18 months in a DDTs-contaminated agricultural site (Zhu et al., 2012). Additionally, the degradation rate of DDTs in the inoculated greenhouse soils was higher than that in inoculated open field soils, which may be attributed to the relatively closed environment of the greenhouse.
3.5. Field demonstration of DDTs biodegradation by strain DDT-1 The biodegradation of p,p′-DDT, p,p′-DDE, and DDTs residues in the strain DDT-1 inoculated soils under field conditions is presented in
Fig. 3. Degradation of DDTs residues in the greenhouse and open field soils after inoculation of strain DDT-1 preparation. B1: open field planted soil; B2: greenhouse/open field planted soil; B3: open field unplanted soil; B4: open field unplanted soil; B5: open field planted soil; B6: greenhouse planted soil without mechanical tillage; B7: greenhouse planted soil with mechanical tillage; B8: greenhouse planted soil; B9: open field planted soil without mechanical tillage; B10: open field planted soil; B11: open field planted soil with mechanical tillage; and B12: open field planted soil. The two curves followed by a different letter in each subfigure are significantly different (p ≤ 0.05).
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Fig. 4. Variations in soil catalase activity (a) and neutral phosphatase activity (b) after inoculation of strain DDT-1 into greenhouse and open field soils. The two curves followed by the same letter in each subfigure are not significantly different (p ≤ 0.05).
3.7. Variation of soil microbial functional diversity after inoculation of strain DDT-1
3.6. Variation of soil enzyme activities after inoculation of strain DDT-1 Soil enzyme activity is related to soil organic matter decomposition and nutrient cycling, reflecting the intensity of soil microbial processes to a certain extent (Trasar-Cepeda et al., 2000; Jin et al., 2009). Changes in enzyme activities in the greenhouse and open field soils after inoculation of strain DDT-1 preparation are shown in Fig. 4. No significant differences in the catalase activity were found between the inoculated field soils and the corresponding un-inoculated field soils throughout the entire bioremediation period (Fig. 4a). A similar pattern in the neutral phosphatase activity was also observed, even though there was a slight reduction at 200 d in the open field planted soils (B5) and a slight increase at 300 d in the open field untilled soils (B9) in inoculation treatments compared to that of the corresponding un-inoculated controls (Fig. 4b). In this study, therefore, the inoculation of strain DDT-1 into the field soils did not significantly affect soil enzyme activities, including catalase and neutral phosphatase activities. Similar to our results, no significant differences in the soil catalase activity were found between strain DXZ9 inoculated-soils and the un-inoculated control soils after treatment for 210 d (Xie, 2013). Huang (2012) also reported that the inoculation of strain DDT-1 into aged DDT-contaminated soils only had a slight stimulation effect on the soil phosphatase activity.
The average well color development (AWCD), which reflects the ability of soil microbial communities to utilize different carbon sources, is commonly used to assess the overall activity of soil microbial communities (Wu et al., 2014). As shown in Fig. 5a, 0 d after inoculation of strain DDT-1, the AWCD values in all inoculated greenhouse and open field soils were higher than those in the corresponding un-inoculated greenhouse and open field soils, suggesting that the overall activities of the soil microbial communities were stimulated in the early stage of inoculation. After inoculation for 210 d and 420 d, no statistically significant difference in the AWCD values was observed between the inoculated greenhouse and open field soils and the corresponding uninoculated control soils (Fig. 5b-c), indicating that the overall soil microbial activities recovered to the control level in the middle and later stages of inoculation. Therefore, the effect of strain DDT-1 inoculation on the overall soil microbial activities gradually disappeared over the inoculation time. Soil microbial diversity indices, including the Simpson index (1/D), Shannon index (H′), and McIntosh index (U), are usually used to evaluate the dominant population, richness and evenness of soil microorganisms, respectively (Fang et al., 2015). Soil microbial diversity indices after inoculation of the strain DDT-1 preparation are summarized in Table 1. Immediately after inoculation of strain DDT-1, the 1/D 150
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Fig. 5. Variations in average well color development (AWCD) at 0 d (a), 180 d (b), and 420 d (c) in the greenhouse and open field soils after inoculation of the strain DDT-1 preparation. The two curves followed by the same letter in each subfigure are not significantly different (p ≤ 0.05).
values in all inoculated greenhouse and open field soils were slightly higher than those in the corresponding un-inoculated control soils, indicating that the dominant soil microbial populations increased in the inoculated greenhouse and open field soils. Subsequently, the 1/D values in all inoculated soils gradually decreased at 180 d after inoculation and then recovered to a level similar to that of the control soils
420 d after inoculation, suggesting that the dominant soil microbial populations recovered to the control level. However, the H′ and U values in all inoculated greenhouse and open field soils were not statistically different from those in the corresponding un-inoculated control soils, indicating that soil microbial evenness and richness were not significantly affected in the middle and later stages of inoculation.
Table 1 Variations in the soil microbial functional diversity indices after inoculation of strain DDT-1 into the greenhouse and open field soils. Time (d)
Treatment
Planted pattern
Shannon index (H′)
Simpson index (1/D)
McIntosh index (U)
0 0 0 0 0 0 0 0 0 0 0 0 180 180 180 180 180 180 180 180 180 180 180 180 420 420 420 420 420 420 420 420 420 420 420 420
B3-CK B3-BR B5-CK B5-BR B6-CK B6-BR B7-CK B7-BR B9-CK B9-BR B11-CK B11-BR B3-CK B3-BR B5-CK B5-BR B6-CK B6-BR B7-MT-CK B7-MT-BR B9-CK B9-BR B11-CK B11-BR B3-CK B3-BR B5-CK B5-BR B6-CK B6-BR B7-CK B7-BR B9-CK B9-BR B11-MT-CK B11-MT-BR
Open field-unplanted Open field-unplanted Open field-planted Open field-planted Greenhouse-unplanted Greenhouse-unplanted Greenhouse-planted Greenhouse-planted Open field-unplanted Open field-unplanted Open field-planted Open field-planted Open field-unplanted Open field-unplanted Open field-planted Open field-planted Greenhouse-untilled Greenhouse-untilled Greenhouse-tilled Greenhouse-tilled Open field-untilled Open field-untilled Open field-tilled Open field-tilled Open field-unplanted Open field-unplanted Open field-planted Open field-planted Greenhouse-untilled Greenhouse-untilled Greenhouse-tilled Greenhouse-tilled Open field-untilled Open field-untilled Open field-tilled Open field-tilled
2.38 2.51 2.58 2.42 2.14 2.08 2.76 2.30 1.92 2.26 2.51 2.43 1.79 2.36 1.52 1.68 1.87 2.00 1.64 1.92 2.24 2.65 2.01 2.59 3.06 3.01 2.77 3.13 3.12 3.02 2.78 2.96 3.16 3.14 2.89 3.04
7.73 ± 1.23 a 11.29 ± 0.76 b 9.21 ± 0.55 a 10.09 ± 0.49 b 4.36 ± 1.47 a 7.28 ± 1.57 b 12.28 ± 1.11 a 12.93 ± 0.26 a 6.58 ± 2.27 a 7.11 ± 1.58 b 10.13 ± 2.67 a 11.78 ± 2.57 a 4.04 ± 2.55 a 9.07 ± 0.50 b 4.70 ± 2.84 a 5.32 ± 3.58 a 6.01 ± 1.51 a 6.50 ± 0.37 a 4.35 ± 1.81 a 5.00 ± 1.53 a 8.12 ± 3.88 a 9.49 ± 2.96 a 6.32 ± 0.83 a 10.44 ± 0.76 b 18.84 ± 2.46 a 21.12 ± 2.32 a 16.85 ± 2.46 a 18.67 ± 2.62 a 16.80 ± 0.78 a 19.44 ± 1.50 a 17.43 ± 2.17 a 18.91 ± 0.83 a 18.97 ± 0.98 a 20.00 ± 1.84 a 18.27 ± 2.17 a 19.87 ± 1.06 a
1.32 2.06 2.12 1.89 0.90 1.81 2.30 1.61 1.78 1.26 1.94 1.87 2.23 1.09 1.09 1.14 1.06 1.38 1.12 0.92 1.51 1.32 1.26 1.75 4.40 4.33 3.09 3.71 3.96 3.28 3.78 3.29 4.32 4.11 3.97 4.00
± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ±
0.35 0.20 0.10 0.04 0.36 0.34 0.10 0.04 0.69 0.15 0.29 0.14 0.29 0.37 0.78 0.78 0.32 0.43 0.21 0.29 1.28 0.28 0.16 0.07 0.24 0.12 0.45 0.25 0.06 0.17 0.39 0.16 0.08 0.15 0.29 0.14
a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a
Data followed by a different letter in the same column are significantly different (p ≤ 0.05). 151
± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ±
0.35 0.31 0.13 0.19 0.58 0.66 0.39 0.23 1.02 0.25 0.23 0.59 1.17 0.94 0.59 0.63 0.17 0.47 0.59 0.26 0.89 0.42 0.31 0.18 0.13 0.34 0.76 0.33 0.13 0.08 0.63 0.35 0.25 0.31 0.55 0.36
a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a
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Fig. 6. Heat maps of the top 30 soil microbial genera at 0 d (a), 180 d (b), and 42 d (c) in the greenhouse and open field soils after inoculation of the strain DDT-1 preparation. 1–12 represent B3-CK (open field unplanted soil-control), B3-BR (open field unplanted soil-inoculation), B5-CK (open field planted soil-control), B5-BR (open field planted soil-inoculation), B6-CK (greenhouse untilled soil-control), B6-BR (greenhouse untilled soil-inoculation), B7-MT-CK (greenhouse soil-mechanical tillage-control), B7-MT-BR (greenhouse soil-mechanical tillage-inoculation), B9-CK (open field untilled soil-control), B9-BR (open field untilled soil-inoculation), B11-MT-CK (open field soil-mechanical tillage-control), and B11-MT-BR (open field soil-mechanical tillage-inoculation), respectively.
later stages of strain inoculation. Therefore, strain DDT-1 might be efficient and ecologically safe for in situ bioremediation of the DDTcontaminated greenhouse and open field soils. Further studies need to be carried out on the effects of seasonal and geographical variation on the colonization and adaptation abilities of strain DDT-1 in the field soils.
3.8. Variation in soil microbial structural diversity after inoculation of strain DDT-1 The heat maps of the top 30 soil microbial genera at 0 d, 180 d and 420 d after inoculation of strain DDT-1 are shown in Fig. 6. After inoculation of strain DDT-1, the dominant soil bacterial genera were Bacillus, Sphingobacterium, Stenotrophomonas, and Nocardioides at 0 d (Fig. 6a); Bacillus, Stenotrophomonas, Sphingobacterium, Nocardioides, and Nitrospira at 180 d (Fig. 6b); and Nitrospira, Nocardioides, Bacillus, Stenotrophomonas, and Sphingobacterium at 420 d (Fig. 6c). Throughout the entire bioremediation period, the soil microbial structural diversity was generally similar between the inoculated greenhouse and open field soils and the corresponding un-inoculated control soils, but several soil microbial genera had some fluctuations in their relative abundances. For instance, an initial increase followed by a decline in the abundance of Stenotrophomonas was observed with the extension of the inoculation time, and that of Nitrospira increased in the later stage of inoculation. The principal component analysis (PCA) of soil microbial communities after inoculation of strain DDT-1 is shown in Fig. S9. The PC1 component contributed more than the PC2 component in the PCA profile throughout the entire bioremediation period. As shown in Fig. S9, a distinctly dispersed pattern was presented at 0 d after strain inoculation, and then a cluster was gradually formed at 180 d after strain inoculation. Finally, a scattered pattern was observed at 420 d after strain inoculation. The results showed that the structural diversity of soil microbial communities was temporarily suppressed in the initial stage of strain inoculation but gradually recovered to the similar level in the un-inoculated controls over the inoculation time.
Acknowledgements This work was supported by the Zhejiang Provincial National Science Foundation of China (No. LY18B070001) and the National Key Research and Development Program of China (Nos. 2016YFD0200205 and 2016YFD0200201). Appendix A. Supporting information Supplementary data associated with this article can be found in the online version at http://dx.doi.org/10.1016/j.ecoenv.2018.04.026. References Baczynski, T.P., Pleissner, D., Grotenhuis, T., 2010. Anaerobic biodegradation of organochlorine pesticides in contaminated soil–significance of temperature and availability. Chemosphere 78 (1), 22–28. Bajaj, A., Mayilraj, S., Mudiam, M.K.R., Patel, D.K., Manickam, N., 2014. Isolation and functional analysis of a glycolipid producing Rhodococcus sp. strain IITR03 with potential for degradation of 1, 1, 1-trichloro-2, 2-bis (4-chlorophenyl) ethane (DDT). Bioresour. Technol. 167, 398–406. Chen, M.J., Cao, F., Li, F.B., Liu, C.S., Tong, H., Wu, H.J., Hu, M., 2013. Anaerobic transformation of DDTs related to biogenic Fe(II) and microbial community activities in paddy soils. J. Agic. Food Chem. 61, 2224–2233. Chen, M.J., Liu, C.S., Chen, P.C., Tong, H., Li, F.B., Qiao, J.T., Lan, Q., 2016a. Dynamics of the microbial community and Fe(III)-reducing and dechlorinating microorganisms in response to pentachlorophenol transformation in paddy soil. J. Hazard. Mater. 312, 97–105. Chen, M.J., Tong, H., Liu, C.S., Chen, D.D., Li, F.B., Qiao, J.T., 2016b. A humic substance analogue AQDS stimulates Geobacter sp. abundance and enhances pentachlorophenol transformation in a paddy soil. Chemosphere 160, 141–148. Fang, H., Dong, B., Yan, H., Tang, F.F., Yu, Y.L., 2010. Characterization of a bacterial strain capable of degrading DDT congeners and its use in bioremediation of contaminated soil. J. Hazard. Mater. 184 (1), 281–289. Fang, H., Han, Y.L., Yin, Y.M., Jin, X.X., Wang, S.Y., Tang, F.F., Cai, L., Yu, Y.L., 2014. Microbial response to repeated treatments of manure containing sulfadiazine and chlortetracycline in soil. J. Environ. Sci. Health Part B-Pestic. Contam. Agric. Wastes
4. Conclusions The inoculation of strain DDT-1 into soils could significantly enhance the degradation of DDTs residues under laboratory and field conditions. The much higher biodegradation percentage and biodegradation rate of DDTs residues were found in the inoculated greenhouse and open field soils compared to the corresponding un-inoculated control soils. No significant effects were found on the soil enzyme activity, microbial functional and structural diversity in the middle and 152
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49 (8), 609–615. Fang, H., Lian, J.J., Cai, L., Yu, Y.L., 2015. Exploring bacterial community structure and function associated with atrazine biodegradation in repeatedly treated soils. J. Hazard. Mater. 286, 457–465. Fang, H., Yu, Y.L., Pan, X., Lin, D.L., Zheng, Y., Zhang, Q., Yin, Y.M., Cai, L., 2016. Biodegradation of DDT by Stenotrophomonas sp. DDT-1: characterization and genome functional analysis. Sci. Rep. 6, 21332. Gao, C.M., Jin, X.X., Ren, J.B., Fang, H., Yu, Y.L., 2015. Bioaugmentation of DDT-contaminated soil by dissemination of the catabolic plasmid pDOD. J. Environ. Sci. 27, 42–50. Grewal, J., Bhattacharya, A., Kumar, S., Singh, D.K., Khare, S.K., 2016. Biodegradation of 1,1,1-trichloro-2,2-bis (4-chlorophenyl) ethane (DDT) by using Serratia marcescens NCIM 2919. J. Environ. Sci. Health Part B-Pestic. Contam. Agric. Wastes 51 (12), 809–816. Han, Y.L., Shi, N., Wang, H.F., Pan, X., Fang, H., Yu, Y.L., 2016. Nanoscale zerovalent iron-mediated degradation of DDT in soil. Environ. Sci. Pollut. Res. 23 (7), 6253–6263. Huang, H.F., Zhang, Y., Chen, W., Chen, W.W., Yuen, D.A., Ding, Y., Chen, Y.J., Mao, Y., Qi, S.H., 2018. Sources and transformation pathways for dichlorodiphenyltrichloroethane (DDT) and metabolites in soils from Northwest Fujian, China. Environ. Pollut. 235, 560–570. Huang, H.G., 2012. Processes and Mechanisms of Agronomic Factors for Enhancing Phytoremediation of Cd-Zn/DDTs Co-contaminated Soils (Ph.D. Dissertation). College of environment and resource science, Zhejiang University, China. Jin, K., Sleutel, S., Buchan, D., Deneve, S., Cai, D.X., Gabriels, D., Jin, J.Y., 2009. Changes of soil enzyme activities under different tillage practices in the Chinese Loess Plateau. Soil Tillage Res. 104 (1), 115–120. Mcglynn, K.A., Quraishi, S.M., Graubard, B.I., Weber, J.P., Rubertone, M.V., Erickson, R.L., 2008. Persistent organochlorine pesticides and risk of testicular germ cell tumors. Natl. Cancer Inst. 100 (9), 663–671. Neitsch, J., Schwack, W., Weller, P., 2016. How do modern pesticide treatments influence the mobility of old incurred DDT contaminations in agricultural soils? J. Agric. Food Chem. 64 (40), 7445–7451. Pan, X., Xu, T.H., Xu, H.Y., Fang, H., Yu, Y.L., 2017. Characterization and genome functional analysis of the DDT-degrading bacterium Ochrobactrum sp DDT-2. Sci. Total Environ. 592, 593–599. Purnomo, A.S., Mori, T., Takagi, K., Kondo, R., 2011. Bioremediation of DDT contaminated soil using brown-rot fungi. Int. Biodeterior. Biodegrad. 65 (5), 691–695. Qu, J., Xu, Y., Ai, G.M., Liu, Y., Liu, Z.P., 2015. Novel Chryseobacterium sp. PYR2 degrades various organochlorine pesticides (OCPs) and achieves enhancing removal and complete degradation of DDT in highly contaminated soil. J. Environ. Manag. 161,
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Biodegradability and ecological safety assessment of Stenotrophomonas sp. DDT-1 in the DDT-contaminated soil
T
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Hua Fang, Yanfei Deng, Qiqing Ge, Jiajia Mei, Houpu Zhang, Huifang Wang, Yunlong Yu Institute of Pesticide and Environmental Toxicology, College of Agriculture & Biotechnology, Zhejiang University, Hangzhou 310058, China
A R T I C LE I N FO
A B S T R A C T
Keywords: DDT Stenotrophomonas Soil Bioremediation Ecological safety assessment
The biodegradability and ecological safety assessment of the previously isolated DDT-degrading bacterial strain Stenotrophomonas sp. DDT-1 were investigated in the DDT-contaminated soil under laboratory and field conditions. Under laboratory conditions, the degradation rates of fresh p,p′-DDT in soil were enhanced by 2.0–3.0-fold with the introduction of the strain DDT-1 compared to those of the control treatments. A similar enhancement in the dissipation of DDTs (p,p′-DDT, p,p′-DDE, p,p′-DDD, and o,p′-DDT) in the aged DDT-contaminated field plot soils resulted from the inoculation with this strain. Meanwhile, the degradation rates of DDTs increased by 2.9–5.5- and 2.8–7.6-fold in the inoculated greenhouse and open field soils, respectively, after field demonstration application of strain DDT-1 preparation. Moreover, no significant differences in the soil enzyme activity, microbial functional diversity, and bacterial community structure were observed between the inoculated and uninoculated field soils, but several soil microbial genera exhibited some fluctuations in abundance. It is concluded that strain DDT-1 could accelerate the removal of DDTs residues in field soils, and furthermore, its inoculation was ecologically safe.
1. Introduction 1,1,1-Trichloro-2,2-bis(4-chlorophenyl) ethane (DDT) has been extensively applied since the 1940s as an insecticidal agent against agricultural pests and arthropod diseases (Ren et al., 2016). DDT can be transformed into some major metabolites, including 1-dichloro-2,2-bis (4-chlorophenyl)ethylene (DDE) and 1,1-dichloro-2,2-bis-(4-chlorophenyl ethane (DDD), which may be more recalcitrant and ecologically toxic (Yu et al., 2011; Chen et al., 2013). Although the use of DDT has been prohibited in most countries since the early 1970s and in China since 1983, DDT residues and its metabolites have still been widely detected in soils, sediments, surface water, and groundwater (Fang et al., 2016). Exposure to DDT residues can cause a wide range of symptoms such as nausea, headache, vomiting, tremors, and confusion (Purnomo et al., 2011; Singh et al., 2016). Meanwhile, the high lipophilicity of DDT allows it to accumulate in food sources and fatty tissues of organisms, disrupting the central neural system and causing reproductive disorders (Mcglynn et al., 2008). Considering these negative effects, it is urgent to seek an effective remediation method to remove residual DDT. Some researchers have reported that soil microorganisms play a key role in the detoxification metabolism of organochlorine compounds (Tong et al., 2014; Chen et al., 2016a, 2016b), and thus bioremediation
⁎
has been recognized as a useful, cost-effective, and safe approach to remove or detoxify DDTs (p,p′-DDT, p,p′-DDE, p,p′-DDD, and o,p′-DDT) residues from contaminated soils (Fang et al., 2010;). The biodegradation of DDTs residues by DDT-degrading strains under laboratory conditions has been well documented (Fang et al., 2016; Pan et al., 2017). Bajaj et al. (2014) reported that a glycolipid-producing Rhodococcus sp. strain IITR03 could degrade 282 µM DDT under aqueous culture conditions. Xiao et al. (2011) found that the white rot fungi Phlebia lindtneri and Phlebia brevispora could remove 70% and 30% of DDT, respectively, in a pure culture of low-nitrogen medium after incubation for 21 d, and further degrade the metabolites DBP and DBH via hydroxylation of the aromatic ring. Ziya and Teresa (2016) demonstrated the ability of the aerobic bacterium Alcaligenes eutrophus A5 to degrade DDT in both aqueous and soil phases. Grewal et al. (2016) observed that approximately 42% of the initial 50 mg/l of DDT disappeared from a culture system after 10-d inoculation with the bacterial strain Serratia marcescens NCIM 2919. However, little information is available on the in situ bioremediation potential and ecological safety using DDT-degrading strains in contaminated field soils. In our previous study, a DDT-degrading bacterial strain DDT-1 was isolated and identified as a member of Stenotrophomonas sp., and this strain could convert DDT into DDE/DDD, DDMU, DDOH, and DDA via dechlorination, hydroxylation, and carboxylation, and ultimately
Corresponding author. E-mail address: [email protected] (Y. Yu).
https://doi.org/10.1016/j.ecoenv.2018.04.026 Received 13 December 2017; Received in revised form 26 March 2018; Accepted 12 April 2018 Available online 24 April 2018 0147-6513/ © 2018 Elsevier Inc. All rights reserved.
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(CFUs)/g and thoroughly intermixed by stirring and sieving to ensure the uniform distribution of strain DDT-1. The soil water content was adjusted to 60% of the maximum water holding capacity and maintained by periodic distilled water additions. Finally, soil samples were transferred into 500-ml plastic cups and immediately covered with aluminum foil with 5 small holes (1 mm diameter). Soil samples amended with the same volume of sterilized water were used as the controls. All plastic cups were placed in an artificial climate chamber at 25 °C in the dark. Each treatment was performed in triplicate, and 25 g of soil samples were randomly collected from each plastic cup using a plastic spoon at 0, 7, 14, 21, and 28 d, respectively, after the inoculation of strain DDT-1 for the determination of p,p′-DDT residues.
mineralized to carbon dioxide (Fang et al., 2016). The objectives of this study were 1) to examine the degradation ability and adaptability of strain DDT-1 to fresh p,p′-DDT residues in different types of soils under laboratory conditions; 2) to investigate the feasibility of using strain DDT-1 to remove or detoxify aged DDTs residues in the field plot soils; 3) to conduct a field demonstration of DDTs biodegradation by strain DDT-1 in the greenhouse and open field soils under different planting and farming conditions; and 4) to assess the ecological safety of in situ bioremediation of the DDT-contaminated soils by strain DDT-1. This study will develop a promising and safe technology for in situ bioremediation of DDT-contaminated soils. 2. Materials and methods
2.5. DDTs residues in the field soil 2.1. Chemicals One hundred forty seven soil samples were collected by a five-point sampling method from a vegetable base located in Cixi, Zhejiang, China. The soils were passed through a 2-mm sieve to remove plant materials and debris after air-drying at room temperature, and then stored at − 20 °C until the determination of the DDTs residues.
Standard samples of p,p′-DDT (purity ≥ 99.5%), p,p′-DDE (purity≥99.5%), p,p′-DDD (purity ≥ 99.5%), and o,p′-DDT (purity ≥ 99.5%) were provided by Dr. Ehrenstorfer (Augsburg, Germany). Anhydrous sodium sulfate of analytical grade was provided by Shuanglin Chemical Co. (Hangzhou, China). Analytical grade of acetone, n-hexane, and sodium chloride were provided by China National Chemical Reagent Corporation (Jiangsu, China). All other chemical reagents used in the experiment were analytical reagent grade.
2.6. Field plot trials To evaluate the feasibility of strain DDT-1 to remove or detoxify DDTs residues in soil, a bioremediation study with a completely randomized block design was performed in the vegetable field located in Cixi, Zhejiang, China. Each block covered an area of 6 m2 (2 m × 3 m) and was separated with PVC boards. A suspension of strain DDT-1 amended with 2% yeast powder was sprayed evenly with water at an inoculation level of approximately 1.0 × l08 CFUs/g using an electric sprayer. Meanwhile, the surface soils (0–10 cm) were tilled to ensure the homogenous distribution of the inocula. The control experiment without inoculation was performed under the same conditions. All plots were planted with corn using the same field management measures. Four successive inoculation treatments were performed every 30 d to ensure the survival and colonization of strain DDT-1 in the field soil. Each plot was performed in triplicate. Each soil subsample consisted of five soil cores, and six surface soil subsamples (0–10 cm, 50 g for each) were randomly collected from each plot using a stainless-steel manual auger. All subsamples from each plot were combined into a single soil sample. Soil samples were collected from all plots at 0, 15, 45, 90, 120, 150, and 210 d, respectively. Five soil subsamples from each plot were thoroughly mixed to obtain a composite sample. Subsequently, all soil samples were sieved (2 mm) to remove plant materials and debris, and then stored at − 20 °C until the determination of the DDTs residues.
2.2. Soils The fresh surface soil samples (0–10 cm) used in this study were collected from two paddy fields located at Xiaoshan (XS) and Jinhua (JH) and a mulberry field located on the Huajiachi (HJC) campus of Zhejiang University, Huangzhou, China. All soil samples were sieved (2 mm) to remove stones and debris after air-drying at room temperature. The residues of DDTs were below limit of detection (LOD) in all soil samples. The physicochemical properties of the soil samples were determined by the Zhejiang Academy of Agricultural Sciences, Hangzhou, China, and are summarized in Table S1. 2.3. Inocula preparation Stenotrophomonas sp. DDT-1 could utilize DDT as its sole carbon and energy source (Fang et al., 2016). Strain DDT-1 was repetitively cultured in the Luria-Bertoni (LB) agar (yeast extract, 5 g; peptone, 5 g; NaCl, 5 g; agar, 20 g; distilled water, 1000 ml; pH 7.2) plates, and a single colony was placed into 100 ml of sterilized LB liquid media and incubated for 24 h on a rotary shaker at 25 °C and 150 r/min in the dark. Then, the colony was transferred into a centrifuge tube and centrifuged for 10 min at 7000 r/min. The supernatant was removed and the remaining was immediately washed 3 times with 40 ml of NaH2PO4-Na2HPO4 buffer (0.1 mol/l, pH 7.0), and ultimately suspended in the same phosphate buffer as the inocula.
2.7. Field demonstration application The field demonstration areas for the strain DDT-1 preparation were divided into 12 experimental zones and designated as B1-B12, including greenhouse and open field soils, and their related information is summarized in Table S2. Each experimental zone included strain DDT-1 inoculation treatment and the corresponding un-inoculated control treatment, and each treatment was replicated at least six times. The suspension of strain DDT-1 supplemented with 2% yeast powder was evenly sprayed with water using an electric sprayer at a dose of 800 L/ ha, and the inoculation level of strain DDT-1 reached 2.0 × 106 CFUs/g dry soil in the greenhouse and open field soils. The addition of yeast powder provided a nitrogen source and energy source for the initial survival and propagation of strain DDT-1 in the inoculated greenhouse and open field soils. Four successive inoculation treatments were performed every 30 d, and mechanical tillage was carried out on the demonstration areas of B7 and B11 during the inoculation of the strain DDT-1 preparation. The un-inoculated areas receiving the same amount of distilled water were used as the controls. Each soil subsample consisted of five soil cores, and six surface soil subsamples (0–10 cm, 100 g
2.4. Biodegradation of DDTs in laboratory soils Soil samples (0.4 kg, dry weight equivalent) were divided into two equal parts. The first part was transferred into a 5-L plastic basin, and the standard solution of p,p′-DDT in acetone at a concentration of 400 mg/l was sprayed on the soil using a small sprinkling can in combination with an appropriate amount of sterile distilled water. Subsequently, the second part was also transferred into this plastic basin. All soil samples were thoroughly stirred using a plastic spoon and passed through a 2-mm sieve to ensure uniform distribution of the added p,p′-DDT at an initial concentration of 1 mg/kg dry weight soil, and were then kept for 1 h on a laminar flow bench to evaporate the solvent. Meanwhile, the strain DDT-1 preparation was inoculated into the soils at an inoculation level of 6.0 × 1010 colony-forming units 146
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2.11. DNA extraction and MiSeq sequencing
for each) were randomly collected from each plot using a stainless-steel manual auger. All subsamples from each experimental zone were combined into a soil sample. Soil samples were collected from all plots at 0, 30, 60, 90, 120, 180, 240, 270, 360, and 400 d, respectively. Subsequently, each sample was sieved (2 mm) to remove plant materials and debris after air-drying at ambient temperature and stored at − 20 °C until the determination of the DDTs residues.
The total DNA was extracted from 0.5 g of soil sample using a nucleic acid extractor and FastDNA SPIN Kit for soil (Qbiogene, Carlsbad, CA, USA) according to the manufacturer's protocol. The purity and concentration of the extracted DNA were determined using a spectrophotometer (NanoDrop ND-1000, Wilmington, DE, USA). The extracted DNA was further analyzed by 0.8% agarose gel electrophoresis (with TAE buffer containing 0.8 µg/ml of ethidium bromide) at a constant voltage of 120 V for 20 min. The 16 S rDNA V4 region was amplified using the universal forward primer 520F (5′-AYTGGGYDTAAAGNG-3′) and the reverse primer 802R (5′-TACNVGGGTATCTAATCC-3′), and a barcode unique to each sample was added to the reverse primer to allow multiplex sequencing. The PCR reaction system contained 25.0 μl, which included 1.0 μl of the 20 ng/μl template, 2.5 μl of buffer (10 ×), 2.0 μl of 2.5 mM dNTP, 1.0 μl of each primer (10 μM), 0.125 μl of Taq DNA polymerase (5 U/μl), and 17.375 μl of ultra-pure water. The PCR running conditions consisted of an initial denaturation at 94 °C for 4 min followed by 30 cycles of 94 °C for 30 s, 50 °C for 30 s, 72 °C for 30 s and a final extension at 72 °C for 5 min. Subsequently, the bacterial 16S amplicon sequencing was performed under the Illumina MiSeq platform (2 × 250 bp, San Diego, CA, USA) by the Personal Biotechnology Co. (Shanghai, China).
2.8. Extraction and determination of DDTs residues in soils The residues of DDTs in the soils were extracted following the method described by Fang et al. (2010). The concentrations of DDTs were determined by an Agilent 6890 N gas chromatography (Agilent Technologies, USA) equipped with an electron capture detector (µ-ECD) and a DB-1701 silica capillary column (30 m × 0.32 mm × 0.25 µm, Agilent Technologies, USA). The oven temperature was initially 160 °C and increased to 220 °C at a rate of 10 °C/min and then rose to 240 °C at a rate of 5 °C/min with maintenance of the final temperature for 2 min. The injector and detector temperatures were set at 230 °C and 280 °C, respectively. The injection mode was splitless and the injection volume was 1.0 μl. The flow rate of the helium carrier gas was set at 2 ml/min. 2.9. Soil enzyme activity assays
2.12. Recovery study
The catalase activity was evaluated based on the recovery rate of hydrogen peroxide. Soil samples (2.0 g dry soil weight) were mixed with 40 ml of distilled water and 5 ml of 0.3% H2O2 in a 100-ml volumetric flask, and then shaken for 30 min at 150 rpm and 30 °C in the dark. After shaking, 5 ml of H2SO4 (1.5 mol/l) was added to the flask to stabilize the un-decomposed hydrogen peroxide, and the sample was filtered through 7-cm quantitative dense filter paper. Subsequently, 25 ml of the filtrates were titrated with 0.02 mol/l KMnO4 and the results are expressed as ml KMnO4/(h·g dry soil). Meanwhile, control experiments without soil samples were conducted using the same procedure. The neutral phosphatase activity was measured referred to the method described by Rahmatpour et al. (2017). Soil samples (1.0 g, dry soil weight), 0.2 ml of toluene, 4 ml of sodium hydrogen phosphatecitric acid buffer (pH 7.0), and 1 ml of p-nitrophenyl sodium hydrogen phosphate solution (0.025 mol/l) were mixed in a 100-ml volumetric flask. The mixture was incubated for 1 h at 35 °C in the dark and the reaction was stopped by adding 1 ml of CaCl2 solution (0.5 mol/l) and 4 ml of NaOH solution (0.5 mol/l). The suspension was shaken for a few seconds and filtered, and the color intensity at 420 nm was determined using a spectrophotometer. The controls were unamended with p-nitrophenyl sodium hydrogen phosphate solution and were processed using the same procedure. The results were modified by the calibration plot of the p-nitrophenol (PNP) standard, and the neutral phosphatase activity is expressed as μg PNP/(h·g dry soil)
To evaluate the validity of the extraction method for DDTs residues, a recovery experiment was performed at spiking levels of 0.1, 1, and 10 mg/kg dry weight soil. The same extraction and detection methods for DDTs residues were used as described above. 2.13. Statistical analysis Analysis of variance (ANOVA) of the degradation half-lives and rates between the inoculated soils and un-inoculated controls was performed using SPSS 19.0 (SPSS, Chicago, IL, USA). The average values and standard deviations of the data were calculated using Microsoft Excel 2007 (Microsoft Corporation, Redmond, WA, USA). The average well color development (AWCD), Shannon index (H′), McIntosh index (U), and Simpson index (1/D) were calculated according to the methods described by Fang et al. (2014). To distinguish the differences in the diversity and abundance of the soil microbial communities after inoculation of strain DDT-1, heat maps of the top 30 soil bacterial genera were visualized using Matlab 7.0 (Mathworks, Natick, MA, USA). 3. Results and discussion 3.1. Evaluation of the recovery The recoveries of DDTs at spiking concentrations (0.1, 1.0 and 10.0 mg/kg) ranged from 91.9% to 100.7% with relative standard deviation (RSD) ≤ 4.4% in the indoor soils, and 87.1–100.7% with RSD ≤ 0.7% in the field soils. The LOD for p,p′-DDT, p,p′-DDE, p,p′-DDD, and o,p′-DDT residues in the soil was 0.005 mg/kg, 0.005 mg/kg, 0.005 mg/kg, and 0.005 mg/kg, respectively. The corresponding limit of quantitation (LOQ) was 0.001 mg/kg, 0.001 mg/kg, 0.001 mg/kg, and 0.001 mg/kg, respectively. These data indicate that the extraction method was satisfactory for the analysis of DDTs residues.
2.10. BIOLOG assay Five grams of the soil samples (dry-weight equivalent) were mixed with 45 ml of physiological saline (0.85%, w/v) in a 100-ml triangular flask and shaken for 1 h on an orbital shaker at 25 °C and 150 r/min in the dark. Subsequently, the mixtures were allowed to settle for 30 min, and a portion of the supernatant was serially diluted using sterile physiological saline in a sterile tube to obtain a 10−3 dilution. Each micropore in the BIOLOG ECO microplate™ (BIOLOG Inc., Hayward, USA) was inoculated with 150 μl of the 10−3 dilution, and all microplates were placed in a thermostat incubator for 168 h at 25 ± 1 °C. Color development in the wells was measured at 4, 24, 48, 72, 96, 120, 144, and 168 h using the BIOLOG reader (BIO-TEK, Winooski, VT, USA) at wavelengths of 750 nm and 590 nm. The output data were recorded by the automatic threshold option through BIOLOG software.
3.2. DDTs residues in the field soils The residual levels of p,p′-DDT, p,p′-DDE, p,p′-DDD, o,p′-DDT and DDTs in the field soils are shown in Figs. S1-S5. p,p′-DDT, p,p′-DDE, p,p′DDD and o,p′-DDT were detected in almost all field soils, and the residual concentration of p,p′-DDE was much higher than other three homologues in concentration, indicating that p,p′-DDT was mainly degraded to p,p′147
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DDE in the field soils under aerobic conditions (Huang et al., 2018; Zhang et al., 2018). Furthermore, the ratio of p,p′-DDT/p,p′-DDE in the field soils was less than 1, with a range of 0.01–0.67, suggesting that DDTs residues may be attributed to its substantial use in cotton fields prior to banning. As shown in Fig. S1, the residual levels of DDTs in the field soils were 0.06–1.02 mg/kg, and the mean residual concentration was 0.31 mg/kg, which showed that this investigation area has been seriously polluted by DDTs. The residues of DDTs in soils have been reported extensively. Fang et al. (2010) reported that the residual level of DDTs was 0.14 mg/kg in the pumpkin-growing field soils located in Cixi, China, and these residues of DDTs may have resulted from extensive use of DDT during the 1960s and 1970s. Zhu et al. (2005) reported that the mean level of DDTs residues was 0.14 mg/kg in the shallow subsurface soils in the outskirts of Beijing. Neitsch et al. (2016) also reported that the total concentration of p,p′-DDT, p,p′-DDE, and p,p′-DDD was 0.12 mg/kg in a conventionally farmed land located in Germany.
Fig. 2. Degradation of DDTs residues in the field plot soils after inoculation of strain DDT-1. Data followed by a different letter in the same treatment time are significantly different (p ≤ 0.05).
3.3. Biodegradation of DDT by strain DDT-1 in different soils under laboratory conditions
degradation percentages of p,p′-DDT by strain DDT-1 were 24.5%, 21.0%, and 20.9% in HJC, XS, and JH soils, respectively, and the corresponding percentages were only 9.5%, 8.9% and 8.4% in the un-inoculated control samples, which showed the much higher degradation percentages of p,p′-DDT in inoculated soils compared with those in the un-inoculated control soils. As shown in Table S3, the degradation rates of p,p′-DDT in strain DDT-1 inoculated HJC, XS, and JH soils were 2.7-, 2.0-, and 3.0-fold higher, respectively, than those in the un-inoculated controls. The degradation half-lives of p,p′-DDT in strain DDT-1 inoculated HJC, XS, and JH soils were 2.4-, 1.5-, and 3.8-fold lower than those in the un-inoculated controls. The significantly higher degradation rate in the inoculation treatment than in the un-inoculated controls indicated that strain DDT-1 could efficiently degrade p,p′-DDT residues in soils, and furthermore, this bacterium could be widely adapted to different types of soil. Similarly, Gao et al. (2015) reported that approximately 60.1%, 61.6%, 63.5%, and 65.0% of the initial p,p′-DDT was degraded by the plasmid donor bacteria E. coli TGI(pDOD-gfp) in JH, JX, XS and HZ soils, respectively. Purnomo et al. (2011) demonstrated the ability of brown-rot fungi G. trabeum to degrade DDT in historically contaminated soils under laboratory conditions. Additionally, the biodegradation of p,p′DDT in the HJC, XS, and JH soils was correlated to the soil physicochemical properties. As shown in Table S1 and Fig. 1, a significant positive correlation (R = 0.997) between the biodegradation rate of p,p′-DDT and soil organic matter content was found in all soils. Han et al. (2016) also found a similar positive correlation between the degradation half-life of p,p′-DDT and soil organic matter content as well as soil clay mineral content.
The degradation of p,p′-DDT by strain DDT-1 in different soils under laboratory conditions is shown in Fig. 1. After inoculation for 28 d, the
3.4. Biodegradation of DDTs residues by strain DDT-1 in field plot soils The biodegradation of DDTs residues in the field plot soils after inoculation of strain DDT-1 is shown in Fig. 2. After inoculation for 210 d, the degradation percentage of DDTs was 38.0%, but the corresponding value was only 11.1% in the un-inoculated controls. As shown in Fig. 2 and Fig. S6, the degradation percentage of DDTs in the inoculated soils was 1.5–3.6-fold higher than that in the un-inoculated control soils. As shown in Table S4, the degradation half-lives of p,p′DDT, p,p′-DDE, p,p′-DDD, o,p′-DDT, and DDTs in the inoculated soils were 47.4%, 18.3%, 33.3%, 68.0%, and 34.0%, respectively, of those in the corresponding un-inoculated soils. The corresponding degradation rates in the inoculated soils were 1.5-, 3.9-, 2.3-, 1.3-, and 2.5-fold higher than those in the un-inoculated soils, which indicated more rapid degradation in the inoculated soils than in the un-inoculated soils. The incorporation of strain DDT-1 into field plot soils could efficiently remove DDTs residues. Several similar studies on the
Fig. 1. Degradation of p,p′-DDT in different types of soils inoculated with strain DDT-1 under laboratory conditions. Data followed by a different letter in the same treatment time from each subfigure are significantly different (p ≤ 0.05). 148
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biodegradation of DDTs in field soils have been conducted. Qu et al. (2015) reported that 80.4% of DDT was removed by strain Chryseobacterium sp. PYR2 in a pilot-scale field soils after inoculation for 45 d. Fang et al. (2010) reported that 56.05% of DDTs were degraded by strain Sphingobacterium sp. D6 in the pumpkin-grown field soils. Wang et al. (2010) also reported that the inoculation of the DDT-degrading strain Pseudoxanthomonas sp. Wax into fine sandy-loam soils at an inoculation level of 108 CFUs/g could thoroughly eliminate 20 mg/kg of DDTs after treatment for 20 d. Some researchers reported that the survival and propagation of the inocula determined the success of DDTs bioremediation. However, the biodegradation activity of the inocula after inoculation in field soils depended largely on some factors, including competition and antagonism by indigenous microorganisms, predation by protozoa, soil physicochemical properties (moisture, pH, aeration, clay content, organic matter content, salinity etc.), environmental conditions (temperature, humidity, rainfall, solar irradiation etc.), and electron acceptors as well as the bioavailability of DDTs (Baczynski et al., 2010; Fang et al., 2010; Purnomo et al., 2011). Additionally, the similar high biodegradation ability of strain DDT-1 for DDTs in the field plot soils and laboratory soils indicated that strain DDT-1 could be applied to the field for the removal of DDTs residues.
Fig. 3 and Figs. S7 and S8. The residual concentration of DDTs did not decrease significantly in any of the un-inoculated controls. After inoculation for 420 d, the mean degradation percentages of p,p′-DDT, p,p′-DDE, and DDTs were 84.9%, 36.7%, and 42.1% in the open field soils, and 76.1%, 38.0%, and 38.7% in the greenhouse soils, respectively. The mean degradation rates of p,p′-DDT, p,p′-DDE, and DDTs in the inoculated greenhouse soils were 1.08-, 1.04-, and 1.08-fold higher than those in the inoculated open field soils. As shown in Tables S5-S7, the degradation half-lives of p,p′-DDT, p,p′-DDE, and DDTs were 15.3–77.0%, 12.6–27.7%, and 10.9–32.5% (open field soils) and 30.8–39.4%, 16.7–28.8%, and 20.6–30.8% (greenhouse soils) shorter in the inoculation treatments than those in the un-inoculated controls. The degradation rates of p,p′-DDT, p,p′-DDE, and DDTs were 1.1–2.4, 2.7–6.3, and 2.8–7.6 (open field soils) and 1.5–1.8, 3.2–5.2, and 2.9–5.5 (greenhouse soils) fold higher in the inoculation treatments than those in the un-inoculated controls. The application of strain DDT-1 preparation could accelerate the degradation of DDTs residues in the greenhouse and open field soils. A similar result has been reported that soil inoculation with strain DDT-1 could decrease the DDTs concentration from 0.489 mg/kg to 0.265 mg/ kg within 18 months in a DDTs-contaminated agricultural site (Zhu et al., 2012). Additionally, the degradation rate of DDTs in the inoculated greenhouse soils was higher than that in inoculated open field soils, which may be attributed to the relatively closed environment of the greenhouse.
3.5. Field demonstration of DDTs biodegradation by strain DDT-1 The biodegradation of p,p′-DDT, p,p′-DDE, and DDTs residues in the strain DDT-1 inoculated soils under field conditions is presented in
Fig. 3. Degradation of DDTs residues in the greenhouse and open field soils after inoculation of strain DDT-1 preparation. B1: open field planted soil; B2: greenhouse/open field planted soil; B3: open field unplanted soil; B4: open field unplanted soil; B5: open field planted soil; B6: greenhouse planted soil without mechanical tillage; B7: greenhouse planted soil with mechanical tillage; B8: greenhouse planted soil; B9: open field planted soil without mechanical tillage; B10: open field planted soil; B11: open field planted soil with mechanical tillage; and B12: open field planted soil. The two curves followed by a different letter in each subfigure are significantly different (p ≤ 0.05).
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Fig. 4. Variations in soil catalase activity (a) and neutral phosphatase activity (b) after inoculation of strain DDT-1 into greenhouse and open field soils. The two curves followed by the same letter in each subfigure are not significantly different (p ≤ 0.05).
3.7. Variation of soil microbial functional diversity after inoculation of strain DDT-1
3.6. Variation of soil enzyme activities after inoculation of strain DDT-1 Soil enzyme activity is related to soil organic matter decomposition and nutrient cycling, reflecting the intensity of soil microbial processes to a certain extent (Trasar-Cepeda et al., 2000; Jin et al., 2009). Changes in enzyme activities in the greenhouse and open field soils after inoculation of strain DDT-1 preparation are shown in Fig. 4. No significant differences in the catalase activity were found between the inoculated field soils and the corresponding un-inoculated field soils throughout the entire bioremediation period (Fig. 4a). A similar pattern in the neutral phosphatase activity was also observed, even though there was a slight reduction at 200 d in the open field planted soils (B5) and a slight increase at 300 d in the open field untilled soils (B9) in inoculation treatments compared to that of the corresponding un-inoculated controls (Fig. 4b). In this study, therefore, the inoculation of strain DDT-1 into the field soils did not significantly affect soil enzyme activities, including catalase and neutral phosphatase activities. Similar to our results, no significant differences in the soil catalase activity were found between strain DXZ9 inoculated-soils and the un-inoculated control soils after treatment for 210 d (Xie, 2013). Huang (2012) also reported that the inoculation of strain DDT-1 into aged DDT-contaminated soils only had a slight stimulation effect on the soil phosphatase activity.
The average well color development (AWCD), which reflects the ability of soil microbial communities to utilize different carbon sources, is commonly used to assess the overall activity of soil microbial communities (Wu et al., 2014). As shown in Fig. 5a, 0 d after inoculation of strain DDT-1, the AWCD values in all inoculated greenhouse and open field soils were higher than those in the corresponding un-inoculated greenhouse and open field soils, suggesting that the overall activities of the soil microbial communities were stimulated in the early stage of inoculation. After inoculation for 210 d and 420 d, no statistically significant difference in the AWCD values was observed between the inoculated greenhouse and open field soils and the corresponding uninoculated control soils (Fig. 5b-c), indicating that the overall soil microbial activities recovered to the control level in the middle and later stages of inoculation. Therefore, the effect of strain DDT-1 inoculation on the overall soil microbial activities gradually disappeared over the inoculation time. Soil microbial diversity indices, including the Simpson index (1/D), Shannon index (H′), and McIntosh index (U), are usually used to evaluate the dominant population, richness and evenness of soil microorganisms, respectively (Fang et al., 2015). Soil microbial diversity indices after inoculation of the strain DDT-1 preparation are summarized in Table 1. Immediately after inoculation of strain DDT-1, the 1/D 150
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Fig. 5. Variations in average well color development (AWCD) at 0 d (a), 180 d (b), and 420 d (c) in the greenhouse and open field soils after inoculation of the strain DDT-1 preparation. The two curves followed by the same letter in each subfigure are not significantly different (p ≤ 0.05).
values in all inoculated greenhouse and open field soils were slightly higher than those in the corresponding un-inoculated control soils, indicating that the dominant soil microbial populations increased in the inoculated greenhouse and open field soils. Subsequently, the 1/D values in all inoculated soils gradually decreased at 180 d after inoculation and then recovered to a level similar to that of the control soils
420 d after inoculation, suggesting that the dominant soil microbial populations recovered to the control level. However, the H′ and U values in all inoculated greenhouse and open field soils were not statistically different from those in the corresponding un-inoculated control soils, indicating that soil microbial evenness and richness were not significantly affected in the middle and later stages of inoculation.
Table 1 Variations in the soil microbial functional diversity indices after inoculation of strain DDT-1 into the greenhouse and open field soils. Time (d)
Treatment
Planted pattern
Shannon index (H′)
Simpson index (1/D)
McIntosh index (U)
0 0 0 0 0 0 0 0 0 0 0 0 180 180 180 180 180 180 180 180 180 180 180 180 420 420 420 420 420 420 420 420 420 420 420 420
B3-CK B3-BR B5-CK B5-BR B6-CK B6-BR B7-CK B7-BR B9-CK B9-BR B11-CK B11-BR B3-CK B3-BR B5-CK B5-BR B6-CK B6-BR B7-MT-CK B7-MT-BR B9-CK B9-BR B11-CK B11-BR B3-CK B3-BR B5-CK B5-BR B6-CK B6-BR B7-CK B7-BR B9-CK B9-BR B11-MT-CK B11-MT-BR
Open field-unplanted Open field-unplanted Open field-planted Open field-planted Greenhouse-unplanted Greenhouse-unplanted Greenhouse-planted Greenhouse-planted Open field-unplanted Open field-unplanted Open field-planted Open field-planted Open field-unplanted Open field-unplanted Open field-planted Open field-planted Greenhouse-untilled Greenhouse-untilled Greenhouse-tilled Greenhouse-tilled Open field-untilled Open field-untilled Open field-tilled Open field-tilled Open field-unplanted Open field-unplanted Open field-planted Open field-planted Greenhouse-untilled Greenhouse-untilled Greenhouse-tilled Greenhouse-tilled Open field-untilled Open field-untilled Open field-tilled Open field-tilled
2.38 2.51 2.58 2.42 2.14 2.08 2.76 2.30 1.92 2.26 2.51 2.43 1.79 2.36 1.52 1.68 1.87 2.00 1.64 1.92 2.24 2.65 2.01 2.59 3.06 3.01 2.77 3.13 3.12 3.02 2.78 2.96 3.16 3.14 2.89 3.04
7.73 ± 1.23 a 11.29 ± 0.76 b 9.21 ± 0.55 a 10.09 ± 0.49 b 4.36 ± 1.47 a 7.28 ± 1.57 b 12.28 ± 1.11 a 12.93 ± 0.26 a 6.58 ± 2.27 a 7.11 ± 1.58 b 10.13 ± 2.67 a 11.78 ± 2.57 a 4.04 ± 2.55 a 9.07 ± 0.50 b 4.70 ± 2.84 a 5.32 ± 3.58 a 6.01 ± 1.51 a 6.50 ± 0.37 a 4.35 ± 1.81 a 5.00 ± 1.53 a 8.12 ± 3.88 a 9.49 ± 2.96 a 6.32 ± 0.83 a 10.44 ± 0.76 b 18.84 ± 2.46 a 21.12 ± 2.32 a 16.85 ± 2.46 a 18.67 ± 2.62 a 16.80 ± 0.78 a 19.44 ± 1.50 a 17.43 ± 2.17 a 18.91 ± 0.83 a 18.97 ± 0.98 a 20.00 ± 1.84 a 18.27 ± 2.17 a 19.87 ± 1.06 a
1.32 2.06 2.12 1.89 0.90 1.81 2.30 1.61 1.78 1.26 1.94 1.87 2.23 1.09 1.09 1.14 1.06 1.38 1.12 0.92 1.51 1.32 1.26 1.75 4.40 4.33 3.09 3.71 3.96 3.28 3.78 3.29 4.32 4.11 3.97 4.00
± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ±
0.35 0.20 0.10 0.04 0.36 0.34 0.10 0.04 0.69 0.15 0.29 0.14 0.29 0.37 0.78 0.78 0.32 0.43 0.21 0.29 1.28 0.28 0.16 0.07 0.24 0.12 0.45 0.25 0.06 0.17 0.39 0.16 0.08 0.15 0.29 0.14
a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a
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± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ±
0.35 0.31 0.13 0.19 0.58 0.66 0.39 0.23 1.02 0.25 0.23 0.59 1.17 0.94 0.59 0.63 0.17 0.47 0.59 0.26 0.89 0.42 0.31 0.18 0.13 0.34 0.76 0.33 0.13 0.08 0.63 0.35 0.25 0.31 0.55 0.36
a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a a
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Fig. 6. Heat maps of the top 30 soil microbial genera at 0 d (a), 180 d (b), and 42 d (c) in the greenhouse and open field soils after inoculation of the strain DDT-1 preparation. 1–12 represent B3-CK (open field unplanted soil-control), B3-BR (open field unplanted soil-inoculation), B5-CK (open field planted soil-control), B5-BR (open field planted soil-inoculation), B6-CK (greenhouse untilled soil-control), B6-BR (greenhouse untilled soil-inoculation), B7-MT-CK (greenhouse soil-mechanical tillage-control), B7-MT-BR (greenhouse soil-mechanical tillage-inoculation), B9-CK (open field untilled soil-control), B9-BR (open field untilled soil-inoculation), B11-MT-CK (open field soil-mechanical tillage-control), and B11-MT-BR (open field soil-mechanical tillage-inoculation), respectively.
later stages of strain inoculation. Therefore, strain DDT-1 might be efficient and ecologically safe for in situ bioremediation of the DDTcontaminated greenhouse and open field soils. Further studies need to be carried out on the effects of seasonal and geographical variation on the colonization and adaptation abilities of strain DDT-1 in the field soils.
3.8. Variation in soil microbial structural diversity after inoculation of strain DDT-1 The heat maps of the top 30 soil microbial genera at 0 d, 180 d and 420 d after inoculation of strain DDT-1 are shown in Fig. 6. After inoculation of strain DDT-1, the dominant soil bacterial genera were Bacillus, Sphingobacterium, Stenotrophomonas, and Nocardioides at 0 d (Fig. 6a); Bacillus, Stenotrophomonas, Sphingobacterium, Nocardioides, and Nitrospira at 180 d (Fig. 6b); and Nitrospira, Nocardioides, Bacillus, Stenotrophomonas, and Sphingobacterium at 420 d (Fig. 6c). Throughout the entire bioremediation period, the soil microbial structural diversity was generally similar between the inoculated greenhouse and open field soils and the corresponding un-inoculated control soils, but several soil microbial genera had some fluctuations in their relative abundances. For instance, an initial increase followed by a decline in the abundance of Stenotrophomonas was observed with the extension of the inoculation time, and that of Nitrospira increased in the later stage of inoculation. The principal component analysis (PCA) of soil microbial communities after inoculation of strain DDT-1 is shown in Fig. S9. The PC1 component contributed more than the PC2 component in the PCA profile throughout the entire bioremediation period. As shown in Fig. S9, a distinctly dispersed pattern was presented at 0 d after strain inoculation, and then a cluster was gradually formed at 180 d after strain inoculation. Finally, a scattered pattern was observed at 420 d after strain inoculation. The results showed that the structural diversity of soil microbial communities was temporarily suppressed in the initial stage of strain inoculation but gradually recovered to the similar level in the un-inoculated controls over the inoculation time.
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