Bioremediation potential of soil contaminated with highly substituted polychlorinated dibenzo-p-dioxins and dibenzofurans: Microcosm study and microbial community analysis

Journal of Hazardous Materials 261 (2013) 351–361

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Bioremediation potential of soil contaminated with highly substituted polychlorinated dibenzo-p-dioxins and dibenzofurans: Microcosm study and microbial community analysis Wei-Yu Chen, Jer-Horng Wu ∗ , Yong-Yu Lin, Hung-Jun Huang, Juu-En Chang Department of Environmental Engineering, National Cheng Kung University, Taiwan, ROC

h i g h l i g h t s • • • •

We demonstrate that near-fully and fully chlorinated dioxins could be biodegraded. We uncover the microbial composition in the dioxin-degrading microcosm. Microbial populations are subjected to great dynamics when dioxins are degraded. We obtain four bacterial cultures that can degrade octachlorodibenzofuran.

a r t i c l e

i n f o

Article history: Received 5 April 2013 Received in revised form 10 July 2013 Accepted 16 July 2013 Available online 24 July 2013 Keywords: Polychlorinated dioxins Octachlorodibenzofuran Bioremediation Microbial community structure

a b s t r a c t Highly chlorinated dibenzo-p-dioxins/dibenzofurans (DD/Fs) are main hazardous dioxins, and ubiquitously distributed in the environment. To study the feasibility of bioremediation for remedying contamination of highly chlorinated dioxins, closed microcosms were constructed with soil from a chronological site under oxygen-stimulated conditions. The results showed that high levels of near-fully and fully chlorinated DD/Fs, particularly octachlorodibenzofuran were effectually reduced without accumulation of less substituted congeners. The clone library analysis of PCR-amplified 16S rRNA gene from the octachlorodibenzofuran-degrading consortia showed that 98.3% of the detected sequences were affiliated with Proteobacteria. The obtained strains with putative aromatic dioxygenase genes and abilities to repetitively grow in octachlorodibenzofuran-containing agars were closely related to members within Actinobacteria, Firmicutes, and Proteobacteria. Among them, certain Rhodococcus, Micrococcus, Mesorhizobium and Bacillus isolates could degrade octachlorodibenzofuran with efficiencies of 26–43% within 21 days. Hierarchical oligonucleotide primer extension analysis further showed that Micrococcus, Rhizobium, Pseudoxanthomonas, and Brevudimonas populations increased largely when high concentrations of octachlorodibenzofuran were reduced. Overall, our results suggest that a distinctive microbial composition and population dynamic could be required for the enhanced degradation of highly chlorinated DD/Fs in the batch microcosm and highlight a potential of bioremediation technologies in remedying polychlorinated dioxins in the polluted sites. © 2013 Elsevier B.V. All rights reserved.

1. Introduction Polychlorinated dibenzo-p-dioxins/dibenzofurans (PCDD/Fs), which encompass a group of 210 congeners, are among the hazardous pollutants of most concern. These compounds, including 17 laterally substituted (2,3,7,8) congeners that are legislatively regulated, display potent endocrine-disrupting activity and are associated with numerous health disorders and carcinogenicity

∗ Corresponding author at: Department of Environmental Engineering, National Cheng Kung University, No. 1, University Road, Tainan City 70101, Taiwan, ROC. Tel.: +886 910385159; fax: +886 6 2752790. E-mail address: [email protected] (J.-H. Wu). 0304-3894/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.jhazmat.2013.07.039

[1]. PCDD/Fs could be formed through various natural and anthropogenic combustion processes, rendering them ubiquitous in the environment [2]. Because of their hydrophobicity, recalcitrance, and strong partition to particles, the concentrations, although usually low in the soil (∼ppb level), build up in soil, sediment, and biota. High concentrations of PCDD/Fs could be inadvertently generated with the manufacture of commercial chlorinated chemicals, such as pentachlorophenol (PCP) and chlorophenoxy pesticides [3]. This could lead to heavy pollution in an area, resulting from mass usage of chemical formulations, improper disposal, and storage and waste, or during accidents [4,5]. Therefore, remediation measures for such contaminated sites are required. The microbiological approach represents an environmental friendly and cost-effective remediation technology for

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PCDD/Fs-contaminated soil [6]. Several studies have demonstrated that chlorinated DD/Fs can be degraded by defined bacterial isolates or microbial consortia [7–11]. Under aerobic conditions, the bacterial catabolism of DD/F and lightly chlorinated counterparts is initiated by various aromatic ring-hydroxylating dioxygenases involved in the upper pathway, from which the catechol related monoaromatic metabolites can be yielded, and then transformed by the catechol dioxygenase in the lower pathway [12,13]. The degradation of highly chlorinated congeners is restricted to a few specialist strains, such as Sphingomonas wittichii RW1 and Pseudomonas veronii PH-03 [14]; however, chlorinated dioxins with 4–6 chlorine substitutions could only be co-metabolized under aerobic conditions or undergo the reductive dechlorination under anaerobic conditions [8]. The underlying knowledge was based on studies that focused mainly on low-chlorinated or the congeners that did not include the 17 toxic counterparts. The field investigations indicated that the highly chlorinated dioxin congeners, specifically hepta- and octa-chloro ones accounted for the majority of toxic dioxins in soil environments regarding total toxicity equivalency quantity (TEQ, the sum of the products of individual concentrations multiplied by their respective toxicity equivalency factors) and molar concentrations [15–18]. Despite a lower toxicity (toxicity equivalency factor is 103 –104 times lower than 2,3,7,8-tetrachlorodibenzo-p-dioxin (2,3,7,8-TCDD)) [19], the biotransformation of highly chlorinated dioxins (>6 chlorines) may lead to more toxic dechlorinated congeners [20]. Recently, simultaneous dechlorination and oxidative degradation of PCDD/Fs could be observed in the less anoxic microcosms [21,22]. However, the relevant microbial community structures were not sufficiently addressed. The An-Shun site located close to the seacoast in the northwest of Tainan, Taiwan, was heavily contaminated with PCDD/Fs due to large production of PCP from 1965 to 1982 [5]. A site investigation uncovered that TEQ ranged from several to hundreds of milligram TEQ per kilogram of soil (mg TEQ/kg) in the hot-spot area, and the dominant forms of congeners were highly chlorinated, including heptachlorodibenzo-p-dioxin (Hp CDDs), heptachlorodibenzofuran (Hp CDFs), octachlorodibenzop-dioxin (OCDD), and octachlorodibenzofuran (OCDF) [23]. These levels of PCDD/Fs are at least 1000 times higher than the regulation standard (1 ␮g TEQ/kg). Because the degradation of fully or nearly fully chlorinated DD/Fs is poorly understood, the application of bioremediation technology to sites such as An-Shun remains challenging. The main purpose of this study was to explore microbial potential for the bioremediation of PCDD/Fs at the An-shun site. This study demonstrates a rapid and effectual degradation of Hp CDD/Fs and OCDD/F in soil microcosms. Culture-dependent and culture-independent biological tools were used to characterize the community structure of microbial consortia that were capable of reducing high concentrations of OCDF. The bacterial populations over the timeframe of degradation were studied by using a quantitative hierarchical oligonucleotide primer extension method [24].

2. Experimental 2.1. Soil samples In this study, soil samples (10–30 cm depth) were collected from the An-Shun site (120◦ 07.467 E, 23◦ 02.173 N) northwest of Tainan, Taiwan, and stored at the ambient temperature. The site was close to the seacoast and was contaminated with high concentrations of PCDD/Fs for decades [2]. The moist soils sieved with a 20-mesh stainless steel sieve (particle size <0.85 mm) were used in the experiments and were characterized as a sandy loam

texture with a pH of 8.0, water content of 9.9%, total organic carbon of 1.17%, total nitrogen of 0.10%, phosphorus of 2.23 mg/kg dry soil, and chloride of 0.19–1.42 g/kg dry soil. 2.2. Batch microcosm assays To inspire the activity of soil microorganisms, the first-batch soil microcosms were prepared by resuspending 10 g (wet weight) of soil with 20 ml of a minimal medium [25] containing (per liter) 2.2 g of Na2 HPO4 , 0.8 g of NH2 PO4 , 1.0 g of NH4 Cl, 0.2 g of MgSO4 ·7H2 O, 10 mg of FeSO4 ·7H2 O, 10 mg of CaCl2 ·2H2 O, and 20 mg of yeast extract (pH 7.2) in 125 ml serum bottles in duplicates. The headspaces of the serum bottles were flushed with a pure oxygen stream for 3 min, capped with Teflon-coated rubber septa, and subsequently sealed with aluminum caps to maintain oxygenrich conditions (approximately 1.5 atm of oxygen) at the onset of the incubation. A total of 12 bottles were prepared and 6 (control bottles) were heat-sterilized twice at 1.21 atm and 121 ◦ C for 30 min to inhibit microbial activity. The bottles were incubated at 28 ◦ C in the dark in an orbital shaker that provided excellent mixing action (300 rpm). During the tested period, every two bottles with and without sterilization were opened at time 0, 6, and 12 weeks for the analysis of dioxin congeners, respectively. Because the bottles were tightly closed, the water content of samples remained relatively constant (about 70%) during incubation. After about 3 months (the bottles capped were kept static at 28 ◦ C in the dark), microbial consortia (50%, v/v) were transferred using sterilized glass pipettes to serum bottles containing a fresh minimal medium and 50 mg of OCDF powder (purity >98%, purchased from AccuStandard Inc., New Haven, USA) for further enrichment. In the second-batch microcosm, the decapped bottles were replenished with oxygen and resealed with Teflon-coated rubber septa and aluminum caps, after the enrichment cultures of 2 ml were sampled using glass pipettes on a weekly basis. The slurry samples for microbial analysis were stored at −20 ◦ C, whereas those for the PCDD/Fs measurement, whole slurry samples were air-dried, homogenized and preserved in the brown glass vials at 4 ◦ C before analysis. 2.3. PCDD/Fs analysis The concentrations of 17 toxic PCDD/Fs congeners (2,3,7,8substituted) were analyzed using the isotope dilution high resolution gas chromatography–high resolution mass spectrometry (HRGC-HRMS) method described in [10,26]. Prior to analysis, the samples were subjected to the treatment procedures including internal standardization, Soxhlet extraction, Synder column concentration, and nitrogen gas vaporization. For the batch microcosm assay, soil samples of approximately 0.1–1 g (dry weight) were extracted for 24 h into a Soxhlet apparatus with HPLC-grade toluene. For the OCDF degradation by the isolated strains (see section 2.6), the whole culture samples were extracted three times (30 min each time) with hexane/acetone (1:1, v/v, HPLC grade), and concentrated on a rotary evaporator. Each extract was re-dissolved in hexane and cleaned on a column with layers of silica impregnated by sulfuric acid. The sample was subsequently split on a Carbopack C/Celite column, and the collected PCDD/Fs were dissolved in toluene. Seventeen stable isotope 13 C12 - or 37 Cl4 -labeled PCDD/Fs (Cambridge Isotope Laboratories and the Wellington Company) were spiked as the internal standards to ensure recovery efficiencies in the pretreatments. Quantification of PCDD/Fs was performed on a gas chromatograph (GC: Agilent 6890 series) equipped with a capillary column, CP-5MS (60 m × 0.250 mm × 0.25 ␮m) and a high-resolution mass spectrometer (HRMS: JMS 700D, JEOL, Japan). The temperature was programmed as follows: the injector was set at 280 ◦ C, the initial oven temperature was 170 ◦ C (maintained for 5 min), heated to 220 ◦ C at 20 ◦ C/min (maintained for 5 min), heated

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to 240 ◦ C at 1 ◦ C/min (maintained for 10 min), and heated to 320 ◦ C at 5 ◦ C/min (maintained for 2 min). The HRMS was operated using electron ionization with selected ion-monitoring (EI-SIM) mode and a resolving power of >10,000. The total TEQs of the dioxin congeners in a sample were calculated based on international toxic equivalency factors (I-TEFs) [27]. 2.4. Soil DNA preparation and microbial composition analysis Total DNAs of weekly samples from the OCDF enriching secondbatch microcosm was recovered using the modified bead-beating method [28]. Briefly, soil slurry of approximately 1 g (wet weight) was centrifuged at 6000 g to remove the supernatant. After washed 3 times with a phosphate buffer (100 mM NaH2 PO4 , pH 8.0), the precipitate was subsequently resuspended in 600 ␮l of a lysis buffer (0.1 mM EDTA, 5% (w/v) SDS, 10 mM Tris–HCl, pH 8.0). The soil DNA was recovered after thermal incubation at 70 ◦ C by using glass bead beating, phenol/chloroform/isoamyl alcohol extraction, and ethanol precipitation. The crude DNA extract was further purified using the DNA Clean-Up System (Promega, USA). Prior to analysis of microbial community structure, the 16S rRNA gene fragments were amplified using PCR with the Bacteria-specific primers 27F/1492R (Table 1), and cloned using a TOPO TA cloning® kit (Invitrogen Inc., CA, US) in accordance with our previous study [29]. The DNA inserts were confirmed using direct PCR with the M13 primer set (Table 1), and screened using the restriction fragment length polymorphism (RFLP) method with 2 tetramer restriction enzymes, AluI and MspI (New England Biolabs Inc., MA). The clones shared a RFLP profile were grouped into one operational taxonomic unit (OTU), and the sequences of each OTU representative were obtained by sequencing with primers Sp6 and T7 (Table 1) on an ABI PRISM® 3130 genetic analyzer and a BigDye Terminator sequencing kit (Applied Biosystem Inc., SFO, US). The nucleotide sequences of the detected OTUs were assessed for chimeric artifacts using Pintail software [30], and then compared with the sequences of relatives in the GenBank using a BLAST search. The phylogenetic tree and bootstrap resampling analysis for 1000 replicates were constructed by applying the neighbor-joining method provided in the MEGA5 program [31] after the sequences were aligned using BioEdit software [32]. 2.5. Hierarchical oligonucleotide primer extension analysis The microbial populations of interest were quantitatively analyzed using the hierarchical oligonucleotide primer extension analysis (HOPE) technique [24]. The HOPE reaction mixture was prepared in a solution (5 ␮l) containing 1.5 ␮mol of each unlabeled primer (Mission Biotech, Taiwan), 10 fmol of the 16S rRNA gene amplicons that were purified by using a QIAquick PCR cleanup kit (Qiagen, Chatsworth, CA, USA), and 2 ␮l of the ready-to-use premix from the SNaPshot® multiplex kit (Applied Biosystems). The singlebase primer extension was performed in a Thermocycler (Biometra, Germany) with 20 cycles of 95 ◦ C for 5 s, 62 ◦ C for 10 s, and 72 ◦ C for 10 s. The extended primers were fluorescently labeled because the SNaPshot premix contained AmpliTaq DNA polymerase, salt, and buffer for extending single fluorescently labeled dye-terminators at the 3 end of the primers. The resulting products were treated with shrimp alkaline phosphatase (USB Affymetrix, US) to minimize the influence of unincorporated dye-terminators, and analyzed using an ABI PRISM® 3130 genetic analyzer. The operation of capillary electrophoresis, collection of fluorescence data, and determination of relative abundances were processed as described elsewhere [33]. The 16S rRNA-targeted oligonucleotide primers (Table 1) used in the HOPE analysis were designed using the ARB package [34] with a sequence database (SSU Ref version 91 from http://www.arb-silva.de/). The primers were newly designed at

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each strand with lengths from 16 to 20 nucleotides (nt). To confer maximal mismatch discrimination efficiencies of primers, the effects of number and position of mismatch bases corresponding to the non-targeted sequences were considered in accordance with the primer design criteria [35]. The specificity of the primers was evaluated in silico against the sequences in the Ribosomal Database Project II (http://rdp.cme.msu.edu/probematch/search.jsp) using the Probe Match tool, and experimentally validated with perfectmatch and mismatch reference DNA. To effectively differentiate the extended primers in the multiplexing detections, primers 338F, 668R mod1, and 585F 8tail were modified with varied lengths of nonsense-sequence (ATGC)n n = 2–8 tails at the 5 end of the primer. 2.6. Bacterial isolation, identification and OCDF-degrading ability Pure bacterial isolates were obtained with OCDF as a cometabolic substrate using the agar plating method. The agar plates were prepared with the minimal [25] and the marine media (per liter: 5 g of peptone, 1 g of yeast extract, 0.1 g of ferric citrate, 19.45 g of NaCl, 5.9 g of MgCl2 , 3.24 g of MgSO4 , 1.8 g of CaCl2 , 0.55 g of KCl, 0.16 g of NaHCO3 , 0.08 g of KBr, 0.034 g of SrCl2 , 0.022 g of HBO3 , 0.004 g of Na2 SiO3 , 0.0024 g of NaF, 0.0016 g of NH4 NO3 , and 0.008 g of Na2 HPO4 ) (Difco, BD Company, USA) with an aim to isolate the bacteria from the oligotrophic and salty environments, respectively. Then, 1 ml of the n-hexane solution containing 0.1 mg OCDF was spread on the surface of agar plates using a glass triangular-end cell spreader. After n-hexane was completely evaporated, approximately 0.1 ml of the diluted suspension (the inoculum was taken after the first-batch microcosm experiment (see experimental Section 2.2), and 10–1000 times diluted with sterilized corresponding medium broth) was spread on the OCDFoverlaying agar plates. The colonies were randomly retrieved after 7 and 14 days of incubation at 28 ◦ C and subsequently streaked on the fresh corresponding OCDF-overlaying agar plates until single homogenous colonies were obtained. The strains were repeatedly subcultured on OCDF-containing minimal medium with a 10times-diluted Luria–Bertani medium (LB-minimal medium) and marine agar plates to confirm their growth ability. For the purity analysis, the bacterial isolates cultivated in the LB-minimal or marine broth media were harvested at the exponential phase and washed 3 times with a phosphate buffer (pH 7.4). Then, the genomic DNA was recovered using a freeze/thaw method [36], and PCR amplification of 16S rRNA gene fragments, cloning, and sequencing were performed as mentioned in the details previously. Bacterial strains with a sequence similarity of >97% were assigned to the same species, as compared to the sequences in the NCBI using BLAST searching. To investigate the OCDF-degrading ability of bacterial isolates, the cells grew in the exponential phase were harvested by centrifugation (6200 g at 4 ◦ C for 5 min), washed twice with phosphate buffer (pH7.4) and resuspended in the corresponding LB-minimal or marine media by adjusting the optical density to 4.0 at 600 nm. The concentrated cell suspension of 1 g (wet weight) was injected into the Teflon-septum-capped brown vials (total volume 40 ml) containing 10 ␮g OCDF in 9 g filter-sterilized LB-minimal or marine media (wet weight), and pure oxygen using the sterile syringes. The vials including those without cells were prepared in duplicate and incubated in the dark (200 rpm, 28 ◦ C) for 21 days, and subjected to the analysis of 17 PCDD/Fs congeners as described previously. 2.7. PCR detection of aromatic dioxygenase-coding genes PCR detection with primers specific to genes coding for aromatic dioxygenases [37] was conducted in a Thermocycler (Biometra, Germany) at the assigned annealing temperatures and

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Table 1 Oligonucleotide primers used in this study. Primer name

Target gene

Specificity

Sequence (5 –3 )a

ddNTP/ Amplicon size (bp)

Annealing temp (◦ C)

Used for

References

27F 1492R 995F 668R mod1

16S 16S 16S 16S

AGAGTTTGATCMTGGCTCAG TACGGYTACCTTGTTACGACTT ATGTTCTCGATCGCCGTA (atgc)2 TACACCGGAAATTCCGCATC

1465

54

G C

62 62

PCR-cloning PCR-cloning HOPE (Set 1) HOPE (Set 1)

[67] [67] This study This study

558F 8tail 822F

16S 16S

(atcg)8 TGGTTCGTTAAGTCTGCC AAGCTAGCCGTTGGCAGA

G T

62 62

HOPE (Set 2) HOPE (Set 2)

This study This study

138F 1000R 1004F 1017R 183R 576R

16S 16S 16S 16S 16S 16S

GTGGTACGGAACAACACG GATCTCTCACGCGGTCCG CGGTTAGGAGAGATCCTT GCCGAACTGAAGGAAAGGA CCTCATGAGAGGTTATGC GACAAACCGCCTACGAAC

G G T T A T

62 62 62 62 62 62

HOPE (Set 2) HOPE (Set 3) HOPE (Set 3) HOPE (Set 3) HOPE (Set 4) HOPE (Set 4)

This study This study This study This study This study This study

338F

16S

Bacteria Bacteria Micrococcus luteus spp. Pseudoxanthomonas spp. Xanthomonas spp. Mesorhizobium amorphae spp. Rhizobium spp. Brevundimonas spp. Sphingomonadales spp. Sphingomonadales spp. Bacillus infantis spp. Rhodococcus erythropolis spp. Bacteria

(gtac)7 ACTCCTACGGGAGGCAG

C

62

HOPE (Set 1/2/3/4)

NAPH-2R

ndo

GTVGAAAAAGASAT

431

51

PCR (Set A)

Modified from [68] [37]

Ac596R

nah

PCR (Set A)

[37]

RieskeF

Rieskeb

PCR (Set B)

[37]

RieskeR

Rieskeb

PCR (Set B)

[37]

MEGC-F11

RHDc

PCR (Set C)

[37]

CSYH-R10

RHDc

PCR (Set C)

[37]

652F

dbfA

PCR (Set D1)

[37]

1208R

dbfA

PCR (Set D1)

[37]

508F

dxnA/dbfA

PCR (Set D2)

[37]

1019R

dxnA/dbfA

PCR (Set D2)

[37]

NidAf

nidA

PCR (Set E)

[37]

Pdo1-r

pdo1

PCR (Set E)

[37]

COMC230F

C23Od

PCR (Set F)

[69]

COMC230R

C23Od

PCR (Set F)

[69]

PCR-screening PCR-screening Sequencing Sequencing

[70] [70] [70] [70]

M13F M13R SP6 T7 a b c d

PAH-degrading Gram positive bacteria PAH-degrading Gram positive bacteria PAH-, toluene- and biphenyl-degrading Gram negative bacteria PAH-, toluene- and biphenyl-degrading Gram negative bacteria Toluene- and biphenyl-degrading bacteria Toluene- and biphenyl-degrading bacteria PCDFs-degrading bacteria PCDFs-degrading bacteria PCDFs-degrading bacteria PCDFs-degrading bacteria PAH-degrading Gram positive bacteria PAH-degrading Gram positive bacteria Monoaromatic hydrocarbonsdegrading bacteria Monoaromatic hydrocarbonsdegrading bacteria pGEM® Vectors pGEM® Vectors pGEM® Vectors pGEM® Vectors

CRGGTGYCTTCCAGTTG TGYCGBCAYCGBGGSAWG

1100

55

CCAGCCGTGRTARSTGCA

ATGGGTGAG GACCCSGT

86

58

CCAGCCGTGGTAGCTGCA

557

GGCGACGACTAYCACGTGCT

54

TCGAAGTTCTCGCCRTCRTC TACAAVGGGCTGRTTTTCGG

512

52

GARAAVTTVGGGAACAC ATGACCACCGAAACAACCGGAACAGC

1366

60

CTGACCCATGTATTCCAGCC CGAGAACGTGCTGGGCATGAAG

561

63

AAGGCGATGTCGTGCGGC

GTAAAACGACGGCCAG CAGGAAACAGCTATGAC ATTTAGGTGACACTATAG TAATACGACTCACTATAGGG

∼1700

55

Degenerate nucleotides: Y = C/T; M = A/C; N = G/A/T/C; K = G/T; R = A/G; S = G/C; W = A/T; V = G/A/C. The low case represents the tail sequences. Rieske, Rieske nonheme iron dioxygenase gene. RHD, ring-hydroxylating dioxygenase gene. Catechol 2,3-dioxygenase gene.

experimental conditions that were previously established (Table 1). The correct amplicons were assessed under UV light after electrophoresis in 2.5% agarose gels stained with ethidium bromide.

3. Results and discussion

2.8. Nucleotide sequence accession numbers

The soil samples with high concentrations of PCDD/Fs collected from the An-Shun site were used in the batch microcosms. To stimulate microbial activities for the degradation of PCDD/Fs in the closed microcosm, oxygen gas and additional nutrients were provided. Fig. 1 shows concentration changes of major

The 16S rRNA gene sequences of clones and strains obtained in the study were deposited in EMBL/GenBank/DDBJ under the accession numbers of JX236680–JX236703.

3.1. Degradation of PCDD/Fs in soil microcosms

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Fig. 1. Degradation of PCDD/Fs in the batch microcosms. (a) Control microcosm (heat-sterilized soil) and (b) soil microcosm in the first-batch experiment. (c) In the second-batch experiment, the microcosm was enriched with soil consortia from (b) and supplemented with 50 mg of octachlorodibenzofuran powder. Symbol: ( ) OCDD; () OCDF; () 1,2,3,4,6,7,8-Hp CDF; () 1,2,3,4,7,8,9-Hp CDF. (d) The profiles of the congeners with lower concentrations detected with weekly samples from the second-batch microcosm experiment. (e) Total toxicity equivalency quantity

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dioxin congeners during the course of soil microcosm experiment. In order to differentiate any potential abiotic degradation, control microcosms containing heat-sterilized soil were monitored, too. The major forms of PCDD/Fs in the soil were fully or near-fully chlorinated, including OCDD, OCDF, and 1,2,3,4,6,7,8Hp CDF, with molar concentrations of 1570–1654, 683–913, and 80–88 ␮mol/kg (time = 0, Fig. 1(a) and (b)), respectively, which contributed to approximately 99% of the total TEQ in the soil. As shown in Fig. 1(a), the molar concentrations of the congeners in the heat-sterilized control bottles remained consistent throughout the 12-week incubation, suggesting little influence of abiotic degradation. However, as shown in Fig. 1(b), after a 6week incubation, approximately 99.9% and 95.8–99.7% of OCDF and Hp CDFs, respectively, were reduced. However, no observable reduction occurred for the OCDD within 12 weeks. It was observed that concentrations of lower chlorinated congeners such as, hexachlorodibenzofurans (Hx CDFs), pentachlorodibenzofuran (PCDF) hexachlorodibenzo-p-dioxins (Hx CDDs), pentachlorodibenzo-pdioxin (PCDD), and TCDD increased in the microcosms (see supplementary Table S1). This observation suggests a dechlorination process of highly chlorinated congeners, which led to the toxicity increase from 1580 ␮g I-TEQ/kg dry soil to 3660 ␮g I-TEQ/kg dry soil. To further validate the degradation of OCDF, microbial consortia from the first-batch experiment were then inoculated to the serum bottles (50%, w/v) containing fresh minimal medium and 50 mg of OCDF powder. In the second-batch experiment, in addition to the additive OCDF, irregular levels of other PCDD/Fs, such as OCDD and 1,2,3,4,6,7,8-Hp CDF could be detected at the onset of incubation, suggesting that the two congeners were likely carried over with the inoculum consortia or impurities added with OCDF, respectively. As shown in Fig. 1(c), the results showed that the major congener, OCDF, decreased from 5096 to 913 ␮mol/kg after 3 weeks, which corresponded to a degradation efficiency of 82.1% and a degradation rate of 199 ␮mol/kg/day. The observed degradation rate was higher than the reported values for highly (tetra- to octa-) chlorinated dioxins degraded by the pure cultures (0.03–2.72 ␮mol/L/day) [38–40] or by the compost microcosm (0.47–1.77 nmol/kg/day) [21]. The speedy OCDF reduction might be attributed to an acclimated microbial activity and a high concentration used. Still, since the initial OCDF concentration was extremely high, the contribution by the abiotic removals should not be excluded [39]. The concentrations of 1,2,3,4,6,7,8-Hp CDF declined from 368 to 46.1 ␮mol/kg, which corresponded to a degradation rate of 15.3 ␮mol/kg/day. Although degradation was not observed in the previous incubation, the OCDD concentration decreased from 110 to 13.4 ␮mol/kg within 3 weeks, which corresponded to 87.9% of the degradation efficiency and 4.6 ␮mol/kg/day of the degradation rate. Likely, the acclimation procedure with easily degradable PCDD/Fs and low initial concentrations of OCDD would be required for inducing the OCDD degradation. However, unlike the first-batch microcosm, as shown in Fig. 1(d), the results showed that the other 13 lower chlorinated congeners present at low concentrations (<10 ␮mol/kg) did not accumulate during the course of the second-batch experiment and also decreased with efficiencies in the ranges of 77.6–89.8%, respectively. The degradation of dioxins was further supported by the detection of the possible dechlorinated metabolites such as salicylic acid and catechol (see supplementary Figs. S1–S2), suggesting an oxidative metabolism enhanced by the weekly supplement with

(TEQs) of 2,3,7,8-chlorinated dioxin congeners detected with weekly samples from the second-batch microcosm experiment. The data points represent the average values derived from duplicate analyses.

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oxygen. The total TEQ of 2,3,7,8-substituted PCDD/Fs in the secondbatch microcosms was reduced with an efficiency of 91% (Fig. 1(e)), demonstrating that OCDF and other higher chlorinated DD/Fs in the second-batch microcosms could be effectively detoxified without accumulation of lower chlorinated congeners at the given conditions.

3.2. Microbial community structure of PCDD/F-degrading consortium The microbial composition inside the OCDF-enriching consortium was analyzed using the 16S rRNA gene clone library. Fig. 2 shows the phylogenetic inference of the detected bacterial

Fig. 2. Phylogenetic tree of bacterial populations based on the 16S rRNA gene sequences retrieved from the clone library, and the isolates obtained in this study. The phylogenetic tree with Aquifex pyrophilus (M83548) as the outgroup was reconstructed using a neighbor-joining algorithm. Bootstrap values greater than 60% are shown at the nodes. The scale bar corresponds to 5 nucleotide substitutions per 100 nucleotides. The inference of taxonomic phyla for the phylotypes is shown on the nodes of the tree. The percentages in parentheses show the relative abundance of each phylotype in the clone library and the obtained isolates, respectively. The specificity of each primer used in HOPE analysis is shown on the right. The primers were arranged into 4 reaction tubes in HOPE analysis, with the Bacteria-level primer (338F) and those targeting the group level distributed across reaction sets 1–4.

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populations in the OCDF-enriching consortium from the third week of incubation. As shown in Fig. 2, 12 OTUs were detected from 124 clones and phylogenetically affiliated with Gammaproteobacteria (62.9% of total clones), Alphaproteobacteria (35.4% of total clones), and Planctomyctes (0.8% of total clones). The finding was in close agreement with the microbial community structures associated with bioremediation of soil contaminated with the chlorinated pollutants [41–43]. Within these found phyla, the detected bacterial populations were related to the species described as efficiently metabolizing chlorinated DD/Fs and aromatic compounds [14,44]. For Gammaproteobacteria, the 4 detected OTUs with a similarity of 99% were assigned to the known bacterial species. Two OTUs, clones En3w21 and En3w116, accounting for 28.2% and 1.6% of total clones, respectively, were similar to Lysobacter daejeonensis and Lysobacter yangpyeongensis. One OTU, represented by clone En3w37 with 17.7% of the total clones, was closely related to Pseudoxanthomonas, whereas the other OTU, represented by clone En3w9 with 15.3% of the total clones, was placed with Xanthomonas sp. T7-07 as the closest neighbor. It has been recognized that the members of Pseudoxanthomonas and Xanthomonas were capable of degrading diesel oil, hexachlorocyclohexane, and polycyclic aromatic hydrocarbon [45–47]. Among the 7 OTUs within Alphaproteobacteria, one (clone En3W2), accounting for 16.1% of total clones in the clone library, was closely affiliated with Sphingopyxis within the order Sphingomonadales, and distinctly branched from an estrogen-degrading bacterium KC9 [48]. Another 6 (clones En3w103, En3w78, En3w134, En3w115, En3w118, and En3w107), each with 0.8%-10.5% of total clones, were closely affiliated with genera Ochrobacterium, Sinorhizobium, Rhizobium, and Brevundimonas. These populations were commonly observed in the soil and rhizosphere environments [49–51]. Within Planctomycetes, one OTU, En3W122, with low abundances (0.8%) in the clone library, was associated with Blastopirellula (99% similarity). In light of this and these studies, it suggests that a microbial community with the bacterial specialists from Alphaproteobacteria and Gammaproteobacteria may be required for enhancing the removal of highly substituted PCDD/Fs.

3.3. Isolation and identification of OCDF-growing bacterial isolates To obtain the microbes having the OCDF-degrading potential, bacterial isolation was performed with the agar plates with primary substrates for bacterial growth and OCDF as a co-metabolic substrate. In addition to the LB-minimum medium, the marine medium that provided the saline conditions was also used, since the geographic location of An-Shun site is close to the seacoast. The results showed that 35 of the 98 isolates obtained were able to repeatedly grow on OCDF-containing agar plates, and were classified to 9 known species and an uncultured bacterium within 4 recognized bacterial lineages at a >97% similarity level (Table 2). Among them, 17 and 18 isolates were isolated with OCDF-containing LB-minimal (coded by B) and marine (coded by M) agar plates, respectively. As shown in Fig. 2 and Table 2, within Actinobacteria, 5 strains, including a representative isolate B43, were similar to Micrococcus luteus, whereas the isolate B11 and another 2 were affiliated with Rhodococcus erythropolis. Previous studies reported that R. erythropolis could use dibenzofuran as the growth substrate [41], and had excellent ability to transform or mineralize lightly chlorinated DD/Fs and polychlorinated biphenyl congeners [42,52–54] with the production of biosurfactants (e.g., Trehalose lipids) to increase substrate bioavailability [55]. For M. luteus, the ability to degrade the PCDD/Fs was unknown, but Ilori and Amund identified its ability to degrade anthracene [56].

357

All isolates of Bacillus and Pseudomonas were obtained with the LB-minimal agar. The Bacillus isolates were classified to Bacillus cereus-like (2 isolates), Bacillus infantis-like (6 isolates), and Bacillus flexus-like (1 isolate) groups. Their phylogenetic relations in the tree were shown using representatives B24, B2, and B25, respectively (Fig. 2). Several studies observed that the Bacillus specialists can degrade polycyclic aromatic hydrocarbon [57], polychlorinated biphenyl [58], and dioxin congeners [44,59]. The Pseudomonas chlororaphis B40-2 strain within Gammaproteobacteria had the known capability to mineralize the chlorinated pollutants such as 1,2,3,4-tetrachlorobenzene [60]. As shown in Table 2, 14 of 17 isolates assigned to Alphaproteobacteria were obtained with the marine agar. These alphaproteobacterial strains, represented by M32, M23, M3, and M2-1, were highly related to Sphingopyxis ginsengisoli (3 isolates), Brevundimonas basaltis (4 isolates), Mesorhizobium amorphae (5 isolates), and the uncultured bacterium within Sphingomonadales (5 isolates), respectively (Fig. 2), suggesting that Alphaproteobacteria was the most abundant phylogenetic group detected using the culture-dependent method. The Sphingopyxis spp. and Brevundimonas spp. were also observed in a high abundance by the cloning and sequencing method (Fig. 2). It has been reported that these two populations were able to degrade the polycyclic aromatic hydrocarbons [61] and chlorinated aromatic compounds [62]. Using the DNA-stable isotope probing technique, Borodina and co-workers observed that Mesorhizobium spp. played a role in the transformation of the halogenated compounds [63]. The isolates related to Sphingomonadales were likely a novel species because 16S rRNA sequences had <96% similarity to the closest known relative, Porphyrobacter sanguineus, an aerobic bacterium capable of degrading biphenyl and dibenzofuran [64]. 3.4. Characterization of OCDF-growing bacterial isolates 3.4.1. PCR detection of aromatic dioxygenase genes To understand whether the obtained bacterial isolates possessed the corresponding genes that can produce key angular dioxygenases involved in the upper pathway and catechol dioxygenase involved in the lower pathway of the PCDD/Fs transformation [65], PCR detection combined with 7 primer sets (primer set A–F) was performed. As shown in Table 2, the results showed that 23 of 35 isolates showed positive detections for 1 to 4 relevant genes. Among the 23 isolates, 12 could be detected with each specific primer set B and C, suggesting that those with the putative gene coding for Rieske-type and/or aromatic ring hydroxylating (RHD) dioxygenases occurred more frequently among the bacterial isolates. In addition, the detected frequency was high for the dbfA and dxnA/dbfA gene encoding for the PCDF-related dioxygenases that were analyzed using primer sets D1 and D2 because correct amplicons could be obtained with 1 and 11 isolates, respectively. This high detected rate could be linked to the use of OCDF as the substrate for isolation. The PCR with primer set A could only reveal a positive result with B40-2 DNA as the template, and that with primer set E failed to detect any DNA from the isolates. Regarding the gene coding for catechol dioxygenase involved in the lower pathway, the primer set F could reveal positive PCR detections with the DNA of 7 isolates from M. amorphae (M3 and M29), Sphingomonadaceae bacterium M9-1, B. infantis B12, and R. erythropolis (B11, B17 and B51). Obviously, these OCDF-growing isolates displayed high genetic divergence on metabolizing aromatic compounds (i.e., different numbers and types of aromatic dioxygenase genes), and this large divergence also varied within species. It was particularly noted that certain isolates of M. amorphae (M3 and M29) and R. erythropolis (B11 and B51) contained the genes coding 2–3 dioxygenase systems (Rieske, RHD, and/or dxnA/dbfA), as well as the

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Table 2 Similarity analysis of 16S rRNA gene and PCR detection of aromatic dioxygenase genes on the bacterial strains obtained in this study. Strain

Closest relative/phylogeny

M3 M3-1 M4 M15 M29

Mesorhizobium amorphae/ Alphaproteobacteria

M23 B24-1 B35 B36

16S similarity (%)

PCR detection of aromatic dioxygenase genesa Set A

Set B

Set C

Set D1

Set D2

Set E

Set F

99 99 98 99 98

− − − − −

+ + − + +

+ + + + +

− − − − −

+ + + + +

− − − − −

+ − − − +

Brevudimonas basaltis/Alphaproteobacteria

100 100 100 100

− − − −

− − − −

+ − − −

− − − −

− − − −

− − − −

− − − −

M2 M2-1 M9-1 M26-2 M34-2

Uncultured Sphingomonadales/Alphaproteobacteria

98 98 98 98 97

− − − − −

+ + − − −

− − − − −

− − − − −

− − − − −

− − − − −

− − + − −

M1 M1-1 M32

Sphingopyxis ginsengisoli/Alphaproteobacteria

99 99 99

− − −

− − −

− − −

− − −

+ + +

− − −

− − −

B40-2

Pseudomonas chlororaphis/Gammaproteobacteria

99

+

−

+

−

−

−

−

B24 B48

Bacillus cereus/Firmicutes

100 100

− −

− −

− −

− −

− −

− −

− −

B2 B8 B10 B12 B28 B36

Bacillus infantis/Firmicutes

99 99 99 99 99 99

− − − − − −

+ + − − − −

− − − − − −

− − − − − −

+ + + − − −

− − − − − −

− − − + − −

B25

Bacillus flexus/Firmicutes

M11-2 B26 B27 B43 B52

Micrococcus luteus/Actinobacteria

B11 B17 B51

Rhodococcus erythropolis/Actinobacteria

99

−

−

−

−

−

−

−

99 100 100 100 100

− − − − −

+ − − − −

+ − − + +

− − − − −

− − − − −

− − − − −

− − − − −

99 99 99

− − −

+ + +

+ − +

+ − −

− − −

− − −

+ + +

a Primer set A for ndo/nah gene, primer set B for Rieske nonheme iron dioxygenase gene, primer set C for ring-hydroxylating dioxygenase gene, primer set D1 for dbfA gene, primer set D2 for dxnA/dbfA gene, primer set E for nid/pdo1 gene, and primer set F for catechol 2,3-dioxygenase gene.

catechol dioxygenase. This finding suggests their potential roles involved in the upper and lower pathways of PCDD/Fs degradation. Nevertheless, most of the strains related to B. basaltis, S. ginsengisoli, Sphingomonadaceae bacterium, and Bacillus spp. usually possessed less than one type of aromatic dioxygenase gene. The occurrence of these distinctive bacterial populations could reflect a functional redundancy for degrading various aromatic intermediates consequent to detoxification of PCDD/Fs, and might be an acclimation probably caused by the long-term selection of soil bacteria with high concentrations of octachlorinated DD/F in the An-Shun site. 3.4.2. Degradation of OCDF by the selected strains According to the results above (Section 3.4.1), eight isolates were selected to evaluate the degradation efficiency of OCDF at the initial concentration of 1 ␮g/g. As shown in Table 3, two isolates of M. amorphae (M3 and M29) that commonly shared 4 genes coding for aromatic dioxygenases could display averaged efficiencies of 34.2–43.4% in the degradation of OCDF. In addition, two R. erythropolis isolates that had identical 16S rRNA sequences but different numbers of aromatic dioxygenase genes were studied. The results showed that the B11 strain displayed OCDF degradation capabilities with an average efficiency of 31.3%, whereas the B51 had inferior ability for OCDF degradation. In the B11 culture, we actually detected the lower chlorinated congeners, including Hp CDFs,

and Hx CDFs, as well as the dechlorinated intermediates such as salicylic acid and catechol (see supplementary Figs. S3–S5). These observations provide support to the ability of certain R. erythropolis strains (such as B11) on the co-metabolic dechlorination and oxidative degradation of OCDF. The Bacillus sp. B2 performed high OCDF degradation capabilities (43.1%) similar to the isolate M3, although they had different numbers of aromatic dioxygenases genes. The Micrococcus sp. B43 only with the RHD gene could still achieve a

Table 3 Biodegradation of octachlorodibenzofuran by the selected bacterial isolates obtained in this study. Bacterial isolates

Growth media

Efficiency (%)

Bacillus infantis-like B2 Rhodococcus erythropolis-like B11 Rhodococcus erythropolis-like B51 Pseudomonas chlororaphis-like B40 2 Micrococcus luteus-like B43 Sphingomonadales sp. M2 1 Mesorhizobium amorphae-like M3 Mesorhizobium amorphae-like M29

LB-minimal LB-minimal LB-minimal LB-minimal LB-minimal Marine Marine Marine

43.1 31.3 −13.6 1.5 26.4 26.8 43.4 34.2

Note: (1) The cells were harvested in the exponential phase (2-day with LB-minimal medium and 4-day with marine medium). The initial cell density, 107 –109 cells/ml. (2) The data represented the average values from duplicated bottles.

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359

Table 4 Relative abundances of specific bacterial populations in the octachlorodibenzofuran-enriching consortium analyzed using the HOPE method. Bacterial targets

Target templates

Primers

Relative abundances of total Bacteria (%)a Week 0 Week 1 Week 2

Rhizobium spp. Brevundimonas spp. Pseudoxanthomonas spp. Micrococcus luteus

En3w134, En3w115 En3w118 En3w37 B43

138F 1000R 668R mod1 995F

N.D. N.D. N.D. N.D.

3.5 ± 0.2 N.D. N.D. N.D.

5.8 ± 0.0 4.4 ± 0.0 19.5 ± 2.2 10.0 ± 0.2

1.8 ± 0.0 0.6 ± 0.5 54.0 ± 1.2 12.5 ± 1.0

0

3.5

39.7

68.9

Total relative abundance of detected bacterial populations

Week 3

Note: The relative abundances of the Sphingomonadale-related populations targeted by primers 1017R and 1004F, Mesorhizobium amorphae targeted by the primer 822F, Xanthomonas targeted by the primer 558F 8tail, Bacillus targeted by the primer 183R and Rhodococcus erythropolis targeted by the primer 576R were below the detection level (ND, <0.1%) in all samples analyzed in this study. a The relative abundances were averaged from triplicate measurements and expressed as means ± standard deviations (n = 3).

OCDF degradation efficiency of 26.4%. Likely the analytical variations, the Pseudomonas sp. B40-2 and Sphingomonadales-related species M2-1 exhibited inconsistent OCDF reduction efficiencies in the duplicate analysis with a range of −32.3–35.3% (1.5%, on average) and 9.6–44.0% (26.8%, on average), respectively, so their abilities for the OCDF degradation cannot be confirmed. Although polychlorinated dioxin congeners are highly recalcitrant to the aerobic degradation, it has been reported that S. wittichii RW1 could degrade a Hx CDD with an efficiency of 10.6% within 5 days [66]. In this study, our results suggest that the members from various bacterial lineages exhibited the catabolic capabilities for highly chlorinated congeners. 3.5. Relative abundance of microbial populations as revealed by HOPE analysis In this study, a quantitative technique, HOPE combined with eleven 16S rRNA-targeted primers (Table 1) was developed to quantify the bacterial populations detected using clone library or isolation methods. These primers, including the Bacteria-specific primer 338F, were assigned to 4 multiplexing HOPE reactions (Table 1). Therefore, the relative abundance of each group detected using specific primers within the total bacteria detected using primer 338F can be determined. Prior to quantification of the samples, the HOPE analysis was optimized to ensure that each primer can be extended with the DNA of the targeting groups, but not extended with the DNA from non-targeting clones or strains. The extension efficiency (color type and fluorescence intensity) of the primers and the electrophoretic distance among the extended primers were carefully validated according to the procedures described in our previous study [24]. In the second-batch microcosm, the microbial populations of interest were monitored weekly. As shown in Table 4, the relative abundance of the targeted populations was below the detection level (<0.1%) of HOPE analysis at the beginning of OCDF-enriching microcosm. After 1 week, the Rhizobium group targeted by primer 138F was detected with relative abundances of 3.5% ± 0.2% with respect to the total bacterial population detected using primer 338F. Subsequently, in addition to the slight growth of the Rhizobium group, the sizes of 3 bacterial populations, including the 1000R-targeting Brevundimonas group, the 668R mod1-targeting Pseudoxanthomonas group, and the 995Ftargeting M. luteus group, were magnified to abundances of 4.4% ± 0.0%, 19.5% ± 2.2%, and 10.0% ± 0.2%, respectively. For the sample in Week 3, the 668R mod1-targeting Pseudoxanthomonas group, accounting for a relative abundance of 54.0% ± 1.2%, became highly predominant in the community. Similarly, the relative abundance of the 995F-targeting Micrococcus group also increased to 12.5% ± 1.0%. By contrast, the average relative abundance of the 138F-targeting Rhizobium group and 1000R-targeting Brevundimonas group decreased by 4.0% and 3.8%, respectively. The 7 remaining bacterial populations were not detected throughout the

experiments by HOPE analysis, suggesting relatively low abundance in the microcosms. Overall, these four detected populations increased from abundance below the detection level (<0.1%) to a total abundance as high as 68.9% of the total bacterial population. The great population dynamics could be mainly attributed to the soil organic carbons and medium nutrients used for bacterial growth in the microcosm. For example, most of the Brevundimonas isolates were not supported to have the genetic potentials for the degradation of dioxin congeners and metabolites (Table 2). However, the increase in relative abundances of the M. luteus group might be in part associated with the degradation of PCDD/Fs due to its degradative ability of OCDF (Table 3). Because we did not obtain any isolates related to the members of Pseudoxanthomonas and Rhizobium groups, their roles in the degradation of OCDF and metabolites are unknown and further studies should be needed. 4. Conclusions In summary, the results of this study showed that hepta- and octa-chlorinated DD/Fs could be effectually reduced with the substantial decrease of the total TEQ by the diversified indigenous microbial populations in a batch microcosm stimulated with oxygen. Our results further suggested for the first time that certain specialists from Rhodococcus, Micrococcus, Bacillus, and Mesorhizobium possessed the capabilities of the OCDF degradation. These findings advance current understanding on the biodegradation of highly chlorinated compounds, and suggest a potential of enhanced biodegradation technologies in remedying the pollution of highly substituted PCDD/Fs at An-Shun. Acknowledgments This work was supported by the joint grants of the National Science Council, Taiwan, and the China Petrochemical Development Corporation Company (100-2628-E-006-026-MY3, 100-2622-E006-010-CC1). The authors thank the Worthies Environmental Engineering Consultants Company for their technical assistance in the analysis of PCDD/Fs. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.jhazmat. 2013.07.039. References [1] K.C. Jones, P. de Voogt, Persistent organic pollutants (POPs): state of the science, Environmental Pollution 100 (1999) 209–221. [2] C. Pappe, Dioxin, patterns and source identification, Fresenius’ Journal of Analytical Chemistry 348 (1994) 63–75. [3] S. Masunaga, T. Takasuga, J. Nakanishi, Dioxin and dioxin-like PCB impurities in some Japanese agrochemical formulations, Chemosphere 44 (2001) 873–885.

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