Article pubs.acs.org/est
Cultivation-Independent Identification of Candidate Dehalorespiring Bacteria in Tetrachloroethylene Degradation Shouhei Yamasaki,† Nobuhiko Nomura,† Toshiaki Nakajima,† and Hiroo Uchiyama*,† †
Graduate School of Life and Environmental Sciences, University of Tsukuba, Tsukuba, Ibaraki, Japan ABSTRACT: Tetrachloroethylene (PCE) is one of the major pollutants and is degraded by dissimilation by dehalorespiring bacteria. The dehalorespiring bacteria are anaerobic, and most cannot be cultured by conventional agar plating methods. Therefore, to identify the dehalorespiring bacteria that dissimilatively degrade PCE, a cultivation-independent method is required. To achieve accurate and detailed analysis of the bacteria, we developed a novel stable isotope probing (SIP) method. This technique involves 2 steps, namely, a labeling step, in which a labeled carbon source is incorporated into the sample’s DNA, and an analysis step, in which the DNA is isolated, fractionated, and analyzed by polymerase chain reaction denaturing gradient gel electrophoresis (PCR-DGGE). Subsequently, 16S rRNA sequencing and phylogenetic analysis were performed to identify the bacteria. Initially, we examined the effectiveness of this method by using Dehalococcoides ethenogenes 195 consortium as a defined model system. The result indicated the method was able to correctly identify the dehalorespiring bacteria D. ethenogenes 195 from the consortium. Moreover, in an artificially contaminated microcosm experiment, we confirmed that the method was able to identify the indigenous dehalorespiring bacteria Dehalobacter sp. Thus, we concluded that this novel method was a feasible tool to identify dehalorespiring bacteria in natural environments.
■
INTRODUCTION Tetrachloroethylene (PCE) is one of the major pollutants causing soil and groundwater contamination. PCE is degraded by the dissimilatory process of dehalogenation under anaerobic conditions. Anaerobic dehalogenating bacteria are classified into 3 types, namely, obligately dehalorespiring bacteria, facultatively dehalorespiring bacteria, and cometabolic dehalogenating bacteria.1 Dehalorespiring bacteria are microbes that rely on the energy acquired by reductive dehalogenation.1,2 Previous studies have reported that dehalorespiring bacteria play an important role in the degradation of PCE, and 2 genera in particular, namely, Dehalococcoides and Dehalobacter, have been identified as the main representatives. The Dehalococcoides ethenogenes 195 bacterium can completely degrade PCE to ethylene,3 and therefore, the Dehalococcoides genus has received much attention as the most important microbe in the degradation of PCE and other chloroethenes. The study of dehalorespiring bacteria has been mainly performed with almost pure liquid culture of strains and the genes of those bacteria.4−7 Therefore, the current knowledge regarding these bacteria has been acquired using culturable strains. However, most dehalorespiring bacteria are obligate anaerobic and difficult to culture.8 In particular, Dehalococcoides has not been reported to be cultured on agar medium. Therefore, to more precisely identify dehalorespiring bacteria, it is important to use cultivation-independent methods. Stable-isotope probing (SIP) is a well-known cultivationindependent method for the study of microorganisms capable of degrading various kinds of pollutants.9 However, SIP is © 2012 American Chemical Society
inadequate for the identification of dissimilate-degrading microbes such as the dehalogenate-degrading microbe group that targets chloroethenes, because SIP is specific for microbes that can use pollutants by assimilation, not dissimilation. To accurately and comprehensively study dechlorinating-degrading microbes, it is important to use a cultivation-independent method for dissimilating-degrading microbes. Unfortunately, an effective cultivation-independent method to identify dissimilate-degrading microbes has not yet been established. The main purpose of this study was to develop a novel cultivation-independent analytical method that can identify obligately and facultatively dehalorespiring bacteria capable of PCE degradation. We tentatively named this method stable isotope probing for dissimilation (SIP-D). In this study, we initially performed the SIP-D method on an array of bacteria including D. ethenogenes 195 as the positive control strain. We confirmed that the SIP-D method successfully identified D. ethenogenes 195 as a dehalorespiring bacterium and, subsequently, applied the SIP-D method to an artificially PCEcontaminated microcosm. Analysis of the microcosm identified the indigenous active dehalorespiring bacteria Dehalobacter sp. as a candidate PCE-dissimilative-degrading bacterium. From these results, we concluded that the SIP-D method is an Received: Revised: Accepted: Published: 7709
April 1, 2012 June 5, 2012 June 18, 2012 June 18, 2012 dx.doi.org/10.1021/es301288y | Environ. Sci. Technol. 2012, 46, 7709−7716
Environmental Science & Technology
Article
could work. The stock culture of D. ethenogenes 195 consortium prepared using previously described methods was used here. Chloroethenes in the stock culture of D. ethenogenes 195 consortium were purged by N2−CO2 (70:30, v/v) gas bubbling. After purging, the chloroethenes in the stock culture was measured to make sure it is free of chloroethanes by gas chromatography (GC). The chloroethene-free stock culture (100 mL) was transferred into a 100-mL vial that is further purged with N2−CO2 (70:30, v/v), the culture was split into two subsamples. After the culture was transferred, the preparation of the WITH-PCE sample entailed the addition of 12 μmol of PCE and 24 μmol of butyric acid as the electron acceptor and hydrogen donor, respectively (Figure 1a). The WITHOUT-PCE sample (Figure 1a) required 24 μmol of butyric acid, but no PCE, to be added to the culture. Following this, both WITH-PCE and WITHOUT-PCE treatments were spiked with 13C-fumarate. Moreover, unlabeled controls for each treatment were prepared using unlabeled (12C) fumarate instead of 13C. All 4 treatments were incubated at 30 °C with gentle shaking. The chloroethene concentration during the incubation was measured by GC as described below. When the PCE concentration dropped to below the detection limit, the WITH-PCE experiments were amended with an additional 12 μmol of PCE and continued. After 7 days incubation, all cultures were filtered, the bacterial cells were trapped on a 0.2μm pore size membrane filter (25-mm diameter; Advantec, Toyo Roshi Kaisha, Ltd., Japan), and the filters were stored at −30 °C until further processing. Application of the SIP-D Method in PCE-Contaminated Microcosms. Application test was carried out to investigate its functionally in environmental sample. Artificially contaminated microcosms described above were used. Chloroethenes in the artificially contaminated microcosm were purged by N2 gas bubbling, and the chloroethene-free artificially contaminated microcosm sample (50 mL) was anaerobically transferred into a 50-mL vial. After the sample was transferred, the WITH-PCE sample was treated with 6 μmol of PCE as an electron acceptor and 13C-fumarate as labeling carbon source. Comparatively, the WITHOUT-PCE sample was treated with only 13C-fumarate. As controls, two other WITH-PCE or WITHOUT-PCE samples were subjected to unlabeled (12C) fumarate instead of 13C. All 4 treatments were incubated at 30 °C with gentle shaking. The chloroethene concentration during the experiment period was measured by GC. When the levels of PCE declined to less than the detection limit, the WITH-PCE treatments were amended with 6 μmol of PCE. After 7-day incubation, each sample containing bacterial cells was collected by centrifugation at 16,000 × g for 10 min. The soils were used in the total DNA extraction experiments. Analytical Methods. For analysis of the chloroethenes, the 100 μL of headspace gas from the vial was injected into the gas chromatograph system (G-3900; Hitachi Science Systems, Ltd., Japan) equipped with a FID detector and silicone DC-200 packed column (2-m long, 3-mm i.d.; GL Science, Tokyo, Japan). Helium (99.99%) was used as a carrier gas, and the injector and detector temperatures were maintained at 200 °C. The column oven temperature was held at 120 °C. DNA Extraction. The filters were transferred into Lysing Matrix E tubes (MP Biomedicals, Santa Ana, CA), and the total DNA was extracted from bacterial cells by using a bead beating and CTAB method.11 The total DNA was dissolved in TE buffer and stored at −30 °C until use.
effective cultivation-independent method to identify microbes capable of PCE degradation by dissimilation.
■
MATERIALS AND METHODS Chemicals. 13C labeled fumaric acid (13C 99%) was purchased from Cambridge Isotope Laboratories (Andover, MA, USA). The 13C labeled fumaric acid was neutralized with sodium hydroxide and used in the experiments. Culture and Growth Conditions. (i). Dehalococcoides ethenogenes 195 Consortium. The D. ethenogenes 195 consortium was kindly provided by Ruth E. Richardson of Cornell University (Ithaca, NY). The D. ethenogenes 195 consortium was grown in the previously described medium.10 The culture medium was prepared in 1-L medium bottles, and the bottle was sealed with a Teflon-coated butyl rubber stopper and an open-hole screw cap. The culture medium contained (per liter) 0.2 g of NH4Cl, 0.1 g of K2HPO4·3H2O, 0.055 g of KH2PO4, 0.2 g of MgCl2·6H2O, 0.001 g of sodium resazurin, 0.1 g of FeCl2·4H2O, 0.5 g of Na2S·9H2O, 6.0 g of NaHCO3, and 10 mL of a trace metal solution; the headspace was replaced with N2−CO2 (70:30, v/v). After autoclaving the bottles, 120 μmol of PCE, 480 μmol of butyric acid, 500 μL of vitamin solution, and 200 μL of pre-fermented yeast extract solution were added per liter of medium. The trace metal solution, vitamin solution, and pre-fermented yeast extract were prepared as previously described.10 The inocula were incubated in the dark at 30 °C with gentle shaking. (ii). Artificially Contaminated Microcosms. The sediment sample used in the artificially contaminated microcosm experiment was collected from a nonpolluted pond in Tsukuba, Ibaraki, Japan. The collected soil was amended with 120 μM of PCE and stored in the dark until use. The artificially contaminated microcosms were prepared in the 1-L medium bottle, and the bottle was sealed with a Tefloncoated butyl rubber stopper and an open-hole screw cap. The culture medium of the artificially contaminated microcosms contained (per liter) 0.5 g of Na2S·9H2O and 0.5 g of L-cysteine hydrochloride·H2O. Following sterilization by autoclave, the final concentration of the medium was adjusted by the addition of 120 μmol of PCE and 100 mL (about 50 g) of PCEamended sediment (per liter of medium). The artificially PCEcontaminated microcosms were incubated in the dark at 30 °C with gentle shaking. Fumarate Assimilation Test. A fumarate assimilation test was done before the test of the SIP-D method for the D. ethenogenes 195 consortia. The fumarate assimilation test involved preparing 50 mL of the defined D. ethenogenes 195 medium, but without pre-fermented yeast extract, in 50-mL vials (actual volume, 69 mL). Basal medium was produced by inoculating the stock culture of D. ethenogenes 195 (1 mL) into the medium and adding 2.5 mmol of fumarate. Fumarate-free medium identical to the basal medium was also prepared but without fumarate. In addition, control media that were not amended with PCE and fumarate (fumarate and PCE-free) or not PCE (PCE-free) were also prepared. All treatments were incubated at 30 °C with shaking. During the degradation of PCE, 2 mL of the subsample was regularly collected to measure the growth of D. ethenogenes 195 as a function of the 16S rRNA gene copy number by quantitative PCR. Evaluation of the Accuracy of the SIP-D Method Using the Consortia Containing D. ethenogenes 195. Evaluation test using the consortia containing D. ethenogenes 195 was carried out to ascertain whether the SIP-D method 7710
dx.doi.org/10.1021/es301288y | Environ. Sci. Technol. 2012, 46, 7709−7716
Environmental Science & Technology
Article
modification; TE buffer was used for final elution instead of DES (DNase/Pyrogen-Free water). The extracted DNA was stored at −30 °C until use. Ultracentrifugation and Fractionation. To separate 13C labeled DNA, the extracted DNA obtained in the SIP-D method was used. The extracted DNA was added to polyallomer Quick-Seal centrifuge tubes (13 × 51 mm, 5.1 mL; Beckman Coulter, Inc., USA) along with a CsCl gradient buffer. The CsCl buffer was prepared by adding 4.5 g of CsCl to 4.4 mL of TE buffer (pH 8.0) and 0.1 mL of EtBr (10 mg/ mL). The tubes were centrifuged at 177,000 × g for 40 h at 20 °C in a Vti 65.2 rotor in an Optima L-70K Ultracentrifuge (Beckman Coulter, Inc., USA). After ultracentrifugation, the gradients were fractionated from the bottom by using a Perista pump (ATTO Corporation, Tokyo, Japan) according to a previously described procedure12 with minor modifications; approximately 250 fractions (20 μL) were collected. The buoyant densities of the 3 fractions selected from the bottom, middle, and top layers were measured, and the standard curve of buoyant density versus fraction number was calculated. Ethidium bromide was extracted from the fractions by TE-saturated 3-methyl-1-butanol. DNA in the fraction was purified by glycogen-assisted PEG precipitation according to a previously described procedure,12 and the purified DNA fractions were resuspended in TE buffer and stored at −30 °C until use. Quantitative PCR. To determine the 16S rRNA gene copy number, quantitative PCR was performed. The bacterial universal primer pairs of 357F (5′-CCTACGGGAGGCAGCAG-3′) and 518R (5′-GTATTACCGCGGCTGG-3′) were used for real-time PCR of the 16S rRNA gene. The reaction was performed using a LightCycler FastStart DNA MasterPLUS SYBER Green I kit (Roche Molecular Biochemicals, Indianapolis, IN, USA) on a LightCycler 1.5 system (Roche Diagnostics, Mannheim, Germany) according to the manual. Calibration curves for the 16S rRNA gene copies were made using serial dilutions of PCR amplicons of E. coli DH5α. The calibration curve and calculation of the copy numbers were determined by LightCycler software version 3.5 (Roche Diagnostics, Mannheim, Germany). The PCR consisted of an initial denaturation step of 10 min at 94 °C, followed by 40 cycles of 30 s at 94 °C, 30 s at 55 °C, and 15 s at 72 °C, with a final extension for 5 min at 72 °C. In the assimilation test of fumarate, the Dehalococcoidesspecific primers of DHC 793f (5′-GGGAGTATCGACCCTCTCTG-3′) and DHC 946r (5′CGTTYCCCTTTCRGTTCACT-3′) were used for real-time PCR. The real-time PCR procedure was performed as described by Yoshida et al.13 PCR-Denaturing Gradient Gel Electrophoresis (PCRDGGE). To identify the candidate unique bacterium in the heavy DNA fraction, PCR-DGGE analysis was performed. For DGGE analysis, the 16S rRNA gene in each buoyant density fraction was amplified. The universal primer pair of 357F-GC (5′-CGCCCGCCGCGCCCCGCGCCCGTCCCGCCGCCCCCGCCCG-CCTACGGGAGGCAGCAG-3′) and 518R was used to obtain PCR products. The PCR consisted of an initial denaturation step of 5 min at 94 °C and a hot start at 94 °C, followed by 28 cycles of 30 s at 94 °C, 30 s at TA °C, and 15 s at 72 °C, with a final extension for 5 min at 72 °C. In the first 20 cycles, the TA decreased by 0.5 °C every cycle, from 65 °C in the first cycle to 55 °C in the twentieth cycle. This touchdown procedure was previously described.14 The PCR
Figure 1. Details of the SIP-D method. The SIP-D method for identifying cultivation-independent dehalorespiring bacteria consists of 2 steps, namely, the labeling step (a) and the analysis step (b). (a) Active bacteria in two subsamples (WITH-PCE and WITHOUTPCE) are labeled with the 13C-labeled general carbon source, and 13Clabeled DNA is extracted and separated, respectively. (b) The 13CDNA prepared in the labeling step was subjected to PCR-DGGE analysis. Comparing the DGGE profiles reveals candidate dehalorespiring bacteria.
DNA extraction of the soils were performed using the soil samples and the Fast DNA Kit (MP Biomedicals, Santa Ana, CA) according to the manufacturers’ manual with a minor 7711
dx.doi.org/10.1021/es301288y | Environ. Sci. Technol. 2012, 46, 7709−7716
Environmental Science & Technology
Article
bank of Japan using the BLAST algorithm. Sequences were aligned, and phylogenetic analysis was performed with the CLUSTALX15 software program using the neighbor-joining method,16 and a bootstrap analysis based on 1,000 replicates was used to place confidence estimates on the tree.
products were purified using the UltraClean PCR Clean-up DNA Purification kit (MO BIO Laboratories, Inc., Carlsbad, CA, USA). The DGGE was performed using a Dcode Universal Mutation Detection system (Bio-Rad Laboratories, Hercules, CA, USA) with a gel acrylamide concentration of 8% and a denaturing gradient of 30%−60%. The electrophoresis was run for 18 h at 36 V, and the gels were subsequently stained with SYBR Gold (Molecular Probes, Eugene, OR) and photographed using a UV transilluminator (AE-691; ATTO Corporation, Japan). Some unique DGGE bands were excised from the gel under UV illumination, placed in 50 μL of TE buffer, and washed by pipetting to remove the contaminating DNA. The gels were resuspended with 50 μL of TE buffer and incubated overnight at 4 °C. The supernatant (0.5 μL) was used as the template DNA in the PCR reamplification using the 357F and 518R primer pair as described above. The PCR product was subsequently purified and used in DNA sequence analysis. 16S rRNA Gene Library of the Artificially Contaminated Microcosm. 16S rRNA gene library was constructed to isolate the candidate dehalorespiring bacteria. The primers 27F (5′-AGAGTTTGATCCTGGCTCAG-3′) and 1494R (5′TGACTGACTGAGGYTACCTTGTTACGACTT-3′) were used for PCR with a template of genomic DNA extracted from the artificially contaminated microcosm. The resulting 1.5kb PCR amplicon was cloned into E. coli DH5α by using the Teasy Vector system I (Promega Corporation, Madison, WI, USA). The clones were screened for the presence of the insert by PCR using the vector-specific primers SP6 (5′-ATTTAGGTGACACTATAG AATA-3′) and T7 (5′-TAATACGACTCACTATAGGG-3′); the amplicons from the 40 positive clones were used in the next step. To confirm that the insert derived from the candidate dehalorespiring bacteria’s 16S rRNA gene obtained by DGGE analysis, the above amplicons were reamplified with the specific primer DhbF (5′-CTGGATTGACGGTACCTACGA-3′) and the universal primer 1494R. The DhbF primer was designed as a specific primer for candidate dehalorespiring bacteria in the artificially contaminated microcosm. The PCR consisted of an initial denaturation step of 5 min at 94 °C and a hot start at 94 °C, followed by 28 cycles of 30 s at 94 °C, 30 s at 58 °C, and 15 s at 72 °C, with a final extension for 5 min at 72 °C. The PCR products were visualized by agarose gel electrophoresis and stained with ethidium bromide. A clone containing about 900 bp of PCR product was identified as the candidate dehalorespiring bacteria’s clone. The 1.5-kb 16S r RNA gene fragment from the clone was used in the DNA sequence analysis. 16S rRNA Sequencing and Phylogenetic Analysis. To determine the phylogenetic position of candidate bacteria, 16S rRNA sequencing and phylogenetic analysis were carried out. Cycle sequencing of the PCR products was performed using the ABI Prism BigDye Terminator Cycle Sequencing Ready Reaction kit (Applied Biosystems, Tokyo, Japan) and the sequencing primers 27F, 357F, 926F (5′-ACTCAAAGGAATTGACGG-3′), 1111F (5′-GCAACGAGCGCAACCC-3′), 518R, 1100R (5′-GGGTTGCGCTCGTTG-3′), and 1494R. The cycle sequencing reaction and purification were performed according to the manufacturers’ protocols. The sequencing analysis was carried out on a 3130 Genetic Analyzer (Applied Biosystems, Japan). In the artificially contaminated microcosm experiment, the gene sequences were compared with those in the DNA data
■
RESULTS Features of the SIP-D Method. The SIP-D method was constructed as a cultivation-independent identification technique for microbes capable of degradation by dissimilation, such as dehalorespiring bacteria. As shown in Figure 1, the SIP-D method consists of 2 steps, namely, the labeling step and the analysis step. In the labeling step, labeled carbon (13C) from the growth carbon source (i.e., organic acid) is incorporated into the biomass (nucleic acid, protein, lipid, etc.) of the active microorganisms in the sample. Two samples, namely, WITHPCE and WITHOUT-PCE, were prepared and incubated with the 13C-labeled carbon source. The presence or absence of PCE was expected to alter the 13C-labeling. That is, in the presence of PCE, the dehalorespiring bacteria grow using dehalorespiration, and, consequently, the dehalorespiring bacteria assimilate the 13C-labeled carbon source. To the contrary, in the absence of PCE, the dehalorespiring bacteria cannot get energy for growth, and, consequently, the dehalorespiring bacteria cannot assimilate the 13C-labeled carbon source. The nondehalorespiring bacteria can assimilate the 13C-labeled carbon source regardless of the presence or absence of PCE. These differences are important in the subsequent analysis step. After 13Clabeling, the total DNA is extracted from the 2 treatments, and the 13C-labeled heavy-DNA is separated from the light-DNA by density gradient ultracentrifugation. The obtained 13C-labeled DNAs are used for the identification of dehalorespiring bacteria in the next analysis step. In the analysis step, the dehalorespiring bacteria are identified by comparing the heavy-DNA band profiles from the DGGE experiments of the WITH-PCE and WITHOUT-PCE treatments (Figure 1b). If the unique band is observed in the heavyDNA fraction of the WITH-PCE treatment, it is deemed to be dehalorespiring bacteria. The SIP-D method effectively deduces dehalorespiring bacteria by this comparison, because the dehalorespiring bacteria can get energy by dehalorespiring in the presence of PCE, which, thereby, causes active cell growth. During growth, the 13C-labeled carbon source is incorporated into the DNA and forms heavy DNA. That is, the activities of dehalorespiring bacteria are controlled by the presence of the respiratory substrate (i.e., PCE). Therefore, we expected that DGGE is a suitable technique for this precise comparison analysis. Examination of Effectiveness of the SIP-D Method Using Consortia Containing D. ethenogenes 195. (i). Confirmation of Suitable Carbon Source for DNA Labeling. To examine the effectiveness of the SIP-D method for the detection of dehalorespiring bacteria, we ascertained whether the SIP-D method could identify the dehalorespiring, PCE-degrading bacterium D. ethenogenes 195 in the consortia. We tested whether fumarate could be used as the carbon source and label the DNA of the above strain. To confirm that fumarate is suitable as a carbon source in the SIP-D method, we investigated whether D. ethenogenes 195 could grow in the presence of fumarate as a carbon source. As shown in Figure 2, D. ethenogenes 195 grew and degraded PCE with fumarate as a carbon source. Without PCE, fumarate was not used as a 7712
dx.doi.org/10.1021/es301288y | Environ. Sci. Technol. 2012, 46, 7709−7716
Environmental Science & Technology
Article
Figure 2. Fumarate assimilation test of Dehalococcoides ethenogenes 195. The assimilation ability was measured by growth of D. ethenogenes 195. D. ethenogenes 195 was cultivated in basal (■), PCE-free basal (●), fumarate-free (□), and PCE and fumarate-free (○) medium. See details in the text. The error bars represent the standard deviation (n = 3). Inset graph shows PCE degradation.
carbon source by D. ethenogenes 195. Therefore, we concluded that fumarate is able to utilize as the 13C-labeled carbon source to label the DNA. ii). DNA Labeling of Active Microbes in the Labeling Step. In the labeling step, after 7-day incubation with or without PCE, all the cultures were filtered, and the total DNA was extracted. The experiment in which 12C-fumate was utilized instead of 13C-fumarate was used as the control. The heavy DNA was separated from the light-DNA by CsCl density gradient ultracentrifugation. After ultracentrifugation, the gradients were fractionated in eppendorf tubes drop-by-drop (∼20 μL) from the bottom to the top. Following this, each DNA fraction was purified, and the 16S rRNA gene copy number was determined by quantitative PCR. Unlabeled lightDNA (12C-DNA) was observed in the buoyant density from approximately 1.575 to 1.580 g/mL. In the presence of 13Cfumarate, the heavy DNA was identified in both the WITHPCE and WITHOUT-PCE samples (Figure 3). The labeled heavy DNA was observed in the buoyant density fractions (≥∼1.580 g/mL). iii). Identification of Dehalorespiring Bacterium by the Analysis Step. The analysis step was performed to identify the active dehalorespiring bacterium. To identify the unique bacterium in the heavy DNA fraction, the fractions ranging in buoyant density from approximately 1.575 to 1.595 g/mL were applied to DGGE. Comparison of the light- and heavy-fraction DGGE profiles enabled the selection of the bands localized to the heavy fractions. In the WITH-PCE samples, 3 bands were unique to the heavy fraction (Figure 4, bands I, II, and III). Comparatively, in the WITHOUT-PCE samples, 2 DNA bands were unique to the heavy fraction (Figure 4, bands IV and V). The bands II and IV were located at the same migration distances, and III and V were also located at the same migration distances. Interestingly, band I was unique to only the heavy fraction of the WITH-PCE sample. Therefore, band I was identified as a dehalorespiring bacterium. To confirm this conclusion, the band I was excised from the gel, and the part of 16S rRNA gene (about 160 bp) encoded in the
Figure 3. 16S rRNA gene distribution of Dehalococcoides ethenogenes 195 in CsCl density gradient. (a) Experiment was performed with PCE (WITH-PCE in Figure 1). The lines indicate the 13C-labeled sample (●) and unlabeled sample (○). (b) Experiment was performed without PCE (WITHOUT-PCE in Figure 1). The lines indicate the 13 C-labeled sample (■) and the unlabeled sample (□). Each symbol represents each fraction. Ratios are based on the number of gene copies in each buoyant density fraction relative to the buoyant density fraction that contained the maximum number of 16S rRNA gene copies in each sample.
extracted DNA was sequenced. These results clearly demonstrate that the band I was derived from D. ethenogenes 195 (100% homology). Application of the SIP-D Method to Environmental Sample. The above results validate the ability of the SIP-D method to detect dehalorespiring bacteria. Therefore, the SIPD method was subsequently utilized to investigate its functionality in environmental samples by using artificially contaminated microcosms. After stable-isotope labeling in the artificially contaminated microcosms, the DNA was extracted and fractionated by CsCl gradient according to the scheme represented in Figure 1a. As shown in Figure 5, the light DNA occupied the fractions ranging in buoyant density from approximately 1.560 to 1.570 g/L in the WITH-PCE and WITHOUT-PCE samples, whereas the heavy DNA occupied the fractions with buoyant density ≥∼1.570 g/L in the WITHPCE sample (Figure 5a). In the WITHOUT-PCE sample, the heavy DNA occupied the fractions with buoyant density of ≥∼1.575 g/L (Figure 5b). The next stage of the analysis step (Figure.1b) revealed the candidate bands of the dehalorespiring bacteria (Figure 6). Unique bands A, B, and C in the WITHPCE sample and D and E in the WITHOUT-PCE sample were observed in each heavy fraction. The bands B and D migrated the same distances, and C and E were also migrated the same distances. Therefore, the B, C, D, and E bands were recognized as nondehalorespiring bacteria because they were not unique to 7713
dx.doi.org/10.1021/es301288y | Environ. Sci. Technol. 2012, 46, 7709−7716
Environmental Science & Technology
Article
Figure 4. DGGE profiles of the 13C-labeled bacterial community in the consortia. a) DGGE profiles of the WITH-PCE samples. The buoyant density gradually increases from lane 1 to 13. b) DGGE profiles of the WITHOUT-PCE samples. The buoyant density gradually increases from lane 14 to 24. Lanes 1 and 14 are approximately 1.5750 g/mL. Lanes 13 and 24 are approximately 1.6000 g/mL. Lane M is the DGGE marker (DGGE Marker II; Nacalai Tesque, Kyoto, Japan). Bands I, II, and III are unique to the heavy fraction in the WITH-PCE sample, and IV and V are unique to the heavy fraction in the WITHOUT-PCE sample. Figure 5. 16S rRNA gene distribution of PCE-contaminated soil microcosm in CsCl density gradient. (a) Analysis of the WITH-PCE sample. The lines indicate the 13C-labeled sample (●) and the unlabeled (12C) sample (○). (b) Analysis of the WITHOUT-PCE sample. The lines indicate the 13C-labeled sample (▲) and the unlabeled (12C) sample (Δ). Ratios are based on the number of gene copies in each buoyant density fraction relative to the buoyant density fraction that contained the maximum number of 16S rRNA gene copies in each sample.
the heavy fraction in the WITH-PCE sample. In contrast, band A was unique to the heavy fraction in the WITH-PCE sample and was recognized as a candidate dehalorespiring bacterium. The band A was excised from the gel, and the part of the 16S rRNA gene (∼160 bp) encoded in the extracted DNA was sequenced. The sequencing suggested that the band A belonged to the genus Dehalobacter. This suggestion was confirmed by comparison with the almost-full-length 16S rRNA gene (1538 bp), including the band A sequence obtained from the gene library of the artificially contaminated microcosm as described in Materials and Methods. The candidate dehalorespiring bacterium was most closely related to Dehalobacter sp. WL (97% homology). A neighbor-joining phylogenetic tree derived from the 16S rRNA gene sequences of dehalorespiring bacteria with the detected Dehalobacter was drawn (Figure 7). The detected Dehalobacter did not participate in the cultured PCE-degrading Dehalobacter cluster.
which comprises obligately dehalorespiring bacteria.17 Although Dehalobacter is a cultivated PCE-degrader, and formed a cluster, the detected Dehalobacter sp. was not involved in it (Figure 7). Therefore, this result suggests that the identified bacterium is a novel strain of PCE-degrading Dehalobacter. Anaerobic dehalogenating bacteria are classified into 3 types, namely, obligately dehalorespiring bacteria (i.e., Dehalococcoides,3,18−20 Dehalobacter17,21−23), facultatively dehalorespiring bacteria (i.e., Desulf itobacterium,24 Sulf urospirillum25), and cometabolic dehalogenating bacteria (i.e., Clostridium26). The SIP-D method is targeted to identify obligately and facultatively dehalorespiring bacteria. In this study, the SIP-D method identified Dehalococcoides and Dehalobacter strains that are obligately dehalorespiring bacteria. Therefore, we have revealed that the SIP-D method is able to detect obligately dehalorespiring bacteria; however, it remains to be demonstrated whether the SIP-D method can detect facultatively dehalorespiring bacteria. In the absence of PCE, facultatively dehalorespiring bacteria can utilize other respiratory substrates (i.e., nitrate, sulfate) for growth. Therefore, if large amounts of other respiratory substrates are present in analytical samples, the SIP-D method might not be able to detect the facultatively dehalorespiring bacteria. Thus, future experiments are necessary to verify the effectiveness of the SIP-D method in the identification of facultatively dehalorespiring bacteria.
■
DISCUSSION In this study, we developed the SIP-D method as a cultivationindependent method to identify active and dissimilative dehalogenate-degrading microbes. The SIP-D method is based on energy acquisition in the dehalorespiration and carbon assimilation of dehalorespiring bacteria. To examine the effectiveness of the SIP-D method, we performed the SIP-D method with a defined consortium containing D. ethenogenes 195, which is a representative dehalorespiring bacterium. As expected, the SIP-D method identified D. ethenogenes 195 in the authentic consortia, and, therefore, we concluded that the SIP-D method is an effective method for detecting dehalorespiring bacteria. In the experiment using the artificially contaminated soil microcosm, the SIP-D method determined that the candidate dehalorespiring bacterium belonged to the Dehalobacter genus, 7714
dx.doi.org/10.1021/es301288y | Environ. Sci. Technol. 2012, 46, 7709−7716
Environmental Science & Technology
Article
rium.24 However, a detailed assimilation test has not been conducted in all dehalorespiring bacteria. Therefore, it is unclear whether a common carbon source is efficiently incorporated into the dehalorespiring bacteria. For effective identification of dehalorespiring bacteria, it may be appropriate to use mixtures of labeled carbon sources or identify a carbon source that is common to dehalorespiring bacteria. Dehalorespiring bacteria are involved in the degradation of many chlorinated hydrocarbons (i.e., polychlorinated biphenyls,28 polychlorinated dibenzo-p-dioxins28,29), and we suggest the SIP-D method is a widely applicable method to identify the degraders. In the future, we will utilize the SIP-D method to identify the pollutant degraders in an authentic contaminated environment.
■
AUTHOR INFORMATION
Corresponding Author
*Phone: +81-29-853-6626. Fax: +81-29-853-6626. E-mail:
[email protected]. Notes
Figure 6. DGGE profiles of 13C-labeled bacterial community in the PCE-contaminated soil microcosm. a) DGGE profiles of the WITHPCE sample. The buoyant density gradually increases from lane 1 to 12. b) The DGGE profiles of the WITHOUT-PCE sample. The buoyant density gradually increases from lane 13 to 24. Lanes 1 and 14 are approximately 1.5645 g/mL, and lanes 12 and 24 are approximately 1.5840 g/mL. The lane M is the DGGE marker. Bands A, B, and C are unique to the heavy fraction in the WITH-PCE sample, and D and E are unique to the heavy fraction in the WITHOUT-PCE sample.
The authors declare no competing financial interest.
■
ACKNOWLEDGMENTS We thank Ruth E. Richardson for providing the Dehalococcoides ethenogenes 195 consortium. This work was supported by a Grant-in-Aid for Scientific Research (B) (no. 21310050, to H.U.).
■
REFERENCES
(1) Hiraishi, A. Biodiversity of dehalorespiring bacteria with special emphasis on polychlorinated biphenyl/dioxin dechlorinators. Microbes Environ. 2008, 23 (1), 1−12. (2) Holliger, C.; Wohlfarth, G.; Diekert, G. Reductive dechlorination in the energy metabolism of anaerobic bacteria. FEMS Microbiol. Rev. 1999, 22, 383−398. (3) Maymo-Gatell, X.; Chien, Y.-T.; Gossett, J. M.; Zinder, S. H. Isolation of a bacterium that reductively dechlorinates tetrachloroethene to ethene. Nature 1997, 276, 1568−1571. (4) Suyama, A.; Yamashita, M.; Yoshino, S.; Furukawa, K. Molecular characterization of the PceA reductive dehalogenase of Desulfitobacterium sp. strain Y51. J. Bacteriol. 2002, 184 (13), 3419−3425. (5) Magnuson, J. K.; Stern, R. V.; Gossett, J. M.; Zinder, S. H.; Burris, D. R. Reductive dechlorination of tetrachloroethene to ethene by a two-component enzyme pathway. Appl. Environ. Microbial. 1998, 64 (4), 1270−1275. (6) Löffler, F. E.; Sun, Q.; Li, J.; Tiedje, J. M. 16S rRNA gene-based detection of tetrachloroethene-dechlorinating Desulf uromonas and Dehalococcoides species. Appl. Environ. Microbiol. 2000, 66 (4), 1369−1374. (7) Cupples, A. M. Real-time PCR quantification of Dehalococcoides population: methods and applications. J. Microbiol. Methods 2008, 72, 1−11. (8) He, J.; Sung, Y.; Krajmalnik-Brown, R.; Ritalahti, K. M.; Löffler, F. E. Isolation and characterization of Dehalococcoides sp. strain FL2, atrichloroethene (TCE)- and 1,2-dichloroethene-respiring anaerobe. Environ. Miclobiol. 2005, 7 (9), 1442−1450. (9) Radajewski, S.; Ineson, P.; Parekh, N. R.; Murrell, J. C. Stableisotope probing as a tool in microbial ecology. Nature 2000, 403, 646− 649. (10) Fennell, D. E. Comparison of alternative hydrogen donors for anaerobic reductive dechlorination of tetrachloroethene. Ph.D. Dissertation, Cornell University, Ithaca, NY, 1998. (11) Wilson, K. Preparation of genomic DNA from bacteria. In Current protocols in molecular biology. Supplement 9; Ausubel, F., Brent, R., Kingston, R., Moore, D., Seidman, J., Smith, J., Struhl, K., Eds.; John Wiley and Sons, Inc.: New York, 1990; pp 241−58.
Figure 7. Neighbor-joining phylogenetic tree derived from the 16S rRNA gene sequences of dehalorespiring bacteria and the Dehalobacter detected in the microcosm experiment. Bar = 2% nucleotide substitution rate (Knuc).
Fumarate was used as a labeled carbon source in this report. Although fumarate is the carbon source of D. ethenogenes 195 (Figure 2), it is uncertain whether it is utilized by other dehalorespiring bacteria. In the environmental samples, several varieties of dehalorespiring bacteria should exist; however, the labeled substrate utilized in the SIP-D method may affect the analytical results. In previous studies of dehalorespiring bacteria, acetic acid and CO2 was utilized as a carbon source for Dehalococcoides,27 and pyruvate was used for Desulf itobacte7715
dx.doi.org/10.1021/es301288y | Environ. Sci. Technol. 2012, 46, 7709−7716
Environmental Science & Technology
Article
of chlorinated dioxins by an anaerobic bacterium. Nature 2003, 421, 357−360.
(12) Neufeld, J. D.; Vohra, J.; Dumont, M. G.; Lueders, T.; Manefield, M.; Friedrich, M. W.; Murrell, J. C. DNA stable-isotope probing. Nature Protoc. 2007, 2 (4), 860−866. (13) Yoshida, N.; Takahashi, N.; Hiraishi, A. Phylogenetic characterization of a polychlorinated-dioxin-dechlorinating microbial community by use of microcosm studies. Appl. Environ. Microbiol. 2005, 71 (8), 4325−4334. (14) Zwart, G.; Huismans, R.; M. P., V.; Peer, Y. V. D.; Rijk, P. D.; Eenhoorn, H.; Muyzer, G.; Hannen, E. J. V.; Gons, H. J.; Laanbroek, H. J. Divergent members of the bacterial division Verrucomicrobiales in a temperate freshwater lake. FEMS Microbiol. Ecol. 1998, 25, 159−169. (15) Thompson, J. D.; Gibson, T. J.; Plewniak, F.; Jeanmougin, F.; Higgins, D. G. The CLUSTAL_X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res. 1997, 25 (24), 4876−4882. (16) Saito, N.; Nei, M. The Neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol. Biol. Evol. 1987, 4 (4), 406−425. (17) Holliger, C.; Schraa, G.; Stams, A. J. M.; Zehnder, A. J. B. A highly purified enrichment culture couples the reductive dechlorination of tetrachloroethene to growth. Appl. Environ. Microbiol. 1993, 59 (9), 2991−2997. (18) Maymo-Gatell, X.; Anguish, T.; Zinder, S. H. Reductive dechlorination of chlorinated ethenes and 1,2-dichloroethane by “dehalococcoides ethenogenes” 195. Appl. Environ. Microbiol. 1999, 65 (7), 3108−3113. (19) Adrian, L.; Szewzyk, U.; Wecke, J.; Görisch, H. Bacterial dehalorespiration with chrolinated benzenes. Nature 2000, 408, 580− 583. (20) He, J.; Ritlahti, K. M.; Yang, K.-L.; Koenigsberg, S. S.; Löffler, F. E. Detoxification of vinyl chloride to ethane coupled to growth of an anaerobic bacterium. Nature 2003, 424, 62−65. (21) Wild, A.; Hermann, R.; Leisinger, T. Isolation of an anaerobic bacterium which reductively dechlorinates tetrachloroethene and trichloroethene. Biodegradation 1996, 7, 507−511. (22) Grostern, A.; Edwards, E. A. 1, 1,1-Trichloroethane-degrading anaerobic mixed microbial culture enhances biotransformation of mixtures of chlorinated ethenes and ethanes. Appl. Environ. Microbiol. 2006, 72 (12), 7849−7856. (23) Holliger, C.; Hahn, D.; Harmsen, H.; Ludwig, W.; Schumacher, W.; Tindall, B.; Vazquez, F.; Weiss, N.; Zehnder, A. J. B. Dehalobacter restrictus gen. nov. and sp. nov., a strictly anaerobic bacterium that reductively dechlorinates tetra- and trichloroethene in an anaerobic respiration. Arch. Microbiol. 1998, 169, 313−321. (24) Suyama, A.; Iwakiri, R.; Kai, K.; Tokunaga, T.; Sera, N.; Furukawa, K. Isolation and characterization of Desulf itobacterium sp. strain Y51 capable of efficient dehalogenation of tetrachloroethene and polychloroethanes. Biosci. Biotechnol. Biochem. 2001, 65 (7), 1474− 1481. (25) Luijten, M. L. G. C.; de Weert., J.; Smidt, H.; Boschker, H. T. S.; de Vos, W. M.; Schraa, G.; Stams, A. J. M. Description of Sulfurospirillum halorespirans sp. nov., an anaerobic, tetrachloroeyhene-respiring bacterium, and transfer of Dehalospirillum multivorans to the geneus Sulfurospirillum as Sulf urospirillum multivorans comb. Nov. Int. J. Syst. Evol. Microbiol. 2003, 53, 787−793. (26) Hata, J.; Miyata, N.; Kim, E.-S.; Takamizawa, K.; Iwahori, K. Anaerobic degradation of cis-1,2-dichloroethylene and vinyl chloride by Clostridium sp. strain DC1 isolated from landfill leachate sediment. J. Biosci. Bioeng. 2004, 97 (3), 196−201. (27) Tang, Y. J.; Yi, S.; Zhuang, W.-Q.; Zinder, S. H.; Keasling, J. D.; Alvarez-Cohen, L. Investigation of carbon metabolism in “Dehalococcoides ethenogenes” strain 195 by use of isotopomer and transcriptomic analyses. J. Bacteriol. 2009, 191 (16), 5224−5231. (28) Yoshida, N.; Ye, L.; Baba, D.; Katayama, A. Reductive dechlorination of polychlorinjated biphenyls and dibenzo-p-dioxins in an enrichment culture containing Dehalobacter species. Microbes Environ. 2009, 24 (4), 343−346. (29) Bunge, M.; Adrian, L.; Kraus, A.; Opel, M.; Lorenz, W. G.; Andreesen, J. R.; Görisch, H.; Lechner, U. Reductive dehalogenation 7716
dx.doi.org/10.1021/es301288y | Environ. Sci. Technol. 2012, 46, 7709−7716
