Environ. Sci. Technol. 2010, 44, 9036–9041
A Quantitative PCR Assay for Aerobic, Vinyl Chloride- and Ethene-Assimilating Microorganisms in Groundwater YANG OH JIN AND TIMOTHY E. MATTES* Department of Civil and Environmental Engineering, 4105 Seamans Center, University of Iowa, Iowa City, Iowa 52242
Received July 2, 2010. Revised manuscript received September 28, 2010. Accepted October 12, 2010.
Vinyl chloride (VC) is a known human carcinogen that is primarily formed in groundwater via incomplete anaerobic dechlorination of chloroethenes. Aerobic, ethene-degrading bacteria (etheneotrophs), which are capable of both fortuitous and growth-linked VC oxidation, could be important in natural attenuation of VC plumes that escape anaerobic treatment. In this work, we developed a quantitative, real-time PCR (qPCR) assay for etheneotrophs in groundwater. We designed and tested degenerate qPCR primers for two functional genes involved in aerobic, growth-coupled VC- and etheneoxidation (etnC and etnE). Primer specificity to these target genes was tested by comparison to nucleotide sequence databases, PCR analysis of template DNA extracted from isolates and environmental samples, and sequencing of qPCR products obtained from VC-contaminated groundwater. The assay was made quantitative by constructing standard curves (threshold cycle vs log gene copy number) with DNA amplified from Mycobacterium strain JS60, an etheneotrophic isolate. Analysis of groundwater samples from three different VC-contaminated sites revealed that etnC abundance ranged from 1.6 × 103 - 1.0 × 105 copies/L groundwater while etnE abundance ranged from 4.3 × 103 - 6.3 × 105 copies/L groundwater. Our data suggest this novel environmental measurement method will be useful for supporting VC bioremediation strategies, assisting in site closure, and conducting microbial ecology studies involving etheneotrophs.
Introduction Bioremediation technologies are gaining acceptance as costeffective and environmentally friendly approaches for cleaning up both organic and inorganic groundwater contamination (1, 2). When employing a bioremediation strategy, it is important to demonstrate that pollutant removal is occurring as a result of microbial activity. For example, at a site undergoing bioremediation, it is common practice to document decreasing pollutant concentration along with the appearance of metabolites or daughter products and that other parameters (e.g., terminal electron acceptors, pH, ORP) are within the appropriate range for bioremediation to occur. Nevertheless, this solely geochemical approach is often insufficient to achieve regulatory approval for site closure. Additional lines of evidence are often needed to confirm * Corresponding author fax: (319) 335-5660; e-mail: tim-mattes@ uiowa.edu. 9036
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that the mediating microorganisms are present, abundant, and active at the site. An emerging means of providing these microbiological lines of evidence is through the application of molecular biology tools. The methodologies employed range from DNA microarrays (3) to proteomics (4-6), but quantitative realtime PCR (qPCR) methods are prevalent, likely because of their sensitivity and potential to yield rapid estimations of microbial abundance. For example, qPCR methods are now commonly used in the field of anaerobic chloroethene bioremediation to measure the abundance of 16S rRNA and key functional genes found in dechlorinating bacteria (e.g., tceA, bvcA, and vcrA) (7-9). An important metabolite produced during anaerobic reductive dechlorination of chloroethenes is vinyl chloride (VC) (10, 11). VC, a known human carcinogen (12), is problematic because it must be formed under anaerobic conditions in order to produce ethene, the desired end product. At some sites, the VC produced can escape anaerobic treatment and form mobile, dilute plumes that pose a threat to human and ecological health. Once a dilute VC plume migrates into groundwater containing oxygen, it is subject to several possible cometabolic and growth-coupled oxidative processes. In groundwater, VC can be oxidized cometabolically in the presence of ammonia, and a variety of aromatic, alkane, and alkene substrates (10). However, methane and ethene are likely to be present in many VC plumes as they are also produced during anaerobic reductive dechlorination of chloroethenes. This suggests that methanotrophs (13, 14) and etheneotrophs (15-18) could play major roles in oxidative VC degradation in groundwater. An absence of cometabolic substrates could also provide selective pressure for etheneotrophs to evolve into VC-assimilating bacteria (10, 16, 17, 19). An important feature of many etheneotrophs and VC-assimilators is their ability to degrade VC at very low oxygen levels (0.02 to 0.1 mg/L) (20, 21), suggesting that these bacteria could be active in groundwater zones considered anaerobic based on low field dissolved oxygen (DO) measurements. The fact that etheneotrophs and VC-assimilators could play a significant role in environmental VC degradation, even under very low oxygen tensions, suggests that rapid methods to quantify their abundance in groundwater (e.g., qPCR) would be valuable for ecological site assessment of VC bioremediation. The objective of this work was to develop and validate qPCR primers capable of quantifying etheneotrophs in groundwater; as such primers are not currently available. Etheneotrophs and VC-assimilators are phylogenetically diverse, likely because VC and ethene biodegradation genes are encoded on large plasmids (10, 22-24). Therefore, quantifying the 16S rRNA gene, a common approach to enumerating various bacterial groups in environmental samples (25, 26), is not appropriate for these bacteria. Instead, we targeted the functional genes etnC, which encodes the alpha subunit of alkene monooxygenase (AkMO), and etnE, which encodes the epoxyalkane coenzyme M transferase (EaCoMT) subunit, for development of qPCR primers. Because AkMO and EaCoMT catalyze the initial reactions in the VC and ethene biodegradation pathways in several isolates (10, 22, 27), it was our rationale that genes encoding them would be effective indicators of etheneotrophs and VC-assimilators in environmental samples. 10.1021/es102232m
2010 American Chemical Society
Published on Web 10/29/2010
TABLE 1. Bacterial Strains and Oligonucleotides Used in This Study bacterial strains
relevant characteristics
substrate
reference
Nocardioides sp. strain JS614 Mycobacterium sp. strain JS60 Mycobacterium sp. strain JS616 Mycobacterium sp. strain JS617 Mycobacterium sp. strain JS621 Mycobacterium sp. strain TM1 Mycobacterium sp. strain TM2 Mycobacterium sp. strain JS622 Mycobacterium sp. strain JS623 Mycobacterium sp. strain JS624 Mycobacterium sp. strain JS625 Xanthobacter autotrophicus Py2 E. coli strain DH5R F′Iq
ATCC BAA-499 ATCC BAA-494 ATCC BAA-496 ATCC BAA-497 ATCC BAA-498 not available in the ATCC not available in the ATCC not available in the ATCC not available in the ATCC not available in the ATCC not available in the ATCC ATCC BAA-1158 F′ proA+B+ lacIq ∆(lacZ)M15 zzf::Tn10 (TetR)/ fhuA2∆(argF-lacZ)U169 phoA glnV44 Ø80 ∆(lacZ)M15gyrA96 recA1 endA1 thi-1 hsdR17
VC VC VC VC VC VC VC ethene ethene ethene ethene propene Luria broth
21 21 21 21 21 21 21 24 24 24 24 24 New England BioLabs
oligonucleotides used to generate qPCR standard products
sequence (5′-3′)
JS60 EtnCF JS60 EtnCR JS614 EtnCF JS614 EtnCR JS623 EtnCF JS623 EtnCR CoM-F1L (etnE) CoM-R2E (etnE)
GATCCATTTCGTTCGATGCT GGCAATTCCTGCAACAAGAT GCGATGGAGAATGAGAAGGA TCCAGTCACAACCCTCACTG GAACTCAGACAGGGCTACGC TACTTCAGCGGGTCCTTCAC AACTACCCSAAYCCSCGCTGGTACGAC GTCGGCAGTTTCGGTGATCGTGCTCTTGAC
qPCR oligonucleotides
sequence (5′-3′)
RTC_F (etnC) RTC_R (etnC) RTE_F (etnE) RTE_R (etnE)
ACCCTGGTCGGTGTKSTYTC TCATGTAMGAGCCGACGAAGTC CAGAAYGGCTGYGACATYATCCA CSGGYGTRCCCGAGTAGTTWCC
Materials and Methods Chemicals, Bacterial Strains, And Growth Conditions. Ethene (99%) was from Airgas, propene (>99%) was from Aldrich, and all other chemicals were reagent grade or better. A total of eleven VC- and ethene-assimilating bacteria (Table 1) were grown on ethene in minimum salts medium (MSMethene) and harvested for DNA extraction as described in (16). Xanthobacter autotrophicus Py2 was grown on MSMpropene as described previously (28) and E. coli DH5R was grown in Luria-Bertani (LB) medium according to New England Biolabs protocols. Site Information and Groundwater Sample Collection. For this study, we selected three different VC-contaminated sites with geochemical conditions favorable for VC oxidation. At the Carver, MA site (4, 29), bioremediation of a dilute VC plume (∼2-29 µg/L) appears to be occurring in response to continuous oxygen injection into groundwater, although ethene amendments have also occurred (29). A portion of the dilute VC plume at NAS Oceana, VA SWMU 2C has been treated with an oxygen-releasing substrate to stimulate VC degradation, and VC concentrations in affected monitoring wells have also trended downward over time (30). At the Soldotna, AK site ongoing efforts to remediate a tetrachloroethene spill has generated a VC plume that exists near the adjacent Kenai River (31). Consistent decreases in VC concentrations in Kenai River sediments and pore water suggest that VC oxidation is occurring (31). Groundwater was collected according to USEPA/540/S95/504 procedures from two monitoring wells at each of the three sites investigated. Groundwater was passed through Sterivex-GP 0.22 µm membrane filter cartridges (Millipore) until the filter clogged (or a maximum of 3 L was filtered). The volume of filtered water was recorded and Sterivex filters were immediately placed on ice and shipped overnight to
product size (bp)
reference
1043
this study
1138
this study
1163
this study
891
22, 24
product size (bp)
reference
106
this study
151
this study
the University of Iowa, where they were stored at -20 °C prior to DNA extraction. Groundwater VC and ethene concentrations at the time of sampling in the selected monitoring wells are reported in Table 2. DNA Extraction from Pure Cultures and Sterivex Filters. DNA was extracted from pure bacterial cultures with a previously reported beadbeating method (16). Prior to DNA extraction, Sterivex filters were thawed and any remaining liquid in the filter housing was eliminated by syringe. Filter housings were opened with a tubing cutter and the filter membrane was aseptically removed and excised into 64 pieces. DNA was extracted from filter pieces with the MO BIO PowerSoil DNA extraction kit according to the manufacturer’s protocol with the following modifications: ∼300 µL (660 mg) of 0.1 mm zirconia/silica beads were added to the beadbeating tube prior to beadbeating with a Bio-Spec MiniBeadbeater-8 (2 min, high speed). Cell lysate (450 µL) was transferred into a clean microcentrifuge tube for the next step in the protocol. Finally, a second elution step (using 20 µL of elution buffer) was used to further increase DNA yield (by 19-20%). These modifications increased DNA yield by 43% in comparison to our previously reported beadbeating method (16). qPCR Primer Design and Analysis. Primer Express 2.0 (Applied Biosystems) was used to design qPCR primers. Default parameters were used, except that the amplicon length was allowed to vary between 100 and 150 bp. Nucleic acid and protein sequence alignments used in primer design were generated with ClustalX (32) and visualized in Bioedit (www.mbio.ncsu.edu/BioEdit/bioedit.html). Specificity of selected primer combinations with respect to the NCBI nonredundant nucleotide collection was tested with Primer BLAST (www.ncbi.nlm.nih.gov/tools/primer-blast). Default program parameters were used except that the misprimed VOL. 44, NO. 23, 2010 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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TABLE 2. Abundance of etnC and etnE in Groundwater Samples Collected from Monitoring Wells at Three VC-Contaminated Sitesa gene copies/L of groundwaterc site (sampling date) Carver, MA (09/29/2009) Soldotna, AK (05/06/2009) NAS Oceana, VA (08/05/2009)
b
b
well
VC (µg/L)
ethene (µg/L)
etnC
etnE
RB46D RB64I MW6 MW40 MW18 MW25
3.9 BDL 9.6 9.4 0.8 46
1.4 BDL 50 72 BDL BDL
7048 ( 2089 1614 ( 110 35 432 ( 1631 100 825 ( 3477 3783 ( 1715 1917 ( 456
121 451 ( 14 298 18 547 ( 2854 629 567 ( 28 857 161 305 ( 4167 4,342 ( 517 23 575 ( 931
a DNA from single Sterivex filter samples was extracted and analyzed from each well. The number of gene copies reported for each sample is the average of three analytical replicates ( the standard deviation. Additional detail concerning sampling, DNA extraction, and analysis of gene copy abundance is provided in Table S7 of the Supporting Information. b These values were obtained via personal communication from Sam Fogel, Bioremediation Consulting, Inc. (Carver), the May 2009 Groundwater Monitoring Report, River Terrace RV Park, Soldotna, AK, Alaska Department of Environmental Conservation, and the 2009 Long-term Monitoring Report for SMWUs 2B, 2C, and 2E, NAS Oceana, Virginia Beach, VA, CH2MHill. BDL: Below detection limit. c The number of gene copies per liter of groundwater was calculated via this formula:
Gene copies 70 µL DNA per filter reaction Gene copies ) (ng DNA per L of ground water) L of groundwater ng DNA per filter 2 µL DNA reaction
(
)(
product size deviation was reduced to 500 bp. The primer specificity stringency was set to 4 total mismatches, and 4 mismatches within the last 7 bps at the 3′ end. End Point PCR Procedures. PCR mixtures (25 µL) contained 12.5 µL of Qiagen HotStart PCR Master Mix, 2 µM of each primer, and 2 ng of template DNA. The thermocycling protocol was 95 °C for 5 min, then 35 cycles of 95 °C (20 s), 55 or 60 °C (20 s) and 72 °C (30 s), followed by a final extension cycle (72 °C, 4 min). PCR products were visualized by agarose gel electrophoresis, then purified with the Qiagen PCR Purification Kit. DNA concentrations were measured on a Qubit fluorometer (Invitrogen) using the Quant-iT dsDNA HS assays (Invitrogen). Selected PCR products were cloned and sequenced as described previously (19). Real-Time PCR Procedures. Real-time PCR was performed with an ABI 7000 Sequence Detection System (Applied Biosystems). Assays proceeded through 3 stages. Stage 1: 1 cycle of 95 °C for 10 min (DNA denaturing), stage 2: 40 cycles of 95 °C (15 s) and 60 °C for 60 s (denaturing, annealing, elongation, and data collection), and stage 3: a PCR product dissociation protocol where the temperature was ramped from 65-95 °C. PCR mixtures were prepared in 25 µL volumes using 12.5 µL of Power SYBR Green PCR Master Mix (Applied Biosystems), 750 nM of each primer (as determined in preliminary qPCR experiments using a dilution series of JS614 DNA template, data not shown), and 2 µL of DNA template (DNA concentration varied 2.0-38.4 ng/µL). To develop standard curves, we designed new primer sets to amplify the etnC from strains JS60 (JS60 EtnCF, JS60 EtnCR; 1043 bp product), JS614 (JS614 EtnCF, JS614 EtnCR; 1138 bp product), and JS623 (JS623 EtnCF, JS623 EtnCR; 1163 bp product) (Table 1). We used the CoMF1L and CoMR2E primer set to produce 891 bp etnE PCR products from JS60, JS614, and JS623 template DNA (Table 1). The number of gene copies per µL of PCR product was calculated by the following: bp *6.022 × 10 ( ng µL ) mole bp bp ng g PCR product size( *1 × 10 *660 gene copy ) g mole bp 23
DNA concentration
9
In some cases, 400 ng/µL Bovine Serum Albumin (BSA; New England Biolabs) was added to qPCR mixtures containing DNA template from environmental samples to alleviate PCR inhibition (33). ABI 7000 System SDS Software (Applied Biosystems) was used to analyze real-time PCR fluorescence data. The “auto 9038
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)(
)
baseline” function was used in all situations. However, the threshold fluorescence value (used for determining the threshold cycle Ct) for each set of standards and samples that were analyzed was optimized manually.
Results qPCR Primer Development and Specificity Testing. An effective primer set is essential for measuring specific gene abundance in environmental DNA. Both etnC (NVC105, NVC106) (34) and etnE (CoMF1L, CoMR2E) (24, 29) primer sets have been described, but the amplicons produced (360-891 bp) are outside the range typically used in qPCR (50-210 bp). Therefore, we developed new qPCR primers that generated products within the 50-210 bp range. Using complete etnC and etnE sequences from Mycobacterium strain JS60, Mycobacterium strain JS623, and Nocardioides strain JS614 (Table S1 of the Supporting Information, SI), we developed a pool of over 200 candidate etnC and etnE qPCR primer sets (hereafter referred to as RTC and RTE, respectively) in Primer Express 2.0 (Applied Biosystems). Concurrently, we generated amino acid and nucleic acid sequence alignments of all etnC and etnE sequences available in Genbank (26 and 15, respectively in June 2009; Table S1 of the SI) to detect conserved regions of each gene. Candidate qPCR primer sets were compared against these sequence alignments to determine the best matches with conserved regions while minimizing required primer degeneracy. This process yielded 8 RTC (expected amplicons of 87-151 bp) and 11 RTE (expected amplicons of 80-151 bp) qPCR primer sets for further analysis. The number of degeneracies ranged from 2-18 (RTC primers) and from 16-128 (RTE primers). The expected RTC and RTE amplicons are located within the regions amplified by previously described etnC primers (NVC105, NVC106 (34)) and etnE primers (CoMF1L, CoMR2E (24)), respectively. Ideally, qPCR primers intended for application to environmental samples should be highly specific while also capturing the sequence diversity present in the community. A series of initial PCR specificity tests (Table S2 of the SI) revealed one etnC primer set (RTC_F and RTC_R; 106 bp product) and one etnE primer set (RTE_F and RTE_R; 151 bp product) (Table 1; patent pending) that displayed the best specificities against DNA templates extracted from both pure cultures and environmental samples. We further tested the specificity of the selected etnC and etnE primer sets using melt-curve analysis, Primer BLAST analysis, and sequencing of qPCR products from clone libraries. Melt-curve analysis
of amplicons following qPCR of environmental DNA templates consistently showed peaks with very similar melting temperatures, suggesting that nonspecific amplicons and primer dimers were minimized (data not shown). Being degenerate, the RTC primers have 16 potential combinations, while the RTE primers have 128 potential combinations. Primer-BLAST indicated that none of the RTC primer combinations have unintended target sequences (data not shown), which suggests that these primers are highly specific to etnC sequences from known etheneotrophs and VC-assimilators. However, Primer-BLAST indicated three possible unintended RTE PCR products with expected sizes of 54 bp (Vitis vinifera, Genbank Acc. No. AM470838.2), 120 bp (Chromohalobacter salexigens peptidoglycan glycosyltransferase, Genbank Acc. No. CP000285.1), and 151 bp (Rhodococcus rhodochrous B-276 EaCoMT gene, Genbank Acc. No. AF426826.1). Generation of the 54 and 120 bp products is unlikely and can be excluded by melt-curve analysis (see above). However, as the etnE from propeneoxidizing Rhodococcus B-276 is closely related to the etnE from VC-assimilating Nocardioides JS614 (22), amplification of related sequences in groundwater DNA could occur. Finally, we prepared clone libraries of etnC and etnE qPCR products amplified from DNA extracted from Carver, MA, Soldotna, AK and NAS Oceana, VA groundwater. BLAST analysis of 35 cloned sequences (21 from the etnC library and 14 from the etnE library) indicated that they were all homologous to the target genes (Tables S3 and S4 of the SI). This provides further evidence that our qPCR primers selectively amplify target genes from groundwater community DNA. BLAST analysis also revealed that etnC and etnE sequence diversity, rather than a single sequence, was being captured from the samples, despite the short amplicon length. Additional information about PCR specificity testing is provided in the Supporting Information. qPCR Standard Curve Construction. With sufficient evidence that our qPCR primers were specific to the target genes, we developed standard curves (i.e., Ct vs log gene copy number) to achieve quantification of gene abundance. Either vectors containing the target DNA region or purified PCR products are typically used as standard qPCR templates. We elected to use PCR products as standard templates in this study as PCR efficiencies, linear response, and dynamic range were within acceptable limits. We also investigated which standard templates (i.e., those amplified from strain JS60, JS614, and JS623 genomic DNA) displayed the best combination of linear dynamic range, PCR amplification efficiency, and R2 values. This comparison, which is described in Table S5 of the SI, led us to use JS60 DNA as the standard template when quantifying etnC and etnE in environmental samples. Quantification of etnC and etnE in Groundwater Samples. Quantification of functional gene copy numbers in groundwater samples is subject to several sources of variability that will affect interpretation of gene abundance data. Among these sources of variability are the efficiency and reproducibility of DNA extraction and recovery from Sterivex filters. During the course of this work, we implemented several modifications to the filter DNA extraction protocol in an effort to improve DNA yields, but despite this we observed that when working with pure cultures, DNA yields varied from strain to strain. For example, DNA yields in quadruplicate extractions from Nocardioides sp. strain JS614 cells on Sterivex filters had a 12% coefficient of variation (CV), while the CV of DNA yields from Mycobacterium sp. strain JS60 and JS623 cells on Sterivex filters were 33 and 34%, respectively. When attempting to quantify gene copy numbers in DNA templates extracted from environmental samples, consideration must also be given to the presence and effect of PCR
inhibitors (35). The presence of PCR inhibitors is normally checked by testing a dilution series of DNA template. If evidence of PCR inhibition is found, then it is common to add BSA as a means of alleviating the effect. We performed qPCR with groundwater DNA extracts (diluted 5, 10, 20, and 40 times) from Carver, MA (RB46D and RB64I), Soldotna, AK (MW6 and MW40), and NAS Oceana, VA (MW18 and MW25) and determined the amplification efficiency for each sample. We also examined the effect that 400 ng/µL BSA had on amplification efficiencies. This procedure revealed that PCR inhibitors were present in nearly all of the DNA extracts we studied and that addition of BSA improved PCR efficiencies in all but one sample (MW40, Soldotna, AK) (Table S6 of the SI). The presence of 400 ng/µL BSA in qPCR standards (with JS60 template) did not adversely affect amplification efficiency when compared to JS60 qPCR standards without BSA. On the basis of these results, 400 ng/µL BSA was used in all subsequent qPCRs. Quantification of etnC and etnE was performed, in triplicate on any qPCR plate, using the groundwater DNA template extracted from filter samples taken from six different wells at three different sites (Table 2). The lowest Ct values in the no template controls (NTCs) were 35.7 (etnC) and 35.2 (etnE). In an attempt to minimize analytical variability, we only quantified gene copies from environmental samples against standards included on the same assay plate (i.e., intraassay standards) rather than from an external (interassay) standard curve as advised in a previous study (36). Despite this, we occasionally observed significant variability in triplicate qPCR assays on the same plate (up to 45% (etnC) and 15% (etnE)). Intra-assay standard PCR efficiencies were 105.5% when using etnC primers (-slope ) 3.21, R2 ) 0.999; Y-intercept ) 38.55, threshold ) 0.21) and 100.0% when using etnE primers (-slope ) 3.32, R2 ) 0.998; Y-intercept ) 38.53, threshold ) 0.24). Using these intra-assay standard curves, along with template DNA concentrations (ng/µL), groundwater sample volume (L), and mass (ng) of DNA per L of groundwater, we calculated etnC and etnE gene copies per liter of groundwater (Table 2). Because we used SYBR Green chemistry, dissociation curves were generated after qPCR to assess amplification quality. In general, dissociation curves of PCR products amplified from groundwater DNA templates displayed single peaks with melting temperatures near those observed in the standards. However, in contrast to the standards, additional minor peaks were often observed (data not shown). We also observed that when using RTC primers, the resulting dissociation curve peak intensities were lower than those with RTE. Despite these observations, the dissociation curves did not indicate any significant formation of primer dimers and nonspecific amplicons.
Discussion Recent studies suggest that etheneotrophs and VC-assimilators could play significant roles in natural attenuation of VC at contaminated sites (4, 20, 21, 29). Here, we describe a novel assay that allows cultivation-independent assessment of etheneotroph presence and abundance in environmental samples within 3 days (assuming sampling and shipping takes 2 days). This is an improvement, in terms of time, in comparison to previously described culture-based microcosm tests (∼60 days) (4, 29) and culture-based semiquantitative genetic assays (∼14 days) (29). In addition to the advantage of time, qPCR methods are quantitative and avoid enrichment bias (37, 38). However, there are other potential sources of bias and variability in during steps in the qPCR workflow (i.e., during sampling and subsequent sample handling (39), DNA extraction, and qPCR analysis) that must be considered. VOL. 44, NO. 23, 2010 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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The limited availability of environmental samples for this work prevented us from thoroughly addressing variability in sampling and DNA extraction that may have occurred with Sterivex filters. However, we observed variability in filter DNA extraction yields from pure etheneotrophic Mycobacterium cultures (33-34% CV). This suggests that the choice of DNA extraction kit could introduce significant variability in estimates of etheneotroph abundance, depending on the distribution of strains present in the community. We must caution that although Mycobacterium strains comprise the majority of etheneotrophs in pure culture there is yet no evidence that they actively degrade VC and ethene in groundwater. VC and ethene biodegradation genes are also known to be plasmid-encoded (22-24), therefore the diversity of etheneotrophs in the environment should extend well beyond Mycobacterium strains. The existence of plasmidborne etheneotrophic functional genes also suggests that etnC and etnE abundance will not equal etheneotroph cell abundance in environmental samples. Studies that aim to identify active ethene- and VC-assimilators in environmental samples via stable-isotope probing and quantify plasmid copy numbers in etheneotrophs are required to address these issues. Even though an absolute quantification approach is taken in this assay, significant loss of DNA template likely occurs during the cell lysis and DNA extraction steps. Assuming that the samples we studied are representative of the groundwater environment from which they were taken, the gene copy numbers reported here likely reflect a fraction of the genes actually present in any particular sample. Further refinement of the method by incorporating internal controls (39) could be used to address this issue. We developed degenerate primers and used SYBR Green PCR chemistry (as opposed to using TaqMan probes) in an effort to capture etnC and etnE diversity present in community DNA. All etnC and etnE sequences available as of June 2009 were utilized to develop these primers; however, the etnC and etnE sequence database remains relatively limited. Therefore, these primers could be biased against the true diversity of these sequences in nature, which could lead to inaccurate estimation of functional gene abundance. We should be able to evaluate this possibility as additional etnC and etnE sequence data becomes available and refine the primers as appropriate. Data in this report was presented in accordance to Minimum Information for the publication of Quantitative real-time PCR Experiments (MIQE) guidelines (40). A particularly relevant MIQE guideline involves testing for PCR inhibition. Co-extraction of substances from environmental samples that inhibit the PCR is yet another source of imprecision. The effect of PCR inhibitors in the samples we studied appeared to be significant. Alleviating PCR inhibition by template dilution, as indicated by apparent improvement in PCR amplification efficiency, was less effective than BSA addition. The MIQE guidelines also suggest providing substantial information concerning qPCR standard curves. When comparing standard curve parameters, we noticed that amplification efficiencies, R2 values, and linear dynamic ranges were similar, but that y-intercepts often varied (Table S5 of the SI). The variability in y-intercepts among different standard curves could be a result of primer bias among different DNA templates (including standards). A comparison of etnC and etnE abundance in samples with VC and ethene concentrations in groundwater at the time of sampling suggests that etnC and etnE abundance is elevated in samples where VC and ethene concentrations were also higher (Table 2). However, more qPCR data from environmental samples is required before firm conclusions can be drawn about the relationships between etnC and etnE abundance and VC and ethene concentrations. Measuring 9040
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both etnC and etnE abundance in environmental samples also allowed us to estimate the etnE/etnC ratio, which ranged from 1.1 to 17.8. This ratio is 2 in Nocardioides strain JS614 (22, 41). Although the etnE/etnC ratio has not yet been determined in other VC- and ethene-assimilating isolates, it seems unlikely that it would vary significantly from 1. Other possible explanations for high etnE/etnC ratios in environmental samples include primer bias and amplification of etnE sequences from propene-assimilating bacteria. For instance, a high etnE/etnC ratio could result if the RTC primers were biased against etnC sequences present in these samples. Conversely, amplification of etnE from propene-assimilating bacteria would lead to an overestimation of etnE abundance in comparison to etnC abundance. Primer-BLAST analysis of the RTE primers suggests that etnE sequences related to those found in propene-assimilating Rhodococcus strain B-276 could be amplified. This is not unexpected as the deduced EtnE in strain B-276 shares 83% amino acid identity to the deduced EtnE in VC-assimilating Nocardioides strain JS614 (22). Overall, the qPCR results presented here clearly indicate the potential for aerobic degradation of VC in and around monitoring wells at each of the three sites studied. This qPCR assay will facilitate detailed studies of microbial community functionality at VC-contaminated sites and help determine if the abundance of these functional genes is correlated with measured VC degradation rates. For example, an increase in etnC and/or etnE abundance in the same well over a relevant time series would suggest that natural attenuation of VC is occurring somewhere in the vicinity of that well’s zone of influence. Ideally, these approaches will be conducted with mRNA extracted from environmental samples to reveal gene expression levels and provide stronger evidence of metabolic functionality. In light of these possible applications, it is important to note that our qPCR method cannot distinguish between VCassimilators and etheneotrophs that cometabolize VC during growth on ethene. Point mutations in the EtnE gene have been observed in response to adaptation of an etheneotroph to VC as a growth substrate (19), however these mutations currently appear to be too subtle to exploit for diagnostic molecular biology tools. Despite this, we expect this qPCR method will be useful for assessing the possibility of aerobic VC biodegradation at contaminated sites. This assay could also be adapted for detailed ecological studies of etheneotrophs in a variety of groundwater systems and in environmental matrices other than groundwater (e.g., soils and sediments).
Acknowledgments We would like to thank Bill Richard (EST Associates, Inc.), Tim McDougall (OASIS Environmental), and Angela Petree (CH2M Hill) for directing groundwater sampling and Sam Fogel (Bioremediation Consulting, Inc.), Jim Begley (MT Environmental Restoration), Laura Cook (CH2M Hill) and Sharon Richmond (Alaska DEC) for sharing geochemical data and coordinating sampling efforts. We also thank Michelle Stolzoff and Stefanie Schmidt for technical assistance. This research was supported by the Strategic Environmental Research and Development Program (SERDP) under project ER-1683.
Supporting Information Available Seven tables with detailed information on qPCR primer specificity testing, standard curve parameters, the effect of BSA on amplification efficiency, and calculation of gene abundance in environmental samples are provided. This information is available free of charge via the Internet at http://pubs.acs.org/.
Literature Cited (1) Freedman, D. L.; Gossett, J. M. Biological reductive dechlorination of tetrachloroethylene and trichloroethylene to ethylene under methanogenic conditions. Appl. Environ. Microbiol. 1989, 55 (9), 2144–2151. (2) Lovley, D. R.; Holmes, D. E.; Nevin, K. P. Dissimilatory Fe(III) and Mn(IV) reduction. Adv. Microb. Physiol. 2004, 49, 219–286. (3) Dennis, P.; Edwards, E. A.; Liss, S. N.; Fulthorpe, R. Monitoring gene expression in mixed microbial communities by using DNA microarrays. Appl. Environ. Microbiol. 2003, 69 (2), 769–778. (4) Chuang, A. S.; Jin, Y. O.; Schmidt, L. S.; Li, Y.; Fogel, S.; Smoler, D.; Mattes, T. E. Proteomic analysis of ethene-enriched groundwater microcosms from a vinyl chloride-contaminated site. Environ. Sci. Technol. 2010, 44 (5), 1594–1601. (5) Morris, R. M.; Fung, J. M.; Rahm, B. G.; Zhang, S.; Freedman, D. L.; Zinder, S. H.; Richardson, R. E. Comparative proteomics of Dehalococcoides spp. reveals strain-specific peptides associated with activity. Appl. Environ. Microbiol. 2007, 73 (1), 320– 326. (6) Morris, R. M.; Sowell, S.; Barofsky, D.; Zinder, S.; Richardson, R. Transcription and mass-spectroscopic proteomic studies of electron transport oxidoreductases in Dehalococcoides ethenogenes. Environ. Microbiol. 2006, 8 (9), 1499–1509. (7) Cupples, A. M. Real-time PCR quantification of Dehalococcoides populations: Methods and applications. J. Microbiol. Methods 2008, 72 (1), 1–11. (8) Johnson, D. R.; Lee, P. K.; Holmes, V. F.; Alvarez-Cohen, L. An internal reference technique for accurately quantifying specific mRNAs by real-time PCR with application to the tceA reductive dehalogenase gene. Appl. Environ. Microbiol. 2005, 71 (7), 3866– 3871. (9) Ritalahti, K. M.; Amos, B. K.; Sung, Y.; Wu, Q.; Koenigsberg, S. S.; Loffler, F. E. Quantitative PCR targeting 16S rRNA and reductive dehalogenase genes simultaneously monitors multiple Dehalococcoides strains. Appl. Environ. Microbiol. 2006, 72 (4), 2765– 2774. (10) Mattes, T. E.; Alexander, A. K.; Coleman, N. V. Aerobic biodegradation of the chloroethenes: pathways, enzymes, ecology, and evolution. FEMS Microbiol. Rev. 2010, 34 (4), 435– 475. (11) Bradley, P. M. History and ecology of chloroethene biodegradation: a review. Bioremed. J. 2003, 7 (2), 81–109. (12) Bucher, J. R.; Cooper, G.; Haseman, J. K.; Jameson, C. W.; Longnecker, M.; Kamel, F.; Maronpot, R.; Matthews, H. B.; Melnick, R.; Newbold, R.; Tennant, R. W.; Thompson, C.; Waalkes, M., Report on Carcinogens, 11th ed.; U.S. Department of Health and Human Services, Public Health Service, National Toxicology Program: 2005. (13) Fogel, M. M.; Taddeo, A. R.; Fogel, S. Biodegradation of chlorinated ethenes by a methane-utilizing mixed culture. Appl. Environ. Microbiol. 1986, 51, 720–724. (14) van Hylckama Vlieg, J. E. T.; de Koning, W.; Janssen, D. B. Transformation kinetics of chlorinated ethenes by Methylosinus trichosporium OB3b and detection of unstable epoxides by online gas chromatography. Appl. Environ. Microbiol. 1996, 62, 3304–3312. (15) Freedman, D. L.; Herz, S. D. Use of ethylene and ethane as primary substrates for aerobic cometabolism of vinyl chloride. Water Env. Res. 1996, 68, 320–328. (16) Jin, Y. O.; Mattes, T. E. Adaptation of aerobic, ethene-assimilating Mycobacterium strains to vinyl chloride as a growth substrate. Environ. Sci. Technol. 2008, 42 (13), 4784–4789. (17) Verce, M. F.; Ulrich, R. L.; Freedman, D. L. Transition from cometabolic to growth-linked biodegradation of vinyl chloride by a Pseudomonas sp. isolated on ethene. Environ. Sci. Technol. 2001, 35 (21), 4242–4251. (18) Koziollek, P.; Bryniok, D.; Knackmuss, H.-J. Ethene as an auxiliary substrate for the cooxidation of cis-dichloroethene and vinyl chloride. Arch. Microbiol. 1999, 172, 240–246. (19) Jin, Y. O.; Cheung, S.; Coleman, N. V.; Mattes, T. E. Association of missense mutations in epoxyalkane coenzyme M transferase with adaptation of Mycobacterium sp. strain JS623 to growth on vinyl chloride. Appl. Environ. Microbiol. 2010, 76 (11), 34133419. (20) Gossett, J. M. Sustained aerobic oxidation of vinyl chloride at low oxygen concentrations. Environ. Sci. Technol. 2010, 44 (4), 1405–1411. (21) Coleman, N. V.; Mattes, T. E.; Gossett, J. M.; Spain, J. C. Phylogenetic and kinetic diversity of aerobic vinyl chlorideassimilating bacteria from contaminated sites. Appl. Environ. Microbiol. 2002, 68 (12), 6162–6171.
(22) Mattes, T. E.; Coleman, N. V.; Gossett, J. M.; Spain, J. C. Physiological and molecular genetic analyses of vinyl chloride and ethene biodegradation in Nocardioides sp. strain JS614. Arch. Microbiol. 2005, 183, 95–106. (23) Danko, A. S.; Luo, M. Z.; Bagwell, C. E.; Brigmon, R. L.; Freedman, D. L. Involvement of linear plasmids in aerobic biodegradation of vinyl chloride. Appl. Environ. Microbiol. 2004, 70 (10), 6092– 6097. (24) Coleman, N. V.; Spain, J. C. Distribution of the coenzyme M pathway of epoxide metabolism among ethene- and vinyl chloride-degrading Mycobacterium strains. Appl. Environ. Microbiol. 2003, 69 (10), 6041–6046. (25) Hristova, K. R.; Lutenegger, C. M.; Scow, K. M. Detection and quantification of methyl tert-butyl ether-degrading strain PM1 by real-time TaqMan PCR. Appl. Environ. Microbiol. 2001, 67 (11), 5154–5160. (26) Bedard, D. L.; Ritalahti, K. M.; Loffler, F. E. The Dehalococcoides population in sediment-free mixed cultures metabolically dechlorinates the commercial polychlorinated biphenyl mixture aroclor 1260. Appl. Environ. Microbiol. 2007, 73 (8), 2513–21. (27) Coleman, N. V.; Spain, J. C. Epoxyalkane:Coenzyme M transferase in the ethene and vinyl chloride biodegradation pathways of Mycobacterium strain JS60. J. Bacteriol. 2003, 185 (18), 5536– 5545. (28) Owens, C. R.; Karceski, J. K.; Mattes, T. E. Gaseous alkene biotransformation and enantioselective epoxyalkane formation by Nocardioides sp. strain JS614. Appl. Microbiol. Biotechnol. 2009, 84 (4), 685–692. (29) Begley, J. F.; Hansen, E.; Wells, A. K.; Fogel, S.; Begley, G. S. Assessment and monitoring tools for aerobic bioremediation of vinyl chloride in groundwater. Remed. J. 2009, 20 (1), 107– 117. (30) Cook, L. J.; Hickman, G.; Chang, A.; Landin, P.; Reisch, T., Comparison of aerobic and anaerobic biotreatments of lowlevel vinyl chloride. In Remediation of Chlorinated and Recalcitrant Compounds; Battelle: Monterey, CA, 2006. (31) Contaminated Sites Program, River Terrace RV Park, Second Five Year Review of the Record of Decision, August 4, 2010 Alaska Department of Environmental Conservation http:// www.dec.state.ak.us/spar/csp/docs/rivterr/DOC009.pdf (Accessed September 21, 2010), (32) Chenna, R.; Sugawara, H.; Koike, T.; Lopez, R.; Gibson, T. J.; Higgins, D. G.; Thompson, J. D. Multiple sequence alignment with the Clustal series of programs. Nucleic Acids Res. 2003, 31 (13), 3497–3500. (33) Kreader, C. A. Relief of amplification inhibition in PCR with bovine serum albumin or T4 gene 32 protein. Appl. Environ. Microbiol. 1996, 62 (3), 1102–1106. (34) Coleman, N. V.; Bui, N. B.; Holmes, A. J. Soluble di-iron monooxygenase gene diversity in soils, sediments and ethene enrichments. Environ. Microbiol. 2006, 8 (7), 1228–1239. (35) Wilson, I. G. Inhibition and facilitation of nucleic acid amplification. Appl. Environ. Microbiol. 1997, 63 (10), 3741–3751. (36) Smith, C. J.; Nedwell, D. B.; Dong, L. F.; Osborn, A. M. Evaluation of quantitative polymerase chain reaction-based approaches for determining gene copy and gene transcript numbers in environmental samples. Environ. Microbiol. 2006, 8 (5), 804– 815. (37) Muniesa, M.; Blanch, A. R.; Lucena, F.; Jofre, J. Bacteriophages may bias outcome of bacterial enrichment cultures. Appl. Environ. Microbiol. 2005, 71 (8), 4269–4275. (38) Dunbar, J.; White, S.; Forney, L. Genetic diversity through the looking glass: Effect of enrichment bias. Appl. Environ. Microbiol. 1997, 63 (4), 1326–1331. (39) Ritalahti, K. M.; Hatt, J. K.; Petrovskis, E.; Lo¨ffler, F. E., Groundwater Sampling for Nucleic Acid Biomarker Analysis. In Handbook of Hydrocarbon and Lipid Microbiology; Timmis, K. N., Ed.; Springer: Berlin, Heidelberg: 2010; pp 3407-3418. (40) Bustin, S. A.; Benes, V.; Garson, J. A.; Hellemans, J.; Huggett, J.; Kubista, M.; Mueller, R.; Nolan, T.; Pfaffl, M. W.; Shipley, G. L.; Vandesompele, J.; Wittwer, C. T. The MIQE guidelines: minimum information for publication of quantitative real-time PCR experiments. Clin. Chem. 2009, 55 (4), 611–622. (41) Chuang, A. S.; Mattes, T. E. Identification of polypeptides expressed in response to vinyl chloride, ethene, and epoxyethane in Nocardioides sp. strain JS614 by using peptide mass fingerprinting. Appl. Environ. Microbiol. 2007, 73 (13), 4368–4372.
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