2009 Lam

Letters in Applied Microbiology ISSN 0266-8254

ORIGINAL ARTICLE

Enhanced biotransformation of TCE using plant terpenoids in contaminated groundwater J.R.-M. Brown1,2, I.P. Thompson3, G.I. Paton2 and A.C. Singer4 1 2 3 4

Centre for Ecology and Hydrology, Oxford, UK School of Biological Sciences, University of Aberdeen, Aberdeen, UK Department of Engineering Science, University of Oxford, Kidlington, Oxfordshire, UK Centre for Ecology and Hydrology, Wallingford, UK

Keywords biostimulation, carvone, cumene, groundwater, plant terpenes, secondary plant metabolites, TCE. Correspondence Andrew C. Singer, Centre for Ecology and Hydrology, Maclean Building, Benson Lane, Crowmarsh Gifford, Wallingford OX10 8BB, UK. E-mail: [email protected]

2009 ⁄ 1000: received 5 June 2009, revised and accepted 9 September 2009 doi:10.1111/j.1472-765X.2009.02738.x

Abstract Aims: To examine plant terpenoids as inducers of TCE (trichloroethylene) biotransformation by an indigenous microbial community originating from a plume of TCE-contaminated groundwater. Methods and Results: One-litre microcosms of groundwater were spiked with 100 lmol 1)1 of TCE and amended weekly for 16 weeks with 20 ll 1)1 of the following plant monoterpenes: linalool, pulegone, R-(+) carvone, S-()) carvone, farnesol, cumene. Yeast extract-amended and unamended control treatments were also prepared. The addition of R-carvone and S-carvone, linalool and cumene resulted in the biotransformation of upwards of 88% of the TCE, significantly more than the unamendment control (61%). The aforementioned group of terpenes also significantly (P < 0Æ05) allowed more TCE to be degraded than the remaining two terpenes (farnesol and pulegone), and the yeast extract treatment which biotransformed 74–75% of the TCE. The microbial community profile was monitored by denaturing gradient gel electrophoresis and demonstrated much greater similarities between the microbial communities in terpene-amended treatments than in the yeast extract or unamended controls. Conclusions: TCE biotransformation can be significantly enhanced through the addition of selected plant terpenoids. Significance and Impact of the Study: Plant terpenoid and nutrient supplementation to groundwater might provide an environmentally benign means of enhancing the rate of in situ TCE bioremediation.

Introduction Accidental and historical release of chlorinated compounds such as TCE has resulted in extensive contamination of the subsurface environment (Major et al. 1991; Shapiro et al. 2004). A combination of TCE’s physical and chemical properties (high solubility and miscibility) (Moran 2007), lends it to the formation of a dense, nonaqueous phase in the subsurface (Lee 2007), resisting biotransformation (Vogel et al. 1987) and dispersing over wide areas. The pump and treat approach is one of the primary mechanisms of TCE-contaminated groundwater remediation. In recent years, bioremediation and biostimulation

have been pursued as a means for remediating TCE, as pump and treat is less cost-effective when applied to lowlevel TCE-contaminated sites (Aulenta et al. 2005; Da Silva et al. 2006). TCE biostimulation has been demonstrated using growth substrates and cometabolites such as toluene (Hopkins and McCarty 1995), phenol (Hopkins et al. 1993) and methanol (DiSpirito et al. 1991; Kang et al. 2001). However, there are concerns about the suitability of using highly toxic chemicals to induce TCE degradation within the environment (http://www.epa.gov/ OGWDW/contaminants/basicinformation/toluene.html, US EPA 2000; Brautbar and Williams Ii 2002). Consequently,

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it would be beneficial to seek alternative substrates that would stimulate TCE-degradation without causing additional environmental concern. A novel avenue for selecting suitable cometabolites is the use of plant secondary metabolites. Plant secondary metabolites are low molecular weight plant products (Hadacek 2002) that are traditionally considered to be nonessential for the basic metabolic processes of the plant (Dixon 2001). Terpenes are the main components of essential oils of plants, such as limonene from lemon oil, carvone from spearmint oil and pinene from pine oil. A wide range of terpenes have already been reported to stimulate microbial degradation of xenobiotic compounds, such as polychlorinated biphenyls (PCB) (Gilbert and Crowley 1997; Singer et al. 2000, 2003; Crowley et al. 2001; Oh et al. 2003). The monoterpene cumene has previously been demonstrated to stimulate TCE degradation in pure culture (Dabrock et al. 1992; Suttinun et al. 2004) and in soil (Suttinun et al. 2004). Hence, we hypothesized that an even wider range of plant secondary metabolites should be applicable to enhancing TCE biodegradation owing to many structural similarities within the terpenoids. In this study, we constructed one-litre microcosms to compare TCE biotransformation by the indigenous microbial community from a TCE-contaminated groundwater plume. Groundwater amended with TCE (100 lmol l)1) was spiked with one of six plant secondary metabolites: 1, 6-octadien-3-ol (linalool), R-(+)p-menth-4 (8)-en-3-one (pulegone), 2-cyclohexen-1-one [R-(+) carvone], 2-cyclohexen-1-one [S-()) carvone], 2,6,10-dodecatrien-1-ol, 3,7,11-trimethyl (farnesol) and isopropylbenzene (cumene), with one treatment receiving only yeast extract and the control remaining unamended. TCE biotransformation was confirmed by headspace analysis-gas chromatography. The microbial communities were monitored by denaturing gradient gel electrophoresis (DGGE; community structure). Materials and methods Materials TCE (99% purity) and S-()) carvone (98% purity), cumene (99% purity), linalool (98% purity) were obtained from Sigma Aldrich (York, UK), R-(+) carvone (98% purity) from Lancaster (Morecambe, UK) and both pulegone (92% purity) and farnesol (96% purity) from Arcos Organics (Leicestershire, UK). Field site Samples were collected in 1000-ml PTFE-lined screwcapped glass bottles with no headspace from boreholes of 770

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a TCE-contaminated site located in southern England. The site covers an area of c. 3 km2 and is characterized by a shallow chalk aquifer, protected by London clay overlain by Silchester Gravels of Eocene age. The groundwater contains a plume of TCE at a concentration between 5 and 620 lmol l)1. A total of 27 boreholes representative of the plume were sampled against a hydraulic gradient within an area of 25 000 m2. The boreholes used in this study contained TCE at a moderate to high level (150–400 lmol l)1). Groundwater samples located outside of the plume served as a control. Microcosm preparation Microcosms were prepared in the 1000-ml sampling bottle by sparging with nitrogen to remove the ‘native’ TCE and any additional co-contaminating volatile organics. All microcosms were spiked with an initial concentration of 100 lmol l)1 of TCE. Each bottle was amended with 20 ll l)1 of substrate: linalool, pulegone, cumene, farnesol, R-(+) carvone, and S-()) carvone. The concentration of 20 ll l)1 was chosen as it was below the concentration of the least soluble of the terpenoids; hence, it was assumed that all microcosms had equivalent exposure to each of the treatment compounds. Control microcosms were set up with no substrate and a sterilize blank which was autoclaved at 121C, 15 psi, for 15 min. A microcosm was also set up with 20 mg l)1 of yeast extract as a terpene-free carbon source control. The nine microcosm treatments were set up in triplicate. Microcosms were kept at 22C in the dark with weekly addition of the respective substrate (20 ll l)1). A small headspace volume (c. 10 ml) was maintained in the bottles to minimize partitioning of TCE between the liquid and gas phase. Hence, microaerophilic and anaerobic conditions likely persisted in the microcosms during the majority of the study except directly following the respiking of the mesocosms each week, at which point each microcosm was vigorously shaken providing limited aeration. Samples were withdrawn using a glass pipette directly after the bottles had been shaken and placed directly into a 20-ml borosilicate gas chromatograph-headspace vial and fitted with a PTFE-lined septum crimp cap. Analytical methods Samples (5 ml) were amended with 50 ll of 5 mol l)1 NaOH to terminate biological activity and 2-ll dibromobenzene as an internal standard. The vial was placed in a headspace sampler (Agilent 7964; Agilent Technologies, Berkshire, UK) in shake mode, with an oven temperature of 70C, loop temperature of 110C and a transfer line temperature of 90C to the gas chromatograph inlet. A

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Molecular biological investigations

TCE concentration mmol l–1

Total nucleic acids from the indigenous microbial community were extracted from 50 ml of groundwater samples by a combination of bead beating, enzymatic lysis and solvent extraction for DGGE analysis, as previously described (Griffiths et al. 2000). A 200-bp product spanning the V3 region of the 16S rDNA was amplified by PCR as previously described (Muyzer et al. 1993), in a total volume of 50 ll containing universal 16S primers 530r (5¢-GTA TTA CCG CGG CTG CTG-3¢) and the GC clamped 338F (5¢-CGC CCG CCG CGC CCC CGC CCC GGC CCG CCG CCC CCG CCC ACT CCT ACG GGA GGC AGC-3¢). Approximately 1 ll of template was amplified with 1Æ5 ll of Taq polymerase (Sigma Chemicals, Dorset, UK), 0Æ5 ll of each primer, 0Æ5 ll of 100 mmol l)1 deoxynucleoside triphosphate on an MJResearch PTC-225 (Peltier Thermal Cycler; MJ Research Instruments, Watertown, MA, USA). The temperature

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programme was as follows: 95C denaturation for 2 min followed by 35 cycles at 95C for 60 s, annealing at 60C for 60 s, 72C extension for 90 s and a single step of 72C for 30 min. DGGE was performed with the Bio-Rad D Code system (Bio-Rad, Hercules, CA, USA). The PCR product was loaded onto a 10% (w ⁄ v) acrylamide gel containing a linear denaturant gradient of 30–60% [100% denaturant consisted of 7 mol l)1 urea and 40% (v ⁄ v) formamide parallel to the direction of electrophoresis]. Gels were run in a 0Æ5· TAE buffer at 60C and 100 V for 16 h. Gels were stained with 0Æ5· TAE buffer containing SyberGold (Molecular Probes, Paisley, UK) and photographed using the VersaDOC Imaging System and the Quantity One software (Bio-Rad). Band and profile analysis were determined using Phoretix 1D analysis software (Phoretix International, Newcastle upon Tyne, UK). Statistical analysis One-way anova with Tukey HSD (honestly significant difference) was performed by using spss 16.0 (SPSS Inc., Chicago, IL). The correlation coefficient r is only stated if their significance level was at least 95% (P < 0Æ05). For the analysis of the relationship between DGGE profiles agglomerative hierarchical clustering was performed by the unweighted-pair group method using arithmetic averages (UPGMA) with squared Euclidean distances which was displayed as a dendrogram. Results Biodegradation experiments were carried out with groundwater samples from a TCE-contaminated aquifer with the substrates (i.e., terpenes and yeast extract). Figure 1 illustrates the biotransformation of TCE in the presence of individual terpenes, yeast extract and the unamended control. Biotransformation was calculated as the per cent loss of TCE when compared to the autoclaved control treatment, while the rate of TCE biotransformation

TCE concentration mmol l–1

75-m · 0Æ53-mm DB-624 capillary column with a 3-lm film (Agilent Technologies) was used to separate TCE. The temperature programme of the Agilent 6890 gas chromatograph (Agilent Technologies) used was as follows: 50C hold for 4 min, 40–280C for 5 min, hold for 6 min at 300C. The electron capture detector (ECD) operated at a temperature of 300C; helium was the carrier gas with a flow rate of 1Æ0 ml min)1. The makeup gas was nitrogen at 60 ml min)1. The data were recorded using the ChemStation Software Rev A.10.02 (Agilent Technologies). Calibration curves were prepared from serial dilution of TCE in methanol. Samples (1 ml) were removed from each microcosm weekly for 16 weeks and analysed for chloride (Chen et al. 2005) using ion chromatography on a Dionex AS50 (Dionex Corporation, Leeds, UK), employing an isocratic pump, an Ionpac ASH-HC column autosampler and electrochemical detector. Results were corrected for background chloride.

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Figure 1 TCE concentration (lmol l)1) in microcosms during the experiment with different terpenoid (left panel) and control (right panel) treatments: (d) linalool; (s) farnesol; (.) pulegone; (4) R-carvone; ( ) S-carvone; (h) cumene; (r) sterilized control; ()) TCE control and ( ) yeast extract. ª 2009 The Authors Journal compilation ª 2009 The Society for Applied Microbiology, Letters in Applied Microbiology 49 (2009) 769–774

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was the difference in TCE removed over the 16-week period (112 days). R-(+) carvone-amended, S-()) carvoneamended, cumene-amended, and linalool-amended groundwater resulted in significantly greater TCE biotransformation after 16 weeks, 82–88% (4–7 nmol TCE ⁄ day; P < 0Æ05), than the treatments with pulegone, farnesol and yeast extract (74–75%; 9–10 nmol TCE ⁄ day) as well as the unamended control (61%; 15 nmol TCE ⁄ day; P < 0Æ001). Chloride originating from dechlorination of TCE increased sharply at week 5 in the unamended control and in the yeast extract-amended treatment, whereas all the other treatments showed a sharp increase in chloride at week 7 (Fig. 2). R-(+) carvone accumulated the highest amount of chloride by week 9 (181 lmol l)1) followed by S-()) carvone and cumene (133 and 122 lmol l)1), respectively. The sterilized control flask showed no degradation of TCE and no chloride accumulation. The DGGE profile of the groundwater community between treatments was expectedly similar at the start of the experiment, but a decline in the number of operational taxonomic units (OTUs) was observed from week 1 through 13 (data not shown). The DGGE demonstrated two major clusters of microbial community fingerprints: (i) the unamended TCE treatment and the yeast extractamended treatment, and (ii) all the terpene-amended treatments (Fig. 3). Within the terpene cluster, there were three subclusters (% TCE biotransfomation): (i) farnesol (74%) and R-(+) carvone (88%); (ii) pulegone (75%), S-()) carvone (86%) and cumene (87%); and (iii) linalool (82%). Notably, the two isomers S-carvone and R-carvone did not cluster together.

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Figure 2 Chloride concentration (lmol l)1) in microcosms during the experiment with different treatments: (d) linalool; (s) farnesol; (.) pulegone; (4) R-carvone; ( ) S-carvone; (h) cumene; (r) sterilized control; ()) TCE control and ( ) yeast extract.

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Figure 3 Unweighted-pair group method using arithmetic averages dendrogram showing the similarity of denaturing gradient gel electrophoresis patterns of 16S rDNA extracted from the different microcosms treatments.

Discussion Terpene addition significantly increase the TCE biotransformation compared to the unamended control. The treatments containing R-(+) carvone, S-(+) carvone, cumene and linalool significantly stimulated the native community to biotransform 82–88% of the TCE present in the respective microcosm treatment compared to the unamended treatment. Dabrock et al. (1992) observed 100% and 71% degradation of TCE with Pseudomonas sp. JR1 and Rhodococcus erythropolis BD1 (isolated from groundwater and TCE ⁄ toluene contaminated soil), using cumene as a growth substrate (Dabrock et al. 1992), a result which is consistent with the findings in this study where 87% of the TCE was biotransformed in the presence of cumene. Other terpenes such as p-cymene, geraniol, citronellol, anise oil, cuminic acid and cinnamyl alcohol were also used by Dabrock et al. (1992), but <10% of TCE degradation was observed. Suttinun et al. (2004) investigated the effect of terpenes such as cumene, limonene, S-()) carvone and pinene at different concentrations on the rates of TCE (10 ppm) degradation in vitro by pure Rhodococcus gordoniae cells. The authors found that all of the terpenes induced greater biotransformation of TCE than the unamended control, with the highest percent biotransformation stemming from the cumene treatment, which achieved 72% TCE removal (Suttinun et al. 2004). The authors also reported significant changes in the amount of TCE removed depending on the concentration of terpene employed; pinene was only effective in the range of 5–10 mg l)1, carvone at 5–25 mg l)1 and cumene at 10–50 mg l)1, while limonene

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yielded significantly greater removal of TCE at all the tested concentrations 5–50 mg l)1. The results from this study are consistent with that of Suttinun et al. (2004) and further extend the spectrum of terpenoids capable of stimulating TCE biotransformation. Chloride accumulation was used as an indication of TCE dechlorination and biotransformation. In comparison to the untreated and yeast extract-amended microcosm, there was a delay in chloride production by 2 weeks in the terpene-amended treatments (Fig. 2). Although the data was not statistically significant, cell counts by flow cytometry suggest that cell growth was greater in terpene-amended microcosms when compared to the control (data not shown). Hence, the delay in TCE dechlorination in terpene-amended treatments might be indicative of competitive exclusion of the TCE for the more energy and carbon-rich terpene substrate. Evident from the DGGE community fingerprints, terpenes enriched for a microbial community distinguishable from the terpene-free treatments. Despite slight shifts in the microbial community within the terpene-amended community fingerprints, this did not appear to correspond to any functional differences with regard to TCE biotransformation. The readily distinguishable microbial communities generated from R-(+) carvone and S-()) carvone addition yielded similarly functioning communities with regard to TCE biotransformation. These results are consistent with a previous report that demonstrated the capacity for river water to evolve into both functionally similar and distinguishable microbial communities after exposure to the terpene isomers R-carvone and S-carvone (Lehmann et al. 2008). In summary, the results from this study provide evidence that secondary plant metabolites such as R-(+) carvone, S-()) carvone, linalool, pulegone and farnesol are readily utilized by groundwater bacteria and can be used as inducers to stimulate TCE biotransformation at a rate in excess of unamended controls. Terpenes are nontoxic, natural alternative substrates and as such can be regarded as benign if released into the environment and are often very effective at low concentrations (Singer et al. 2003). To our knowledge, this is the first study to use pulegone, linalool and farnesol to stimulate degradation of TCE. Acknowledgements The authors would like to thank the EPSRC and the AWE plc for their financial support, Sean Amos of AWE plc for providing groundwater samples, Ken Killham and Graeme I. Paton for their support, Andrew Whiteley for his help with flow cytometry and Hong Li for his help with chloride analysis.

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