Ecotoxicology DOI 10.1007/s10646-014-1205-y

Survival of prokaryotes in a polluted waste dump during remediation by alkaline hydrolysis Marie Bank Nielsen • Kasper Urup Kjeldsen Mark Alexander Lever • Kjeld Ingvorsen

•

Accepted: 27 January 2014 Ó Springer Science+Business Media New York 2014

Abstract A combination of culture-dependent and culture-independent techniques was used to characterize bacterial and archaeal communities in a highly polluted waste dump and to assess the effect of remediation by alkaline hydrolysis on these communities. This waste dump (Breakwater 42), located in Denmark, contains approximately 100 different toxic compounds including large amounts of organophosphorous pesticides such as parathions. The alkaline hydrolysis (12 months at pH [12) decimated bacterial and archaeal abundances, as estimated by 16S rRNA gene–based qPCR, from 2.1 9 104 and 2.9 9 103 gene copies per gram wet soil respectively to below the detection limit of the qPCR assay. Clone libraries constructed from PCR-amplified 16S rRNA gene fragments showed a significant reduction in bacterial diversity as a result of the alkaline hydrolysis, with preferential survival of Betaproteobacteria, which increased in relative abundance from 0 to 48 %. Many of the bacterial clone sequences and the 27 isolates were related to known xenobiotic degraders. An archaeal clone library from a non-hydrolyzed sample showed the presence of three main clusters, two representing methanogens and one representing marine aerobic ammonia oxidizers. Isolation of Electronic supplementary material The online version of this article (doi:10.1007/s10646-014-1205-y) contains supplementary material, which is available to authorized users. M. B. Nielsen (&)  K. Ingvorsen Department of Bioscience, Microbiology, Aarhus University, Ny Munkegade 116, Building 1540, 8000 Aarhus C, Denmark e-mail: [email protected] K. U. Kjeldsen  M. A. Lever Department of Bioscience, Center for Geomicrobiology, Aarhus University, Ny Munkegade 116, Building 1540, 8000 Aarhus C, Denmark

alkalitolerant bacterial pure cultures from the hydrolyzed soil confirmed that although alkaline hydrolysis severely reduces microbial community diversity and size certain bacteria survive a prolonged alkaline hydrolysis process. Some of the isolates from the hydrolyzed soil were capable of growing at high pH (pH 10.0) in synthetic media indicating that they could become active in in situ biodegradation upon hydrolysis. Keywords Parathion  p-Nitrophenol  Alkaline hydrolysis  Alkalitolerance  Microbial survival at high pH  Biodegradation

Introduction Organophosphorous pesticides such as parathion and methyl parathion are used extensively worldwide to control insect pests. These pesticides act as acetylcholinesterase inhibitors and thus disrupt neurotransmission in target insects as well as non-target organisms including humans (Cho et al. 2006; Horne et al. 2002; Qiu et al. 2006). Parathion and methyl parathion are considered to be relatively non-persistent in the environment since they are usually quite rapidly degraded by microorganisms in soil and water when applied at rates necessary for insect control (Butler et al. 1981; Cook et al. 1978; Lichtenstein and Schulz 1964). However these compounds are very persistent in soil when present at high concentrations as a result of spillage or disposal (Butler et al. 1981; Wolfe et al. 1973). Breakwater 42 is a highly polluted 90,000 m3 waste dump located at Harboøre Tange in the North-Western part of Jutland, Denmark. In the period from 1953 to 1967 large amounts of chemical waste from a pesticide-producing

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factory were dumped at this site. The waste dump contains approximately 100 different chemicals, the most important ones being parathion, methyl parathion and sulfotep, which collectively comprise an estimated 225 tons of a total of 250 tons of waste. The distribution of the chemicals in the waste dump is very irregular. For example, the concentration of ethyl parathion varies between 0.4 and 31,000 mg/kg (wet weight) in soil samples within the waste dump (Andersen et al. 2007). Likewise in situ pH values ranging from 2.6 to 8.0 also indicate an irregular pattern of distribution of geochemical parameters with respect to location and depth (Andersen et al. 2007). In two previous remediation efforts, a total of 6,850 tons of chemicals and contaminated sand were removed by excavation. The remaining part of the pollution is located in the soil, between 6 and 8.5 m below ground surface (bgs). In 2006 the contaminated area was enclosed by a wall of hydraulically tight steel sheet piling, and covered with a plastic membrane to prevent spreading of the pollution. The pollution is believed to be contained downwards by a thin clay layer at approx. 8.5 m bgs but pollution has sporadically been found below this layer. A current remediation project aims to remove the remaining 250 tons of pollution using a combination of alkaline hydrolysis and bioremediation. The strategy is to first apply alkaline hydrolysis by infiltrating caustic soda into the soil, and thereby raising the pH to C12. Alkaline hydrolysis is expected to convert some of the hydrophobic compounds, some of which may be tightly associated with soil particles or present in free phase, to smaller, more water-soluble components, which are more bio-available and -degradable. For example, parathions will spontaneously hydrolyze to para-nitrophenol (PNP) and diethyl or dimethylthiophosphoric acid resulting in a decreased toxicity (Horne et al. 2002). The hydrolyzed chemicals will be pumped into a wastewater treatment plant for removal by biodegradation. Eventually, the expectedly small amounts of chemicals remaining in the waste depot after the alkaline hydrolysis will be removed by in situ bioremediation. Alkaline hydrolysis has been shown to effectively remediate TNT and RDX in ex situ water and soil experiments (Hwang et al. 2005, 2006; Emmrich 1999) and has been successfully implemented in full-scale remediations of TNT- and RDX-contaminated soils (Britto 2010). To our knowledge, alkaline hydrolysis has not previously been implemented in the remediation of complex mixtures of pollutants on a large scale or in combination with bioremediation. Thus, the effect of alkaline hydrolysis on the survival and community composition of indigenous microbial communities remains unknown. According to established ecological models, stress associated with toxic compounds will decrease the community diversity (Odum 1985; Reardon et al. 2004).

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Indeed, bacterial communities in polluted environments have been found to be less diverse than communities in non-polluted environments (Ro¨ling et al. 2001). Nonetheless, considerable bacterial diversity has been found in polluted environments (Gremion et al. 2003, Militon et al. 2010) and available data indicate, that the ability to degrade anthropogenic pollutants, is broadly distributed among a diverse range of microbial taxa. Several taxonomic groups of bacteria are known to degrade pollutants including members of the genera Pseudomonas (Gammaproteobacteria), Burkholderia (Betaproteobacteria), Sphingomonas (Alphaproteobacteria), Rhodococcus (Actinobacteria), Bacillus (Firmicutes) and Clostridia (Firmicutes) (Abulencia et al. 2006; Gremion et al. 2003; Lillis et al. 2009; Mahmood et al. 2005; Militon et al. 2010; Salehi et al. 2010; Samanta et al. 2000; Tomei et al. 2006). Although such pollutant degraders are known to survive stressful conditions, the effect of an alkaline hydrolysis treatment on the microbial community composition in a contaminated environment has so far not been investigated. To that end it would be very advantageous if a fraction of the indigenous prokaryotic xenobiotic degraders survives alkaline hydrolysis and subsequently can serve as founder community for in situ bioremediation. The extant prokaryotic communities at the Breakwater 42 site have been exposed to heavy chemical pollution for almost 60 years, which has exerted a strong selection pressure for survival and degradation of xenobiotics. This waste dump therefore constitutes a unique study site from a bioremediation perspective. The major purpose of the present study was to investigate the prokaryotic community composition in both soil and water samples. Another objective was to investigate how alkaline hydrolysis affects microbial abundance and diversity. This was done using both cultivation-dependent and cultivation-independent approaches.

Materials and methods Site description and sample collection Samples were collected from the contaminated subsurface soil at the Breakwater 42 waste dump (Harboøre Tange, Denmark; 56° 390 37.8500 N, 8° 100 14.2200 E) in October 2009. Two soil core samples, F2 and F3, were collected by sonic drilling at two highly contaminated sites situated 7 m apart. Samples were taken with a sterile spatula from the center of the cores from 7.5 to 8.2 m bgs in F2 and from 6.6 to 8.25 m bgs in F3 and stored in sterile 50 mL Falcon tubes. The soil had a very strong smell and in some core sections pollutants were visibly present as free phase. The pH of the soil was measured on site with pH sticks

Survival of prokaryotes in a polluted waste dump

(Merck). Samples for the hydrolysis experiment and isolation procedures were stored on ice until returning to the laboratory within \6 h, and subsequently stored at 4 °C until used for analyses (1 day). Water samples were collected in February 2009 from three monitoring wells (Well1, Well2 and Well5) by the use of a Whale inline pump (12 V). The water was allowed to flow for some time before collection of the samples. All three wells were situated within a pilot test area, where alkaline hydrolysis had been carried out for 3 months from October to December 2008. During the pilot test experiment, the pH in these wells was increased from approx. 6 to 12.4 by the addition of 0.44 M NaOH solution. After 3 months, the pH had declined to 11.8, and by the time of sampling, an additional 2 months later, the pH was *8.5–9.0. All water samples were greatly discolored appearing brownish/yellowish. This discoloration indicates the successful action of alkaline hydrolysis in the test area as the yellow color most likely represents PNP. The water was kept on ice and transported to the laboratory in sterile blue-cap bottles, and subsequently stored at 9 °C. Alkaline hydrolysis: laboratory scale Alkaline hydrolysis was simulated in mesocosm laboratory experiments with soil from site F2 and F3. Subsamples of soil were homogenized with a sterile spatula. 12 g (wet weight) of soil were incubated in sterile 50-mL Falcon tubes with 15 mL sterile NaOH solution (0.44 M) and *28 mL headspace of atmospheric air, resulting in a final pH of *13. Non-hydrolyzed soil was incubated under the same conditions. After incubation in duplicate for 12 months in the dark at 4 °C, subsamples were removed and used for DNA extraction and as inoculum for isolation of pure cultures. The pH of the soil at the time of sampling was 12.7 for hydrolyzed soil from both sites, and 7.5 (F2) and 5.5 (F3) for the non-hydrolyzed soil. DNA extraction DNA extraction from soil samples F2 and F3 before and after alkaline hydrolysis was compared using four different methods: (1) The PowerSoilTM DNA Isolation Kit (Mo Bio Laboratories Inc.); (2) The FastDNAÒ SPIN Kit for Soil (MP Biomedicals LLC); (3) The grind, freeze–thaw, SDS method of Zhou et al. (1996) and (4) a new DNA extraction method developed during this study which uses a modified version of the whole cell extraction technique of Kallmeyer et al. (2008) combined with lysis and purification steps; i. e. ten aliquots (0.5 cm3 each) of soil were added to ten sterile 2 mL Eppendorf tubes and mixed with 600 lL 3 % (w/v) NaCl, 100 lL detergent solution [3 % NaCl (w/v), 100 mM disodium EDTA hydrate, 100 mM sodium pyrophosphate

decahydrate, 1 % (v/v) Tween 80 (Kallmeyer et al. 2008)] and 100 lL 99.8 % methanol. The tubes were shaken for 1 h using a Vortex Genie 2 with a 2.0 mL Screw-cap Microtube Holder (Scientific Industries, Inc.). After passive settling of sand grains (approx. 10 min) the supernatants containing suspended cells were transferred to a new tube. The cell extraction procedure of the soil samples was repeated once, however this time including an additional sonication step (15 W for 30 s on ice; Sonopuls HD2070 tip sonicator; Bandelin Electronic) before the vortexing. After removing the supernatant a final cell extraction step was performed by adding 800 lL 3 % NaCl to each tube and vortexing for 10 s followed by removal of the supernatant. Each of the three cell extractions (supernatants) were mixed with 1/10 of a volume of 20 % w/v SDS (sodium dodecyl sulfate solution) and proteinase K (Sigma) solution (1449 U/mL); then incubated for 3 9 1 h at 50 °C under gentle agitation (600 rpm) on an Eppendorf Thermomixer with freezing at -20 °C after each hour. Finally, the temperature was increased to 65 °C followed by an additional 1 h incubation. DNA was precipitated after combining the three cell extracts in a 50 mL Falcon tube by adding one volume of isopropanol and 0.1 volume of 5 M sodium chloride and incubating at -20 °C for 2 h followed by centrifugation at 10,0009g for 30 min at room temperature and washing (one time) with cold 70 % ethanol. The pelleted DNA was resuspended in 100 lL dH2O (Sigma-Aldridge). All DNA extractions were made in duplicates and were subsequently pooled. Due to discoloration, extracts were subjected to a final purification step using the MO BIO PowerCleanÒ DNA Clean-up Kit using a 100 lL final elution volume. DNA from water samples from Well1, Well2 and Well5 was extracted using a method modified from Massana et al. (1997) and Schmidt et al. (1991). Water was filtered through a sterile 0.22 lm vacuum filter unit (Jet Biofil) until the filter clogged (approx. 600 mL). Filters were cut in smaller pieces and mixed with 4.5 mL lysis buffer (40 mM ethylenediamine-tetraacetic acid, 50 mM trizma hydrochloride, 750 mM sucrose) in 50 mL Falcon tubes and subsequently shaken for 10 min at room temperature. Subsequently, proteinase K solution [3 mg Proteinase K (Boehringer) in 400 lL lysis buffer] and 600 lL 10 % w/v SDS were added, and the samples were incubated in a 60 °C water bath for 2 h with gentle inversions every 15–20 min. The lysate was transferred to a new tube and the filter extracted one more time by incubating with 2 mL lysis buffer for an additional 10 min. The two lysates were combined and mixed with 1.3 mL 5 M NaCl. After the mixing, 930 lL 10 % w/v CTAB (cetyltrimethylammonium bromide) in 0.7 M NaCl was added and the tubes were incubated in a 65 °C water bath for 20 min. The lysates were mixed with an equal volume of chloroformisoamyl alcohol (24:1, v/v) and centrifuged at 3,2209g for

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15 min at room temperature. The aqueous phase was recovered and DNA precipitated with a 0.69 volume of isopropanol at room temperature for 30 min followed by centrifugation at 3,2209g for 30 min at room temperature. The pelleted DNA was washed once with cold 70 % ethanol and resuspended in 100 lL dH2O (Sigma-Aldridge). Due to discoloration, extracts were purified using the MO BIO PowerCleanÒ DNA Clean-up Kit using a 100 lL final elution volume. PCR and cloning Bacterial 16S rRNA genes were PCR amplified from water and soil DNA extracts using the bacterial primer 26F (50 AGAGTTTGATCCTGGCTCA-30 (modified from Hicks et al. 1992)) and the prokaryotic primer 1492R (50 GGYTACCTTGTTACGACTT-30 (Loy et al. 2002)). Archaeal 16S rRNA genes were amplified using the archaeal primer ARC-8F [50 -TCCGGTTGATCCTGCC-30 (Teske et al. 2002)] and the prokaryotic primer 1492R (Loy et al. 2002). PCR reaction mixtures consisted of 10 lL of Hot Star Taq Master Mix (Qiagen), 0.4 lL of each primer (10 pmol/lL, MWG Biotech AG, Ebersberg, Germany), 0.8 lL bovine serum albumin (BSA, 10 lg/lL, Amersham Biosciences) and 8.4 lL template solution. Thermal cycling included 95 °C for 15 min, then 30 (bacteria) or 40 (archaea) cycles of 95 °C for 45 s, 57 °C for 45 s and 72 °C for 90 s. Cycling was completed after a final elongation step at 72 °C for 10 min. PCR products were purified using the GenEluteTM PCR Clean-Up Kit (Sigma) and cloned using the pGEMÒ-T vector system (Promega) with chemical competent E. coli JM 109 cells according to the manufacturer’s specifications. Inserts of purified plasmids were sequenced on one strand using the 26F or ARC-8F primer (Macrogen, Seoul, Korea). The 16S rRNA genes from bacterial pure cultures were either PCR-amplified directly from colony cell material dissolved in dH2O, or from DNA extracts obtained with the PowerSoilTM DNA Isolation Kit (Mo Bio Laboratories Inc.). The PCR primers and thermal cycling were as described above with the following PCR reaction mixture: 25 lL of Hot Star Taq Master Mix (Qiagen), 1 lL of each primer (10 pmol/lL, MWG Biotech AG), 2 lL BSA (10 lg/lL, Amersham Biosciences), 20 lL dH2O and 1 lL template solution. The PCR products were sequenced on both strands using the 26F/1492R primers (Macrogen, Seoul, Korea). qPCR Bacterial and archaeal 16S rRNA gene abundance in soil DNA extracts was quantified by qPCR using a LightCycler 480 Instrument (Roche). The PCR amplification mixture

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included: 10 lL SYBR Green I Master 29 (Roche), 2 lL BSA (10 mg/L, Amersham Biosciences), 1 lL of forward and reverse primer (10 lM each) and 6 lL template. The following primer pairs were used for the bacterial and archaeal assay respectively: Bac8Fmod (50 -AGAGTTTGA TYMTGGCTCAG-30 , modified from Loy et al. 2002)/ Bac338Rabc (50 -GCWGCCWCCCGTAGGWGT-30 , modified from Daims et al. 1999) and Arch806F (50 -ATTAGA TACCCSBGTAGTCC-30 , Takai and Horikoshi 2000)/ Arch958R (50 -YCCGGCGTTGAMTCCAAT-30 , DeLong 1992). Thermal cycling consisted of an initial denaturation step at 95 °C for 5 min followed by 50 cycles consisting of 95 °C for 30 s, 55 °C for 30 s and 72 °C for 15 s. At the end of each cycle (at 80 °C) the fluorescent signal was measured at 530 nm. Standard curves were prepared from serial dilutions of plasmids pGEMÒ-T containing an environmental bacterial or archaeal 16S rRNA gene insert generated with the primer pairs 26F/1492R or ARC-8F/ 1492R, respectively. The standard curves were linear from 4.5 9 103 - 4.5 9 109 gene copies per reaction for Bacteria and 3.1 9 102 - 3.1 9 109 gene copies per reaction for Archaea. The number of cells per gram wet soil was calculated assuming 4.08 bacterial and 1.76 archaeal gene copies per cell according to Lee et al. 2009.

Analysis of sequence data 16S rRNA gene sequences were aligned using the SINA Web Aligner of the SILVA rRNA database project (Pruesse et al. 2007) and added to the Silva SSURef release 102 ARB database (Pruesse et al. 2007; Ludwig et al. 2004). All alignments were refined manually. Phylogenetic trees and distance matrices were calculated using the neighbor-joining algorithm implemented in the ARB program package (Ludwig et al. 2004). Distance matrices for bacterial and archaeal sequences were calculated without distance correction and respectively using the bacterial and archaeal positional variability filters included in the SSURef release 102 ARB database for selecting alignment positions. The distance matrices served as input files for Mothur (Schloss et al. 2009), which was used for grouping sequences into operational taxonomic units (OTUs) according to a furthest neighbor principle, for rarefaction analyses and for inter clone library comparisons. Higher order taxonomic profiles of bacterial 16S rRNA gene sequences were generated using the online classifier of the Ribosomal Database Project (Wang et al. 2007). The sequences determined in this study have been deposited in the GenBank database under accession numbers KF641187–KF641788.

Survival of prokaryotes in a polluted waste dump

Isolation of pure cultures

Results

Inocula for isolation of pure cultures from non-hydrolyzed and hydrolyzed soil samples were prepared by mixing 1 g soil (non-hydrolyzed soil or hydrolyzed soil slurry) with 8.8 mL saline water (0.9 % w/v NaCl, pH 6.4) and 200 lL 100 mM Na-pyrophosphate followed by 15 min vortexing and ultrasonication on ice for 2 9 20 s at 30 W using a Sonopuls HD2070 tip sonicator (Bandelin Electronic, Berlin, Germany). From this mixture tenfold serial dilutions (up to 10-6) were made in sterile saline water (0.9 % w/v NaCl, pH 6.4) and 100 lL aliquots were plated in duplicate on four different types of solid medium (see below). Furthermore, slurries of hydrolyzed soil and hydrolyzed soil mixed in a 1:1 ratio with saline water (0.9 %, pH 6.4) were spread directly on the four solid media. Isolation of pure cultures from the soil samples was performed on the following solid media: R2A (DifcoTM) (pH 7.2), R2A with 15 g/L NaCl (pH 7.2), enrichment medium (EM) with 10 g/L NaCl (pH 7.5) (Nielsen et al. 2011) and tryptic soy broth (Scharlau, Spain) (pH 7.3). Plates were incubated under both oxic and anoxic conditions at 15 °C. In addition, soil (1.5 g wet weight) from the hydrolyzed samples was inoculated into 100 mL liquid ISP2 medium (Nielsen et al. 2011) at pH 8.5 and 9.6 and incubated at room temperature under oxic conditions. Pure cultures were obtained by successive subculturing on plates. For the isolation of pure cultures from water samples, 250 mL of water was filtered through a 0.22 lm vacuum filter unit (Jet Biofil, Canada JET Biochemicals) and the filter was incubated in 100 mL ISP2 medium adjusted to pH 8.5 (Nielsen et al. 2011) in 500 mL Erlenmeyer flasks. The flasks were incubated on a shaker (100 rpm) at atmospheric oxygen levels at 15 °C and at room temperature (22–23 °C). Cultures exhibiting growth (increased turbidity) were diluted as described above and plated on ISP2 agar plates (pH 8.5). Pure cultures were obtained as described above.

Contaminated subsurface samples are usually not accessible for cultivation-independent studies of microbial community structure due to the extremely low cell counts which makes it difficult to isolate sufficient DNA even for PCR-based analyses (Abulencia et al. 2006). Soil from Breakwater 42 proved to be very recalcitrant to DNA extraction. Despite numerous attempts and modifications of published protocols, DNA could only be extracted from soil samples using the newly developed method number four, which is based on an initial whole cell extraction step. Using this method DNA extracts adequate for PCR could was obtained from subsamples from sample F3 (before and after alkaline hydrolysis). The amount of DNA extracted from the two types of subsamples was too low to be detected on 1 % agarose gels, but near full length bacterial 16S rRNA gene fragments could be PCR amplified from both subsamples, and archaeal 16S rRNA gene fragments were PCR-amplifiable from the DNA extracts of the nonhydrolyzed subsample.

Growth of isolates at high pH The ability of isolates from hydrolyzed soil to grow at high pH was tested in 10 mL R2A medium in glass test tubes. The pH of the medium was adjusted after sterilization by aseptic addition of a Na-sesquicarbonate solution (42 g/L NaHCO3 and 53 g/L Na2CO3). The highest pH obtained with this solution was pH 10.0 (by the addition of 60 mL/ L). When testing growth at higher pH values, the medium was titrated to pH 10.8 and 11.1 using 3 M NaOH. The tubes were incubated on a rotary shaker (100 rpm) at room temperature.

qPCR qPCR analysis based on the DNA extracted from the site F3 soil showed the presence of a very low number of bacterial and archaeal 16S rRNA gene copies in the soil. For the non-hydrolyzed soil subsample, the number of bacterial gene copies per gram wet soil was 2.1 9 104. For the hydrolyzed soil, the number of copies was below the qPCR detection limit (*4.5 9 103 copies per reaction), indicating a decrease in bacterial abundance. For the Archaea, the number of gene copies per gram was 2900 for non-hydrolyzed soil and below the qPCR detection limit (3.1 9 102 gene copies per reaction) for the hydrolyzed soil, again indicating a decrease in total abundance. Prokaryotic community structure of Breakwater 42 soil Clone libraries were constructed from each of the resulting PCR amplicons and the plasmid inserts of 207 (nonhydrolyzed soil) and 176 (hydrolyzed soil), and 84 (nonhydrolyzed soil) clones were sequenced from the two bacterial and the single archaeal clone library, respectively. Rarefaction analysis of the clone library sequence data shows that alkaline hydrolysis caused a pronounced decrease in bacterial diversity with 71 detectable OTUs before alkaline hydrolysis and 37 after alkaline hydrolysis (based on a 97 % sequence identity cut off) (Fig. 1; Online Resource 2). Only 16 OTUs were shared between the two libraries (Online Resource 2). According to a test by the Libshuff method (Singleton et al. 2001), the phylotype composition of the two clone libraries was significantly

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M. B. Nielsen et al. Table 1 Taxonomic affiliation of archaeal 16S rRNA gene sequence OTUs retrieved from sediment core F3 OTU number

Abundancea

Closest characterized relativeb

Identity (%)b

Phylum Thaumarchaeota Family Nitrosopumilaceae 1

1/84

Candidatus Nitrosoarchaeum koreensis MY1 (HQ331116)

93

Family Nitrososphaeraceae 2

26/84

Nitrososphaera viennensis EN76 (FR773157)

96

3

2/84

95

4

1/84

Nitrososphaera viennensis EN76 (FR773157) Nitrososphaera viennensis EN76 (FR773157)

93

Miscellaneous crenarchaeotic group Fig. 1 Rarefaction curves calculated for bacterial 16S rRNA gene clone libraries derived from soil site F3 before (upper curve) and after (lower curve) alkaline hydrolysis. Operational taxonomic unit (OTU) grouping was based on a 97 % sequence identity cut off

different (p \ 0.0001 for both pairwise comparisons). Archaeal diversity was low, with only seven OTUs detectable among the 84 sequenced clones (Table 1; Online Resource 1). The clone library derived from the non-hydrolyzed F3 soil was dominated by sequences representing Gram positive bacteria, primarily Clostridia (38 %) and Bacilli (9 %) (Fig. 2). Alpha-, Gamma- and Deltaproteobacteria were also present with Alphaproteobacteria, making up the largest group (16 %) while Acidobacteria comprised 13 % of the clone sequences. None of the remaining clone sequences comprised more than 6 % of the clone library. Most clone sequences derived from the hydrolyzed soil belonged to the Betaproteobacteria, making up almost half of the sequences (48 %). This class of bacteria was not detected in the non-hydrolyzed soil. By contrast, the Deltaproteobacteria, which made up 11 % of the sequences in the non-hydrolyzed soil, were not represented in the hydrolyzed soil. The Gram positive Bacilli and Clostridia were also present in the hydrolyzed-soil, but at a somewhat lower relative abundance compared to the non-hydrolyzed soil. The clone sequences in the hydrolyzed soil belonged to eight different classes or phyla, compared to 12 in the non-hydrolyzed soil. Hence, the alkaline hydrolysis treatment caused a clear decrease in diversity, which was also demonstrated by the decrease in OTUs (Fig. 1). The majority of the archaeal clone sequences belonged to the phylum Euryarchaeota (62 %) and the rest belonged to the phylum Thaumarchaeota (38 %) (Table 1). Archaeal 16S rRNA gene PCR amplicons could not be obtained from the hydrolyzed soil indicating a marked reduction in

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5

2/84

None

–

Phylum Euryarchaeota/class Methanomicrobia Family Methanocellaceae 6

28/84

Methanocella arvoryzae MRE50 (AM114193)

Family Methanosarcinaceae 7 24/84 Methanosarcina siciliae DSM3028T (FR733698)

96

98

According to phylogenetic analyses based on the Silva SSURef_111 arb database and the NCBI taxonomy of the closest characterized relative OTUs based on a sequence identity cut off of 97 % a

Relative abundance in clone library expressed as number of clones belonging to a given OTU out of a total of 84 clones

b

According to phylogenetic analyses based on the Silva SSURef_111 arb database

the abundance of Archaea due to the alkaline hydrolysis treatment. Bacterial community structure of Breakwater 42 water samples DNA extracted from the water samples was used to construct a clone library comprised of a total of 108 sequences; 22 clones from Well1, 54 clones from Well2 and 32 clones from Well5. The diversity in the wells differed both between the three wells, and between the wells and the soil. In Well1 a large part of the clone sequences belonged to the Deltaproteobacteria (54 %) and 18 % of the sequences belonged to the Clostridia (Fig. 2). In Well2 the majority of the clone sequences belonged to the Gammaproteobacteria (48 %) and the Actinobacteria (46 %). Both of these classes were absent in Well5, where the majority of clone sequences belonged to the Alphaproteobacteria (16 %), Betaproteobacteria (28 %),

Survival of prokaryotes in a polluted waste dump

Fig. 2 Taxonomic distribution and relative clone library abundance of bacterial 16S rRNA gene sequence clones derived from soil site F3 before and after alkaline hydrolysis treatment as well as from three

nearby contaminated ground water wells. The detailed phylogenetic relationship of the sequence data is shown in Online Resource 2. n = number of sequences analyzed

Deltaproteobacteria (31 %) and Epsilonproteobacteria (16 %). LIBSHUFF analyses showed that sequence identity compositions of the clone libraries constructed for Well1 and Well2, and Well2 and Well5 are significantly different (all p values \0.0001), while the compositions of Well1 and Well5 were not significantly different (pairwise p values = 0.1856 and 0.0648). LIBSHUFF analyses comparing the sequence identity composition of the clone libraries constructed for wells with those of the soil showed that the three wells were significantly different from both the nonhydrolyzed and hydrolyzed soil (all p values\0.0001).

inoculated with the soil/water mix. No anaerobic strains were isolated, but a total of 11 aerobic isolates were obtained and identified. Total cell counts using SYBR Gold and DAPI were attempted with soil samples from Breakwater 42 to confirm this low bacterial abundance, but unfortunately efficient separation of the cells from the soil and free phase organics failed, hindering a precise cell count. Together the 27 isolates (Table 2) represented a taxonomically diverse range of Bacteria; ten isolates were affiliated with Gammaproteobacteria, two with Betaproteobacteria, four with Alphaproteobacteria, three with Actinobacteria and eight with Firmicutes. Most isolates shared high sequence similarities (98–100 %) with cultivated Bacteria and with phylotypes detected in the 16S rRNA gene clone libraries (Table 2). The isolates from the nonhydrolyzed soil were dominated by Gammaproteobacteria (9 out of 15 isolates), which were only represented by one isolate in the hydrolyzed sample. Similar to the clone library results (see below), isolates affiliated with the Betaproteobacteria were not found in the non-hydrolyzed soil, but appeared in the hydrolyzed soil (2 isolates).

Isolation of pure cultures No growth was observed on plates inoculated with F2 soil. For the non-hydrolyzed F3 soil samples, growth was observed on all of the tested media. In general, very few colonies appeared on the inoculated plates with the TSB plates producing the fewest. The maximum number of colonies was observed on EM ? NaCl (17 colonies on the 10-1 dilution plates incubated under oxic conditions). Anoxic incubation produced a maximum of three colonies on the 10-1 dilution plates. A total of ten aerobic bacterial strains and five anaerobic bacterial strains were isolated and identified by 16S rRNA gene sequencing from the nonhydrolyzed F3 soil sample. For the F3 hydrolyzed soil samples (F3H1), most colonies were observed on R2A plates, with a maximum of two colonies on plates

Growth of pure cultures at high pH Eleven isolates were obtained from the hydrolyzed soil three of which were closely related to phylotypes detected in the clone libraries derived from the hydrolyzed soil

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M. B. Nielsen et al. Table 2 Isolates from non-hydrolyzed F3 soil, hydrolyzed F3 soil and Well2. For each isolate, the accession number, the closest relative according to BLAST search, the isolation conditions and high pH Isolate

Accession no.

Closest relativea

growth conditions, the degree of similarity and the taxonomic affiliation are given

Isolation conditions/high pH growth conditions

Identity (%)a

Taxonomic affiliation

Phylotype presence in clone librariesb F3-B

F3-H

Well 2

Non-hydrolyzed soil R2A, pH 7.4

99

Gammaproteobacteria

97 %

0

0

EM ? NaCl, pH 7.5

99

Firmicutes

0

0

0

Stenotrophomonas sp. (GU391493)

R2A, pH 7.4

99

Gammaproteobacteria

97

0

0

Sphingobium yanoikuyae (EU307932) Acinetobacter iwoffii (FJ860882)

R2A, pH 7.4

99

Alphaproteobacteria

0

0

0

R2A, pH 7.4

99

Gammaproteobacteria

0

0

97 %

KF641659

Pseudomonas fluorescens (GU198112)

TSB, pH 7.3

100

Gammaproteobacteria

0

0

99 %

F3_a7

KF641660

Edwardsiella tarda (FJ405303)

EM ? NaCl, pH 7.5

99

Gammaproteobacteria

0

0

99 %

F3_a8

KF641661

Pseudomonas sp. (GU391489)

R2A, pH 7.4

99

Gammaproteobacteria

0

0

99 %

F3_a9

KF641662

Pseudomonas fluorescens (GU198101)

EM ? NaCl, pH 7.5

99

Gammaproteobacteria

0

0

99 %

F3_a10

KF641663

Acinetobacter iwoffii (FJ860882)

R2A, pH 7.4

99

Gammaproteobacteria

0

0

97 %

F3_an1

KF641664

Bacillus sp. (FR727703)

EM ? NaCl, pH 7.5

99

Firmicutes

99 %

99 %

0

F3_an2

KF641665

Bacillus sp. (FR727703)

EM ? NaCl, pH 7.5

99

Firmicutes

99 %

99 %

0

F3_an3

KF641666

Acinetobacter radioresistens (GU145275)

EM ? NaCl, pH 7.5

99

Gammaproteobacteria

0

0

97 %

F3_an4

KF641667

Cellulomonas cellasea (NR_037077)

R2A, pH 7.4

97

Actinobacteria

0

0

0

F3_an5

KF641668

Cellulomonas cellasea (NR_037077)

R2A, pH 7.4

98

Actinobacteria

0

0

0

F3_a1

KF641654

F3_a2

KF641655

F3_a3

KF641656

F3_a4

KF641657

F3_a5

KF641658

F3_a6

Hydrolyzed soil F3H1_a1 KF641669

Stenotrophomonas sp. (AM745261) Bacillus pumilus (GU191914)

Bacillus sp. (JF309254)

R2A, pH 7.4/R2A, pH 10.0

Firmicutes

0

0

0

F3H1_a2

KF641670

Naxibacter haematophilus (EU554441)

R2A, pH 7.4

98

Betaproteobacteria

0

96 %

0

F3H1_a3

KF641671

Herbaspirillum seropedicae (HQ406764)

R2A, pH 7.4

99

Betaproteobacteria

0

96 %

0

F3H1_a4

KF641672

Sphingomonas aerolata (FR691420)

R2A, pH 7.4

99

Alphaproteobacteria

0

0

0

F3H1_a5

KF641673

Pseudomonas stutzeri (CP002622)

R2A, pH 7.4/R2A, pH 10.0

99

Gammaproteobacteria

0

0

97 %

F3H1_a6

KF641674

Bacillus novalis (AJ542512)

TSB, pH 7.3

98

Firmicutes

0

0

0

F3H1_a7

KF641675

Planococcus rifietoensis (KF749392)

R2A, pH 7.4/R2A, pH 10.0

98

Firmicutes

0

0

0

F3H1_a8

KF641676

Methylobacterium sp. (EU912446)

R2A, pH 7.4

99

Alphaproteobacteria

0

0

0

123

100

Survival of prokaryotes in a polluted waste dump Table 2 continued Isolate

Accession no.

F3H1_a9

KF641677

F3H1_a10

KF641678

F3H1_a11

KF641679

KF641680

Closest relativea

Isolation conditions/high pH growth conditions

Identity (%)a

Taxonomic affiliation

Phylotype presence in clone librariesb F3-B

F3-H

Well 2

R2A, pH 7.4

98

Alphaproteobacteria

0

0

0

R2A, pH 7.4

99

Actinobacteria

0

0

0

Planococcus sp. (AY745836)

ISP2, pH9.6/R2A, pH 10.0

99

Firmicutes

0

0

0

Aerococcus viridians (HQ425688)

ISP2, pH 8.5

99

Firmicutes

0

0

0

Sphingomonas humi (AB220146) Nocardioides sp. (AJ565419)

Well2 W2_a1 a

According to BLAST searches of the NCBI (http://www.ncbi.nlm.nih.gov) nucleotide sequence database

b

Presence is indicated by the pairwise identity shared with the most closely related phylotype in the 16S rRNA gene clone libraries respectively derived from soil samples F3 before (F3-B) and after (F3-H) alkaline hydrolysis and from one of the three sampled wells. Zero indicates that no phylotype sharing [95 % pairwise identity was detected

sample and from well2 (Table 2). Surviving 12 months at an elevated pH of 12.7–13 may imply a certain physiological adaptation to high pH values, although the four isolates belonging to the Firmicutes may have survived due to their ability to form endospores. Growth of the isolates at high pH was therefore tested. Of the 11 isolates, the following four grew at pH 10: F3H1_a1 (Firmicute endospore-former) (final DOD600 = 0,380), F3H1_a5 (Pseudomonas) (final DOD600 = 0,335), F3H1_a7 (Firmicute endospore-former) (final DOD600 = 0.677) and F3H1_a11 (Firmicute endospore-former) (final DOD600 = 1.202). Growth at higher pH values (10.8 and 11.1) was also tested. Two isolates (F3H1_a7 and F3H1_a11) exhibited growth at these initial high pH values, but the pH decreased to ten during growth. Nonetheless, isolation of bacteria from hydrolyzed soil indicate that certain bacteria in the Breakwater 42 waste dump are capable of surviving alkaline hydrolysis for an extended period of time, and they may also be capable of growth at high pH values.

Discussion The waste dump at Breakwater 42 was established in 1953 and contains 250 tons of approximately 100 different chemicals, many of which are highly toxic to humans, animals and microorganisms. The objective of the current remediation effort is to detoxify the site to a point where the threat to the surrounding aquatic environment is eliminated or greatly reduced. The strategy is to use a novel combination of alkaline hydrolysis and bioremediation. To the best of our knowledge the impact of an alkaline hydrolysis process on the indigenous microbial community

has never been reported and nothing is known about the microbial community in this unique environment. We analyzed the prokaryotic community in soil samples before and after a 1-year alkaline hydrolysis treatment by a cultivation-isolation-dependent approach and a culture-independent 16S rRNA gene-dependent approach. The prokaryotic community in Breakwater 42 Total prokaryotic community size in the soil was very low with 5,100 bacterial and 1,600 archaeal cells detected per gram wet soil respectively by qPCR. These numbers represent minimum estimates due to difficulties in extraction of DNA from the soil matrix, which resulted in successful DNA extraction with only one of the four methods tested. Of these four methods, two commercial soil DNA extraction kits and a published DNA extraction method for soils failed to provide amplifiable DNA, whereas a new method designed here resulted in successful isolation of amplifiable DNA. Based on published comparisons on soils and other sample types it is not unusual—but in fact the norm—to observe quantitative and phylogenetic differences in DNA extraction efficiency between extraction methods and sample types due to differences in cell lysis and purification methods (e.g. Zhou et al. 1996; Hurt et al. 2001; Sørensen et al. 2004; Delmont et al. 2011; Flores et al. 2012; Willner et al. 2012)—with so far not one method working best on all sample types. Though we do not know why our protocol was successful here, a conspicuous difference to the other three protocols is that we perform a cell isolation step prior to cell lysis, whereas the three other protocols rely on direct DNA lysis in the presence of soil. This initial isolation step is likely to have (1) minimized

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DNA sorption to particles, which can result in significant loss of DNA during direct extraction on low-biomass samples, and (2) reduced the background of co-extracted organic matter, e.g. humic acids, which may interfere downstream with (a) DNA recovery during DNA purification and (b) PCR amplification, if they are insufficiently removed during DNA purification. The alkaline treatment reduced the community size even further as illustrated by the decrease in the number of both bacterial and archaeal cells per gram wet soil to values below the qPCR detection limit (4.5 9 103 and 3.1 9 102 gene copies per reaction for Bacteria and Archaea, respectively). This is equivalent to 6.9 9 103 and 4.8 9 102 gene copies per gram wet soil for Bacteria and Archaea, respectively. For comparison, the number of cells in sediment samples from an aquifer (10–12 m bgs) contaminated with chlorinated ethenes varied between 5 9 106 to 1 9 107 cells per gram sediment (Davis et al. 2002) which are 10–100 times lower than typical cell abundances in pristine soil. Microscopic investigation of F2 and F3 soil samples was very difficult due to autoflourescens but indicated a very low cell abundance, which was in accordance with qPCR data and low plate counts from F3 (approx. 1,800 CFU/g wet soil) obtained on R2A, pH 7.2. The existence of low cell numbers is supported by results obtained for the F2 soil sample, from which no isolates and no DNA extract could be obtained. Together this suggests that the degree of pollution in the waste dump is highly inhibitory to microbial growth resulting in cell densities several orders of magnitude below those of pristine surface soils and sediments. The environmental conditions at 7–9 m bgs in the Breakwater 42 waste dump are very heterogeneous. The isolation of aerobic and anaerobic strains and the presence of many clone sequences respectively related to both known aerobic and anaerobic taxa is a strong indication of the presence of both oxic and anoxic niches in the soil. In accordance, significant heterogeneity in chemical parameters (pH, salinity, oxygen tension, types and concentrations of pollutants) have been observed within the Breakwater 42 waste dump (Andersen et al. 2007) and are likely to have a pronounced effect on prokaryotic abundance and community structure. The clone sequences from the non-hydrolyzed soil mainly belong to the Alphaproteobacteria, Deltaproteobacteria and the Clostridia (Fig. 2), which are common soil bacteria that have often been found in contaminated soil environments (e.g. Abulencia et al. 2006; Gremion et al. 2003; Lillis et al. 2009; Reardon et al. 2004). Members of the Clostridia are known pollutant degraders, e.g. being able to degrade PNP (Suresh et al. 2007) and 2,4,6-trinitrotoluene (TNT) (Watrous et al. 2003). The ability of

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Clostridia to form endospores might facilitate their survival in the waste dump; however, DNA extraction procedures are typically unlikely to lyse endospores at high efficiency as compared to vegetative cells indicating that the anaerobic clostridial phylotypes represent active-vegetative cells. Many of the sequences belonging to the Clostridia are closely related to the sulfate-reducing genera Desulfosporosinus and Desulfotomaculum. Other clone sequences within the Deltaproteobacteria are related to known sulfate reducers like members of the genera Desulfobulbus, Desulfococcus and Desulfovibrio (Online Resource 2). This indicates the presence of sulfate and anoxic conditions suitable for sulfate reduction, which is likely in a seawaterimpacted environment like Breakwater 42. The competition between sulfate reducers and methanogens depends on the concentration of sulfate and the electron donors available. Typically sulfate reducers outcompete methanogens over a range of concentrations of sulfate and common substrates such as H2 (Kristjansson et al. 1982; Lovley et al. 1982). Methanogens are, however, frequently present in sulfate-containing environments (Lovley et al. 1982) and in the Breakwater 42 soil a large part of the Archeal clone sequences belonged to the methanogenic Methanosarcina (29 %) and Methanocella (34 %) (Table 1). Methanosarcina use methylated compounds, acetate, formate, and H2/ CO2, and are commonly found to coexist with sulfate reducers, because most sulfate reducers are unable to utilize methylated substrates. The high abundance of Methanocella, which uses only H2/CO2, indicate that sulfate depleted areas are present within the waste dump. In accordance with the finding of methanogens and thus methane production, some of the Alphaproteobacterial clone sequences are related to members of the methanotrophic genera Methylocystis and Methylobacterium. Strains of Methylobacterium have also been found to degrade xenobiotics like explosives (Van Aken et al. 2004) and dichloromethane (Doronina et al. 2000). Another indication of the presence of reduced one-carbon compounds is the affiliation of some of the Alphaproteobacterial sequences with methylotrophic Hyphomicrobium species. According to BLAST searches many of the clone sequences obtained from the Breakwater 42 waste dump were most closely related to organisms isolated from contaminated sites or to sequences of uncultured organisms from such environments (results not shown). These sites include a radioactive waste site (Field et al. 2010), coal-tarwaste-contaminated aquifer waters (Bakermans and Madsen 2002), a benzene degrading enrichment culture (Kunapuli et al. 2007), a reactor treating monochlorobenzene contaminated groundwater (Alfreider et al. 2002), hydrocarbon-contaminated soil (Militon et al. 2010), uranium contaminated soil (Brodie et al. 2006), asphalt permeated

Survival of prokaryotes in a polluted waste dump

soil (Kim and Crowley 2007), arsenic contaminated soil (Karnachuk et al. 2009) and coal-tar contaminated water (Yagi et al. 2010). The exposure of the soil for 12 months to alkaline hydrolysis conditions (pH 12.7–13) had a marked effect on the diversity of the microbial community (cf. Figs. 1, 2, Online Resource 1, Table 2). Alkaline hydrolysis was associated with an abrupt decrease in the diversity as shown by the decrease in recoverable OTUs from 71 to 37 at the 97 % sequence identity cut-off and a marked shift in the community composition towards betaproteobacterial predominance (Figs. 1, 2). The appearance of Betaproteobacteria is puzzling, and although we cannot rule out that the results are influenced by DNA extraction bias, the results may be explained by: (1) cell growth during the alkaline hydrolysis treatment; however, the two isolates from the hydrolyzed soil belonging to the Betaproteobacteria were not capable of growing at high pH in the laboratory, or (2) the initial abundance of the Betaproteobacteria before the hydrolysis was too low to be detected in the clone libraries, but due to preferential survival of these bacteria during the hydrolysis, their relative abundance increased, making them abundant in the clone libraries. Notably many of the betaproteobacterial genera represented in the clone library of the hydrolyzed soil contain species described as degraders of pollutants. Especially Ralstonia and Burkholderia are well known degraders of numerous pollutants; for example Ralstonia degrading PNP (Samanta et al. 2000; Tomei et al. 2006; Salehi et al. 2010), 3-Methyl-4-Nitrophenol (breakdown product of the organophosphorous insecticide fenitrothion) (Bhushan et al. 2000) chloroaromatic compounds (Matus et al. 2003), 4-aminobenzenesulfonate (Gan et al. 2011), and carbazole (Schneider et al. 2000), and Burkholderia degrading polychlorinated biphenyls (Chain et al. 2006), PNP (Prakash et al. 1996), trichloroethene (Barth et al. 2002), fenitrothion (Hayatsu et al. 2000) and polycyclic aromatic hydrocarbons (Kang et al. 2003). Clone sequences belonging to Delftia, Variovorax, Diaphorobacter and Herbaspirillum were also found in the hydrolyzed soil and have been described to degrade polycyclic aromatic hydrocarbons (Klankeo et al. 2009; Saul et al. 2005) and herbicides (Dejonghe et al. 2003; Leibeling et al. 2010; Sørensen et al. 2008). The fact that these Bacteria are aerobic needs to be considered when implementing in situ bioremediation at the Breakwater 42 waste dump. The initial oxygen consumption will probably be low due to reduced microbial biomass but as the pH and the concentration of the toxic compounds are lowered and the conditions in the waste dump becomes more favorable for bacterial growth, the oxygen consumption will probably increase. Therefore, aeration may be necessary in order to exploit the biodegradative potential of these Bacteria. An unique feature of the water samples as compared to the F3 soil samples is the presence of a large fraction of

Actinobacteria in Well2 (48 % of the clone sequences). Actinobacteria were not detected in the non-hydrolyzed soil, Well1 and Well5, and was only represented by two clone sequences in the hydrolyzed soil samples. Interestingly, all clone sequences from Well2 (pH = 8.5–9.0) belonging to the Actinobacteria (26 sequences) were members of the genus Citricoccus, the closest relative being Citricoccus nitrophenolicus (21 out of 26 clone sequences share a 99–100 % sequence similarity with this species). C. nitrophenolicus was previously isolated from the waste water treatment plant at the pesticide producing factory situated next to the Breakwater 42 waste dump (Nielsen et al. 2011). It is capable of degrading PNP at pH 10 (Nielsen and Ingvorsen 2013), and can survive long periods of high pH, i.e. a pure culture of the strain survived at least 30 days at pH values between 11.8 and 12.8 (unpublished data). Also, the water samples were collected from an area subjected to a small-scale alkaline hydrolysis experiment. Presence of the strain in the water indicates its ability to survive alkaline conditions. Hence, this strain might survive a large-scale alkaline hydrolysis process and when pH decreases after termination of the hydrolysis the strain might be capable of degrading the PNP produced in situ by the abiotic hydrolysis of parathions. The strain may, therefore, be a potential candidate for in situ or ex situ bioremediation of the Breakwater 42 site. Isolates Most of the isolated bacterial strains belong to genera with members described as degraders of various pollutants. Two bacterial isolates from the hydrolyzed soil (F3H1_a7 and F3H1_a11) belong to the Planococcus genus (Table 2). Members of the Planococcus genus are aerobic, non-spore forming bacteria, often found in marine environments. The species P. alkanoclasticum is capable of degrading hydrocarbons (Engelhardt et al. 2001). Four Pseudomonas strains were isolated from the soil. Three of the isolates from non-hydrolyzed soil (F3_a6, F3_a8, F3_a9) had a 99–100 % sequence similarity to P. fluorescens (Table 2), which degrade for example 3,4-dihaloanilines (Travkin et al. 2003) and petroleum hydrocarbons (Barathi and Vasudevan 2001). Other Pseudomonas species have been shown to degrade parathions (e.g. Chaudhry et al. 1988; Choi et al. 2009). The fourth Pseudomonas strain (F3H1_a5; Table 2) was isolated from hydrolyzed soil. Two isolates (F3H1_a1 and F3H1_6) show 98–100 % sequence similarity to Bacillus species (Table 2). Bacillus species have been reported to degrade for example PNP (Kadiyala and Spain 1998). Their ability to form spores facilitates survival under unfavorable conditions like high pH and high pollutant concentrations. Two Betaproteobacterial species (F3H1_a2, F3H1_a3) belonging to the

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genera Naxibacter and Herbaspirillum, respectively, were isolated from the hydrolyzed soil, while no Betaproteobacteria were isolated from the non-hydrolyzed soil (Table 2). Herbaspirillum chlorophenolicum is known to degrade phenol and chlorophenol (Im et al. 2004) while Naxibacter alkalitolerans is capable of growing at pH 12 (Xu et al. 2005). Two Alphaproteobacterial isolates belonging to Sphingomonas were obtained from the hydrolyzed soil (F3H1_a4 and F3H1-a9); this group of Bacteria possesses versatile metabolic capabilities and has been shown to degrade pentachlorophenol (e.g. Nohynek et al. 1996; Ederer et al. 1997), PNP (Leung et al. 1997), polycyclic aromatic hydrocarbons like fluorine (Wattiau et al. 2001), benzo[a]pyrene (Rentz et al. 2008), chlorpyrifos (Li et al. 2007) and parathion (Kumar and D’Souza 2010). Finally, an actinobacterial isolate belonging to the Nocardioides was isolated from the hydrolyzed soil; a species within this genus has been shown to degrade PNP (Yoon et al. 1999). The ability of Bacteria to survive 12 months at pH 12.7–13 is interesting. It might be due to the fact that these bacteria reside in biofilms, in dense aggregates or adhere to solid phase organics, which offer protection from the harsh chemical conditions in the environment. For example, the biofilm mode of growth has been shown to confer resistance to environmental stressors, such as heavy metals (Teitzel and Parsek 2003), which are known to be present in the Breakwater 42 waste dump. The isolation of 11 bacterial strains from hydrolyzed soil (upon 12 months of incubation at pH 12.7–13) suggests that a fraction of the indigenous bacterial population will survive the upcoming infusion of the waste depot with NaOH solution. Subsequent testing revealed that only four of these isolates, namely F3H1_a1, F3H1_a5, F3H1_a7 and F3H1_a11, were able to grow in R2A medium adjusted to pH 10.0 (Table 2). Two of the strains (F3H1_a7 and F3H1_a11) also grew at higher initial pH values (10.8 and 11.1) and even though pH decreased during growth to a final value of ten this indicates that they might be capable of growing at pH values higher than ten. Our results thus suggest that part of the indigenous bacterial population at Breakwater 42 is able to survive extremely high pH values in situ for at least 12 months, and is able to resume growth when subjected to lower pH values. Therefore it is likely that some pollutant degrading microorganisms will survive exposure to the large-scale alkaline hydrolysis planned to take place at the Breakwater 42 depot, and resume activity when the pH is subsequently lowered to more neutral pH values. The bioremediation project The results of this study show that alkaline hydrolysis has an adverse effect on the indigenous prokaryotic community

123

in the waste dump. The implementation of alkaline hydrolysis prior to bioremediation will lower the bacterial numbers and the diversity in the soil. Some bacteria do survive the hydrolysis, and some of them are potentially capable of degrading pollutants, but it needs to be demonstrated whether such putative alkalitolerant or alkaliphilic species can proliferate and carry out significant in situ decontamination of pollutants in a large scale remediation process of the Breakwater 42 waste depot. The exclusive appearance of aerobic bacteria after alkaline hydrolysis deserves consideration as this may necessitate aeration and/or NO3- injection in order to affect in situ bioremediation. Due to the very low cell numbers in the soil it may be necessary to implement bioaugmentation for successful remediation of the waste dump. Acknowledgments The research was supported by a joint grant from Cheminova A/S, Central Denmark Region and The Aarhus University Research Foundation (AUFF). Thanks to Loren Mark Ramsay (Alectia), Morten Bondgaard (Central Denmark Region), Børge Hvidberg (Central Denmark Region) and Lars Ernst (Central Denmark Region) for providing geochemical data and helping with soil and water sampling. K.U. Kjeldsen and M.A. Lever were funded by the Danish National Research Foundation. Conflict of interest of interest.

The authors declare that they have no conflict

References Abulencia CB, Wyborski DL, Garcia JA, Podar M, Chen WQ, Chang SH, Chang HW, Watson D, Brodie EL, Hazen TC, Keller M (2006) Environmental whole-genome amplification to access microbial populations in contaminated soils. Appl Environ Microbiol 72:3291–3301 Alfreider A, Vogt C, Babel W (2002) Microbial diversity in an in situ reactor system treating monochlorobenzene contaminated groundwater as revealed by 16S ribosomal DNA analysis. System Appl Microbiol 25:232–240 Andersen L, Jørgensen C, Holm J (2007) In situ biologisk nedbrydning som oprensningsmetode ved Høfde 42. Danish Ministry of the Environment. http://www2.mst.dk/Udgiv/publikationer/ 2007/978-87-7052-608-1/pdf/978-87-7052-609-8.pdf Bakermans C, Madsen EL (2002) Diversity of 16S rDNA and naphtalene dioxygenase genes from coal-tar-contaminated aquifer waters. Microbiol Ecol 44:95–106 Barathi S, Vasudevan N (2001) Utilization of petroleum hydrocarbons by Pseudomonas fluorescens isolated from a petroleumcontaminated soil. Environ Int 26:413–416 Barth JAC, Slater G, Schu¨th C, Bill M, Downey A, Larkin M, Kalin RM (2002) Carbon isotope fractionation during aerobic biodegradation of trichloroethene by Burkholderia cepacia G4: a tool to map degradation mechanisms. Appl Environ Microbiol 68:1728–1734 Bhushan B, Samanta SK, Chauhan A, Chakraborti AK, Jain RK (2000) Chemotaxis and biodegradation of 3-methyl-4-nitrophenol by Ralstonia sp. SJ98. Biochem Biophys Res Commun 275: 129–133 Britto R (2010) Full-scale Alkaline Hydrolysis of Organic Explosives in Soil. www.tetratech.com

Survival of prokaryotes in a polluted waste dump Brodie EL, DeSantis TZ, Joyner DC, Baek SM, Larsen JT, Andersen GL, Hazen TC, Richardson PM, Herman DJ, Tokunaga TK, Wan JM, Firestone MK (2006) Application of a high-density oligonucleotide microarray approach to study bacterial population dynamics during uranium reduction and reoxidation. Appl Environ Microbiol 72:6288–6298 Butler LC, Staiff DC, Davis JE (1981) Methyl parathion persistence in soil following simulated spillage. Arch Environ Contem Toxicol 10:451–458 Chain PS, Denef VJ, Konstantinidis KT, Vergez LM, Agullo L, Reyes VL, Hauser L, Cordova M, Gomez L, Gonzalez M, Land M, Lao V, Larimer F, LiPuma JJ, Mahenthiralingam E, Malfatti SA, Marx CJ, Parnell JJ, Ramette A, Richardson P, Seeger M, Smith D, Spilker T, Sul WJ, Tsoi TV, Ulrich LE, Zhulin IB, Tiedje JM (2006) Burkholderia xenovorans LB400 harbors a multi-replicon, 9.73-Mbp genome shaped for versatility. Proc Natl Acad Sci USA 103:15280–15287 Chaudhry GR, Ali AN, Wheeler WB (1988) Isolation of a methyl parathion-degrading Pseudomonas sp. that possesses DNA homologous to the opd gene from a Flavobacterium sp. Appl Environ Microbiol 54:288–293 Cho CMH, Mulchandani A, Chen W (2006) Functional analysis of organophosphorus hydrolase variants with high degradation activity towards organophosphate pesticides. Protein Eng Des Sel 19:99–105 Choi M-K, Kim K-D, Ahn K-M, Shin D-H, Hwang J-H, Seong CN, Ka J-O (2009) Genetic and phenotypic diversity of parathiondegrading bacteria isolated from rice paddy soils. J Microbiol Biotechnol 19:1679–1687 Cook AM, Daughton CG, Alexander M (1978) Phosphorus-containing pesticide breakdown products: quantitative utilization as phosphorus sources by bacteria. Appl Environ Microbiol 36:668–672 Daims H, Bru¨hl A, Amann R, Schleifer K-H, Wagner M (1999) The domain-specific probe EUB338 is insufficient for the detection of all bacteria: development and evaluation of a more comprehensive probe set. System Appl Microbiol 22:434–444 Davis JW, Odom JM, DeWeerd KA, Stahl DA, Fishbain SS, West RJ, Klecka GM, DeCarolis JG (2002) Natural attenuation of chlorinated solvents at area 6, Dover air force base: characterization of microbial community structure. J Contam Hydrol 57:41–59 Dejonghe W, Berteloot E, Goris J, Boon N, Crul K, Maertens S, Hofte M, De Vos P, Verstraete W, Topp EM (2003) Synergistic degradation of linuron by a bacterial consortium and isolation of a single linuron-degrading Variovorax strain. Appl Environ Microbiol 69:1532–1541 Delmont TO, Robe P, Cecillon S, Clark IM, Constancias F, Simonet P, Hirsch PR, Vogel TM (2011) Accessing the soil metagenome for studies of microbial diversity. Appl Environ Microbiol 77:1315–1324 DeLong EF (1992) Archaea in coastal marine environments. Proc Natl Acad Sci USA 89:5685–5689 Doronina NV, Trotsenko YA, Tourova TP, Kuznetsov BB, Leisinger T (2000) Methylopila helvetica sp. nov. and Methylobacterium dichloromethanicom sp. nov.: Novel aerobic facultatively methylotrophic bacteria utilizing dichloromethane. System Appl Microbiol 23:210–218 Ederer MM, Crawford RL, Herwig RP, Orser CS (1997) PCP degradation is mediated by closely related strains of the genus Sphingomonas. Mol Ecol 6:39–49 Emmrich M (1999) Kinetics of the alkaline hydrolysis of 2,4,6trinitrotoluene in aqueous solution and highly contaminated soils. Environ Sci Technol 33:3802–3805 Engelhardt MA, Daly K, Swannell RPJ, Head IM (2001) Isolation and characterization of a novel hydrocarbon-degrading, Gram-

positive bacterium, isolated from intertidal beach soil, and description of Planococcus alkanoclasticus sp. nov. J Appl Microbiol 90:237–247 Field EK, D’Imperio S, Miller AR, VanEngelen MR, Gerlach R, Lee BD, Apel WA, Peyton BM (2010) Application of molecular techniques to elucidate the influence of cellulosic waste on the bacterial community structure at a simulated low-level-radioactive-waste site. Appl Environ Microbiol 76:3106–3115 Flores GE, Henley JB, Fierer N (2012) A direct PCR approach to accelerate analyses of human-associated microbial communities. PLoS ONE 7:e44563 Gan HM, Shahir S, Ibrahim Z, Yahya A (2011) Biodegradation of 4-aminobenzenesulfonate by Ralstonia sp. PBA and Hydrogenophaga sp. PBC isolated from textile wastewater treatment plant. Chemosphere 82:507–513 Gremion F, Chatzinotas A, Harms H (2003) Comparative 16S rDNA and 16S rRNA sequence analysis indicates that actinobacteria might be a dominant part of the metabolically active bacteria in heavy metal-contaminated bulk and rhizosphere soil. Environ Microbiol 5:896–907 Hayatsu M, Hirano M, Tokuda S (2000) Involvement of two plasmids in fenitrothion degradation by Burkholderia sp. Strain NF100. Appl Environ Microbiol 66:1737–1740 Hicks RE, Amann RI, Stahl DA (1992) Dual staining of natural bacterioplankton with 40 ,6-diamidino-2- phenylindole and fluorescent oligonucleotide probes targeting kingdom-level 16S rRNA sequences. Appl Environ Microbiol 58:2158–2163 Horne I, Sutherland TD, Harcourt RL, Russell RJ, Oakeshott JG (2002) Identification of an opd (organophosphate degradation) gene in an Agrobacterium isolate. Appl Environ Microbiol 68:3371–3376 Hurt RA, Qiu X, Wu L, Roh Y, Palumbo AV, Tiedje JM, Zhou J (2001) Simultaneous recovery of RNA and DNA from soils and sediments. Appl Environ Microbiol 67:4495–4503 Hwang S, Ruff TJ, Bouwer EJ, Larson SL, Davis JL (2005) Applicability of alkaline hydrolysis for remediation of TNTcontaminated water. Water Res 39:4503–4511 Hwang S, Felt DR, Bouwer EJ, Brooks MC, Larson SL, Davis JL (2006) Remediation of RDX-contaminated water using alkaline hydrolysis. J Environ Eng 132:256–262 Im W-T, Bae H-S, Yokota A, Lee ST (2004) Herbaspirillum chlorophenolicum sp. nov., a 4-chlorophenol-degrading bacterium. Int J Syst Evol Microbiol 54:851–855 Kadiyala V, Spain JC (1998) A two-component monooxygenase catalyzes both the hydroxylation of p-nitrophenol and the oxidative release of nitrite from 4-nitrocatechol in Bacillus sphaericus JS905. Appl Environ Microb 64:2479–2484 Kallmeyer J, Smith DC, Spivack AJ, D’Hondt S (2008) New cell extraction procedure applied to deep subsurface sediments. Limnol Oceanogr Methods 6:236–245 Kang H, Hwang SY, Kim YM, Kim E, Kim Y-S, Kim S-K, Kim SW, Cerniglia CE, Shuttleworth KL, Zylstra GJ (2003) Degradation of phenanthrene and naphthalene by a Burkholderia species strain. Can J Microbiol 49:139–144 Karnachuk OV, Gerasimchuk AL, Banks D, Frengstad B, Stykon GA, Tikhonova ZL, Kaksonen A, Puhakka J, Yanenko AS, Pimenov NV (2009) Bacteria of the sulfur cycle in the soils of gold mine tailings, Kuznetsk Basin, Russia. Microbiology 78:483–491 Kim J-S, Crowley DE (2007) Microbial diversity in natural asphalts of the Rancho La Brea Tar Pits. Appl Environ Microbiol 73:4579–4591 Klankeo P, Nopcharoenkul W, Pinyakong O (2009) Two novel pyrene-degrading Diaphorobacter sp. and Pseudoxanthomonas sp. isolated from soil. J Biosci Bioeng 108:488–495 Kristjansson JK, Scho¨nheit P, Thauer RK (1982) Different Ks values for hydrogen of methanogenic bacteria and sulfate reducing

123

M. B. Nielsen et al. bacteria: an explanation for the apparent inhibition of methanogenesis by sulfate. Arch Microbiol 131:278–282 Kumar J, D’Souza SF (2010) An optical microbial biosensor for detection of methyl parathion using Sphingomonas sp. immobilized on microplate as a reusable biocomponent. Biosens Bioelectron 26:1292–1296 Kunapuli U, Lueders T, Meckenstock RU (2007) The use of stable isotope probing to identify key iron-reducing microorganisms involved in anaerobic benzene degradation. ISME J 1:643–653 Lee ZM-P, Bussema C III, Schmidt TM (2009) rrn DB: documenting the number of rRNA and tRNA genes in bacteria and archaea. Nucleic Acids Res 37(SUPPL. 1):D489–D493 Leibeling S, Schmidt F, Jehmlich N, von Bergen M, Mu¨ller RH, Harms H (2010) Declining capacity of starving Delftia acidovorans MC1 to degrade phenoxypropionate herbicides correlates with oxidative modification of the initial enzyme. Environ Sci Technol 44:3793–3799 Leung KT, Tresse O, Errampalli D, Lee H, Trevors JT (1997) Mineralization of p-nitrophenol by pentachlorophenol-degrading Sphingomonas spp. FEMS Microbiol Lett 155:107–114 Li X, He J, Li S (2007) Isolation of a chlorpyrifos-degrading bacterium, Sphingomonas sp. strain Dsp-2, and cloning of the mpd gene. Res Microbiol 158:143–149 Lichtenstein EP, Schulz KR (1964) The effects of moisture and microorganisms on the persistence and metabolism of some organophosphorous insecticides in soil, with special emphasis on parathion. J Econ Entomol 57:618–627 Lillis L, Doyle E, Clipson N (2009) Comparison of DNA- and RNAbased bacterial community structures in soil exposed to 2,4dichlorophenol. J Appl Microbiol 107:1883–1893 Lovley DR, Dwyer DF, Klug MJ (1982) Kinetic analysis of competition between sulfate reducers and methanogens for hydrogen in soils. Appl Environ Microbiol 43:1373–1379 Loy A, Lehner A, Lee N, Adamczyk J, Meier H, Ernst J, Schleifer K-H, Wagner M (2002) Oligonucleotide microarray for 16S rRNA gene-based detection of all recognized lineages of sulfatereducing prokaryotes in the environment. Appl Environ Microbiol 68:5064–5081 Ludwig W, Strunk O, Westram R, Richter L, Meier H, Yadhukumar Buchner A, Lai T, Steppi S et al (2004) ARB: a software environment for sequence data. Nucleic Acids Res 32:1363–1371 Mahmood S, Paton GI, Prosser JI (2005) Cultivation-independent in situ molecular analysis of bacteria involved in degradation of pentachlorophenol in soil. Environ Microbiol 7:1349–1360 Massana R, Murray AE, Preston CM, DeLong EF (1997) Vertical distribution and phylogenetic characterization of marine planktonic archaea in the Santa Barbara channel. Appl Environ Microbiol 63:50–56 Matus V, Sa´nchez MA, Martı´nez M, Gonza´lez B (2003) Efficient degradation of 2,4,6-trichlorophenol requires a set of catabolic genes related to tcp genes from Ralstonia eutropha JMP134(pJP4). Appl Environ Microbiol 69:7108–7115 Militon C, Boucher D, Vachelard C, Perchet G, Barra V, Troquet J, Peyretaillade E, Peyret P (2010) Bacterial community changes during bioremediation of aliphatic hydrocarbon-contaminated soil. FEMS Microbiol Ecol 74:669–681 Nielsen MB, Ingvorsen K (2013) Biodegradation of para-nitrophenol by Citricoccus nitrophenolicus strain PNP1T at high pH. Biodegradation 24:79–87 Nielsen MB, Kjeldsen KU, Ingvorsen K (2011) Description of Citricoccus nitrophenolicus sp. nov., a para-nitrophenol degrading actinobacterium isolated from a wastewater treatment plant and emended description of the genus Citricoccus Altenburger et al. 2002. Antonie Leeuwenhoek 99: 489–499

123

Nohynek LJ, Suhonen EL, Nurmiaho-Lassila E-L, Hantula J, Salkinoja-Salonen M (1996) Description of four pentachlorophenol-degrading bacterial strains as Sphingomonas chlorophenolica sp. nov. Syst Appl Microbiol 18:527–538 Odum EP (1985) Trends expected in stressed ecosystems. Bioscience 35:419–422 Prakash D, Chauhan A, Jain RK (1996) Plasmid-encoded degradation of p-nitrophenol by Pseudomonas cepacia. Biochem Biophys Res Commun 224:375–381 Pruesse E, Quast C, Knittel K, Fuchs B, Ludwig W, Peplies J, Glo¨ckner FO (2007) SILVA: a comprehensive online resource for quality checked and aligned ribosomal RNA sequence data compatible with ARB. Nuc Acids Res 35:7188–7196 Qiu XH, Bai WQ, Zhong QZ, Li M, He FQ, Li BT (2006) Isolation and characterization of a bacterial strain of the genus Ochrobactrum with methyl parathion mineralizing activity. J Appl Microbiol 101:986–994 Reardon CL, Cummings DE, Petzke LM, Kinsall BL, Watson DB, Peyton BM, Geesey GG (2004) Composition and diversity of microbial communities recovered from surrogate minerals incubated in an acidic uranium-contaminated aquifer. Appl Environ Microbiol 70:6037–6046 Rentz JA, Alvarez PJJ, Schnoor JL (2008) Benzo[a]pyrene degradation by Sphingomonas yanoikuyae JAR02. Environ Pollut 151:669–677 Ro¨ling WFM, van Breukelen BM, Braster M, Lin B, van Verseveld HW (2001) Relationships between microbial community structure and hydrochemistry in a landfill leachate-polluted aquifer. Appl Environ Microbiol 67:4619–4629 Salehi Z, Vahabzadeh F, Sohrabi M, Fatemi S, Znad HT (2010) Statistical medium optimization and biodegradative capacity of Ralstonia eutropha toward p-nitrophenol. Biodegradation 21:645–657 Samanta SK, Bhushan B, Chauhan A, Jain RK (2000) Chemotaxis of a Ralstonia sp. SJ98 toward different nitroaromatic compounds and their degradation. Biochem Biophys Res Commun 269:117–123 Saul DJ, Aislabie JM, Brown CE, Harris L, Foght JM (2005) Hydrocarbon contamination changes the bacterial diversity of soil from around Scott Base, Antarctica. FEMS Microbiol Ecol 53:141–155 Schloss PD, Westcott SL, Ryabin T, Hall JR, Hartmann M, Hollister EB, Lesniewski RA, Oakley BB, Parks DH, Robinson CJ, Sahl JW, Stres B, Thallinger GG, Van Horn DJ, Weber CF (2009) Introducing mothur: open-source, platform-independent, community-supported software for describing and comparing microbial communities. Appl Environ Microbiol 75:7537–7541 Schmidt TM, Delong EF, Pace NR (1991) Analysis of a marine picoplankton community by 16 S ribosomal-RNA gene cloning and sequencing. J Bacteriol 173:4371–4378 Schneider J, Grosser RJ, Jayasimhulu K, Xue W, Kinkle B, Warshawsky D (2000) Biodegradation of carbazole by Ralstonia sp. RJGII.123 isolated from a hydrocarbon contaminated soil. Can J Microbiol 46:269–277 Singleton DR, Furlong MA, Rathbun SL, Whitman WB (2001) Quantitative comparisons of 16S rRNA gene sequence libraries from environmental samples. Appl Environ Microbiol 67:4373–4376 Sørensen KB, Lauer A, Teske A (2004) Archaeal phylotypes in a metal-rich and low-activity deep subsurface sediment of the Peru Margin, ODP Leg 201, Site 1231. Geobiology 2:151–161 Sørensen SR, Albers CN, Aamand J (2008) Rapid mineralization of the phenylurea herbicide diuron by Variovorax sp. Strain SRS16 in pure culture and within a two-member consortium. Appl Environ Microbiol 74:2332–2340

Survival of prokaryotes in a polluted waste dump Suresh K, Prakash D, Rastogi N, Jain RK (2007) Clostridium nitrophenolicum sp. nov., a novel anaerobic p-nitrophenoldegrading bacterium, isolated from a subsurface soil sample. Int J Syst Evol Microbiol 57:1886–1890 Takai K, Horikoshi K (2000) Rapid detection and quantification of members of the archaeal community by quantitative PCR using fluorogenic probes. Appl Environ Microbiol 66:5066–5072 Teitzel GM, Parsek MR (2003) Heavy metal resistance of biofilm and planktonic Pseudomonas aeruginosa. Appl Environ Microbiol 69:2313–2320 Teske A, Hinrichs KU, Edgcomb V, Gomez AD, Kysela D, Sylva SP, Sogin ML, Jannasch HW (2002) Microbial diversity of hydrothermal sediments in the Guaymas basin: evidence for anaerobic methanotrophic communities. Appl Environ Microbiol 68:1994–2007 Tomei MC, Rossetti S, Annesini MC (2006) Microbial and kinetic characterization of pure and mixed cultures aerobically degrading 4-nitrophenol. Chemosphere 63:1801–1808 Travkin VM, Solyanikova IP, Rietjens IMCM, Vervoort J, Van Berkel WJH, Golovleva LA (2003) Degradation of 3,4-dichloroand 3,4-difluoroaniline by Pseudomonas fluorescens 26-k. J Environ Sci Health B 38:121–132 Van Aken B, Yoon JM, Schnoor JL (2004) Biodegradation of nitrosubstituted explosives 2,4,6-trinitrotoluene, hexahydro-1,3,5-trinitro-1,3,5-triazine, and octahydro-1,3,5,7-tetranitro-1,3,5-tetrazocine by a phytosymbiotic Methylobacterium sp. associated with poplar tissues (Populus deltoids nigra DN34). Appl Environ Microbiol 70:508–517 Wang Q, Garrity GM, Tiedje JM, Cole JR (2007) Naive bayesian classifier for rapid assignment of rRNA sequences into the new bacterial taxonomy. Appl Environ Microbiol 73:5261–5267

Watrous MM, Clark S, Kutty R, Huang S, Rudolph FB, Hughes JB, Bennett GN (2003) 2,4,6-Trinitrotoluene reduction by an Feonly hydrogenase in Clostridium acetobutylicum. Appl Environ Microbiol 69:1542–1547 Wattiau P, Bastiaens L, van Herwijnen R, Daal L, Parsons JR, Renard ME, Springael D, Cornelis GR (2001) Fluorene degradation by Sphingomonas sp. LB126 proceeds through protocatechuic acid: a genetic analysis. Res Microbiol 152:861–872 Willner D, Daly J, Whiley D, Grimwood K, Wainwright CE, Hugenholtz P (2012) Comparison of DNA extraction methods for microbial community profiling with an application to pediatric bronchoalveolar lavage samples. PLoS ONE 7:e34605 Wolfe HR, Staiff DC, Armstrong JF, Comer SW (1973) Persistence of parathion in soil. Bull Environ Contam Toxicol 10:1–9 Xu P, Li W-J, Tang S-K, Zhang Y-Q, Chen G-Z, Chen H–H, Xu L-H, Jiang C-L (2005) Naxibacter alkalitolerans gen. nov., sp. nov., a novel member of the family ‘Oxalobacteraceae’ isolated from China. Int J Sys Evol Microbiol 55:1149–1153 Yagi JM, Neuhauser EF, Ripp JA, Mauro DM, Madsen EL (2010) Subsurface ecosystem resilience: long-term attenuation of subsurface contaminants supports a dynamic microbial community. ISME J 4:131–143 Yoon J-H, Cho Y-G, Lee Taik S, Suzuki K-I, Nakase T, Park Y-H (1999) Nocardioides nitrophenolicus sp. nov., a p-nitrophenoldegrading bacterium. Int J Syst Bacteriol 49:675–680 Zhou JZ, Bruns MA, Tiedje JM (1996) DNA recovery from soils of diverse composition. Appl Environ Micobiol 62:316–322

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