Microbial Community Changes in Response to Ethanol or Methanol ...

APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Sept. 2010, p. 5728–5735 0099-2240/10/$12.00 doi:10.1128/AEM.00308-10 Copyright © 2010, American Society for Microbiology. All Rights Reserved.

Vol. 76, No. 17

Microbial Community Changes in Response to Ethanol or Methanol Amendments for U(VI) Reduction䌤 Tatiana A. Vishnivetskaya,1 Craig C. Brandt,1 Andrew S. Madden,2 Meghan M. Drake,1 Joel E. Kostka,3 Denise M. Akob,3,4 Kirsten Ku ¨sel,4 and Anthony V. Palumbo1* Oak Ridge National Laboratory, Oak Ridge, Tennessee 378311; The University of Oklahoma, College of Earth and Energy, School of Geology and Geophysics, Norman, Oklahoma 730192; Florida State University, Tallahassee, Florida 323063; and Institute of Ecology, Friedrich Schiller University Jena, Dornburger Strasse 159, D-07743 Jena, Germany4 Received 4 February 2010/Accepted 22 June 2010

Microbial community responses to ethanol, methanol, and methanol plus humics amendments in relationship to U(VI) bioreduction were studied in laboratory microcosm experiments using sediments and ground water from a uranium-contaminated site in Oak Ridge, TN. The type of carbon source added, the duration of incubation, and the sampling site influenced the bacterial community structure upon incubation. Analysis of 16S rRNA gene clone libraries indicated that (i) bacterial communities found in ethanol- and methanolamended samples with U(VI) reduction were similar due to the presence of Deltaproteobacteria and Betaproteobacteria (members of the families Burkholderiaceae, Comamonadaceae, Oxalobacteraceae, and Rhodocyclaceae); (ii) methanol-amended samples without U(VI) reduction exhibited the lowest diversity and the bacterial community contained 69.2 to 92.8% of the family Methylophilaceae; and (iii) the addition of humics resulted in an increase of phylogenetic diversity of Betaproteobacteria (Rodoferax, Polaromonas, Janthinobacterium, Methylophilales, and unclassified) and Firmicutes (Desulfosporosinus and Clostridium). such as Clostridium spp. (20). These data suggest that U(VI) can be reduced by many microorganisms once suitable electron donors are available. The purpose of this study was to analyze the ability of various amendments to stimulate the reduction of U(VI) by the indigenous microbial communities found in subsurface sediments collected from a uranium-contaminated site. A previous publication from this project (42) gave a very limited analysis of the microbial community. Here we present a detailed phylogenetic analysis of the bacterial community structure and link community structure to capability of U(VI) reduction in sediments stimulated with ethanol and methanol. This study was designed to explore whether microbial communities that demonstrate U(VI) reduction after stimulation with different alcohols show a similar structure. Also, it was designed to detect differences between the methanol-stimulated communities that were capable of U(VI) reduction and those that were not capable of U(VI) reduction. Since humic substances have been reported to promote U(VI) reduction (10, 34), we also examined the effects of humics on the community structure and reduction of U(VI).

The use of uranium in nuclear research, fuel production, and weapons manufacturing has resulted in environmental contamination at production, manufacturing, and storage sites throughout the United States. Although all of the common isotopes of uranium (238U [99.27%], 235U [0.72%], and 234U [0.005%]) are radioactive, it is the chemical toxicity of uranium that is usually of greatest concern when it is present as a contaminant. The U.S. Department of Energy (DOE) has ongoing efforts to identify and remediate contaminated areas under its control. Stimulating the in situ metabolism of microorganisms capable of reduction of U(VI) to U(IV), producing the insoluble mineral uraninite which precipitates and renders uranium immobile in ground water, has been proposed as an environmentally safe and a potentially cost-effective remediation method (37). Typically, an organic substrate is added to stimulate microbial growth and promote the development of anaerobic conditions, under which the reduction of U(VI) is favored (67). Various substrates (e.g., acetate, ethanol, glucose, and methanol) have been used either in the field or in microcosm studies, and most were capable of stimulating microbial U(VI) reduction (1, 8, 42, 43, 47, 60); however, the addition of methanol did not always result in U(VI) reduction (49). Many microorganisms are known to reduce U(VI) in pure culture, including a hyperthermophilic archaeon (28), a thermophilic bacterium Thermoterrabacterium ferrireducens (29), the mesophilic dissimilatory metal-reducing bacteria Geobacter and Shewanella (67) and Anaeromyxobacter dehalogenans (71), the sulfate-reducing bacterium Desulfovibrio sp. (61), and fermentative bacteria

MATERIALS AND METHODS Microcosm experiments. Bulk sediments contaminated with U(VI) were collected from two sites, designated here P4 and P5, located at the Subsurface Biogeochemistry Research program’s Oak Ridge Field Research Center (ORFRC). The ORFRC is located at the Y-12 National Security Complex on the DOE Oak Ridge Reservation in Oak Ridge, TN. The sites are near the S-3 ponds, which were historically used as a disposal area for low-level liquid radioactive waste. The acidic (pH 3 to 5) sediments and groundwater from the P4 and P5 sites contained 4.3 and 5.4 ␮M uranium (VI), 2,727.9 and 985.9 ppm nitrate, and 22.6 and 57.6 ppm sulfate, respectively, and various minor inorganic, radionuclide, and organic constituents (for a detailed site characterization see reference 45). Uncontaminated samples were collected from a nearby location out-

* Corresponding author. Mailing address: Biosciences Division, Oak Ridge National Laboratory, Oak Ridge, TN 37831-6342. Phone: (865) 481-8914. Fax: (865) 576-0524. E-mail: [email protected]. 䌤 Published ahead of print on 2 July 2010. 5728

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TABLE 1. Characteristics and diversity estimates for 16S rRNA gene clone libraries derived from the eight microcosm experiments and background Result based on 99% and 97% identities Expt label

P5EU42 P5EU137 P5MU78 P5MhU78 P4EU16 P4EU38 P4MN10 P4MN38 BG

Site

Amendment

P5

Ethanol

P4

Methanol Methanol ⫹ humics Ethanol Methanol No

Incubation (days)

42 137 78 78 16 38 10 38 0

No. of replicates

3 3 2 2 2 2 2 2 2

Decrease in soluble U(VI)

⫹ ⫹ ⫹ ⫹ ⫹ ⫹ ⫺ ⫺ ND

DNA yield (␮g g⫺1)

0.7–1.5 0.05–0.3 1.8–2.1 1.5–1.8 0.7–1.1 0.2–3.0 0.7–0.8 1.0–1.4 1.9–2.1

No. of clonesa

274 276 164 136 187 185 178 165 182

No. of OTUsb

Shannon (H’)c

Simpson (1/D)d

Homologous coveragee (%)

99%

97%

99%

97%

99%

97%

99%

97%

47 60 42 57 62 52 23 27 28

38 47 29 41 51 47 17 16 19

1.98 3.19 2.76 3.66 3.26 3.07 1.93 1.72 2.53

1.88 2.93 2.34 3.21 2.93 2.91 1.61 1.29 1.74

2.63 13.34 9.62 32.25 13.34 10.75 3.85 2.70 8.20

2.56 11.23 6.67 19.23 10.0 10.0 3.13 2.17 3.14

89.7 89.5 81.7 76.5 75.4 80.9 92.1 89.6 84.6

93.4 92.0 89.6 85.3 82.4 83.8 94.4 95.2 89.5

a

Total number of clones sequenced for each experiment. Number of operational taxonomic units based on 99% and 97% identities of the partial 16S rRNA gene sequences. Shannon index; higher numbers represent greater diversity. d Reciprocal of Simpson’s index; higher numbers represent greater diversity. e The equation for calculating coverage for a single sample (X) is CX ⫽ 1 ⫺ (NX/n), where NX is the number of OTUs in the sample and n is the total number of sequences. A lower value for homologous coverage indicates a higher number of OTUs or unique sequences in the population. b c

side the ORFRC site. The sediments were briefly exposed to air (20.9% oxygen) during sampling. The bulk sediment samples were homogenized anaerobically in a glove bag (N2:H2, 97:3; COY Laboratory Products Inc., Grass Lake, MI). Approximately 20 g of sediment and 80 ml of ORFRC groundwater were added to 0.5-liter serum bottles, and the pH was adjusted to 7.0 using anoxic sodium bicarbonate. The concentrations of electron donors were selected relative to a single electron transfer (65): methanol (40 mM), industrial-grade ethanol (20 mM), glucose (10 mM), acetate (30 mM), lactate (20 mM), pyruvate (24 mM), glycerol (17 mM) (42). Natural humic material, extracted from uncontaminated topsoil obtained at the ORFRC, was added in a concentration of 30 mg liter⫺1 to one of the methanol treatments. Detailed extraction procedures and characteristics of isolated humics have been given elsewhere (23, 40). Four replicate microcosms were prepared for each treatment. Unamended controls were included in each experiment. The microcosms were incubated at ambient temperature under anaerobic conditions in a glove bag (N2:H2, 97:3). Samples for aqueous chemical constituents and DNA were collected from the microcosms by using sterile needles and syringes in an anaerobic glove bag. For uranium, nitrate, and sulfate measurements the microcosm P4 was sampled every other day from day 0 through day 38, while microcosm P5 was sampled every other day from day 0 through day 12 and every sixth day from day 18 through day 137. Changes in aqueous uranium, nitrate, and sulfate concentrations from the microcosm experiments have been presented previously (42). Microbial community analysis is presented for microcosms amended with methanol (P5M and P4M), methanol plus humics (P5Mh), and ethanol (P5E and P4E) to allow comparisons between conditions where uranium reduction was observed to those where it was not. Samples for DNA extraction were taken (i) when the sediments turned from light yellow to dark black, which indicated iron reduction and likely the precipitation of biologically reduced U in sediments, and (ii) at the final time point. Approximately 1.5 ml of groundwater/sediment (4/1, vol/wt) was used for DNA extraction. 16S rRNA gene clone libraries. The total community genomic DNA (cgDNA) was extracted from duplicate or triplicate samples (see Table 1) using the PowerSoil DNA isolation kit (MO BIO Laboratories, Inc., Carlsbad, CA). Taq polymerase was used to amplify the purified cgDNA with primers targeted to Escherichia coli 16S rRNA positions 8 to 27 (5⬘-AGA GTT TGA TCC TGG CTC AG-3⬘) and 1510 to 1492 (5⬘-GGT TAC CTT TTA CGA CTT-3⬘). The forward (27f) primer, based on recent critical evaluation, amplifies about 70.9% of known bacterial species (21). After amplification the PCR products were extracted and purified from UltraPure agarose (Invitrogen, Carlsbad, CA) by using a QIAquick gel extraction kit (Qiagen Inc., Valencia, CA). Following purification, PCR products were ligated in pCR 2.1-TOPO vectors (Invitrogen, Carlsbad, CA), transformed into One Shot Mach1-T1R (Invitrogen, Carlsbad, CA) chemically competent E. coli, and plated onto LB agar containing 50 ␮g ml⫺1 kanamycin and 5-bromo-4-chloro-3-indolyl-␤-D-galactopyranoside. White colonies (typically 90 to 100) resulting from growth of transformants incubated overnight at 37°C

were selected. The selected colonies were grown at 37°C with aeration in LB broth containing 50 ␮g ml⫺1 kanamycin. Clones were sequenced using the BigDye Terminator v3.1 cycle sequencing kit (Applied Biosystems, Inc., Foster City, CA) and a reverse primer targeting positions 536 to 519 of the E. coli 16S rRNA gene [5⬘-G(A/T)A TTA CCG CGG C(G/T)G CTG-3⬘] (33). An Applied Biosystems 3730 automated system (Foster City, CA) was used for sequencing. A total of 1,728 clones were sequenced, trimmed, and filtered for quality, resulting in 1,565 high-quality 16S rRNA gene sequences. Phylogenetic and statistical analyses. The 16S rRNA gene sequences were taxonomically assigned using the Naïve Bayesian rRNA classifier of the Ribosomal Database Project II (RDP) (68). Sequences from this study were subsequently aligned using the ClustalW multiple alignment tool (66) of the program BioEdit v7.0.5.3 (25). The program DNADIST v3.5c in the BioEdit program was used to compute a distance matrix from the aligned nucleotide sequences (17). The distance matrix was input into the DOTUR program (v1.53) to assign the sequences to operational taxonomic units (OTUs) using the furthest-neighbor clustering algorithm (57) at 99% and 97% identity. Rarefaction curves were produced by standard calculations using the total number of clones obtained compared to the number of clones representing each unique OTU. The Shannon index and Simpson’s diversity index were calculated using the DOTUR program (57). A neighbor-joining tree was created using MEGA version 4 software (64). The bootstrap data represented 1,000 samplings. Multiple environments were simultaneously analyzed by comparing the microbial communities phylogenetically using weighted UniFrac to conduct a principal coordinates analysis (38, 39). The neighbor-joining tree generated for input to UniFrac was limited to 999 sequences and rooted with the Methanosarcina mazei 16S rRNA gene sequence (EF452664). Initially, sequences from each clone library were aligned separately, and OTUs were identified at 97% identity. One representative sequence was selected for most OTUs. The environmental input file for UniFrac contained the count of how many times the selected sequence appeared in the clone library. The UniFrac significance test with abundance weights was used to determine which communities were significantly different. P values were corrected for multiple comparisons by multiplying the calculated P value by the number of comparisons made (Bonferroni correction) (38, 39). Nucleotide sequence accession numbers. The sequences obtained in this study were deposited into the NCBI GenBank database under accession numbers GU949620 through GU950618.

RESULTS Population size and clone libraries descriptions. The DNA yield ranged from 0.05 to 3.0 ␮g g⫺1 of wet sediment (Table 1). Assuming that the predicted effective genome size of the soil bacterial/archaeal population is 4.7 Mb, as estimated from

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FIG. 1. Rarefaction analysis of the 16S rRNA gene clone library coverage of microbial diversity. OTUs were defined at 97% sequence identity.

metagenomics data (4, 52), and a genome of this size would be 4.05 fg (16), the theoretical size of the prokaryotic cell population based on the total DNA recovered ranged from 1.1 ⫻ 107 to 5.4 ⫻ 108 cells g⫺1 of wet sediment. The most probable number of Archaea in the ORFRC sediments was estimated to be ⬍102 cells g⫺1 (41). We did not include the eukaryotes in the population size estimate because they were usually associated with the sediments containing lower levels of contamination (58); however, inclusion of the eukaryotic component would reduce the size estimate of the cell population by 25% (52). Clone libraries derived from methanol-amended P5M microcosms exhibited the highest diversity, followed by clone libraries from ethanol-amended P4E and P5E and with lowest diversity occurring in P4M microcosms (Table 1; Fig. 1). Most of the clones belonged to the Alpha-, Beta-, Delta-, and Gammaproteobacteria. The remaining clones were from Firmicutes, Acidobacteria, Bacteroidetes, Actinobacteria, and other and unclassified bacteria (Table 2). The bacterial communities detected in alcohol-stimulated P4 and P5 microcosms were different from the community in the uncontaminated background (BG) site, where Gammaproteobacteria were dominant (Shi-

FIG. 2. Results of a PCoA obtained with weighted and normalized UniFrac using the 16S rRNA gene sequences isolated from the ethanol- and methanol-amended samples. The neighbor-joining tree was rooted with an archaeal outgroup. The ends of the alignment were trimmed prior to the analysis so that all of the aligned sequences were the same length. See Table 1 for descriptions of the samples.

gella, Pseudomonas), followed by Actinobacteria (Rhodococcus), Betaproteobacteria (Alcaligenes), and Alphaproteobacteria (Tables 1 and 2). Community analysis. Principal coordinates analysis (PCoA) in UniFrac revealed that 55.2% of the total variance was explained by the first two coordinate axes (Fig. 2). Factor 1 accounted for 31.38% of the variation and correlated with the origin of the samples (P4 or P5). Factor 2 explained 23.82% of the variation and correlated with the type of electron donor (ethanol or methanol). Addition of humics did not affect the community

TABLE 2. Relative contributions of 16S rRNA clones from background and ethanol- and methanol-amended samplesa Total clones (%, mean ⫾ SE) Phylum

Ethanol-amended experiments

Methanol-amended experiments

Background P5EU42

P5EU137

P4EU16

P4EU38

P5MU78

P5MhU78

P4MN10

P4MN38

Proteobacteria Alphaproteobacteria Betaproteobacteria Deltaproteobacteria Gammaproteobacteria

75.7 ⫾ 26.8 5.4 ⫾ 2.9 7.5 ⫾ 7.4 0.0 ⫾ 0.0 62.8 ⫾ 16.5

89.2 ⫾ 4.4 59.0 ⫾ 8.9 0.4 ⫾ 0.7 0.7 ⫾ 0.6 84.4 ⫾ 5.0 36.3 ⫾ 12.8 4.4 ⫾ 2.8 16.2 ⫾ 8.7 0.0 ⫾ 0.0 5.7 ⫾ 8.1

69.7 ⫾ 18.2 7.5 ⫾ 1.7 58.5 ⫾ 15.7 3.8 ⫾ 0.8 0.0 ⫾ 0.0

89.2 ⫾ 2.6 22.0 ⫾ 17.6 9.1 ⫾ 6.5 0.0 ⫾ 0.0 58.2 ⫾ 8.5

66.5 ⫾ 0.9 3.0 ⫾ 0.9 45.1 ⫾ 5.2 18.3 ⫾ 3.4 0.0 ⫾ 0.0

65.2 ⫾ 10.8 4.4 ⫾ 0.2 47.0 ⫾ 4.3 13.8 ⫾ 6.7 0.0 ⫾ 0.0

98.9 ⫾ 1.5 0.0 ⫾ 0.0 69.2 ⫾ 2.5 0.0 ⫾ 0.0 29.8 ⫾ 1.0

94.5 ⫾ 0.1 0.7 ⫾ 1.0 92.8 ⫾ 0.4 0.0 ⫾ 0.0 1.1 ⫾ 1.5

Firmicutes Acidobacteria Bacteroidetes Actinobacteria Othersb Unclassified

1.6 ⫾ 2.3 0.0 ⫾ 0.0 0.0 ⫾ 0.0 21.0 ⫾ 29.7 0.6 ⫾ 0.8 1.1 ⫾ 1.5

7.6 ⫾ 3.0 10.5 ⫾ 2.6 0.7 ⫾ 0.6 23.6 ⫾ 8.4 1.1 ⫾ 0.1 2.9 ⫾ 0.6 0.0 ⫾ 0.0 0.0 ⫾ 0.0 0.0 ⫾ 0.0 0.0 ⫾ 0.0 1.4 ⫾ 1.6 4.0 ⫾ 1.6

4.3 ⫾ 1.4 5.9 ⫾ 0.9 0.0 ⫾ 0.0 3.2 ⫾ 3.0 9.0 ⫾ 9.6 8.0 ⫾ 5.1

1.6 ⫾ 0.8 2.7 ⫾ 0.7 0.6 ⫾ 0.8 1.1 ⫾ 1.6 3.2 ⫾ 4.5 1.6 ⫾ 0.7

2.4 ⫾ 1.7 0.0 ⫾ 0.0 28.0 ⫾ 1.7 0.0 ⫾ 0.0 0.0 ⫾ 0.0 3.0 ⫾ 0.9

17.7 ⫾ 2.8 0.0 ⫾ 0.0 15.6 ⫾ 5.8 0.0 ⫾ 0.0 0.0 ⫾ 0.0 1.5 ⫾ 2.1

0.5 ⫾ 0.8 0.0 ⫾ 0.0 0.0 ⫾ 0.0 0.0 ⫾ 0.0 0.0 ⫾ 0.0 0.5 ⫾ 0.8

4.8 ⫾ 0.8 0.0 ⫾ 0.0 0.7 ⫾ 1.0 0.0 ⫾ 0.0 0.0 ⫾ 0.0 0.0 ⫾ 0.0

a b

See Table 1 for descriptions of the experimental labels. “Others” were from the phyla Chloroflexi, Verrucomicrobia, Gemmatimonadetes, Planctomycetes, and Nitrospira and OD1 and OP10.

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FIG. 3. Bacterial composition of the clone libraries based on the Naïve Bayesian rRNA classifier. The Betaproteobacteria were divided into five groups: BCO, Burkholderiaceae, Comamonadaceae, and Oxalobacteraceae; Un, unclassified, which had a ⬍80% confidence bootstrap value to any known family within the phylum; R, Rhodocyclales; N, Neisseriales; NM, Nitrosomonadales and Methylophilales. The Gammaproteobacteria were divided into two groups: X, Xanthomonadales; EP, Enterobacteriales and Pseudomonadales. The “other” group includes the phyla Chloroflexi, Verrucomicrobia, Gemmatimonadetes, Planctomycetes, and Nitrospira and OD1, and OP10. Clones that could not be assigned with the ⬎80% confidence bootstrap value were labeled as “bacteria unclassified.”

composition (P ⫽ 0.16), as indicated by the UniFrac significance test and the proximity of the P5MhU77 and P5MU77 samples in the PCoA plot (Fig. 2). Site effects. The abundance levels of Beta- and Deltaproteobacteria were greater than Alpha- and Gammaproteobacteria in the P5 clone libraries. In the P4 clone libraries of Alpha-, Beta-, and Gammaproteobacteria were abundant, while Deltaproteobacteria were present at lower percentages. Actinobacteria and others were detected in some of the P4 samples but not in any of the P5 samples (Table 2). Microcosms from both sites contained Betaproteobacteria (Burkholderiales and Rhodocyclales) and Deltaproteobacteria. However, there were site-specific differences in the communities at a lower phylogenetic level. The major genera were Acidovorax, Diaphorobacter, Azoarcus, and Geobacter in P5EU and Ottowia, Dechloromonas, and Myxococcales in P4EU. Sequences with 92% identity to both Geobacter sp. FRC-32 and Geobacter thiogenes (formerly Trichlorobacter thiogenes [13, 46]) were recovered from the P5EU, P4EU, and P5MU microcosms. Sequences related to other Geobacter species (e.g., Geobacter metallireducens, Geobacter uraniireducens) were found in the P5EU and P5MU micro-

cosms, but sequences related to Myxococcales appeared only in P4MU. In the case of Acidobacteria, Geothrix spp. were present in P5EU but Acidobacteriaceae Gp[6/3/7/16] was only detected in P4EU. Amendment effects. The reduction of nitrate and sulfate in ethanol-amended microcosms from both sites was faster than in methanol-amended microcosms. The complete nitrate reduction was observed by day 8 (P5EU), 14 (P5MU, P5MhU, and P4EU), or 16 (P4MN) (42; also unpublished data). Sulfate reduction was slower, with final values at day 31 in ethanolamended microcosms and at day 42 in methanol-amended microcosms. The methanol-amended microcosms from P5 (P5MU and P5MhU) showed U(VI) reduction, but the P4 methanol-amended microcosm (P4MN) did not. The P4MN samples had a higher percentage of Proteobacteria than P5M, whereas the abundances of Firmicutes and Bacteroidetes were greater in P5M than in P4MN samples (Table 2; Fig. 3). Methanol added at the start was slowly consumed and was measured at a high level of 12.12 to 13.28 mM through day 12 (Table 3) in both P5MU and P5MhU microcosms, which were dominated by Betaproteobacteria (mostly Rhodoferax and Polaromo-

TABLE 3. Electron donors and acceptors measured in the experiments Final concn (mM) for indicated microcosma Incubation (days)

0 4 6 10 12 38 42 65 a b c

P4EUb

P5EU

P5MU

P5MhU

P4MN

Acetate

Methanol

Ethanol

Acetate

Ethanol

Methanol

Methanol

Butyrate

0 NDc 0.21 ND 2.29 ND 0 0

0 ND 0 ND 3.17 ND 3.04 0

8.75 ND 3.51 ND 1.87 ND 0 0

0 0 0 0.66 ND 2.32 ND ND

2.52 1.68 0.47 0.66 ND 0 ND ND

13.39 ND 15.22 ND 12.12 ND 0 0

12.79 ND 13.86 ND 13.28 ND 0 0

0.21 0.16 0.17 0 ND 0 ND ND

P5EU, P5 ethanol; P4EU, P4 ethanol; P5MU, P5 methanol; P5MhU, P5 methanol plus humics; P4MN, P4 methanol-amended microcosm. Methanol was not measured in the P4EU and P4MN experiments. ND, not done.

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nas), followed by Bacteroidetes (Roseivirga-like), Deltaproteobacteria (Geobacter), Firmicutes (Desulfosporosinus), and Alphaproteobacteria (Brevundimonas) (Table 2; Fig. 3). The majority of Betaproteobacteria clones in P5MU and P5MhU were 97 to 99% similar to Rhodoferax ferrireducens, a facultatively anaerobic iron-reducing bacterium. The methanolamended P5 microcosms showed 93% U(VI) reduction in 90 days with a high reduction rate of 0.95 ␮mol liter⫺1 day⫺1 (42). The addition of humics did not change the U(VI) reduction rate but had a positive influence on the enrichment of Firmicutes (Clostridium) and diversity of Betaproteobacteria (Janthinobacterium, Methylophilales, and unclassified) and a negative influence on the enrichment of Bacteroidetes (Table 2; Fig. 3). The diversity in the methanol-amended P4MN microcosms was low, with two dominant classes, Beta- and Gammaproteobacteria (Table 2), and methanol was not measured (Table 3). Based on the RDP classifier (80% confidence threshold), the majority of the Betaproteobacteria sequences from P4MN microcosms were identified as environmental clone sequences. The bulk of these sequences grouped together phylogenetically, and the closest cultured relatives from GenBank were from the family Methylophilaceae, which are obligate methanol-utilizing heterotrophs that are not known to reduce Fe(III) and U(VI). Other Proteobacteria were represented by Pseudomonas, Enterobacteriaceae, and single clones in the family Burkholderiales. Clones found in low abundance were affiliated with Firmicutes (Desulfosporosinus, Clostridium), Alphaproteobacteria (Rhizobiaceae), Bacteroidetes, and unclassified bacteria (Table 2; Fig. 3). Ethanol stimulated an enrichment of the same bacterial classes in samples from the two sites (P4E and P5E) (Table 2; Fig. 3). Proteobacteria represented by the Alpha-, Beta-, Delta-, and Gammaproteobacteria classes constituted 59.0 to 89.2% of the total clones. The remaining clones were affiliated with Firmicutes, Acidobacteria, Bacteroidetes, Actinobacteria, and others and unclassified bacteria (Table 2; Fig. 3). The most noticeable difference between amendments was the presence of Acidobacteria in the ethanol-amended microcosms (Table 2). Ethanol-amended P4 or P5 samples showed U(VI) reduction rates of 1.0 and 0.8 ␮mol liter⫺1 day⫺1, respectively (42). Incubation time effects. One factor which could affect bacterial community structure is incubation time. The UniFrac significance test indicated a significant difference (P ⫽ 0.02) in community composition at 137 (P5EU137) versus 42 (P5EU42) days. Most of the ethanol added at the start of the P5 experiment was utilized by day 42, with acetate (2.29 mM) and methanol (3.17 mM) as the primary products at day 12 (Table 3). Acetate was consumed by day 42 and methanol by day 65 (Table 3). The aerophilic Betaproteobacteria Neisseriales (94% identity to Vogesella indigofera) was detected at high abundance at day 42 but was almost completely replaced by obligate anaerobes Acidobacteria (Geothrix), Gammaproteobacteria (Enterobacteriaceae), and Deltaproteobacteria (Geobacter) by day 137 (Fig. 3). The ethanol-amended P4E bacterial community was significantly affected by incubation time (P ⬍ 0.001) based on the UniFrac significance test. Similar to the P5E experiment, most of the initial ethanol in P4EU was utilized before day 10 and was completely depleted by day 38, with acetate at about 2.32 mM as the primary product (Table 3). Samples showed a


decrease in bacterial diversity at 38 days (P4EU38) compared to 16 days (P4EU16) of incubation (Table 1; Fig. 1). A shift in the Proteobacteria community, characterized by an increased abundance of Alpha- and Gammaproteobacteria and a decrease in Beta- and Deltaproteobacteria, was observed (Table 2; Fig. 3). The P4 samples amended with methanol (P4MN10 and P4MN38) exhibited the lowest phylogenetic diversities (Fig. 1). Butyrate (about 0.2 mM) was detected at day 0, day 3, and day 6 of the P4MN incubation but was completely consumed by day 10 (Table 3). No other metabolites (formate or acetate) were detected in these samples, and methanol was not measured. The UniFrac significance test (P ⫽ 0.43) indicated that there was no significant difference in community structure at 38 versus 10 days. DISCUSSION The Deltaproteobacteria have often been detected in conjunction with U(VI) reduction at the ORFRC. Following in situ biostimulation of U(VI) reduction, North et al. (47) observed an increase from 5% to 40% of Deltaproteobacteria 16S rRNA gene sequences, with a significant rise in Geobacter-type sequences. After biostimulation Amos et al. (2) found an enriched Geobacter lovleyi SZ, which is known to reduce U(VI). In our study, sequences related to Geobacter spp. were recovered only from U(VI)-reducing microcosms. The observed increase in Geobacter clones in the ethanol-amended microcosms agreed with previous studies that showed an increase in Geobacter spp. in uranium-contaminated subsurface sediments amended with ethanol in the field (47) or in microcosms (44). Geobacter species can use ethanol either as a carbon source or as an electron donor when metals [e.g., Fe(III), Mn(IV) or (III), Cr(VI), and U(VI)] are available as electron acceptors (36, 59). Geobacter spp. were also linked to U(VI) reduction at field sites where acetate was added as the electron donor (3, 9, 27). While Geobacter species in pure cultures do not metabolize methanol directly (12, 62), they may process acetate produced by anaerobic enrichment cultures (42). Acetate production was likely at levels that were rapidly consumed during the experiment, as it was not detected in the methanol-amended samples. Anaeromyxobacter, for which U(VI) reduction supports growth (55), has also been detected along with U(VI)reducing activity (8, 47, 51). U(VI) reduction by Anaeromyxobacter spp. requires H2 as an electron donor, which cannot be replaced by acetate (71). Thus, acetate and H2 may serve as electron donors for U(VI) reduction by species of Geobacter and Amaeromyxobacter (55, 71). In the results reported here, Firmicutes clones (Clostridium and Desulfosporosinus) were recovered from all microcosms, with higher abundances found in the microcosms exhibiting U(VI) reduction. Both Clostridium spp. and Desulfosporosinus spp. are capable of U(VI) reduction, and Desulfosporosinus can also reduce sulfate (20, 63). Some Firmicutes, such as Clostridium formicoaceticum (15), can directly utilize ethanol and methanol for acetogenesis, and the reductive synthesis of acetate could be the preferred electron-accepting process during the early stages of incubation (14, 30). In the ethanol-amended microcosms, we observed the accumulation of acetate and concurrent disappearance of ethanol. Although acetate is not the only intermediate of ethanol oxidation (56), it was the primary

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product we could detect. Propionate and other volatile fatty acids can be produced in the presence of ethanol and sulfate, but they may not accumulate at a high level (31). The presence of butyrate in methanol-amended microcosms was consistent with the presence of Clostridium clones. Clostridium species (e.g., C. methylotrophicum) can metabolize methanol and form butyrate, which can be further oxidized to acetate and H2 (54). The order Burkholderiales was detected with a higher relative contribution in the U(VI)-reducing microcosms. In methanol-amended P5M microcosms the majority of clones were affiliated with Rhodoferax ferrireducens, which is unable to utilize methanol but can oxidize acetate with the reduction of Fe(III) (19). Betaproteobacteria, such as Acidovorax, Diaphorobacter, and Janthinobacterium, can contribute to U(VI) reduction (7, 8, 48), and they were detected in ethanol-amended microcosms. Clones related to Rhodocyclales (Azoarcus and Dechloromonas) were abundant in the ethanol-amended samples. These anaerobic bacteria can utilize ethanol under denitrifying conditions and contribute to metal reduction (1, 53). Neisseriales, with Vogesella indigofera as a major component, were also found in high abundance in P5E. V. indigofera is capable of denitrification and can use ethanol as a carbon source under aerobic conditions (22). Under anaerobic conditions, members of the Neisseriales can oxidize Fe(II) coupled to nitrate reduction (69, 70). Anaerobic nitrate-dependent microbial Fe(II) oxidation is characterized by rapid removal of heavy metals and radionuclides, including U(VI) (32). Diverse Acidobacteria sequences were detected in uranium-contaminated subsurface sediments from DOE sites in Tennessee and Colorado (5, 8); however, the metabolic capabilities and the ecological role these bacteria play in contaminated sites are still unknown. Geothrix fermentans, a culturable species within Acidobacteria, is a strictly anaerobic Fe(III)-reducing bacterium that does not use ethanol as a carbon source but does utilize acetate (11). Geothrix spp. found in ORFRC sediments could contribute to indirect U(VI) reduction through the production of bioreduced Fe(II) (72). It is still unknown if Acidobacteria Gp3 capable of ethanol- and methanol-induced growth can reduce Fe(III) or U(VI) (50). Other bacteria that may contribute to U(VI) reduction are Alphaproteobacteria (Rhizobiales), Gammaproteobacteria (Enterobacteriales, Pseudomonadales, and Xanthomonadales), Actinobacteria, and Bacteroidetes. Similar clones or isolates have been previously detected in the uranium-contaminated ethanol-biostimulated sites at the ORFRC (1, 18, 60). The increase of biodiversity upon addition of humics may be a result of an excess of recalcitrant organic compounds that were available as a nutrient source or as a terminal electron acceptor (34). Microbially reduced humics can donate electrons to Fe(III), U(VI), Hg(II), and Cr(VI), thereby increasing the reduction rates (24, 34). This process has been demonstrated for Geobacter metallireducens, Shewanella alga, and Geothrix, and Wolinella species (10, 35). Methanol addition does not always lead to U(VI) reduction, but if it occurs it results in nearly complete reduction (49). The methanol-amended microcosms with U(VI) reduction yielded diverse bacterial populations, while communities in nonU(VI)-reducing microcosms were dominated by obligate methylotrophs of Betaproteobacteria (Methylophilaceae) and

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potential methanol-consuming Gammaproteobacteria (Pseudomonas and Enterobacteriaceae) (6). Certain obligately methylotrophic bacteria are capable of methanol oxidation under denitrifying conditions due to the presence of a unique electron transfer chain involving both processes (26). Use of the supplied amendment (e.g., methanol) as an electron donor or as a carbon source may determine the structure of a bacterial community. This study showed that different microbial communities are associated with the type of electron donor added to the system, duration of incubation, and origin of the sediments. Reduction of U(VI) can be accomplished directly by bacteria such as Deltaproteobacteria (e.g., Geobacter spp. and Desulfovibrio spp.), abiotically by bioreduced products [e.g., sulfide produced by Desulfosporosinus spp. and Fe(III) reduction by Geothrix spp.], or by both. The metabolic diversity of the natural microbial community is sufficiently large and redundant for the bioreduction of U(VI) to occur, regardless of the type of electron donor added. ACKNOWLEDGMENTS This work was funded by the U.S. Department of Energy’s Office of Science Biological and Environmental Research, Environmental Remediation Sciences Program. Oak Ridge National Laboratory is managed by UT-Battelle, LLC, for the U.S. Department of Energy under contract DE-AC05-00OR22725. We thank Marilyn Kerley for help with Sanger sequencing, Dave Watson and Lisa Fagan for help with sampling, and Tommy Phelps for useful discussions. REFERENCES 1. Akob, D. M., H. J. Mills, T. M. Gihring, L. Kerkhof, J. W. Stucki, A. S. Anastacio, K. J. Chin, K. Kusel, A. V. Palumbo, D. B. Watson, and J. E. Kostka. 2008. Functional diversity and electron donor dependence of microbial populations capable of U(VI) reduction in radionuclide-contaminated subsurface sediments. Appl. Environ. Microbiol. 74:3159–3170. 2. Amos, B. K., Y. Sung, K. E. Fletcher, T. J. Gentry, W. M. Wu, C. S. Criddle, J. Zhou, and F. E. Loffler. 2007. Detection and quantification of Geobacter lovleyi strain SZ: implications for bioremediation at tetrachloroethene- and uranium-impacted sites. Appl. Environ. Microbiol. 73:6898–6904. 3. Anderson, R. T., H. A. Vrionis, I. Ortiz-Bernad, C. T. Resch, P. E. Long, R. Dayvault, K. Karp, S. Marutzky, D. R. Metzler, A. Peacock, D. C. White, M. Lowe, and D. R. Lovley. 2003. Stimulating the in situ activity of Geobacter species to remove uranium from the groundwater of a uranium-contaminated aquifer. Appl. Environ. Microbiol. 69:5884–5891. 4. Angly, F. E., D. Willner, A. Prieto-Davo ´, R. A. Edwards, R. Schmieder, R. Vega-Thurber, D. A. Antonopoulos, K. Barott, M. T. Cottrell, C. Desnues, E. A. Dinsdale, M. Furlan, M. Haynes, M. R. Henn, Y. Hu, D. L. Kirchman, T. McDole, J. D. McPherson, F. Meyer, R. M. Miller, E. Mundt, R. K. Naviaux, B. Rodriguez-Mueller, R. Stevens, L. Wegley, L. Zhang, B. Zhu, and F. Rohwer. 2009. The GAAS metagenomic tool and Its estimations of viral and microbial average genome size in four major biomes. PLoS Comput. Biol. 5:e1000593. 5. Barns, S. M., E. C. Cain, L. Sommerville, and C. R. Kuske. 2007. Acidobactetia phylum sequences in uranium-contaminated subsurface sediments greatly expand the known diversity within the phylum. Appl. Environ. Microbiol. 73:3113–3116. 6. Bellion, E., M. E. Kent, J. C. Aud, M. Y. Alikhan, and J. A. Bolbot. 1983. Uptake of methylamine and methanol by Pseudomonas sp. strain Am1. J. Bacteriol. 154:1168–1173. 7. Byrne-Bailey, K. G., K. A. Weber, A. H. Chair, S. Bose, T. Knox, T. L. Spanbauer, O. Chertkov, and J. D. Coates. 2010. Completed genome sequence of the anaerobic iron-oxidizing bacterium Acidovorax ebreus strain TPSY. J. Bacteriol. 192:1475–1476. 8. Cardenas, E., W. M. Wu, M. B. Leigh, J. Carley, S. Carroll, T. Gentry, J. Luo, D. Watson, B. Gu, M. Ginder-Vogel, P. K. Kitanidis, P. M. Jardine, J. Zhou, C. S. Criddle, T. L. Marsh, and J. A. Tiedje. 2008. Microbial communities in contaminated sediments, associated with bioremediation of uranium to submicromolar levels. Appl. Environ. Microbiol. 74:3718–3729. 9. Chang, Y. J., P. E. Long, R. Geyer, A. D. Peacock, C. T. Resch, K. Sublette, S. Pfiffner, A. Smithgall, R. T. Anderson, H. A. Vrionis, J. R. Stephen, R. Dayvault, I. Ortiz-Bernad, D. R. Lovley, and D. C. White. 2005. Microbial

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