AEM
Home Help [Feedback] [For Subscribers] [Archive] [Search] [Contents]
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Supplemental material
Right arrow Other Versions of this Article:
AEM.01194-07v1
74/4/1019    most recent
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowReprints and Permissions
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Blöthe, M.
Right arrow Articles by Küsel, K.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Blöthe, M.
Right arrow Articles by Küsel, K.
Agricola
Right arrow Articles by Blöthe, M.
Right arrow Articles by Küsel, K.

 Previous Article  |  Next Article 

Applied and Environmental Microbiology, February 2008, p. 1019-1029, Vol. 74, No. 4
0099-2240/08/$08.00+0     doi:10.1128/AEM.01194-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

pH Gradient-Induced Heterogeneity of Fe(III)-Reducing Microorganisms in Coal Mining-Associated Lake Sediments{triangledown} ,{dagger}

Marco Blöthe,1,{ddagger} Denise M. Akob,2 Joel E. Kostka,2 Kathrin Göschel,1 Harold L. Drake,1 and Kirsten Küsel1,3*

Department of Ecological Microbiology, University of Bayreuth, 95440 Bayreuth, Germany,1 Department of Oceanography, Florida State University, Tallahassee, Florida 32306,2 Limnology Research Group, Friedrich Schiller University Jena, 07743 Jena, Germany3

Received 29 May 2007/ Accepted 29 November 2007


   ABSTRACT
Top
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Lakes formed because of coal mining are characterized by low pH and high concentrations of Fe(II) and sulfate. The anoxic sediment is often separated into an upper acidic zone (pH 3; zone I) with large amounts of reactive iron and a deeper slightly acidic zone (pH 5.5; zone III) with smaller amounts of iron. In this study, the impact of pH on the Fe(III)-reducing activities in both of these sediment zones was investigated, and molecular analyses that elucidated the sediment microbial diversity were performed. Fe(II) was formed in zone I and III sediment microcosms at rates that were approximately 710 and 895 nmol cm–3 day–1, respectively. A shift to pH 5.3 conditions increased Fe(II) formation in zone I by a factor of 2. A shift to pH 3 conditions inhibited Fe(II) formation in zone III. Clone libraries revealed that the majority of the clones from both zones (approximately 44%) belonged to the Acidobacteria phylum. Since moderately acidophilic Acidobacteria species have the ability to oxidize Fe(II) and since Acidobacterium capsulatum reduced Fe oxides at pHs ranging from 2 to 5, this group appeared to be involved in the cycling of iron. PCR products specific for species related to Acidiphilium revealed that there were higher numbers of phylotypes related to cultured Acidiphilium or Acidisphaera species in zone III than in zone I. From the PCR products obtained for bioleaching-associated bacteria, only one phylotype with a level of similarity to Acidithiobacillus ferrooxidans of 99% was obtained. Using primer sets specific for Geobacteraceae, PCR products were obtained in higher DNA dilutions from zone III than from zone I. Phylogenetic analysis of clone libraries obtained from Fe(III)-reducing enrichment cultures grown at pH 5.5 revealed that the majority of clones were closely related to members of the Betaproteobacteria, primarily species of Thiomonas. Our results demonstrated that the upper acidic sediment was inhabited by acidophiles or moderate acidophiles which can also reduce Fe(III) under slightly acidic conditions. The majority of Fe(III) reducers inhabiting the slightly acidic sediment had only minor capacities to be active under acidic conditions.


   INTRODUCTION
Top
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Very phylogenetically diverse prokaryotes are capable of dissimilatory Fe(III) reduction or respiratory growth with Fe(III) as the sole electron acceptor (49, 50). The thermodynamic energy available from Fe(III) reduction varies based on a variety of parameters, including the Fe(III) form, crystallinity, and pH. The energetics of Fe(III) reduction under neutral-pH conditions differ substantially from those under acidic conditions (72). While Fe(III) exists predominantly in the solid phase as oxyhydroxide minerals at circumneutral pH, Fe(III) is more soluble and should be an electron acceptor for microbial growth under low-pH conditions. Reduction of both solid and soluble forms of Fe(III) becomes more thermodynamically favorable with decreasing pH. Since proton concentrations select for acidophilic microorganisms at low pH, it is not surprising that Fe(III) reduction in neutral-pH and acidic environments is carried out by different microbial populations (50, 72). During the last two decades, Fe(III) reduction in neutral-pH environments and neutrophilic Fe(III)-reducing microorganisms have been studied intensively (13, 14, 47, 49, 51). However, the process in acidic environments has received little attention (26, 34, 38, 41, 60).

Some acidophilic bacteria have the capacity to reduce Fe(III) (39, 63). The chemolithoautotrophs Acidithiobacillus thiooxidans and Acidithiobacillus ferrooxidans can couple the anaerobic oxidation of elemental sulfur to the reduction of Fe(III) (11, 20, 62). Heterotrophic acidophiles belonging to the genus Acidiphilium can also catalyze the reduction of Fe(III) and appear to be widely distributed in metal-rich acidic environments (30, 31, 32, 39, 41, 42). In anoxic environments these organisms may contribute to iron cycling by redissolving Fe(III) minerals which precipitate when the pH is increased by mixing with groundwater or surface water. However, the rate and extent of Fe(III) reduction vary significantly in the isolates that have been examined (30). Acidiphilium cryptum JF-5 was the first acidophile isolated under Fe(III)-reducing conditions at pH 3 from an acidic coal mining-associated lake (41). Coal mining-associated lakes receive high concentrations of Fe(II), protons, and sulfate due to the oxidation of pyrite in the mine tailings (27, 59). Subsequently, Fe(II) is oxidized biologically (e.g., by A. ferrooxidans) and precipitates as poorly crystalline Fe(III) oxyhydroxysulfate to the sediment (59). Whereas most studies have shown that the optimum pH for growth of Acidiphilium is 3 to 4, A. cryptum ATCC 33463 was recently shown to reduce small amounts of solid-phase Fe(III) at pH 5 (8). Since most cultivated Fe(III)-reducing prokaryotes are neutrophilic and have a negligible capacity to reduce Fe(III) at acidic pHs (pH < 5.5), we have only a marginal understanding of the microorganisms that drive the reduction of Fe(III) under moderately acidic conditions. Thus, the objectives of this study were to (i) explore the microbial diversity of coal mining-associated lake sediments, (ii) investigate the impact of pH on Fe(III)-reducing potentials, and (iii) elucidate the Fe(III)-reducing microbial community in acidic (pH 3) and slightly acidic (pH 5) sediment zones.


   MATERIALS AND METHODS
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Field site description and sampling.
The acidic coal mining-associated lake that we studied (Lake 77) is located in the Lusatian mining area in east central Germany. The pH of the lake water and the maximum summer temperature of the upper sediment were approximately 3 and 12°C, respectively (59). Sediment cores were obtained using a boat and a gravity corer in Plexiglas tubes (inside diameter, 5.9 cm), always at the deepest location (7 m). On each sampling date (November 2001, July 2003, November 2003, and February 2004) 5 to 12 cores were obtained, transported to the laboratory, and sectioned based on sediment stratification under an N2 atmosphere within 24 h. Sediments were fairly uniform horizontally in the area of the lake examined.

Visually and geochemically, the sediment could be sectioned into four different zones. The upper orange sediment zone (zone I, 0 to 7 cm) had a pH of 2.8 to 3.1 and was enriched with reactive iron, mainly schwertmannite [Fe8O8(OH)x(SO4)y] (40, 59). In the next zone, a yellowish-brownish sediment zone (zone II, 7 to 10 cm), the pH varied from 3.4 to 4.7. The third zone (zone III, 10 to 16 cm) had a pH of 5.2 to 5.7 and was enriched in goethite ({alpha}-FeOOH), which is consistent with the fact that schwertmannite is solved or transformed in more stable minerals, such as goethite, under changing physicochemical conditions (7). Below 16 cm the sediment pH ranged from 5.3 to 6.1. Only small amounts of reactive iron were present in this zone, zone IV; thus, we focused on the upper three zones. The sediment dry weights were approximately 8.2, 14.6, 22.4, and 36.3% of the wet weights in zones I, II, III, and IV, respectively. More detailed geochemical descriptions of the sediment have been given elsewhere (40, 59).

Preparation of sediment microcosms.
For zones I and III, sediment from replicate cores was pooled under anoxic conditions, and 40 g (wet weight) of sediment was transferred to sterile 150-ml infusion bottles (Merck ABS, Dietikon, Switzerland) inside a Mecaplex O2-free chamber (100% N2 gas phase). The bottles were closed with rubber stoppers and screw-cap seals, flushed with sterile argon for 15 min, and incubated in the dark at 15°C with an initial overpressure of 20 to 25 kPa argon at room temperature. No electron donors were added. The pH was adjusted with sterile solutions of 10 N HCl and 10 N NaOH. Three replicates of all microcosms were prepared. Samples were removed with sterile, argon-flushed syringes. Rates of Fe(II) formation were calculated only for the time period when a linear increase in the Fe(II) concentration was observed.

Media and enumeration studies.
One representative sediment core obtained in November 2001 was sectioned to obtain 1.5-cm sediment depth zones under an N2 atmosphere. Subsamples were taken from every 1.5-cm sediment depth zone with sterile syringes to determine the pH and sulfate and Fe(II) contents of the pore water. One milliliter of wet sediment was transferred into a sterile 50-ml polycarbonate tube, diluted fourfold with sterile filtered 4% paraformaldehyde, and incubated for 2 h at 4°C. Fixed samples were diluted 500-fold in sterile filtered 1x phosphate-buffered saline (130 mM NaCl, 10 mM NaPi; pH 7.4), homogenized, and incubated for 15 min in an ultrasonic water bath. Five microliters of 4',6-diamidino-2-phenylindole (DAPI) (100 µg ml–1) and 0.5 ml of 1x phosphate-buffered saline were added to the fixed samples, which were then incubated for 15 min on ice. Stained samples were filtered on black polycarbonate filters (Millipore GTBP 047000) and washed with 10 to 20 ml of filter-sterilized distilled H2O. For counting, each filter was covered with Citifluor, and cells were examined using an epifluorescence microscope (Zeiss Axioskop) with a high-pressure mercury bulb (50 W) and 02 filter set (G365, FT395, LP420; Zeiss).

For pure-culture studies, A. ferrooxidans DSM 583 and Acidobacterium capsulatum DSM 11244 were used. Cells of A. ferrooxidans were cultivated in Thio medium (62) under a CO2 gas phase. Autoclaved, washed flowable sulfur (Stoller Chemical Company, Inc., Houston, TX) was added to a final 10 mM concentration. Cells of A. capsulatum were cultivated in Acido medium that contained (per liter) 2.0 g of (NH4)2SO4, 0.5 g of KH2PO4, 0.5 g of MgSO4·7H2O, and 0.1 g of KCl. The pH was adjusted with NaOH to obtain a final pH between 2.2 and 5. The incubation temperature was 30°C. A. ferrooxidans and A. capsulatum cultures were grown first under oxic conditions with FeSO4 (10 mM) and glucose (2 mM) as the electron donors, respectively. After 2 weeks, cells were centrifuged, washed three times, and transferred to anoxic media to obtain a final optical density at 660 nm of 0.3 to 0.4. The optical density at 660 nm was determined with a Spectronic 501 photometer (Bausch & Lomb Inc., Rochester, NY). Soluble Fe(III) was added as ferric sulfate [Fe(III)2(SO4)3] from a sterile, anoxic stock solution. Goethite, schwertmannnite, and FeOOH were prepared as previously described (43) and added at a final concentration of 40 mM.

Numbers of cultured Fe(III) reducers were determined by the most-probable-number (MPN) technique using a 10-fold dilution series with three replicate tubes per dilution and incubation at 15°C in selective media. Zone III sediment (pH 5.6) from two cores obtained in November 2003 was sectioned and pooled under an N2 atmosphere. Five grams (wet weight) was transferred to a bottle containing 95 ml of dilution buffer (43) and some 0.2-mm-diameter glass beads. The bottle was sealed and mixed using an end-over-end shaker for 1 h. Dilution series were prepared, and media were inoculated using the dilutions. The medium contained (per liter) 40 mM amorphous ferric hydroxide [Fe(OH)3] (51), 2.0 g of (NH4)2SO4, 0.5 g of K2HPO4, 0.5 g of MgSO4, 0.1 g of KCl, 5 ml of a trace metal solution (23), and 5 ml of a vitamin solution (23). The pH of each medium was adjusted to pH 5.5 with 0.5 N HCl or 0.1 M NaOH. The gas phase was N2-CO2 (80:20, vol/vol). Either ethanol (10 mM), lactate (5 mM), or H2 (10 ml) in combination with succinate as a carbon source (1 mM) was added as an electron donor. Tubes containing H2 were incubated horizontally. Tubes were considered positive based on the formation of Fe(II) and the consumption of electron donors compared to uninoculated controls. MPN values were calculated from standard MPN tables and were within 95% certainty (3).

Analytical techniques.
The reduction of Fe(III) was determined by determining the amount of Fe(II) formed after acid extraction. Aliquots (0.2 ml) of the medium or the sediment suspension were removed with sterile syringes, transferred to 9.8 ml of 0.5 N HCl, and then incubated for 1 h at room temperature (41). The Fe(II) formed was measured after addition of acetate by the phenanthroline method (73). The pH was determined with an Ingold U457-S7/110 combination pH electrode (Ingold-Meβtechnik, Switzerland). Short-chain aliphatic acids and alcohols were measured with Hewlett-Packard 1090 series II high-performance liquid chromatographs (44). Headspace gases (H2 and CO2) were measured with 5890 series II gas chromatographs (Hewlett-Packard Co., Palo Alto, CA) (43).

DNA extraction.
DNA was extracted from 10 g of sediment from zones I, II, and III by first adjusting the pH to 9 with sterile 5 N NaOH and incubating the preparations overnight at 4°C in 100 ml of sterile distilled H2O to dissolve humic substances. After incubation, the sediment was centrifuged at 8,000 x g for 20 min, the supernatant was discarded, and the pellet was resuspended in 4 ml of sterile extraction buffer (100 mM EDTA, 10 mM Tris [pH 8.0], 1% sodium dodecyl sulfate). Samples were incubated at 70°C for 30 min with occasional mixing and then centrifuged at 10,000 x g for 15 min. Each supernatant was transferred to a clean centrifuge tube, and each pellet was washed once with 10 ml of extraction buffer. The supernatant was retained and pooled with the supernatant resulting from the previous centrifugation. Nucleic acids were precipitated overnight at –20°C with isopropanol and centrifuged to pellet nucleic acids. The solution was centrifuged again. Potassium acetate was added to the pellet to a final concentration of 0.5 M, and the solution was incubated on ice for 2 h and centrifuged at 5,000 x g for 5 min. The DNA yield was determined by gel electrophoresis on 1.5% agarose gels. All gels were stained for 20 min with SYBR gold (Molecular Probes, Hamburg, Germany) and documented using a scanner (Storm 860 molecular imager; Molecular Dynamics, Sunnyvale, CA) and the ImageQuant 5.0 software (Molecular Dynamics, Sunnyvale, CA). DNA was extracted from MPN culture cell pellets using an Ultra Clean soil DNA kit according to the manufacturer's instructions (Mo Bio Laboratories, Solana Beach, CA).

PCR amplification of 16S rRNA genes.
Aliquots of DNA from zone I, II, and III sediment samples and MPN cultures were PCR amplified using Bacteria domain-specific (57) and group-specific (14, 21, 22, 25, 28, 70) 16S rRNA gene primers (see Table S1 in the supplemental material). DNA extracts from zone I and III sediment samples were PCR amplified with primers GM3 and GM4 in a reaction mixture consisting of 20 pmol of each primer, 200 µM deoxynucleoside triphosphates, 300 µg of bovine serum albumin, 1x PCR buffer, and 1 U of Taq DNA polymerase (Eppendorf, Hamburg, Germany). Template DNA (4 µl) was preheated to 70°C prior to addition to the PCR mixture. Thermocycling was performed with a T-Gradient cycler (Biometra, Göttingen, Germany), and the program consisted of 35 cycles of 95°C for 60 s, 42°C for 60 s, and 72°C for 90 s, followed by a final extension step of 72°C for 7 min. PCR amplification of DNA extracts from zone I, II, and III sediments using group-specific primers for Acidiphilium, bioleaching-associated bacteria, Geobacter, Geothrix, and Shewanella was performed as described above using an annealing temperature of 55°C. The primer set for acidophilic bioleaching-associated bacteria is specific for six bacterial phylotypes (A. ferrooxidans, A. thiooxidans, Acidithiobacillus caldus, Sulfobacillus thermosulfidooxidans, Leptospirillum ferrooxidans, A. cryptum, and Acidiphilium organovorum) which are involved in the bioleaching of mineral ores (21). DNA extracts from MPN cultures were PCR amplified using the Bacteria domain-specific primers 8F and 1392R in a mixture containing 10 to 50 ng of DNA, 1x LA PCR buffer II (TaKaRa Mirus Bio, Madison, WI), 2.5 mM deoxynucleoside triphosphates, 0.5 µM forward primer, 0.5 µM reverse primer, 2.0 µg of bovine serum albumin, and 0.03 U of LA Taq polymerase (TaKaRa Mirus Bio, Madison, WI). Thermocycling was performed as follows: incubation at 95°C for 3 min, followed by 30 cycles of 94°C for 30 s, 55°C for 30 s, and 72°C for 30 s and then a final extension step of 72°C for 10 min.

Clone library construction.
PCR amplicons produced with the Bacteria domain-specific primer pair GM3/GM4 and group-specific 16S rRNA gene primers from zone I and III sediments were cloned using the pGEM-T vector and Escherichia coli JM109 competent cells according to the manufacturer's instructions (Promega, Madison, WI). Zone I (pH 3) and zone III (pH 5.5) were selected due to the difference in pH. Clones in libraries constructed with primers GM3 and GM4 were screened by denaturing gradient gel electrophoresis (DGGE) (48) and grouped into phylotypes based on migration behavior. Clone libraries constructed with group-specific amplicons were screened by restriction fragment length (RFLP) analysis. Cloned inserts were PCR amplified with the group-specific primer pair and verified by gel electrophoresis on a 1.5% agarose gel (Roth, Darmstadt, Germany). Five microliters of PCR product was digested in a single reaction with 1 U of restriction enzymes HaeIII and CfoI (Promega, Madison, WI) at 35°C for 2 h. The digested PCR products were separated by gel electrophoresis in 7% polyacrylamide (Bio-Rad Laboratories, Hercules, CA) in 1x TAE buffer (40 mM Tris-acetate [pH 7.4], 20 mM sodium acetate, 1 mM EDTA) at 23°C at 100 V for 3.5 h with the D-Code system (Bio-Rad Laboratories, Hercules, CA).

16S rRNA gene amplicons from MPN cultures were purified using a QIAquick PCR purification kit (Qiagen, Valencia, CA) and were cloned into the TOPO TA cloning vector pCR 2.1 according to the manufacturer's instructions (Invitrogen, Carlsbad, CA). Cloned inserts were PCR amplified using the vector-specific primers M13F and M13R and were digested with restriction enzyme HaeIII (0.25 U µl–1; New England Biolabs, Beverly, MA) for 2 h at 37°C. The digested PCR products were separated by gel electrophoresis on a 3.5% MetaPhor agarose gel (Cambrex, Rockland, ME) in 1x Tris-borate-EDTA buffer at 4°C for 3 h. All clones screened using RFLP analysis were grouped into phylotypes on the basis of the RFLP banding patterns.

DGGE fingerprinting.
Community DNA extracted from zone I, II, and III sediments was fingerprinted using DGGE. DNA extracts were PCR amplified with the universal primers GM5-clamp and 907RM (56) as described above for primer pair GM3/GM4 with an annealing temperature of 55°C. PCR products were separated on a 7% polyacrylamide gel with a 40 to 60% denaturant gradient (100% denaturant was 7 M urea and 40% formamide) in 1x TAE buffer using the D-Code system (Bio-Rad Laboratories, Hercules, CA). DGGE gels were run for 18 h at 60°C and 100 V.

Dilution PCR analysis of sediment DNA extracts.
The relative levels of members of the genus Acidiphilium, the bioleaching-associated bacteria, the family Geobacteraceae, the genus Geothrix, and the genus Shewanella in sediment samples were determined using a modified MPN-PCR technique (61). In contrast to quantitative PCR, the dilution PCR method provided only rough comparative estimates for PCR products from different sediment zones amplified with the same primer sets. DNA extracts from zones I, II, and III were adjusted to a concentration of 10 µg ml–1 and serially diluted 10-fold (10° to 10–4) in sterile water. PCRs were performed as a single replicate for each dilution as described above using an annealing temperature of 55°C and 4 µl of each dilution as the template. PCR amplicons were detected by 1.5% (wt/vol) agarose gel electrophoresis using TAE buffer at 80 V for 45 min. Gels were stained with SYBR gold for 20 min and documented with the Storm 860 molecular imager. The corresponding dilution level was considered positive if a PCR product was present.

Phylogenetic and statistical analyses.
Representative clones for each RFLP phylotype were sequenced bidirectionally using a Big-Dye Terminator v3.1 cycle sequencing kit (Applied Biosystems, Foster City, CA) and an Applied Biosystems 3100 genetic analyzer with capillary electrophoresis. Sequences were assembled using Sequencher v4.5 (Gene Codes Corp., Ann Arbor, MI), and prior to phylogenetic analysis, vector sequences flanking the 16S rRNA gene inserts were removed. Previously identified sequences with high sequence similarity to the clones obtained in this study were determined using the BLAST algorithm with the GenBank database available from the National Center for Biotechnology Information (4). Clone sequences were checked for chimeras using the program CHIMERA_CHECK from Ribosomal Database Project II. All clone sequences and reference sequences were aligned with the ARB software package using the Fast Aligner algorithm, incorporating ribosomal secondary structure data. Dendrograms were constructed with the ARB software package by adding 16S rRNA sequences to the distance matrix tree using PARSIMONY_INTERAKTIV without changing the overall tree topology (52). The coverage of the clone libraries was calculated by using the equation described by Singleton et al. (69), and the sampling efficiency in clone libraries was assessed using the Analytica Rarefaction 1.3 software (http://www.uga.edu/strata/software/) originally described by Heck et al. (33).

Nucleotide sequence accession numbers.
The 16S rRNA gene sequences determined in this study have been deposited in the EMBL database under accession numbers AM712138 to AM712178 and AM713378 to AM713401.


   RESULTS
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Vertical geochemical profiles.
The pore water pH increased from 3 in the upper 7.5 cm of the sediment to 5.5 at a depth of 8 to 10 cm (Fig. 1). The concentrations of Fe(II) and sulfate increased with increasing depth from approximately 20 to 50 mM and from 8 to 15 mM, respectively. Similar profiles were obtained with other sediment cores (40, 59). Due to the distinct vertical geochemical gradients, the sediment was separated into an upper acidic zone (pH 3; zone I), a transition zone (zone II), and a slightly acidic zone (pH 5.5; zone III). The highest density of DAPI-stained cells (~1.5 x 109 cells ml–1) was detected in zone I, whereas the cell density decreased to ~0.75 x 108 cells ml–1 in zone III (Fig. 1).


Figure 1
View larger version (18K):
[in this window]
[in a new window]

  		FIG. 1. Vertical biogeochemical characterization of the sediment of acidic coal mining-associated Lake 77. One representative core obtained in November 2001 was used. (A) Symbols: {circ}, Fe(II); {blacksquare}, sulfate. (B) Symbols: {blacktriangleup}, pH; {triangleup}, cell counts. The cell count data are the means ± standard deviations for three replicate sediment samples.

 
16S rRNA gene-based community analysis.
PCR products obtained from zones I, II, and III produced different DGGE patterns (see Fig. S1 in the supplemental material). Fourteen, 11, and 13 bands were differentiated for zones I, II, and III, respectively. Only five bands appeared to have similar migration behaviors in all zones, suggesting that there were differences in the microbial community structures of the three zones.

Clone libraries of Bacteria domain-specific amplicons (obtained with GM3/GM4) contained 185 and 95 clones from zone I and III sediments, respectively. DGGE screening of the clone libraries (48) revealed the presence of 40 different phylotypes in zone I, whereas zone III contained 42 different phylotypes (data not shown). Bacterial 16S rRNA gene clone libraries derived from zone I and III sediments showed 85 and 65% coverage, respectively. No PCR product was obtained with Archaea domain-specific primers. Many of the clones (45% of the clones from zone I and 43% of the clones from zone III) belonged to the phylum Acidobacteria (Fig. 2). Other sequences detected in both zones were related to gene sequences of Nitrospira, Cytophagales, and Alpha-, Gamma-, and Deltaproteobacteria groups along with some 18S rRNA gene sequences. Sequences related to Firmicutes, Actinomyces, and Betaproteobacteria were obtained only from zone I. Sequences related to Verrumicrobia subdivision 5, Thermus/Deinococcus, Bacillus/Clostridium, and an uncultured group were detected only in zone III.


Figure 2
View larger version (55K):
[in this window]
[in a new window]

  		FIG. 2. Frequencies of bacterial phylogenetic lineages detected in 16S rRNA gene-based clone libraries derived from zones I and III. Calculations were based on the total number of clones associated with phylotypes of sequenced representatives.

 
Effect of pH on microbial formation of Fe(II) in sediments and by pure cultures.
Fe(II) was formed in zone I sediment microcosms prepared from sediment cores obtained in July 2003 at a rate of linear production that was approximately 710 nmol cm–3 day–1 (Fig. 3A). When the pH was changed from the in situ pH (pH 2.9) to 5.3, Fe(II) was formed after 5 days of incubation at a rate of 1,550 nmol cm–3 day–1. In zone III sediment microcosms at the in situ pH, Fe(II) was formed at a rate of approximately 895 nmol cm–3 day–1 (Fig. 3B). When the pH was decreased to 2.9, Fe(II) was formed after 5 days of incubation at a rate of 557 nmol cm–3 day–1. Thus, a shift to slightly acidic conditions increased Fe(II) formation in zone I sediment by a factor of 2, but a shift to acidic conditions in zone III sediment inhibited Fe(II) formation, although the soluble pool of Fe(III) that was assumed to be easily bioavailable should have been larger at pH 2.9 than at pH 5.3. Similar results were obtained with sediment cores obtained in October 2000 (data not shown).


Figure 3
View larger version (10K):
[in this window]
[in a new window]

  		FIG. 3. Effect of pH on microbe-catalyzed Fe(II) formation in zones I (A) and III (B). Sediment from replicate cores obtained in July 2003 was used. Symbols: •, Fe(II) formation at the in situ pH (pH 2.9 in zone I and pH 5.3 in zone III); {circ}, Fe(II) formation under altered pH conditions. The data are the means ± standard deviations of three replicates.

 
The rates of Fe(II) formation under in situ pH conditions calculated on a dry weight basis were approximately 8.7 and 1.9 nmol mg–1 day–1 in zones I and III, respectively, due to the smaller amount of sediment (on a dry weight basis) in zone I than in zone III. The rates of Fe(II) formation under in situ pH conditions for sediment cores obtained in November 2001 were approximately 595 and 1,405 nmol cm–3 day–1 for zones I and III, respectively, which equaled 7.3 and 3.0 nmol mg–1 day–1 on a dry weight basis (data not shown). These rates were determined in long-term incubation experiments. Fe(II) formation was linear for 45 and 58 days of incubation for zones I and III, respectively, and resulted in maximum Fe(II) concentrations of 38 and 108 mM in zones I and III, respectively.

The total amounts and rates of Fe(II) formation from schwertmannite by cultures of A. ferroxidans decreased with increasing pH (Fig. 4A). At pH 3, 4, and 5, the rates were approximately 84, 21, and 3 µmol liter–1 day–1, respectively. Similar results were obtained with FeOOH (data not shown). Small amounts of goethite were reduced at pH 3 but not at pH 5 by A. ferrooxidans. The rates of Fe(II) formation from FeOOH by cultures of A. capsulatum supplemented with 2 mM glucose under anoxic conditions were similar (approximately 57 µmol liter–1 day–1) at initial pH values of 3 and 5 (Fig. 4B). Fe(II) formation from goethite was delayed at pH 5. However, similar total amounts of Fe(II) were present at the end of incubation at pH 3 and 5. Fe(II) was also formed from soluble Fe(III)2(SO4)3 at pH 2.2. In general, the number of DAPI-stained cells did not increase during Fe(II) formation (data not shown). Glucose was fermented to acetate, ethanol, and trace amounts of succinate and lactate. The recovery of reducing equivalents theoretically obtained from the oxidation of glucose varied between 6 and 14%.


Figure 4
View larger version (16K):
[in this window]
[in a new window]

  		FIG. 4. Effect of pH on microbe-catalyzed Fe(II) formation from schwertmannite by A. ferrooxidans DSM 583 at pH 3 (•), pH 4 ({circ}), and pH 5 ({blacksquare}) with elemental sulfur (10 mM) as the electron donor (A) and by A. capsulatum DSM 11244 at pH 5 (filled symbols) and pH 3 (open symbols) from FeOOH (• and {circ}) or goethite ({blacktriangleup} and {triangleup}) or from soluble Fe(III)2(SO4)3 at pH 2.2 ({square}) with glucose (2 mM) as the electron donor (B). The data are the means ± standard deviations of three replicates.

 
Dilution PCR analysis.
Dilution PCR performed with sediment samples from zones I, II, and III yielded PCR products with 16S rRNA primers specific for acidophiles belonging to the genus Acidiphilium or the bioleaching-associated bacteria and for neutrophiles belonging to the genus Geobacter (Table 1). PCR products of acidophiles were obtained at the same dilution for all three sediment zones. For bioleaching-associated bacteria, a PCR product was obtained only with undiluted DNA. PCR products of neutrophiles with specific primer sets GM3/Geo825R and Gb564/Gb1290 could be obtained at higher dilutions (10–2) with zone III sediments. Use of Gb564/Gb1290 also yielded a PCR product at the 10–2 dilution for zone II sediments.


View this table:
[in this window]
[in a new window]

  		TABLE 1. Dilution PCR products obtained with specific primer sets for Acidiphilium, bioleaching bacteria, Geothrix, Shewanella, and Geobacter from DNA extracted from sediment zones I, II, and III

 
When a primer set specific for Geothrix was used, a PCR product was obtained only with zone II sediment. No PCR products were obtained with a primer set specific for Shewanella. A PCR product was obtained with the undiluted DNA only when nested PCR was performed after a first cycle with universal bacterial primer set GM3/GM4.

RFLP and comparative sequence analysis.
A total of 50 Acidiphilium 16S rRNA gene clones (22 clones from zone I and 28 clones from zone III) were screened by RFLP analysis, and eight different phylotypes were differentiated. Four of these eight phylotypes were present in zones I and III, while four phylotypes were present only in zone III. Comparative sequence analyses indicated that the four phylotypes present in both zones had 99% similarity at the nucleotide level to cultured Acidiphilium species. The remaining four sequences from zone III were 95% similar to cultured Acidiphilium or Acidisphaera species sequences. Comparative phylogenetic analysis of the sequences revealed two distinct groups (Fig. 5). For the bioleaching-associated bacteria, 20 clones from zone I and 28 clones from zone III were screened. Although the primer set used for bioleaching-associated bacteria obtains sequences specific for A. thiooxidans, A. caldus, S. thermosulfidooxidaans, L. ferrooxidans, and other groups (21), only one phylotype with 99% similarity to A. ferrooxidans was obtained. Nonetheless, other mesophilic bacteria potentially involved in bioleaching might be not amplified with this specific primer set.


Figure 5
View larger version (17K):
[in this window]
[in a new window]

  		FIG. 5. Phylogenetic tree of Acidiphilium-related 16S rRNA gene sequences (indicated by boldface type) determined by phylogenetic distance analysis with a maximum likelihood algorithm.

 
When primer pair Gb564/Gb1290 specific for Geobacter was used, screening yielded 38 and 52 phylotypes from zones I and III, respectively. Twenty clones from zone I and 40 clones from zone III were sequenced. Only one short sequence which was derived from zone III showed 97% similarity to Geobacter spp. The remaining sequences were similar to Acidobacterium sp. (89 to 97%), Streptococcus (90 to 99%), Propionibacterium (97%), Ferrimicrobium (92%), and Actinomyces (94%). When primer pair GM3/Geo825R, also specific for Geobacter, was used, 33 different phylotypes were obtained. Comparative sequence analysis revealed that four of the sequences obtained showed high sequence identity to Geobacteraceae sequences; one showed 90% similarity to Geobacter grbicium (15), and three were related to Geobacter sp. strain CdA-2 (93 to 97%) (Fig. 6). The remaining sequences were similar to Syntrophobacteraceae (91 to 96%), Desulfobacteraceae (91 to 93%), Magnetobacterium sp. (85%), Acidobacterium sp. (92%), and Actinomyces sp. (91%).


Figure 6
View larger version (25K):
[in this window]
[in a new window]

  		FIG. 6. Phylogenetic tree of Deltaproteobacteria-related 16S rRNA gene sequences determined by phylogenetic distance analysis with a maximum-likelihood algorithm. The designations for clone sequences derived from dilution PCR analysis with group-specific primers begin with "D-PCR," and the designations for clone sequences from sediment community clone libraries begin with "Common."

 
When the primer pair specific for Geothrix was used, 12 clones were obtained from zone II, and 9 of them were sequenced without previous RFLP screening. Only one sequence was related to Geothrix fermentans (95%). When the primer pair specific for Shewanella was used, eight white clones were sequenced without previous RFLP screening. One sequence was related to Shewanella algae (93%), and the remaining 7 clones were related to Sterolibacterium denitrificans (~95%) and Chromobacterium sp. (89 to 95%).

Enumeration and characterization of Fe(III) reducers.
Pooled sediment from three cores obtained in March 2003 was used to enumerate the Fe(III)-reducing microorganisms in zone III capable of growing at pH 5.5. The MPN of Fe(III)-reducing microorganisms were approximately 104 to 105 cells g (wet weight)–1, i.e., approximately 0.5% of the total DAPI-stained microorganisms present in zone III. The following MPN values were obtained for Fe(III) reducers from sediment zone III cultured at pH 5.5: for H2-utilizing Fe(III) reducers, 2.3 x 104 cells g (wet weight) of sediment–1 (confidence limits, 4.9 x 103 to 1.1 x 105 cells g–1); for ethanol-utilizing Fe(III) reducers, 4.0 x 105 cells g (wet weight) of sediment–1 (8.6 x 104 to 1.9 x 106 cells g–1); and for lactate-utilizing Fe(III) reducers, 2.3 x 104 cells g (wet weight) of sediment–1 (4.9 x 104 to 1.1 x 105 cells g–1). In MPN tubes scored positive for Fe(II) formation, the consumption of ethanol and lactate yielded acetate as the main organic end product. Four MPN enrichment cultures that exhibited high Fe(III)-reducing activities were selected for further 16S rRNA gene-based community analysis [one MPN 10–4 dilution and one MPN 10–3 dilution of ethanol-utilizing Fe(III) reducers, one MPN 10–3 dilution of lactate-utilizing Fe(III) reducers, and one MPN 10–1 dilution of H2-utilizing Fe(III) reducers]. Rarefaction curves indicated that there was saturation of sampling (data not shown). The majority (63%) of sequenced 16S rRNA gene clones of the four clone libraries were most closely related to members of the Betaproteobacteria (Fig. 7), primarily to species of Thiomonas (see Table S2 in the supplemental material), a genus which is closely related to the family that contains Thiobacillus. Members of the Deltaproteobacteria and Firmicutes comprised 19 and 18% of all the clones, respectively. Phylotypes related to the class Deltaproteobacteria included some clones related to Geobacter species, whereas most phylotypes related to the Firmicutes were related to Desulfosporosinus. Phylotypes related to Acidobacteria were detected only in H2-supplemented enrichment cultures and showed 96% similarity to G. fermentans.


Figure 7
View larger version (19K):
[in this window]
[in a new window]

  		FIG. 7. Frequencies of bacterial phylogenetic lineages detected in 16S rRNA gene clone libraries from four MPN enrichment cultures of Fe(III)-reducing bacteria derived from zone III.

 

   DISCUSSION
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Raising the water table after open-cast lignite mines were closed led to the formation of numerous acidic mining-associated lakes in wide areas of Germany, Poland, and the Czech Republic (67). Coal mining-associated lakes are characterized by moderate temperatures, small amounts of toxic metals, and a pH not less than 2 (27). The more extreme acid mine drainage (AMD)-impacted habitats contain fewer prokaryotic lineages (two archaeal and eight bacterial divisions) than many other environments (5, 9, 74). This study demonstrated that coal mining-associated lake sediments were inhabited by a diverse bacterial community that differed from the communities in extreme AMD environments. No PCR product was obtained with Archaea domain-specific primers, while 10 and 12 bacterial lineages were detected in sediment zones I and III, respectively. Phylogenetic analyses of 16S rRNA genes placed many of the sequences obtained in the Proteobacteria, Nitrospira, Firmicutes, and Acidobacteria groups, similar to the results for AMD and other bioleaching sites (5, 74) and uranium-contaminated sediments (2, 70). However, in contrast to the previous studies, many of the clones (46 and 45% in zones I and III, respectively) were related to sequences which clustered within the Acidobacteria. Many of the phylotypes were related to G. fermentans, an Fe(III)-reducing heterotroph, and to one of the three cultured and described representatives of the phylum Acidobacteria (12, 16). Members of the Acidobacteria have been detected in a great variety of ecosystems, including soils, peatlands, geothermal vents, AMD sites, and uranium-contaminated environments (6, 12, 36, 54, 66). Although the physiological capabilities of the vast majority of Acidobacteria remain unclear, some moderate acidophilic Acidobacterium spp. have been shown to oxidize Fe(II) (31). A. capsulatum formed Fe(II) under anoxic conditions at pHs ranging from 2.2 to 5 during glucose fermentation. Since the cell counts of A. capsulatum did not increase and only small amounts of reducing equivalents were transferred to Fe(III), the dissimilatory reduction of Fe(III) appeared not to be coupled with growth. Nonetheless, Acidobacteria seemed to play a role in the cycling of iron in coal mining-associated lake sediments. Clones related to Betaproteobacteria, Firmicutes, and Actinomycetes were detected only in zone I, whereas clones related to Verrucomicrobia, Thermus/Deinococcus, or the Bacillus-Clostridium group were detected only in zone III. The diversity was greater in zone III than in zone I, which might have been due to the less acidic conditions. The low coverage (65%) of the rarefaction analysis indicated that the sampling efficiency was low; thus, the diversity in zone III might be even higher.

The dominant lithoautotrophic Fe(II) oxidizer in coal mining-associated lake sediments appeared to be A. ferrooxidans. Oligonucleotide probe-based studies have indicated that Leptospirillum strains often dominate microbial communities in hot and extremely acidic AMD environments (9, 24, 68), although some species are mesophiles and have been isolated from cold and cool environments. Nonetheless, coal mining-associated lake sediments appear to be a more suitable habitat for A. ferrooxidans due to their moderately acidophilic nature (37).

Large amounts of amorphous Fe(OH)3, goethite, and Fe(III) oxyhydroxysulfates, like schwertmannite or jarosite, precipitate to the sediments of coal mining-associated lakes (43, 59). Reduction of Fe(III) is often the dominant electron-accepting process for the oxidation of organic matter, whereas sulfate reduction is restricted to sediments with a pH greater than 5 (40, 53, 59). Although the primary production rate in these lakes is low (29, 58), Fe(III) reduction occurred at high rates and was linear for up to 60 days of incubation, indicating that both the pool of bioavailable Fe(III) and the pool of labile carbon were sufficient.

Culture-dependent techniques used in previous studies demonstrated that heterotrophic and autotrophic acidophilic bacteria capable of reduction of Fe(III) were present in the sediments (41, 53). Due to the absence of elemental sulfur in the upper sediment zone (59), heterotrophic species appear to be responsible for the reduction of Fe(III) under acidic conditions. Surprisingly, PCR products of Acidiphilium-related strains were obtained to the same DNA dilution level in all three sediment zones, and a higher number of phylotypes related to cultured Acidiphilium or Acidisphaera species were obtained from zone III than from zone I. Acidisphaera rubrifaciens is a moderately acidophilic aerobic heterotroph that was isolated from an AMD site (35). The optimum pH for growth of A. cryptum JF-5 is approximately pH 3.2 (41), but six members of the genus Acidiphilium are adapted to pH values from 1.5 to 6.0 (37). A. cryptum ATCC 33463 has the capacity to reduce small amounts of Fe(III) at pH 5 (8), and A. capsulatum formed Fe(II) at pHs ranging from 2.2 to 5. Although the metabolic roles of the phylotypes detected only in zone III remain unknown, we can speculate that these organisms might be moderate acidophiles or well adapted to pH 5 conditions.

Acidophilic Fe(III) reducers likely do not have to attach to the surface of Fe(III) minerals to transfer electrons to Fe(III); instead, they can utilize the small pool of soluble Fe(III) for reduction (10). Thus, the reduction of Fe(III) should be less favorable at higher pHs, because the solubility of the Fe(III) minerals decreases with increasing pH. The concentration of soluble Fe(III) can reach 10 mM under extremely acidic conditions, whereas only 10–15 mM Fe(III) is present under neutral-pH conditions (46). In addition, the amount of Fe(II) adsorbed to mineral surfaces (17, 47, 64, 65) should increase at elevated pHs. This advanced surface passivation would inhibit the microbial reductive dissolution at higher pHs. Nonetheless, Fe(II) formation increased in zone I after a small lag phase when the pH was increased from 2.9 to 5.3. This lag phase could have resulted from acclimation to different chemical conditions caused by pH adjustment or could indicate a shift in the microbial Fe(III)-reducing population. Thus, the capacity of some acidophiles to reduce Fe(III) at pH 5 combined with the activity of moderate acidophilic Fe(III) reducers might be responsible for the elevated rates of Fe(III) reduction in zone I at pH 5.3. The increase in pH might also have resulted in better availability of carbon sources. At higher pHs, schwertmannite transforms to goethite, which leads to a reduction in the surface area. An increase in pH alters the charge of the iron oxide surfaces to more negative values so that negatively charged organic matter can desorb (45) and more surface Fe(III) might be accessible for Fe(III) reduction.

Zone I was highly enriched by freshly precipitated schwertmannite resulting from high Fe(III) sedimentation rates (570 g m–2 day–1) (59). The Fe(III) content of the solid phase in zone I was approximately 200 to 350 g kg–1, and 60 to 100% was reactive iron. In zone III, less than 90 g kg–1 was present, and the main Fe(III) oxide was goethite. When the pH of zone III was decreased from 5.3 to 2.9, Fe(II) formation was reduced by 30 to 40% despite the better solubility of Fe(III) under low-pH conditions. The lower rate of Fe(III) reduction suggested that the majority of Fe(III) reducers inhabiting zone III had minor capacities to reduce Fe(III) under acidic conditions and/or acidophilic Fe(III) reducers did not dominate this Fe(III)-reducing community. The pH and geochemical gradients in the sediment appeared to be responsible for the occurrence of two distinct Fe(III)-reducing communities.

PCR products obtained using primer sets specific for Geobacteraceae were obtained with higher DNA dilutions from zone III than from zone I. However, dilution PCR allowed only rough comparative estimates for zone III and zone I. Only a few sequences related to cultured Fe(III) reducers were detected. With primer pair Gb564/Gb1290, one sequence that was related to the family Geobacteraceae was obtained from 31 clones, and four sequences were obtained from 45 clones with primer pair GM3/Geo825R. The latter four sequences were related to an isolate obtained from a mining-impacted sediment designated Geobacter sp. strain CdA-2 that can reduce Fe(III) at a broad pH range (pH 5.5 to 8.1) (19). Both specific clone libraries revealed that these PCR primer sets are not specific for the Geobacteraceae group, as reported previously (18). Similar observations for acidic (pH < 4) uranium-contaminated sediments revealed that members of the Betaproteobacteria accounted for a large portion of the microbial community, whereas the levels of Geobacteraceae were below the detection limit (2). Geobacteraceae were detected only in enrichment cultures of acidic uranium-contaminated sediments incubated under neutral-pH conditions (60). Firmicutes-related clones had high sequence similarity to the spore-forming genus Desulfosporosinus and the endospore-forming genera Paenibacillus and Sporomusa (1, 71, 75). Using both cultivation-dependent and -independent techniques, these three genera have been shown to inhabit acidic coal mining-associated lake sediments (43, 53) and acidic uranium-contaminated sediments (2, 60).

Shewanella or Geothrix species appeared to not contribute substantially to the reduction of Fe(III) in coal mining-associated lake sediments. Clonal analysis of the highest positive dilutions of Fe(III)-reducing communities enriched from zone III at pH 5.5 revealed that the majority of the clones were related to Thiomonas species (55), independent of the electron donor used. Some clone sequences obtained from our cultures were related to known Fe(III) reducers, like Geobacter sp. clone Fe_P3-19 or G. fermentans, but they were also related to cultured sulfate reducers, like Desulfosporosinus species, which are known to inhabit acidic coal mining-associated lake sediments (43, 53). Sulfate reducers can reduce Fe(III) either directly or indirectly through generation of sulfide (50). To obtain a better understanding of the cycling of Fe in other slightly acidic environments, more suitable cultivation techniques should be developed to study the ecophysiology of new acidophilic or acid-tolerant isolates.


   ACKNOWLEDGMENTS
 
We thank Winfrid Gade for his help with collecting the sediment cores and Stefan Peiffer for constructive discussions.

Financial support for this study was provided by the Deutsche Forschungsgemeinschaft (grant KU 1367/1-2) and the German Ministry of Education, Science, Research, and Technology (grant PT BEO 51-0339476C).


   FOOTNOTES
 
* Corresponding author. Mailing address: Limnology Research Group, Institute of Ecology, Friedrich Schiller University Jena, Dornburgerstrasse 159, 07743 Jena, Germany. Phone: (49) (0)3641-949461. Fax: (49) (0)3641-949462. E-mail: Kirsten.Kuesel{at}uni-jena.de Back

{triangledown} Published ahead of print on 14 December 2007. Back

{dagger} Supplemental material for this article may be found at http://aem.asm.org/. Back

{ddagger} Present address: Department of Geology and Geophysics, University of Wisconsin—Madison, Madison, WI 53706. Back


   REFERENCES
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Achouak, W., P. Normand, and T. Heulin. 1999. Comparative phylogeny of rrs and nifH genes in the Bacillaceae. Int. J. Syst. Bacteriol. 49:961-967.[Abstract/Free Full Text]
  2. Akob, D. M., H. J. Mills, and J. E. Kostka. 2007. Metabolically active microbial communities in uranium contaminated subsurface sediments. FEMS Microbiol. Ecol. 59:95-107.[CrossRef][Medline]
  3. Alef, K. 1991. Methodenhandbuch Bodenmikrobiologie: Aktivitäten, Biomasse, Differenzierung. Ecomed Verlagsgesellschaft mgH, Landsberg/Lech, Germany.
  4. Altschul, S. F., W. Gish, W. Miller, E. W. Myers, and J. D. Lipman. 1990. Basic local alignment search tool. J. Mol. Biol. 215:403-410.[CrossRef][Medline]
  5. Baker, B. J., and J. F. Banfield. 2003. Microbial communities in acid mine drainage. FEMS Microbiol. Ecol. 44:139-152.[CrossRef]
  6. Barns, S. M., E. C. Cain, L. Sommerville, and C. R. Kuske. 2007. Acidobacteria phylum sequences in uranium-contaminated subsurface sediments greatly expand the known diversity within the phylum. Appl. Environ. Microbiol. 73:3113-3116.[Abstract/Free Full Text]
  7. Bigham, J. M., U. Schwertmann, S. J. Traina, R. L. Winland, and M. Wolf. 1996. Schwertmannite and the chemical modeling of iron in acid sulfate waters. Geochim. Cosmochim. Acta 60:2111-2121.[CrossRef]
  8. Bilgin, A. A., J. A. Silverstein, and J. D. Jenkins. 2004. Iron respiration by Acidiphilium cryptum at pH 5. FEMS Microbiol. Ecol. 49:137-143.[CrossRef]
  9. Bond, P. L., S. P. Smriga, and J. F. Banfield. 2000. Phylogeny of microorganisms populating a thick, subaerial, predominantly lithotrophic biofilm at an extreme acid mine drainage site. Appl. Environ. Microbiol. 66:3842-3849.[Abstract/Free Full Text]
  10. Bridge, T. M., and D. B. Johnson. 2000. Reductive dissolution of ferric iron minerals by Acidiphilium SJH. Geomicrobiol. J. 17:193-206.[CrossRef]
  11. Brock, T. D., and J. Gustafson. 1976. Ferric iron reduction by sulfur- and iron-oxidizing bacteria. Appl. Environ. Microbiol. 32:567-571.[Abstract/Free Full Text]
  12. Buckley, D. H., and T. M. Schmidt. 2002. Exploring the diversity of soil—a microbial rain forest, p. 183-208. In J. T. Staley and A. Reysenbach (ed.), Biodiversity of microbial life: foundation of Earth's biosphere. Wiley-Liss, Inc., New York, NY.
  13. Canfield, D. E., R. Raiswell, and S. Bottrell. 1993. The reactivity of sedimentary iron minerals toward sulfide. Am. J. Sci. 292:659-683.
  14. Coates, J. D., and L. A. Achenbach. 2001. The biogeochemistry of aquifer systems, p. 719-727. In C. J. Hurst, G. R. Knudsen, M. J. McInerney, L. D. Stetzenbach, and M. W. Walter (ed.), Manual of environmental microbiology, 2nd ed. ASM Press, Washington, DC.
  15. Coates, J. D., V. Bhupathiraju, L. A. Achenbach, M. J. McInerney, and D. R. Lovley. 2001. Geobacter hydrogenophilus, Geobacter chapelleii, and Geobacter grbicium, three new, strictly anaerobic, dissimilatory Fe(III)-reducers. Int. Syst. Evol. Microbiol. 51:581-588.
  16. Coates, J. D., D. J. Ellis, C. V. Gaw, and D. R. Lovley. 1999. Geothrix fermentans gen. nov., sp. nov., a novel Fe(III)-reducing bacterium from a hydrocarbon-contaminated aquifer. Int. J. Syst. Bacteriol. 49:1615-1622.[Abstract/Free Full Text]
  17. Coughlin, B. R., and A. T. Stone. 1995. Nonreversible adsorption of divalent metal ions (MnII, CoII, NiII, CuII, and PbII) onto goethite: effects of acidification, FeII addition, and picolinic acid addition. Environ. Sci. Technol. 29:2445-2455.
  18. Cummings, D. E., O. L. Snoeyenbos-West, D. T. Newby, A. M. Niggemyer, D. R. Lovley, A. Achenbach, and R. F. Rosenzweig. 2003. Diversity of Geobacteraceae species inhabiting metal-polluted freshwater lake sediments ascertained by 16S rDNA analyses. Microb. Ecol. 46:257-269.[Medline]
  19. Cummings, D. E., A. W. March, B. Bostick, S. Spring, J. F. F. Caccavo, S. Fendorf, and R. F. Rosenzweig. 2000. Evidence for microbial Fe(III) reduction in anoxic, mining impacted lake sediments (Lake Coeur d'Alene, Idaho). Appl. Environ. Microbiol. 66:154-162.[Abstract/Free Full Text]
  20. Das, A., and A. K. Mishra. 1996. Role of Thiobacillus ferrooxidans and sulphur (sulphide)-dependent ferric-iron-reducing activity in the oxidation of sulphide minerals. Appl. Microbiol. Biotechnol. 45:377-382.[CrossRef]
  21. De Wulf-Durand, P., L. J. Bryant, and L. I. Sly. 1997. PCR-mediated detection of acidophilic, bioleaching-associated bacteria. Appl. Environ. Microbiol. 63:2944-2948.[Abstract]
  22. Dojka, M. A., P. Hugenholtz, S. K. Haack, and N. R. Pace. 1998. Microbial diversity in a hydrocarbon- and chlorinated-solvent-contaminated aquifer undergoing intrinsic bioremediation. Appl. Environ. Microbiol. 64:3869-3877.[Abstract/Free Full Text]
  23. Drake, H. L. 1994. Acetogenesis, acetogenic bacteria, and the acetyl-CoA "Wood/Ljungdahl" pathway: past and current perspectives, p. 3-60. In H. L. Drake (ed.), Acetogenesis. Chapman and Hall, Inc., New York, NY.
  24. Edwards, K. J., T. M. Gihring, and J. F. Banfield. 1999. Seasonal variations in microbial populations and environmental conditions in an extreme acid mine environment. Appl. Environ. Microbiol. 65:3627-3632.[Abstract/Free Full Text]
  25. Embley, T. M., B. J. Finlay, R. H. Thomas, and P. L. Dyal. 1992. The use of rRNA sequences and fluorescent probes to investigate the phylogenetic positions of the anaerobic ciliate Metopus palaeformis and its archaeabacterial endosymbionts. J. Gen. Microbiol. 138:1479-1487.[Medline]
  26. Fortin, D., B. Davis, and T. J. Beveridge. 1996. Role of Thiobacillus and sulfate-reducing bacteria in iron biocycling in oxic and acidic mine tailings. FEMS Microbiol. Ecol. 21:11-24.[CrossRef]
  27. Geller, W., H. Klapper, and M. Schultze. 1998. Natural and anthropogenic sulfuric acidification of lakes, p. 3-14, In W. Geller, H. Klapper, and W. Salomons (ed.), Acid mining lakes—acid mine drainage, limnology and reclamation. Environmental Science Series. Springer, Berlin, Germany.
  28. Goebel, B. M., and E. Stackebrandt. 1994. Cultural and phylogenetic analysis of mixed microbial populations found in natural and commercial bioleaching environments. Appl. Environ. Microbiol. 60:1614-1621.[Abstract/Free Full Text]
  29. Gyure, R. A., A. Konopka, A. Brooks, and W. Doemel. 1987. Algal and bacterial activities in acidic (pH 3) strip mine lakes. Appl. Environ. Microbiol. 53:2069-2076.[Abstract/Free Full Text]
  30. Hallberg, K. B., and D. B. Johnson. 2001. Biodiversity of acidophilic prokaryotes. Adv. Appl. Microbiol. 49:37-84.[Medline]
  31. Hallberg, K. B., and D. B. Johnson. 2003. Novel acidophiles isolated from moderately acidic mine drainage waters. Hydrometallurgy 71:139-148.[CrossRef]
  32. Harrison, A. P., Jr. 1984. The acidophilic Thiobacilli and other acidophilic bacteria that share their habitat. Annu. Rev. Microbiol. 38:265-292.[CrossRef][Medline]
  33. Heck, K. L., G. van Belle, and D. Simberloff. 1975. Explicit calculation of the rarefaction diversity measurement and the determination of sufficient sample size. Ecology 56:1459-1461.[CrossRef]
  34. Herlihy, A. T., and A. L. Mills. 1985. Sulfate reduction in freshwater sediments receiving acid mine drainage. Appl. Environ. Microbiol. 49:179-186.[Abstract/Free Full Text]
  35. Hiraishi, A., Y. Matsuzawa, T. Kanbe, and N. Wakao. 2000. Acidisphaera rubrifaciens gen. nov., sp. nov., an anaerobic bacteriochlorophyll-containing bacterium isolated from acidic environments. Int. J. Syst. Evol. Microbiol. 50:1539-1546.[Abstract]
  36. Hugenholtz, P., B. M. Goebel, and N. R. Pace. 1998. Impact of culture-independent studies on the emerging phylogenetic view of bacterial diversity. J. Bacteriol. 18:4765-4774.
  37. Johnson, D. B., and F. F. Roberto. 1997. Heterotrophic acidophiles and their role in the bioleaching of sulfide materials, p. 259-279. In D. E. Rawlings (ed.), Biomining: theory, microbes and industrial processes, Springer, New York, NY.
  38. Johnson, D. B., M. A. Ghauri, and S. McGinness. 1993. Biogeochemical cycling of iron and sulphur in leaching environments. FEMS Microbiol. Rev. 11:63-70.[Medline]
  39. Johnson, D. B., and S. McGinness. 1991. Ferric iron reduction by acidophilic heterotrophic bacteria. Appl. Environ. Microbiol. 57:207-211.[Abstract/Free Full Text]
  40. Küsel, K., and T. Dorsch. 2000. Effect of supplemental electron donors on the microbial reduction of Fe(III) and sulfate in coal mining-impacted freshwater lake sediments. Microb. Ecol. 40:238-249.[Medline]
  41. Küsel, K., T. Dorsch, G. Acker, and E. Stackebrandt. 1999. Microbial reduction of Fe(III) in acidic sediments: isolation of Acidiphilium cryptum JF-5 capable of coupling the reduction of Fe(III) to the oxidation of glucose. Appl. Environ. Microbiol. 65:3633-3640.[Abstract/Free Full Text]
  42. Küsel, K., U. Roth, and H. L. Drake. 2002. Microbial reduction of Fe(III) in the presence of oxygen under low pH conditions. Environ. Microbiol. 4:414-421.[CrossRef][Medline]
  43. Küsel, K., U. Roth, T. Trinkwalter, and S. Peiffer. 2001. Effect of pH on the anaerobic microbial cycling of sulfur in mining-impacted freshwater lake sediments. Environ. Exp. Bot. 46:213-223.[CrossRef]
  44. Küsel, K., and H. L. Drake. 1995. Effects of environmental parameters on the formation and turnover of acetate by forest soils. Appl. Environ. Microbiol. 61:3667-3675.[Abstract]
  45. Laskov, C., W. Amelung, and S. Peiffer. 2002. Organic matter preservation in the sediment of an acidic mining lake. Environ. Sci. Technol. 36:4218-4221.
  46. Lindsay, W. L., and A. P. Schwab. 1982. The chemistry of iron in soils and its availability to plants. J. Plant Nutr. 5:1531-1536.
  47. Liu, C., S. Kota, J. M. Zachara, J. K. Fredrickson, and C. K. Brinkmann. 2001. Kinetic analysis of the bacterial reduction of goethite. Environ. Sci. Technol. 35:2482-2490.
  48. Liu, W. T., C. L. Huang, J. Y. Hu, L. F. Soon, S. L. Ong, and W. J. Ng. 2002. Denaturing gradient gel electrophoresis polymorphism for rapid 16S rDNA clone screening and microbial diversity estimation. J. Biosci. Bioeng. 93:101-103.
  49. Lovley, D. R. 2006. Dissimilatory Fe(III)- and Mn(IV)-reducing prokaryotes, p. 635-658. In M. Dworkin et al. (ed.), The prokaryotes, 3rd ed., vol. 2. Springer, New York, NY.
  50. Lovley, D. R., D. E. Holmes, and K. P. Nevin. 2004. Dissimilatory Fe(III) and Mn(IV) reduction. Adv. Microb. Physiol. 49:212-286.
  51. Lovley, D. R., and E. J. P. Phillips. 1986. Organic matter mineralization with reduction of ferric iron in anaerobic sediments. Appl. Environ. Microbiol. 51:683-689.[Abstract/Free Full Text]
  52. Ludwig, W., O. Strunk, W., R. Westram, L. Richter, H. Meier, Yadhukumar, A. Buchner, T. Lai, S. Steppi, G. Jobb, W. Förster, I. Brettske, S. Gerber, A. W. Ginhart, O. Gross, S. Grumann, S. Hermann, R. Jost, A. König, T. Liss, R. Lüβmann, M. May, B. Nonhoff, B. Reichel, R. Strehlow, A. Stamatakis, N. Stuckmann, A. Vilbig, M. Lenke, T. Ludwig, A. Bode, and K.-H. Schleifer. 2004. ARB: a software environment for sequence data. Nucleic Acids Res. 32:1363-1371.[Abstract/Free Full Text]
  53. Meier, J., H.-D. Babenzien, and K. Wendt-Potthoff. 2004. Microbial cycling of iron and sulfur in sediments of acidic and pH-neutral mining lakes in Lusatia (Brandenburg, Germany). Biogeochemistry 67:135-156.
  54. Meisinger, D. B., J. Zimmermann, W. Ludwig, K.-H. Schleifer, G. Wanner, M. Schmid, P. C. Bennett, A. S. Engel, and N. M. Lee. 2007. In situ detection of novel Acidobacteria in microbial mats from a chemolithoautotrophically based cave ecosystem (Lower Kane Cave, WY, USA). Environ. Microbiol. 9:1523-1534.[CrossRef][Medline]
  55. Moreira, D., and R. Amils. 1997. Phylogeny of Thiobacillus cuprinus and other mixotrophic thiobacilli: proposal for Thiomonas gen. nov. Int. J. Syst. Evol. Microbiol. 47:522-528.[Abstract/Free Full Text]
  56. Muyzer, G., E. C. de Waal, and A. G. Uitterlinden. 1993. Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S rRNA. Appl. Environ. Microbiol. 59:695-700.[Abstract/Free Full Text]
  57. Muyzer, G., S. Hottenträger, A. Teske, and C. Waver. 1995. Denaturing gradient gel electrophoresis of PCR amplified 16S rDNA—a new molecular approach to analyse the genetic diversity of mixed microbial communities, p. 1-22. In A. D. Akkermanns, J. D. van Elsas, and F. J. de Bruijn (ed.), Molecular microbial ecology manual 3.4.4. Kluwer, Dordrecht, The Netherlands.
  58. Nixdorf, B., and M. Kapfer. 1998. Stimulation of phototrophic pelagic and benthic metabolism close to sediments in acidic mining lakes. Water Air Soil Pollut. 108:317-330.[CrossRef]
  59. Peine, A., A. Tritschler, K. Küsel, and S. Peiffer. 2000. Electron flow in an iron-rich acidic sediment—evidence for an acidity-driven iron cycle. Limnol. Oceanogr. 45:1077-1087.
  60. Petrie, L., N. N. North, S. L. Dollhopf, D. L. Blackwill, and J. E. Kostka. 2003. Enumeration and characterization of iron(III)-reducing microbial communities from acidic subsurface sediments contaminated with uranium(VI). Appl. Environ. Microbiol. 69:7467-7479.[Abstract/Free Full Text]
  61. Picard, C., C. Ponsonnet, E. Paget, X. Nesme, and P. Simonet. 1992. Detection and enumeration of bacteria in soil by direct DNA extraction and polymerase chain reaction. Appl. Environ. Microbiol. 58:2717-2722.[Abstract/Free Full Text]
  62. Pronk, J. T., J. C. De Bruyn, P. Bos, and J. G. Kuenen. 1992. Anaerobic growth of Thiobacillus ferrooxidans. Appl. Environ. Microbiol. 58:2227-2230.[Abstract/Free Full Text]
  63. Pronk, J. T., and D. B. Johnson. 1992. Oxidation and reduction of iron by acidophilic bacteria. Geomicrobiol. J. 10:153-171.
  64. Roden, E. R., and J. M. Zachara. 1996. Microbial reduction of crystalline iron(III) oxides: influence of oxide surface area and potential for cell growth. Environ. Sci. Technol. 30:1618-1628.
  65. Roden, E. R., M. M. Urrutia, and J. M. Carroll. 2000. Bacterial reductive dissolution of crystalline Fe(III) oxide in continuous-flow column reactors. Appl. Environ. Microbiol. 66:1062-1065.[Abstract/Free Full Text]
  66. Sait, M., K. E. R. Davis, and P. H. Janssen. 2006. Effect of pH on isolation and distribution of members of subdivision 1 of the phylum Acidobacteria occurring in soil. Appl. Environ. Microbiol. 72:1852-1857.[Abstract/Free Full Text]
  67. Schnoor, J. L., J. N. Galloway, and B. Moldan. 1997. East central Europe: an environment in transition. Environ. Sci. Technol. 31:412-416.
  68. Schrenk, M. O., K. J. Edwards, R. M. Goodman, R. J. Hamers, and J. F. Banfield. 1998. Distribution of Thiobacillus ferrooxidans and Leptospirillum ferrooxidans: implications for generation of acidic mine drainage. Science 279:1519-1522.[Abstract/Free Full Text]
  69. Singleton, D. R., M. A. Furlong, S. L. Rathbun, and W. B. Whitman. 2001. Quantitative comparisons of 16S rRNA gene sequence libraries from environmental samples. Appl. Environ. Microbiol. 67:4374-4376.[Abstract/Free Full Text]
  70. Snoeyenbos-West, O. L., K. P. Nevin, R. T. Anderson, and D. R. Lovley. 2000. Enrichment of Geobacter species in response to stimulation of Fe(III) reduction in sandy aquifer sediments. Microb. Ecol. 39:153-167.[CrossRef][Medline]
  71. Spring, S., and F. Rosenzweig. 2006. The genera Desulfitobacterium and Desulfosporosinus: taxonomy, p. 771-786. In M. Dworkin et al. (ed.), Th