Anaerobic Consumers of Monosaccharides in a Moderately Acidic Fen

Abstract

16S rRNA-based stable isotope probing identified active xylose- and glucose-fermenting Bacteria and active Archaea, including methanogens, in anoxic slurries of material obtained from a moderately acidic, CH₄-emitting fen. Xylose and glucose were converted to fatty acids, CO₂, H₂, and CH₄ under moderately acidic, anoxic conditions, indicating that the fen harbors moderately acid-tolerant xylose- and glucose-using fermenters, as well as moderately acid-tolerant methanogens. Organisms of the families Acidaminococcaceae, Aeromonadaceae, Clostridiaceae, Enterobacteriaceae, and Pseudomonadaceae and the order Actinomycetales, including hitherto unknown organisms, utilized xylose- or glucose-derived carbon, suggesting that highly diverse facultative aerobes and obligate anaerobes contribute to the flow of carbon in the fen under anoxic conditions. Uncultured Euryarchaeota (i.e., Methanosarcinaceae and Methanobacteriaceae) and Crenarchaeota species were identified by 16S rRNA analysis of anoxic slurries, demonstrating that the acidic fen harbors novel methanogens and Crenarchaeota organisms capable of anaerobiosis. Fermentation-derived molecules are conceived to be the primary drivers of methanogenesis when electron acceptors other than CO₂ are absent, and the collective findings of this study indicate that fen soils harbor diverse, acid-tolerant, and novel xylose-utilizing as well as glucose-utilizing facultative aerobes and obligate anaerobes that form trophic links to novel moderately acid-tolerant methanogens.

Wetlands harbor approximately 30% of the global reserves of soil carbon; they are characterized by highly dynamic O₂ gradients and temporarily negligible concentrations of inorganic electron acceptors, such as sulfate or nitrate (e.g., 39, 43, 50); and they are one of the most important global sources of CH₄ (2). Wetlands account for about 20% of the global annual emission of the greenhouse gas CH₄ (19), which increased by 70% between 1970 and 2004 (42). CH₄ is the expected end product of the mineralization of organic matter when CO₂ is the sole electron acceptor (37). The organic matter of peat consists of 20 to 50% lignin and lignin-derived substances (e.g., lignocellulose [48]). Lignocellulose is the main component of the cell walls of plant tissue (1, 35) and consists of cellulose, hemicellulose, and lignin (1). The main components of hemicellulose are the polymers xylan (which consists mainly of xylose) and cellulose (which consists mainly of glucose). Xylose and glucose are released via the hydrolysis of xylan and cellulose by extracellular enzymes of primary fermenters (26). Hence, xylose and glucose are conceived to be intermediates in the “intermediary ecosystem metabolism” of methanogenic wetlands (i.e., in the metabolic processes that precede methanogenesis in wetlands), which can be converted by fermenters to organic acids (e.g., acetate), alcohols, CO₂, and H₂. Acetate and H₂-CO₂ are major substrates for methanogenesis (7, 37, 50, 51). Thus, fermentation is the prerequisite for the production of CH₄.

Although 16S rRNA gene pools of a few acidic wetlands have been evaluated (8, 21, 27), the microbial community composition of active fermenters involved in the carbon flow of fens is still largely unresolved. Since fermenters do not form monophyletic groups, it is not possible to accurately identify fermenters based on 16S rRNA gene sequences alone. Although methanogens in acidic wetlands have been assessed by 16S rRNA gene and methyl coenzyme M reductase alpha (mcrA) subunit analyses (e.g., see references 13, 14, and 18), the active fraction of methanogens has thus far not been identified. The objectives of this study were to employ different technologies to identify (i) xylose- and glucose-utilizing fermenters, (ii) intermediates in the carbon flow to methane, and (iii) active, moderately acid-tolerant Archaea in a northern temperate acidic fen.

MATERIALS AND METHODS

Sampling site and sampling.

The Schlöppnerbrunnen fen is located in the Lehstenbach catchment in the Fichtelgebirge, in northeastern Bavaria, Germany (50°07′53"N, 11°52′51"E), and is 700 m above sea level. The fen emits up to approximately 530 μmol CH₄ m⁻² h⁻¹ (S. Goldberg, K.-H. Knorr, and G. Gebauer, unpublished data). Norway spruce (Picea abies) is the dominant tree of the catchment. Thirty percent of the catchment is covered by fens or intermittent seeps. The annual precipitation varies between 900 and 1,160 mm year⁻¹, and the average annual temperature range is 5 to 8°C (43). Schlöppnerbrunnen is completely overgrown by Molinia caerulea, Eriophorum vaginatum, Carex canescens, and Juncus effusus (43). The fen soil is a fibric histosol type, with a pH of approximately 5. Soil samples were taken on 2 November 2005 from the upper 20 cm of the fen, transported in air-tight plastic bags, and stored at 2°C until processed (within approximately 2 h). The dry weight of the collected soil was approximately 15% (wt/wt).

Incubations for stable isotope probing.

An anoxic mineral salt solution was prepared using a modified Hungate technique (20). The sterile anoxic mineral solution contained mineral salts [12.6 mg/liter (NH₄)₂SO₄; 13.5 mg/liter CaCl₂·2H₂O; 10 mg/liter MgCl₂·6H₂O; 4 mg/liter KH₂PO₄; and 10 mg/liter FeCl₂·4H₂O; modified from the solution described in ref. 28] and trace elements [15 mg/liter nitrilotriacetic acid; 5 mg/liter MnSO₄·1H₂O; 1 mg/liter FeSO₄·7H₂O; 1 mg/liter CoCl₂·6H₂O, CaCl₂·2H₂O; 1 mg/liter ZnSO₄·H₂O; 0.1 mg/liter CuSO₄·H₂O; 2 mg/liter AlK(SO₄)₂·12H₂O; 0.1 mg/liter H₃BO₃; and 0.1 mg/liter Na₂MoO₄·2H₂O]. The solution was boiled and subsequently cooled under a 100% N₂ gas phase. After the solution was autoclaved, the pH was adjusted to 4.5 with HCl, and a filter-sterilized (0.2-μm-pore size) vitamin solution (10 ml per liter) containing 0.02 mg/liter biotin, 0.02 mg/liter folic acid, 0.1 mg/liter pyridoxine-HCl, 0.05 mg/liter thiamine-HCl, 0.05 mg/liter riboflavin, 0.05 mg/liter niacin, 0.05 mg/liter dl-Ca-pantothenic acid, 0.001 vitamin B₁₂, 0.05 mg/liter p-aminobenzoic acid, and 0.05 mg/liter lipoic acid was added anoxically. A soil slurry was prepared by adding 690 ml of the sterile anoxic mineral salt solution to 275 g (fresh weight) of fen soil in a sterile screw-cap, butyl rubber-stoppered flask (1.1 liter), and the gas phase was adjusted to 100% N₂ in order to maximize anaerobic activities and minimize incubation times. Fen soil was subsequently preincubated without any substrates to minimize the amount of electron acceptors other than CO₂. The preincubation was performed in an end-over-end shaker for 8 days in the dark, until nitrate and sulfate were depleted and most of the Fe³⁺ was converted to Fe²⁺ (based on Fe²⁺ reaching a stable, maximum end concentration). The slurry was divided into 11 butyl rubber-stoppered flasks (350 ml) in an anaerobic chamber (Mecaplex, Switzerland) under an N₂ atmosphere. [¹³C]xylose or [¹³C]glucose as well as [¹²C]xylose or [¹²C]glucose was provided as a supplement from sterile, anoxic stock solutions to a final concentration of approximately 0.4 mM. Controls were unsupplemented slurries. Three replicates were performed for both the ¹²C-labeled substrate-containing slurry and the control slurry. Substrates were refed when their concentration was below 0.2 mM. The flasks were incubated at 15°C in the dark. Gases (H₂, CO₂, and CH₄), monosaccharides, alcohols, and organic acids were analyzed.

Analytical techniques.

The moisture content of the fen soil was determined by weighing the soil before and after it was dried at 60°C for 72 h. pH was measured with a model U457-S7/110 combination pH electrode (Ingold, Steinbach, Germany). Nitrate, sulfate, and Fe²⁺ were measured spectrophotometrically (46, 47, 49). Gases were measured with a 5980 series II model gas chromatograph, with thermal conductivity and flame ionization detection, and with a 3396 series II model integrator (Hewlett Packard, Palo Alto, CA). Monosaccharides, organic acids, and alcohols were analyzed with a 1090 series II high-performance liquid chromatograph with a refractive index detector (Hewlett Packard, Palo Alto, CA [29, 36]).

Nucleic acid extraction.

RNA was extracted from 0.5 g of soil by bead-beating lysis, organic solvent extraction, and precipitation (16). The purification of RNA from coextracted DNA was performed with a Qiagen RNA/DNA mini-kit (Qiagen, Hilden, Germany) according to the manufacturer's protocol. DNA was extracted from fen material with a Power Soil DNA isolation kit (MoBio Laboratories, Carlsbad, CA), according to the manufacturer's protocol.

Separation of ¹³C- and ¹²C-labeled RNA.

A gradient solution (buoyant density, 1.78 ± 0.005 g ml⁻¹) of cesium trifluoroacetate (83.3%), gradient buffer (pH 8; 100 mM Tris; 100 mM KCl; 1 mM EDTA; 13.5%), and formamide (3.2%) was added to 500 ng of RNA and filled into 13-mm-by-51-mm polyallomer Quick Seal tubes (Beckman). RNA fractions were resolved by equilibrium centrifugation (∼130,000 × g at 20°C for 62 h) (11, 32). Fractions (400 μl) were collected with syringes with 21-gauge needles. The densities of the gradient solutions of all fractions were determined by weighing them at 25°C. The fractionated RNA was recovered by the addition of an equal volume of isopropanol to the sample and centrifugation at 13,000 × g (at 4°C for 15 min). The RNA pellet was washed with 70% ice-cold ethanol, air dried, and dissolved in 20 μl of diethyl pyrocarbonate-treated water.

Reverse transcriptase PCR and PCR.

Reverse transcription of RNA was performed with random hexamers and SuperScript II reverse transcriptase according to the manufacturer's protocol (Invitrogen, Karlsruhe, Germany). cDNA was amplified by the bacterial and archaeal 16S rRNA-specific primer sets 27F/907RM (30) and ARC4F/915R (17), respectively. PCR was performed in a PeqLab Primus 96 model thermal cycler (PeqLab, Erlangen, Germany). A denaturation step of 5 min at 95°C was performed at the beginning of PCR amplifications. Denaturation and elongation were performed at 95°C (5 min) and 72°C (90 s), respectively. For the primer 27F/907RM, a 4-cycle pre-PCR procedure with annealing at 40°C (1 min) was performed, followed by 30 cycles with an annealing temperature at 50°C (for 30 s). A step of 31 PCR cycles with annealing at 55°C (50 s) was used for the primer ARC4F/915R. For denaturing gradient gel electrophoresis (DGGE) analysis, PCR was performed with the primers GM5-clamp/907RM for Bacteria and 344Fa-Clamp/915R for Archaea, for 33 cycles at an annealing temperature of 56°C (18, 40). PCR with the primer M13F/M13R was performed per the published protocol (38).

DGGE.

16S rRNA genes were analyzed by DGGE (18, 40) using a PhorU system (Ingeny, Goes, The Netherlands) on 6% polyacrylamide gels, with a denaturant gradient from 35 to 70% (100% denaturant was 42% [wt/vol] urea and 40% [vol/vol] formamide). Electrophoresis was performed for 18 h at 100 V and 60°C. Gels were stained with SYBR Green (Invitrogen, Karlsruhe, Germany), visualized on a fluorescence imager unit (Storm 860; GE Healthcare, Piscataway, NJ), and analyzed with ImageQuant 5.0 software (GE Healthcare, Piscataway, NJ).

T-RFLP analysis.

For terminal restriction fragment length polymorphism (T-RFLP) analysis, PCR was performed with primers 27F and ARC915R labeled with IRD700 (MWG, Ebersberg, Germany). T-RFLP was performed per the published protocol (31). Amplified and IRD700-labeled bacterial cDNA were restricted with mung bean nuclease (New England Biolabs, Frankfurt am Main, Germany) to remove single-stranded DNA and minimize the formation of pseudo- terminal restriction fragments (pseudo-T-RFs) (12). The restricted cDNA was purified with Millipore PCR₉₆ cleanup plates (Millipore Corporation, Bedford, MA) and digested with the restriction endonuclease MspI (Fermentas, St. Leon-Rot, Germany). Gel electrophoresis was performed with a NEN model 4300 DNA analyzer (Licor, Lincoln, NE). The polyacrylamide gel consisted of 15 g of urea, 3.75 ml of 40% acrylamide-bis solution (37.5:1; 2.6% C; Bio-Rad, Hercules, CA), 12 ml of 2.5× Tris-borate-EDTA buffer (AppliChem GmbH, Darmstadt, Germany), and 3.25 ml of double-distilled H₂O. For the stabilization of the comb region of the gel, a bind-silane solution (1:1 bind-silane [PlusOne; GE Healthcare, Piscataway, NJ] and 10% acetic acid) was applied to the glass plates. The gel was poured according to the manufacturer's protocol. Electrophoresis was performed for 4 h at 1,200 V and 45°C. Gels were analyzed with GelQuest (Sequentix, Klein Raden, Germany).

Cloning, sequencing, phylogenetic analysis, and in silico T-RFLP analysis.

PCR products were ligated in a pGEM-T Easy vector and transformed into Escherichia coli JM109 competent cells according to the manufacturer's protocol (Promega, Madison, WI). Clones were screened for the correct insert with M13F/M13R (38). RFLP analysis of cloned bacterial and archaeal 16S rRNA genes was performed with the restriction enzymes HaeIII (Fermentas, St. Leon-Rot, Germany) and TacI (New England BioLabs, Ipswich, MA), respectively. One representative sequence for each of the different RFLP patterns was obtained commercially from Macrogen (Seoul, South Korea). Sequences were analyzed with ARB software (34) and WU-BLAST2 (http://www.ncbi.nlm.nih.gov/BLAST/). Sequences were aligned with FastAligner software (included in ARB software) (http://www.arb-home.de [34]), and the alignment was refined manually. Tree calculations were based upon 16S rRNA sequences (700 to 800 bp) and performed using parsimony, neighbor-joining, and maximum likelihood methods. Consensus trees were drawn using consistent branching of all three methods (33). The outgroups Thermocrinis albus (EBI accession no. AJ278895) and Thermoplasma acidophilum (EBI accession no. M20822) were used for calculations of the bacterial and archaeal trees, respectively. In silico T-RFs were calculated with T-RFcut software (44) implemented in ARB and compared to experimentally obtained T-RFs to associate T-RFs with 16S rRNA sequences.

Nucleotide sequence accession numbers.

16S rRNA sequences obtained in this study were deposited in the European Bioinformatics Institute (http://www.ebi.ac.uk) under the accession numbers AM712557 to AM712594, AM773593, AM773619, AM773938, AM773944, AM773947, AM773964, AM773974, AM773982, AM773991, and AM773995.

RESULTS

Effects of xylose and glucose on fermentation and methanogenesis.

Anoxic slurries of fresh fen soil were preincubated for 8 days to ensure that electron acceptors other than CO₂ were consumed prior to the addition of monosaccharides. CH₄ was produced at rates of 16 ± 0.01 μmol day⁻¹ g_dw⁻¹ (where g_dw is grams [dry weight]) in unsupplemented controls after the preincubation period, indicating that (i) electron acceptors other than CO₂ had been consumed or were not inhibiting methanogenesis and (ii) fermentative microbes would very likely be the primary consumers of monosaccharides after the preincubation period. Xylose and glucose were initially consumed within 5 and 4 days, respectively (Fig. 1). The cumulative amounts of xylose and glucose that were consumed within 14 days were 18 μmol g_dw⁻¹ and 20 μmol g_dw⁻¹, respectively. Formate accumulated transiently up to approximately 2 μmol g_dw⁻¹ and was consumed. In xylose- and glucose-supplemented fen slurries, acetate accumulated to approximately 4.4 and 9.5 μmol g_dw⁻¹, respectively, and butyrate to approximately 3.3 and 6.3 μmol g_dw⁻¹, respectively (Fig. 1A2 and B2), indicating that fermentation occurred in the slurries. Approximately 40 μmol CO₂ g_dw⁻¹ was produced in both treatments (Fig. 1A3 and B3). H₂ and CH₄ accumulated in both the xylose-supplemented slurry (approximately 4.0 μmol g_dw⁻¹ and 1.6 μmol g_dw⁻¹, respectively; Fig. 1A3) and the glucose-supplemented slurry (approximately 3.6 μmol g_dw⁻¹ and 1.2 μmol g_dw⁻¹, respectively; Fig. 1B3). H₂ was not detected in unsupplemented slurries. At day 5 of the xylose experiment, 75% of the electrons were recovered in fermentation products (acetate, butyrate, formate, and CO₂ [27%]) and 1% in CH₄; at day 12, electron recoveries from fermentation products (acetate, butyrate, propionate, and CO₂ [50%]) and CH₄ approximated 89% and 2%, respectively. At day 3 of the glucose experiment, 70% of the electrons were recovered from fermentation products (acetate, butyrate, propionate, and CO₂ [37%]) and 0.4% from CH₄; at day 12, electron recoveries from fermentation products (acetate, butyrate, propionate, and CO₂ [48%]) and CH₄ approximated 98% and 2%, respectively. Since the bulk of recovered electrons were accounted for in fermentation products, it can be concluded that fermentation was significant during electron flow.

FIG. 1.

Effects of supplemental xylose (A1) and glucose (B1) on the production of organic acids (A2 and B2) and gases (A3 and B3) by anoxic fen slurries (n = 3). Concentrations were corrected with values obtained from control slurries (i.e., unsupplemented anoxic slurries). Gases are given in mM, i.e., mmol produced per liter of liquid phase. Arrows indicate the addition of monosaccharides. Dotted lines indicate early and late sampling time points for RNA gradients.

Fingerprints of xylose and glucose consumers.

Isopycnic centrifugations to separate ¹³C- and ¹²C-labeled RNA were performed with RNA extracted after days 2 and 4 (early time points, before monosaccharides were completely consumed; Fig. 1A1 and B1) from xylose- and glucose-supplemented slurries, respectively, and at day 13 (a late time point) for both slurries. The buoyant densities of all fractions (1-12) followed a linear gradient from 1.84 to 1.76 g ml⁻¹, indicating that the isopycnic centrifugation was successful. The buoyant densities of fractions 4 and 5 (“heavy”) and 10 (“light”) were 1.816 to 1.81 g ml⁻¹ and 1.768 g ml⁻¹, respectively, indicating that these fractions contained [¹³C]RNA and [¹²C]RNA, respectively (32). DGGE profiles of bacterial 16S rRNA fragments amplified from “heavy” fractions 4 and 5 were different from those of “light” fractions 6 to 11 (Fig. 2 and data not shown). These results indicate that certain bacterial 16S rRNAs in the “heavy” fractions were distinct from those of the “light” fractions and that a sufficient amount of ¹³C label was incorporated into the RNA to allow a successful separation of [¹³C]RNA from [¹²C]RNA at the early and late time points.

FIG. 2.

DGGE profile of fractions 4 to 11 of bacterial RNA gradients obtained from the [¹³C]xylose experiment at day 13. Numbers at the top of the lanes correspond to fractions 4 to 11. Arrows identify bands that occur only in the “heavy” fractions (4 and 5) or are more intensive than in the “light” fractions (6-11).

T-RFLP analysis of density-resolved “heavy” bacterial rRNA in fraction 4 obtained from the incubations with [¹³C]xylose and [¹³C]glucose at early and late time points showed up to 31 T-RFs (Fig. 3), indicating that (i) diverse microorganisms had metabolized the labeled substrates and (ii) a sufficient amount of ¹³C label was incorporated into the RNA at the early time point. In the [¹³C]xylose experiment, T-RFLP profiles of the heavy fraction yielded a higher relative abundance of 18 T-RFs (Fig. 3A1 and A2, T-RFs 116, 140, 150, 166, 170, 190, 266, 326, 420, 440, 454, 468, 492, 496, 520, 544, 562, and 592 bp; these T-RFs refer to those detected at day 4 and/or 13) than the T-RFLP profiles of the “light” fraction (Fig. 3A1 and A2). Eleven of these T-RFs (116, 140, 150, 166, 190, 266, 468, 492, 520, 544, and 592 bp) occurred at both day 4 and day 13, and three of these T-RFs (166, 266, and 440 bp) became more prominent by day 13. In the [¹³C]glucose experiment, T-RFLP profiles of the “heavy” fraction yielded a higher relative abundance of 15 T-RFs (Fig. 3B1 and B2, 126, 140, 150, 158, 170, 190, 224, 326, 400, 420, 468, 496, 520, 544, and 562 bp; these T-RFs refer to those detected at day 2 and/or 13) than the T-RFLP profiles of the “light” fraction. Five of these T-RFs (158, 326, 420, 468, and 562 bp) occurred at both day 2 and day 13, and three of these T-RFs (140, 290, and 496 bp) became more prominent by day 13.

FIG. 3.

T-RFLP profiles of bacterial cDNA of “heavy” fraction 4 and “light” fraction 10 obtained from the [¹³C]xylose experiment (A) or the [¹³C]glucose experiment (B) at days 2 (A1), 4 (B1), and 13 (A2 and B2).

Phylogenetic analysis of active prokaryotic populations.

16S rRNA gene libraries of the “heavy” RNA fractions were generated to elucidate the phylogenetic identity of anaerobic xylose and glucose consumers. A total of 190 clones were screened by RFLP. Coverage of gene libraries as determined by RFLP analysis ranged from 86% to 96% (see Fig. S1 in the supplemental material). Fourteen different operational taxonomic units (OTUs; i.e., different RFLP patterns) were detected in the “heavy” fractions that were obtained from the early time points of the xylose and glucose incubations. Ten and six different OTUs were retrieved from the “heavy” RNA of the xylose and of the glucose incubations, respectively, at day 13 (data not shown). These 16S rRNA sequences were affiliated with classes Alpha-, Beta-, Gamma-, and Deltaproteobacteria, as well as with phyla Actinobacteria, Acidobacteria, and Clostridia (Fig. 4), indicating that phylogenetically diverse bacteria metabolized xylose and glucose in the moderately acidic fen soil slurries.

FIG. 4.

Consensus phylogenetic tree of bacterial 16S rRNA sequences obtained in this study (bold). Accession numbers are provided after sequence identifiers, and the relative abundance levels (percentages) of OTUs in gene libraries are given in parentheses. Codes: X4 and X13, sequences retrieved from “heavy” fractions at days 4 and 13, respectively, of the [¹³C]xylose experiment; G2 and G13, sequences retrieved from “heavy” fractions at days 2 and 13, respectively, of the [¹³C]glucose experiment; B, sequences obtained by direct 16S rRNA analysis of fen soil; n.d., not detected. Scale bar represents 10% sequence dissimilarity. The outgroup is Thermocrinis albus (accession no. AJ278895).

T-RFs detected in RNA density gradients from the [¹³C]xylose experiment were assigned to the cloned 16S rRNA genes either from fraction 4 or from direct 16S rRNA gene analysis (Table 1 and data not shown). T-RFs indicative of Acidobacteriaceae (96 and 116 bp), Acidothermaceae (140 bp), Sphingomonadaceae and Aeromonadaceae (both 150 bp), Pseudomonadaceae (492 bp), and Enterobacteriaceae and Aeromonadaceae (both 496 bp), as well as from Clostridiaceae (520 bp) organisms, were identified in the “heavy” fraction at the early time point (Fig. 3A1). These T-RFs had a lower relative abundance at the late sampling time point (Fig. 3A2). Relative abundance levels of T-RFs indicative of Syntrophobacteraceae, Rhodobiaceae (both 166 bp), and Acidobacteriaceae (266 bp) species were higher at day 13 than at day 4 (Fig. 3A1 and A2). The 440-bp T-RF indicative of Rhodospirillaceae and Chlorobiaceae organisms occurred only at day 13 (Fig. 3A2). T-RFs detected in RNA density gradients from the [¹³C]glucose experiment were assigned to families Sphingomonadaceae and Aeromonadaceae (both 150 bp) and to the class Clostridia (520 bp). Relative abundance levels of such T-RFs were already higher in the “heavy” fraction than in the “light” fraction after 2 days (Fig. 3B1), indicating that these community members had consumed [¹³C]glucose during the initial incubation period. The relative abundance levels of these T-RFs decreased during the incubation period (Fig. 3B2). The relative abundance levels of T-RFs indicative of Enterobacteriaceae and Aeromonadaceae species (both 496 bp) were higher at day 13 than at day 2. The T-RF indicative of members of the family Acidaminococcaceae (290 bp) occurred only at day 13 (Fig. 3B2).

TABLE 1.

Phylogenetic affiliations of representative 16S rRNA gene sequences and corresponding T-RFs^a

Organism affiliationb		Sequence,c accession no. (no. of sequences)	Organism with the closest related sequence (accession no.)	Similarity (%)	Closest cultured relative (accession no.)	Similarity (%)	T-RF length (bp)
Class	Family
Alphaproteobacteria	Sphingomonadaceae	X4A5, AM712585 (2)	Sphingomonas sp. (AF385529)	92	Sphingomonas echinoides (AJ012461)	95	150
	Rhodobiaceae	B2-16, AM773982 (3)	Uncultured bacterium (AJ583166)	95	Roseospirillum parvum (AJ011919)	90	166
	Rhodospirillaceae	B2-8, AM773974	Uncultured bacterium (AJ295647)	96	Skermanella parooensis (X90760)	90	440
Deltaproteobacteria	Syntrophobacteraceae	B1-92, AM773964	Uncultured bacterium (AJ306805)	97	Syntrophobacter wolinii (X70905)	96	166
Gammaproteobacteria	Enterobacteriaceae	X13A3, AM712574 (2)	Shigella sonnei (X80726)	>99	Shigella flexneri (X96963)	>99	496
	Aeromonadaceae	G2A6, AM712567	Bacterium H4 (AY345546)	99	Tolumonas auensis (X92889)	98	496
	Aeromonadaceae	G2A13, AM712565	Uncultured bacterium (AJ318149)	92	Tolumonas auensis (X92889)	93	150
	Pseudomonadaceae	X13A11, AM712542 (2)	Pseudomonas sp. (AF098466)	100	Pseudomonas putida (AJ308312)	99	492
Actinobacteria	Acidothermaceae	X4A11, AM712581	Uncultured actinomycete (Z73383)	94	Acidothermus cellulolyticus (X70635)	90	140
Acidobacteria	Acidobacteriaceae	X4A10, AM712580 (16)	Uncultured bacterium (AY043842)	93	Acidobacterium capsulatum (D26171)	92	96
	Acidobacteriaceae	B1-69, AM773947	Uncultured bacterium (AJ000986)	94	Acidobacterium capsulatum (D26171)	93	116
	Acidobacteriaceae	B2-33, AM773995 (2)	Uncultured bacterium (AF523892)	97	Acidobacterium capsulatum (D26171)	94	266
Sphingobacteria	Flexibacteraceae	B1-26, AM773619	Uncultured bacterium (AY218620)	85	Flexibacter canadensis (M62793)	86	166
Clostridia	Clostridiaceae	G13A6, AM712563 (5)	Uncultured bacterium (AY212768)	>99	Clostridium puniceum (X71857)	>99	520
	Acidaminococcaceae	G13A4, AM712561 (2)	Uncultured bacterium (AY524568)	95	Acetonema longum (M61919)	89	290
Chlorobia	Chlorobiaceae	B1-60, AM773938	Uncultured bacterium (AF445728)	89	Chlorobium limicola (Y101640)	78	440

^aThe phylogenetic affiliations of representative 16S rRNA gene sequences and the corresponding T-RFs are those of organisms shown in Fig. 3 and 4.

^bSequence length, ∼700 to 800 bp.

^cFor an explanation of sequence identifiers, see the legend to Fig. 4.

An archaeal 16S rRNA gene library was generated from RNA preparations after 13 days of incubation, to identify potentially active Archaea. Sixteen clones were screened by RFLP, and six representative clones were sequenced (Fig. 5). Archaeal 16S rRNA sequences clustered with the 16S rRNA sequences of Methanobacterium subterraneum (accession no. X99044; 95% sequence similarity), Methanosarcina thermophila (accession no. M59140; 98% sequence similarity), and Methanosarcina barkeri (accession no. AJ012094; 97% sequence similarity) and the phylum Crenarchaeota group I.3 (94 to 99% sequence similarity [Fig. 5]).

FIG. 5.

Consensus phylogenetic tree of archaeal 16S rRNA sequences (bold) obtained at day 13 of the [¹³C]xylose experiment. Accession numbers are provided after sequence identifiers, and relative abundance levels (percentages) of OTUs in gene library X13 are given in parentheses. Scale bar represents 10% sequence dissimilarity. The outgroup is Thermoplasma acidophilum (accession no. M20822).

DISCUSSION

Most of the phylogenetic groups detected by RNA-based stable isotope probing (RNA-SIP) in this study (Table 1 and Fig. 3 and 4) are widespread in peatlands (e.g., 8, 21, 27). Species of Acidobacteria accounted for 23%, Deltaproteobacteria for 17%, and Alphaproteobacteria for 16% of the 16S rRNA gene sequences detected in soil from the Ljubljana Marsh (Slovenia) and are hypothesized to represent the most abundant phylogenetic groups there (27). The main groups of 16S rRNA gene sequences retrieved from an acidic sphagnum peat bog (pH 3.9 to 4.5) in western Siberia were affiliated with organisms of Acidobacteria, Alphaproteobacteria, Verrucomicrobia, Actinobacteria, and Deltaproteobacteria (8). Thus, Acidobacteria species are readily detectable and, hence, potentially abundant in peatlands. Indeed, many sequences obtained in the present study from “heavy” RNA during the SIP were affiliated with Acidobacteria (Table 1, Fig. 4). However, little is known about their function in the flow of carbon in wetlands. The nearest cultured relative (based on 16S rRNA gene similarities) of the Acidobacteria sequences detected was Acidobacterium capsulatum, an organism that can use xylose and glucose as the sole carbon sources at moderately acidic pH levels (3-6) but that grows only aerobically (23). The relative abundance of a 266-bp T-RF indicative of Acidobacterium sp. increased in “heavy” RNA during the incubation of fen material with [¹³C]xylose (Fig. 3A1 and A2). Such findings suggest that certain Acidobacteriaceae species survived for a minimum of 13 days and/or utilized xylose under anoxic conditions.

A novel facultative, moderately acid-tolerant (growth at pH 4 to 7) species within the family Rhodospirillaceae was isolated from a mesotrophic Siberian fen (45). Sequences affiliated with Rhodospirillaceae were retrieved from the “heavy” fraction of RNA in the [¹³C]glucose experiment. A T-RF indicative of such organisms (440 bp; Table 1, Fig. 3 and 4) had higher relative abundance levels after prolonged incubation in both the [¹³C]glucose and [¹³C]xylose experiments.

The relative abundance of Actinomycetes-related T-RFs (140 bp; Fig. 3 and 4) is consistent with the occurrence of anaerobic and facultative Actinomycetes species (15). Other recognized families that are capable of hexose and/or pentose fermentation include Acidaminococcaceae (obligate anaerobes), Aeromonadaceae (facultative aerobes), Clostridiaceae (obligate anaerobes), and Enterobacteriaceae (facultative aerobes) (4, 9, 15). Sequences indicative of these organisms were detected by RNA-SIP (Fig. 3 and 4). Some members of the family Acidaminococcaceae are capable of glucose fermentation and production of acetate (10, 22). For example, the acetogen Acetonema longum converts glucose stoichiometrically to acetate via the acetyl-coenzyme A pathway (10, 22). 16S rRNA sequences retrieved from the “heavy” RNA of the [¹³C]glucose experiment affiliated with A. longum and T-RFs indicative of such sequences had higher relative abundance levels at the late time point (Fig. 3B1 and B2), indicating that members of the family Acidaminococcaceae are likely involved in the metabolism of glucose in anoxic fen slurries. No sequences affiliated with A. longum were retrieved from “heavy” RNA in the [¹³C]xylose experiment (Fig. 3A1 and A2), an observation that agrees with the fact that A. longum is unable to use xylose (22).

Most of the 16S rRNA sequences retrieved from the “heavy” RNA of the [¹³C]glucose and [¹³C]xylose incubations were affiliated with the genus Clostridium. Species of this genus are capable of the production of acetate, butyrate, and H₂/CO₂ from glucose and xylose (e.g., see reference 9) and occur in acidic bogs (e.g., see reference 28), which is consistent with the fermentation products observed with monosaccharide-supplemented fen slurries (Fig. 1) and with the high relative abundance level of T-RFs indicative of Clostridia organisms in “heavy” RNA (520 bp; Fig. 3A1, A2, and B1). A significant drift of clostridial [¹²C]RNA molecules into the “heavy” fractions is unlikely because clostridial species have a relatively low G+C content (9, 15).

Products indicative of mixed acidic and butyrate fermentations (i.e., formate, acetate, butyrate, CO₂, and H₂) were formed in xylose- and glucose-supplemented fen slurries. Shigella and Tolumonas species are known to ferment sugars (4). Shigella- and Tolumonas-related RNAs were labeled with [¹³C]xylose and [¹³C]glucose (Fig. 3 and 4), indicating that these monosaccharides were degraded by these organisms in anoxic fen slurries. Sequences closely related to those of Shigella sonnei and Tolumonas auensis were detected (Table 1). S. sonnei is a facultative, acid-resistant member of the family Enterobacteriaceae that utilizes mixed acid fermentation to convert monosaccharides to formate, acetate, and ethanol without the production of gases (4). Formate, acetate, and ethanol are also fermentation products of T. auensis, which is a facultative member of the family Aeromonadaceae (4).

T-RFs of 150 bp, indicative of the family Sphingomonadaceae, had a high relative level of abundance in the [¹³C]RNA-dominated fractions of the xylose and glucose experiments at the early time points (Fig. 3 and 4). The nearest cultured relative (based on 16S rRNA gene similarities) of the detected Sphingomonadaceae sequences was Sphingomonas echinoides (95% similarity; Table 1), indicating that a potentially new genus of the family Sphingomonadaceae was detected. Species of Sphingomonas grow on monosaccharides (e.g., xylose and glucose) under oxic conditions (5), and one genus within the family Sphingomonadaceae (Zymomonas sp.) ferments monosaccharides (e.g., glucose [5]). Thus, it is hypothesized that new species of this family assimilated xylose- and glucose-derived carbon in anoxic fen slurries.

The acidic fen slurries produced amounts of CH₄ that were similar to those of another acidic peat (0.4 to 1.7 μmol g_dw⁻¹ day⁻¹) from the same region in the Fichtelgebirge (18). The transient production and subsequent consumption of formate were concomitant to the accumulation of CH₄ (Fig. 1), suggesting that formate was a precursor of CH₄. These results indicate that fermentation processes and methanogenesis were active in anoxic fen slurries and were trophically linked. RNA of Methanobacteriaceae, Methanosarcinaceae, and Crenarchaeota organisms was detected at day 13 in the [¹³C]xylose experiment, indicating that the anoxic fen slurries harbor active acid-tolerant methanogens and Crenarchaeota species. DNA but not RNA of Methanobacteriaceae or Methanosarcinaceae species has also been detected in other acidic wetlands (e.g., 6, 18, 25). Methanosarcinaceae species reduce acetate to CH₄ (3), indicating that a net consumption of acetate might have occurred during the [¹³C]xylose experiment. Knowledge about the physiology of mesophilic Crenarchaeota species is incomplete, mainly because only a single mesophilic crenarchaeal isolate (i.e., an aerobic, autotrophic ammonia oxidizer) has been cultivated (24). Detection of the phylum Crenarchaeota group 1.3 RNA in fen slurries after 13 days of anoxic incubation (Fig. 5) is consistent with the widespread occurrence of this crenarchaeal group in anoxic habitats (e.g., flooded soils and sediments) (41) and suggests that members of this crenarchaeal group are capable of anaerobiosis.