Characterization of a nitrate-respiring bacterial community using the nitrate reductase gene (narG) as a functional marker

Lisa G. Gregory{dagger}, Philip L. Bond, David J. Richardson and Stephen Spiro

School of Biological Sciences, University of East Anglia, Norwich, NR4 7TJ, UK

Correspondence
Stephen Spiro
s.spiro{at}uea.ac.uk


   ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Bacterial cultures capable of reducing nitrate to nitrite, or of complete denitrification, were established from 5, 10, 15 and 20 cm depths of a freshwater sediment. Taxonomic analysis of the 56 isolates using 16S rRNA gene sequences revealed an unexpected species richness, which included representatives of the {gamma}-Proteobacteria, Bacillus spp., Staphylococcus spp. and members of the Actinobacteria. Gram-positive species tended to predominate in the lower depths of the sediment, where there was evidence of active sulphate respiration. Sequences (from the narG gene) potentially encoding the catalytic subunit of the membrane-associated nitrate reductase were successfully amplified from 46 of the isolates, using a nested PCR with four degenerate primers. NarG sequences clustered into three major groupings that were supported by alternative phylogenetic analyses. The NarG sequences from Gram-positive isolates (according to rRNA gene phylogeny) clustered together within sequences from the low-G+C Gram-positive bacteria. However, this cluster also included two sequences from members of the genus Pseudomonas. Another group contained mostly NarG sequences from the Proteobacteria (according to rRNA gene phylogeny), but also included five sequences from Gram-positive species. The third group of NarG sequences contained three sequences from Gram-positive species. Thus, the NarG-derived phylogeny is not entirely consistent with 16S rRNA-based taxonomy, precluding the use of the narG gene as a taxonomically useful tool for the characterization of nitrate-respiring bacteria. Total DNA was also extracted from the four depth intervals of the sediment sample and used in similar narG amplifications. Most sequences amplified directly from environmental DNA clustered in the Gram-negative group, and none was in the predominantly Gram-positive group. The study also revealed a degree of spatial organization of a nitrate-respiring community in terms of both microbiology and narG sequences.

Abbreviations: LGCGP, low-G+C Gram-positive bacteria

The EMBL accession numbers for the nar sequences reported in this paper are AJ314921–AJ314996; the EMBL accession numbers for the 16S rRNA gene sequences are AJ489332–AJ489384.

{dagger}Present address: Department of Chemical Engineering, Yale University, New Haven, CT 06520-8286, USA.


   INTRODUCTION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Denitrification, the respiratory reduction of nitrate to gaseous products, is an important component of the nitrogen cycle, which influences soil and water fertility, atmospheric chemistry, and waste and water treatment processes (Knowles, 1982; Zumft, 1992). Complete denitrification requires the sequential action of four enzymes, nitrate reductase, nitrite reductase, nitric oxide (NO) reductase and nitrous oxide (N2O) reductase. In at least three cases, biochemically distinct enzymes catalyse the same reaction in the denitrification pathway. Respiratory nitrate reduction can be catalysed either by a membrane-associated or by a soluble periplasmic nitrate reductase; some bacteria express both enzymes, which can have different physiological roles (Richardson et al., 2001). Respiratory nitrite reduction (to NO) is catalysed by a copper nitrite reductase or a cytochrome cd1 nitrite reductase, but no species is known that can express or has the genes for both enzymes (Watmough et al., 1999). NO reduction can be catalysed by at least three distinct enzymes, two-subunit enzymes that accept electrons either from soluble c-type cytochromes or from menaquinol and a single-subunit enzyme that accepts electrons from the quinol pool (Watmough et al., 1999; Suharti et al., 2001; Cramm et al., 1999). Denitrification is modular, in the sense that it is common for bacteria to have only a part of the pathway. Reduction of nitrate only to nitrite is not denitrification sensu stricto since it does not result in a gaseous product, but is the most commonly occurring variant in the use of N-oxyanions and oxides as terminal electron acceptors (Zumft, 1992). Reduction of nitrate or nitrite as far as N2O is also common, indeed denitrification is a major contributor to the atmospheric N2O budget. Denitrification occurs in many distantly related species of the Bacteria and Archaea (Zumft, 1992), which precludes the use of 16S rRNA-based methods for the identification and characterization of bacteria with this physiological capability. Thus, considerable effort in recent years has been put into the development of techniques to allow the genes encoding denitrification enzymes to be used as functional markers for ecological studies.

Primer pairs and PCR protocols have been developed for the amplification of genes encoding the membrane-associated and periplasmic nitrate reductases, the copper and cytochrome cd1 nitrite reductases, and the nitrous oxide reductase (Braker et al., 1998, 2000; Flanagan et al., 1999; Gregory et al., 2000; Hallin & Lindgren, 1999; Petri & Imhoff, 2000; Scala & Kerkhof, 1998, 1999). Amplifications directly from environmental DNA tend to reveal a greater degree of sequence diversity in nitrite and nitrous oxide reductase genes than is apparent in the same genes from cultured isolates (Braker et al., 2000; Scala & Kerkhof, 1998, 1999). Of the two nitrate reductases, the membrane-associated enzyme is typically involved with nitrate respiration under anoxic conditions and probably has a greater role to play in the environmental nitrogen cycle (Richardson et al., 2001). Hence, this study has focussed on the membrane-associated nitrate reductase and makes use of previously developed PCR primer systems that successfully amplify fragments of the narG gene that encodes the catalytic molybdenum-cofactor-containing subunit of the enzyme (Gregory et al., 2000). The goals of the work were two-fold. First, to determine whether phylogenies based on narG sequences are consistent with 16S rRNA-based taxonomy and thus whether nitrate reductase sequences contain useful taxonomic information about nitrate-respiring bacteria. The second objective was to exploit the primers to determine whether there is spatial organization in the nitrate-respiring community in a depth profile through a freshwater sediment. A recent study using nitrite reductase genes has suggested that there is rather little spatial variability along the vertical axis of sediments, perhaps because of mixing events (Braker et al., 2001). However, spatial organization might be expected given the gradients in, for example, oxygen, nitrate and sulphate concentrations and redox potential that typically exist in sediments.


   METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Sediment sampling and analysis.
Three replicate sediment cores (25x9 cm) were collected in November 1998 from an artificial freshwater lake (grid reference TG191071) that was dug on the University of East Anglia campus in the 1960s. The cores were stored at 4 °C in plastic bags under an atmosphere of nitrogen for no more than 16 h prior to subsequent manipulations. Cores were separated into 5 cm depth intervals (0–5, 5–10, 10–15 and 15–20 cm), and the pH and redox potential measured in each segment using a portable meter (Mettler Toledo) fitted with probes designed for field measurements. Pore water collected from each sediment depth with a sterile syringe and needle was filtered through a 0·45 µm membrane, degassed and stored at 4 °C for no more than 24 h. Concentrations of nitrate, nitrite and sulphate were determined by ion exchange chromatography (Dionex DX-100 with IonPac AS4A analytical conductivity detector).

Isolation and growth of bacteria.
For bacterial isolations, 3 ml sediment samples were taken from each depth interval of the sediment profile, resuspended in 3 ml sterile 0·1 % (w/v) sodium cholate by gentle shaking for 30 min at 4 °C, before serial dilution in the same buffer. Samples were plated onto a phosphate-buffered basal medium (pH 7·4) based on that described by Harms et al. (1985) and supplemented with KNO3 (5 g l-1), sodium succinate (13·5 g l-1), NaFeEDTA (2 mg l-1), MgCl2.6H2O (25 mg l-1), H3BO3 (2·8 mg l-1), MnSO4.H2O (1·6 mg l-1), NaMoO4.H2O (0·8 mg l-1), ZnSO4.7H2O (0·24 mg l-1) and Cu(NO3)2.3H2O (0·04 mg l-1). Plates were incubated anaerobically for 3 days at 30 °C under an atmosphere of 10 % H2, 10 % CO2 and 80 % N2. Colonies were chosen (to be representative of all colony types that could be distinguished visually) for further analysis and were purified by anoxic growth on the same medium supplemented with 0·1 % (w/v) yeast extract. Incubation of the original isolation plates for a further 48 h did not reveal any additional slow-growing colonies. Ten, 18, 15 and 13 isolates from the 0–5, 5–10, 10–15 and 15–20 cm sediment depth intervals, respectively, were used in further analyses; these were given the prefixes Lgg5, Lgg10, Lgg15 and Lgg20. Isolates were tested for their ability to evolve gas from nitrate, reaction to Gram stain and production of fluorescent pigments on King's A and B media (Stolp & Gadkari, 1992). The ability to accumulate nitrite from nitrate was tested using a chemical nitrite assay based on that described by Coleman et al. (1978). Cultures were grown anaerobically to stationary phase, and 1 µl samples were mixed with 89 µl of 1 % (w/v) sulphanilamide in 1 M HCl and 10 µl of 0·2 % (w/v) aqueous N-naphthylethylene diamine dihydrochloride in 96-well microtitre plates. The plates were incubated at room temperature for 25 min and the absorbance read at 540 nm in a microtitre plate reader. Nitrite concentrations were estimated by comparison to standards.

Determination of rRNA gene sequences.
Chromosomal DNA extracted from stationary phase cultures using Wizard Genomic DNA purification kits (Promega) was used in PCRs with 16S rRNA primers pA (nucleotides 8–28 in the Escherichia coli 16S rRNA) and pH' (nucleotides 1542–1522; Edwards et al., 1989). Reaction mixes (50 µl) contained 10–100 ng genomic DNA, 10 pmol each primer, 0·2 mM dNTP mix (Bioline), 3 mM MgCl2, 1x times; Expand High Fidelity PCR Buffer (Roche) and 1·3 U Expand High Fidelity Taq DNA polymerase (Roche). Reactions were denatured at 94 °C for 5 min, then cycled 26 times at 94 °C for 40 s, 55 °C for 1 min and 72 °C for 2 min, prior to a final extension at 72 °C for 10 min. PCR products were gel-purified using the QIAEX II Agarose Gel Extraction Kit (Qiagen) and sequenced using the Big Dye Terminator Reaction Mix (Amersham) with primers pC' (nucleotides 361–341) and pD' (nucleotides 536–519; Edwards et al., 1989). Reactions (20 µl) contained 30–90 ng PCR product, 10 pmol primer and 4 µl of the reaction mix, and were subjected to 30 cycles of 96 °C for 30 s, 45 °C for 5 s and 60 °C for 4 min. The reactions were 2-propanol-precipitated, dried and resolved on an ABI automated sequencer at the John Innes Centre (Norwich, UK).

Extraction of DNA from sediment samples.
A method based on that described by Bruce et al. (1992) was used to extract total DNA from the same sediment samples from which the bacterial isolates had been cultured. Two grams (wet weight) of sediment sample were mixed with 5 ml extraction buffer (1 % SDS in 0·12 M Na2HPO4, pH 8·0) and incubated at 70 °C for 1 h with occasional shaking. The sample was centrifuged at 2800 g for 10 min at 4 °C and the resulting supernatant was stored on ice. The pellet was resuspended in 5 ml fresh extraction buffer and incubated as before. This extraction process was repeated and the three supernatant fractions were pooled and then centrifuged at 8000 g for 30 min. The supernatant was added to an equal volume of 30 % (w/v) polyethylene glycol in TE buffer (10 mM Tris/HCl, 1 mM Na2EDTA, pH 8·0), and 0·1 volume of 5 M NaCl was added. After overnight precipitation at 4 °C, the sample was centrifuged (5000 g, 10 min); the resulting pellet was dissolved in 8 ml TE buffer containing 8 g CsCl and 100 µl ethidium bromide (10 mg ml-1). The sample was centrifuged at 50 000 r.p.m. for 18 h at 18 °C in a Beckmann Ti70 rotor. Under UV light, a single band of DNA was visible and was withdrawn using a sterile syringe. The ethidium bromide was removed by extraction with CsCl-saturated 2-propanol, and the DNA was dialysed against TE buffer, extracted with an equal volume of phenol/chloroform and concentrated by ethanol precipitation. The solution containing the DNA was straw-coloured due to contaminating humic substances that co-extract with DNA (Tsai & Olson, 1992). Humic substances (which can inhibit PCRs) were removed by chromatography through a Sephacryl-100 HR gel matrix (Tsai & Olson, 1992). Five millilitres of Sephacryl-100 HR gel matrix (Sigma-Aldrich) equilibrated in TE buffer were packed into a 5 ml sterile syringe plugged with 1·5 cm of glass wool. The column was centrifuged (1100 g, 10 min) in a swing-bucket rotor. An aliquot of the DNA solution (50 µl) was loaded onto the column and centrifuged (1100 g, 10 min at room temperature). The eluent was collected and pooled with other cleaned fractions from the original DNA sample and residual eluent was collected after a final column spin (1100 g, 10 min at room temperature). The DNA solution was filtered through a Centricon 100 concentrator spin column (Amicon-Fisher) according to the manufacturer's instructions; the eluent (50 µl) containing the DNA was stored at 4 °C.

Isolation of the nar and nap gene fragments.
Fragments of the narG gene were amplified from the genomic DNA of cultured isolates, or from total DNA extracted from sediment, by nested PCR using the primers T37, T38, T39 and W9 and reaction conditions that have been described previously (Gregory et al., 2000). PCR products were blunt-ended with T4 DNA polymerase (Roche), phosphorylated with T4 polynucleotide kinase (Roche) and ligated into SmaI-digested and dephosphorylated pUC18 (Pharmacia). Ligation mixtures were transformed into E. coli JM83 [ara {Delta}(lac–proAB) rpsL {phi}80 lacZ{Delta}M15] and plasmids purified from recombinant colonies with the Wizard Plus SV Minipreps DNA purification system (Promega). Clones were sequenced on both strands using the cycle sequencing protocol (above) with vector-specific ‘universal’ and ‘reverse’ primers, and were given prefixes 5, 10, 15 and 20, corresponding to the depth of the sediment from which the total DNA had been extracted. Fragments of the napA gene were amplified from the genomic DNA of cultured isolates, using the primers V16, V17, V66 and V67 and reaction conditions that have been described previously (Flanagan et al., 1999).

Phylogenetic analysis of rRNA genes.
Phylogenetic affiliations of the partial sequences were initially estimated using the program BLAST (basic local alignment search tool; Altschul et al., 1997) and available nucleotide databases. Gene sequences were reduced to unambiguously alignable positions in ARB (a software environment for sequence data; Strunk & Ludwig, 2002; http://www.arb-home.de/). Gaps and missing data were excluded, resulting in a dataset of 81 taxa and 335 nt. Evolutionary analyses of alignments were performed by distance methods using ARB (Strunk & Ludwig, 2002) and PAUP (Swofford, 1996), and by parsimony and maximum-likelihood algorithms in PAUP. Distances were calculated in ARB according to the substitution algorithm of Jukes & Cantor (1969), and phylogenetic trees were assembled by neighbour joining. Maximum-likelihood used the HYK model (Kishino & Hasegawa, 1989) with a transition-to-transversion ratio of 2. Heuristic searching was used in the parsimony and maximum-likelihood analyses.

Phylogeny of NarG amino acid sequences.
Sequences were compiled and aligned in ARB. A protein distance matrix was calculated in ARB, and identical sequences were grouped for the analyses. Gaps and missing sequence were excluded, yielding a dataset of 68 taxa and 106 aa for comparisons. Distances were calculated in ARB using the PAM substitution matrix and phylogenetic trees were inferred by neighbour joining. Bootstrapping of the distance analyses was performed in ARB. Parsimony inference of the amino acid data was performed in PAUP by heuristic search and compilation of a consensus tree.


   RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Isolation and characterization of nitrate-respiring bacteria
Three replicate 20 cm cores were cut from a freshwater sediment and divided into 5 cm intervals. The upper 3 cm was composed of small stones and sand. Particle size decreased down the core; the lowest 5 cm section was a fine black silt with a sulphurous smell, indicative of sulphate respiration. The nitrate concentration was 28±2 µM in the overlying water and approximately 5 µM at all depths of the core; nitrite was undetectable throughout. The pH of the pore water varied between 6·0 and 7·8, and was not correlated with depth.

From each sediment depth, enrichment cultures for nitrate-respiring bacteria were established, using solid media containing a non-fermentable carbon source (succinate) and 50 mM nitrate. Between 10 and 18 isolates were chosen from each depth, representing the different colony types that were distinguishable by visual examination (Table 1). Five isolates were true denitrifiers (as judged by their ability to evolve gas from nitrate), most of the remainder appeared to accumulate nitrite from nitrate, indicating an incomplete denitrification pathway (Table 1). These isolates can presumably utilize nitrate as a terminal electron acceptor (reducing it only as far as nitrite) to allow growth on the non-fermentable carbon source. Some of the cultures grew to rather low final densities (OD600<0·2 in stationary phase) in liquid cultures; most of these isolates also accumulated relatively high concentrations of nitrite (Table 1), which was possibly inhibiting further growth. For most of the isolates, approximately 400 nt of 16S rRNA gene sequence information was generated, which was compared to other rRNA gene sequences in the GenBank database. In most cases the sequences were>97 % identical to previously reported sequences. Phylogenetic analysis of the 16S rRNA gene sequences (see below) revealed considerable species diversity. Gram-positive species tended to predominate at the lower sediment depths and Gram-negative species predominated in the upper sections.


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Table 1. Characteristics of strains isolated from the freshwater sediment

 
Characterization of narG sequences
Of the 56 strains used in this study, 46 were positive in the narG PCR amplification; all of those that were negative were from Gram-positive genera (Table 1). Amplification products were cloned and at least 366 nt were sequenced on both strands. Comparison with sequence databases verified that these clones potentially encoded nitrate reductase sequences. The sequences were conceptually translated and used to assemble an alignment of Nar sequences, which included all other known and predicted Nar sequences from laboratory strains and from genome sequences. The alignment was used to construct a dendrogram, using distance matrix methods, and the tree topology was scrutinized by parsimony and bootstrap analyses (Fig. 1). Major lineages supported by the analyses were the archaeal lineage containing Aeropyrum pernix and Pyrobaculum aerophilum, the Actinobacteria containing Mycobacterium tuberculosis and Streptomyces coelicolor, the clade representing the low-G+C Gram-positive bacteria (LGCGP) division, and the Proteobacteria lineage (Fig. 1). The dendrogram shows that most of the NarG sequences obtained in this study fall into the LGCGP and the Proteobacteria. These clades were supported by both parsimony and distance phylogenetic inferences and bootstrap values (Fig. 1). Within the Proteobacteria, subclades that are well supported include those containing the E. coli NarA and NarZ sequences (E. coli has two isoenzymes of the membrane-associated nitrate reductase).



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Fig. 1. (on facing page) Phylogenetic relationship of NarG amino acid sequences with representative nitrate reductase sequences from the databases. The dendrogram was generated using distance-matrix and neighbour-joining methods, and rooted thereafter at the branch, making Aeropyrum pernix and P. aerophilum the outgroup. Reference strain names are followed by accession numbers. The scale bar represents 0·1 substitutions per site. Values at nodes represent the percentage of replicate trees that support the branching order; only bootstrap values greater than 59 % are shown. Branch points supported by distance and parsimony estimations are indicated by open circles. Cultured isolates are prefixed Lgg5, Lgg10, Lgg15 and Lgg20, corresponding to enrichments from the 5, 10, 15 and 20 cm sediment depths, respectively. Clones amplified directly from environmental DNA are designated with the prefixes clone 5, clone 10, clone 15 and clone 20, likewise indicating the depth of the sediment from which DNA was extracted. Identical sequences are indicated as multiple entries on the nodes, except the sequence clone 10.6, which represents clone sequences 5.1, 5.2, 5.3, 5.5, 5.8, 5.9, 5.10, 5.11, 5.12, 10.8, 10.9 and 15.9. Phylogenetic placement of isolates that differ at the interphyla level, according to the NarG and 16S rRNA analyses, is indicated by shaded boxes.

 
Comparison of the NarG and 16S rRNA gene phylogenies
Phylogenetic comparisons were performed on the partial 16S rRNA gene sequences generated from the isolates obtained in this study (Fig. 2). Representative sequences utilized in these analyses included those from the same strains used in the NarG sequence phylogeny. The tree is well supported by the three phylogenetic methods used. Well-supported major clades include those containing the Proteobacteria, the {gamma}-Proteobacteria, the LGCGP and the Actinobacteria (Fig. 2). Comparison of the 16S rRNA and the NarG phylogenies reveals that the two are partially congruent. Of the {gamma}-Proteobacteria, 87 % (13 out of 15) were correctly predicted as such by narG sequence analysis. Of the LGCGP, 69 % (18 out of 26) were correctly predicted. Therefore, narG sequence information cannot be used with confidence to predict the taxonomic position of organisms as defined by 16S rRNA gene sequences. Some strains have identical NarG sequences but are well separated by 16S rRNA phylogeny (including the sets of strains Lgg10.10, 10.15 and 10.16, Lgg10.14 and 20.9, and the set Lgg10.7 and 20.7). The isolate Lgg15.6, placed in the Actinobacteria by 16S rRNA phylogeny, clustered in an unaffiliated clade by narG sequence analyses. Other less-contrasting discrepancies between the two phylogenies occurred. For example, in contrast to 16S rRNA analyses, Lgg5.11 and Lgg15.7 were both placed within what may be a Bacillus cereus clade (no narG reference available) by narG sequence analyses. Horizontal transfer of the narG gene might provide an explanation for the occurrence of very similar narG alleles in distantly related bacteria. Such evidence for gene transfer could be verified by including more reference narG sequences and additional gene sequence data. The 16S rRNA and narG sequences of Lgg5.3, 10.7, 5.12, 10.6, 10.10, 10.14, 10.15 and 15.1 were re-amplified and re-sequenced, confirming that their positions on the dendrogram were not due to sequencing errors.



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Fig. 2. Phylogenetic relationship of 16S rRNA gene sequences with representative sequences from the databases. Representative sequences used in Figs 1 and 2 are taken from the same strain. The dendrogram was generated using distance-matrix and neighbour-joining methods, and rooted thereafter at the branch with P. aerophilum and Archaeoglobus fulgidus as the outgroup. Accession numbers for sequences from reference strains are indicated. The scale bar represents 0·1 substitutions per site. Branch points supported by distance, maximum-likelihood and parsimony estimations are indicated by solid circles. Marginally supported branch points (supported by two of the phylogenetic analyses) are indicated by open circles. Branch points without circles are not supported by the majority of analyses. Cultured isolates are prefixed Lgg5, Lgg10, Lgg15 and Lgg20, corresponding to enrichments from the 5, 10, 15 and 20 cm sediment depths, respectively. Phylogenetic placement of isolates that differ at the interphyla level, according to the NarG and 16S rRNA analyses, is indicated by shaded boxes.

 
Characterization of narG sequences in community DNA
Total DNA was extracted from each of the sediment core sections and was used as the template in PCRs for the amplification of narG sequences. A total of 28 clones selected at random were sequenced, and the translated sequences were included in the alignment and dendrogram (Fig. 1). Eighteen of the sequences were closely related to one another and fell in a group containing the NarA nitrate reductase of E. coli (Fig. 1). With one exception, all 18 sequences were from the 5 or 10 cm depths and were also closely related to two sequences from cultured isolates (Lgg5.12 and 5.1). The remaining sequences were more diverse, a further seven were spread within the Proteobacteria group, and four (all from the 15 cm depth of sediment) formed a deep branching clade not affiliated with any known NarG sequences (Fig. 1). The sequences from the 15 cm depth were more diverse than those from the 5 and 10 cm depths. Interestingly, of all the sequences cloned from environmental DNA, none fell into the LGCGP group (Fig. 1).

Detection of napA genes
The bulk of nitrate respiration in anoxic environments is usually attributed to the membrane-bound nitrate reductase, though the contribution that is made by the periplasmic enzyme is not known and has not been extensively studied. To evaluate the abundance of the periplasmic nitrate reductase gene, napA, in the sediment nitrate-respiring community, genomic DNAs were used as templates for the amplification of napA fragments (Flanagan et al., 1999). A total of 12 isolates from the 5 and 10 cm depths gave napA amplification products (Table 1), including three from Gram-positive species (Lgg5.1, 5.11 and 10.6). Only two Gram-negative isolates from the lower two depths were positive in the napA amplification reaction (Lgg15.13 and 15.14). It is possible that strains potentially expressing the periplasmic enzyme might predominate in the upper layers of the sediment, since this enzyme would allow the bacteria to derive a physiological benefit from nitrate respiration in the presence of low concentrations of oxygen.


   DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
An enrichment protocol designed to isolate denitrifying bacteria from a freshwater sediment led to the culture of bacteria that, in most cases, could reduce nitrate only as far as nitrite. This is consistent with the observation of Zumft (1992) that reduction of nitrate only as far as nitrite is the mostly widely distributed variant in the use of N-oxyanions and oxides as electron acceptors for respiration. A rationale for the predominance of this physiology in the sediment environment might be provided by the high concentration of sulphate, and activity of sulphate-reducing bacteria (as evidenced by black colouration and sulphurous odour). Sulphide, the product of sulphate respiration, is believed to inhibit the reduction of NO to N2O in denitrifying bacteria (Knowles, 1982) and so might limit the abundance of species with the complete denitrification pathway.

This study has revealed a somewhat unexpected species diversity and spatial organization in a sediment community potentially capable of nitrate reduction. Spatial organization extends to the type of nitrate reductase genes detected, since the napA gene encoding the periplasmic nitrate reductase was more abundant in the upper layers of the sediment. This enzyme has a variety of physiological roles in different organisms, including catalysing the first step of a true anaerobic denitrification pathway, scavenging low concentrations of nitrate, providing the apparatus for nitrate respiration in the presence of oxygen and allowing for the disposal of reducing equivalents during growth on reduced carbon substrates (Richardson et al., 2001). The physico-chemical make-up of the upper layers of the sediment may therefore provide an environment in which nitrate respiration catalysed by Nap is favoured. Analyses of denitrifying communities using PCR primers targeted against the nirS and nirK genes encoding nitrite reductase and the nitrous oxide reductase gene nosZ have also considerably expanded the previously known sequence diversity (Braker et al., 1998, 2000; Hallin & Lindgren, 1999; Petri & Imhoff, 2000; Scala & Kerkhof, 1998, 1999; Rösch et al., 2002).

In all known cases, the bacterial membrane-bound nitrate reductase is encoded in a polycistronic transcription unit that includes the narH gene that encodes an iron–sulphur protein (Richardson et al., 2001). A recent study has examined the relatedness of narH sequences in nitrate-respiring bacteria and concluded that narH- and 16S rRNA-based phylogenies are largely congruent (Petri & Imhoff, 2000), unlike the narG and 16S rRNA phylogenies. The reason for the different outcomes from the narH and narG analyses is not clear, though rather more Gram-positive sequences are included in the narG tree (26) than the four examined by Petri & Imhoff (2000). Results for narG are consistent with the possibility of this gene being subject to horizontal transfer. Interestingly, genes encoding the membrane-bound nitrate reductase are plasmid-encoded in Thermus thermophilus and can move between Thermus strains by conjugation (Ramirez-Arcos et al., 1998). Thus, there is some evidence to indicate that horizontal transfer of nar genes is possible. Amongst other genes involved in the respiratory reduction of nitrogen compounds, there is some evidence for lateral transfer of the nirK gene between denitrifying bacteria, though not between ammonia oxidizers (Casciotti & Ward, 2001). For genes involved in other reactions of the nitrogen cycle, a phylogeny derived from the amoA gene encoding the ammonia monooxygenase of nitrifying bacteria was largely consistent with 16S rRNA taxonomy (Aakra et al., 2001). For nitrogen fixation genes, there is considerable discussion as to whether or not nif gene phylogenies provide evidence for horizontal gene transfer (Hirsch et al., 1995). One of the goals of this work was to determine whether the narG gene could be used to infer taxonomic information about nitrate-respiring and denitrifying bacteria. The lack of congruence between the narG and 16S rDNA trees suggests that this cannot be done with confidence.


   ACKNOWLEDGEMENTS
 
We are grateful to Hans Schutten for help with the sediment sampling. L. G. G. was the recipient of a studentship from the Natural Environment Research Council.


   REFERENCES
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
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Received 4 July 2002; revised 18 September 2002; accepted 23 September 2002.



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