School of Biological Sciences, University of East Anglia, Norwich, NR4 7TJ, UK
Correspondence
Stephen Spiro
s.spiro{at}uea.ac.uk
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Abbreviations: LGCGP, low-G+C Gram-positive bacteria
The EMBL accession numbers for the nar sequences reported in this paper are AJ314921AJ314996; the EMBL accession numbers for the 16S rRNA gene sequences are AJ489332AJ489384.
Present address: Department of Chemical Engineering, Yale University, New Haven, CT 06520-8286, USA.
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
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 |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
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 05, 510, 1015 and 1520 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 828 in the Escherichia coli 16S rRNA) and pH' (nucleotides 15421522; Edwards et al., 1989). Reaction mixes (50 µl) contained 10100 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 361341) and pD' (nucleotides 536519; Edwards et al., 1989
). Reactions (20 µl) contained 3090 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
(lacproAB) rpsL
80 lacZ
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 |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
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.
|
|
|
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 |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
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 ironsulphur 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 |
---|
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Altschul, S. F., Madden, T. L., Schaffer, A. A., Zhang, J., Zhang, Z., Miller, W. & Lipman, D. J. (1997). Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res 25, 33893402.
Braker, G., Fesefeldt, A. & Witzel, K.-P. (1998). Development of PCR primer systems for amplification of nitrite reductase genes (nirK and nirS) to detect denitrifying bacteria in environmental samples. Appl Environ Microbiol 64, 37693775.
Braker, G., Zhou, J., Wu, L., Devol, A. H. & Tiedje, J. M. (2000). Nitrite reductase genes (nirK and nirS) as functional markers to investigate diversity of denitrifying bacteria in Pacific Northwest marine sediment communities. Appl Environ Microbiol 66, 20962104.
Braker, G., Ayala-del-Rio, H. L., Devol, A. H., Fesefeldt, A. & Tiedje, J. M. (2001). Community structure of denitrifiers, bacteria, and archaea along redox gradients in Pacific Northwest marine sediments by terminal restriction fragment length analysis of amplified nitrite reductase (nirS) and 16S rRNA genes. Appl Environ Microbiol 67, 18931901.
Bruce, K. D., Hiorns, W. D., Hobman, J. L., Osborn, A. M., Strike, P. & Ritchie, D. A. (1992). Amplification of DNA from native populations of soil bacteria by using the polymerase chain reaction. Appl Environ Microbiol 58, 34133416.[Abstract]
Casciotti, K. L. & Ward, B. B. (2001). Dissimilatory nitrite reductase genes from autotrophic ammonia-oxidizing bacteria. Appl Environ Microbiol 67, 22132221.
Coleman, K. J., Cornish-Bowden, A. & Cole, J. A. (1978). Purification and properties of nitrite reductase from Escherichia coli K12. Biochem J 175, 483493.[Medline]
Cramm, R., Pohlmann, A. & Friedrich, B. (1999). Purification and characterization of the single-component nitric oxide reductase from Ralstonia eutropha H16. FEBS Lett 460, 610.[CrossRef][Medline]
Edwards, U., Rogall, T., Blocker, H., Emde, M. & Bottger, E. C. (1989). Isolation and direct complete nucleotide determination of entire genes. Characterization of a gene coding for 16S ribosomal RNA. Nucleic Acids Res 17, 78437853.[Abstract]
Flanagan, D. A., Gregory, L. G., Carter, J. P., Karakas-Sen, A., Richardson, D. J. & Spiro, S. (1999). Detection of genes for periplasmic nitrate reductase in nitrate respiring bacteria and in community DNA. FEMS Microbiol Lett 177, 263270.[CrossRef][Medline]
Gregory, L. G., Karakas-Sen, A., Richardson, D. J. & Spiro, S. (2000). Detection of genes for membrane-bound nitrate reductase in nitrate-respiring bacteria and in community DNA. FEMS Microbiol Lett 183, 275279.[CrossRef][Medline]
Hallin, S. & Lindgren, P.-E. (1999). PCR detection of genes encoding nitrite reductase in denitrifying bacteria. Appl Environ Microbiol 65, 16521657.
Harms, N., de Vries, G. E., Maurer, K., Veltkamp, E. & Stouthamer, A. H. (1985). Isolation and characterization of Paracoccus denitrificans mutants with defects in the metabolism of one-carbon compounds. J Bacteriol 164, 10641070.[Medline]
Hirsch, A. M., McKhann, H. I., Reddy, A., Liao, J., Fang, Y. & Marshall, C. R. (1995). Assessing horizontal transfer of nifHDK genes in eubacteria: nucleotide sequence of nifK from Frankia strain HFPCcI3. Mol Biol Evol 12, 1627.[Abstract]
Jukes, T. H. & Cantor, C. R. (1969). Evolution of protein molecules. In Mammalian Protein Metabolism, pp. 21132. Edited by H. N. Munro. New York: Academic Press.
Kishino, H. & Hasegawa, M. (1989). Evaluation of the maximum likelihood estimate of the evolutionary tree topologies from DNA sequence data, and the branching order in Hominoidea. J Mol Evol 29, 170179.[Medline]
Knowles, R. (1982). Denitrification. Microbiol Rev 46, 4370.
Petri, R. & Imhoff, J. F. (2000). The relationship of nitrate reducing bacteria on the basis of narH gene sequences and comparison of narH and 16S rDNA based phylogeny. Syst Appl Microbiol 23, 4757.[Medline]
Ramirez-Arcos, S., Fernandez-Herrero, L. A., Marin, I. & Berenguer, J. (1998). Anaerobic growth, a property horizontally transferred by an Hfr-like mechanism among extreme thermophiles. J Bacteriol 180, 31373143.
Richardson, D. J., Berks, B. C., Russell, D. A., Spiro, S. & Taylor, C. J. (2001). Functional, biochemical and genetic diversity of prokaryotic nitrate reductases. Cell Mol Life Sci 58, 165178.[Medline]
Rösch, C., Mergel, M. & Bothe, H. (2002). Biodiversity of denitrifying and dinitrogen-fixing bacteria in an acid forest soil. Appl Environ Microbiol 68, 38183829.
Scala, D. J. & Kerkhof, L. J. (1998). Nitrous oxide reductase (nosZ) gene-specific PCR primers for detection of denitrifiers and three nosZ genes from marine sediments. FEMS Microbiol Lett 162, 6168.[CrossRef][Medline]
Scala, D. J. & Kerkhof, L. J. (1999). Diversity of nitrous oxide reductase (nosZ) genes in continental shelf sediments. Appl Environ Microbiol 65, 16811687.
Stolp, H. & Gadkari, D. (1992). Non-pathogenic members of the genus Pseudomonas. In The Prokaryotes, 2nd edn, pp. 719741. Edited by A. Balows, H. G. Trüper, M. Dworkin, W. Harder & K.-H. Schleifer. New York: Springer-Verlag.
Strunk, O. & Ludwig, W. (2002). ARBA software environment for sequence data. Department of Microbiology, Technical University of Munich, Germany.
Suharti Stampraad, M. J., Schröder, I. & de Vries, S. (2001). A novel copper A containing menaquinol NO reductase from Bacillus azotoformans. Biochemistry 40, 26322639.[CrossRef][Medline]
Swofford, D. L. (1996). PAUP. Phylogenetic analysis using parsimony, version 4.0b, 5th edn. Sunderland, Massachusetts: Sinauer Associates.
Tsai, Y.-L. & Olson, B.H. (1992). Rapid method for separation of bacterial DNA from humic substances in sediments for polymerase chain reaction. Appl Environ Microbiol 58, 22922295.[Abstract]
Watmough, N. J., Butland, G., Cheesman, M. R., Moir, J. W. B., Richardson, D. J. & Spiro, S. (1999). Nitric oxide in bacteria: synthesis and consumption. Biochim Biophys Acta 1411, 456474.[Medline]
Zumft, W. G. (1992). The denitrifying prokaryotes. In The Prokaryotes, 2nd edn, pp. 554582. Edited by A. Balows, H. G. Trüper, M. Dworkin, W. Harder & K.-H. Schleifer. New York: Springer-Verlag.
Received 4 July 2002;
revised 18 September 2002;
accepted 23 September 2002.
HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
INT J SYST EVOL MICROBIOL | MICROBIOLOGY | J GEN VIROL |
J MED MICROBIOL | ALL SGM JOURNALS |