| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
1 Department of Rhizosphere Biology, GSF – National Research Centre for Environment and Health, Ingolstaedter Landstrasse 1, D-85764 Neuherberg, Germany
2 Institute of Soil Ecology, GSF – National Research Centre for Environment and Health, Ingolstaedter Landstrasse 1, D-85764 Neuherberg, Germany
Correspondence
Anton Hartmann
anton.hartmann{at}gsf.de
 |
ABSTRACT |
|---|
 TOP ABSTRACT MAIN TEXTREFERENCES |
|---|
7c, C16 : 0 and C18 : 17c, and the DNA G+C content ranged from 60.9 to 61.5 mol%. Therefore these three isolates should be classified within a novel species, for which the name Herbaspirillum hiltneri sp. nov. is proposed. The type strain is N3T (=DSM 17495T=LMG 23131T).
The GenBank/EMBL/DDBJ accession numbers for the 16S rRNA gene sequences of strains N3T, N5 and N9 are DQ150563–DQ150565, respectively, and those for the 23S rRNA gene sequences of strains N3T, N5 and N9 are DQ150552–DQ150554.
A similarity matrix based on 16S and 23S rRNA gene sequences of Herbaspirillum species is available as supplementary material in IJSEM Online.
Present address: Institute of Biochemical Plant Pathology, GSF – National Research Centre for Environment and Health, Ingolstaedter Landstrasse 1, D-85764 Neuherberg, Germany. 
|
MAIN TEXT |
|---|
|
TOPABSTRACTMAIN TEXTREFERENCES |
|---|
). The genus was first described with the species Herbaspirillum seropedicae (Baldani et al., 1986), which has been isolated from rice, maize and sorghum plants. After detailed taxonomic studies, the mildly plant-pathogenic [Pseudomonas] rubrisubalbicans was reclassified to the genus Herbaspirillum as Herbaspirillum rubrisubalbicans (Baldani et al., 1996). Another species of this genus, Herbaspirillum frisingense, was isolated from the C4-fibre plants Miscanthus spp., Spartina spectinata and Pennisetum purpureum by Kirchhof et al. (2001). From root nodules of garden beans (Phaseolus vulgaris), several strains representing one distinct species were isolated and described as Herbaspirillum lusitanum (Valverde et al., 2003). These root-colonizing Herbaspirillum species were detected not only on the surface of the root, but also as intra- and intercellular colonizers in the root interior (Olivares et al., 1997). These bacteria seem to prefer plants of the family Gramineae as hosts (Kirchhof et al., 2001), but they are also found on other plant species (Baldani et al., 1996). According to Döbereiner et al. (1993), the close association of endophytically colonizing herbaspirilli results in an additional supply of bacterially fixed nitrogen for the host plant. Experimental evidence for this thesis was provided by Elbeltagy et al. (2001) using rice (Oryza officinalis) inoculated with Herbaspirillum sp. strain B501.
The organisms listed so far all display the ability to fix atmospheric nitrogen, but, according to the latest taxonomic data, this can no longer be considered a common feature within the genus Herbaspirillum. In Korea, a 4-chlorophenol-degrading strain was isolated from river sediments near an industrial plant (Bae et al., 1996). Because of its morphological and physiological attributes, this isolate was first thought to belong to the species Comamonas testosteroni (Bae et al., 1996), but further phylogenetic analyses based on 16S rRNA gene sequences, as well as DNA–DNA hybridization experiments, indicated that this classification was incorrect; the name Herbaspirillum chlorophenolicum was proposed by Im et al. (2004) for this non-plant-associated strain. Ding & Yokota (2004) discovered a novel species in well water, and described it as Herbaspirillum putei; in the same study, the authors proposed the transfer of [Aquaspirillum] autotrophicum and [Pseudomonas] huttiensis to the genus Herbaspirillum as Herbaspirillum autotrophicum and Herbaspirillum huttiense. All three isolates were unable to fix nitrogen under laboratory conditions; only in H. putei was the nifH gene required for nitrogen fixation detected by PCR-based methods.
In this publication, we characterize the three isolates N3T, N5 and N9, which originated from surface-sterilized wheat roots (Triticum aestivum). On the basis of 16S and 23S rRNA gene sequence data, as well as DNA–DNA hybridization results and phenotypic features, these isolates belong to a novel species within the genus Herbaspirillum.
Isolates N3T, N5 and N9 were derived from 4- to 8-week-old wheat plants (T. aestivum var. Naxos) grown in agricultural soil from Neumarkt (Oberpfalz, Germany). Surface sterilization was carried out with chloramine T (1 % w/v) for 10 min. After surface sterilization, roots were washed three times with 1x PBS, crushed and then plated on NB agar [nutrient broth no. 4 (Fluka) solidified with 16 g agar l–1] at appropriate dilutions to obtain single colonies. Colonies were picked and singled out on new plates. With the exception of H. lusitanum P6-12T (a gift from J. Igual, Instituto de Recursos Naturales y Agrobiología, Salamanca, Spain), reference strains of Herbaspirillum were obtained from the Deutsche Sammlung von Mikroorganismen und Zellkulturen (DSMZ) (Braunschweig, Germany) or the IAM Culture Collection (Institute of Applied Microbiology, The University of Tokyo, Japan). All Herbaspirillum strains were grown at 30 °C on NB medium. For nitrogen-free microaerobic growth conditions, semi-solid JNFb medium was used as described by Döbereiner (1995).
Oxidase activity was determined from the oxidation of 1 % N,N-dimethyl-p-phenylenediamine hemioxalate. The presence of catalase activity was demonstrated by bubble formation in a 3 % (v/v) H2O2 solution. Metabolism of various substrates was tested in at least three replicates with Biolog GN2 Microplates according to the protocol provided by the manufacturer. Temperature and pH growth optima were determined in NB liquid medium (nutrient broth no. 4) after 16 h growth at 30 °C. Culture medium (5 ml) was inoculated with 10 µl preculture grown overnight. After incubation, the cell density was measured photometrically at 436 nm. The pH was adjusted by adding 0.05 M MES or MOPS buffer (Sigma-Aldrich).
Chromosomal DNA was isolated with a NucleoSpinTissue kit (Macherey Nagel). A PCR was performed in a Thermocycler Primus 25 (MWG-Biotech) in a total volume of 50 µl per reaction, according to standard protocols. The annealing temperature was 50 °C for all primers. The universal 16S primer 616-V (5'-AGAGTTTGATYMTGGCTCAG-3') and 630-R (5'-CAKAAAGGAGGTGATCC-3'), as well as the universal 23S primers 118-V (5'-CCGAATGGGGRAACCC-3') and 985-R (5'-CCGGTCCTCTCGTACT-3') were used for the PCR and sequencing. Additionally, the internal 23S primer 1019-V (5'-TAGCTGGTTCTYYCCGAA-3') was used for sequencing only. Amplified DNA fragments were cloned with the TOPO TA cloning kit (Invitrogen). Clones were sequenced using the Big Dye Terminator labelling kit (Applera) with an ABI PRISM 3730 DNA analyser (Applied Biosystems). For phylogenetic analyses, the 16S and 23S rRNA gene sequences obtained were aligned with the FastAligner version 1.03 tool implemented in the ARB software package (Strunk & Ludwig, 1997; Ludwig et al., 2004). Phylogenetic tree construction was performed by using the maximum-likelihood (Olsen et al., 1994), neighbour-joining (Saitou & Nei, 1987) and maximum-parsimony (Felsenstein, 1993) methods.
For fluorescence in-situ hybridization (FISH), 1 ml pure overnight culture was fixed with 4 % paraformaldehyde for at least 1 h at 4 °C (Amann et al., 1990). Hybridization with fluorochrome (fluorescein, Cy3)-labelled oligonucleotide probes was performed according to the protocols described by Manz et al. (1992) and Amann et al. (1992). New genus- and species-specific oligonucleotide probes based on 16S rRNA gene sequence data were created using the probe-design and probe-match tools of the ARB software package (Strunk & Ludwig, 1997; Ludwig et al., 2004). Hybridization conditions for the in-situ application of the newly developed probes had to be optimized by gradually increasing the formamide concentration in the hybridization buffer, as described by Manz et al. (1992). To determine the correct stringency conditions for hybridization, non-target bacterial species with only one or two mismatches in the target sequence were included in each hybridization.
Light-microscopic observation of cell morphology and size, as well as wavelength-specific detection of FISH-labelled cells, was performed with an Axioplan 2 epifluorescence microscope (Zeiss) equipped with filter sets 01, 09 and 15.
For scanning electron microscopy, cells were grown overnight in NB medium and then harvested from 1 ml cell suspension by centrifugation at 5000 g for 5 min. The cells were washed twice with 1x PBS and fixed with 2 ml 1 % glutaraldehyde in PBS (pH 7.4) overnight at 4 °C. The fixed cells were dehydrated through a series of ethanol solutions at increasing concentrations (50, 70, 80, 95 and 100 % ethanol). Ethanol was replaced with liquid CO2, and the samples were dried in a critical-point dryer. Cells were sputter-coated with platinum and examined in a scanning electron microscope (JSM 6300F; JEOL).
PCR amplification of the nifH and nifD genes was carried out by using bacterial genomic DNA as a template. Various primers and reaction conditions were used as described by Zehr & McReynolds (1989), Kloos et al. (1995), Stoltzfus et al. (1997) and Kirchhof et al. (2001).
For DNA–DNA hybridizations, about 3 g cell material was produced for each isolate by centrifuging a 2 l overnight culture for 25 min at 5000 g. After two washing steps with 1x PBS, cells were resuspended in 10 ml 50 % 2-propanol. All further steps were carried out by the DSMZ according to the protocols of Cashion et al. (1977) and De Ley et al. (1970), with the modifications described by Huß et al. (1983) and Escara & Hutton (1980). Renaturation rates were calculated with the program TRANSFER.BAS of Jahnke (1992).
The DNA G+C contents of the isolates were determined by the method described by Mesbah & Whitman (1989) with an HPLC system (LaChrom; Merck/Hitachi) using an L7400 UV detector.
The analysis of cellular fatty acids was performed according to a slightly modified procedure described by Gattinger et al. (2002). Briefly, cells grown overnight on NB medium were washed twice with 1x PBS. Lipids were extracted from these fresh cells, according to the Bligh–Dyer method, using phosphate buffer (0.05 M, pH 7.4), methanol and chloroform as extraction solvents. The resulting lipid material was subjected to mild alkaline methanolysis (Zelles & Bai, 1993) to yield fatty acid methyl esters (FAME). One aliquot of the FAME sample was derivatized with 1-trimethylsilyl imidazole (TMSI) and the other was derivatized with dimethyl disulfide (DMDS) (Zelles & Bai, 1993) to generate analytes suitable for GC and the localization of functional groups in the FAME molecule. GC/MS was performed with a Hewlett Packard 5971A mass selective detector combined with a 5890 series II GC system, equipped with an HP 5 capillary column (50 m length, 0.25 mm internal diameter; coated with a cross-linked 5 % phenylmethyl rubber phase with a film thickness of 0.3 µm), with helium as the carrier gas, at operating conditions as described elsewhere (Zelles & Bai, 1993). The identification and quantification of individual components were achieved using chromatography software (HP ChemStation; SOLVIT) and comparison with a FAME standard mixture. Under these GC conditions FAMEs containing 10 or more carbon atoms could be determined.
The three isolates N3T, N5 and N9 were Gram-negative, motile, slightly curved rods that formed circular, smooth, opaque and convex colonies after overnight growth on NB agar at 30 °C. They were positive for oxidase and catalase and showed optimal growth at temperatures between 26 and 34 °C and at pH values between 6 and 8. For examination of cell size and flagella, scanning electron micrographs were prepared (Fig. 1a, b). Up to three unipolar flagella could be detected, but most cells had two flagella. The size of single cells ranged between 1.6 and 2.0 µm in length and between 0.5 and 0.6 µm in diameter. The isolates were unable to grow on semi-solid, nitrogen-free JNFb medium (Kirchhof et al., 2001) and detection of nifD and nifH genes was not possible with PCR-based methods using various specific primers. According to these results, the isolates do not possess the ability to fix atmospheric nitrogen. As the isolates were derived from surface-sterilized wheat roots, they seem to be very closely associated with the plant host and perhaps even possess the potential for endophytic colonization.
|
. The major fatty acids detected were C16 : 17c, C16 : 0 and C18 : 17c; smaller proportions of C17 : 0 cyclo and C18 : 0 were also found. All of the other fatty acids detected were present at <1 mol% (Table 2). Similar results were obtained for other Herbaspirillum species, which showed the same dominant fatty acids with only slight differences in mole ratios (Ding & Yokota, 2004).
|
|
), these results provided evidence that the three isolates belong to the genus Herbaspirillum. The highest sequence similarity (99.9 %) was detected with H. lusitanum P6-12T, which was an indication that N3T, N5 and N9 were representatives of this species. However, all species of the genus Herbaspirillum exhibit a high degree of 16S rRNA gene sequence similarity, which ranges between 97.0 and 99.9 % (Supplementary Table S1). This is characteristic of species whose divergence has begun relatively recently and which are therefore hard to differentiate by 16S rRNA gene sequence-based methods (Fox et al., 1992). In this case, analysis of 16S rRNA gene sequences cannot provide sufficient evidence for species affiliation (Rossello-Mora & Amann, 2001).
In order to clarify the species affiliation of the novel strains, 2.5 kb fragments of the 23S rRNA genes of all the Herbaspirillum species were sequenced and phylogenetically analysed. The three isolates showed similarities of 99.8–99.9 % with respect to each other (see Supplementary Table S1). The sequence similarities between H. lusitanum P6-12T and the three isolates were 98.9–99.0 %, while the similarities within the genus ranged from 96.3 to 99.7 %. Since there are not as many 23S rRNA gene sequences available as there are for the 16S rRNA gene, the threshold values for species and genus level are not so clearly defined using the former (Rossello-Mora & Amann, 2001). However, as the type strains of H. huttiense and H. putei show 23S rRNA gene sequence similarity of 99.7 %, which is higher than the similarity between our isolates and H. lusitanum P6-12T, these results provide a first indication that N3T, N5 and N9 represent a distinct species within the genus Herbaspirillum. With the higher resolution of the 23S rRNA gene sequence data, the construction of a phylogenetic tree became possible (Fig. 2): it showed two distinct clusters of Herbaspirillum species. One cluster consisted of the type strains of H. seropedicae, H. rubrisubalbicans, H. huttiense, H. putei, H. frisingense and H. chlorophenolicum, while the other contained the type strains of H. autotrophicum and H. lusitanum and isolates N3T, N5 and N9. The closely related genus Collimonas is difficult to separate from the genus Herbaspirillum by phylogenetic analysis based on 16S rRNA or 23S rRNA gene sequence data, but it is clearly distinct when biochemical and physiological characteristics are considered (de Boer et al., 2004).
|
). Although this guideline value cannot be considered as absolute and is extended to 50 % in some exceptional cases (Stackebrandt & Goebel, 1994), the affiliation of isolate N3T to the species H. lusitanum could be ruled out according to these results. A DNA–DNA hybridization value as low as this is unusual in the context of a 16S rRNA gene sequence similarity of almost 100 %, but there are several comparable examples in the literature. Fox et al. (1992) detected similarly low DNA–DNA hybridization values when comparing several Bacillus globisporus and Bacillus psychrophilus strains, although a 99.5 % 16S rRNA gene sequence similarity, as well as various phenotypic traits, pointed towards an affiliation of the strains to the same species. Jaspers & Overmann (2004) isolated 11 strains from a bacterial freshwater community whose members had identical 16S rRNA gene sequences but which exhibited great genetic diversity and specific utilization of different carbon sources. Further examples exist in species of the genus Aeromonas (Martinez-Murcia et al., 1992) and Halobacillus (Amoozegar et al., 2003). There is no exact threshold for genus definition based on DNA–DNA hybridization values, but if the values between the individual Herbaspirillum species (10–45 %; Ding & Yokota, 2004) are considered, the data presented unambiguously show that the isolates should be allocated to the genus Herbaspirillum. The values for DNA–DNA hybridization between isolate N3T and the other Herbaspirillum species were as follows: H. autotrophicum DSM 732T, 32 %; H. rubrisubalbicans DSM 9440T, 30 %; H. huttiense DSM 10281T, 28 %; H. chlorophenolicum IAM 15024T, 25 %; H. seropedicae DSM 6445T, 20 %; H. frisingense GSF30T, 17 %; and H. putei IAM 15032T, 14 %. Results were obtained from three independent measurements, with a standard deviation of 5 %.
Kirchhof et al. (2001) designed specific 16S rRNA gene-targeted probes for the detection of all members of the genus Herbaspirillum that had validly published names at that time. The highly variable region at Escherichia coli positions 446–463 made differentiation possible down to the species level. Using the latest 16S rRNA gene sequence data available from the NCBI database (National Center for Biotechnology Information, US National Library of Medicine, Bethesda, USA) and sequences obtained within the scope of this work, the same binding position could be used for the development of oligonucleotide probes that were specific for the newly described Herbaspirillum species. At the appropriate formamide concentration (given in Table 3), all probes allowed specific detection of their respective target organisms. In the case of probe Hhilu446, differentiation of H. lusitanum and the three isolates (N3T, N5 and N9) was not possible, because of their high levels of 16S rRNA gene sequence similarity. The same is true for the pair H. huttiense and H. putei, which are both detected by Hhupu446. In addition, this probe has only one weakly destabilizing mismatch from the respective H. rubrisubalbicans target region. Therefore, in order to discriminate H. rubrisubalbicans, the application of an unlabelled oligonucleotide of Hrubri446 as a competitor is necessary when Hhupu446 is used.
|
Description of Herbaspirillum hiltneri sp. nov.
Herbaspirillum hiltneri (hilt'ne.ri. N.L. gen. masc. n. hiltneri of Hiltner, in honour of Professor Lorenz Hiltner, who coined the rhizosphere concept in 1904).
Cells are Gram-negative, slightly curved rods that are motile due to (generally) two unipolar flagella. Circular, smooth, opaque and convex colonies are formed on NB agar after overnight growth at 30 °C. They are oxidase- and catalase-positive and the optimal growth temperature is 26–34 °C. The pH optimum lies between 6 and 8. Single cells are 1.6–2.0 µm in length and 0.5–0.6 µm in diameter. A wide variety of sugars and alcohols, as well as fatty acids and some amino acids, are metabolized. In contrast to all other Herbaspirillum species, H. hiltneri sp. nov. is able to use L-phenylalanine as a carbon source. Disaccharides and trisaccharides are not metabolized. Shows 98.9 % 23S rRNA gene sequence similarity to its closest relative, H. lusitanum, but is not distinguishable from H. lusitanum on the basis of 16S rRNA gene sequence data. Values for DNA–DNA hybridization to the other Herbaspirillum species range between 32 and 14 %; values for hybridization to isolates N5 and N9 exceed 95 %. DNA G+C content is 60.9–61.5 mol%. Major fatty acids are C16 : 17c, C16 : 0 and C18 : 17c.
The type strain, N3T (=DSM 17495T=LMG 23131T), was isolated in close association with roots of Triticum aestivum. Its DNA G+C content is 60.9 mol%.
|
ACKNOWLEDGEMENTS |
|---|
|
REFERENCES |
|---|
|
TOPABSTRACTMAIN TEXTREFERENCES |
|---|
Amann, R. I., Zarda, B., Stahl, D. A. & Schleifer, K. H. (1992). Identification of individual prokaryotic cells by using enzyme-labeled, rRNA-targeted oligonucleotide probes. Appl Environ Microbiol 58, 3007–3011.
Amoozegar, M. A., Malekzadeh, F., Malik, K. A., Schumann, P. & Sproer, C. (2003). Halobacillus karajensis sp. nov., a novel moderate halophile. Int J Syst Evol Microbiol 53, 1059–1063.
Bae, H. S., Lee, J. M., Kim, Y. B. & Lee, S. T. (1996). Biodegradation of the mixtures of 4-chlorophenol and phenol by Comamonas testosteroni CPW301. Biodegradation 7, 463–469.[CrossRef][Medline]
Baldani, J. I., Baldani, V. L. D., Seldin, L. & Döbereiner, J. (1986). Characterization of Herbaspirillum seropedicae gen. nov., sp. nov., a root-associated nitrogen-fixing bacterium. Int J Syst Bacteriol 30, 86–93.
Baldani, J. I., Pot, B., Kirchhof, G. & 8 other authors (1996). Emended description of Herbaspirillum; inclusion of [Pseudomonas] rubrisubalbicans, a mild plant pathogen, as Herbaspirillum rubrisubalbicans comb. nov.; and classification of a group of clinical isolates (EF group 1) as Herbaspirillum species 3. Int J Syst Bacteriol 46, 802–810.
Brosius, J., Dull, T. L., Sleeter, D. D. & Noller, H. F. (1981). Gene organization of primary structure of a ribosomal operon from Escherichia coli. J Mol Biol 148, 107–127.[CrossRef][Medline]
Cashion, P., Holder-Franklin, M. A., McCully, J. & Franklin, M. (1977). A rapid method for the base ratio determination of bacterial DNA. Anal Biochem 81, 461–466.[CrossRef][Medline]
de Boer, W., Leveau, J. H., Kowalchuk, G. A., Klein Gunnewiek, P. J., Abeln, E. C., Figge, M. J., Sjollema, K., Janse, J. D. & van Veen, J. A. (2004). Collimonas fungivorans gen. nov., sp. nov., a chitinolytic soil bacterium with the ability to grow on living fungal hyphae. Int J Syst Evol Microbiol 54, 857–864.
De Ley, J., Cattoir, H. & Reynaerts, A. (1970). The quantitative measurement of DNA hybridization from renaturation rates. Eur J Biochem 12, 133–142.[Medline]
Ding, L. & Yokota, A. (2004). Proposals of Curvibacter gracilis gen. nov., sp. nov. and Herbaspirillum putei sp. nov. for bacterial strains isolated from well water and reclassification of [Pseudomonas] huttiensis, [Pseudomonas] lanceolata, [Aquaspirillum] delicatum and [Aquaspirillum] autotrophicum as Herbaspirillum huttiense comb. nov., Curvibacter lanceolatus comb. nov., Curvibacter delicatus comb. nov. and Herbaspirillum autotrophicum comb. nov. Int J Syst Evol Microbiol 54, 2223–2230.
Döbereiner, J. (1995). Isolation and identification of aerobic nitrogen-fixing bacteria from soil and plants. In Methods in Applied Soil Microbiology and Biochemistry, pp. 134–141. Edited by K. Alef & P. Nannipieri. London: Academic Press.
Döbereiner, J., Reis, V. M., Paula, M. A. & Olivares, F. L. (1993). Endophytic diazotrophs in sugar cane, tuber plants and cereals. In New Horizons in Nitrogen Fixation, pp. 671–676. Edited by R. Palacios, J. Mora & W. E. Newton. Dordrecht: Kluwer.
Elbeltagy, A., Nishioka, K., Sato, T., Suzuki, H., Ye, B., Hamada, T., Isawa, T., Mitsui, H. & Minamisawa, K. (2001). Endophytic colonization and in planta nitrogen fixation by a Herbaspirillum sp. isolated from wild rice species. Appl Environ Microbiol 67, 5285–5293.
Escara, J. F. & Hutton, J. R. (1980). Thermal stability and renaturation of DNA in dimethyl sulfoxide solutions: acceleration of the renaturation rate. Biopolymers 19, 1315–1327.[CrossRef][Medline]
Felsenstein, J. (1993). PHYLIP (phylogeny inference package), version 3.5c. Distributed by the author. Department of Genome Sciences, University of Washington, Seattle, USA.
Fox, G. E., Wisotzkey, J. D. & Jurtshuk, P., Jr (1992). How close is close: 16S rRNA sequence identity may not be sufficient to guarantee species identity. Int J Syst Bacteriol 42, 166–170.
Gattinger, A., Schloter, M. & Munch, J. C. (2002). Phospholipid etherlipid and phospholipid fatty acid fingerprints in selected euryarchaeotal monocultures for taxonomic profiling. FEMS Microbiol Lett 213, 133–139.[Medline]
Huß, V. A. R., Festl, H. & Schleifer, K.-H. (1983). Studies on the spectrophotometric determination of DNA hybridization from renaturation rates. Syst Appl Microbiol 4, 184–192.
Im, W. T., Bae, H. S., Yokota, A. & Lee, S. T. (2004). Herbaspirillum chlorophenolicum sp. nov., a 4-chlorophenol-degrading bacterium. Int J Syst Evol Microbiol 54, 851–855.
Jahnke, K. D. (1992). Basic computer program for evaluation of spectroscopic DNA renaturation data from GILFORD System 2600 spectrometer on a PC/XT/AT type personal computer. J Microbiol Methods 15, 61–73.
Jaspers, E. & Overmann, J. (2004). Ecological significance of microdiversity: identical 16S rRNA gene sequences can be found in bacteria with highly divergent genomes and ecophysiologies. Appl Environ Microbiol 70, 4831–4839.
Kirchhof, G., Eckert, B., Stoffels, M., Baldani, J. I., Reis, V. M. & Hartmann, A. (2001). Herbaspirillum frisingense sp. nov., a new nitrogen-fixing bacterial species that occurs in C4-fibre plants. Int J Syst Evol Microbiol 51, 157–168.[Abstract]
Kloos, K., Fesefeldt, A., Gliesche, C. G. & Bothe, H. (1995). DNA-probing indicates the occurrence of denitrification and nitrogen fixation genes in Hyphomicrobium. Distribution of denitrifying and nitrogen fixing isolates of Hyphomicrobium in a sewage treatment plant. FEMS Microbiol Ecol 18, 205–213.[CrossRef]
Ludwig, W., Strunk, O., Klugbauer, S., Klugbauer, N., Weizenegger, M., Neumaier, J., Bachleitner, M. & Schleifer, K. H. (1998). Bacterial phylogeny based on comparative sequence analysis. Electrophoresis 19, 554–568.[CrossRef][Medline]
Ludwig, W., Strunk, O., Westram, R. & 29 other authors (2004). ARB: a software environment for sequence data. Nucleic Acids Res 32, 1363–1371.
Manz, W., Amann, R., Ludwig, W., Wagner, M. & Schleifer, K.-H. (1992). Phylogenetic oligodeoxynucleotide probes for the major subclass of Proteobacteria: problems and solutions. Syst Appl Microbiol 15, 593–600.
Martinez-Murcia, A. J., Benlloch, S. & Collins, M. D. (1992). Phylogenetic interrelationships of members of the genera Aeromonas and Plesiomonas as determined by 16S ribosomal DNA sequencing: lack of congruence with results of DNA–DNA hybridizations. Int J Syst Bacteriol 42, 412–421.
Mesbah, M. & Whitman, W. B. (1989). Measurement of deoxyguanosine/thymidine ratios in complex mixtures by high-performance liquid chromatography for determination of the mole percentage guanine + cytosine of DNA. J Chromatogr 479, 297–306.[CrossRef][Medline]
Olivares, F. L., James, E. K., Baldani, J. I. & Döbereiner, J. (1997). Infection of mottled stripe disease-susceptible and resistant sugar cane varieties by the endophytic diazotroph Herbaspirillum. New Phytol 135, 723–737.[CrossRef]
Olsen, G. J., Matsuda, H., Hagström, R. & Overbeek, R. (1994). fastDNAmL: a tool for construction of phylogenetic trees of DNA sequences using maximum likelihood. Comput Appl Biosci 10, 41–48.
Rossello-Mora, R. & Amann, R. (2001). The species concept for prokaryotes. FEMS Microbiol Rev 25, 39–67.[Medline]
Saitou, N. & Nei, M. (1987). The neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol Biol Evol 4, 406–425.[Abstract]
Schmid, M., Baldani, J. I. & Hartmann, A. (2005). The Genus Herbaspirillum. In The Prokaryotes: an Evolving Electronic Resource for the Microbiological Community, 3rd edn, release 3.19. Edited by M. Dworkin. New York: Springer. http://link.springer-ny.com/link/service/books/10125/
Stackebrandt, E. & Goebel, B. M. (1994). Taxonomic note: a place for DNA-DNA reassociation and 16S rRNA sequence analysis in the present species definition in bacteriology. Int J Syst Bacteriol 44, 846–849.
Stoltzfus, J. R., So, R., Malarvithi, P. P., Ladha, J. K. & de Bruijn, F. J. (1997). Isolation of endophytic bacteria from rice and assessment of their potential for supplying rice with biologically fixed nitrogen. Plant Soil 194, 25–36.[CrossRef]
Strunk, O. & Ludwig, W. (1997). ARB: a software environment for sequence data. http://www.arb-home.de/
Valverde, A., Velázquez, E., Gutiérrez, C., Cervantes, E., Ventosa, A. & Igual, J.-M. (2003). Herbaspirillum lusitanum sp. nov., a novel nitrogen-fixing bacterium associated with root nodules of Phaseolus vulgaris. Int J Syst Evol Microbiol 53, 1979–1983.
Wayne, L. G., Brenner, D. J., Colwell, R. R. & 9 other authors (1987). International Committee on Systematic Bacteriology. Report of the ad hoc committee on reconciliation of approaches to bacterial systematics. Int J Syst Bacteriol 37, 463–464.
Zehr, J. P. & McReynolds, L. A. (1989). Use of degenerate oligonucleotides for amplification of the nifH gene from the marine cyanobacterium Trichodesmium thiebautii. Appl Environ Microbiol 55, 2522–2526.
Zelles, L. & Bai, Q. Y. (1993). Fractionation of fatty acids derived from soil lipids by soil phase extraction and their quantitative analysis by GC-MS. Soil Biol Biochem 25, 130–134.
This article has been cited by other articles:
|
|
 |
|
  S.-Y. Jung, M.-H. Lee, T.-K. Oh, and J.-H. Yoon Herbaspirillum rhizosphaerae sp. nov., isolated from rhizosphere soil of Allium victorialis var. platyphyllum Int J Syst Evol Microbiol, October 1, 2007; 57(10): 2284 - 2288. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| INT J SYST EVOL MICROBIOL | MICROBIOLOGY | J GEN VIROL |
| J MED MICROBIOL | ALL SGM JOURNALS | |
-hydroxybutyric acid, 
-ketobutyric acid, 