| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
1 DSMZ – German Collection of Microorganisms and Cell Cultures GmbH, Inhoffenstr. 7b, 38124 Braunschweig, Germany
2 Helmholtz Centre for Infection Research, Inhoffenstr. 7, 38124 Braunschweig, Germany
Correspondence
Erko Stackebrandt
erko{at}dsmz.de
 |
ABSTRACT |
|---|
 TOP ABSTRACT MAIN TEXTREFERENCES |
|---|
7c. Polyhydroxybutyrate and polyphosphate granules as well as unidentified enterosomes and a polar organelle are visible by electron microscopy. Comparative 16S rRNA gene sequence analysis indicated that the isolates were placed within the class Betaproteobacteria, remotely related to Chitinibacter tainanensis DSM 15459T, Silvimonas terrae KCTC 12358T, Formivibrio citricus DSM 6150T and Iodobacter fluviatilis DSM 3764T. On the basis of phylogenetic and phenotypic distinctness, we propose a novel genus, Deefgea gen. nov., with Deefgea rivuli sp. nov. as the type species. The type strain of Deefgea rivuli is strain WB 3.4-79T (=DSM 18356T=CIP 109326T).
The GenBank/EMBL/DDBJ accession number for the 16S rRNA gene sequence of strain WB 3.4-79T is AM397080.
Results of EELS and a 16S rRNA gene sequence-based maximum-likelihood tree are available as supplementary material in IJSEM Online.
|
MAIN TEXT |
|---|
|
TOPABSTRACTMAIN TEXTREFERENCES |
|---|
40 %) and many of the slime-producing bacteria were members of the genus Flavobacterium (Brambilla et al., 2007
; Cousin et al., 2007), while other strains with the slimy phenotype were affiliated to various classes of the Proteobacteria on the basis of partial 16S rRNA gene sequences (unpublished results). Phylogenetic analysis demonstrated the close affiliation of most water isolates to described species (>98 % 16S rRNA gene sequence similarity), and only a few were more distantly related. In this communication we present the characterization of two of these strains which are affiliated to the class Betaproteobacteria.
Strains WB 3.4-79T and WB 3.3-25 were isolated from sampling site 3 (Brambilla et al., 2007) of the Westerhöfer Bach on the western slopes of the Harz Mountains in Lower Saxony, Germany. Creek water was spread on R2A plates (Difco) which were incubated at 10 or 18 °C for 4 days. Single slimy colonies on the plates were purified by transferring them onto new plates and incubating again under the same conditions. The isolates was cultured routinely on R2A agar at 28 °C and maintained as a glycerol suspension (20 %, w/v) at –70 °C. Strains were preserved in the long term as freeze-dried cultures or in liquid nitrogen.
Ultrastructural analysis
Mid-exponential phase cells of strain WB 3.4-79T were prepared as ‘whole-mount’ samples either shadow-cast (Fig. 1a, b) or negatively stained (Fig. 1c), following a general protocol (Yakimov et al., 1998; Golyshina et al., 2000). Cells were also embedded in epoxy resin (Spurr, 1969) and ultrathin-sectioned. Energy-filtered transmission electron microscopy (EFTEM) was done with a CEM 902 instrument (Zeiss) at 80 kV in the elastic bright-field mode (Lünsdorf et al., 2001, 2006).
|
). Individual cells were surrounded by a slime layer, variable in thickness. In negatively stained cells, the slime layer can be visualized as an electron-dense halo (Fig. 1c) and in a platinum-shadowed preparation as a greyish amorphous rim at the cell periphery that occasionally showed radiant extrusions (Fig. 1a, open arrowheads).
Analysis of ultrathin sections showed the cell wall of strain WB 3.4-79T to be outlined by an outer and cytoplasmic membrane (Fig. 1e), characteristic of Gram-negative bacteria. In contact with the cytoplasmic membrane at the cytoplasmic face, a regularly structured layer was occasionally found, which is isomorphic to ‘polar organelles’. These have been described in Sphaerotilus natans (and other bacteria), and have been assumed to play a role in energy supply in this polytrichous monopolarly flagellated bacterium, based on their ATPase and cytochrome oxidase activities (Tauschel, 1985). Additionally, three morphotypes of intracellular inclusions, polyhydroxybutyrate (phb), polyphosphate (pp) and enterosomes (es) (Fig. 1d, e), are present. Electron-transparent polyhydroxybutyrate inclusions filled most of the cellular volume and were often torn out during the sectioning process. Polyphosphate granules, intensely stained and small in size, were interspersed as small clusters within the cytoplasm or were associated with the surface of large, electron-dense inclusions, 170–200 nm in size. Electron energy loss spectroscopy (EELS) revealed distinctly the presence of phosphorus in both small and large dark cellular inclusions (see Supplementary Fig. S1 in IJSEM Online). EELS also revealed the electron density to be based mainly on uranium, added as a stain during the dehydration process. A closer view of these large inclusions revealed the interior to be filled with a fine particulate matrix that, after low-frequency filtering or fast Fourier transformation, revealed a faint structural order, indicative of residual crystallinity (see Supplementary Fig. S1f–h). Residual crystallinity can also be deduced from the polyhedral contours of these inclusions. EELS and high-resolution elemental mapping showed the presence of phosphorus and nitrogen as the main constituents of these inclusions, supplemented by small amounts of calcium (Supplementary Fig. S1b–e). Nitrogen, as an indicator element of proteins, and the weak structural order of the particulate matrix indicate that these dark inclusions can be considered ‘enterosomes' in the general sense (Cannon et al., 2001). The presence of a distinct outer proteinaceous shell could not be observed and thus they differ at the structural level from typical ‘polyhedral bodies’. Further biochemical and physiological analysis has to be done to differentiate whether these inclusions represent ‘carboxysomes' in a specific sense (Cannon et al., 2001) or, because of the presence of calcium and phosphate, analogues to the ‘acidocalcisomes' of Agrobacterium tumefaciens (Seufferheld et al., 2003) or general ‘metabolosomes' (Brinsmade et al., 2005).
Genomic and phylogenetic characterization
Genomic DNA was extracted using the DNeasy Tissue kit (Qiagen) following the manufacturer's instructions. The 16S rRNA gene was amplified as described by Rainey et al. (1996) using the primers 10–30 forward and 1500 reverse. PCR products were purified with the QIAquick PCR purification kit (Qiagen) and sequenced directly by using the CEQ Dye Terminator cycle sequencing kit and on a CEQ 8000 Genetic Analysis system. 16S rRNA gene sequences were aligned with corresponding sequences from the DSMZ database using the ae2 editor (Maidak et al., 1997). Evolutionary distances were calculated by the method of Jukes & Cantor (1969). A distance analysis dendrogram was reconstructed by the neighbour-joining and maximum-likelihood algorithms (Felsenstein, 1993) and by the algorithm of De Soete (1983). Bootstrap analysis was used to evaluate the tree topology by performing 500 resamplings (Felsenstein, 1985).
Nearly complete 16S rRNA gene sequences (1508 nt) were obtained from strains WB 3.4-79T and WB 3.3-25. As the two sequences were identical, only that of strain WB 3.4-79T was analysed further. Preliminary comparison against the 16S rRNA gene sequences in GenBank by BLAST (Altschul et al., 1997) indicated that strain WB 3.4-79T belonged to the Chitinibacter–Iodobacter branch of the family Neisseriaceae, class Betaproteobacteria. The sequence of strain WB 3.4-79T was then included in the DSMZ database of 16S rRNA gene sequences and similarity values were determined. On the basis of these values, the closest described type strains were Chitinibacter tainanensis BCRC 17254T (93.8 %), Formivibrio citricus DSM 6150T (92.6 %), Silvimonas terrae KCTC 12358T (91.9 %) and Iodobacter fluviatilis ATCC 33051T (92.3 %). The 16S rRNA gene sequence similarity between strain WB 3.4-79T and other members of the order Neisseriales was less than 90 %. Strain WB 3.4-79T and the four type strains indicated above formed a monophyletic clade with a high bootstrap value (99 %), which was in accordance with the different treeing algorithms employed. While strain WB 3.4-79T branched adjacent to Chitinibacter tainanensis BCRC 17254T according to the neighbour-joining and De Soete analyses (Fig. 2), maximum-likelihood analysis showed strain WB 3.4-79T and Formivibrio citricus DSM 6150T to be phylogenetic neighbours (Supplementary Fig. S2).
|
) indicate that strains may belong to different genospecies even if they share 100 % 16S rRNA gene sequence similarity. Therefore, the genomic relatedness between the two novel strains was determined by the spectrophotometric DNA–DNA reassociation method. DNA was isolated as described by Marmur (1961) and was fragmented by ultrasonication (Bandelin Sonoplus) for 2 min at 170 µmss. DNA–DNA hybridization was carried out as described by De Ley et al. (1970), with the modifications of Huß et al. (1983) and Escara & Hutton (1980), using a model Cary 100 Bio UV/VIS spectrophotometer equipped with a Peltier-thermostatted 6x6 multicell changer and a temperature controller with in-situ temperature probe (Varian). The G+C content of the DNA of the two strains was determined according to Mesbah et al. (1989). The DNA relatedness of strains WB 3.4-79T and WB 3.3-25 was 100 %. The G+C content of strain WB 3.4-79T was 48.5 mol% (two determinations).
Analyses of chemotaxonomic properties
Except for fatty acid analysis, biomass for chemotaxonomic studies was prepared by growing the strain in shake flasks in marine broth 2216 (Difco) at 28 °C for 2 days. Cultures were checked for purity, harvested by centrifugation and freeze-dried. Polar lipids and isoprenoid quinones were extracted following the procedure of Minnikin et al. (1984). The polar lipids were separated by TLC using the standard method (Minnikin et al., 1984), whereas the quinone extract was filtered and reduced to dryness by a stream of dry nitrogen. The dried preparation was dissolved in 200 µl isopropanol and filtrated using a Dyna Gard 0.2 µl syringe filter. Aliquots (5 µl) were separated by HPLC following a described method (Kroppenstedt & Mannheim, 1989). Fatty acids of Chitinibacter tainanensis DSM 15459T, Formivibrio citricus DSM 6150T and Silvimonas terrae DSM 18233T were extracted and analysed (Miller, 1982; Sasser, 1990) according to the standard protocol of the Microbial Identification System (MIDI Inc., 1999). As Formivibrio citricus DSM 6150T does not grow anaerobically on TSBA medium, medium DSM 505 (DSMZ, 2001) was used to obtain biomass for fatty acid extraction. The same protocol was also reported for Silvimonas terrae KCTC 12358T (Yang et al., 2005). Extracts were analysed by using a Hewlett Packard model HP6890A gas chromatograph equipped with a flame-ionization detector as described by Kämpfer & Kroppenstedt (1996).
Ubiquinone 8 was the only isoprenoid quinone present. Phosphatidylethanolamine and phosphatidylglycerol were the major polar lipids. Diphosphatidylglycerol occurred in smaller amounts. The fatty acid composition is compiled in Table 1.
|
). The data obtained are indicated in the species description and in Table 2. Accumulation of poly-
-hydroxybutyrate was determined as described by Ostle & Holt (1982). Physiological and biochemical tests were performed on the two isolates and Chitinibacter tainanensis DSM 15459T at 28 °C using API 20NE, API 20E and API 50CH strips (bioMérieux) and Biolog GN microplates (Oxoid). All tests were inoculated with cells grown on R2A. API strips were used according to the manufacturer's instructions; utilization reactions were observed for 5 days. Acid production from carbohydrates was tested on API 50CH strips with bioMérieux medium E. Utilization of carbohydrates was determined on API 50CH strips with modified AUX medium in which growth factors and amino acids were replaced by a solution containing [(l distilled water)–1]: phosphate buffer, 40 mM; (NH4)2SO4, 2 g; MgSO4.7H2O, 6 g; CaCl2.2H2O, 0.066 g; agar, 1.5 g; yeast extract, 0.1 g; trace element solution, 1 ml; vitamin solution, 5 ml. Results of Biolog GN plates did not change over incubation times of 1–4 days. Wells showing a photometric value above 25 or 80 units were scored as weak and positive, respectively.
|
The pH range for growth was tested in buffered R2A medium at 28 °C between pH 5.1 and 9.5, with steps of about 0.3 pH. Growth of strain WB 3.4-79T occurred at pH 5.8–8.5, with an optimum around pH 7.3–7.6.
Salt tolerance of strain WB 3.4-79T was tested on R2A supplemented with 0.5, 1, 2, 4, 6, 8 and 10 % (w/v) NaCl at 28 °C. Good growth was observed after 3 days incubation at 0.5 %, while growth was weak on agar containing 1 % NaCl. No growth occurred at 2 % NaCl or above.
Tests for chemolithoautotrophic growth were done in mineral medium 81 (DSMZ, 2001) at 28 °C for 7 days under reduced (H2/N2/CO2/air; 70 : 10 : 10 : 10 by vol.) and aerobic conditions. Hydrogen and oxygen served as electron donor and acceptor, respectively. Growth was positive under both conditions and cells grew to similar densities.
Antibiotic-sensitivity tests were performed by using filter-paper discs containing 15 different antibiotics and concentrations. Discs were placed on R2A plates spread with WB 3.4-79T culture and were then incubated at 28 °C for 3 days. Susceptibility was scored as positive at zone diameters above 13 mm, intermediate susceptibility at 10–12 mm and resistance at less than 10 mm. The reactions are indicated in the species description.
Strains WB 3.4-79T and WB 3.3-25 are virtually identical in all taxonomic criteria used. They were isolated from the same site and may be considered clones, enriched on different growth media. The novel taxon can be distinguished from all neighbours by its significantly lower G+C content and in addition from Chitinibacter tainanensis (Chern et al., 2004) by the lack of chitin and gelatin hydrolysis, lack of growth at 40 °C, lack of C16 : 0 3-OH and C19 : 0 methyl fatty acids and by some phenotypic reactions. It differs from Silvimonas terrae (Yang et al., 2005) by the lack of hydrolysis of chitin, starch and cellulose, lack of growth at 40 °C, lack of C17 : 0 cyclo and C14 : 0 3-OH fatty acids and in the quantities of C16 : 0 fatty acid and those summed in features 4 and 7. While strains WB 3.4-79T and WB 3.3-25 are facultatively anaerobic, Formivibrio citricus (Tanaka et al., 1991) is strictly anaerobic. Due to this feature, only the API 20A (anaerobe) tests could be performed on the latter organism. Based on phylogenetic, cultural and physiological distinctness, we propose the genus Deefgea gen. nov. and Deefgea rivuli sp. nov. as new taxa, with strain WB 3.4-79T as the type strain of Deefgea rivuli.
Description of Deefgea gen. nov.
Deefgea [De.ef.ge'a. N.L. fem. n. Deefgea arbitrary name derived from the acronym DFG for Deutsche Forschungsgemeinschaft (German Science Foundation)].
Cells are Gram-negative, rod-shaped (0.7–0.8x3–4 µm) and facultatively anaerobic. They occur mostly singly (Fig. 1a) and are motile by means of a single flagellum (Fig. 1b) or rarely two polar flagella. Colonies on R2A are circular, convex and pearl white. Catalase and oxidase are positive. Contain polyhydroxybutyrate and polyphosphate granules. The predominant quinone is Q-8. Polar lipids are phosphatidylethanolamine and phosphatidylglycerol. Major cellular fatty acids (>10 %) are C16 : 0 and C16 : 17c. The G+C content of the DNA is 49 mol%. Based on 16S rRNA gene sequence analysis, Deefgea belongs to the class Betaproteobacteria, family Neisseriaceae, showing a distant relatedness to Chitinibacter tainanensis, Silvimonas terrae and Formivibrio citricus. The type species is Deefgea rivuli.
Description of Deefgea rivuli sp. nov.
Deefgea rivuli (ri'vu.li. L. gen. masc. n. rivuli of/from a rivulet, a small brook).
In addition to the characters that define the genus, it has the following characteristics. Cells have one round end and one pointed end; in older cultures, larger cells (4–7 µm) are occasionally present. Cells are highly motile at temperatures below 10 °C. Colonies on R2A agar are 1–2.5 mm in diameter, opaque with a smooth, shiny surface and butyrous with entire margins. Similar characteristics define colonies formed on Columbia agar (grey–white, umbonate, 3.2–4 mm in diameter), H3P (cream–white, 0.5–0.7 mm) and BTS (cream–white, 1.0–2.2 mm). Unspecified enterosomes and a polar organelle are present in the cytoplasm. No acid production in OF test under oxic or anoxic conditions. Grows at 4 °C but not above 32 °C. No growth above 2 % NaCl (w/v). pH optimum is between pH 7.3 and 7.6. Grows chemolithoautotrophically under aerobic and reduced conditions. Positive in API 20NE strips for reduction of nitrate to nitrite and utilization of glucose, N-acetylglucosamine and gluconate (all substrates utilized weakly). Tests positive in API 20E strips for acidification of glucose (only transiently after 24 h; realkalinization after 48 h) and acid production from sucrose. Substrates utilized in API 50CH with AUX medium are ribose, glucose, fructose, N-acetylglucosamine, sucrose and gluconate. Substrates positive in Biolog plates are dextrin, N-acetylglucosamine, D-fructose and -D-glucose; weakly positive for D-mannose, methylpyruvate and -hydroxybutyric acid. No degradation of chitin, cellulose, casein, starch or xylan. Resistant (per disc) to penicillin G (10 IU), oxacillin (5 µg), ampicillin (10 µg), cephalothin (30 µg), cefotaxime (30 µg), pipemidic acid (20 µg), erythromycin (15 µg), vancomycin (30 µg) and lincomycin (15 µg); intermediately susceptible to norfloxacin (10 µg), ofloxacin (5 µg) and colistin (10 µg); susceptible to tetracycline (30 µg), chloramphenicol (30 µg), imipenem (10 µg) and doxycycline (30 µg).
The type strain, WB 3.4-79T (=DSM 18356T=CIP 109326T), was isolated from water of the Westerhöfer Bach, near Westerhof, Lower Saxony, Germany (51° 45' 49'' N 10° 05' 31.7'' E).
|
ACKNOWLEDGEMENTS |
|---|
|
REFERENCES |
|---|
|
TOPABSTRACTMAIN TEXTREFERENCES |
|---|
Brambilla, E., Päuker, O., Cousin, S., Steiner, U., Reimer, A. & Stackebrandt, E. (2007). High phylogenetic diversity of Flavobacterium spp. in a hardwater creek, Harz Mountain, Germany. Org Divers Evol (in press).
Brinsmade, S. R., Paldon, T. & Escalante-Semerena, J. C. (2005). Minimal functions and physiological conditions required for growth of Salmonella enterica on ethanolamine in the absence of the metabolosome. J Bacteriol 187, 8039–8046.
Cannon, G. C., Bradburne, C. E., Aldrich, H. C., Baker, S. H., Heinhorst, S. & Shively, J. M. (2001). Microcompartments in prokaryotes: carboxysomes and related polyhedra. Appl Environ Microbiol 67, 5351–5361.
Chern, L.-L., Stackebrandt, E., Lee, S.-F., Lee, F.-L., Chen, J.-K. & Fu, H.-M. (2004). Chitinibacter tainanensis gen. nov., sp. nov., a chitin-degrading aerobe from soil in Taiwan. Int J Syst Evol Microbiol 54, 1387–1391.
Cousin, S., Päuker, O. & Stackebrandt, E. (2007). Flavobacterium aquidurense sp. nov. and Flavobacterium hercynium sp. nov., from a hard-water creek. Int J Syst Evol Microbiol 57, 243–249.
De Ley, J., Cattoir, H. & Reynaerts, A. (1970). The quantitative measurement of DNA hybridization from renaturation rates. Eur J Biochem 12, 133–142.[Medline]
De Soete, G. (1983). A least squares algorithm for fitting additive trees to proximity data. Psychometrika 48, 621–626.[CrossRef]
DSMZ (2001). Catalogue of strains, 7th edn. Braunschweig, Germany: Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH.
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. (1985). Confidence limits on phylogenies: an approach using the bootstrap. Evolution 39, 783–791.[CrossRef]
Felsenstein, J. (1993). PHYLIP – Phylogeny Inference Package, version 3.5c. Distributed by the author. Department of Genome Sciences, University of Washington, Seattle, USA.
Golyshina, O. V., Pivovarova, T. A., Karavaiko, G. I., Kondrat'eva, T. F., Moore, E. R. B., Abraham, W. R., Lünsdorf, H., Timmis, K. N., Yakimov, M. M. & Golyshin, P. N. (2000). Ferroplasma acidiphilum gen. nov., sp. nov., an acidophilic, autotrophic, ferrous-iron-oxidizing, cell-wall-lacking, mesophilic member of the Ferroplasmaceae fam. nov., comprising a distinct lineage of the Archaea. Int J Syst Evol Microbiol 50, 997–1006.[Abstract]
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.
Jukes, T. H. & Cantor, C. R. (1969). Evolution of protein molecules. In Mammalian Protein Metabolism, pp. 21–132. Edited by H. N. Munro. New York: Academic Press.
Kämpfer, P. & Kroppenstedt, R. M. (1996). Numerical analysis of fatty acid patterns of coryneform bacteria and related taxa. Can J Microbiol 42, 989–1005.
Kroppenstedt, R. M. & Mannheim, W. (1989). Lipoquinones in members of the family Pasteurellaceae. Int J Syst Bacteriol 39, 304–308.
Lünsdorf, H., Strömpl, C., Osborn, A. M., Bennasar, A., Moore, E. R. B., Abraham, W.-R. & Timmis, K. N. (2001). Approach to analyze interactions of microorganisms, hydrophobic substrates, and soil colloids leading to formation of composite biofilms, and to study initial events in microbiogeological processes. Methods Enzymol 336, 317–331.[Medline]
Lünsdorf, H., Kristen, I. & Barth, E. (2006). Cationic hydrous thorium dioxide colloids – a useful tool for staining negatively charged surface matrices of bacteria for use in energy-filtered transmission electron microscopy. BMC Microbiol 6, 59.[CrossRef][Medline]
Maidak, B. L., Larsen, N., McCaughey, M. J., Overbeek, R., Olsen, G. J. & Woese, C. R. (1997). The ribosomal database project. Nucleic Acids Res 25, 109–111.
Marmur, J. (1961). A procedure for the isolation of deoxyribonucleic acid from microorganisms. J Mol Biol 3, 208–218.
Mesbah, M., Premachandran, U. & Whitman, W. B. (1989). Precise measurement of the G+C content of deoxyribonucleic acid by high-performance liquid chromatography. Int J Syst Bacteriol 39, 159–167.
MIDI Inc. (1999). Sherlock Microbial Identification System, Operating Manual, version 3.0. Newark, DE: MIDI, Inc.
Miller, L. T. (1982). A single derivatization method for bacterial fatty acid methyl esters including hydroxy acids. J Clin Microbiol 16, 584–586.
Minnikin, D. E., O'Donnell, A. G., Goodfellow, M., Alderson, G., Athalye, M., Schaal, A. & Parlett, J. H. (1984). An integrated procedure for the extraction of isoprenoid quinines and polar lipids. J Microbiol Methods 2, 233–241.[CrossRef]
Ostle, A. G. & Holt, J. G. (1982). Nile blue A as a fluorescent strain for poly--hydroxybutyrate. Appl Environ Microbiol 44, 238–241.
Rainey, F. A., Ward-Rainey, N., Kroppenstedt, R. M. & Stackebrandt, E. (1996). The genus Nocardiopsis represents a phylogenetically coherent taxon and distinct actinomycete lineage; proposal for Nocardiopsaceae fam. nov. Int J Syst Bacteriol 46, 1088–1092.
Sasser, M. (1990). Identification of bacteria by gas chromatography of cellular fatty acids. USFCC Newsl 20, 16.
Seufferheld, M., Vieira, M. C. F., Ruiz, F. A., Rodrigues, C. O., Moreno, S. N. J. & Docampo, R. (2003). Identification of organelles in bacteria similar to acidocalcisomes of unicellular eukaryotes. J Biol Chem 278, 29971–29978.
Smibert, R. M. & Krieg, N. R. (1994). Phenotypic characterization. In Methods for General and Molecular Bacteriology, pp. 607–655. Edited by P. Gerhardt, R. G. E. Murray, W. A. Wood & N. R. Krieg. Washington, DC: American Society for Microbiology.
Spurr, A. R. (1969). A low-viscosity epoxy resin embedding medium for electron microscopy. J Ultrastruct Res 26, 31–43.[CrossRef][Medline]
Stackebrandt, E. & Ebers, J. (2006). Taxonomic parameters revisited: tarnished gold standards. Microbiol Today 33, 152–155.
Tanaka, K., Nakamura, K. & Mikami, E. (1991). Fermentation of S-citramalate, citrate, mesaconate, and pyruvate by a gram-negative strictly anaerobic non-spore-former, Formivibrio citricus gen. nov., sp. nov. Arch Microbiol 155, 491–495.[CrossRef]
Tauschel, H. D. (1985). ATPase and cytochrome oxidase activities at the polar organelle in swarm cells of Sphaerotilus natans: an ultrastructural study. Arch Microbiol 141, 303–308.[CrossRef][Medline]
Yakimov, M. M., Golyshin, P. N., Lang, S., Moore, E. R. B., Abraham, W. R., Lünsdorf, H. & Timmis, K. N. (1998). Alcanivorax borkumensis gen. nov., sp. nov., a new, hydrocarbon degrading and surfactant-producing marine bacterium. Int J Syst Bacteriol 48, 339–348.
Yang, H.-C., Im, W.-T., An, D.-S., Park, W.-S., Kim, I. S. & Lee, S.-T. (2005). Silvimonas terrae gen. nov., sp. nov., a novel chitin-degrading facultative anaerobe belonging to the ‘Betaproteobacteria’. Int J Syst Evol Microbiol 55, 2329–2332.
This article has been cited by other articles:
|
|
 |
|
  S.-C. Chang, W.-M. Chen, J.-T. Wang, and M.-C. Wu Chitinilyticum aquatile gen. nov., sp. nov., a chitinolytic bacterium isolated from a freshwater pond used for Pacific white shrimp culture Int J Syst Evol Microbiol, December 1, 2007; 57(12): 2854 - 2860. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
S. Muurholm, S. Cousin, O. Pauker, E. Brambilla, and E. Stackebrandt Pedobacter duraquae sp. nov., Pedobacter westerhofensis sp. nov., Pedobacter metabolipauper sp. nov., Pedobacter hartonius sp. nov. and Pedobacter steynii sp. nov., isolated from a hard-water rivulet Int J Syst Evol Microbiol, October 1, 2007; 57(10): 2221 - 2227. [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 | |
