| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
Note |
Institute of Technical Microbiology, Technical University Hamburg-Harburg, Kasernenstr. 12, 21073 Hamburg, Germany
Correspondence
Garabed Antranikian
antranikian{at}tuhh.de
 |
ABSTRACT |
|---|
 TOP ABSTRACT MAIN TEXTREFERENCES |
|---|
-subclass of the Proteobacteria and belongs to the genus Psychromonas. On the basis of DNA–DNA hybridization experiments, chemotaxonomic studies and phenotypic characterization, strain Pull 5.3T (=CECT 5674T =DSM 14288T) clearly represents a novel species, for which the name Psychromonas arctica sp. nov. is proposed.
Published online ahead of print on 19 September 2002 as DOI 10.1099/ijs.0.02182-0.
The GenBank/EMBL/DDBJ accession number for the 16S rRNA gene sequence of P. arctica Pull 5.3T is AF374385.
|
MAIN TEXT |
|---|
|
TOPABSTRACTMAIN TEXTREFERENCES |
|---|
). In comparison, psychrotolerant micro-organisms, sometimes also referred to as ‘psychrotrophs’, generally have optimum and maximum growth temperatures of 20 °C or above (Ingraham & Stokes, 1959).
Cold-adapted micro-organisms are found in both permanently and temporarily cold habitats, which comprise more than 80 % of the Earth's biosphere. Oceans, covering three-quarters of the Earth, polar regions (14 % of the Earth's surface), high mountains and deep lakes provide various aquatic and terrestrial cold environments where the temperature seldom or never reaches 5 °C (Gounot, 1999). The ability of micro-organisms to grow at low temperatures is not restricted to prokaryotes. A wide variety of micro-organisms, including bacteria, archaea, yeast, fungi and algae, is found in cold environments. These micro-organisms are free-living in soil and fresh and saline waters or are associated with plants and cold-blooded animals such as fish or crustaceans. Among the bacteria, almost all types have been identified either after isolation (Ravenschlag et al., 1999; Bowman et al., 1997) or by detection in their natural habitats using a 16S rRNA approach (Fuhrman et al., 1993; DeLong et al., 1994; Vetriani et al., 1998). Unlike hyperthermophiles, they do not belong to new phyla. The majority of psychrophiles studied to date belong to the Gram-negative Proteobacteria. This is not surprising, since Gram-negatives are predominant in marine waters, where most investigations have been performed.
Psychrophiles and psychrotolerant micro-organisms have a wide range of adaptations, including alterations in the protein and lipids of their membrane, energy-generation systems, protein synthesis and hydrolytic enzymes (Russell, 1998). Higher specific activity at low temperatures and thermosensitivity of cold-active enzymes provide a valuable source for exploration of novel biotechnological processes (Feller & Gerday, 1997). Since it has become clear that psychrophiles also represent a naturally occurring model for investigating protein adaptation to cold (Aghajari et al., 1998), an increasing number of novel bacterial strains from different Arctic and Antarctic environments have been isolated.
In this study, employing a broad screening programme, a number of heterotrophic bacteria was isolated from sea-ice and sea-water samples obtained from the area of Spitzbergen, in the Arctic. Among the novel isolates, a group of strains was found to be similar to, but distinct from in a number of characteristics, a previously described psychrophilic species, Psychromonas antarctica, isolated from sediment of a high-salinity pond on the McMurdo Ice Shelf, Antarctica (Mountfort et al., 1998). Taxonomic and physiological analysis of the newly isolated strains demonstrated that they represent a novel species of the genus Psychromonas, and we propose the name Psychromonas arctica sp. nov. for the species represented by the type strain Pull 5.3T.
Arctic sea-ice and sea-water samples were collected in 1998 during an expedition to Svalbard, Spitzbergen. Samples were collected and transported to the laboratory at ambient temperature, 2–10 °C. An aliquot (0·5 ml) of liquid samples or a piece of membrane filter (cellulose acetate, pore size 0·2 µm) through which sea water was filtered were used for inoculation of 10 ml complex marine medium containing starch as a carbon source in 20 ml tubes. The complex medium contained (l-1): NaCl, 28·13 g; KCl, 0·77 g; CaCl2.2H2O, 0·02 g; MgSO4.7H2O, 0·5 g; NH4Cl, 1·0 g; iron ammonium citrate, 0·02 g; yeast extract, 0·5 g; 10-fold-concentrated trace element solution (DSM 141), 1 ml; 10-fold-concentrated vitamin solution (DSM 141), 1 ml; KH2PO4, 2·3 g; Na2HPO4.2H2O, 2·9 g; starch, 5 g. The pH was adjusted with NaOH to 7·2. In order to identify and isolate the producers of two main starch-degrading enzymes, 
-amylase (hydrolyses -1,4-glycosidic bonds) and pullulanase (specifically cleaves -1,6-glycosidic bonds), enrichment cultures were screened on agar plates containing complex medium and 0·5 % dyed amylopectin (Jørgensen et al., 1997) or dyed pullulan (Megazyme). The degradation of dyed amylopectin indicates the presence of -amylase activity, whereas halo formation on dyed pullulan is due to pullulanase (hydrolyses -1,6-linkage in pullulan) or pullulan hydrolase (hydrolyses -1,4-linkage in pullulan). Colonies that formed clearing zones on any of the dyed substrates were selected and transferred to liquid cultures. For the isolation of pure cultures, serial dilution and plating techniques were applied.
Enrichment cultures were obtained by inoculation of the complex medium containing starch as a carbon source with sea-ice and sea-water samples from Spitzbergen. After incubation for 6 weeks at 4 °C, stable enrichment cultures were obtained. Isolation was achieved by serial dilution and subsequent cultivation on agar plates containing dyed substrate (amylopectin or pullulan) at 4 °C. A total of 12 pure cultures were obtained, including seven strains isolated on dyed pullulan (Pull 1.5; Pull 5.3T; Pull 6.3; Pull 6.5; Pull 7.2; Pull 15.3; Pull 16.2) and five strains isolated on dyed amylopectin (Amyl 8.2; Amyl 18.6; Amyl 18.7; Amyl 20.2; Amyl 20.3). Interestingly, all strains isolated on dyed pullulan were also able to hydrolyse dyed amylopectin. The strains isolated on dyed amylopectin were not able to degrade dyed pullulan, however.
Gram staining and catalase tests were performed as described by Smibert & Krieg (1994). Cytochrome oxidase activity was determined with the Bacident Oxidase assay (Merck). Growth was measured by determining the optical density at 600 nm (1 cm path length) using a Shimadzu UV 1602 spectrophotometer.
To determine the salt requirement for growth, media were prepared with ten different NaCl concentrations between 0 and 6 % (w/v). The concentrations of other salts were kept constant. The vitamin requirement was tested after at least ten transfers on medium without vitamins. The pH optimum for growth was tested between pH 4 and 11. Growth with different electron donors (yeast extract, Casamino acids, starch, xylan, chitin, cellulose, gelatin, glycogen, acetate, ethylene glycol, betaine, mannitol, sorbitol, valine, isoleucine, L-histidine, L-arginine, L-serine, alanine, D-fructose, glucose, lactose, maltose, mannose, sucrose, xylose, glycerol, citrate, fumarate, propionate, lactate, pyruvate, malate and succinate) was tested using oxygen as electron acceptor on complex media lacking starch and yeast extract. Tubes without electron donors were inoculated and served as negative controls. Growth tests on different electron acceptors were made under anaerobic conditions in sulfate- and nitrate-free medium that was supplemented with glucose as the carbon source. The following electron acceptors were tested: thiosulfate (10 mM), elemental sulfur, nitrate (5 mM), nitrite (2 mM) or iron(III) citrate (30 mM). All test tubes were inoculated with sulfate- and nitrate-free pre-culture. The same pre-culture was used as the inoculum for growth experiments without electron acceptors using glucose, starch, lactate, pyruvate, fumarate, malate or propionate as a carbon source at a final concentration of 10 mM. Growth was measured by direct cell counting under the phase-contrast microscope. All tests were incubated at least in triplicate at the optimal temperature for growth of 20 °C.
Fatty acid methyl ester (FAME) analysis was performed according to the modified method of Lepage & Roy (1984). Total lipids were extracted according to Bligh & Dyer (1959). For derivatization, an aliquot of 3 mg total lipid extract was resolved in 2 ml methanol/hexane (4 : 1, v/v) plus pyrogallol and was methylated with 200 µl acetylchloride at 100 °C for 1 h; 5 ml 6 % K2CO3 was added and the mixture was centrifuged for 10 min at 2200 g. The upper, hexane phase containing the FAMEs was removed and dried with Na2SO4. FAMEs were analysed by capillary GC performed on an LS 32 GC (Chrompack; Kohn, 1996). For separation of fatty acid species, a fused silica capillary column (D23, 40 m; Fisons) was used. The chromatographic conditions were as follows: injector temperature (PTV), 65–270 °C; split ratio, 15 : 1; carrier gas, helium at a flow rate of 40 cm s-1. The column oven temperature profile was: initial temperature, 60 °C for 0·1 min; from 60 to 180 °C at 40 °C min-1; 180 °C for 2 min; from 180 to 210 °C at 2 °C min-1; 210 °C for 3 min; from 210 to 240 °C at 3 °C min-1; 240 °C for 10 min. Spectra were recorded by a flame-ionization detector at 280 °C.
Chromosomal DNA of the newly isolated strains was prepared according to the method described by Ausubel et al. (1992) and used as template DNA (5–100 ng) in a 100 µl PCR. Primers OPA-3, OPA-4 and OPA-13 and a combination of ERIC1R and ERIC2 were respectively used for RAPD- and ERIC-PCR (Versalovic et al., 1991; Rippere et al., 1998). PCRs were performed as described previously (Könönen et al., 1998; Versalovic et al., 1991) with a Perkin-Elmer Gene Amp PCR System 2400 thermal cycler. Negative controls with water instead of DNA showed no amplification. Following the reaction, 10 µl aliquots of PCR products were separated on a 1·5 % agarose gel stained with ethidium bromide.
The 12 starch-degrading isolates were arranged in similarity groups based upon the results of RAPD and ERIC fingerprinting (data not shown). Analysis of DNA fragments delineated five different patterns, independently of the method used. Whereas strains isolated on dyed pullulan showed only two distinct genotypes, strains isolated on dyed amylopectin were more diverse and showed three different genotypes. Based on preliminary physiological and phylogenetic characterization, the largest group (group B), containing six pullulan-degrading strains, was selected for further study. Strain Pull 5.3T was used as a representative of the group for detailed taxonomic and physiological characterization.
Plasmids were prepared by the alkaline lysis procedure with QIAprep Spin Miniprep kit (Qiagen) following the instructions of the manufacturer. From 50 µl isolated plasmid, 5 µl was used for 1 % agarose gel electrophoresis followed by staining with ethidium bromide.
All isolates of group B were found to contain small extrachromosomal DNA elements, as revealed by electrophoretic analysis of chromosomal DNA (Fig. 1). An identical plasmid profile was found in all strains within this group, confirming the results of fingerprinting. No plasmid bands were found in starch-degrading strains from other groups. Extrachromosomal DNA elements have been reported in a wide variety of bacteria, permitting their bacterial hosts to survive better in adverse environments or to compete better with other micro-organisms occupying the same ecological niche (Helinski et al., 1985). However, little is known about plasmids in psychrophiles. Since there is a considerable interest in the properties of cold adapted host-plasmid expression systems for the overproduction of heat-sensitive proteins (Tutino et al., 2000), the presence of plasmids in the novel isolates may be significant in recombinant DNA technology, particularly as a cloning vehicle for foreign genes.
|
). The G+C content of genomic DNA was determined by HPLC (Mesbah et al., 1989). The spectrophotometric renaturation rate procedure was performed as described previously (De Ley et al., 1970; Huß et al., 1983; Escara & Hutton, 1980) using a Gilford System 2600 spectrophotometer equipped with a Gilford 2527-R thermal programmer and plotter. DNA–DNA reassociation values were determined by using the program TRANSFER.BAS (Jahnke & Bahnweg, 1992).
Cells of strain Pull 5.3T were harvested from 2 ml culture samples by centrifugation and resuspended in 100 µl water. A subsample of 1 µl was used as a template for the amplification of 16S rDNA with primers 9–27f and 1492–1515r (Buchholz-Cleven et al., 1997). The PCR incorporated a hot start at 94 °C for 5 min and at 80 °C for 1 min before the addition of HiFi DNA polymerase mixture, followed by 30 cycles of 94 °C for 1·5 min, 46 °C for 1·5 min and 68 °C for 1·5 min. The amplification was performed with reagents from the Expand High Fidelity PCR System kit (Roche Diagnostics) following the recommendations of the manufacturer and using a Gene Amp PCR System 2400 thermal cycler. Negative controls with water instead of DNA showed no amplification. Amplicons were separated on a 1·0 % agarose gel stained with ethidium bromide and purified by the QIAquick PCR purification kit (Qiagen). The Taq DyeDeoxy Terminator cycle sequencing kit (Applied Biosystems) was used to sequence the purified PCR product directly. Sequencing reactions were analysed on an Applied Biosystems 373S DNA sequencer. Both strands of the amplification product were sequenced using primers 7F, 787F, 787R, 1175R, 1099F and 1492R (Buchholz-Cleven et al., 1997) (primer nomenclature refers to the 5' end of the respective target on the 16S rDNA according to Escherichia coli numbering).
Almost the complete sequence of the 16S rRNA gene of strain Pull 5.3T was determined. To determine the closest relatives of the novel isolate, preliminary searches in the EMBL database were performed with the program FASTA. Reference sequences utilized in phylogenetic analysis were retrieved from the EMBL database and aligned with the newly determined sequence of the novel isolate by using CLUSTAL W. Software from PHYLIP version 3.57c (Felsenstein, 1993) and MEGA version 2.0 (Kumar et al., 2001) was used for phylogenetic and molecular evolutionary analyses. DNADIST, with the maximum-likelihood option, was employed to analyse sequence similarities and NEIGHBOR (Kimura's two-parameter correction) was used to create a phylogenetic tree. The 16S rRNA sequence of Bacillus subtilis TB11 (AF058766) was used as the outgroup.
The G+C content of the DNA from strain Pull 5.3T was 40·1 mol%. The complete 16S rRNA gene sequence of strain Pull 5.3T (1528 nt, E. coli positions 8–1534) showed that the novel isolate belongs to the -subclass of the Proteobacteria (Fig. 2) and is related to species of the genera Colwellia, Moritella, Vibrio and Shewanella with sequence similarities of 89·0–89·8 %. The closest relative is P. antarctica, sharing 94·8 % 16S rDNA similarity, which, at the time of writing, is the only species described in the genus Psychromonas (Mountfort et al., 1998). The relative binding ratio for DNA–DNA hybridization of strain Pull 5.3T and P. antarctica is 25·0 %, well below the threshold value of 70 % accepted for the distinction of different species (Wayne et al., 1987).
|
a). In old cultures (2–3 days), the cells became more pleomorphic and non-motile coccoid cells, 1·3–1·7 µm in diameter, were observed. Electron microscopic observations revealed the presence of blebs on the cell surface (Fig. 3b) when the cells were grown at the optimal growth temperature of 20 °C. Similar membrane structures have been reported for other Gram-negative and Gram-positive bacteria (Antranikian et al., 1987). Although the function of the blebs of strain Pull 5.3T is still not clear, these structures could be connected with instability of membranes caused by the high incubation temperature. Cytoplasmic glycogen-like inclusions were found by microscopic investigation to be abundant in actively growing cells (Fig. 3a).
|
). Subsequent cultivation of the strain, independent of conditions used, resulted in complete loss of its ability to form biofilms within 2 months.
No spores were detected in either actively growing or old cultures, regardless of whether the newly isolated strain was cultivated on the regular complex medium or on the medium of Duncan & Strong (1968).
A culture grown at 4 °C was used for the determination of total cellular fatty acid composition. The novel isolate contained 16 : 1
7c as a major fatty acid (nearly 50 %). Other dominant fatty acids included 16 : 0, 16 : 17t and 18 : 17 as determined by GC (7·0–16·2 %). On the other hand, 12 : 0 and 14 : 15t represented relatively minor components (2·74–5·22 %). A fatty acid with an equivalent chain length of 14·33 could not be identified. The fatty acid profile of isolate Pull 5.3T resembles those of other marine bacteria characterized by the predominance of 16 : 1 and other monounsaturated species and short-chain fatty acids (Herbert, 1981). The conversion of cis- to trans-unsaturated fatty acids is believed to change the membrane fluidity in response to an environmental stimulus and was also reported for a heterogenic group of bacteria that included psychrophilic pollutant-degrading bacteria (Keweloh & Heipieper, 1996). The ability to synthesize trans-unsaturated fatty acid isomers could be an ecological advantage for Pull 5.3T, similar to a phenomenon described for the marine psychrophilic Vibrio strain ABE-1 (Okuyama et al., 1991). This allows the organism to react to changes in temperature and salinity much faster than by changing the degree of membrane saturation (Keweloh & Heipieper, 1996). Polyunsaturated fatty acids, which have been reported recently for a number of marine psychrophilic bacteria (Nichols 1999; Russell & Nichols et al., 1999), could not be detected.
Strain Pull 5.3T grew aerobically at temperatures between 0 and 25 °C (data not shown). Maximum growth rate as well as an increasing cell size, decrease in membrane stability and lower final cell yield were observed at 20 °C. No growth was observed at 27 °C or above. The organism grew well at salt concentrations of 1–7 % (w/v) with an optimum at 2 % (data not shown). The pH range for growth was 6·5–9·8, with an optimum at pH 8·8 (data not shown). Under optimal conditions, using starch or glucose as a carbon source, the growth rate was 0·68 h-1.
Of a number of polymeric substrates tested (starch, xylan, chitin, cellulose, gelatin, glycogen), only starch was found to support growth. However, the monosaccharides fructose, glucose, mannose and mannitol as well as the disaccharides lactose, maltose and sucrose were utilized. Acetate, pyruvate, succinate, fumarate, glycerol and alanine supported growth, but propionate, malate, citrate, lactate and xylose did not. Isolate Pull 5.3T fermented glucose and starch, resulting in the formation of acetate, ethanol, formate, lactate and CO2. No hydrogen, 1-propanol, 1-butanol, propionate or succinate could be detected as fermentation products. Nitrate, nitrite, sulfate, sulfite, iron(III) and elemental sulfur were tested as possible electron acceptors during growth of the strain on starch or glucose. Additional electron acceptors had little or no effect on growth or on the formation of fermentation products. At the end of growth, the concentration of electron acceptors had not decreased and no reduction products were found.
Although the fermentation pattern of Pull 5.3T and its ability to utilize carbohydrates are similar to those reported for P. antarctica, the novel isolate differs from its closest relative in a number of characteristics (Table 1). Differences in morphology, temperature and pH optima, the ability to form biofilms and the low DNA–DNA hybridization value indicate that the novel isolate Pull 5.3T represents a novel species of the genus Psychromonas, for which we propose the name Psychromonas arctica sp. nov.
|
Cells are 0·7–1·7 µm wide and 1·3–2·6 µm long. In old cultures, cells become more pleomorphic, with non-motile coccoid cells that are 1·3–1·7 µm in diameter. The pH optimum for growth is 8·5–8·8. Growth occurs in the presence of 1–7 % NaCl (w/v) with an optimum at 2 % NaCl. The temperature range for growth is 0–25 °C, with an optimum at 20 °C. Does not survive at temperatures above 27 °C for more than 4 h. The G+C content is 40·1 mol%. Sucrose, fructose, glucose, mannose, mannitol, lactose, maltose, acetate, pyruvate, succinate, fumarate, glycerol and alanine serve as carbon sources. Glucose and starch are fermented to acetate, ethanol, formate, lactate and CO2. Xylan, chitin, gelatin, propionate, malate, citrate, lactate and xylose do not support growth. Vitamins are not required for growth. The type strain, strain Pull 5.3T (=CECT 5674T =DSM 14288T), was isolated from a sea-water sample taken near Svalbard, Spitzbergen.
|
ACKNOWLEDGEMENTS |
|---|
|
REFERENCES |
|---|
|
TOPABSTRACTMAIN TEXTREFERENCES |
|---|
-amylase from Alteromonas haloplanctis in its native form and complexed with an inhibitor. Protein Sci 7, 564–572.[Medline]
Antranikian, G., Herzberg, C., Mayer, F. & Gottschalk, G. (1987). Changes in the cell envelope structure of Clostridium sp. strain EM1 during massive production of -amylase and pullulanase. FEMS Microbiol Lett 41, 193–197.
Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D. D., Seidman, J. G., Smith, J. A. & Struhl, K. (1992). Short Protocols in Molecular Biology, 2nd edn. New York: Greene–Wiley.
Bligh, E. G. & Dyer, W. J. (1959). A rapid method of total lipid extraction and purification. Can J Biochem Physiol 37, 911–917.
Bowman, J. P., McCammon, S. A., Brown, M. V., Nichols, D. S. & McMeekin, T. A. (1997). Diversity and association of psychrophilic bacteria in Antarctic sea ice. Appl Environ Microbiol 63, 3068–3078.[Abstract]
Buchholz-Cleven, B. E. E., Rattunde, B. & Straub, K. L. (1997). Screening for genetic diversity of isolates of anaerobic Fe(II)-oxidizing bacteria using DGGE and whole-cell hybridization. Syst Appl Microbiol 20, 301–309.
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]
Costerton, J. W., Lewandowski, Z., Caldwell, D. E., Korber, D. R. & Lappin-Scott, H. M. (1995). Microbial biofilms. Annu Rev Microbiol 49, 711–745.[CrossRef][Medline]
De Ley, J., Cattoir, H. & Reynaerts, A. (1970). The quantitative measurement of DNA hybridization from renaturation rates. Eur J Biochem 12, 133–142.[Medline]
DeLong, E. F., Wu, K. Y., Prezelin, B. B. & Jovine, R. V. (1994). High abundance of archaea in Antarctic marine picoplankton. Nature 371, 695–697.[CrossRef][Medline]
Duncan, C. L. & Strong, D. H. (1968). Improved medium for sporulation of Clostridium perfringens. Appl Microbiol 16, 82–89.[Medline]
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]
Feller, G. & Gerday, C. (1997). Psychrophilic enzymes: molecular basis of cold adaptation. Cell Mol Life Sci 53, 830–841.[CrossRef][Medline]
Felsenstein, J. (1993). PHYLIP (Phylogenetic inference package) version 3.57c. Distributed by the author. Department of Genetics, University of Washington, Seattle, USA.
Fuhrman, J. A., McCallum, K. & Davis, A. A. (1993). Phylogenetic diversity of subsurface marine microbial communities from the Atlantic and Pacific Oceans. Appl Environ Microbiol 59, 1294–1302.
Gounot, A. M. (1999). Microbial life in permanently cold soils. In Cold-adapted Organisms: Ecology, Physiology, Enzymology and Molecular Biology, pp. 3–17. Edited by R. Margesin & F. Schinner. Heidelberg: Springer-Verlag.
Helinski, D. R., Cohen, S. N., Clewell, D. B., Jackson, D. A. & Hollaender, A. (editors) (1985). Plasmids in Bacteria (Basic Life Sciences series, vol. 30). New York: Plenum.
Herbert, R. A. (1981). Low temperature adaptation in bacteria. In Effects of Low Temperature on Biological Membranes, pp. 41–54. Edited by G. J. Morris & A. Clarke. London: Academic Press.
Huß, V. A. R., Festl, H. & Schleifer, K. H. (1983). Studies on the spectrometric determination of DNA hybridization from renaturation rates. Syst Appl Microbiol 4, 184–192.
Ingraham, J. L. & Stokes, J. L. (1959). Psychrophilic bacteria. Bacteriol Rev 23, 97–108.
Jahnke, K.-D. & Bahnweg, G. (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.
Jørgensen, S., Vorgias, C. E. & Antranikian, G. (1997). Cloning, sequencing, characterization, and expression of an extracellular -amylase from the hyperthermophilic archaeon Pyrococcus furiosus in Escherichia coli and Bacillus subtilis. J Biol Chem 272, 16335–16342.
Keweloh, H. & Heipieper, H. J. (1996). Trans unsaturated fatty acids in bacteria. Lipids 31, 129–137.[Medline]
Kohn, G., van der Ploeg, P., Mobius, M. & Sawatzki, G. (1996). Influence of the derivatization procedure on the results of the gas-chromatographic fatty acid analysis of human milk and infant formulae. Z Ernährwiss 35, 226–234.[CrossRef][Medline]
Könönen, E., Mättö, J., Väisänen-Tunkelrott, M.-L., Frandsen, E. V. G., Helander, I., Asikainen, S., Finegold, S. M. & Jousimies-Somer, H. (1998). Biochemical and genetic characterization of a Prevotella intermedia/nigrescens-like organism. Int J Syst Bacteriol 48, 39–46.
Kumar, S., Tamura, K., Jakobsen, I. B. & Nei, M. (2001). MEGA2: Molecular Evolutionary Genetics Analysis software. Bioinformatics 17, 1244–1245.
Lepage, G. & Roy, C. C. (1984). Improved recovery of fatty acid through direct transesterification without prior extraction or purification. J Lipid Res 1, 1391–1396.
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.
Morita, R. J. (1975). Psychrophilic bacteria. Bacteriol Rev 39, 144–167.
Mountfort, D. O., Rainey, F. A., Burghardt, J., Kaspar, H. F. & Stackebrandt, E. (1998). Psychromonas antarcticus gen. nov., sp. nov., a new aerotolerant anaerobic, halophilic psychrophile isolated from pond sediment of the McMurdo Ice Shelf, Antarctica. Arch Microbiol 169, 231–238.[CrossRef][Medline]
Nichols, D., Bowman, J. P., Sanderson, K., Nichols, C. M., Lewis, T., McMeekin, T. & Nichols, P. D. (1999). Developments with Antarctic microorganisms: culture collections, bioactivity screening, taxonomy, PUFA production and cold-adapted enzymes. Curr Opin Biotechnol 10, 240–246.[CrossRef][Medline]
Okuyama, H., Okajima, N., Sasaki, S., Higashi, S. & Murata, N. (1991). The cis/trans isomerization of the double bound of a fatty acid as a strategy for adaptation to changes in ambient temperature in the psychrophilic bacterium Vibrio sp. strain ABE-1. Biochim Biophys Acta 1084, 13–20.[Medline]
Ravenschlag, K., Sahm, K., Pernthaler, J. & Amann, R. (1999). High bacterial diversity in permanently cold marine sediments. Appl Environ Microbiol 65, 3982–3989.
Rippere, K. E., Tran, M. T., Yousten, A. A., Hilu, K. H. & Klein, M. G. (1998). Bacillus popilliae and Bacillus lentimorbus, bacteria causing milky disease in Japanese beetles and related scarab larvae. Int J Syst Bacteriol 48, 395–402.
Russell, N. J. (1998). Molecular adaptations in psychrophilic bacteria: potential for biotechnological applications. Adv Biochem Eng Biotechnol 61, 1–21.[Medline]
Russell, N. J. & Nichols, D. S. (1999). Polyunsaturated fatty acids in marine bacteria – a dogma rewritten. Microbiology 145, 767–779.
Smibert, R. M & Krieg, N. R. (1994). Phenotypic characterization. In Methods for General and Molecular Microbiology, pp. 607–654. Edited by F. Gerhardt, R. G. E. Murray, W. A. Wood & N. R. Krieg. Washington, DC: American Society for Microbiology.
Tutino, M. L., Duilio, A., Moretti, M. A., Sannia, G. & Marino, G. (2000). A rolling-circle plasmid from Psychrobacter sp. TA144: evidence for a novel rep subfamily. Biochem Biophys Res Commun 274, 488–495.[CrossRef][Medline]
Versalovic, J., Koeuth, T. & Lupski, J. R. (1991). Distribution of repetitive DNA sequences in eubacteria and application to fingerprinting of bacterial genomes. Nucleic Acids Res 19, 6823–6831.
Vetriani, C., Reysenbach, A.-L. & Doré, J. (1998). Recovery and phylogenetic analysis of archaeal rRNA sequences from continental shelf sediments. FEMS Microbiol Lett 161, 83–88.[CrossRef][Medline]
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.
This article has been cited by other articles:
|
|
 |
|
  S. Hosoya, J.-H. Jang, M. Yasumoto-Hirose, S. Matsuda, and H. Kasai Psychromonas agarivorans sp. nov., a novel agarolytic bacterium Int J Syst Evol Microbiol, June 1, 2009; 59(6): 1262 - 1266. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
S. Hosoya, M. Yasumoto-Hirose, K. Adachi, A. Katsuta, and H. Kasai Psychromonas heitensis sp. nov., a psychrotolerant bacterium isolated from seawater in Japan Int J Syst Evol Microbiol, October 1, 2008; 58(10): 2253 - 2257. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
M. Miyazaki, Y. Nogi, Y. Fujiwara, and K. Horikoshi Psychromonas japonica sp. nov., Psychromonas aquimarina sp. nov., Psychromonas macrocephali sp. nov. and Psychromonas ossibalaenae sp. nov., psychrotrophic bacteria isolated from sediment adjacent to sperm whale carcasses off Kagoshima, Japan Int J Syst Evol Microbiol, July 1, 2008; 58(7): 1709 - 1714. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
Y. Nogi, S. Hosoya, C. Kato, and K. Horikoshi Psychromonas hadalis sp. nov., a novel piezophilic bacterium isolated from the bottom of the Japan Trench Int J Syst Evol Microbiol, June 1, 2007; 57(6): 1360 - 1364. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
A. J. Auman, J. L. Breezee, J. J. Gosink, P. Kampfer, and J. T. Staley Psychromonas ingrahamii sp. nov., a novel gas vacuolate, psychrophilic bacterium isolated from Arctic polar sea ice. Int J Syst Evol Microbiol, May 1, 2006; 56(Pt 5): 1001 - 1007. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
E. P. Ivanova, S. Flavier, and R. Christen Phylogenetic relationships among marine Alteromonas-like proteobacteria: emended description of the family Alteromonadaceae and proposal of Pseudoalteromonadaceae fam. nov., Colwelliaceae fam. nov., Shewanellaceae fam. nov., Moritellaceae fam. nov., Ferrimonadaceae fam. nov., Idiomarinaceae fam. nov. and Psychromonadaceae fam. nov. Int J Syst Evol Microbiol, September 1, 2004; 54(5): 1773 - 1788. [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 | |
