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1 Department of Microbiology, University of Georgia, Athens, GA 30602-2605, USA
2 Department of Natural Sciences, Clayton College and State University, Morrow, GA 30260-1250, USA
3 Center for Biomarker Analysis, University of Tennessee, Knoxville, TN 37932-2575, USA
4 Institute of Ecology, University of Georgia, Athens, GA 30602-2360, USA
Correspondence
William B. Whitman
Whitman. whitman{at}arches.uga.edu
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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Published online ahead of print on 9 September 2002 as DOI 10.1099/ijs.0.02438-0.
The GenBank/EMBL/DDBJ accession number for the nearly complete 16S rRNA gene sequence of strain B33D1T is AY039806.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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) and Rubrobacter radiotolerans [originally Arthrobacter radiotolerans (Yoshinaka et al., 1973); later reclassified by Suzuki et al. (1988)]. Both of these organisms are notable for their radiation tolerance, rare pigmentation and thermophily. In contrast to the extreme environments that were the sources of the described Rubrobacter species, a large number of related 16S rRNA gene sequences have been recovered from moderate, terrestrial environments (Furlong et al., 2002; Holmes et al., 2000; McCaig et al., 1999; Rheims et al., 1996; Ueda et al., 1995), suggesting a wider range of habitat for the group. In this paper, we characterize an organism that is closely related to these common, uncultured soil bacteria within the Rubrobacteridae. |
METHODS |
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), on a plate composed of 50 % Difco nutrient broth medium (NB; pH 7·0, 
23 °C). Isolates were allowed to grow for 2 weeks before colonies were picked. A single pink colony was selected from a dilution series and maintained on the same medium. The organism was stored at -70 °C in medium containing 15 % glycerol.
Growth conditions.
Unless otherwise indicated, the optimization of growth conditions for B33D1T was carried out in undiluted Difco nutrient broth or NB medium. The temperature range was determined using a temperature gradient incubator (Scientific Industries Inc.). The pH range of B33D1T was determined by buffering NB with 25 mM MES (pH 5·5, 6·0 and 6·5), HEPES (pH 7·0, 7·5 and 8·0), EPPS (pH 8·5) or CHES (pH 9·0, 9·5, 10·0). The pH of the medium did not change during growth. Growth in various salt concentrations was tested by adding 0–10 % NaCl to NB. Growth under anaerobic conditions was tested by sparging NB with N2 gas for 30 min to remove O2 and incubating cultures in Balch tubes (Balch et al., 1979). To test growth under microaerophilic conditions, 1 % (v/v) air was added to N2-sparged tubes of NB. The effect of increased CO2 concentrations on B33D1T was tested by adding 1–5 % (v/v) CO2 to air in the headspace of NB medium in Balch tubes.
Biochemical properties.
The ability of various compounds to serve as sole carbon sources was tested in minimal medium with the carbon source added to a final concentration of 0·2 % (w/v). The minimal medium consisted of 1 mM K2HPO4, 2 mM NH4NO3, 1 mM MgSO4, 1 % (v/v) trace mineral solution (Whitman et al., 1986) and 1·0 % (v/v) iron solution (Whitman et al., 1986) adjusted to pH 7·0. In some tests (for example, the carbon utilization of lignin-associated compounds), the high concentration of iron produced a precipitate, and a 10-fold lower concentration was used. Tubes were incubated at 30 °C for 2 weeks before growth results were recorded.
The oxidase reaction of B33D1T was tested by applying a few drops of BBL Oxidase test reagent (Becton-Dickinson) to cells on a piece of filter paper. Other tests were performed as described by Furlong et al. (2002).
Desiccation resistance.
Strain B33D1T and Deinococcus radiodurans ATCC 35073 were grown in liquid media (NB for B33D1T, NB+10 % glucose for D. radiodurans) at 30 °C until growth was apparent. Wild-type Escherichia coli ATCC 9637 was grown in NB. Samples of the cultures (1–3 ml) were centrifuged for 3 min and the pellets were resuspended in 0·4 ml PBS (pH 7·2). Samples (0·1 ml) were then aliquotted into microfuge tubes, the tubes were centrifuged for 3 min and the buffer was discarded. The open microfuge tubes containing the cell pellets were placed upright in a sealed Mason jar containing Drierite absorbent for up to 25 days. Tubes were removed at various intervals and the cell pellets were resuspended and serially diluted in PBS. D. radiodurans was spread onto plates of NB+10 % glucose. E. coli was spread onto NB plates. For enumeration of B33D1T, 1 ml cell suspension was added to 9 ml molten agar medium (NB+1·5 % agar, 55 °C) and poured into a sterile Petri dish. Once the plates had solidified, they were sealed with Parafilm and incubated at 30 °C until colonies became visible.
Fatty acid analysis.
A sample of isolate B33D1T was analysed to determine the phospholipid fatty acid (PLFA) content of the organism. The cell paste was extracted with the single-phase chloroform/methanol buffer system of Bligh & Dyer (1954), as modified by White et al. (1979). The total lipid extract was fractionated into neutral lipids, glycolipids and polar lipids by silicic acid column chromatography (Guckert et al., 1985). The polar lipids were transesterified to the fatty acid methyl esters by a mild alkaline methanolysis (Guckert et al., 1985). The fatty acid methyl esters were then analysed by capillary GC with flame-ionization detection on a Hewlett Packard 5890 Series 2 chromatograph with a 50 m non-polar column (0·2 mm i.d., 0·11 µm film thickness). The injector and detector were respectively maintained at 270 and 290 °C. The column temperature was programmed at 60 °C for 2 min, ramped at 10 °C min-1 to 150 °C and finally ramped to 312 °C at 3 °C min-1. Preliminary peak identification was by comparison of retention times with known standards. Detailed identification of peaks was by GC/MS of selected samples using a Hewlett Packard 5890 series 2 GC interfaced to a Hewlett Packard 5971 mass-selective detector using the same column and temperature program as described previously. Mass spectra were determined by electron impact at 70 eV. Methyl nonadecanoate (C19 : 0) was used as the internal standard and amounts of PLFAs were expressed as equivalent peak responses to the internal standard.
Microscopy.
The cellular morphology of B33D1T was observed by phase-contrast microscopy, negative staining with negrosin dye and electron microscopy. A Nikon TE 300 inverted microscope was used to observe the cellular morphology of B33D1T. Digital photomicrographs were taken through the IP Lab Spectrum software package (version 3.4.5) using an attached Princeton Instruments MicroMax high-resolution, cooled CCD camera. Samples for scanning electron microscopy (SEM) were prepared according to the method of Hahn et al. (1998) except that 4 % glutaraldehyde was used to fix the cells. A Leo 982 field emission SEM was used to examine the cells. To test for the presence of a capsule, an India ink suspension, as well as a standard capsule stain, were used (Benson, 1990). The Gram reaction of isolate B33D1T was determined by a common staining technique and observed using light microscopy (Benson, 1990).
Phylogeny.
The nearly complete 16S rRNA sequence of B33D1T was obtained by amplification of the genomic DNA with primers 27f and 1392r (Lane et al., 1991; Furlong et al., 2002). Sequencing reactions were performed using a Big Dye sequencing kit (Perkin-Elmer) with primers 27f, 1392r, 533f, 519r and 907r (Lane, 1991). The sequencing reaction products were run on an ABI 377 Automated Sequencer (Perkin-Elmer). To construct the phylogenetic trees, the sequence was first aligned with reference organisms and environmental clone sequences using the PILEUP program included in the GCG software package. The evolutionary distances between aligned sequences were determined using a Jukes–Cantor algorithm in the DNADIST program of the PHYLIP software package (Felsenstein, 1989). The tree topology was determined using the FITCH program and the tree robustness was testing using 100 replicate trees as generated by SEQBOOT within the PHYLIP set of programs. The nearest phylogenetic neighbours of isolate B33D1T were determined by FASTA searches (Pearson & Lipman, 1988) of GenBank.
DNA G+C content.
The G+C content of isolate B33D1T was determined by the method of Mesbah et al. (1989). The value reported was the mean of five replicates.
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RESULTS AND DISCUSSION |
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). Older cultures generally had uniformly shorter rods. In liquid culture, cells were often observed with an indentation near the centre of the cell, presumably due to preparation for cellular division. Also in liquid culture, cells often grew in long chains that wrapped around each other, producing large aggregates. No capsule was observed. Cells of strain B33D1T stained Gram-positive. No motility was observed and no spores were apparent by phase-contrast microscopy.
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28 °C) often initially produced less pigment, although the deep-pink colour appeared over time. Strain B33D1T had a low growth efficiency upon streaking on agar plates, and only a few or no colonies resulted upon spread-plating on solid agar surfaces. However, B33D1T did grow in a liquid culture or embedded in soft, nutrient agar. The low viability of cells on the surface of an agar plate was at least partially explained by their desiccation sensitivity. Strain B33D1T cells rapidly lost viability upon desiccation when compared with E. coli (which has normal sensitivity to desiccation) and D. radiodurans (which is desiccation resistant; Fig. 2). Strain B33D1T was also sensitive to high temperature. Following incubation of a cell suspension containing 106 c.f.u. in PBS at 70 °C for 30 min, no c.f.u. were detected by pour plating. Although phylogenetically related Rubrobacter species are resistant to high levels of radiation (Ferreira et al., 1999), the apparent desiccation and heat sensitivity of B33D1T may indicate poor radiation resistance, as the phenotypes are often correlated (e.g. Billi & Potts, 2002; Mattimore & Battista, 1996; Sanders & Maxcy, 1979).
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).
Nutritional characteristics
A variety of compounds were tested as sole carbon and energy sources for strain B33D1T. Strain B33D1T generally grew to a low cellular density even in a rich medium, where the maximum OD600 was near 0·1. Growth was observed on a variety of sugars, including fructose, galactose, glucose, lactose, mannose, sorbitol, sucrose and xylose. Growth was not observed on cellobiose or mannitol. Strain B33D1T utilized pyruvate and appeared to grow weakly on acetate, but did not grow on citrate, malate or succinate. Casamino acids provided good growth. When tested individually, only the amino acids alanine, arginine and lysine supported growth. Glycerol was the only alcohol tested that could support the growth of B33D1T. Other alcohols that did not support growth were methanol, ethanol, 1-propanol, 2-propanol, butanol, isobutyl alcohol and isoamyl alcohol. Because strain B33D1T was isolated from soil, a variety of products of lignin degradation were tested as possible carbon sources. Of the compounds tested, only chlorogenic acid supported growth. Anthranilic acid, benzoic acid, catechol, protocatechuic acid, p-coumaric acid, gentisic acid, ferulic acid, p-hydroxybenzoic acid, syringic acid and vanillic acid did not produce observable growth after 2 weeks.
Strain B33D1T utilized ammonium as a sole nitrogen source and could grow on Casamino acids as a sole source of carbon and nitrogen. It did not reduce nitrate or produce urease.
Other growth conditions
A variety of pH and temperature conditions were tested. Strain B33D1T grew optimally at 28–30 °C, but could grow at between 19 and 38 °C. Strain B33D1T could grow at pH 6–7·5, with an optimum around pH 6–6·5. Strain B33D1T did not grow in the absence of oxygen but grew when 1 % (v/v) air was added to the headspace. Because the concentration of CO2 is often higher in soil than in the atmosphere, the growth of strain B33D1T under these conditions was tested. Increased CO2 levels (1–5 %, v/v) did not enhance the growth rate or yield significantly. While Rubrobacter strains could grow in media with increased salt concentrations (Ferreira et al., 1999; Carreto et al., 1996; Suzuki et al., 1988), strain B33D1T did not grow when as little as 1 % NaCl was added to the growth medium.
Biochemical and chemotaxonomic properties
Strain B33D1T was catalase-positive and oxidase-negative. Casein and Tween 80 were not hydrolysed. Strain B33D1T was sensitive to polymyxin, ampicillin, tetracycline and streptomycin. No haemolysis was observed on blood agar.
The fatty acid profile of B33D1T revealed no unusual compounds. The major PLFAs were i16 : 0 (54 %) and 18 : 1
9c (36 %). Trace amounts of 16 : 0 (4 %), 19 : 112c (2 %), 16 : 17c (1 %) and br17 : 1 (1 %) were also detected. No polyunsaturated PLFAs were found. Phylogenetically related Rubrobacter isolates contain only small amounts, if any, of the PLFAs that were abundant in strain B33D1T (Carreto et al., 1996).
The G+C content of the genomic DNA of strain B33D1T was 71·8±0·2 mol% (mean±SD; n=5), which is slightly higher than the value of 67·6 mol% reported for R. xylanophilus (Carreto et al., 1996) but not unexpected, given the high G+C content common for members of the phylum Actinobacteria.
Phylogeny
Based on 16S rRNA sequence analyses, strain B33D1T grouped within the subclass Rubrobacteridae of the phylum Actinobacteria, more specifically within subgroup 2 as defined by Holmes et al. (2000) (Fig. 3). No other isolates have been reported from this particular subgroup, although a number of environmental 16S rRNA genes have been described. The most closely related sequence in GenBank by FASTA analysis was an uncultivated clone (YNPFFP1; accession no. AF391984) from a thermal soil (93 % identity over 1364 bases), while the closest characterized organism was R. xylanophilus (86 % over 1372 bases). The other cloned sequences in subgroup 2 originated from terrestrial or sediment sources, as did strain B33D1T. Given the apparently ubiquitous distribution of this group in soil samples in various parts of the world, there is little reason to assume an association of this organism with earthworms, even though it was originally isolated from earthworm burrow soil.
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). The lower temperature optimum and sensitivity to NaCl appear to be the most important distinguishing features. Lastly, the poor plating efficiency and sensitivity to desiccation may explain why this taxon is so poorly represented in culture collections. Presumably, more representatives might be easily isolated from soil by pour plating.
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Cells are Gram-positive rods of medium length. Non-motile. Spores are not formed. Aerobic and mesophilic. Catalase-positive and oxidase-negative. Grow well on common sugars and a few amino acids as sole carbon sources. Sensitive to desiccation. Phylogenetically, placed within the subclass Rubrobacteridae of the Actinobacteria. The type species is Solirubrobacter pauli.
Description of Solirubrobacter pauli gen. nov., sp. nov.
Solirubrobacter pauli (pau'li. L. gen. n. pauli of Paulus, named for the prominent soil microbiologist Eldor A. Paul).
Cells are rods, 1·4x0·7 µm, and grow in long chains. Colonies are round, convex and pink in colour. The G+C content of the type strain is 71·8 mol%. The major fatty acids are i16 : 0 (54 %) and 18 : 19c (36 %). The type strain grows at pH 6·0–7·5 and 19–38 °C, with optima at pH 6·5 and 28–30 °C. Fructose, galactose, glucose, lactose, mannose, sorbitol, sucrose, xylose, pyruvate, acetate, Casamino acids, alanine, arginine, lysine, glycerol and chlorogenic acid support growth. Does not grow on cellobiose, mannitol, citrate, malate, succinate, methanol, ethanol, 1-propanol, 2-propanol, butanol, isobutyl alcohol, isoamyl alcohol, anthranilic acid, benzoic acid, catechol, protocatechuic acid, p-coumaric acid, gentisic acid, ferulic acid, p-hydroxybenzoic acid, syringic acid or vanillic acid. The type strain is strain B33D1T (=ATCC BAA-492T =DSM 14954T).
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ACKNOWLEDGEMENTS |
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REFERENCES |
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Benson, H. J. (1990). Microbiological Applications: a Laboratory Manual in General Microbiology. Dubuque, IA: Wm. C. Brown.
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Ferreira, A. C., Nobre, M. F., Moore, E., Rainey, F. A., Battista, J. R. & da Costa, M. S. (1999). Characterization and radiation resistance of new isolates of Rubrobacter radiotolerans and Rubrobacter xylanophilus. Extremophiles 3, 235–238.[CrossRef][Medline]
Furlong, M. A., Singleton, D. R., Coleman, D. C. & Whitman, W. B. (2002). Molecular and culture-based analyses of prokaryotic communities from an agricultural soil and the burrows and casts of the earthworm Lumbricus rubellus. Appl Environ Microbiol 68, 1265–1279.
Guckert, J. B., Antworth, C. P., Nichols, P. D. & White, D. C. (1985). Phospholipid ester-linked fatty acid profiles as reproducible assays for changes in prokaryotic community structure of estuarine sediments. FEMS Microbiol Ecol (FEMS Microbiol Lett Special Issue) 31, 147–158.[CrossRef]
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Holmes, A. J., Bowyer, J., Holley, M. P., O'Donoghue, M., Montgomery, M. & Gillings, M. R. (2000). Diverse, yet-to-be-cultured members of the Rubrobacter subdivision of the Actinobacteria are widespread in Australian arid soils. FEMS Microbiol Ecol 33, 111–120.[CrossRef][Medline]
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, 
), E. coli (
) and D. radiodurans (
). Multiple lines for strain B33D1T represent separate experiments.