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1 Institut für Mikrobiologie und Genetik, Universität Wien, A-1030 Vienna, Austria
2 Department of Biology and Geology, University of South Carolina Aiken, Aiken, SC, USA
3 Institut für Bakteriologie, Mykologie und Hygiene, Veterinärmedizinische Universität, A-1210 Vienna, Austria
4 DSMZ – Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH, Braunschweig, Germany
5 Institut für Zoologie, Abteilung Ultrastrukturforschung, Universität Wien, A-1090 Wien, Austria
6 Department of Microbiology and Molecular Medicine, Clemson University, Clemson, SC 29634, USA
7 Department of Biological Sciences, Florida International University, Miami, FL 33199, USA
Correspondence
Ewald B. M. Denner
ewald.denner{at}univie.ac.at
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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7c, along with C19 : 0 cyclo 8c and C16 : 0. Other fatty acids present in smaller amounts were C17 : 0, C18 : 0, C16 : 17c, C20 : 17c and C18 : 1 2-OH. The major polar lipids were phosphatidylethanolamine, phosphatidylglycerol and phosphatidylcholine. Minor amounts of diphosphatidylglycerol, phosphatidylmonomethylethanolamine and phosphatidyldimethylethanolamine were present. The G+C content of the genomic DNA was 66·3 mol%. Phylogenetic analysis of the 16S rRNA gene sequence showed that WP1T represents a separate subline of descent within the order ‘Rhizobiales’ of the ‘Alphaproteobacteria’. The new line of descent falls within the group of families that includes the Rhizobiaceae, Bartonellaceae, Brucellaceae and ‘Phyllobacteriaceae’, with no particular relative within this group. The 16S rRNA gene sequence similarity to all established taxa within this group was not higher than 92·0 % (to Mesorhizobium mediterraneum). To accommodate this emerging coral pathogen, the creation of a new genus and species is proposed, Aurantimonas coralicida gen. nov., sp. nov. (type strain WP1T=CIP 107386T =DSM 14790T).
Published online ahead of print on 13 December 2002 as DOI 10.1099/ijs.0.02359-0.
The GenBank/EMBL/DDBJ accession number for the 16S rRNA gene sequence of Aurantimonas coralicida strain WP1T is AJ065627.
Present address: Marine Biomedicine and Environmental Sciences, Medical University of South Carolina, Charleston, SC 29412, USA. 
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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). The disease was designated white plague type II because of its similarity to an earlier epizootic characterized as plague, or white plague, that occurred on the same reefs in the 1970s (Dustan, 1977). Both epizootics emerged as a sudden occurrence of diseased coral colonies on Florida's reefs. The diseased corals exhibited active coral tissue death, in which a sharp line was present between freshly exposed coral skeleton and apparently healthy coral tissue and which migrated across coral colonies, eventually resulting in colony death. No pathogen was isolated in the original plague (now designated type I) outbreak.
Microbiological studies conducted as part of the documentation of the 1995 white plague type II outbreak revealed that the disease was caused by a single Gram-negative bacterium, isolated as strain WP1T (Richardson et al., 1998a). At this time, it was demonstrated that pure cultures of WP1T readily initiated disease activity in healthy corals in the laboratory, thus satisfying the procedures of Koch's postulates (Richardson et al., 1998b). Based on a BLAST search comparing a 300 bp sequence of the 16S rRNA gene sequence (accession no. AF143861) and a limited number of phenotypic tests, the coral pathogen WP1T was identified as a possibly novel Sphingomonas species (Richardson et al., 1998a). Beyond it, a replication sequence comparison between the re-determined 16S rRNA gene sequence in the study presented here and the originally deposited sequence revealed that there was no significant degree of similarity (
82 %) between the two sequences.
In order to exclude any strain confusion, we have followed the history and distribution of strain WP1T among our different laboratories carefully, but we were not able to identify any problem. Also, pathogenic, physiological and biochemical traits of the early WP1T and the strain that we are now working with agree perfectly. The similarity between the 16S rRNA gene sequence of WP1T determined independently in Vienna and in South Carolina is 99·7 %, confirming that the two laboratories are working with the same strain. Therefore, we conclude that we are working with the same strain originally isolated and described by Richardson et al. (1998a) and that the problem with the low sequence similarity is related to the sequence originally deposited in GenBank. The polyphasic study presented here, which included molecular, chemosystematic and standard bacteriological analyses, reports on the actual taxonomic position of this emerging coral pathogen.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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Standard bacteriological characterization.
Strain WP1T was routinely cultivated aerobically on Bacto marine agar 2216 at 28 °C or otherwise as indicated in the text. To verify growth on different bacteriological media, WP1T was streaked onto Luria–Bertani agar (Atlas, 1993), Tryptone soy agar (TSA), MacConkey agar and R2A agar (Oxoid). The media were used in their original formulations and as marine versions containing 3·2 % (w/v) sea salts (Sigma). The ability to grow anaerobically was tested by means of a commercial atmosphere-generation system (AnaeroGen; Oxoid). Cell morphology was examined by phase-contrast microscopy (Leitz, Diaplan) from shake-flask cultures (150 r.p.m.) grown overnight in Bacto marine broth 2216 (Difco). Scanning electron microscopy (SEM) was performed on a Hitachi S4700 field emission scanning electron microscope at 5·0 kV. SEM samples were prepared by fixing intact colonies of the isolate in a 3·5 % glutaraldehyde solution (in 0·1 M sodium cacodylate buffer) for 18 h. The samples were dehydrated using a series of 30-min immersions in five different ethanol solutions (50, 70, 85, 95 and 100 %). Dehydrated samples were critical-point dried (CO2), mounted using carbon tape and sputter-coated with gold. For transmission electron microscopy (TEM), 2 µl bacterial suspension was placed onto a carbon-coated 400-mesh Ni grid (ATHENE SIRA, diameter 3·05 mm; Smethurst High-light). After 20 min absorption time, the grid was fixed in a 2·5 % (w/v) glutaraldehyde-cacodylate buffer (pH 7·4). Subsequently, the grid was rinsed three times in distilled water and stained with 1 % (w/v) uranyl acetate (pH 4·2). Excess stain was removed by touching the rim of the grid with a filter paper and the grid was then air-dried at room temperature. TEM samples were examined on a Philips EM 902 transmission electron microscope.
Growth at different temperatures was tested on Bacto marine agar 2216 plates incubated between 4 and 45 °C for as long as 2 weeks. Sensitivity against various antimicrobial agents (Table 1) was tested by the disc diffusion method using commercial antibiotic-impregnated discs (Oxoid). Briefly, 100 µl cell suspension (McFarland standard 0·5) in sterile 3·2 % sea water was plated onto Bacto marine agar 2216; after 48–72 h incubation at 28 °C, any sign of growth inhibition was scored as sensitivity. Resistance was indicated if no inhibition zone was observed. Assay for cytolytic properties was performed on sea-water-supplemented (3·2 %, w/v) TSA plates containing 5 % (v/v) defibrinated sheep blood. Biochemical characterization was carried out by following the standard methods of Smibert & Krieg (1994) supplemented by API 20E, API 20NE and API ZYM galleries (bioMérieux). API test systems were used according to the manufacturer's instructions except that (i) bacterial suspensions were prepared in autoclaved artificial sea water [40 g sea salts l-1 (Sigma) in demineralized water] and (ii) the reading was done after 5 h (API ZYM) and up to 7 days (API 20E, API 20NE). The presence of cytochrome c oxidase was tested with Bactident-oxidase test strips (Merck). Metabolic fingerprinting was carried out using the Biolog system. Briefly, five subcultures of WP1T were grown for 3 days on GASW agar (Smith & Hayasaka, 1982). Subsequently, cell suspensions were prepared in sterile (3·2 %, w/v) artificial sea water to an OD600 of 0·146. Microtitre plates (96-well; Biolog GN1 MicroPlate) containing 95 different carbon sources were inoculated with 150 µl of the adjusted cell suspension in each well and incubated at 30 °C for 3 days. A positive colorimetric reaction (the result of utilization of the sole carbon source and concomitant reduction of tetrazolium dye) was measured at 490 nm on an automated microplate reader (Molecular Devices; EMAX model). For this assay, any absorbance greater than 40 % of the control well (sterile sea water) was considered as positive.
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100 mg) with methanol/hexane (2 : 1, v/v) and were analysed by HPLC as described by Tindall (1990). Polyamines were extracted as described by Busse & Auling (1988) and were analysed according to Busse et al. (1997). Fatty acid methyl esters were extracted and prepared by the standard protocol of the MIDI/Hewlett Packard Microbial Identification System (Sasser, 1990). The fatty acid methyl ester profile was analysed by GLC using a GC-14A gas chromatograph (Shimadzu) as described by Groth et al. (1996). Polar lipids were extracted from 100 mg lyophilized cell material by the modified Folch procedure devised by Bligh & Dyer (1959) and resolved in 250 µl chloroform/methanol (2 : 1, v/v). Two-dimensional TLC was carried out as described by Denner et al. (2001). For pigment analysis, 100 mg cell material grown on Bacto marine agar 2216 was scraped from the agar surface and placed into a small (5 ml) Teflon-sealed glass vial. Subsequently, methanol (2 ml) was added to extract methanol-soluble pigments; after centrifugation (10 000 g, 4 °C, 5 min), the supernatant was scanned (300–800 nm) on a Hitachi S-2000 absorbance spectrophotometer. For DNA G+C content analysis, genomic DNA was isolated from lyophilized cell material and purified on hydroxyapatite according to the procedure of Cashion et al. (1977). The DNA was hydrolysed with P1 nuclease and the nucleotides were dephosphorylated with bovine alkaline phosphatase (Mesbah et al., 1989). The resultant deoxyribonucleosides were then analysed by HPLC (Tamaoka & Komagata, 1984). G+C content of DNA was calculated from the ratio of deoxyguanosine and thymidine according to Mesbah et al. (1989).
Determination and analysis of the 16S rRNA gene sequence.
Preparation of genomic DNA, enzymic amplification of 16S rDNA, PCR and sequencing were performed as described by Denner et al. (2001). Initial database searching was done by FASTA analysis (Pearson & Lipman, 1988; Pearson, 1990). Subsequently, relevant nucleotide sequences were retrieved from EMBL and GenBank databases, aligned manually using the program PILEUP (Devereux et al., 1984) and edited to remove nucleotide positions of ambiguous alignment and gaps. A continuous stretch of 1301 nucleotides in the alignment was used in the pairwise evolutionary distance estimation (Jukes & Cantor, 1969). Phylogenetic dendrograms were constructed using the neighbour-joining method (Saitou & Nei, 1987) and confidence in the tree topology was determined using 1000 bootstrapped trees. Phylogenetic analyses were performed using the programs included in the PHYLIP software package (Felsenstein, 1995). Taxonomic nomenclature was used according to Boone et al. (2001).
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RESULTS AND DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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). The highest 16S rRNA gene sequence similarity (92·0 %) was found to Mesorhizobium mediterraneum UPM-Ca36T. Dendrograms of phylogenetic relationships inferred from neighbour-joining (Fig. 1) and both maximum-likelihood and maximum-parsimony analysis (data not shown), including a subset of ‘Alphaproteobacteria’ species, showed that WP1T represents a separate subline of descent within the ‘Rhizobiales’. Bootstrap analysis gave 100 % confidence for this position. The new line of descent falls within the group of families that includes the Rhizobiaceae, Bartonellaceae, Brucellaceae and ‘Phyllobacteriaceae’, with no particular relative within this group (Fig. 1); strain WP1T may represent a novel family within the ‘Rhizobiales’.
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; Yokota et al., 1992; Busse et al., 1999). Fingerprinting of the cellular lipids by two-dimensional TLC revealed a complex composition consisting of diphosphatidylglycerol (DPG), phosphatidylethanolamine (PE), phosphatidylmonomethylethanolamine (PME), phosphatidyldimethylethanolamine (PDE), phosphatidylglycerol (PG), phosphatidylcholine (PC) and several unidentified lipids (Fig. 2). DPG, PE and PG are widely distributed amongst bacteria and are thus of little value for chemotaxonomic purposes (Wilkinson, 1988). The N-methylated PE derivatives PME, PDE and PC are of considerable taxonomic interest. Most of the bacteria described to contain N-methylated derivatives of PE are actinomycetes or are Gram-negatives, particularly bacteria belonging to the class ‘Alphaproteobacteria’ (Wilkinson, 1988; Busse et al., 1999). The polar lipid compositions of members of the ‘Rhizobiales’ lineage are not entirely clear. All species investigated so far contain PE and PC in comparable amounts; DPG is also consistently present (Kaneshiro & Marr, 1962; Thiele et al., 1968; Bunn & Elkan, 1970; Thiele & Schwinn, 1973; Thompson et al., 1983; Kämpfer et al., 1999; Choma & Komaniecka, 2002). Either or both of PME and PDE may also be formed: this applies to Agrobacterium tumefaciens (Goldfine & Ellis, 1964), Rhizobium leguminosarum (Faizova et al., 1971), Brucella spp. (Thiele et al., 1968; Thiele & Schwinn, 1973; Kulikov & Dranovskaia, 1988), Sinorhizobium meliloti (Thompson et al., 1983), Aminobacter aminovorans, Pseudoaminobacter spp. (Kämpfer et al., 1999) and Mesorhizobium spp. (Choma & Komaniecka, 2002).
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7c (76·9 %) along with C19 : 0 cyclo 8c (10·5 %) and C16 : 0 (6·7 %). Additional fatty acids detected in smaller amounts included C18 : 0 (1·5 %), C16 : 17c (1·3 %), C17 : 0 (0·6 %) and C20 : 17c (0·5 %). The sole hydroxylated fatty acid was C18 : 1 2-OH (2·0 %). The predominance of octadecenoic acids together with significant amounts of a cyclic C19 : 0 fatty acid is a typical feature of members of the ‘Rhizobiales’ lineage (Wilkinson, 1988; Moreno et al., 1990; Lechner et al., 1995; Jarvis et al., 1996; Dunfield et al., 1999; Kämpfer et al., 1999; Tighe et al., 2000). The closest match in the MIDI fatty acid database was Ochrobactrum anthropi; however, the similarity index was low (0·722). The hydroxylated fatty acid C18 : 1 2-OH detected in the fatty acid profile of WP1T is not common amongst bacteria, but, interestingly, this compound has been found in small quantities in Agrobacterium biovars 1 and 2, Mesorhizobium huakuii, Mesorhizobium loti, R. leguminosarum, Rhizobium hainanense, Rhizobium tropici (Jarvis et al., 1996; Dunfield et al., 1999; Tighe et al., 2000), O. anthropi (Lechner et al., 1995) and Phyllobacterium myrsinacearum (Mergaert et al., 2002).
The main component [15·3 µmol (g dry weight)-1] in the cellular polyamine pattern of WP1T was sym-homospermidine. Smaller amounts of spermidine [7·0 µmol (g dry weight)-1] and putrescine [4·8 µmol (g dry weight)-1] were also present. Generally, the polyamine patterns of all species of the ‘Rhizobiales’ that have been examined so far are dominated by sym-homospermidine (Busse & Auling, 1988; Auling et al., 1991; Hamana & Matsuzaki, 1992; Hamana & Takeuchi, 1998; Kämpfer et al., 1999). This polyamine pattern is not consistent, however. For example, both O. anthropi and Defluvibacter lusatiensis have either spermidine as the dominant compound or a combination of both putrescine and spermidine (Lechner et al., 1995; Fritsche et al., 1999).
Cultural, physiological and biochemical characteristics
Colonies of the coral pathogen WP1T on Bacto marine agar 2216 appeared opaque, circular, entire, convex, smooth and golden-orange in colour. Prior to pigment development (typically after 2 days of growth), colonies were translucent. Pigment extraction (methanol) yielded peaks at 
max 447 and 470–471 nm and showed a slight inflexion at 424–427 nm. This spectral characteristic is indicative of carotenoids (Schmidt et al., 1994). Cells were Gram-negative, rod-shaped (1x1·5–2·5 µm) with polar polytrichous flagella (Fig. 3). Interestingly, cells of WP1T exhibited a branching rod morphology (Fig. 4) that is usually found among non-spore-forming, high-G+C-content, Gram-positive bacteria such as Arthrobacter and Corynebacterium (Holt et al., 1994). This morphotype is relatively rare among Gram-negative bacteria, but has been found in some species of aerobic anoxygenic phototrophic bacteria (Yurkov & Beatty, 1998). Olson et al. (2002) recently isolated several Gram-negative bacteria that also displayed a branching rod morphology from marine sponges.
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. Strain WP1T exhibited a sharp clear zone of 
-haemolysis on blood agar. This is of potential importance in the aetiology of white plague type II, because this assay is indicative of cytolytic toxins (Rowe & Welch, 1994). Biochemical testing revealed further that WP1T is strongly ureolytic, since the urease test was positive after a few hours of incubation. The formation of ammonia by the coral pathogen may potentially contribute to pathogenesis (specifically by causing coral bleaching). Inhibition of photosynthesis by ammonia is well established (Warren, 1961; Cohen et al., 1975; Abeliovich & Azov, 1976). Ammonia acts as an uncoupler of photosynthesis by passing across membranes, thereby destroying the pH gradient across the thylakoid membrane (Smith & Raven, 1979; Velthuys, 1980). Recently, Banin et al. (2001) demonstrated that the toxin P from the coral pathogen Vibrio shilonii (Kushmaro et al., 2001) rapidly inhibits photosynthesis of zooxanthellae of Oculina patagonica in the presence of NH3.
Conclusions
Based upon the polyphasic taxonomic data obtained in this study, we conclude that the coral bacterial pathogen WP1T is a representative of a hitherto unknown marine taxon of the order ‘Rhizobiales’ of the class ‘Alphaproteobacteria’. We propose the name Aurantimonas coralicida gen. nov., sp. nov. for the strain.
The ‘Rhizobiales’ lineages include several bacteria such as Bradyrhizobium, Mesorhizobium, Rhizobium, Phyllobacterium, Brucella and Bartonella spp. that are known to form symbiotic or pathogenic associations with plants and animals (Holt et al., 1994). In this respect, the capacity of Aurantimonas coralicida to initiate disease in corals is a further indication that specific prokaryotic–eukaryotic association may be an important property shared by this group. To our knowledge, this is the first description of a species of the ‘Rhizobiales’ that is pathogenic for marine invertebrates. There is some evidence from our sequence database search by FASTA that at least three unidentified bacterial strains closely related to Aurantimonas coralicida have been isolated. These are strain SI85-l9A1, a marine manganese-oxidizing bacterium (Caspi et al., 1996), strain Eplume 4.J1, an isolate from a Pacific hydrothermal plume (Kaye & Baross, 2000), and strain R7951, isolated from the polar sea (Mergaert et al., 2001). The 16S rRNA gene sequence similarity of SI85-9A1 and R7951 to Aurantimonas coralicida WP1T was respectively 98·9 % (1293 ungapped positions) and 98·8 % (1292 ungapped positions). The phylogenetic relationship between strains SI85-9A1, R7951 and Aurantimonas coralicida WP1T is shown in Fig. 1. Strain Eplume 4.J1, of which only a partial (348 nt) 16S rRNA gene sequence is available (accession no. AF251774), exhibited 99·7 % sequence similarity to WP1T. These three isolates may represent another species of Aurantimonas or, at least, are novel strains of Aurantimonas coralicida.
Description of Aurantimonas gen. nov.
Aurantimonas (Au.ran.ti.mo'nas. M.L. adj. aurantiaca orange-coloured; Gr. fem. n. monas a unit; N.L. fem. n. Aurantimonas orange-coloured unicellular organism).
Gram-negative; endospores are not formed. Strictly aerobic. Catalase- and oxidase-positive. Intracellular pigments (carotenoids) are produced; the visible absorption spectrum of the pigment (methanol extract) shows peaks at max 447 and 470–471 nm and a slight inflexion at 424–427 nm. Sole respiratory lipoquinones present are ubiquinones, with Q-10 predominating; Q-9 may account for about 1 % of the total. The main cellular polyamine is sym-homospermidine; minor amounts of putrescine and spermidine are present. Major polar lipids are PE, PG and PC. DPG, PME and PDE are present as secondary components. The predominant fatty acid is C18 : 17c, along with C19 : 0 cyclo 8c and C16 : 0. Other fatty acids are C17 : 0, C18 : 0, C16 : 17c and C20 : 17c. The sole hydroxylated fatty acid is C18 : 1 2-OH. The G+C content of the DNA of the type species is 66·3 mol% (by HPLC). The type species is Aurantimonas coralicida.
Description of Aurantimonas coralicida sp. nov.
Aurantimonas coralicida [co.ra.li'ci.da. L. n. coralium (red) coral; L. masc./fem. suffix -cida murderer, killer; N.L. masc./fem. n. coralicida coral-killer].
Cells are rods (average 1·5–2·5x1 µm) with a bulbous branching rod morphology. Motile by means of polar polytrichous flagella. Colonies on marine agar 2216 are opaque, golden-orange-coloured, circular, entire, convex and smooth. Prior to pigment development, colonies are translucent. Urease is present. Details of nutritional, physiological and biochemical features are specified in Table 1. Chemotaxonomic characteristics are the same as those given in the genus description. The type strain, strain WP1T (=CIP 107386T =DSM 14790T), was isolated from a diseased colony of the scleractinian coral Dichocoenia stokesi (elliptical star coral) in the Florida Keys.
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ACKNOWLEDGEMENTS |
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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