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1 Division of Entomology, Commonwealth Scientific and Industrial Research Organization, Canberra ACT 2601, Australia
2 Ecologie microbienne des Insectes et Interaction Hôte–pathogène Unité EMIP INRA 1133, Université Montpellier II, 34095 Montpellier CEDEX 5, France
3 Department of Microbiology and Immunology, University of Melbourne, Vic. 3010, Australia
4 Gold Coast Hospital, Southport Qld 4215, Australia
Correspondence
R. J. Akhurst
Ray.Akhurst{at}csiro.au
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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; Boemare et al., 1993). The bacteria are transported into the haemocoel of a host insect within the intestinal tract of infective-stage nematodes, where they multiply, killing the host and producing conditions that are favourable for the nematodes' maturation and reproduction. Two phases are recognized in Photorhabdus, with phase I being isolated from wild-type Heterorhabditis infective juveniles and phase II appearing in laboratory cultures (Akhurst, 1980; Boemare & Akhurst, 1988). Photorhabdus–Heterorhabditis associations have become important biopesticides, particularly as the nematodes are mobile and can locate insect pests in cryptic habitats (Bedding & Miller, 1981).
Farmer et al. (1989) reported the isolation of Photorhabdus from human clinical specimens in the USA. Six isolates were recovered from several states over the period 1977–1989. The route of infection could not be ascertained and no further isolations have been reported from the USA. These isolates were very homogeneous and were shown to constitute a fifth DNA hybridization group, distinct from nematode-symbiotic Photorhabdus (Farmer et al., 1989). Fischer-Le Saux et al. (1999) included these isolates in their polyphasic examination of the taxonomy of the genus Photorhabdus and concluded that they constituted a separate species from any of the nematode-symbiotic Photorhabdus spp. They designated the novel species Photorhabdus asymbiotica. No phase I isolates of this species have been reported.
Peel et al. (1999) subsequently reported the isolation of Photorhabdus sp. from four human infections in two Australian states between 1994 and 1998. PCR-RFLP analysis of the 16S rRNA gene showed that these Australian isolates were also highly homogeneous, but distinct from the American isolates. It was not clear from the analysis whether the American isolates were more similar to the Australian isolates or to some symbiotic Photorhabdus isolates (Peel et al., 1999). Two more isolations were performed subsequently (Gerrard et al., 2003).
A polyphasic approach to the taxonomic study of the Australian clinical isolates was conducted, in order to determine their relationship to clinical and symbiotic bacteria of the genus Photorhabdus. The isolates were characterized phenotypically by biochemical and morphological tests. DNA–DNA relatedness was assessed by measurement of reassociation and 
Tm of heterologous DNA. In addition to sequencing the 16S rRNA gene for comparison with sequences in various databases, the gyrB gene of the Australian clinical isolates and Photorhabdus isolates that are representative of the taxonomic groups described by Fischer-Le Saux et al. (1999) was sequenced for phylogenetic analysis. The sequence of the gyrB gene has proven to be useful for investigating the intrageneric relationships of disparate groups of bacteria (Venkateswaran et al., 1999; Kasai et al., 2000; Yamamoto et al., 2000). The polyphasic taxonomic study indicated that human clinical isolates of Photorhabdus fall into two distinct subspecies of P. asymbiotica.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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) were provided by the Microbiological Diagnostic Unit of the University of Melbourne, the Gold Coast Hospital and the Centers for Disease Control and Prevention (USA). Symbiotic isolates were obtained from the CSIRO Entomology collection.
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. Acid production from and assimilation of carbon sources was tested with API kits (bioMérieux). Luminescence was assessed visually by observation after 10 min dark-adaptation and by analysis of a just visibly turbid aqueous suspension of cells from a 24 h nutrient broth (Oxoid) culture in a Beckman LS2800 scintillation counter, on the tritium setting. Insect pathogenicity was assessed by the insect pin test of Peel et al. (1999). Maximum temperature for growth was determined in nutrient broth that was inoculated from a 24 h nutrient broth culture that had been incubated in a water bath with accuracy of ±0·2 °C for 48 h; the five P. asymbiotica isolates from the USA (Table 1
) were tested simultaneously with the seven Australian clinical isolates.
DNA isolation.
DNA was isolated from cells harvested from 24 h nutrient broth (Oxoid) cultures by using a Qiagen DNeasy kit.
PCR amplification and sequencing of the 16S rRNA gene.
The prokaryote-specific primers of Fischer-Le Saux et al. (1999) were used in PCR amplification of the 16S rRNA gene. PCR was performed on 0·3 µg target DNA in 100 µl Gibco PCR mix that contained 4 mM MgCl2, 1 mM each dNTP, 25 pM each primer, 100 µg cresol red, 13 % (w/v) sucrose and 2·5 U Taq DNA polymerase (Gibco). Reactions were run on a Corbett FTS 320 thermocycler, with 35 cycles of denaturation at 94 °C for 1 min, annealing at 55 °C for 1 min and extension at 72 °C for 2·5 min, followed by a final extension at 72 °C for 3 min and a renaturation step at 25 °C for 10 min. The completed reactions were electrophoresed on 1·5 % (w/v) agarose gel at 5 V cm–1. The band was excised and the amplicon was purified with a NucleoTrap Gel Extraction kit (Clontech), cloned into a pGEM-T Easy vector (Promega) and transformed into Escherichia coli strain DH10
by electroporation. In order to identify and eliminate nucleotide changes that were introduced during PCR, four individual clones recovered from each ligation were sequenced with the primer set described by Fischer-Le Saux et al. (1999). Sequencing reactions were performed with an ABI Prism Dye Terminator cycle sequencing ready reaction kit and analysed with an ABI 373A automated sequencer, under the conditions recommended by the manufacturer (Perkin Elmer). The consensus sequence of the four clones was used in subsequent phylogenetic analysis.
PCR amplification and sequencing of the gyrB gene.
PCR amplification of gyrB genes was performed with the universal primers of Yamamoto & Harayama (1995). PCR was carried out on 50 ng target DNA in 20 µl Dynazyme buffer that contained 2·5 mM each dNTP, 10 µM each primer and 1 U Dynazyme EXT DNA polymerase. Reactions were run on a Corbett FTS 320 thermocycler, with a 2 min denaturation step at 94 °C, followed by 34 cycles of denaturation at 94 °C for 30 s, annealing at 58 °C for 30 s and extension at 72 °C for 75 s. A final extension was conducted at 72 °C for 2 min and was followed by a renaturation step at 25 °C for 5 min.
gyrB amplicons were sequenced with primers UP-1S and UP-2Sr (Yamamoto & Harayama, 1995) and UP-3Sr (5'-AGAAGAGAATTTTGGATCAG-3'). Sequencing reactions were performed with an ABI Prism Dye Terminator cycle sequencing ready reaction kit and analysed with an ABI 373A automated sequencer, under the conditions recommended by the manufacturer (Perkin Elmer).
Phylogenetic analyses.
16S rRNA and gyrB gene sequences were aligned against homologous sequences (see Figs 1 and 2) by using the program CLUSTAL X version 1.81 (Thompson et al., 1997). This alignment was then checked manually and corrected, and regions of uncertain alignment were eliminated, with the software SEAL version 1.d1 (A. Rambaut, Department of Zoology, University of Oxford, UK). Further analyses were restricted to the unambiguously aligned regions, totalling 1400 positions of the 16S rRNA gene and 1129 positions of gyrB.
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). Evolutionary distances between pairs of micro-organisms were determined with the correction of Jukes & Cantor (1969), implemented in the DNADIST program, and phylograms were derived with the FITCH and NEIGHBOR programs, employing least-squares (Fitch & Margoliash, 1967) and neighbour-joining (Saitou & Nei, 1987) algorithms, respectively. Significance of nodes was tested by bootstrap analysis that generated evolutionary distances and used neighbour-joining to produce 1000 trees, and then by compiling a consensus tree, using the programs SEQBOOT, DNADIST, NEIGHBOR or FITCH and CONSENSE. Maximum-likelihood and maximum-parsimony analyses of the sequence data employed the programs DNAML and DNAPARS, respectively. Sequences used as outgroups were the 16S rRNA gene sequences of E. coli (GenBank accession no. J01695) and Proteus vulgaris (X07652) and the gyrB sequences of E. coli (X04341) and Xenorhabdus nematophila isolate AN6 (AY322431). Trees were represented graphically with the software TREEVIEW version 1.4 (R. D. M. Page, Division of Environmental and Evolutionary Biology, University of Glasgow, Glasgow, UK).
DNA labelling.
High-molecular-mass genomic DNA was sheared by sonication with a Branson sonifier 250 on 50 % duty cycle at power level 5 for a total of 5 min. This produced DNA fragments with an average size of 0·5 kb. The sheared DNA was labelled with 32P by random-primed synthesis using an NEBlot kit (New England Biolabs). Labelled DNA was separated from unincorporated label by gel filtration using Probe-Quant G-50 Micro columns (Amersham Pharmacia Biotech).
DNA–DNA hybridization.
Measurements of reassociation between genomes were performed by a modified version of the competitive hybridization method of De Ley et al. (1970). Nitrocellulose membranes (Nitropure, MSI), to which 2 µg denatured, high-molecular-mass genomic DNA was bound, were incubated with 0·2 µg sheared, denatured [32P]DNA of the same isolate, in the presence or absence of a 250-fold excess of competitor DNA. Competitor DNA was 50 µg unlabelled, sheared, denatured DNA of either the same isolate or another isolate. Hybridization was performed in a buffer of 2x SSC (Sambrook et al., 1989)+30 % (v/v) formamide for 18 h at 50 °C. After five brief rinses in 2x SSC+0·1 % (w/v) SDS at room temperature, filters were washed twice in the same solution at 50 °C for 30 min. Filter-bound radioactivity was measured by using a Beckman LS2800 scintillation counter. Reassociation (%) was calculated as 100x(counts in the absence of competing DNA–counts with heterologous competing DNA)/(counts in the absence of competing DNA–counts with homologous competing DNA).
Thermal stability of DNA–DNA hybrids was determined after hybridizing, as described above, filter-bound DNA and [32P]DNA of either the same isolate or another isolate. After hybridization, filters were washed five times in 2x SSC+0·1 % SDS at room temperature and then incubated in successive 2 ml volumes of 2x SSC+30 % (v/v) DMSO at a range of temperatures between 50 and 95 °C. The amount of radioactivity released from the filters at each temperature was measured by Cerenkov radiation, with tritium discriminators on a Beckman LS2800 scintillation counter.
Tm, defined as the temperature at which 50 % of the radioactive probe remains bound to the filter, was calculated for each hybridization, and the difference in melting temperature between homologous and heterologous duplexes (Tm) was used as a measure of the similarity between DNA sequences.
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RESULTS AND DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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Phenotypic characterization
Each of the six Australian clinical isolates was Gram-negative, rod-shaped and motile with peritrichous flagella. All were phase II variants; they were not noticeably pigmented on nutrient agar, but were either yellow (9800946), off-white (GCH001) or cream on egg-yolk agar. None adsorbed dye from MacConkey agar or showed antibiotic activity against Micrococcus luteus. Isolate GCH001 showed a weak lecithinase reaction on egg-yolk agar; all others were negative. Few cells produced inclusions. Luminescence was detected in all isolates by scintillation counting, but not usually by the dark-adapted eye, whether the bacteria were in liquid or on solid media. All isolates were positive for catalase, lipase (Tween 20) and gelatin hydrolysis (Kohn's) and were negative for nitrate reduction, oxidase, Voges–Proskauer reaction, hydrogen sulfide production, -galactosidase activity with o-nitrophenyl -D-galactopyranoside, urease, tryptophan deaminase, indole, lysine and ornithine decarboxylases and arginine dihydrolase. All isolates were totally or partially haemolytic on sheep and horse blood agar; annular haemolysis (Akhurst et al., 1996) was observed on sheep blood agar for isolates MB, 9800946 and GCH001, but not for the four isolates that were obtained from one patient (9802336, etc.; see Table 1); none of the isolates displayed annular haemolysis on horse blood agar. Isolate 9800946 exhibited total haemolysis on sheep blood agar and partial haemolysis on horse blood agar, whereas the four isolates from the same patient exhibited partial and total haemolysis on sheep and horse blood agar, respectively. Total haemolysis was recorded for the MB isolate on both sheep and horse blood agar. Maximum temperature for growth of all Australian clinical isolates was 40 °C, whereas the maximum growth temperature for USA isolates of P. asymbiotica was 38 °C.
Acid, but not gas, was produced from glucose by all isolates. Other carbon sources from which acid was produced by all isolates were: fructose, glycerol, N-acetyl-D-glucosamine, mannose, maltose and ribose. No acid was produced by any isolate from adonitol, aesculin, amygdalin, L-arabinose, D- or L-arabitol, arbutin, cellobiose, dulcitol, erythritol, D-fucose, galactose, gentiobioise, glycogen, inulin, 2-ketogluconate, lactose, D-lyxose, mannitol, melezitose, melibiose, methyl D-glucoside, methyl D-mannoside, raffinose, rhamnose, sorbitol, sorbose, starch, sucrose, D-tagatose, trehalose, D-turanose, xylitol, -D-xyloside or D- or L-xylose. All isolates assimilated the following: glucose, mannose, maltotriose, maltose, ribose, glycerol, L-malate, citrate, N-acetyl-D-glucosamine, D-gluconate, succinate, fumarate, L-aspartate, L-glutamate, L-proline, L-alanine and L-serine. None of the following compounds was assimilated by any isolate: DL-
-amino-n-butyrate, adonitol, aesculin, DL--amino-n-valerate, L-arabinose, D- or L-arabitol, benzoate, betaine, caprate, caprylate, D-cellobiose, m-coumarate, dulcitol, i-erythritol, ethanolamine, D-(+)-galactose, D-galacturonate, -gentiobiose, glutarate, DL-glycerate, histamine, L-histidine, m-hydroxybenzoate, DL--hydroxybutyrate, hydroxyquinoline--glucuronide, itaconate, 2-keto-D-gluconate, 5-keto-D-gluconate, DL-lactate, -lactose, lactulose, D-lyxose, malonate, maltitol, D-mannitol, D-melezitose, -D-(+)-melibiose, methyl -D-glucopyranoside, methyl -galactopyranoside, methyl -D-glucopyranoside, methyl -galactopyranoside, methyl D-glucopyranose, mucate, 2-oxoglutarate, palatinose, phenyl 3-propionate, phenylacetate, p-hydroxybenzoate, protocatechuate, putrescine, quinate, D-(+)-raffinose, -L-rhamnose, D-saccharate, D-sorbitol, L-(+)-sorbose, sucrose, D-tagatose, D-(–)-tartrate, L-(+)-tartrate, meso-tartrate, trans-aconitate, tricarballylate, trigonelline, tryptamine, L-tryptophan, D-turanose, xylitol or D-xylose.
Each of the six Australian clinical isolates killed Galleria mellonella larvae when injected.
Characters that varied between isolates are presented in Table 2.
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, who characterized the same isolates, with the exception of GCH001. For example, Peel et al. (1999) reported that the isolates were all positive for urease and negative for DNase and acidification of maltose, whereas this study found no urease activity, that all Australian clinical isolates acidified maltose and that only isolate 9800946 was negative for DNase. Different protocols, involving different temperatures and times of incubation, were employed for many tests in these two studies. Peel et al. (1999) conducted almost all their biochemical tests at 35 °C, whereas the corresponding tests reported here were conducted at 28 °C. The choice of protocols and conditions in the study reported here was made to remain consistent with those used previously to characterize isolates of the genus Photorhabdus (Boemare & Akhurst, 1988; Fischer-Le Saux et al., 1999). This allowed comparison of the data gathered in this study with those obtained in earlier studies, to test whether the novel taxon suggested by sequence and DNA–DNA hybridization analyses could be differentiated from existing taxa in the genus Photorhabdus on the basis of phenotypic characters.
DNA–DNA hybridization
DNA of isolate MB reassociated most strongly with that of the P. asymbiotica isolate 3265-86T (Table 3) and its reassociation with isolates from other Photorhabdus spp. was always <55 %. Tm for isolate MB was >6 °C for all isolates, except for 3265-86T (4·1 °C) and Q614(A) (4·9 °C). DNA of isolate Q614(A) had 52 % reassociation with that of isolate MB and only 31 % with that of isolate 3265-86T of P. asymbiotica.
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. These groupings were the same in all analyses. The relationship of these subspecies-level groupings with each other, i.e. their organization into recognized species, was not supported by these analyses. Each different analysis gave different groupings of the subspecies into higher-level clades, and few of these higher-level groupings were given any statistical support (Fig. 1). The 16S rRNA gene sequences of the four Australian clinical isolates were slightly more similar to those from isolates of P. asymbiotica (mean Jukes–Cantor-corrected distance, 2·95 %; range, 2·91–2·99 %) than to sequences from members of other clusters (mean Jukes–Cantor-corrected distance, 3·66 %; range, 3·29–4·19 %).
Phylogenies based on 16S rRNA gene sequences have previously been used to distinguish several groups within the genus Photorhabdus that were consistent with DNA–DNA hybridization and phenotypic data (Szállás et al., 1997; Fischer-Le Saux et al., 1999). However, 16S rRNA gene sequence analysis proved less reliable with the additional set of sequences used in this study. Although phylogenetic analysis of 16S rRNA gene sequences supported the grouping of isolates into the subspecies groupings identified by Fischer-Le Saux et al. (1999), these subspecies groupings did not form consistent groups at the species level. Isolates of Photorhabdus luminescens subsp. luminescens did not cluster closely with the other two subspecies of P. luminescens, and P. asymbiotica could not be distinguished as a separate species (Fig. 1). The inconsistency of the species-level groupings in these analyses suggests that 16S rRNA gene sequence data are not suitable for the definition of species within the genus Photorhabdus, although they are useful at the subspecies level. This is because sequence differences are restricted to a few variable regions, and so the information content is not sufficient to provide reliable species-level groupings. Members of each subspecies displayed sequences that were very similar, even in variable regions, thus drawing the members of the subspecies together.
In contrast to 16S rDNA data, the gyrB gene provides useful data for investigating bacterial intrageneric phylogenies (Venkateswaran et al., 1999; Kasai et al., 2000; Yamamoto et al., 2000). The gyrB sequence data for Photorhabdus provided a good match with DNA–DNA hybridization and phenotypic data at both subspecies and species levels (Fig. 2). This is because the proportion of invariant nucleotide positions is lower than in the 16S rRNA gene. Each of the four clustering methods tested in the analysis of the gyrB data grouped isolates into the subspecies recognized by Fischer-Le Saux et al. (1999), and then the subspecies into recognized species in a highly reproducible (all four methods) and statistically significant (bootstrap values of 99 % or greater) manner. It is clear from these analyses that the species and subspecies identified by Fischer-Le Saux et al. (1999) are valid.
There may be more than the three subspecies of P. luminescens that were proposed by Fischer-Le Saux et al. (1999). In analysis of gyrB sequences, although the two Chinese isolates C8404 and C8406 clustered consistently with P. luminescens, the level of sequence divergence suggests that they may not fall into any of the recognized subspecies. Akhurst et al. (1996) showed that these isolates had 64–82 % DNA–DNA reassociation with P. luminescens subsp. luminescens isolate Hb and P. luminescens subsp. akhurstii isolate D1. Isolate C8406 forms a discrete PCR-RFLP profile, with greatest similarity to those of the P. luminescens subsp. luminescens group (Brunel et al., 1997; Fischer-Le Saux et al., 1999) and isolate C8404 produces a novel 16S rRNA gene PCR-RFLP profile that is most similar to that of the P. luminescens subsp. akhurstii group (S. Pages, personal communication). The gyrB, DNA relatedness and PCR-RFLP data suggest that, whilst they are members of the species P. luminescens, isolates C8404 and C8406 do not fall into any recognized subspecies. The gyrB analysis also suggests that they may not belong to a common subspecies.
In all analyses of gyrB sequences, two main clusters of Photorhabdus temperata isolates were formed and isolate NZH3 always branched more deeply, but coherently, with these. The mean Jukes–Cantor-corrected distance between members of different clusters was 4·95 % (range, 4·10–6·57 %). This suggests that there are three genetically distinct lineages of P. temperata, given that levels of sequence divergence between these clusters were similar to those for recognized subspecies of P. luminescens (mean Jukes–Cantor-corrected distance between members of different clusters was 5·06 %, with a range of 4·56–5·70 %).
Isolates MB, GCH001, 9800946 and 9802892T always grouped together and formed a sister group to the cluster of P. asymbiotica isolates. Isolate Q614(A) always branched more deeply, but reproducibly, with isolates of P. asymbiotica and the Australian clinical isolates. The mean Jukes–Cantor-corrected distance between any of the American isolates of P. asymbiotica and any of the four Australian clinical isolates was 5·57 % (range, 5·23–6·18 %). Coherence of the branching pattern in all analyses, statistical support for the groupings and the level of sequence divergence between any of the American or Australian clinical isolates and isolate Q614(A) (5·51–7·54 %) all suggested strongly that there are at least three genetically distinct lineages of P. asymbiotica. In analyses of gyrB sequences, isolate Q614(A) always grouped with the four Australian clinical isolates and the P. asymbiotica isolates; the bootstrap value for this grouping was 100 %.
Non-luminescent isolate Q614
The unexpected grouping of the non-luminescent isolate Q614 (Akhurst & Boemare, 1986) with a subgroup of P. luminescens based on comparative 16S rRNA gene sequence analysis (Szállás et al., 1997) was investigated. An earlier DNA–DNA hybridization study had shown that isolate Q614, recovered from a nematode host and characterized by two of the authors (R. J. A. and N. E. B), had <50 % reassociation with the DNA of two P. luminescens isolates (Boemare et al., 1993). The correct identity of the original isolate from the CSIRO Culture Collection was demonstrated by its lack of bioluminescence when tested by scintillation counter, as it is the only non-luminescent isolate of Photorhabdus in the CSIRO collection. The 16S rRNA gene sequence of that isolate, labelled Q614(A), had only 95·5 % sequence similarity to the sequence submitted by Szállás et al. (1997), labelled Q614(S). It is evident that the isolate used by Szállás et al. (1997) was identified incorrectly as the non-luminescent isolate Q614 and that the sequence deposited with GenBank accession no. Z76749 is annotated incorrectly. Consequently, the phylogenetic positioning of isolate Q614 in subcluster IV (Szállás et al., 1997) is incorrect.
The verified isolate Q614(A) did not branch consistently with any subspecies grouping in different analyses of 16S rRNA gene sequences, which is consistent with our observation that comparative analysis of this gene is not able to group isolates into recognized species. In contrast, analyses of gyrB sequences always grouped Q614(A) with the four Australian clinical isolates and with isolates of P. asymbiotica; the bootstrap value for this grouping was 100 % (Fig. 2). This suggests that Q614(A) may represent a novel subspecies of P. asymbiotica. However, DNA–DNA hybridization data suggest that Q614(A) is not conspecific with P. asymbiotica, as it had only 52 and 31 % reassociation with isolates MB and 3265-86T, respectively, and a Tm of nearly 5 °C with isolate MB (Table 2). As Q614(A) is the only representative of a putative subgroup, it would be prudent to wait until more isolates are available for investigation before a formal decision on its taxonomic position can be made.
Clinical isolates of Photorhabdus
Both the gyrB phylogeny and DNA–DNA hybridization data show that the Australian and USA clinical isolates are closely related (Fig. 2, Table 2). As isolate MB had 76 % DNA–DNA reassociation and a Tm of 4·1 °C with P. asymbiotica isolate 3265-86T, they meet the definition of Wayne et al. (1987) for being regarded as conspecific. In the phylogenetic analysis conducted with gyrB sequence data, the USA clinical isolates, identified previously as P. asymbiotica, and the Australian clinical isolates formed a discrete group, albeit with Q614(A). Further, with each of the four clustering methods employed in the gyrB analysis, the Australian clinical isolates and the USA clinical isolates formed discrete subgroups. Moreover, DNA–DNA reassociation and Tm values for the MB/3265-86T combination were near the species definition limits of Wayne et al. (1987) of 70 % and 5 °C, respectively, and the two subgroups could be differentiated on the basis of several phenotypic characters (Table 4). On the basis of these differences, we propose that the Australian and USA clinical isolates of the genus Photorhabdus should be assigned to different subspecies of P. asymbiotica.
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Cells are Gram-negative rods of 2–3x0·5–1·0 µm that are motile by peritrichous flagella. Maximum temperature for growth in nutrient broth is 40 °C. Yellow, brown or no pigment. Catalase-positive; oxidase-negative; nitrate is not reduced. Negative for o-nitrophenyl -D-galactopyranoside activity and the Voges–Proskauer test. Bioluminescent. No phase I isolates have been detected; isolates do not absorb dyes, are negative for lecithinase on egg-yolk agar and are, at most, weakly antibiotic. Positive for Simmons' citrate. Negative for H2S from triple-sugar iron agar, tryptophan deaminase, lysine and ornithine decarboxylases, arginine dihydrolase and indole. Acid is produced from fructose, N-acetylglucosamine, glucose, glycerol, maltose, mannose and ribose. No gas is produced from glucose. Proteinaceous inclusions are produced poorly. Annular haemolysis is usually evident on sheep blood agar at 25 °C. Does not use L-fucose, DL-lactate or mannitol.
The type strain is 3265-86T (=ATCC 43950T=CIP 106331T=DSM 15149T). Natural habitat is uncertain; all isolates were obtained from human clinical specimens. The GenBank accession numbers for the 16S rRNA and gyrB gene sequences of the type strain are Z76755 (Szállás et al., 1997) and AY278494, respectively.
Description of Photorhabdus asymbiotica subsp. asymbiotica subsp. nov.
Photorhabdus asymbiotica subsp. asymbiotica (a.sym.bi.o'ti.ca. Gr. pref a not; N.L. fem. adj. symbiotica living together; N.L. fem. adj. asymbiotica not symbiotic).
Maximum temperature for growth in nutrient broth is 37–38 °C. Yellow or brown pigment. Positive for urease and aesculin hydrolysis; weakly positive for Simmons' citrate. Negative for DNase. Acid is produced from trehalose and usually from aesculin; variable for gluconate. Proteinaceous inclusions are produced poorly. Annular haemolysis occurs on sheep blood and horse blood agars. Positive for Tween 60 and Tween 80 esterases; Tween 40 esterase activity is variable.
The type strain is 3265-86T (=ATCC 43950T=CIP 106331T=DSM 15149T). Natural habitat is uncertain; all isolates were obtained from human clinical specimens in the USA. The GenBank accession numbers of the 16S rRNA and gyrB gene sequences of the type strain are Z76755 (Szállás et al., 1997) and AY278494, respectively.
Description of Photorhabdus asymbiotica subsp. australis subsp. nov.
Photorhabdus asymbiotica subsp. australis (aus.tra'lis. L. fem. adj. australis southern; this subspecies was detected in the southern hemisphere).
Maximum temperature for growth is 40 °C. Yellow or no pigment; weakly pigmented. Most isolates are positive for DNase and most are negative for aesculin hydrolysis. Negative for urease. Most isolates produce acid from gluconate; variable for acid production from aesculin and negative for trehalose. Proteinaceous inclusions are rare. Annular haemolysis is variable on sheep blood and horse blood agars. Most isolates are negative for Tween 60 and Tween 80 esterases and most grow on myo-inositol.
The type strain is 9802892T (=CIP 108025T=ACM 5210T). Natural habitat is uncertain; all isolates were obtained from human clinical specimens in Australia. The GenBank accession numbers of the 16S rRNA and gyrB gene sequences of the type strain are AY280572 and AY278496, respectively.
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ACKNOWLEDGEMENTS |
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REFERENCES |
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Akhurst, R. J. (1983). Taxonomic study of Xenorhabdus, a genus of bacteria symbiotically associated with insect pathogenic nematodes. Int J Syst Bacteriol 33, 38–45.
Akhurst, R. J. (1987). Use of starch gel electrophoresis in the taxonomy of the genus Heterorhabditis (Nematoda: Heterorhabditidae). Nematologica 33, 1–9.
Akhurst, R. J. & Boemare, N. E. (1986). A non-luminescent strain of Xenorhabdus luminescens (Enterobacteriaceae). J Gen Microbiol 132, 1917–1922.
Akhurst, R. J., Bedding, R. A., Bull, R. M. & Smith, D. R. J. (1992). An epizootic of Heterorhabditis spp. (Heterorhabditidae: Nematoda) in sugar cane scarabaeids (Coleoptera). Fundam Appl Nematol 15, 71–73.
Akhurst, R. J., Mourant, R. G., Baud, L. & Boemare, N. E. (1996). Phenotypic and DNA relatedness between nematode symbionts and clinical strains of the genus Photorhabdus (Enterobacteriaceae). Int J Syst Bacteriol 46, 1034–1041.
Bedding, R. A. & Miller, L. A. (1981). Use of a nematode, Heterorhabditis heliothidis, to control black vine weevil, Otiorhynchus sulcatus, in potted plants. Ann Appl Biol 99, 211–216.
Bleakley, B. & Nealson, K. H. (1988). Characterization of primary and secondary forms of Xenorhabdus luminescens strain Hm. FEMS Microbiol Ecol 53, 241–250.
Boemare, N. E. & Akhurst, R. J. (1988). Biochemical and physiological characterization of colony form variants in Xenorhabdus spp. (Enterobacteriaceae). J Gen Microbiol 134, 751–761.
Boemare, N. E., Akhurst, R. J. & Mourant, R. G. (1993). DNA relatedness between Xenorhabdus spp. (Enterobacteriaceae), symbiotic bacteria of entomopathogenic nematodes, and a proposal to transfer Xenorhabdus luminescens to a new genus, Photorhabdus gen. nov. Int J Syst Bacteriol 43, 249–255.
Brunel, B., Givaudan, A., Lanois, A., Akhurst, R. J. & Boemare, N. (1997). Fast and accurate identification of Xenorhabdus and Photorhabdus species by restriction analysis of PCR-amplified 16S rRNA genes. Appl Environ Microbiol 63, 574–580.[Abstract]
De Ley, J., Cattoir, H. & Reynaerts, A. (1970). The quantitative measurement of DNA hybridization from renaturation rates. Eur J Biochem 12, 133–142.[Medline]
Farmer, J. J., III, Jorgensen, J. H., Grimont, P. A. D. & 8 other authors (1989). Xenorhabdus luminescens (DNA hybridization group 5) from human clinical specimens. J Clin Microbiol 27, 1594–1600.
Felsenstein, J. (1989). PHYLIP – Phylogeny inference package (version 3.2). Cladistics 5, 164–166.
Fischer-Le Saux, M., Viallard, V., Brunel, B., Normand, P. & Boemare, N. E. (1999). Polyphasic classification of the genus Photorhabdus and proposal of new taxa: P. luminescens subsp. luminescens subsp. nov., P. luminescens subsp. akhurstii subsp. nov., P. luminescens subsp. laumondii subsp. nov., P. temperata sp. nov., P. temperata subsp. temperata subsp. nov. and P. asymbiotica sp. nov. Int J Syst Bacteriol 49, 1645–1656.
Fitch, W. M. & Margoliash, E. (1967). Construction of phylogenetic trees. Science 155, 279–284.
Gerrard, J. G., McNevin, S., Alfredson, D., Forgan-Smith, R. & Fraser, N. (2003). Photorhabdus species: bioluminescent bacteria as emerging human pathogens? Emerg Infect Dis 9, 251–254.[Medline]
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.
Kasai, H., Tamura, T. & Harayama, S. (2000). Intrageneric relationships among Micromonospora species deduced from gyrB-based phylogeny and DNA relatedness. Int J Syst Evol Microbiol 50, 127–134.[Abstract]
Peel, M. M., Alfredson, D. A., Gerrard, J. G., Davis, J. M., Robson, J. M., McDougall, R. J., Scullie, B. L. & Akhurst, R. J. (1999). Isolation, identification, and molecular characterization of strains of Photorhabdus luminescens from infected humans in Australia. J Clin Microbiol 37, 3647–3653.
Poinar, G. O., Jr, Thomas, G. M. & Hess, R. (1977). Characteristics of the specific bacterium associated with Heterorhabditis bacteriophora (Heterorhabditidae: Rhabditida). Nematologica 23, 97–102.
Poinar, G. O., Jr, Thomas, G. M., Haygood, M. & Nealson, K. H. (1980). Growth and luminescence of the symbiotic bacteria associated with the terrestrial nematode, Heterorhabditis bacteriophora. Soil Biol Biochem 12, 5–10.
Poinar, G. O., Jr, Jackson, T. & Klein, M. (1987). Heterorhabditis megidis sp. n. (Heterorhabditidae: Rhabditida), parasitic in the Japanese beetle, Popillia japonica (Scarabaeidae: Coleoptera), in Ohio. Proc Helminthol Soc Wash 54, 53–59.
Saitou, N. & Nei, M. (1987). The neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol Biol Evol 4, 406–425.[Abstract]
Sambrook, J., Fritsch, E. F. & Maniatis, T. (1989). Molecular Cloning: a Laboratory Manual, 2nd edn. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory.
Smits, P. H. & Ehlers, R.-U. (1991). Identification of Heterorhabditis spp. by morphometric characters and RFLP and of their symbiotic bacteria Xenorhabdus by species-specific DNA probes. IOBC/WPRS Bull 14, 195–201.[Medline]
Szállás, E., Koch, C., Fodor, A., Burghardt, J., Buss, O., Szentirmai, A., Nealson, K. H. & Stackebrandt, E. (1997). Phylogenetic evidence for the taxonomic heterogeneity of Photorhabdus luminescens. Int J Syst Bacteriol 47, 402–407.
Thompson, J. D., Gibson, T. J., Plewniak, F., Jeanmougin, F. & Higgins, D. G. (1997). The CLUSTAL_X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res 25, 4876–4882.
Venkateswaran, K., Moser, D. P., Dollhopf, M. E. & 10 other authors (1999). Polyphasic taxonomy of the genus Shewanella and description of Shewanella oneidensis sp. nov. Int J Syst Bacteriol 49, 705–724.
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.
Yamamoto, S. & Harayama, S. (1995). PCR amplification and direct sequencing of gyrB genes with universal primers and their application to the detection and taxonomic analysis of Pseudomonas putida strains. Appl Environ Microbiol 61, 1104–1109.[Abstract]
Yamamoto, S., Kasai, H., Arnold, D. L., Jackson, R. W., Vivian, A. & Harayama, S. (2000). Phylogeny of the genus Pseudomonas: intrageneric structure reconstructed from the nucleotide sequences of gyrB and rpoD genes. Microbiology 146, 2385–2394.
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