mitted by the infective dauer larvae and converts into phase. II when cultured in vitro (1). The phase variants of all. Xenorhabdus spp.can be distinguished by ...
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0099-2240/90/010181-06$02.00/0 Copyright © 1990, American Society for Microbiology
Development and Application of Oligonucleotide Probes for Molecular Identification of Xenorhabdus Species JORN PUTZ,l FRANK MEINERT,l URS WYSS,2 RALF-UDO EHLERS,2 AND ERKO STACKEBRANDTL* Institut fur Allgemeine Mikrobiologiel and Institut fur Phytopathologie,2 Christian-Albrechts-Universitat, Olshausenstr. 40, 2300 Kiel, Federal Republic of Germany Received 13 July 1989/Accepted 11 October 1989
Synthetic deoxyoligonucleotide probes that hybridized against the region at positions 455 through 480 of 16S rRNA were developed for the identification of all five Xenorhabdus species. Sequence variation in the respective rRNA region between two strains of Xenorhabdus luminescens in addition allowed the construction of two strain-specific probes. Of 27 isolates determined to be Xenorhabdus strains by phenotypic characterization, 24 could be assigned to four of the five species. Two strains (HL-1 and HL-2) isolated from a Heterorhabditis sp. and a single strain (D-1.1) isolated from Steinernema affinis showed no hybridization signal with any of the five species-specific probes. With regard to the available species descriptions of nematodes, the results presented here confirm that, except for Steinernema affinis, the individual nematode hosts harbor only a single Xenorhabdus species.
is a symbiont of Steinernema bibionis, S. affinis, and S. kraussei. The purpose of this study was to develop nucleic acid probes for a genetically based identification of strains of the five species of Xenorhabdus to introduce a reliable and rapid method to separate the species and to distinguish them from other enterobacteria as well as from pseudomonads and flavobacteria. The last two groups of organisms show not only similar colony morphology on diagnostic media but often occur as contaminants in in vitro cultures for mass production of nematodes (Ehlers, unpublished observation). Furthermore, if the specificity between host and symbiont were exclusive, identification of the bacterial species would contribute to nematode classification.
Bacteria of the genus Xenorhabdus are mutualistically associated with entomopathogenic rhabditid nematodes of the genera Steinernema (syn. Neoaplectana) and Heterorhabditis (3, 24). The nematode-bacterium complex exhibits great potential as a biological agent for insect pest control in cryptic environments (16, 20). The free-living L3 dauer larva embeds cells of the symbiotic bacterium in an intestinal vesicle (7) and releases them after penetrating into the host insect's hemocoel. The parasitic nematode stages feed on multiplied bacteria and disintegrated insect tissues (17, 21, 22). Xenorhabdus isolates tend to produce two colony forms, designated phases I and II (primary and secondary forms, respectively). The unstable phase I is preferentially transmitted by the infective dauer larvae and converts into phase II when cultured in vitro (1). The phase variants of all Xenorhabdus spp. can be distinguished by absorption of bromothymol blue or neutral red from agar media (1, 4) and among isolates of species and strains by different biochemical and physiological characters (8). However, the published phenotypic descriptions of Xenorhabdus strains very often contradict each other (3, 9, 14, 24, 25). It has been suggested (8) that one of the reasons for the inconsistency of results may be the different reactions of the phase variations (5, 6, 8), which make classification of Xenorhabdus spp. a difficult and time-consuming task, especially because some strains of Xenorhabdus luminescens do not exhibit phase II characters, even when subcultured biweekly for half a year (Ehlers, unpublished observation). The number of validly described species has recently been increased from two (Xenorhabdus nematophilus and X. luminescens) to five by elevating three subspecies of X. nematophilus to species status (5). Every described Xenorhabdus sp. can be appointed to a different host nematode species; X. luminescens is a symbiont of Heterorhabditis spp., X. nematophilus is a symbiont of Steinernema feltiae, Xenorhabdus poinarii is a symbiont of Steinernema glaseri and Steinernema sp. strain NC513, Xenorhabdus beddingii is a symbiont of a new genus of the family Steinernematidae and unnamed Steinernema spp., and Xenorhabdus bovienii *
MATERIALS AND METHODS Isolation of Xenorhabdus strains. Symbionts of Steinernema sp. strain Q58, S. bibionis T228, and S. glaseri G1 were obtained from the Deutsche Sammlung von Mikroorganismen, Braunschweig, Federal Republic of Germany (DSM strains). One isolate each of X. nematophilus and S. feltiae DD-136 and two X. luminescens strains (one from Heterorhabditis bacteriophora HB and the other from H. heliothidis NC1) were obtained from the American Type Culture Collection, Rockville, Md. (ATCC strains). Symbiotic bacteria other than the DSM and ATCC strains were isolated from L3 dauer larvae of their associated nematode by the method of Akhurst (1). Table 1 is a compilation of the bacterial strains investigated, their host nematodes, and the geographical sources. Several gram-negative reference strains (IFAM strains) were taken from the Culture Collection of the Institut fur Allgemeine Mikrobiologie, Kiel, Federal Republic of Germany. Phase I symbionts were separated from phase II variants as described previously
(1-3).
Isolation of crude rRNA and sequence analysis. Protocols for mechanical lysis of 4 g (wet weight) of cells with glass beads, isolation of crude rRNA, and sequence analysis of parts of the 16S rRNA primary structure with reverse transcriptase and selective cDNA primers were as described previously (13, 19). Synthesis and labeling of oligodeoxynucleotides. DNA oli-
Corresponding author. 181
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TABLE 1. Bacterial strains and nematode hosts used in this study Reference no. in figures
1 2 3 4 5 6 7
Associated nematode
Bacterium
X. beddingii DSM 4764T X. beddingii DSM 4765 X. nematophilus ATCC 19061T DD-1 DD-2 P-1
P-5
Geographical
Steinernema sp. strain Q58 Steinernema sp. strain Q58
Australia Australia
S. feltiae S. feltiae S. feltiae S. feltiae S. feltiae
United States United States United States Poland Poland
DD-136 DD-136 DD-136 Pieridarum Pieridarum
source
8 9 10 11 12 13 14 15 16 17 18 19 20
X. bovienli DSM 4766T DSM 4767 G-1 G-3 S-1.1 S-2.1 S-3.1 0-2 0-7 D-1.1 D-2.1 H-1 H-2
S. bibionis T228 S. bibionis T228 S. bibionis N8 S. bibionis N8 S. bibionis SF1 S. bibionis SF2 S. bibionis SF3 S. bibionis OBSIII S. bibionis OBSIII S. affinis Dl S. affinis D2 Heterorhabditis heliothidis NC1 H. heliothidis NC1
Australia Australia Czechoslovakia Czechoslovakia Federal Republic of Germany Federal Republic of Germany Federal Republic of Germany Netherlands Netherlands Federal Republic of Germany Federal Republic of Germany United States United States
21 22 23 24 25 26 27 28 29 30 31
X. luminescens ATCC 29304 ATCC 29999T HL-1 HL-2 HB-5 NZ-1 NZ-5 HW-1 R-1 V-1 V-16
H. heliothidis NC1 H. bacteriophora HB Heterorhabditis sp. strain Heterorhabditis sp. strain H. bacteriophora HB Heterorhabditis sp. strain Heterorhabditis sp. strain Heterorhabditis sp. strain Heterorhabditis sp. strain Heterorhabditis sp. strain Heterorhabditis sp. strain
United States Australia Netherlands Netherlands Australia New Zealand New Zealand Netherlands A? Australia Australia
32 33 34 35
X. poinarii UQM 2216T N-1 N-2 SG-1
S. glaseri Gi Steinernema sp. strain NC513 Steinernema sp. strain NC513 S. glaseri Gi
36
A. hydrophila IFAM 1223
37 38 39 40 41 42
Enterobacter aerogenes IFAM 1907 E. coli IFAM 1671 Enterobacter carotovora IFAM 1226 P. vulgaris IFAM 1011 Pseudomonasfluorescens IFAM 930 Serratia marcescens IFAM 947
gonucleotides were synthesized with an automatic synthesizer (Applied Biosystems, Weiterstadt; model 318A) as described by Beaucage and Caruthers (6). Oligomers were removed from the matrix with 3 ml of 33% (wt/vol) ammonium hydroxide for 16 h at 55°C. The solvent was evaporated in a vacuum centrifuge (SpeedVac concentrator; Savant), and the pellet was dissolved in 200 ,ul of water. The quality of the oligonucleotides was checked by fast-protein liquid chromatography. 5' labeling of probes and their purification by polyacrylamide gel electrophoresis were done as described previously (18). The sequences of the probes are shown in Fig. 2.
HL81 HL81 NZ NZ HW79 ROLANDO V16 V16
United United United United
States States States States
Hybridization. Depending on the experiment, the amount of crude rRNA loaded on Hybond-N membranes (Amersham) ranged between 1 ng and 5 ,ug. rRNAs were dissolved in 100 ,u1 of O.lx SSC (lx SSC is 0.15 M NaCl-0.015 M trisodium citrate [pH 7.2]), applied to the filters by a dot blot apparatus (Minifold SRC-96; Schleicher & Schuell, Dassel), and fixed to the filters by UV light (5 min at 254 nm). Hybridization was carried out in plastic bags. Conditions of prehybridization (11) and hybridization and washing steps (E. Stackebrandt and 0. Charfreitag, J. Gen. Microbiol., in press) have been documented. Hybridization temperatures, which depend upon the nucleotide composition of the oligo-
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40
C.CGGCAGGCCUAACACAUGCUAGUCGGACGCUAACAGGAAACAGCUUGCUI!ltCiUGCUGACCAGUGCGCGACGGGUl;C.AUIAUGUCUUCCCUGUUUGC GGAUNGGAUUAGCUNGUAGGCGCGGUGAUUGCCCACCUAGCG.ACCAUCCGUNGCUt!GUCUGAGAGAUUACCAGCCAACAUCUGGGCUGAGUACACGCCCI'C
C?GCCAUGGAGCGGGAUAACCACUGGAAACGGUNGCUAAUACCGCAUGACCUCUUGGCGAGUUAAGUCGGGACCUUCGCGCCUCACGCCAUCGUIGACCCACA
}!GACUCCUACGGGAGGCANUUGU}1GCGOAAUUUGCACAAUGGGCGCAAGCCUGAUGCAC CCAUGCCGGCGUGUAUGAAGAAGGCIIIItCGGC-UUGUAAAGU CUUUC AGCGCGGAGGAAGGCG UN AGUCUNA
gA&A00CVUACOLUgACGUUACCCGC AGAAG AAGC ACCGGCCU AACUC CC UC CCAGC AG C CC COG UAAU A
CGGAGGGUGCNAGCGUUAAUCGGAAUUACUGGGCGUtAAGCGCACGACAGGCGGUCAAUUAAGUUCGAtAGUGAAAUCCCCCCCUCUUAACCCCG.1'I!ACGCCA CseAcAGG CC tCd CCUAG At AUGrUGr AG G AAnUrAsCCGG UGG A AAdU AAUUoCCACGneapsCC9G0GG AG ArGGGRGA UCCAACrACUNGUUGGCUeIG AGUCUCGUtAG
CCCCUGGACGAAG ACUG ACGCUNAGGUGCGH AAGCGUGGGG AG CAAAC AGGAUUIJG AU ACCCUCGUl'G UCCA CGCUGUAAACGCAUG UCG AUUUGAG GCU IIUG CCCUUGAGGCNJM!}NNN NC CGG AGCUAACGCGUU AAAUCG ACCGCCUGGCG!GVGfNCGGCCG C1ACGGUU A AA A CUCAA AUG A AUUG ACGGCGGG CCC COA
CAAGCGGUNGAGCAUGUNGUUUAAUUCGAUGIIAACGCGAAGAACCUUACCUACUCUUGACAUCCACGCAAUUCCGCAGA.GAUGCCGGACGUCCUUCGGGA oooo*.UGAUAAACCGGAGGAAGGUGGGGAAGACGUCUAGUCAUCAUGCCCC ............................. ACCGUNJAGACAGGUGCUGCAUtlGCUNUCGUCAGCUCCU? UUACGAGUAGGGCUACACACGUGCUACAAUGGCAGAUACAAAGAGAAGCGACCUCGCGAGAGCAAGCGGAACUCAUCAAGUCUOUCGUAGUCCGGAUUGG ... AGUCUGCAACUCGACUCCAUGAAGUCCGAAUCGCUAGUAAUCGUAGAUI!AGAAUGCUACOGUGAAUACGUUCCCGGGCCUItCUACACACCG.1401 FIG. 1. Partial sequence of the 16S rRNA of X. nematophilus ATCC 19061 as determined by the reverse transcriptase sequencing method (13, 19). Numbers indicate the termini of the two fragments by the E. coli numbering system (10). The boxed region marks the target site of the strain-specific oligonucleotide probe. Dots indicate the unsequenced region. N, Nucleotide composition not determined.
nucleotide (23), are given in the figure legends. Autoradiography was for 12 to 24 h with intensifying screens (Dupont Cronex, Quanta III). RESULTS AND DISCUSSION Subcultures of strains used for the isolation of nucleic acids were checked for the presence of phase I and phase II characters. Most but not all strains of X. beddingii, X. nematophilus, X. bovienii, and X. poinarii had converted into phase II forms during the several passages of growth before biomass production. Phase II variants were also found for X. bovienii DSM 4766 and X. beddingii DSM 4764, originally deposited as phase I forms by Akhurst and Boemare (5) as UQM 2210 and UQM 2871, respectively (UQM strains are from the Culture Collection of the department of Microbiology, University of Queensland, St. Lucia, Queensland, Australia). With the exception of X. luminescens ATCC 29304 and ATCC 29999 and the strains H-1 and H-2, all other Heterorhabditis isolates still showed phase I characteristics. Comparison of partial sequences of the 16S rRNA from X. nematophilus ATCC 19061 (Fig. 1) with homologous counterparts from Escherichia coli and Proteus vulgaris (data not shown) revealed the presence of two regions (positions 455 through 485 and 1005 through 1025, according to the E. coli X. beddingil DSN 4764 probe X. nematophilus ATCC 19061 probe X. bovienih DSN 4766 X. X. X.
X.
probe luminescens ATCC 29999 probe luminescens ATCC 29304 probe luminescens consensus probe poinarhi UQN 2216 probe
E. coli P. vulgaris
numbering system [10]) of substantial sequence variation. The primary structure of these regions was determined for the type strains of the other four Xenorhabdus spp. as well. Because sequences of the region around position 470 showed a much higher degree of variation than did those around position 1015, this portion was selected as a target for synthetic oligonucleotide probes. The sequences of both the target and the probes are shown in Fig. 2. With a 32P-labeled "universal" probe that hybridizes to positions 787 through 803 of all eubacterial 16S rRNAs, as little as 1 ng of crude rRNA could be detected (Fig. 3). Assuming that 16S rRNA contributes to crude rRNA of about 30% (1,500 nucleotides for 16S rRNA versus 3,000 nucleotides for 23S rRNA, not taking into account 5S rRNA and tRNAs) the detectable amount was even less. For routine application 500 ng of each of the 42 strains tested was blotted on filters in a 7-by-6 matrix with continuous numeration (Table 1 and Fig. 4). Each experiment was done in duplicate, one with the universal probe to check the availability of rRNA at each spot (data not shown) and one with the species-specific probe (Fig. 4). In cases where weak signals with the universal probe revealed the presence of a substantially lower amount of rRNA than the average, the yield of the respective rRNA was redetermined and a new filter was prepared. The influence of the hybridization and washing tempera-
51CAGCG :CGGAGGAAGGCGUGGACCUGAAUACGGUUCACGAUUCACGUUACCCG 3'
3'CTOGACTTATOCCAAGTOCTA 5'
5'CAGCG ;GGGAGGAAOGCGUN;ACGICUtIAACAGGOCUUACGAUUGACGUUACCCG 3' 3'GTCCCGAATGCTAAC 5'
51CAGCC CGGGAGGAAGGCAACAGCGUAAAUAGCOCUGUUGAUUGACGIJUItCCCG 3' 3'TCGCATTTATCGCGACAACTA 5'
5'CAGCG:GGGAGOAAOGGUCCAGCCUGAAGAGGGUUAGACUUUGACGUUACCCG 3' 3'TTCTCCCAATCTCAAA 5' 51CACCG OGCGAGGAAGGCUAUCCCCUGAAGAGGGCGAUAGCUUGACGUUACCCG 3' 3'TTCTCCCGCTATCGAA 5'
3' CCGGACTTCTCCCGC 5'
5'CAGCGoGGGAGGAAGGCCGGAOCCUGAAUAAGGUUGGCGUUUGACGUUACCCG 3' 3'TCGGACTTATTCCAACCGCAA 5'
5'CAGCG;GGGACGAAGGGACUAAAGUUAAUACCUUUGCUCAUUGACGUUACCCG 3'
5'CAV'CG COGGAGGAACGUGAUAAACUUAAUACCPtUUGUCAAUUCACGUUACCCG
3'
FIG. 2. Partial sequences of 16S rRNA of type strains of Xenorhabdus species containing a variable region (positions 455 through 485) against which oligonucleotide probes were synthesized. Homologous sequences of E. coli and P. vulgaris are given for comparison.
184
PUTZ ET AL.
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w
APPL. ENVIRON. MICROBIOL.
I1
riL e
5-
rL s
W
EJ I ElJ
L0
rMk
50
gri
I*
a _mi--
5 00
rIL
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Jlfn
f5-
tx
FIG. 3. Autoradiogram of a dot blot hybridization snowinig the sensitivity of the detection system. Different amounts of crude rRNA from the isolates 0-7 (no. 16 in Table 1) (a) and V-1 (no. 30 in Table 1) (b) were hybridized with a 5'-32P-labeled probe in a buffer consisting of 6x SSC, 3x Denhardt solution, and 0.1% sodium bisphosphate at 28°C. After 2 h, filters were washed with 6x SSC at 20°C for 3 min and at 28°C for 10 min. The probe was a 17-mer oligonucleotide with the sequence 5' CTACCAGGGTATCTAAT 3'
(19).
a
d
EIi~~~~XjE1
ture on the specificity of the probe is illustrated in Fig. 5. A filter hybridized with a probe against X. luminescens ATCC 29999 at 20°C below the calculated melting point of the homologous hybrid was autoradiographed; almost all rRNA dots showed a more or less significant signal, including those of the enterobacterial reference organisms (no. 36 through 42). Washing of the filter at a stringent temperature (10°C below the Tm) removed the unspecific binding, leaving strong signals only for dots no. 19 through 21 and 24 through 31. Results of hybridization with the species-specific probes are shown in Fig. 4. Except for three strains D-1.1 (no. 17), HL-1 (no. 22), and HL-2 (no. 23), all other isolates could be assigned to one of four of the five species. None of the strains belonged to X. beddingii (no. 1 and 2). Four isolates (no. 4 through 7) were found to be strains of X. nematophilus. Eight strains (no. 10 through 16 and 18) gave positive results with the probe developed against X. bovienii. Three strains (no. 33 through 35) were assigned to X. poinarii. The hybridization signals were slightly weaker than those of the homologous reaction, which possibly indicates a single mismatch somewhere in the middle part of the target sequence of the symbionts of Steinernema sp. strain NC513. This nematode resembles S. glaseri in morphological and physi-
ological characteristics (Ehlers, unpublished observation). Three probes were constructed for X. luminescens. A consensus probe was developed for strains ATCC 29999 and ATCC 29304, which were found to be rather unrelated in a recent phylogenetic study (12). This probe detected both described strains and all but two isolates (HL-1 and HL-2) that in biochemical tests reacted like X. luminescens. The probe was exactly complementary to the target of strain
b
dl
j]n9EW j]f
E15Ui
6
c
d2
EfWIW
nF,iI n
e
ILfEE
FIG. 4. Autoradiograms of dot blot hybridization between crude rRNA of various Xenorhabdus strains and species-specific oligonucleotide probes. Conditions are as in the legend to Fig. 3, except for hybridization and washing temperatures, respectively, which are indicated within parentheses: a, probe X. beddingii (42 and 52°C); b, probe X. nematophilus (28 and 38°C); c, probe X. bovienii (40 and 50°C); d, probe X. luminescens (32 and 42°C; see also Fig. 5); d1, probe ATCC 29999 (26 and 36°C); d2, probe ATCC 29304 (28 and 38°C); e, probe X. poinarii (42 and 52°C).
MOLECULAR IDENTIFICATION OF XENORHABDUS SPECIES
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b
a
iSi!liESiSEE FIG. 5. Autoradiogram of a dot blot hybridization showing the influence of the washing temperature on the specificity of the probe-target hybrid. Crude rRNA (500 ng) from each strain (no. 1 through 42) was hybridized with a 15-mer oligonucleotide specific for X. luminescens strains (consensus probe in Fig. 1). Numbers 1 through 35 refer to Xenorhabdus strains as indicated in Table 1. Numbers 36 through 42 are enterobacterial reference organisms. (a) Hybridization and washing temperature at 32°C, all other conditions as in the legend to Fig. 3; (b) washing under stringent conditions, i.e., at 42°C (10°C below the Tm of the hybrid), which removed all unspecific signals.
ATCC 23904, whereas the respective sequence of strain ATCC 29999 differs in three positions: a G residue of the probe bound to a U residue, and there were two differences at the termini that contributed insignificantly only to the thermal stability of the hybrids. The other two X. luminescens probes were strain specific. The probe for strain ATCC 29304 (no. 21) also identified the two isolates NC19 H-1 and H-2, whereas the probe for strain ATCC 29304 (no. 24) gave positive, although in most cases weaker, hybridization signals with rRNA of an additional seven isolates from a different geographical area (no. 25 through 31). None of the classical methods used so far (5, 14) has been sufficiently powerful to detect the presence of strain clusters within this species. The probes do not discriminate between rRNAs isolated from either phase I or phase II variants. This is obvious from Fig. 4; the probe developed against a phase II strain (ATCC 29999) gave hybridization signals of the same intensity as those given by rRNA from a phase I variant. The results of the probe application confirm the high specificity of the nematode-bacterium relationship. The only exceptions refer to the isolates D-1.1 and D-1.2 of S. affinis. Although rRNA of the former isolate gave no signal with any of the five probes, rRNA of the latter isolate hybridized with the X. bovienii probe. It may therefore be speculated that S. affinis, which lives associated with S. bibionis (both nematode species were isolated from the same location at Dannau, Schleswig-Holstein, Federal Republic of Germany), contains two different symbionts. Strain D-1.1, which is specific for S. affinis, may actually constitute a new Xenorhabdus species. Strain D-2.1, on the other hand, is a strain of X. bovenii that might have been taken up from S. bibionis when both nematode and Xenorhabdus strains had multiplied in the same insect larva. The other two isolates that gave negative hybridization results may also be representatives of a new species. These strains (HL-1 and HL-2) are symbionts of a yet unidentified species of Heterorhabditis. As in the case for strain D-1.1, sequence information and a thorough phenotypic characterization of more isolates are needed to confirm their species status.
In conclusion, for the allocation of new isolates to described species of moderate relationships, the use of nucleic acid probes directed against a gene product (16S rRNA) of
proven genetic stability is superior to classification based on traditional phenotypic characterization. For Xenorhabdus strains the phenotype-based taxonomy is not satisfying because the data available for the description of species are not consistent (3, 8, 9, 14, 15, 24, 25), causing problems when new isolates need to be classified. Moreover, application of probes is definitely faster than checking a broad spectrum of properties of diagnostic value (5), especially when a large number of isolates is included in a taxonomic survey. ACKNOWLEDGMENTS This study was supported by a joint grant from the Bundesministerium fur Forschung und Technologie, Bonn, and Badische Anilin und Sodafabrik (BASF), Ludwigshafen. We thank A. J. Peisker for excellent technical assistance. LITERATURE CITED 1. Akhurst, R. J. 1980. Morphological and functional dimorphism in Xenorhabdus spp., bacteria symbiontically associated with 2.
3. 4.
5.
6. 7.
8. 9.
insect pathogenic nematodes Neoaplectana and Heterorhabditis. J. Gen. Microbiol. 121:303-309. Akhurst, R. J. 1982. Antibiotic activity of Xenorhabdus spp., bacteria symbiotically associated with insert pathogenic nematodes of the families Heterorhabditidae and Steinernematidae. J. Gen. Microbiol. 128:3061-3066. Akhurst, R. J. 1983. Taxonomic study of Xenorhabdus, a genus symbiotically associated with insect pathogenic nematodes. Int. J. Syst. Bacteriol. 33:38-45. Akhurst, R. J. 1986. Xenorhabdus nematophilus subsp. poinarii: its interaction with insect pathogenic nematodes. Syst. Appl. Microbiol. 8:142-147. Akhurst, R. J., and N. E. Boemare. 1988. A numerical taxonomic study of the genus Xenorhabdus (Enterobacteriaceae) and proposed elevation of the subspecies of X. nematophilus to species. J. Gen. Microbiol. 134:1835-1845. Beaucage, S. L., and M. H. Caruthers. 1981. Deoxynucleoside phosphoramidites: a new class of key intermediates for deoxypolynucleotide synthesis. Tetrahedron Lett. 22:1859-1862. Bird, A. F., and R. J. Akhurst. 1983. The nature of the intestinal vesicle in nematodes of the family Steinernematidae. Int. J. Parasitol. 13:599-606. Boemare, N., and R. J. Akhurst. 1988. Biochemical and physiological characterization of colony form variants in Xenorhabdus spp. (Enterobacteriaceae). J. Gen. Microbiol. 134:751-761. Brenner, D. J. 1984. Enterobacteriaceae, p. 408-420. In N. R. Krieg and J. G. Holt (ed.), Bergey's manual of systematic
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bacteriology, vol. 1. The Williams & Wilkins Co., Baltimore. 10. Brosius, J., J. L. Palmer, J. P. Kennedy, and H. F. Noller. 1978. Complete nucleotide sequence of a 16S ribosomal RNA gene from Escherichia coli. Proc. Natl. Acad. Sci. USA 75:48014805. 11. Denhardt, D. T. 1966. A membrane filter technique for the detection of complementary DNA. Biochem. Biophys. Res. Commun. 23:641-646. 12. Ehlers, R., U. Wyss, and E. Stackebrandt. 1988. 16S rRNA cataloguing and phylogenetic position of the genus Xenorhabdus. Syst. Appl. Microbiol. 10:121-125. 13. Embley, M. T., J. Smida, and E. Stackebrandt. 1988. The phylogeny of mycolate-less wall chemotype IV actinomycetes and description of Pseudonocardiaceae fam. nov. Syst. Appl. Microbiol. 11:44-52. 14. Grimont, P. A. D., A. G. Steigerwalt, N. Boemare, F. W. Hickman-Brenner, C. Deval, F. Grimont, and D. J. Brenner. 1984. Deoxyribonucleic acid relatedness and phenotypic study of the genus Xenorhabdus. Int. J. Syst. Bacteriol. 34:378-388. 15. Hotchkin, P. G., and H. K. Kaya. 1984. Electrophoresis of soluble proteins from two species of Xenorhabdus, bacteria mutualistically associated with nematodes Steinernema spp. and Heterorhabditis spp. J. Gen. Microbiol. 130:2725-2731. 16. Kaya, H. K. 1985. Entomogenous nematodes for insect control in IPM systems, p. 283-302. In M. A. Hoy and D. C. Herzog (ed.), Biological control in agricultural IPM systems. Academic Press, Inc., Orlando, Fla. 17. Kondo, E., and N. Ishibashi. 1988. Histological and SEM observations on the invasion and succeeding growth of entomogenous nematode, Steinernema feltiae, (str. DD-136), in
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Spodoptera litura (Lepidoptera: Noctuidae) larvae. Appl. Entomol. Zool. 23:88-96. Krupp, G., and J. Gross. 1983. Sequence analysis of in vitro 32P-labeled RNA (II), p. 11-58. In P. F. Agris and R. A. Kopper (ed.), The modified nucleosides of transfer RNA (II). Alan R. Liss, Inc., New York. Lane, D. J., B. Pace, G. Olsen, D. A. Stahl, M. L. Sogin, and N. R. Pace. 1985. Rapid determination of 16S ribosomal RNA sequences for phylogenetic analyses. Proc. Natl. Acad. Sci. USA 82:6955-6959. Poinar, G. O., Jr. 1979. Nematodes for biological control of insects. CRC Press Inc., Boca Raton, Fla. Poinar, G. O., Jr., and P. T. Himsworth. 1967. Neoaplectana parasitism of larvae of the greater wax moth, Galleria mellonella. J. Invertebr. Pathol. 9:241-246. Poinar, G. O., Jr., and G. M. Thomas. 1966. Significance of Achromobacter nematophilus POINAR and THOMAS (Achromobacteriaceae: Eubacteriales) in the development of the nematode, DD-136 (Neoaplectana sp., Steinernematidae). Parasitology 56:385-390. Szostak, J. W., J. I. Stiles, B. K. Tye, P. Chin, F. Sherman, and R. Wu. 1979. Hybridisation with synthetic oligonucleotides. Methods Enzymol. 68:419-428. Thomas, G. M., and G. 0. Poinar, Jr. 1979. Xenorhabdus gen. nov., a genus of entomopathogenic nematophilic bacteria of the family Enterobacteriaceae. Int. J. Syst. Bacteriol. 29:352-360. Thomas, G. M., and G. 0. Poinar, Jr. 1983. Amended description of the genus Xenorhabdus Thomas & Poinar. Int. J. Syst. Bacteriol. 33:878-879.
