Heterorhabditis gerrardi n. sp

Journal of Helminthology (2009) 83, 309–320 q Cambridge University Press 2009

doi:10.1017/S0022149X09222942

Heterorhabditis gerrardi n. sp. (Nematoda: Heterorhabditidae): the hidden host of Photorhabdus asymbiotica (Enterobacteriaceae: g-Proteobacteria) K.L. Plichta1, S.A. Joyce2, D. Clarke2, N. Waterfield3 and S.P. Stock1* 1

Department of Entomology, University of Arizona, 1140 E. South Campus Dr., Tucson, AZ 85750, USA: 2Department of Microbiology, University College Cork, Ireland: 3University of Bath, Bath, United Kingdom (Accepted 7 December 2008; First Published Online 16 February 2009) Abstract

A new entomopathogenic nematode species from Australia, Heterorhabditis gerrardi n. sp. (Nematoda: Heterorhabditidae) is described. Morphological and molecular studies together with cross-hybridization tests indicated that this nematode represents a new undescribed species, closely related to members in the ‘indica-group’. However, the new species can be distinguished from other species in this genus by a combination of several qualitative and quantitative morphological traits. Key diagnostic features include: body size and excretory pore position of the third-stage infective juveniles; male bursa with a reduction of bursal rays, usually affecting the terminal set of papillae, with symmetrical or asymmetrical loss of one or two pairs; vulva of hermaphrodites more anteriorly located than in other species in the indica-group (V% average: 43), with non-protruding or slightly protruding lips, and longer tail length (average: 106 mm). The new species can be further characterized by molecular traits of sequence data from the internal transcribed spacer (ITS) region of ribosomal DNA. Additionally, the bacterial symbiont of this new species, Photorhabdus asymbiotica Kingscliff strain, was phenotypically characterized and compared with other P. asymbiotica strains. The Kingscliff strain revealed many characters not present in other strains of this species. We hypothesize that the newly found traits may contribute to the maintenance of this mutualistic association of the bacterium with its nematode host.

Introduction Entomopathogenic nematodes in the family Heterorhabditidae are obligate pathogens of a wide range of insect pests (revised by Shapiro-Ilan et al., 2002) and have been recovered from diverse climates and geographic regions of the world (revised by Adams et al., 2006). These nematodes are mutualistically associated with Photorhabdus bacteria (g-Proteobacteria: Enterobacteriaceae). The bacterial symbionts are harboured in the *E-mail: [email protected]

anterior portion of the intestine of the only free-living stage of these nematodes, also known as the infective juvenile or third-stage infective juvenile (IJ). The nematodes regurgitate the bacteria, which kill the insects and provide a food source for the nematodes to grow and multiply. At present more than a dozen species of Heterorhabditis nematodes have been described, and only three Photorhabdus species have been identified. Of them, P. asymbiotica is the only species known to be a human pathogen. Indeed, this species invades soft tissues and disseminates bacteraemic infections in human patients both in the United States and Australia (Akhurst & Smith, 2002; Gerrard et al., 2006). Originally, this bacterium was

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not found in association with nematodes, and because of this it was given the species name ‘asymbiotica’ ( ¼ not a symbiont). Gerrard et al. (2006) isolated a Heterorhabditis nematode from sandy soil samples taken at Kingscliff beach, New South Wales, Australia, a site where a patient with P. asymbiotica infection had been reported. Preliminary observations depicted this nematode as an undescribed Heterorhabditis species. In this study we provide a thorough morphological description of this new species, which is supplemented with internal transcribed spacer (ITS) rDNA sequence data and cross-hybridization assays. Evolutionary relationships of this nematode with other members of Heterorhabditis are presented and discussed. Moreover, the bacterial symbiont of this new Heterorhabidtis species was phenotypically characterized and compared with other existing P. asymbiotica strains.

Materials and methods Nematode isolation and rearing Soil samples (650 g) were collected from the property where the reported patient contracted the bacterial infection, in Kingscliff, Australia. Five mealworm larvae, Tenebrio mollitor L. (Coleoptera: Tenebrionidae) were placed in each container as baits according to procedures described by Kaya & Stock (1997). Dead insects were removed from the containers 5 days later and placed in modified White traps to recover the nematodes (Kaya & Stock, 1997). Third-stage infective juveniles (IJs) began to emerge 15 days after exposure of mealworm larvae to nematodes in the soil samples. Emerging IJs were surface sterilized in 0.4% Hyamine (Sigma, St. Louis, Missouri, USA) for 15 min and washed three times in sterile phosphate-buffered saline (PBS). Surface sterilized nematodes were used to infect Galleria mellonella larvae (100 IJs per insect) to confirm Koch’s postulates for pathogenicity (Kaya & Stock, 1997). Morphological characterization Differential interference contrast microscopy For morphological studies, first-generation hermaphrodites and second-generation adults were obtained by dissecting infested cadavers on days 4– 5 and 7 – 8 after initial infection. Third-stage infective juveniles were obtained upon emergence and during the first 2 days. A total of 20 specimens from each life stage were considered for evaluation of qualitative and quantitative data. Nematodes were examined live and/or heat-killed in 608C Ringer’s solution. Heat-killed nematodes were placed in triethanolamine – formalin (TAF) fixative (Kaya & Stock, 1997) and processed to anhydrous glycerine for mounting (Seinhorst, 1959). Measurements and digital light microscopy images were made from live and mounted specimens using an Olympus IX51 microscope equipped with differential interference contrast (DIC) optics and a digital video image system. Specimen measurements were made using AnalySIS Image software (Soft Imaging System Corp., California, USA). The following abbreviations for morphological features are used in the text and tables: ABW ¼ anal body width,

D% ¼ (EP/ES) £ 100, E% ¼ (EP/TL) £ 100, EP ¼ distance from anterior end to excretory pore, ES ¼ distance from anterior end to oesophagus base, GL ¼ gonad length, GS ¼ GuL/SpL, GuL ¼ gubernaculum length, MBW ¼ maximum body width, NR ¼ nerve ring position, StL ¼ stoma length, StW ¼ stoma width, SpL ¼ spicule length, SW ¼ SpL/GuL, TBL ¼ total body length, TL ¼ tail length, TRF ¼ testis reflexion, V% ¼ (TBL/length to vagina) £ 100, WA ¼ width at anus/cloaca. Scanning electron microscopy (SEM) Adult males and third-stage infective juveniles were rinsed three times in M9 buffer for 5 min per rinse. All nematodes were relaxed and heat-killed as above, then fixed in 8% glutaraldehyde buffered in cacodylate at pH 7.33 overnight. Fixed nematodes were rinsed in distilled water three times, post-fixed in OsO4 for 1 h, rinsed again in distilled water and serially dehydrated at 15-min intervals in ethanol (McClure & Stowel, 1978). Specimens were then critical point dried in liquid CO2, mounted on SEM stubs, coated twice with gold and scanned using a Philips XL series microscope at 20 kV accelerating voltage. Molecular characterization Nucleic acid preparations used for polymerase chain reaction (PCR) amplifications of the new Heterorhabditis sp. were extracted from pools of 10 – 50 third-stage infective juveniles. Nucleic acids were extracted from the digestion supernatant using phenol –chloroform enrichment, ethanol/ammonium acetate precipitation (Ausbel, 1989). The resulting pellet was washed with 70% ethanol, resuspended in Tris-EDTA (TE) buffer (pH 8.0), treated with 50 mg of RNAse A (1 h at 378C), and DNA recovered following reprecipitation with ethanol. DNA was quantified by spectrophotometry, and 100– 200 ng used per PCR reaction. PCR was used to amplify a region within the 50 -end of the nuclear internal transcribed spacer (ITS) region. Typical PCR reactions included 0.5 mM of each primer, 200 mM deoxynucleoside triphosphates, and MgCl2 concentration of 2 mM in a total reaction volume of 25 ml. PCR conditions, e.g. annealing temperature and MgCl2 concentration, were adjusted empirically as needed to optimize reaction specificity for individual isolates. Typical conditions included denaturation at 948C for 3 min, followed by 33 cycles of 948C for 30 s, 608C for 30 s, and 728C for 1.15 min, followed by a post-amplification extension at 728C for 5 min (Stock et al., 2001). PCR primers considered were 94, forward (50 – TTGAACCGGGTAAAAGTCG) and 93, reverse (50 – TTAGTTTCTTTTCCTCCGCT), as described by Stock et al. (2001). One microlitre of each PCR product was used for agarose gel electrophoresis (1.3% agarose in 1 £ Tris-borate-EDTA (TBE) buffer) to confirm amplicon size and yield. PCR products were prepared for direct sequencing using enzymatic treatment with EXOSAP-IT (USB Corp., Cleveland, Ohio, USA). Sequencing reactions were performed using dye-terminator cycle sequencing chemistry, and reaction products were separated and detected using an ABI 3730 automated

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DNA sequencer. Sequences for each species were completely double-stranded for accuracy, using the PCR primers and two internal sequencing primers. The forward internal primer was 264 (50 -GGTTTTTCATCGATACGCG), and the reverse internal primer 389 (50 -TGC AGACGCTTAGAGTGGTG). Sequence analysis and phylogenetic analysis Contig assembly and sequence conflict resolution was performed with the aid of EditSeq (DNAStar software, Lasergene Corp., Madison, Wisconsin, USA). Sequence regions corresponding to the PCR amplification primers were removed prior to multiple sequence alignment and phylogenetic analysis, because primer incorporation during amplification masks potential mismatches (substitutions) that may occur in PCR priming sites. Sequences were aligned initially using CLUSTAL X v.1.53b (Thompson et al., 1997), and the resulting output was adjusted manually to improve homology statements using MacClade software v.4.0 (Maddison & Maddison, 2002). ITS sequences of H. gerrardi n. sp. were deposited in GenBank under the accession number FJ1525445. Sequence data were analysed by unweighted maximum parsimony (MP) using PAUP* v. 4.0b3a (Swofford, 2002). Unrecoded gaps were treated as missing data. Tree searches for these ITS datasets were performed using heuristic methods with TBR (tree-bisection-reconnection) branch swapping, and a minimum of 1000 replicates of random stepwise addition. Reported consistency indices (CI) do not include uninformative characters. Bootstrap parsimony analyses were performed using heuristic searches (simple stepwise addition, TBR branchswapping, MULPARS) and 2000 pseudoreplicates.

Cross-hybridization tests Cross-breeding tests were carried out on lipid agar plates according to Kaya & Stock (1997). The Kingscliff isolate of the new Heterorhabditis species was crossed with two H. indica strains T1 (Thailand) and CR4 (Costa Rica). One female and one male of the appropriate strains were placed on the agar plates (35 mm diameter) and incubated at 258C. Controls consisted of one male and one female of the same isolate/species. Ten plates per isolate were tested. Progeny production was checked daily and for a period of three consecutive days according to Stock & Kaya (1996). Hybridization assays were repeated three times. Isolation and characterization of bacteria A motorized pestle was used to crush five individual IJs in 100 ml of Luria broth (LB). Thirty-five microlitres were plated on to LB and neutral bromothymol (NBT) agars and incubated at 308C for 48 h. Single colonies were replicated on to LB agar for incubation at 30 and 378C and allowed to grow for an additional 48 h and then examined for bioluminescence. Nematode-borne bacteria were analysed by MLST (multi-locus sequence typing) (Ed Feil, pers. comm.). Bacteria were also characterized phenotypically following procedures described by Joyce & Clarke (2003). Mutualistic exclusivity Members of all three characterized P. asymbiotica strains were assessed for their ability to support the growth and development of different nematode species. Bacteria, representative of all three known strains, were seeded

Table 1. Morphometric traits of adult stages and third-stage infective juveniles of H. gerrardi n. sp. Measurements – range and average (in parentheses) – are in micrometres. Characters TBL MBW EP NR ES V% SpL GuL SW% GS TL WA Ratio a Ratio b Ratio c D% E%

Hermaphrodite (n ¼ 20)

Female (n ¼ 20)

Male (n ¼ 20)

Third-stage infective juvenile (n ¼ 20)

2049–4288 (3073 ^ 646.2) 93.4–208.9 (136.2 ^ 2.3) 102.6–288.1 (189.8 ^ 43.7) 81.8– 210.4 (127.3 ^ 26.8) 145.5–316.8 (177.5 ^ 32.6) 40 –48 (43.5 ^ 2) – – – – 90.1–195.5 (106.5 ^ 20.5) 40 –80 (50 ^ 12) – – – 90 –147 (108 ^ 25.4) 100 –287 (181 ^ 46.7)

1428–2533 (1960 ^ 284) 71–161 (111.5 ^ 20.2) 107.5–157 (131.5 ^ 10.6) 73–141 (96 ^ 15) 120–182 (138 ^ 12) 43–55 (49 ^ 2) – – – – 66–95 (80.0 ^ 9) 22–38 (32 ^ 3.5) – – – 74–112 (95.5 ^ 9.5) 125–201 (165.5 ^ 21.5)

508– 916 (745 ^ 160) 33.8–48.4 (42.3 ^ 3.9) 92.5–140.8 (125 ^ 11.2) 53.8–87.4 (68.8 ^ 11.0) 78.3–114.8 (103.2 ^ 9.3) – 34.4–47.9 (42.7 ^ 3.6) 16.0–27.2 (21.8 ^ 3.0) 138– 274 (183 ^ 28) 40– 69 (50.5 ^ 7.5) 28.4–45.8 (37.6 ^ 4.6) 15.1–26.9 (23.6 ^ 2.4) – – – 99.7–171.7 (121.2 ^ 19) 270– 472 (335 ^ 49.1)

551– 683 (604 ^ 39) 17.5– 28.9 (23.4 ^ 3.4) 92.0– 110.8 (98.8 ^ 5.5) 81.2– 105.0 (93.9 ^ 18.0) 110.0–130.0 (123.8 ^ 4.6) – – – – – 76.1– 141 (101.9 ^ 14.1) 11.7–21.2 (14.5 ^ 2.9) 22.5– 32 (13.4 ^ 2.6) 0.16– 0.23 (0.21 ^ 0.02) 0.11–0.21 (0.17 ^ 0.03) 73–92 (80 ^ 0.5) 73–138 (99 ^ 1.7)

ABW, anal body width; D%, (EP/ES) £ 100; E%, (EP/TL) £ 100; EP, distance from anterior end to excretory pore; ES, distance from anterior end to oesophagus base; GS, GuL/SpL; GuL, gubernaculum length; MBW, maximum body width; NR, nerve ring position; ratio a, TBL/MBW; ratio b, TBL/ES; ratio c, TBL/TL; SpL, spicule length; SW%, (SpL/GuL) £ 100; TBL, total body length; TL, tail length; V%, (TBL/length to vagina) £ 100; WA, width at anus/cloaca.

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Fig. 1. Heterorhabditis gerrardi n. sp. line drawings. (A) hermaphrodite, anterior end, lateral view; (B) hermaphrodite vulval region, lateral view; (C) hermaphrodite tail region, lateral view; (D) female tail region, lateral view; (E) female vulval region, lateral view; (F, G) male tail ventral view showing numerical variation of bursal rays; (H) spicule, lateral view; (I) gubernaculum lateral view; (J, K) third-stage infective juvenile: (J) anterior end (lateral view), (K) tail region (lateral view). Scale bars: (A) 27 mm; (B) 43 mm; (C) 28.5 mm; (D, F, G) 23 mm; (E) 33 mm; (H) 16 mm; (I) 12.5 mm; (J) 30 mm; (K) 20 mm.

separately on to lipid agar plates in triplicate and incubated at 288C for 48 h. Surface sterilized Kingscliff infective juveniles (IJs) were added (day 1) and plates were examined and scored for nematode growth and development from day 5 to 18 using a stereomicroscope with a magnification of £ 50. When infective juveniles appeared on plates, they were collected, surface sterilized and crushed individually, as described above, to determine whether repackaging of that particular

bacterial species had occurred, thereby completing mutualism with the Kingscliff nematode.

Results and discussion Description of Heterorhabditis gerrardi n. sp.* The morphological traits of adult stages and third-stage infected juveniles of H. gerrardi are given in table 1.


313

Fig. 2. Heterorhabditis gerrardi n. sp. DIC and SEM micrographs of hermaphrodite and second-generation female. (A) Female, second generation, in toto; (B) hermaphrodite, anterior end, lateral view; (C) hermaphrodite, lateral view showing cardias (c) and anterior portion of intestine; (D) vulva opening of hermaphrodite, lateral view; (E) tail of hermaphrodite, lateral view, showing post-anal swelling (arrow); (F) vulva opening of second-generation female, lateral view; (G) tail of second-generation female, lateral view, showing post-anal swelling (arrow). Scale bars: (A) 186 mm; (B) 12 mm; (C, F) 14 mm; (D, E) 15 mm; (G) 32 mm.

Hermaphrodite C-shaped after transverse striae. (figs 1A and 2B). lips, each bearing

heat killed. Cuticle with fine Head region slightly truncated Labial region with six prominent one papilla. Cephalic papillae not

observed with SEM. Amphidial opening small, porelike. Stoma funnel-shaped, with refractile cheilorhabdions (figs 1A and 2B). Oesophagus with a cylindrical corpus. Isthmus present. Basal bulb pyriform or slightly rounded with small valve. Cardia present

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Fig. 3. Heterorhabditis gerrardi n. sp. DIC and SEM micrographs of male and third-stage infective juvenile. (A –C) Light microscopy images of tail of male, showing spicules and arrangement and number of bursal rays (A, lateral view; B,C, ventral views); (D, E) scanning electron micrographs of male tail showing bursal rays; (F) spicule, lateral view; (G) En face view of male head showing stomal opening, labial papillae (lp) and amphid (arrow), also Photorhabdus asymbiotica bacteria are shown inside the stoma; (H) third-stage infective juvenile showing dorsal cuticular tooth (arrow); (I) tail of third-stage infective juvenile showing anal opening (arrow). Scale bars: (A–C) 14 mm; (D) 30 mm; (E) 20 mm; (F) 13 mm; (G) 5 mm; (H, I) 19.5 mm.

(fig. 2C). Nerve ring located in the middle of the isthmus. Gonad didelphic, amphidelphic. Vulva located near the mid-body region (fig. 2A). Vulval lips slightly protruding or non-protruding and symmetrical (figs 1A and 2D). Copulatory plug not observed. Tail conoid with slightly protruding postanal swelling (figs 1C and 2E). Phasmids inconspicuous.

Male Body curved ventrally when heat killed. Cuticle with fine longitudinal striae. Head truncated or slightly rounded. Six labial papillae. Amphidial opening conspicuous. Mouth opening funnel-shaped, stoma short. Oesophagus with cylindrical procorpus and metacarpus. Isthmus distinct. Basal bulb pyriform, with reduced valve. Nerve ring surrounding isthmus or just

315


Table 2. Comparison of morphometric traits (range and mean) of infective juveniles of Heterorhabditis gerrardi n. sp. (in bold) with other Heterorhabditis spp. All measurements in micrometres. Third-stage infective juvenile

H. poinarii

1

H. taysaerae 2 H. indica

3

H. baujardi 4 H. floridensis

5

H. brevicaudis 6 H. mexicana

7

H. bacteriophora 8 H. amazonensis

9

H. georgiana 10 H. safricana 11 H. gerrardi n. sp. H. marelatus 12 H. zealandica 13 H. megidis 14 H. downesi

15

Male

TBL

MBW

EP

TL

RF

D%

E%

SpL

GuL

D%

NA 350– 410 418 332– 499 528 479– 573 551 497– 595 562 554– 609 572 528– 632 578 530– 620 588 512– 671 589 567– 612 598 547– 651 600 550– 676 604 551– 682 654 588– 700 685 570– 740 768 736– 800 637 588– 692

NA 18–22 19 17–23 20 19–22 20 18–22 21.2 19–23 22 20–24 23 20–24 23 18–31 23 20–24 27 23–34 28.6 24.8–31.8 23.4 17.5–29 28 24–32 27 22–30 29 27–32 35 29–48

NA

NA

NP

NA

NA

55 44 –70 101 93 –109 90 83 –97 63 48 –68 76 68 –80 99 91 –106 98 82–112 107 91–115 98 86 –108 93 86 –108 101.9 76 –141 107 99–117 102 87–119 119 112–128 68 62 –74

NP

82 71 –96 84 79 –90 84 76 –92 81 71 –90 84 78 –88 81 72 –86 84 79 –90 88 83 –92 82 70 –93 84 80 –90 90 NA 77 60 –86 80 73 –92 85 81 –91 85 76 –98

180 110–230 94 83–103 108 98 –114 105 95–134 147 NA 104 87–111 112 103–130 100 89–109 NA

NA 24 –32 18 14 –21 21 18 –23 20 18 –22 23 17 –30 22 20 –24 23 18 –32 20 18 –25 21 19 –23 25 20 –28 24 19 –27 22 16 –27 19 18 –22 22 19 –25 21 17 –24 18 17–19.5

NA

90 74 –113 98 88–107 97 91–103 109 101–122 111 104 –116 102 83–109 103 87 –110 107 85 –115 104 97 –113 110 103–122 98.8 91.9–110.8 102 81 –113 112 94–123 131 123–142 115 96–128

NA 43 –55 39 30 –42 43 35 –48 40 33 –45 42 36 –46 47 44 –48 41 30–-47 40 36 –44 41 35 –45 44 41 –49 45 35 –54 43 34 –48 45 42 –50 51 48 –55 49 46 –54 43 41 –47

NP NP NP NP NP NP NP NP NP NP NP NP NP P

119 99–133 99 73–138 96 86 –110 108 103–109 110 103–120 170 160–180

NA 122 NA NA 112 105 –119 88 NA 129 114–149 117 NA 103 95–109 110 100–122 117 92–133 121 99.7– 172 113 NA 118 NA 122 NA NA

After: 1Kakulia & Mikaia, 1997; 2Shamseldean et al., 1996; 3Poinar et al., 1992; 4Phan et al., 2003; 5Nguyen et al., 2006; 6Liu, 1994; 7Nguyen et al., 2004; 8Poinar, 1976; 9Andalo´ et al., 2007; 10Nguyen et al., 2008; 11Malan et al., 2008; 12Liu & Berry, 1996; 13Poinar, 1990; 14Poinar et al., 1987; 15Stock et al., 2002. D%, (EP/ES) £ 100; E%, (EP/TL) £ 100; EP, distance from anterior end to excretory pore; GuL, gubernaculum length; MBW, maximum body width; NA, not available; NP, not present; P, present; RF, tail refractile spine; SpL, spicule length; TBL, total body length; TL, tail length.

above the basal bulb. Excretory pore usually posterior to basal bulb. Testis single, reflexed anteriorly. Vas deferens well developed. Spicules paired, separate, with rostrum (figs 1H and 3A). Gubernaculum thin, straight or slightly curved ventrally. Tail conoid with peloderan bursa, slightly curved ventrally. Nine pairs of genital papillae (bursal rays) with the typical Heterorhabditis formula: 1 2 () 3 3 (fig. 3B – E). Pairs 4 and 7 curved outward (laterally) (fig. 2E). Pairs 7, 8 and 9 form a terminal group at the end of the bursa. Approximately 30% of the examined specimens (total 40 males) showed variable number of papillae in the two terminal sets (i.e. pairs 4 – 6 and 7 – 9). In some instances pair 6 was missing on one side of the bursa, whereas in other specimens pairs 8 and 9 or 9 were missing on both sides or only one side of the bursa (fig. 3B and C). Female Body C-shaped after heat-killed. Cuticle with fine transverse striae. Head region, lips and oesophageal region

similar to that of the hermaphrodites, but smaller. Gonads didelphic, amphidelphic. Vulva, a transverse slit, located in the mid-body region. Copulatory plug not observed. Vulval lips non-protruding. Vagina short. Tail conical with or without post-anal swelling (figs 1D and 2G). Infective juvenile Body long and slender, enclosed in the second-stage juvenile cuticle. Cuticle of exsheathed IJ with fine annulations. Lateral field with two longitudinal ridges. Anterior end rounded. Labial region with a conspicuous dorsal cuticular tooth (fig. 3H). Amphidial aperture inconspicuous. Mouth and anus are obliterated. Oesophagus narrow. Isthmus long and narrow. Basal bulb pyriform, with small valve (fig. 1J). Intestinal cardia visible and wide, harbouring symbiotic bacteria. Hemizonid, when observed, located 2 – 3 annules anterior to excretory pore. Nerve ring located at the isthmus level. Excretory pore located at the level of the basal bulb

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Fig. 4. Evidence of internal transcribed spacer region (ITS) of ribosomal DNA lineage independence for Heterorhabditis gerrardi n. sp. based on maximum parsimony analysis. Numbers in bold indicate bootstrap values. Numbers after species names correspond to GenBank accession numbers.

Cross-hybridization tests

(fig. 1J). Phasmids inconspicuous. Tail long and pointed terminus (fig. 1K and 3I).

Host Type host.

No progeny was produced when the Kingscliff isolate was crossed with the two H. indica isolates. Positive controls consisting of second-generation males and females of the Kingscliff isolate yielded fertile progeny.

Unknown.

Type locality. Australia.

Isolated from sandy soil in Kingscliff,

Diagnosis and relationships Morphological and molecular evidence suggest that the new Heterorhabditis species is related to species in the indica-group, which currently comprises six species, namely: H. indica Poinar, 1992, H. amazonensis Anadalo´ et al., 2007, H. baujardi Phan et al., 2003, H. floridensis Nguyen et al., 2006, H. mexicana Nguyen et al., 2004 and H. taysearae Shamseldean et al., 1996. However, H. gerrardi n. sp. can be differentiated from these species by a combination of morphological and/or morphometric

Type material. Holotype male deposited in the USDA Nematode Collection, Beltsville, Maryland, USA. Paratypes males (five specimens), females (five specimens), and thirdstage infective juveniles (five paratypes) were deposited at the UC Davis Nematode Collection. Etymology. This species is named after John Gerrard who first isolated and reported the bacterial symbiont of this nematode.

Table 3. Adjusted character distance matrix based on ITS rDNA sequence comparison between Heterorhabditis spp.

H. gerrardi n. sp. H. indica H. safricana H. marelatus H. downesi H. megidis H. gergiana H. zealandica H. bacteriophora HP88 H. bacteriophora NC1 H. amazonensis H. baujardi H. floridensis H. mexicana H. taysearae Pellioditis typica

1

2

3

4

5

6

7

8

9

10

11

12

13

14

15

16

– 8 105 104 103 114 120 119 56 65 51 49 57 59 59 133

– 105 104 103 114 118 117 57 67 49 51 55 57 57 131

– 13 25 33 103 102 39 49 109 107 111 113 111 125

– 24 30 100 99 36 46 108 106 110 112 112 127

– 18 102 101 38 48 107 105 109 111 109 128

– 112 111 43 55 113 112 114 116 114 130

– 1 14 23 116 114 120 125 123 119

– 13 22 115 113 119 124 122 119

– 21 53 51 56 60 58 107

– 63 51 66 70 68 126

– 1 6 9 11 134

– 6 11 12 117

– 5 7 136

– 6 137

– 137

–

þ þþ þ þþ þ þþ þ þþ þ CAS, chrome azurol S; LB, Luria broth; EMB, Eosin Methylene Blue (Oxoid); NBT, neutral bromothymol.

Red Green White þ þþ þ þþ White

þþ þ

þ þþ þ þþ þ þþ þ þþ þ Red Green White þ þþ þ þþ White

þþ þ

þ þþ þ þ þ þþ þ þþ þ þ þþ þ þþ þ 2 þþ þ þþ þ 2 þþ þ Red Pink Red Green Green Green Pink White White þ þþ 2 þ þþ þ þþ þ þþ þ þþ

P. luminescens (TT01 wild type) P. asymbiotica (USA ATCC) P. asymbiotica Kingscliff isolate (from insect cadaver) P. asymbiotica Kingscliff isolate (from human patient) P. asymbiotica Kingscliff isolate (from nematode host)

Orange White White

þþ þ 2 þþ þ

EMB Antibiotic Stilbene CAS Motility LB Isolate

Table 4. Phenotypic characterization of Photorhabdus asymbiotica isolates recovered from various sources.

NBT

McConkey

Protease

Lipase

Catalase

Bioluminescence


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traits of adult and third-stage juvenile stages (table 2). For example, third-stage infective juveniles of H. gerrardi n. sp. have a larger body size (average: 604 mm) than any of the species in the indica-group. Similarly the tail (average: 107 mm) of the third-stage infective juveniles is longer than in other species of the indica-group. Except for H. amazonenis, the excretory pore of the new species is more posteriorly located than in other species members of this clade. As in most Heterorhabditis spp., males of H. gerrardi n. sp. have a peloderan bursa usually supported by nine pairs of bursal rays which are arranged according to the formula: 1 2 3 () 3. However, a reduction of this formula, usually affecting the terminal set of three papillae, with symmetrical or asymmetrical loss of one or two pairs, is characteristic in the new species. Reduction of bursal rays has also been observed in other members of the indicagroup, including H. indica, H. amazonensis, H. floridensis, H. georgiana Nguyen et al., 2008 and H. mexicana, but usually affecting the terminal set of papillae and not pairs 4 – 6. Hermaphrodites of H. gerrardi n. sp. can be distinguished from other species in the indica-group by having the vulval opening more anteriorly located (V% average: 43), and with non-protruding or slightly protruding lips. Hermaphrodites of the new species can be separated from H. baujardi, H. floridensis, H. taysearae and H. indica, by having a longer tail (average: 106 mm). The new species can also be separated from other Heterorhabditis species by the size of the infective juveniles, which are much shorter than those of H. zealandica, H. downesi Stock et al., 2002 and H. megidis Poinar et al., 1987, H. gerrardi n. sp. can also be differentiated from H. bacteriophora Poinar, 1976 by the value of D% in the males (average: 121 versus 117) and from H. brevicaudis Liu, 1994 by having third-stage infective juveniles with a longer tail (average: 107 versus 76 mm). The new species can be further characterized by molecular traits of sequence data from ITS region of ribosomal DNA. Analysis of ITS rDNA sequences yielded three equally parsimonious trees with a tree length of 714 steps and a consistency index (CI), excluding uninformative characters, of 0.83. A total of 321 parsimony informative traits were depicted for this analysis. ITS analysis placed H. gerrardi n. sp. (GenBank accession no. FJ152545) as a member of the indica-clade and as sister taxon of H. indica. Bootstrap re-sampling provided strong support for this topology (100%) and the 50% majority rule consensus tree is given (fig. 4). Results from this analysis indicate H. gerradi n. sp. differs from H. indica by 8 base pairs out of 321 compared base pairs (table 3). These molecular observations provide further evidence for the distinctiveness of this species. Bionomics Heterorhabditis gerrardi n. sp. was found in sandy soils in Kingscliff beach, Australia. This new species behaves as other Heterorhabditis spp., its life cycle is completed after 15 days of initial infection of an insect host. Firstgeneration hermaphrodites are fully mature on days 4 – 5 after initial infection. Second-generation adults can be recovered on days 7 – 9 after initial infection. Cadavers infected by this new species do not have the typical brick-red/burgundy colouration. Instead

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Fig. 5. Phenotypic characteristics of Photorhabdus asymbiotica (Kingscliff strain) grown in different culture media. LB, Luria broth; CAS, chrome azurol S; Fizz, catalase test using H2O2; NBT, neutral bromothymol.

cadavers turn grey with blotchy pink spots. Once infection has progressed, cadavers remain with a greyish-pink colouration. Isolation and characterization of bacteria Nematodes were observed to contain a monoculture of non-pigmented bacteria. Cultures incubated at 308C were weakly bioluminescent while those incubated at 378C were strongly bioluminescent. MLST analysis confirmed the symbiotic bacteria as identical to bacteria isolated from the infected human wound and from the infected insects, and also identified as the Australian group of Photorhabdus asymbiotica (Gerrard et al., 2006).

Interestingly, phenotypic tests revealed differences between the human pathogenic strains of this Photorhabdus species. For example, the Kingscliff isolate of P. asymbiotica retained the ability to produce proteases and lipases and is highly bioluminescent when compared to P. asymbiotica USA/ATCC strain, which appears not to produce these factors (table 4, fig. 5). Joyce & Clarke (2003) previously proposed these factors as symbiosis factors that are regulated by the repressor HexA. More interestingly, the USA/ATCC strain produces no detectable siderophores and stilbene production is severely reduced. Both of these factors are required to support normal nematode growth and development (Gerrard et al., 2006; Joyce et al., 2008). Indeed P. asymbiotica USA/

Fig. 6. Recovery of Heterorhabditis gerrardi n. sp. infective juveniles (IJs) grown in lipid agar in the presence of representatives of other Photorhabdus species and/or strains.


ATCC did not support nematode growth and development of H. gerrardi n. sp., nor any nematode species when this bacterium was used in growth and development assays. Isolates of P. asymbiotica bacteria from all sources (i.e. nematode, human and insect associated) considered in this study were also tested for their ability to support nematode growth and development in vitro on lipid agar. All three bacterial isolates were capable of supporting recovery and development of the Kingscliff nematodes, and the bacteria were retained by the nematode. Furthermore three members of Photorhabdus luminescens: LN2 (origin H. indica), INAH23 (origin H. indica) and UK211 (origin H. megidis) supported growth and development of the Kingscliff nematode and yielded infective juveniles. However, when nematodes were surface sterilized and individually crushed, bacteria were not retained in any of these nematode species (fig. 6). In similar studies, it has been shown that other Photorhabdus species, such as P. temperata, support growth and development of Heterorhabditis species that naturally harbour P. luminescens; however, bacteria are not retained by non-natural nematode hosts (Joyce, pers. comm.). Furthermore Photorhabdus strains isolated from H. mexicana and H. taysearae (both members of the ‘indica-group’), and Photorhabdus luminescens HP88 and TT01 strains, were toxic to H. gerrardi n. sp., causing death of IJs.

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