Multigene Sequence-Based and Phenotypic Characterization Reveals the Occurrence of a Novel Entomopathogenic Nematode Species, Steinernema anantnagense n. sp.

Steinernema anantnagense n. sp. Steiner_6, Steiner_7, and Steiner_8 nematodes were isolated from soil samples collected in the Pir Panjal Range of Kashmir Valley, India using Corcyra cephalonica Stainton (Lepidoptera: Pyralidae) larvae as a bait insect. The isolates Steiner_6, Steiner_7, and Steiner_8 were collected in the Waghama area of Bijbehara Anantnag of the union territory of Jammu and Kashmir (GPS coordinates: 33.828914, 75.100091; 1606 m above the sea level) from soils around roots of willow, walnut, and apple intercrops, respectively, in areas adjoining district Anantnag, India. The insect cadavers recovered from soil samples were washed with ddH₂O, sterilized with 0.1% NaOCl₂, and nematode IJs were recovered from them by the White trap method (White, 1927). The IJs were sterilized with 0.1% NaOCl₂ and stored in 250 mL tissue culture flasks in Biological Oxygen Demand incubator at 15°C. The new species has been registered in the ZooBank at urn:lsid:zoobank.org:pub:210D5242-2C15-437F-8D57-B00EECD98B85.

Different life stages of S. anantnagense n. sp. were obtained from infected Galleria mellonella larvae exposed to 100 IJs/insects in a 15 cm-diameter Petri dish lined with moistened Whatman number 1 filter paper and kept in the dark at 25°C. The wax moth larvae died within 48 h after inoculation. After they died, the insect cadavers were transferred to a modified White trap (Kaya & Stock, 1997) and incubated at 25°C until IJs emerged. First- and second-generation adult nematodes were obtained by dissecting infected G. mellonella cadavers in Ringer’s solution after 3–4 and 6–7 days of infection, respectively. Infective juveniles (IJs) were collected after they emerged from G. mellonella cadavers in White traps (White, 1927). Nematodes were killed with water at 60°C, fixed in 4% formalin solution (4 mL formaldehyde, 1 mL Glycerol, 95 mL ddH₂O), dehydrated by the Seinhorst method (Seinhorst, 1959), and transferred to anhydrous glycerin. Nematodes were, after that, picked with a peacock feather and mounted on permanent glass slides with extra layers of paraffin wax to prevent the flattening of the nematodes as described (Bhat et al., 2022). Morphometric measurements were taken using the Nikon DS-L1 image acquisition software mounted on a phase-contrast microscope (Nikon Eclipse 50i) in μm. Light microscopy photographs were captured using a Nikon Eclipse 80i microscope (Olympus, Tokyo, Japan) equipped with differential interference contrast optics (DIC) and a Nikon Digital Sight DS-U1 camera. For the scanning electron microscopy (SEM), nematodes preserved in 4% formalin were re-hydrated in distilled water, dehydrated in a graded ethanol-acetone series, critical point dried with liquid CO₂, mounted on SEM stubs with a carbon tape, coated with gold in sputter coater, and observed with a Zeiss Merlin microscope (5 kV) (Zeiss, Oberkochen, Germany) (Abolafia, 2015). All micrographs were processed using Adobe® Photoshop® CS. Morphological characters of closely related species were taken from the original publications. The terminology used for the morphology of stoma and spicules follows the proposals by De Ley et al. (1995) and Abolafia and Peña-Santiago (2017a), respectively, and the terminology for pharynx follows the proposals by Bird and Bird (1991) and Baldwin and Perry (2004).

Self-crossing and cross-hybridization experiments were carried out using G. mellonella larvae hemolymph as described by Kaya & Stock, 1997 with minor modifications. To this end, drops of hemolymph obtained from surface-sterilized G. mellonella larvae were placed in sterile Petri dishes (35×10 mm). Hemolymph drops were treated with a small amount of phenylthiourea to prevent melanization. Then 40–60 surface-sterilized IJs (0.1% NaOCl for 30 min, followed by thrice rinse through sterile distilled water) were added to the hemolymph drops. Then, Petri dishes were wrapped in moistened tissue paper and kept in plastic bags at 25°C (room temperature). Petri dishes were observed daily for the presence of males and virgin females. Then, males and virgin females in the ratio of 3:3 were placed separately in fresh hemolymph drops and were crossed with adults of the opposite sex of the other species. Controls consist of crosses of identical isolates; some females were kept without males to check their virginity (n=30). The Petri dishes were observed daily for 15 days to determine the production of offspring. Experiments were conducted twice under the same conditions. The following species were crossed: Steinernema anantnagense n. sp. (Steiner_6, Steiner_7, and Steiner_8), S. ichnusae Sardinia, S. litorale Aichi, S. weiseri, S. akhursti Akh, S. citrae, S. cholashanense GARZE, S. feltiae P1, S. silvaticum, S. africanum RW14-M-C2a-3, and S. xueshanense DEQ.

Genomic DNA was extracted from single virgin females as described (Bhat et al., 2023). Briefly, several virgin females were first washed with Ringer’s solution and then with PBS buffer and then individually transferred into sterile PCR tubes (0.2 mL), each containing 20 μL extraction buffer (17.6 μL nuclease-free dH₂O, 2 μL 5X PCR buffer, 0.2 μL 1% Tween, and 0.2 μL proteinase K). The buffers with single virgin females were frozen at −20°C for 60 min or overnight and then immediately incubated in a water bath at 65°C for 1.2 h, followed by incubation at 95°C for 10 min. The lysates were cooled on ice and centrifuged at 6500 × g for 2 min. The following primers were used for PCR reactions: the internal transcribed spacer regions (ITS1-5.8S-ITS2) were amplified using primers 18S: (5′-TTGATTACGTCCCTGCCCTTT-3′) (forward), and 28S: (5′-TTTCACTCGCCGTTACTAAGG-3′) (reverse) (Vrain et al., 1992). The D2D3 regions of 28S rRNA were amplified using primers D2F: 5′-CCTTAG TAACGGCGAGTGAAA-3′ (forward) and 536: 5′-CAGC TATCCTGAGGAAAC-3′ (reverse) (Nadler et al., 2006). The 12S mitochondrial gene was amplified using the primers 505F: 5′-GTTCCAGAATAATCGGCTAGAC-3′ (forward) and 506R: 5′-TCTACTTTACTACAACTTACT CCCC-3′ (reverse) (Nadler et al., 2006) and the cytochrome oxidase subunit I (COI) gene was amplified using the universal primers LCO-1490 (5′-GGTCAACAAA TCATAAAGATATTGG-3′) (forward) and HCO-2198 (5′-TAAACTTCAGGGTGACCAAAAAATCA-3′) (reverse) (Folmer et al., 1994). The 25 µL PCR reactions consisted of 12.5 µL of Dream Taq Green PCR Master Mix (Thermo Scientific, USA), 0.5 µL of each forward and reverse primer at 10 µm, 2 µL of DNA extract and 9.5 µL of nuclease-free distilled water. The PCR reaction was performed using a thermocycler with the following settings: for ITS and D2-D3 markers, 1 cycle of 5 min at 94°C followed by 37 cycles of 30 sec at 94°C, 30 sec at 50°C, 1 min 30 s at 72°C, and by a single final elongation step at 72°C for 10 min. For the 12S marker, the PCR protocol included initial denaturation at 94°C for 3 min, followed by 30 cycles of 94°C for 30 s, 50°C for 30 s, and 72°C for 45 s, followed by a final extension at 72°C for 15 min. For the COI marker, the PCR program was as follows: one cycle of 94°C for 2 min followed by 37 cycles of 94°C for 30 s, 51°C for 45 s, 72°C for 2 min, and a final extension at 72°C for 12 min. PCR was followed by electrophoresis (40 min, 130 V) of 10 μL of PCR products in a 1% TBA (tris–boric acid–EDTA) buffered agarose gel stained with SYBR Safe DNA Gel Stain (Invitrogen, Carlsbad, California, USA) (Bhat et al., 2020b). PCR products were purified using QIAquick PCR Purification Kit (Qiagen, Valencia, CA) and sequenced using reverse and forward primers by Sanger sequencing (Bioserve Ltd., Hyderabad, India). Obtained sequences were manually edited and trimmed using BioEdit and deposited in the NCBI under the accession numbers: OQ40749, OQ407497, and OQ407497 for ITS; OQ407498, OQ407499, and OQ407500 for 28S; OQ404917, OQ407535, and OQ407536 for mtCOI; and OQ407491, OQ407492, and OQ407493 for mt12S.

To obtain genomic sequences of nematodes that belong to all the validly described species closely related to S. anantnagense n. sp., we searched the database of the National Center for Biotechnology Information (NCBI) using the Basic Local Alignment Search Tool (BLAST) (Altschul et al., 1990). Steinernema monticola (AB698756, GU395647, AY943994, and AY944020) was used as an outgroup in ITS, D2D3, mtCOI, and mt12S based phylogenetic trees. The resulting sequences were aligned with MUSCLE (v3.8.31) (Edgar, 2004) and used to reconstruct phylogenetic relationships by the Maximum Likelihood method based on the following nucleotide substitution models: Hasegawa-Kishino-Yano model (HKY+G) (ITS), Tamura–Nei (TN93+G+I) (D2–D3 & COI), and Tamura 3-parameter (T92+G) (12S). To select the best substitution models, best-fit nucleotide substitution model analyses were conducted in MEGA 11 (Nei & Kumar, 2000; Tamura et al., 2021). The trees with the highest log likelihood are shown. The percentages of trees where the associated taxa clustered together are displayed next to the branches. Initial tree(s) for the heuristic search were obtained automatically by applying Neighbor–Join and BioNJ algorithms to a matrix of pairwise distances estimated using the Maximum Composite Likelihood (MCL) approach, and then selecting the topology with superior log likelihood value. In some cases, a discrete Gamma distribution (+G) was used to model evolutionary rate differences among sites and the rate variation model allowed for some sites to be evolutionarily invariable (+I). The trees are drawn to scale, with branch lengths measured in the number of substitutions per site. Graphical representation and edition of the phylogenetic trees were performed with Interactive Tree of Life (v3.5.1) (Chevenet et al., 2006; Letunic and Bork, 2016).

The Xenorhabdus entomopathogenic bacteria associated with S. anantnagense n. sp. Steiner_6, Steiner_7, and Steiner_8 nematodes were isolated as described previously (Machado et al., 2018; Machado et al., 2019). Briefly, G. mellonella (n = 10) larvae were exposed to 100 nematode infective juveniles. Two to three days later, insect cadavers were surface–sterilized with 0.1% sodium hypochlorite solution and cut open with a sharp blade. Sterile polypropylene inoculation loops were inserted into the cadaver, and the loops were then streaked on LB agar plates and incubated at 28°C for 24–48 h. Xenorhabdus–like colonies were sub-cultured until monocultures were obtained. The strains were further sub-cultured and maintained on LB agar plates at 28°C. To establish their taxonomic identities, we reconstructed phylogenetic relationships based on whole genome sequences of the isolated bacteria and all the different species of the genus Xenorhabdus (Machado et al., 2023) and genomic sequences were obtained as described by Machado et al. (2021). Genome sequences were deposited in the National Centre for Biotechnology Information, and accession numbers are listed in Table S3. Phylogenetic relationships were reconstructed based on the assembled genomes and the genome sequences of all validly published species of the genus with publicly available genome sequences as described by Machado et al. (2023). Whole genome sequence similarities were calculated by the digital DNA-DNA hybridization (dDDH) method using the recommended formula 2 of the genome-to-genome distance calculator (GGDC) web service of the Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH (DSMZ) (Auch et al., 2010, 2020; Meier-Kolthoff et al., 2013, 2014).

Body 2.3–4.9 mm long, and C-shaped after heat relaxation and fixation. Cuticle with poorly visible annuli, with fine transversal incisures. Lateral fields absent. Labial region rounded, and continuous with the adjacent part of the body. Labial plate with six lips that are fused together, each with one labial papilla at the tip and one lower cephalic papilla each except for the lateral lips. Amphid openings present at the lateral lips, close to the labial papilla, with a small transversal slit. Stoma funnel-shaped, shallow, short, and wider at the anterior part. Cheilostom short with rounded and refringent rhabdia; gymnostom scarcely developed with a minute rhabdia; stegostom robust, slightly wider than long, with a funnel-shaped lumen and walls with very minute rhabdia. Pharynx muscular with a subcylindrical procorpus, a somewhat swollen metacorpus, a short and robust isthmus, and a spheroid basal bulb with reduced valves. Nerve ring surrounds the posterior part of the isthmus. Secretory-excretory pore circular, located at the anterior part of the isthmus. Deirids inconspicuous. Cardia short, conoid, and surrounded by intestinal tissue. Intestine tubular without differentiation, with thinner walls at the anterior end. Reproductive system didelphic, amphidelphic, and ovaries are reflexed in dorsal position. Oviducts well developed with glandular spermatheca, and uteri tubular with numerous uterine eggs. Vagina short with muscular walls, vulva protruding in the form of a transverse slit. Rectum 0.3–0.6 times the anal body diameter, with three small rectal glands. Anus well developed. Tail conoid, shorter than body anal diameter, with an acute terminus. Phasmids located at the posterior part of the tail, at 25–30% of the tail length.

Similar to first generation females, but smaller, measuring 1.8–2.4 mm in length. Tail conoid, with an acute terminus, longer than the first generation females.

Body 1.2–1.9 mm long, ventrally curved posteriorly, C- or J-shaped when heat killed. General morphology similar to that of females. Reproductive system monorchic, with the testis ventrally reflexed. Spicules paired, symmetrical, ventrally curved with a well-developed manubrium, either rounded or spoon-shaped. Calamus short and narrower, lamina ventrad curved at the anterior part and bears longitudinal ribs, ending in a blunt terminus. Velum indistinct, does not reach the spicule tip, and with no rostrum or retinaculum. Gubernaculum with a rounded manubrium, a fusiform corpus and a narrower and elongated tip, 0.5–0.7 times spicules length. Tail conoid with a rounded terminus bearing a fine mucron. Bursa absent. 11 pairs of genital papillae and a single mid-ventral papilla present, arranged as follows: five pairs (GP1-GP5) subventral precloacal, one pair (GP6) lateral precloacal, one single (MP) midventral precloacal, two pairs (GP7-GP8) sub-ventral ad-cloacal, one pair (GP9) subdorsal postcloacal and two pairs (GP10-GP11) postcloacal at terminus. Phasmids terminal, located laterally between the last pair of genital papillae.

Morphology of second generation males similar to that of the first generation males, but smaller, 0.8–1.2 mm in length. Tail with long, straight and robust mucron. Spicules curved ventrally, with a rhomboid shaped manubrium, slightly broader than the calamus, and a lamina curved ventrally at the anterior part. Ventral velum very reduced, and two longitudinal lateral ribs present. Gubernaculum with a slightly ventrad curved manubrium that is rounded and ventrad bent, slightly fusiform corpus and a narrower and slender terminus. Arrangement of genital papillae and phasmids similar to that of first generation males.

IJ body 0.7–0.8 mm long, almost straight or slightly curved body shape when heat-killed. Body tapers gradually at both extremes, cuticle with transverse incisures, well-developed annuli. Lateral fields begin as a single line close to the anterior end, and increase to eight ridges before gradually reducing to five and then two near the anus and phasmid levels, respectively. Lip region slightly narrower than the adjacent part of the body, six amalgamated lips; with smaller lateral lips, six reduced labial and four prominent cephalic papillae. Amphidial apertures pore-like, oral opening triangular with a noticeable margin. Stoma reduced and tubular with a small lumen, consisting of a short cheilostom and an elongated gymno-stegostom. Pharynx elongated and narrow, with a very long corpus; a slightly narrower isthmus, and a pyriform basal bulb with reduced valves. Nerve ring surrounds the isthmus, excretory pore located at the metacorpus level. Hemizonid present. Deirids inconspicuous. Cardia conoid. Intestine bears a bacterial sac at the anterior part. Rectum long, almost straight, with very short cuticular and elongated cellular parts, anus distinct. Genital primordium located at the equatorial region, tail conoid, tapering gradually to a pointed terminus, with a longer cellular part than the hyaline part and an irregular cellular-hyaline junction. Phasmids located at 37–45% of the tail length.

Steinernema anantnagense n. sp. is a highly pathogenic nematode species that can be easily reared on G. mellonella larvae at a temperature ranging from 18–24°C. The life cycle of this new species is similar to the life cycle of other Steinernema species. When G. mellonella larvae are exposed to 50–100 infective juveniles (IJs), they die within 24–48 h. The first- and second-generation adults of S. anantnagense n. sp. can be found in the insect cadavers 3–4 and 5–6 days after infection, respectively. The pre-infective juveniles leave the host body, mature for a few days, and then migrate to the water traps after 10–15 days.

The type hosts of Steinernema anantnagense n. sp. are unknown as the nematodes of this genus can infect different species of insects, and were obtained from soil samples using the insect baiting technique (Bedding and Akhurst 1975; White 1927). Steinernema anantnagense n. sp. Steiner_6, Steiner_7, and Steiner_8 nematodes were isolated, using the Corcyra cephalonica baiting method, from soil samples collected around the roots of willow, walnut, and apple trees in the Anantnag district of the Union Territory of Jammu and Kashmir, India (GPS coordinates: Lat. 33.828914°, Long. 75.100091°, 1606 m above sea level).

The type material for Steinernema anantnagense n. sp. are Steiner_7 nematode populations. For each stage (holotype and paratypes), including first-generation males and females, second-generation males and females, and infective juveniles, six permanent slides were prepared and deposited in the National Nematode Collection of India, located at the Indian Agricultural Research Institute (IARI) in New Delhi, India. Additionally, some permanent slides (paratypes) (n = 15) were deposited at the nematode collection of the Department of Animal Biology, Plant Biology and Ecology at the University of Jaén in Spain (IND001-01 – IND001-15). Live cultures of these nematodes are maintained at the Division of Entomology, Faculty of Agriculture, Sher-e-Kashmir University of Agricultural Science & Technology of Kashmir, India.

The species name is derived from the location Anantnag, a District in the Union Territory of Jammu and Kashmir, India, where the nematode specimens used in this study to describe the new species were obtained.

Mating experiments were carried out to determine the reproductive isolation of S. anantnagense n. sp. by pairing males and females of this species with individuals from other Steinernema species, including S. ichnusae, S. litorale, S. weiseri, S. akhursti, S. citrae, S. cholashanense, S. feltiae, S. silvaticum, S. africanum, and S. xueshanense. No offspring were produced when S. anantnagense n. sp. nematodes were allowed to interact with nematodes of the any of the above mentioned species, indicating that S. anantnagense n. sp. is reproductively isolated. Cross tests were also conducted between males and females of Steiner_6, Steiner_7, and Steiner_8 to determine their conspecific status. The results showed that fertile offspring were produced, confirming that they belong to the same species. Controls were also carried out, which included self-crossed species, and offspring were observed in all of them. However, no progeny were observed in the single-female control plates.

Steinernema anantnagense n. sp. is characterized by adults with a short stoma, rounded cheilorhabdia, and a robust pharynx with a round basal bulb. Females of the first generation are between 2.3–4.9 mm in length, with didelphic-amphidelphic reproductive system, and possess a shorter conoid tail bearing a short mucron in the first generation (c = 61–122, c′ = 0.5–0.8) and a longer conoid tail with thin mucron in the second generation (c = 43–67, c′ = 0.8–1.3). Males are smaller, between 1.2–1.9 mm in length, with a reproductive system that is monorchid and that has ventrally curved spicules bearing rounded manubrium in the first generation and rhomboid manubrium in the second generation, gubernaculum is fusiform in the first generation and anteriorly ventrad bent in the second generation, tail is conoid and slightly ventrally curved with a minute mucron in the first generation (c = 34–64, c′ = 0.8–1.6) and with a longer and more robust mucron in the second generation (c = 31–52, c′ = 0.6–1.4). The infective juveniles have a nearly straight body (0.7–0.8 mm length), poorly developed stoma and pharynx, lateral fields with eight longitudinal ridges and a conoid-elongate tail (49–66 µm, c = 12–16, c′ = 1.6–2.1) with a hyaline posterior part.

Steinernema anantnagense n. sp. belongs to a group of species known as the “Kushidai-clade”, and presents several traits common to this group. Several of the morphological and morphometric traits of the IJs and adults overlap with those of other species in the “Kushidai-clade”. However, several distinct morphological and morphometrical characteristics can differentiate S. anantnagense n. sp. from these closely related species (Tables 2–4).

Steinernema anantnagense n. sp. and S. akhursti (Qiu et al., 2005) morphologically differ in several traits. In the case of IJs, the distance from the head to the nerve ring is shorter in S. anantnagense n. sp. (54–71) compared to S. akhursti (83–95 μm), and the tail is shorter (49–66 μm) in S. anantnagense n. sp. than in S. akhursti (68–75 μm). The ratio of body length to tail (c) is greater in S. anantnagense n. sp. (12–16) than in S. akhursti (10–12), while the ratio of tail to body length (c′) is lower in S. anantnagense n. sp. (1.6–2.1) than in S. akhursti (3.3–3.7). Additionally, S. anantnagense n. sp. has a smaller H% value (20–36) compared to S. akhursti (49–56) (Table 2). The first-generation males of S. anantnagense n. sp. have a larger body diameter (167–211 μm) and much shorter spicule and gubernaculum (56–70 μm and 31–43 μm, respectively) compared to S. akhursti (body diameter: 115–150 μm, spicule: 85–100 μm, gubernaculum: 58–68 μm) (Table 3). The first-generation females of the two species also differ in some morphometric measurements (Table 4).

Steinernema anantnagense n. sp. differs from S. populi (Tain et al., 2022) in IJ body length (0.75–0.83 vs. 0.97–1.17 mm), the distance from anterior end to excretory pore (45–62 vs. 70–86 μm) and to nerve ring (54–71 vs. 98–113) μm), tail length (49–66 vs. 55–72 μm) and lower a, b, c and c′ ratios and lower D% (Table 2). The first-generation males of the new species differ from those of S. populi in body diameter (167–211 vs. 66–95 μm), tail length (29–39 vs. 39–68 μm), lower a and c ratios, and mucron (always present vs. present or absent) (Table 3). The first-generation females of the new species differ from those of S. populi in mucron (present vs. absent) and other characters (Table 4).

Steinernema anantnagense n. sp. can be distinguished from S. kushidai (Mamiya, 1988) by several morphological features. The body length of IJs in S. anantnagense n. sp. is longer (0.75–0.83 mm) than in S. kushidai (0.42–0.66 mm), and the distance from the anterior end to the nerve ring is shorter (54–71 μm) in S. anantnagense n. sp. compared to S. kushidai (70–84 μm). Additionally, the neck length is longer (120–143 μm), and the ratio c is higher (12–16) in S. anantnagense n. sp. than in S. kushidai (neck length: 106–120 μm, c: 10–13) (Table 2). The first-generation males of S. anantnagense n. sp. can be distinguished from S. kushidai by having a larger body diameter (167–211 vs. 75–156 μm), a shorter tail (29–39 vs. 40 μm), and the presence of a mucron, while S. kushidai does not have a mucron (Table 3). The first-generation females of S. anantnagense n. sp. also have a larger body diameter (314–409 μm), a longer distance from the anterior end to the nerve ring (143–182 μm), and the presence of a mucron, which are all different from S. kushidai (54–59 μm, 111–144 μm, and absent mucron, respectively) (Table 4).

In comparison to S. sangi (Phan et al., 2001), S. anantnagense n. sp. has a longer body length of IJs (0.75–0.83 vs. 0.70–0.78 mm), a shorter distance from the anterior end to the nerve ring (54–71 vs. 78–97 μm), a shorter tail length (49–66 vs. 76–89 μm), and greater c ratio, greater E%, and lower H% (12–16 vs. 9–10, 74–113 vs. 56–70, and 20–36 vs. 44–52, respectively) (Table 2). There are also minor differences in some characters between the first-generation males and females of the new species and those of S. sangi, which are presented in Tables 3 and 4, respectively.

The IJs of S. anantnagense n. sp. displays several distinguishing features from other related species. In comparison to S. cholashanense (Nguyen et al., 2008), the position of the nerve ring is more anterior (54–71 μm vs. 72–97 μm)), and the c′ ratio is lower (1.6–2.1 vs. 3.5–5.0). When compared to S. hebeiense (Chen et al., 2006), S. anantnagense n. sp. has a greater body length (0.75–0.83 vs. 0.61–0.71 mm), larger body diameter (32–42 vs. 23–48 μm), a more anterior position of the nerve ring (54–71 μm vs. 73–83 μm), a longer neck length (120–143 vs. 100–111 μm), a lower a ratio (19–24 vs. 24–28), and a higher c ratio (12–16 vs. 9.4–11). When compared to S. tielingense (Ma et al., 2012), S. anantnagense n. sp. has a shorter body length (0.75–0.83 vs. 0.82–0.98 mm), a more anterior position of the excretory pore and nerve ring (45–62 μm and 54–1 μm, respectively, as opposed to 64–73 μm and 90–105 μm, respectively), a shorter tail (49–66 vs. 74–85 μm), smaller ratios of a, b, and c′ and H% (19–24, 5.5–6.7, 1.6–2.1, and 20–36, respectively, as opposed to 27–31, 6.7–7.9, 3.5–4.6, and 53–64, respectively), but a longer c ratio (12–16 vs. 10–12). Compared to S. xinbinense (Ma et al., 2012), S. anantnagense n. sp. has a greater body length (0.75–0.84 vs. 0.64–0.74 mm), larger body diameter (32–42 μm vs. 28–31 μm), a smaller distance from the anterior end to the nerve ring (54–71 μm vs. 75–90 μm), a shorter tail (49–66 vs. 65–78 μm), a longer c ratio (12–16 vs. 8–11), a smaller c′ ratio (1.6–2.1 vs. 3–5), and a longer E% (74–113 vs. 65–78). Steinernema anantnagense n. sp. can be differentiated from S. xueshanense (Mrácek et al., 2009) by a smaller distance from the anterior end to the excretory pore and nerve ring (45–62 vs. 60–72 μm and 54–71 vs. 81–96 μm, respectively), a shorter tail length (49–66 vs. 80–92 μm), lower ratios of a and c′ (19–24 vs. 26–32 and 1.6–2.1 vs. 3.8–5.1, respectively), and lower H% (20–36 vs. 46–55). In addition, the position of the IJs nerve ring in the new species is more anterior (54–71 μm) compared to S. feltiae (Nguyen, 2007) (108–117 μm), and it also has a shorter tail length (49–66 vs. 81–89 μm) (Table 3).

The ITS regions of S. anantnagense n. sp. (Steiner_6, Steiner_7, and Steiner_8) are each 730 bp in length, consisting of ITS1 (278 bp), 5.8S (157 bp), and ITS2 (295 bp). Compared to other related species, the ITS region of S. anantnagense n. sp. shows differences of 19–117 bp, resulting in sequence similarity values of 78–97% (Table 5). Similarly, the D2-D3 expansion segments of the 28S rRNA gene of S. anantnagense n. sp. differ from those of other species by 5–35 bp, resulting in sequence similarity values of 95–99% (Table 6). In addition, the mitochondrial COI exhibit differences of 65–90 bp, resulting in 82–87% sequence similarity values, respectively (Table S1). Further, the mitochondrial 12S genes also exhibit 33–82 bp differences, resulting in sequence similarity values of 79–92%, respectively (Table S2). When these sequences were compared with sequences in the NCBI database using BLAST search, we observed that the top hit record for the ITS was 97.24% with S. akhursti (DQ375757) from China, for the D2D3 was 99.42% with S. akhursti (KF289902) from China, for mtCOI was 88.23% with S. sangi (MF621239) from India, and for mt12S rRNA was 92.34% with S. kushidai (AP017467) from Japan. Taken together, these observations suggest that S. anantnagense n. sp. represents a new taxonomic entity within the “Kushidai” clade, as evidenced by the lower sequence similarity scores between this species and all other known species, thus supporting its novel taxonomic status.

Phylogenetic reconstructions based on the nucleotide sequences of the internal transcribed spacer (ITS) marker of the rRNA gene, D2D3 expansion segments of the 28S rRNA gene, the cytochrome oxidase subunit I (COI), and the mitochondrial 12S rRNA gene show that S. anantnagense n. sp. Steiner_6, Steiner_7, and Steiner_8 are conspecific and belong to the “Kushidai” clade and the “Feltiae–Kushidai–Monticola” superclade (Figs. 6 and 7). Phylogenetic analyses of all four abovementioned markers clearly separate S. anantnagense n. sp. from all other species. In addition, these phylogenetic reconstructions show that S. anantnagense n. sp. is closely related to other Asian species, including S. akhursti, S. kushidai, and S. populi. No phylogenetic tree was built using the 18S rRNA genetic region because insufficient 18S rRNA gene sequences are publicly available. However, the resulting sequences were deposited in the NCBI databank under the following accession numbers: OQ407498 (Steiner_6), OQ407499 (Steiner_7), and OQ407500 (Steiner_8).

Figure 6:

Maximum-likelihood phylogenetic tree between the newly described Steinernema anantnagense n. sp. and other closely related species of Steinernema species based on nucleotide sequences of: (A) the Internal Transcribed Spacer (ITS1-5.8S-ITS2) rRNA, flanked by primers 18S and 26S, and (B) the D2-D3 expansion segments of the large subunit (28S) of rRNA flanked by primers D2F and 536. Numbers at nodes represent bootstrap values based on 100 replications. Bars represent average nucleotide substitutions per sequence position. NCBI accession numbers of the nucleotide sequences used for the analyses are shown next to the species names. The scale bar shows the number of substitutions per site.

Figure 7:

Maximum-likelihood phylogenetic tree between the newly described Steinernema anantnagense n. sp. and other closely related species of Steinernema species based on the nucleotide sequences of: (A) the COI region of the mitochondrial gene, flanked by primers LCO-1490 and HCO-2198, and (B) the mitochondrial 12S rRNA gene, flanked by primers 505F and 506R. Numbers at nodes represent bootstrap values based on 100 replications. Bars represent average nucleotide substitutions per sequence position. NCBI accession numbers of the nucleotide sequences used for the analyses are shown next to the species names (accession numbers in bold font are the sequences newly generated in this study). The scale bar shows the number of substitutions per site.

Phylogenetic reconstructions based on whole genome sequences show that the bacterial symbiont isolated from S. anantnagense n. sp. Steiner_7, named here XENO-2, is closely related to X. japonica DSM 16522^T and X. vietnamensis VN01^T (Fig. 8). The digital DNA–DNA hybridization (dDDH) values between XENO-2 and X. japonica DSM 16522^T, and between XENO-2 and X. vietnamensis VN01^T are 51.8% and 40.0%, respectively. These values are below the 70% divergence threshold for prokaryotic species delineation, indicating that XENO-2^T represents a novel species within the genus Xenorhabdus (Wayne et al., 1987). This species is formally described elsewhere.

Figure 8:

Phylogenetic reconstruction based on core genome sequences of Xenorhabdus bacterial strains. 1719910 nucleotide positions (1501 core genes) were used in the analysis. Numbers at the nodes represent SH-like branch supports. Bar represents 0.05 nucleotide substitutions per sequence position. Accession numbers of the genome sequences used for the reconstruction are shown in Table S3.

The term “monticolum” was introduced by Stock et al. (1997) to refer to the geographic origin of the nematodes studied, which were collected in Mount Jiri (Sancheong, Gyeongnam province, Korea). This term is a combination of “monti” referring to “mountain” and “colum” derived from the Latin suffix “cola” meaning “that lives in a place.” However, it should be noted that, as the suffix “cola” is a masculine noun in Latin, it does not have gender variations. Therefore, the correct term to use is “monticola.” The correct usage of this term has been discussed in detail by Nicolson, 1987. In light of this, we propose to refer to this species as Steinernema monticola, as was first used by Choo et al. (1998).