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Introduction
Entomopathogenic nematodes (EPN) of the family Steinernematidae Filipjev, 1934 (Rhabditida) are soil-dwelling organisms that form a mutualistic relationship with bacteria of the genus Xenorhabdus. Steinernematids have a worldwide distribution (Hominick Reference Hominick and Gaugler2002), and to date, more than 110 species have been described from all continents except Antarctica (Půža and Machado Reference Půža and Machado2024). This number is growing every year. Infective juveniles (IJs) are the only free-living stage of nematodes capable to locate the host and penetrate through its natural body openings. Once inside the host haemocoel, the bacteria are released by the nematode, whereupon they start to develop exponentially and kill the host and prevent invasion by secondary organisms by producing toxins and secondary metabolites (Stock Reference Stock2019). The nematodes feed on the lysed tissues of the host and bacterial colonies (Griffin et al. Reference Griffin, Boemare, Lewis, Grewal, Ehlers and Shapiro-Ilan2005). Nematodes complete 1–3 generations within the host cadaver, though the number of generations depends on the host’s size. As food resources are depleted, new IJs are produced which re-associate with the symbiotic bacteria and disperse in search of new hosts (Poinar Reference Poinar, Gaugler and Kaya1990; Půža Reference Půža and Lugtenberg2015).
In India, research on EPNs begun in the 1960s, mainly focusing on the evaluation of exotic EPN species on economically important insect pests; unfortunately, limited success was achieved due to the poor adaptability of exotic EPN species to local climatic conditions. Therefore, the focus on isolating indigenous EPNs has intensified, and this heightened emphasis has led to the successful isolation of nine native species of Steinernema nematodes (Patil et al. Reference Patil, Linga, Mhatre, Gowda, Rangasamy and Půža2023). More recently, several surveys have been conducted to document the diversity of EPNs in the country (Bhat et al. Reference Bhat, Machado, Abolafia, Askary, Půža, Ruiz-Cuenca, Ameen, Rana, Sayed and Al-Shuraym2023; Patil et al. Reference Patil, Linga, Mhatre, Gowda, Rangasamy and Půža2023). This has resulted in the description of three Steinernema species from India, including S. indicum Patil, Linga, Mhatre, Gowda, Rangasamy and Půža Reference Patil, Linga, Mhatre, Gowda, Rangasamy and Půža2023; S. anantnagense Bhat, Machado, Abolafia, Askary, Půža, Ruiz-Cuenca, Ameen, Rana, Sayed and Al-Shuraym Reference Bhat, Machado, Abolafia, Askary, Půža, Ruiz-Cuenca, Ameen, Rana, Sayed and Al-Shuraym2023; and S. shori Soni, Patil, Linga, Mhatre, Gowda, Ganguli and Půža Reference Soni, Patil, Linga, Mhatre, Gowda, Ganguli and Půža2023.
In the last few years, field sampling has been done in different ecosystems in Kerala, India, to determine the occurrence and distribution of EPN. From these samples, 19 strains of EPNs were successfully isolated from the rhizosphere of the coconut palm (Cocos nucifera), and one undescribed species of Steinernema strain S0804 was also identified (Anes et al. Reference Anes, Babu, Sivadasan and Josephrajkumar2018, Reference Anes, Babu, Sivadasan and Josephrajkumar2020). Morphological, morphometric, and molecular data prove that Steinernema strain S0804 isolated in the present study is a new species which is named herein as Steinernema keralense n. sp. This is the fourth species of the genus Steinernema described from India.
Materials and methods
Nematode isolation and rearing
Soil samples were collected from the rhizosphere of a coconut palm (Cocos nucifera) in ICAR-Central Plantation Crops Research Institute (CPCRI), Regional Station, research farm located in Kayamkulam (9°09′109.0864′′N, 76°30′0.48.0852′E with 3.05 m above the mean sea level) city of Alappuzha district in Kerala state, India. The soil type was sandy loam. Soil samples were taken from surface to a depth of 15 cm. Collected soil samples were placed into plastic bags for transport to the laboratory. In the laboratory, three last-instar Galleria mellonella (L.) larvae were placed in a 100 ml plastic container and then filled with moistened soil from each sample. Soil samples were checked daily for dead larvae, which were removed and placed in modified White trap (White Reference White1927). Nematodes that emerged from any individual cadaver were considered a separate isolate and used to infect live G. mellonella larvae to confirm Koch’s postulates (Kaya and Stock Reference Kaya, Stock and Lacey1997). For taxonomic studies, last-instar G. mellonella larvae were inoculated with infective juveniles (IJs) (200 IJs per G. mellonella) in a 9.0 cm-diameter Petri dish lined with a moistened filter paper and kept in the dark at 28 ± 2°C. First- and second-generation adult nematodes were obtained by dissecting EPNs infected dead G. mellonella larvae at 3 and 5 days, respectively, after the death of host. The cadaver dissection was carried out in Ringer’s solution. To obtain IJs, dead G. mellonella larvae were transferred to White trap and incubated at 25°C until IJs emerged. Infective juveniles emerged from the cadaver 7 days after the death of host.
Differential interference contrast microscopy
For light microscopy, the specimens of different stages were heat-killed, fixed in formaldehyde-glycerine fixative (Hooper Reference Hooper and Southey1970) for 24 h, and then transferred to glycerine-alcohol (5 parts glycerine: 95 parts 30% alcohol; Seinhorst Reference Seinhorst1959) for slow dehydration in a desiccator. Dehydrated specimens were mounted in anhydrous glycerine on glass slides using the wax ring method (De Maeseneer and D’Herde Reference De Maeseneer and D’ Herde1963). Morphometric analysis of the nematode specimens was done for 20 individuals of the adult stages of both generations and IJs, using Carl Zeiss Axio imager Z2 microscope fitted with DIC optics (Jena, Germany), a digital camera (Zeiss Axiocam 503 colour camera), and the image analysing software Zen 2 Blue edition.
Scanning electron microscopy (SEM)
Adults of both generations were obtained by dissecting G. mellonella larvae in Ringer’s solution (pH 7.3). They were rinsed three times for 3 min in Ringer’s solution. All the nematodes were heat-killed and then fixed in 4% formalin buffered with 0.1 M Phosphate buffer at pH 7.2 for 24 h at 4–6°C. They were post-fixed with a 2% osmium tetroxide solution for 12 h at 25°C, and then dehydrated at 15 min intervals through 20%, 30%, 50%, 70%, 90%, 95%, and 100% ethanol. Then, they were critical point-dried with liquid CO2, mounted on SEM stubs, and coated with gold (Nguyen and Smart Reference Nguyen and Smart1995, Reference Nguyen and Smart1997). The mounts were examined with a Carl Zeiss EVO-18 scanning electron microscope (Jena, Germany).
Molecular characterization
Total DNA was extracted from a single adult female nematode following the CTAB method in a CTAB buffer (CTAB 2%, 100 mM Tris pH 8.3, 20 mM EDTA, 1.4 M NaCl, and Mercaptoethanol 1%). The obtained DNA of the nematode was used as a template for the amplification of ITS region (including small subunit ribosomal RNA gene, partial sequence; internal transcribed spacer 1, 5.8S ribosomal RNA gene, and internal transcribed spacer 2, complete sequence; and large subunit ribosomal RNA gene, partial sequence) using 18S: 5’-TTGATTACGTCCCTGCCCTTT-3’ and 26S: 5’-TTTCACTCGCCGTTACTAAGG-3’ primers (Vrain et al. Reference Vrain, Wakarchuk, Levesque and Hamilton1992). Similarly, the D2–D3 expansion segments of the 28S rDNA gene were amplified using primers D2F: 5’-CCTTAGTAACGGCGAGTGAAA-3’ and 536: 5’-CAGCTATCCTGAGGAAAC-3’ (Nguyen Reference Nguyen, Nguyen and Hunt2007), and the cytochrome oxidase I (COI) gene was amplified using primers LCO 1490: 5′-GGTCAACAAATCATAAAGATATTGG-3′ and HCO 2198: 5′-TAAACTTCAGGGTGACCAAAAAATCA-3′ (Folmer et al. Reference Folmer, Black, Hoeh, Lutz and Vrijenhoek1994). All the PCR reactions were carried out separately in a thermocycler following separate cycling profiles such as for ITS region: 1 cycle of 94°C for 7 min followed by 35 cycles of 94°C for 60 s, 50°C for 60 s, 72°C for 60 s, and a last step of 72°C for 10 min.; for 28S rDNA: 1 cycle of 95°C for 5 min followed by 35 cycles of 94°C for 45 s, 54°C for 30 s, 72°C for 60 s, and a final extension at 72°C for 10 min; and for COI: 1 cycle of 94°C for 5 min followed by 35 cycles of 94°C for 60 s, 55°C for 60 s, 72°C for 90 s, and a final extension at 72°C for 10 min. The amplified PCR products were visualized in 1% agarose gel and documented under Gel Doc XR (Bio-Rad Laboratories India Pvt. Ltd.). Further PCR products were purified with a QIAquick PCR purification kit and sequenced in both directions by Bio Kart India Pvt. Ltd., Bengaluru (Karnataka, India). The obtained sequences were analyzed against NCBI BLASTn and deposited in GenBank with accession numbers MT139646 (ITS sequences), PQ249115 (28S sequence), and PQ248558 (COI sequence).
Entomopathogenic bacteria isolation and molecular characterization
The symbiotic bacteria associated with the nematode was isolated by following the procedure described by Akhurst (Reference Akhurst1980). The IJs of EPNs were surface sterilized using 0.1% Hyamine 10X. After 15–30 min, the IJs were washed three times with sterile distilled water to remove traces of Hyamine. The surface-sterilized IJs were crushed in a small 1.5 ml Eppendorf tube. A loop full of this crushed suspension was then streaked on the NBTA (nutrient agar supplemented with 0.004% (w/v) triphenyltetrazolium chloride and 0.0025% (w/v) bromothymol blue), and the plates were incubated at 28°C for 2–3 days. Single colonies were inoculated with a sterile toothpick on 10 ml of nutrient broth medium and incubated on an orbital shaker (180 rpm) at 28°C. Bacterial DNA was extracted from a one-day-old culture using the QIAamp DNA mini kit (Qiagen India Pvt. Ltd.) following the manufacturer’s instructions. The obtained DNA was used as a template for the amplification of 16S rRNA gene using the primers F27: 5´-AGAGTTTGATCCTGGCTCAG -3´ and R1492: 5´-TACGGYTACCTTGTTACGACTT-3´ (Heuer et al. Reference Heuer, Krsek, Baker, Smalla and Wellington1997). Similarly, the Recombinase A gene (recA) was amplified using primers RecA1F: 5′-GCTATTGATGAAAATAAACA-3′ and RecA2R: 5′-RATTTTRTCWCCRTTRTAGCT-3′ (Tailliez et al. Reference Tailliez, Laroui, Ginibre, Paule, Pagès and Boemare2010), and Gyrase B gene (gyrB) was amplified using primers 1200F gyrB: 5′-GATAACTCTTATAAAGTTTCCG-3′ and 1200R gyrB: 5′-CGGGTTGTATTCGTCACGGCC-3′ (Tailliez et al. Reference Tailliez, Laroui, Ginibre, Paule, Pagès and Boemare2010). All the PCR reactions were carried out separately in a thermocycler following separate cycling profiles, such as for 16S rRNA: 1 cycle of 94°C for 1 min followed by 33 cycles of 94°C for 60 s, 55°C for 60 s, 72°C for 2 min, and a final extension at 72°C for 3 min; recA: 1 cycle at 94°C for 2 min followed by 35 cycles at 94°C for 30 s, 49.5°C for 35 s, 72°C for 60 s, and a final extension at 72°C for 2 min and; gyrB: 1 cycle at 94°C for 2 min followed by 35 cycles at 94°C for 30 s, 56.5°C for 35 s, 72°C for 60 s, and a final extension at 72°C for 2 min. All the PCR primers were synthesized by Eurofins Pvt. Ltd., Bengaluru (Karnataka, India), and the PCR products were sequenced by Bio Kart India Pvt. Ltd., Bengaluru (Karnataka, India). The obtained sequences were analyzed against NCBI BLASTn, and subsequently, the sequences were deposited in GenBank with accession numbers PQ584826 (16S rRNA sequence), PQ594193 (recA sequence), and PQ594192 (gyrB sequence).
Phylogenetic analysis
The sequences were edited and compared with those present in GenBank using the Basic Local Alignment Search Tool (BLAST) of the National Centre for Biotechnology Information (NCBI). Alignments of our samples with sequences from species of the glaseri-group were created for each amplified DNA region using default ClustalW parameters in MEGA 7.0 (Kumar et al. Reference Kumar, Stecher and Tamura2016) and manually optimized in BioEdit (Hall 1999). Pairwise distances were calculated in MEGA 7.0 (Kumar et al. Reference Kumar, Stecher and Tamura2016).
Phylogenetic trees were constructed using the Minimum Evolution method (Rzhetsky and Nei Reference Rzhetsky and Nei1992) in MEGA 7.0 (Kumar et al. Reference Kumar, Stecher and Tamura2016) with Steinernema sacchari Nthenga, Knoetze, Berry, Tiedt and Malan Reference Nthenga, Knoetze, Berry, Tiedt and Malan2014 and Steinernema unicornum Edgington, Buddie, Tymo, France, Merino and Hunt Reference Edgington, Buddie, Tymo, France, Merino and Hunt2009 serving as outgroup taxa. The Minimum Evolution tree was obtained using the Close-Neighbour-Interchange (CNI) algorithm (Nei and Kumar Reference Nei and Kumar2000), and the neighbour-joining algorithm (Saitou and Nei Reference Saitou and Nei1987) was used to generate the initial tree. Evolutionary distances were computed using the p-distance method (Nei and Kumar Reference Nei and Kumar2000) and are expressed as the number of base differences per site.
Results
Description of Steinernema keralense n. sp. (Figures 1–3)
Steinernema keralense n. sp. Line drawings. A: First-generation male, tail with spicules and gubernaculum; B: Second-generation male, tail with spicules and gubernaculum; C, D: First-generation male with arrangement of genital papillae; E: Vulval region of first-generation female; F: First-generation female tail; G: Vulval region of second-generation female; H–F: Second-generation female tail; I: Anterior region of infective juvenile showing excretory pore and nerve ring; J: Infective juvenile tail showing anus and hyaline region; K: Infective juvenile lateral field showing ridges numbered from 1 to 7; L: First-generation male spicule. Scale bar: (A–B, E–J, L) 20 μm; (C, K) 10 μm; (D) 30 μm.
Steinernema keralense n. sp. Scanning electron microscopy photographs of infective juvenile, male and female. A: Infective juvenile showing cuticle with prominent transverse striations; B: Infective juvenile lateral field in mid-body (ridges numbered 1–7); C: First-generation female tail showing three projections on tip of the tail (arrow); D: First-generation female tail showing projections on tip of the tail (arrow) and anus (arrow); E: Vulval region of first-generation female; F: Tail region of infective juvenile with anus (arrow); G: First-generation male tail with genital papillae (numbered); H: First-generation male tail with a part of genital papillae (numbered) and single papilla (s).
Steinernema keralense n. sp. Light microscopy photographs of infective juvenile, male and female. A: Infective juvenile showing excretory pore (EP) and nerve ring (NR); B: Infective juvenile tail with anus (AN) and hyaline region (H); C, D: First-generation female showing tail variation; E: First-generation male tail with spicules and gubernaculum; F: Second-generation male tail with spicules, gubernaculum, and mucron (M); G: First-generation female showing asymmetrical vulval lips; H: Second-generation female with symmetrical vulval lips. I: First-generation male spicules.
Measurements
The dimensions of the holotype and paratype specimens are provided in Table 1.
Morphometrics of Steinernema keralense n. sp. All measurements are in μm and in the form: mean ± s.d. (range).
* Measuring along the chord
Description
Infective juvenile. Body of heat-relaxed specimens almost straight or gently curved and slightly narrowed at the anterior and posterior ends. Body 1,067 μm long on average (Table 1), ensheathed in the second-stage cuticle immediately after emergence, which is easily lost during storage. Cephalic region gently rounded, continuous with body shape, with four distinct cephalic papillae and a pair of pore-like amphidial apertures located laterally. Stoma closed. Cuticle marked with prominent transverse striations (Figure 2A). Deirids not observed. Hemizonid distinct ca 6 μm long, at level of anterior margin of basal bulb. Pharynx with a thin corpus of almost uniform diameter, slightly swollen metacorpus, an isthmus surrounded by the nerve ring and a distinct basal bulb (Figure 3A). Excretory pore located slightly posterior to midpharynx (D% = 58) (Table 1). Cardia present. Bacterial vesicle usually not well seen. Lateral field prominent, with seven ridges (8 incisures) in the mid-body region (Figure 2B). Lateral field beginning anteriorly with a cuticular depression and at excretory pore level two ridges become five. Near anus, number of ridges reducing to six. After anal region reduced to two ridges extended almost to tail tip (Figure 2F). Formula for lateral field = 2, 5, 7, 6, 2. Rectum long and narrow. Anus distinct (Figure 3B). Tail conoid with pointed terminus. Hyaline portion occupying 47% of tail length (Table 1 and Figure 3B).
First-generation male. Body curved posteriorly, mostly J-shaped when heat-relaxed. Cuticle smooth under light microscope, but with faint transverse striation visible under SEM (Figure 2H). Head round and continuous with body contour. Cephalic region with six labial, four cephalic papillae, and two amphidal apertures. Cephalic papillae are usually larger than labial papillae. Stoma shallow, broad, and usually cuticularized. Pharynx with cylindrical procorpus and slightly swollen metacorpus. Narrow isthmus surrounded by nerve ring located just anterior to basal bulb. Cardia prominent, intestine generally with a wide lumen. Excretory pore located posterior to midpharynx but anterior to nerve ring (ca 76% of distance from anterior body end to base of pharynx) (Table 1). Single reflexed testis, consisting of a germinal growth zone leading to a seminal vesicle. Spicules paired, curved, golden-brown in colour, ca 68 μm long, and spicule tip sharp (Figure 3I). Head of spicule usually elongate (head length/head width of 1.4:1). Shaft indistinct, lamina with two internal ribs, well curved. Blade with two internal ribs, well curved. Velum present. Gubernaculum arcuate, ca 80% of spicule length, and boat-shaped in lateral view. Tail short and rounded, mucron absent (Figure 3E). Twenty-five genital papillae comprising 12 pairs and a single midventral papilla located just anterior to cloacal aperture. Paired papillae arranged as follows: seven pairs subventral precloacal, one pair lateral precloacal, one pair adcloacal subventral, and three pairs postcloacal (one pair subdorsal and two pairs subventral terminal) (Figures G, H).
Second-generation male. Second-generation male similar to the first-generation male in morphology, but smaller in both body and organ sizes. Tail with a well-developed mucron (Figure 2F).
First-generation female. Body usually C-shaped, variable in length and usually coiled on heat relaxation. Head rounded, continuous with body. Stoma shallow, six labial papillae more prominent than four cephalic papillae, slit like amphidial apertures observed under SEM. Deirids not observed. Nerve ring surrounding isthmus, just anterior to basal bulb. Excretory pore located anterior to nerve ring at about mid-point of pharynx. Cardia prominent. Vagina short, leading into paired uteri. Genital system amphidelphic, reflexed ovaries, always containing many eggs. Vulva is asymmetrical, transverse slit without epiptygmata (Figure 3G), with vulval lips usually protruding ca 22 μm from the body surface. Rectum narrow, anal opening distinct. Postanal swelling not observed in most of the mature females. Lateral field and phasmids not observed. Tail conoid to dome-shaped (Figures 3C, D), bearing three minute projections (Figure 2C). Tail length much shorter than anal body diameter (Figure 3D).
Second-generation female. Similar to the first-generation in general morphology, but most of the morphometric measurements smaller. Excretory pore located more posteriorly than in the first-generation female. Tail conical, longer than the anal body diameter, with a pointed tip and no mucron. Tail length slightly longer than anal body diameter.
Taxonomic summary
Type material. Holotype first-generation male, paratype males (first-generation, 20 slides with 20 specimens; second-generation, 20 slides with 20 specimens), paratype females (first-generation, 20 slides with 20 specimens; second-generation, 20 slides with 20 specimens), and IJ (20 slides with 80 specimens) deposited at National Insect Museum, ICAR-National Bureau Agricultural insect resources (NBAIR), Bengaluru, Karnataka, India. Live culture of Steinernema keralense n. sp maintained at the division of germplasm collection and characterisation, ICAR-NBAIR, Karnataka, India and the division of crop protection, ICAR-CPCRI, Regional Station, Kayamkulam, India.
Type host. The natural host is unknown.
Type locality. Steinernema keralense n. sp. was recovered by baiting with G. mellonella larvae from soil samples collected from the rhizosphere of a coconut (Cocos nucifera) palm in ICAR-Central Plantation Crops Research Institute, Regional Station, research farm located in Kayamkulam city (9°09′109.0864′′N, 76°30′0.48.0852′E with 3.05 m above the mean sea level), Alappuzha district in Kerala state, India.
Etymology. * The specific epithet is named after Kerala state, where the type population was recovered.
Diagnosis and relationships
Steinernema keralense n. sp. is characterized by combination of the morphological and morphometric traits of infective juveniles and adults (Table 1). The IJ of the new species can be recognised by the body length of 1067 µm (914–1268 μm), maximum body diameter of 39 µm (30–43 μm), and tail length of 91 µm (76–102 μm). The excretory pore is located quite posteriorly (EP = 82 µm (73–92 μm)). Hyaline layer occupies approximately half of tail length, lateral fields with seven ridges (eight incisures) in mid-body region (Figure 2B). Male spicules curved, with a sharp tip, golden brown in colour, manubrium elongate with a length to width ratio of 1.4:1. The SW% ratio of the first-generation males is 136 (114–169). First-generation males lack a mucron on the tail tip (Figure 3E), while the second-generation males bear a 4 μm (2.3–6.2 μm) mucron (Figure 3F). The first-generation males are further characterised by very short spicules of 68 μm (60–72 μm) in length (Table 1). Genital papillae with 12 pairs and a single mid ventral papilla located just anterior to cloacal aperture (Figures 2G, H). The tail of the first-generation female is 24 μm (14–39 μm) in length and the anal body diameter is 61 μm (42–91 μm), while the tail of the second-generation female is 41 μm (30–47 μm) in length and the anal body diameter is 37 μm (31–44 μm) (Table 1). First-generation females possess asymmetrical vulval lips (Figure 3G), whereas the second-generation females possess symmetrical vulval lips (Figure 3H).
Steinernema keralense n. sp. can be distinguished from other Steinernema species by a combination of morphological and morphometric characteristics of males and infective juveniles. Phylogenetic analyses placed S. keralense n. sp. in the longicaudum-clade of glaseri-group within the Steinernematidae family. In the diagnosis, special emphasis will be given to the representatives of longicaudum clade that are phylogenetically closest relatives of S. keralense n. sp. – namely, S. adamsi Baniya, Subkrasae, Ardpairin, Anesko, Vitta and Dillman Reference Baniya, Subkrasae, Ardpairin, Anesko, Vitta and Dillman2024, S. guangdongense Qiu, Fang, Zhou, Pang and Nguyen Reference Qiu, Fang, Zhou, Pang and Nguyen2004; S. hermaphroditum Stock, Griffin and Chaerani Reference Stock, Griffin and Chaerani2004; S. lamjungense Khatri-Chhetri, Waeyenberge, Spiridonov, Manandhar and Moens Reference Khatri-Chhetri, Waeyenberge, Spiridonov, Manandhar and Moens2011; S. longicaudum Shen and Wang Reference Shen and Wang1992; S. pui Qiu, Zhao, Wu, Lv and Pang Reference Qiu, Zhao, Wu, Lv and Pang2011; and S. taiwanense Tseng, Hou and Tang Reference Tseng, Hou and Tang2018.
The IJs of S. keralense n. sp. can be distinguished from those of S. adamsi, S. hermaphroditum and S. lamjungense by their greater body length of 1067 μm (914–1268 μm) vs. 880 μm (775–957 μm), 929 μm (700–950 μm), and 832 μm (690–950 μm), respectively. The IJs of S. keralense n. sp. further differ from S. hermaphroditum and S. lamjungense by larger body diameter of 39 μm (30–43 μm) vs. 29 μm (25–32.5 μm), 27 μm (23–31 μm), respectively. The IJs of S. keralense n. sp. also differ from those of S. adamsi, S. hermaphroditum, and S. lamjungense in possessing longer distance from anterior end to excretory pore at 82 μm (73–92 μm) vs. 70 μm (64–75 μm), 65 μm (62.5–82.5 μm), and 68 μm (61–73 μm), respectively. The distance from anterior end to nerve ring in the IJs of S. keralense n. sp., at 105 μm (91–118 μm) is greater than that in S. adamsi (83 μm (75–92 μm)) and S. lamjungense (93 μm (82–100 μm)), and lower in comparison with S. pui (117 μm (100–125 μm)) and S. taiwanense (124 μm (120–127 μm)). The IJs of S. keralense n. sp. also differ from those of S. adamsi and S. lamjungense in possessing longer pharynx length of 141 μm (127–153 μm) vs. 103 μm (90–112 μm) and 127 μm (111–142 μm), respectively. With the length of 91 µm (76–102 μm), the tail of S. keralense n. sp. is much longer than those of S. adamsi (73 µm (62–82 μm)) and S. pui (69 µm (60–80 μm)). The b ratio of 7.6 (6.7–8.9) in S. keralense n. sp. is greater in comparison to 6.2 (5.6–7.1) in S. hermaphroditum and 6.6 (5.8–7.2) in S. lamjungense (Table 3).
Furthermore, the c ratio of S. keralense n. sp. of 12 (11–13) is greater than that of S. hermaphroditum (10.6 (9.3–11)) and S. lamjungense (9.6 (7.7–10.8)), but smaller than that of S. pui (15 (13–16)). The IJs of S. keralense n. sp. can also be distinguished from other closely related species of glaseri-group based on D% of IJs being 58 (54–63), which is lower than S. adamsi (68 (61–78)) and higher than S. hermaphroditum (50 (47–55)). The E% of S. keralense n. sp. is lower than S. pui (Table 3). The IJs of S. keralense n. sp. are distinct from all other members of the longicaudum clade by having a maximum of seven ridges in its lateral field (Figure 2B). In comparison, the species that have been described to date within this clade possess eight ridges in their lateral fields.
The first-generation males of S. keralense n. sp. can be distinguished from those of S. guangdongense, S. lamjungense, S. longicaudum, S. pui, and S. taiwanense by having shorter spicules being 68 μm (60–72 μm) long. The gubernaculum length of the first-generation males of S. keralense n. sp. is 55 μm (50–63 μm), which is higher than in S. hermaphroditum with 48 μm (47–50 μm), but lower than in S. longicaudum with 60 μm (54–65 μm), S. pui with 62 μm (58–65 μm), and S. taiwanense with 68 μm (65–70 μm). The first-generation males of S. keralense n. sp. can also be distinguished from those of S. adamsi, S. guangdongense, and S. taiwanense by having a larger body diameter of 138 μm (120–156 μm). The D% ratio of S. keralense n. sp. of 76 (62–89) is greater in comparison to S. hermaphroditum with 48 (44–50). Furthermore, the first-generation males of S. keralense n. sp. differ from those of S. guangdongense, S. hermaphroditum, S. lamjungense, S. longicaudum, S. pui, and S. taiwanense by having lower SW% ratio of 136 (114–169). In addition, the GS% ratio of S. keralense n. sp. is 82 (72–92), which is higher than in S. hermaphroditum, S. lamjungense, S. longicaudum, S. taiwanense, and S. pui (Table 2).
Comparative morphometrics of first-generation males of Steinernema keralense n. sp. and other species of the glaseri-group. All the measurements are in μm and in the form: mean (range). Abbreviations as defined in Table 1.
– Data not available.
Comparative morphometrics of infective juveniles of Steinernema keralense n. sp. and other species of the glaseri-group. All data are in μm and in the form: mean (range). Abbreviations as defined in Table 1.
– Data not available. ∗ Calculated from original description.
Molecular characterization and phylogenetic analysis
Analysis of sequences from the ITS region of the rDNA and the D2-D3 region of the 28S rDNA confirmed that S. keralense n. sp. is a distinct steinernematid species within the glaseri-group (sensu Spiridonov et al. Reference Spiridonov, Reid, Podrucka, Subbotin and Moens2004) (Figures 4 and 5). Phylogenetic reconstruction and sequence similarity analysis (Table 4) indicate that S. keralense n. sp. is closely related to species within the longicaudum clade (sensu Spiridonov and Subbotin Reference Spiridonov, Subbotin, Hunt and Nguyen2016). However, neither sequence provides a clear or well-supported resolution of phylogenetic relationships within the glaseri-group. The new species is further distinguished by the sequence of the mitochondrial cytochrome oxidase I (COI) gene, which shows a 9.5–12.9% divergence from other members of the glaseri-group with available COI sequences (Table 5).
Phylogenetic relationships of Steinernema keralense n. sp. and other members of the glaseri-group of Steinernema based on analysis of ITS rDNA regions. Steinernema sacchari and S. unicornum were used as outgroup taxa. The percentages of replicate trees in which the associated taxa clustered together in the bootstrap test (10,000 replicates) are shown next to the branches. Branch lengths indicate evolutionary distances and are expressed in the units of number of base differences per site.
Phylogenetic relationships of Steinernema keralense n. sp. and other members of the glaseri-group of Steinernema based on analysis of D2D3 expansion segments of the 28S rDNA. Steinernema sacchari and S. unicornum were used as outgroup taxa. The percentages of replicate trees in which the associated taxa clustered together in the bootstrap test (10,000 replicates) are shown next to the branches. Branch lengths indicate evolutionary distances and are expressed in the units of number of base differences per site.
Pairwise nucleotide similarities (below diagonal) and number of base differences (above diagonal) of ITS and D2D3 regions among species of the longicaudum clade of the glaseri-group and selected other glaseri-group species.
Pairwise nucleotide distances (%) of COI gene sequences among species within the glaseri-group.
Bacterial symbiont
Based on phylogenetic analysis (Figure 6), the symbiotic bacterium associated with S. keralense n. sp. (bacterial strain S0804) appears to represent a new Xenorhabdus species, closely related to X. griffiniae and X. ehlersii. The recA and gyrB gene sequence of Xenorhabdus sp. S0804 show approximately 96% and 96.3%, and 96.4% and 97% similarity, respectively, to those of X. griffiniae and X. ehlersii.
Phylogenetic relationships of Xenorhabdus sp. strain S0804 isolated from Steinernema keralense n. sp. and other closely related species of Xenorhabdus, based on the analysis of concatenated recA and gyrB gene sequences. Photorhabdus temperata was used as outgroup. The percentages of replicate trees in which the associated taxa clustered together in the bootstrap test (1,000 replicates) are given at each node. Branch lengths indicate evolutionary distances and are expressed in units of number of base differences per site.
This finding aligns with previously published data on nematodes from the longicaudum clade. S. hermaphroditum harbours X. griffiniae (Tailliez et al. Reference Tailliez, Pages, Ginibre and Boemare2006), the symbiont of S. adamsi is identical or closely related to X. griffiniae (Baniya et al. Reference Baniya, Subkrasae, Ardpairin, Anesko, Vitta and Dillman2024), and S. longicaudum harbours the closely related X. ehlersii (Lengyel et al. Reference Lengyel, Lang, Fodor, Szállás, Schumann and Stackebrandt2005). Based on current knowledge, nematodes from longicaudum clade associate exclusively with bacteria from this specific Xenorhabdus clade. However, the symbiont of S. guangdongense, S. lamjungense, S. pui, and S. taiwanense remain unidentified (Půža and Machado Reference Půža and Machado2024). However, the bacterium X. griffiniae has also been found to associate with the nematodes from the karii clade (Dreyer et al. Reference Dreyer, Malan and Dicks2018; Patil et al. Reference Patil, Linga, Mhatre, Gowda, Rangasamy and Půža2023; Půža et al. Reference Půža, Nermut, Mráček, Gengler and Haukeland2017).
Acknowledgements
The authors thank the directors, ICAR – Central Plantation Crops Research Institute, Kasaragod and ICAR-National Bureau of Agricultural Insect Resources, Bengaluru, for providing the research facilities, and the Central Instrumentation Facility (CIF), University of Agricultural Sciences, Bengaluru (UASB), GKVK, Bengaluru. We thank the National Higher Education Project (NAHEP) of the Indian Council of Agricultural Research, New Delhi, for supporting the CIF facility, and Dr. Nataraja Karaba, Head of the Central Instrumentation Facility, for expert assistance with the analysis.
Author contribution
K.M. Anes and J. Patil contributed equally to this work: conceptualization, methodology, software, validation, investigation, writing (original draft), review, editing, and visualization. K.M. Anes, A. Josephrajkumar, and J.V. Sajan: soil sample collection and formal analysis. M. Babu, B.S. Gotyal, and N. Aarthi: molecular characterization. J. Patil and V. Půža: molecular analysis, visualization, writing (review), and editing. P.H. Mhatre: nematodes drawing. M.T. Gowda: formal analysis. S.N. Sushil: visualization and investigation.
Financial support
This work was supported by the Indian Council of Agricultural Research (ICAR) New Delhi, ICAR-Central Plantation Crops Research Institute, Kasaragod, Kerala and ICAR-National Bureau of Agricultural Insect Resources, Bengaluru, Karnataka and Coconut Development Board, Kochi.
Competing interest
None.
Ethical standard
The authors assert that all procedures contributing to this work comply with the ethical standards of the relevant national and institutional guides on the care and use of laboratory animals.
