Taxonomic and molecular characterization of a new entomopathogenic nematode species, Heterorhabditis casmirica n. sp., and whole genome sequencing of its associated bacterial symbiont
Parasites & Vectors volume 16, Article number: 383 (2023)
Nematodes of the genus Heterorhabditis are important biocontrol agents as they form a lethal combination with their symbiotic Photorhabdus bacteria against agricultural insect pests. This study describes a new species of Heterorhabditis.
Six Heterorhabditis nematode populations were recovered from agricultural soils in Jammu and Kashmir, India. An initial examination using mitochondrial and nuclear genes showed that they belong to a new species. To describe this new species, a variety of analyses were conducted, including reconstructing phylogenetic relationships based on multiple genes, characterizing the nematodes at the morphological and morphometric levels, performing self-crossing and cross-hybridization experiments, and isolating and characterizing their symbiotic bacteria.
The newly discovered species, Heterorhabditis casmirica n. sp., shares 94% mitochondrial cytochrome C oxidase subunit I gene (COI) sequence identity with Heterorhabditis bacteriophora and Heterorhabditis ruandica, and 93% with Heterorhabditis zacatecana. Morphologically, it differs from H. bacteriophora in its infective juvenile phasmids (present vs. inconspicuous) and bacterial pouch visibility in the ventricular portion of the intestine (invisible vs. visible); genital papilla 1 (GP1) position (at manubrium level vs. more anterior), and in its b ratio (body length/neck length), c ratio (tail length/bulb width), and D% [(excretory pore/neck length) × 100]. Other morphological differences include anterior end to the nerve ring distance (77–100 vs. 121–130 μm), V% [(anterior end of vulva/body length) × 100] (46–57 vs. 41–47) in hermaphroditic females; rectum size (slightly longer than the anal body diameter vs. about three times longer), phasmids (smaller vs. inconspicuous), body length (0.13–2.0 vs. 0.32–0.39 mm), body diameter (73–150 vs. 160–220 μm), anterior end to the excretory pore distance (135–157 vs. 174–214 μm), and demanian ratios in amphimictic females. Morphological differences with H. ruandica and H. zacatecana were also observed. Furthermore, H. casmirica n. sp. did not mate or produce fertile progeny with other Heterorhabditis nematodes reported from India. It was also discovered that H. casmirica n. sp. is associated with 'Photorhabdus laumondii subsp. clarkei symbiotic bacteria.
The discovery of H. casmirica n. sp. provides novel insights into the diversity and evolution of Heterorhabditis nematodes and their symbiotic bacteria. This new species adds to the catalog of entomopathogenic nematodes in India.
Entomopathogenic nematodes belonging to the families Heterorhabditidae and Steinernematididae are highly effective biocontrol agents against agricultural pests. These nematodes have independently evolved mutual associations with insect pathogenic bacteria of the genera Photorhabdus and Xenorhabdus, respectively [1,2,3,4]. At the infective juvenile (IJ) stage, these nematodes, which reside in the soil, actively search for insect hosts . When an appropriate host is located, the IJs penetrate the insect body through natural openings or by directly breaking through the cuticle. They then release their bacterial symbionts upon sensing unknown chemical cues in the hemolymph [6, 7]. The bacteria multiply and produce virulence factors and toxins that kill the infected host [8,9,10]. Furthermore, the bacteria secrete exoenzymes that degrade the insect tissues and produce several metabolites essential for nematode growth, development, and reproduction [11, 12]. The bacteria also produce potent secondary metabolites that act as antibiotics and deter scavenging arthropods. Upon resource depletion, the new generation of nematodes disperses in search of new hosts [9, 13].
Heterorhabditis species are generally more virulent than those of Steinernema . However, they are less speciose than Steinernema [15, 16]. Despite this, new valid species of Heterorhabditis are often described and added to the list. The genus Heterorhabditis comprises 21 valid species, including two recently described species, Heterorhabditis ruandica from Rwanda and Heterorhabditis zacatecana from Mexico [15, 17]. Most of the valid species described so far have been molecularly characterized, except for Heterorhabditis egyptii  and Heterorhabditis hambletoni , which have only been morphologically characterized. The genus Heterorhabditis is globally distributed, although some species are only reported in certain geographic regions. In India, for instance, three species of Heterorhabditis have been documented so far: Heterorhabditis indica [20, 21], Heterorhabditis bacteriophora , and Heterorhabditis baujardi . Heterorhabditis indica, described by Poinar et al. , is the only new species of the genus Heterorhabditis reported from India to date.
In this study, we present the discovery of, and characterize, a new entomopathogenic nematode species, Heterorhabditis casmirica n. sp., and its symbiotic bacteria, recovered from the union territory of Jammu and Kashmir, India. Our study contributes to the characterization of soil biodiversity in general and advances our efforts to understand the biodiversity of an important group of biological control agents, which are essential tools for eco-friendly and sustainable agricultural practices.
Six populations of nematodes, namely HM, HM8, HP1, HPH, HH1, and HH4, were obtained from soil samples collected in the northwestern part of the union territory of Jammu and Kashmir, India. The samples were collected from soils around the roots of walnut and willow trees in the Anantnag district (Global Positioning System coordinates 33.828914, 75.100091; altitude 1606 m above sea level). Each one of these six populations was isolated from different soil samples. Each soil sample was separated by about 2 km from each other. Nematodes were isolated from soil samples using Corcyra cephalonica as a bait insect. Insects with nematode infestation symptoms were washed with double distilled H2O, sterilized with 0.1% NaOCl2, and then placed in White traps to recover the new generation of IJs . Recovered nematodes were reared using Galleria mellonella larvae as hosts under laboratory conditions [25, 26]. The IJs were stored in 250-mL tissue culture flasks in a biological oxygen demand incubator at 15 °C [27, 28]. The new species has been registered at ZooBank under urn:lsid:zoobank.org:pub:BBFC7CC6-7294-4548-AA7F-5CD5293E4103.
Nematode morphological and morphometric characterization, light and scanning electron microscopy
Hermaphroditic females, males and amphimictic females were obtained by dissecting G. mellonella cadavers in Ringer’s solution 4 and 6 days after infestation, respectively [26, 28]. The IJs were collected from White traps after emerging from the G. mellonella cadavers. The nematodes were then killed with hot water, fixed in TAF solution (2 mL triethanolamine, 7 mL of 40% commercial formaldehyde solution, and 91 mL distilled water), transferred to anhydrous glycerin, and mounted on permanent glass slides with additional layers of paraffin wax to prevent flattening during microscopy [29, 30]. Morphological measurements (in micrometers) were taken using Nikon DS-L2 image acquisition software on a phase-contrast microscope (Nikon Eclipse 80i). Twenty specimens at each developmental stage were measured. Light microscopy (LM) and scanning electron microscopy (SEM) photographs were obtained using various nematological techniques detailed by Abolafia . In brief, nematodes fixed in 4% formalin solution were processed to anhydrous glycerin using Siddiqi’s method with lactophenol-glycerin solutions . Subsequently, the nematodes were permanently mounted on glass microscope slides using the glycerin-paraffin method [33, 34]. The LM photographs were captured using a Nikon Eclipse 80i microscope (Olympus, Tokyo, Japan) with differential interference contrast optics and a Nikon Digital Sight DS-U1 camera. For SEM, nematodes preserved in glycerin were removed from permanent microscope slides by removing the cover glass, rehydrated in distilled water, dehydrated in a graded ethanol–acetone series, critically point dried with liquid CO2, mounted on SEM stubs with copper tape, coated with gold in a sputter coater, and finally observed with a Zeiss Merlin microscope (5 kV) (Zeiss, Oberkochen, Germany) . The LM and SEM micrographs, obtained at different magnifications for each structure, were processed and combined using Adobe Photoshop Creative Suite (Microsoft, Redmond, WA).
Comparisons were made between all the valid described species of Heterorhabditis based on morphological, morphometric and molecular characters, using the keys published by Machado et al. . Demanian indices and other ratios were calculated following the method outlined by de Man . The stoma morphology was described using the terminology provided by De Ley et al. , the spicule and gubernaculum morphology was described using the terminology established by Abolafia and Peña-Santiago  and the terminology for pharynx follows the proposals of Bird and Bird  and Baldwin and Perry .
Self-crossing and cross-hybridization experiments
Self-crossing and cross-hybridization experiments were carried out on lipid agar plates following the methodology described by Dix et al. . Heterorhabditis casmirica n. sp. isolates HM, HM8, HP1, HPH, HH1, and HH4 were crossed with each other and allowed to interact with Indian populations of H. bacteriophora (P4, P5 and KAS), H. indica (TH7, TH8 and TH9) and H. baujardi (HeTD4) nematodes. Control experiments were also conducted by self-crossing all the nematode species/strains. In each experiment, 20 second-generation males and 20 second-generation virgin females of each species were placed on 35-mm-diameter lipid agar plates and incubated at 25 °C. Progeny production was observed daily for 7 consecutive days. The experiments were conducted twice under the same conditions.
Nematode molecular characterization and phylogenetic relationships
Genomic DNA was extracted from individual hermaphroditic females isolated from insect cadavers infested with H. casmirica n. sp. HM, HM8, HP1, HPH, HH1, or HH4, as described . Briefly, individual virgin females were washed separately with Ringer’s solution and then washed in phosphate-buffered saline (pH 7.2). Virgin females were then individually transferred to sterile polymerase chain reaction (PCR) tubes (0.2 mL) containing 20 μL extraction buffer (17.6 μL nuclease-free distilled H2O, 2 μL of 5X PCR buffer, 0.2 μL 1% Tween, and 0.2 μL proteinase K). Samples were frozen at −20 °C for 60 min or overnight and then immediately incubated in a PCR thermocycler 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 3 min. The resulting supernatants were used as DNA templates to amplify different taxonomically relevant gene markers. A fragment of ribosomal rRNA (rRNA) containing the internal transcribed spacer (ITS) regions (ITS1-5.8S-ITS2) was amplified using primers 18S (5′-TTGATTACGTCCCTGCCCTTT-3′) (forward) and 28S (5′-TTTCACTCGCCGTTACTAAGG-3′) (reverse) . A fragment of rRNA containing the D2–D3 regions of the 28S rRNA was amplified using primers D2F (5′-CCTTAG TAACGGCGAGTGAAA-3′) (forward) and 536 (5′-CAGCTATCCTGAGGAAAC-3′) (reverse) . The 12S mitochondrial gene was amplified using primers 505F (5′-GTTCCAGAATAATCGGCTAGAC-3′) (forward) and 506R (5′-TCTACTTTACTACAACTTACT CCCC-3′) (reverse)  and the mitochondrially encoded cytochrome oxidase subunit I gene (MT-COI) was amplified using primers HCF (5′-TTACATGATACTTATTATG-3′) (forward) and HCF (5′-CTGATAACTGTGACCAAATACATA-3′) (reverse) . The PCR reactions consisted of 2 µL of DNA extract, 12.5 µL of DreamTaq Green PCR Master Mix (Thermo Scientific, USA), 0.75 µL of each forward and reverse primer at 10 µM and 9 µL of nuclease-free distilled H2O. The PCR reactions were performed using a thermocycler (Applied Biosystems Veriti 96-Well Thermal Cycler) with the following settings: (i) for ITS, D2–D3 and 12S—one cycle of 3 min at 94 °C followed by 35 cycles of 30 s at 94 °C, 30 s at 50 °C, 1 min 30 s at 72 °C, followed by a single final elongation step at 72 °C for 20 min; (ii) for the MT-COI gene—one cycle of 3 min at 94 °C followed by 38 cycles of 10 s at 94 °C, 30 s at 40 °C, 60 s at 72 °C, followed by a single final elongation step at 72 °C for 10 min . PCR was followed by electrophoresis (45 min, 100 V) of 5 μL of PCR products in a 1% Tris–boric acid–ethylenediaminetetraacetic acid-buffered agarose gel stained with SYBR Safe DNA Gel Stain (Invitrogen, Carlsbad, CA). PCR products were purified using the FastGene Gel/PCR extraction kit (Nippon Genetics, Japan) and sequenced using reverse and forward primers by Sanger sequencing (Bioserve, Hyderabad, India). The obtained sequences were manually curated, trimmed and deposited at the National Center for Biotechnology Information (NCBI) under the accession numbers given in Additional file 1: Table S4. To complete this data set and to obtain genomic sequences of nematodes that belong to all the valid described species of the genus Heterorhabditis, we searched the database of the NCBI by using the Basic Local Alignment Search Tool and the accession numbers of the sequences obtained previously [17, 47]. The resulting sequences were used to reconstruct phylogenetic relationships by the maximum likelihood method based on the following nucleotide substitution models: Tamura–Nei (TN93+G+I) (MT-COI) and Kimura 2-parameter (K2+G) (D2–D3) (ITS). To select the best substitution models, best-fit nucleotide substitution model analyses were carried out in MEGA 11 [48,49,50,51]. Sequences were aligned with MUSCLE (v3.8.31) . The trees with the highest log likelihood are shown. The percentage of trees in which the associated taxa clustered is shown next to the branches. Initial tree(s) for the heuristic search were obtained automatically by applying neighbor–joining and BIONJ algorithms to a matrix of pairwise distances estimated using the maximum composite likelihood approach, and selecting the topology with a superior log-likelihood value. In some cases, a discrete gamma distribution (+G) was used to model evolutionary rate differences between 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 [53, 54].
The Photorhabdus entomopathogenic bacteria associated with the different H. casmirica n. sp. nematode populations were isolated as described previously [55, 56]. Briefly, larvae of G. mellonella (Lepidoptera: Pyralidae) were exposed to 100 nematode IJs. Three to 4 days later, insect cadavers were surface sterilized and cut open with a surgical blade. Bacteria-digested internal organs were spread onto Luria—Bertani (LB) agar plates and incubated at 28 °C for 24–48 h. Photorhabdus—like colonies were then streaked on fresh LB agar plates until monocultures were obtained. A single primary form colony was then selected and used for further experiments. Bacteria primary forms were determined by examining colony morphology, colony texture, pigment production, and bioluminescence. The strains were further subcultured and maintained on LB agar plates at 28 °C. An initial molecular characterization, using 16S rRNA gene sequences, was carried out to determine the taxonomic affiliation of the obtained bacterial cultures as described previously [3, 4, 17, 56]. Phylogenetic reconstruction and sequence comparisons based on whole genome sequences were carried out to confirm the taxonomic affiliation of the obtained bacterial cultures as described previously [3, 55, 56]. Briefly, genomic DNA was extracted and purified using the GenElute Bacterial Genomic DNA Kit (Sigma-Aldrich, Switzerland) following the manufacturer’s instructions. The resulting DNA was used for library preparation using the TruSeq DNA PCR-Free LT Library Prep (FC-121-3003) kit. Indexed libraries were then pooled at equimolar concentrations and sequenced [2 × 150 base pairs (bp)] on an Illumina HiSeq 3000. Raw Illumina reads were quality trimmed using Trimmomatic 0.39 . The resulting reads were assembled with SPAdes 3.14.1 (k-mer sizes of 31, 51, 71, 91, and 111 bp) . Scaffolds with a mean read depth smaller than 20% of the median read depth of the longer scaffolds (≥ 5000 bp) as well as scaffolds that were shorter than 200 bp were removed. The final assemblies were polished using Pilon 1.22 . Phylogenetic relationships were reconstructed based on the assembled genomes and the genome sequences of all valid published species of the genus [3, 55, 56]. For this, core genome alignments were created using Roary 3.6.2 . Based on this alignment, a maximum likelihood tree was constructed using Fasttree 2.1.10 based on the Jukes–Cantor plus CAT nucleotide evolution model .
Results and discussion
Six populations of Heterorhabditis nematodes (HM, HM8, HP1, HPH, HH1, and HH4) were isolated from agricultural soils in Kashmir, India. Initial molecular and morphological characterization showed that they are genetically identical, morphologically very similar, and represent a novel species closely related to H. bacteriophora. The nematode population HM was chosen as the type material to describe this newly discovered species.
Heterorhabditis casmirica n. sp.
Hermaphroditic female body C-shaped when heat relaxed, body robust, always containing many juveniles, in some specimens a few eggs were visible. Cuticle almost smooth, about 0.8 to 1.6 µm thick. Lateral fields and phasmids not distinguishable under LM. Anterior end tapering anteriorly. Labial region with six prominent lips, each with a terminal conoid labial papilla. Cephalic papillae not observed with LM. Amphidial apertures pore-like. Stoma rhabditoid type, 1.1–1.7 times the lip region width, with a short cheilostom with a hardly visible refringent rounded cheilorhabdia, gymnostom with refringent bar-like rhabdia, well-developed, and funnel-shaped stegostom surrounded by the pharyngeal collar and bearing minute rhabdia. Pharynx with sub-cylindrical procorpus, slightly swollen metacorpus, robust isthmus, and poorly developed, spheroid basal bulb with inconspicuous valves. Nerve ring surrounding the isthmus, at 55–74% of neck length. Excretory pore at basal bulb level or intestine level, at 94–120% of neck length. Cardia conoid. Reproductive system didelphic–amphidelphic. Ovaries well developed, reflexed. Oviducts poorly differentiated. Uteri with numerous embryonated eggs. Vagina short. Vulva a transverse slit, with smooth top and scarcely prominent lips, close to mid-body. Rectum slender, about 0.9–1.4 times the anal body diameter. Anal region swelling posteriorly. Tail conoid with narrower pointed terminus, lacking a mucron. Phasmids inconspicuous.
Body arcuate with general morphology similar to that of hermaphroditic females. Body tapering toward anterior end; labial papillae acute and prominent. Reproductive system didelphic–amphidelphic with ovaries well developed, reflexed, oviducts and uteri poorly visible, vagina very short, and vulva small with a transverse slit opening. Rectum slightly longer than that of hermaphroditic females, about 1.7–1.9 times longer than the anal body diameter. Anal lips usually prominent. Tail conoid longer than that of hermaphroditic females, with acute tip lacking a mucron. Phasmids very small, located at 50–62% of tail length.
Body curved ventrally (open C-shape) or sometimes straight when heat relaxed. Anterior end truncate. Lip region with six scarcely separated lips and six conoid liplets at oral margin; six labial papillae at liplet tips and four cephalic papillae at the base of the dorsal and ventral lips. Amphidial aperture pore-like, just posterior to the lateral lips. Stoma 0.8–1.4 times the lip region width, with short cheilostom and hardly visible refringent rounded cheilorhabdia, short gymnostom with refringent bar-like rhabdia, and long, funnel-shaped stegostom surrounded by the pharyngeal collar and bearing minute rhabdia. Pharynx with subcylindrical procorpus, scarcely swollen metacorpus, isthmus robust and slightly narrower than metacorpus, and basal bulb poorly developed, spheroid, with poorly developed valvular apparatus. Nerve ring located surrounding isthmus, at 55–69% of neck length. Excretory pore located at basal bulb or intestine level, at 99–107% of neck length. Cardia conoid, protruding into intestine. Intestine without differentiation although with narrower walls at anterior end. Reproductive system monorchid, with testis anteriorly reflexed and vas deferens well developed. Spicules well developed, separate, with small, almost quadrangular manubrium with very refringent dorsal and ventral walls, frequently smaller at the left spicule, calamus developed, and almost straight lamina with acute tip, poorly developed dorsal hump, and ventral velum slightly developed. Gubernaculum robust, straight or slightly curved ventrally, 40–63% of spicule length, with manubrium visibly hook-like. Tail conoid with acute tip, ventrally curved posteriorly, flanked by the bursa. Bursa peloderan bearing nine pairs of bursal papillae 1 + 2/3 + 3: three precloacal and six postcloacal, with genital papilla 4 (GP4) and genital papilla 7 (GP7) open outside.
Infective sheathed juveniles (third-stage juvenile ensheathed in cuticle of second-stage juvenile)
Body straight when heat relaxed. Sheath (second-stage cuticle) present. Cuticle with longitudinal ridges except for the anterior part of the body, with annuli at the lip region and with tessellate pattern posterior to the lip region. Lip region lacking differentiated lips, bearing six labial papillae and cephalic papillae not visible. Amphidial aperture pore-like, having a cuticular dimple-like structure at its anterior part. Oral opening triradiate, closed. Stoma tubular, about twice as wide as the lip region. Pharynx slender, with corpus subcylindrical, isthmus narrower and slender, and basal bulb pyriform without developed valves. Nerve ring surrounding the isthmus, at 64–76% of neck length. Excretory pore at isthmus level, at 81–94% of neck length. Hemizonid clearly visible. Cardia conoid, surrounded by the intestinal tissue. Bacterial pouch not visible. Lateral fields not well differentiated from cuticle. Rectum narrow, not clearly discernible. Anus not well developed. Tail conoid-elongate with finely rounded terminus, without mucron. Terminal hyaline part 30–45% of tail length. Phasmids not visible.
Infective non-sheathed juveniles (third-stage juvenile)
Body with habitus straight when heat relaxed. Cuticle with transversal striae (annuli). Lateral field with two prominent longitudinal ridges. Lip region rounded, lacking differentiated lips, and labial and cephalic papillae not visible. Amphidial apertures oval. Oral opening rounded, closed, bearing a small dorsal tooth. Stoma, pharynx, nerve ring and excretory pore location similar to the sheathed stage. Hemizonid well developed. Cardia conoid, surrounded by intestinal tissue. Rectum narrow and hardly visible. Anus closed. Tail conoid with refringent acute tip without mucron. Phasmids very small, located at posterior part of tail.
Diagnosis of H. casmirica n. sp.
Heterorhabditis casmirica n. sp. is characterized by having females and males with six conoid oral liplets, pore-like amphids and a robust pharynx, pharynx slender in juveniles, nerve ring surrounding the isthmus and excretory pore at basal bulb or intestine level in adults and at isthmus level in juveniles. Hermaphroditic females 2.8–4.2 mm long, with conoid tail (72–114 µm long, c = 56–84, c′ = 1.4–2.2) with narrower tip; amphimictic females 1.2–2.0 mm long, with conoid tail (64–83 µm long, c = 16–31, c′ = 1.6–2.5); males 0.6–0.9 mm long, with ventrally curved tail (16–32 µm long, c = 24–32, c′ = 1.1–1.6), bursa with nine bursal papillae, spicules 38–48 µm long with manubrium with refractive walls, frequently smaller at the left spicule, gubernaculum 18–26 µm long with hook-like manubrium; juvenile with a tubular stoma and narrow and slender pharynx, second-stage juvenile (J2) 0.4–0.5 µm long, with cuticle with longitudinal ridges and conoid-elongate tail with finely rounded tip, and third-stage juvenile (J3) 0.5–0.6 µm long, with transversal annuli, a lateral field with two longitudinal ridges, oral opening with dorsal tooth and conoid tail with refringent acute tip.
Morphological relationships of H. casmirica n. sp. with other closely related species
Heterorhabditis casmirica n. sp. shares morphological similarities with Heterorhabditis bacteriophora , Heterorhabditis beicherriana , Heterorhabditis egyptii , Heterorhabditis georgiana , Heterorhabditis ruandica , and Heterorhabditis zacatecana . However, several distinct morphological and morphometric characteristics can be used to differentiate H. casmirica n. sp. from these closely related species (Tables 2, 3, 4, 5).
IJs of H. casmirica n. sp. can be differentiated from those of H. bacteriophora by differences in the c ratio (4.7–6.4 vs. 5.7–7.0), the presence of a bacterial sac (invisible vs. visible in the ventricular portion of the intestine), and size of phasmids (very small at the posterior part of the tail vs. inconspicuous). Compared to H. beicherriana IJs, those of H. casmirica n. sp. differ in the shape of amphidial apertures (oval vs. inconspicuous), the position of the excretory pore (at isthmus level vs. at the beginning of the basal bulb), visibility of the bacterial sac (invisible vs. visible), and the size of phasmids (very small at the posterior part of the tail vs. inconspicuous). When compared to H. egyptii, H. casmirica n. sp. differs in IJ tail length (85–115 vs. 53–75 µm), anterior end to excretory pore distance (98–129 vs. 81–94 µm), c ratio (4.7–6.4 vs. 6.8–9.1), and D% (83–97 vs. 74–82). When compared to H. georgiana, H. casmirica n. sp. IJs exhibit distinctions in visibility of the bacterial cell (invisible vs. visible posterior to cardia), with that in J2 and J3 occupying more than one half of the tail length (vs. about one half), and in phasmid size (very small at the posterior part of the tail vs. inconspicuous). When compared to H. ruandica, H. casmirica n. sp. IJs can be distinguished by the anterior end to nerve ring distance (79–94 vs. 52–64 µm), the position of the excretory pore (at isthmus level vs. at or just posterior to the basal bulb), tail length (85–115 vs. 49–64 µm), neck length (114–138 vs. 75–102 µm), c ratio (4.7–6.4 vs. 3.4–5.8), and presence of a cephalic tooth (small vs. large). When compared to H. zacatecana, the IJs of H. casmirica n. sp. differ in maximum body diameter (17–24 vs. 23–27 µm), the position of the excretory pore (at isthmus level vs. at or just posterior to the basal bulb), the anterior end to nerve ring distance (79–94 vs. 69–72 µm), neck length (114–138 vs. 78–99 µm), tail length (85–115 vs. 52–63 µm), c ratio (4.7–6.4 vs. 8.2–10), and c′ ratio (5.1–8.0 vs. 4.3–6.7). A detailed comparison of the morphology of the IJs of H. casmirica n. sp. with those of other Heterorhabditis species is given in Table 2.
The males of H. casmirica n. sp. can be distinguished from those of H. bacteriophora based on the neck length (106–118 vs. 99–105 µm), b ratio (6.4–8.2 vs. 9.1), c′ ratio (1.1–1.6 vs. 1.8), D% (99–107 vs 117 µm), spicules with a rectangular manubrium with strongly refringent walls (vs rectangular with scarcely refringent walls), gubernaculum more than a half of the spicule length (vs. shorter) and GP1 at the level of the manubrium (vs. more anterior in the type population). In comparison to male H. beicherriana, differences include body size (0.6–0.9 vs. 0.9–1.2 mm), maximum body diameter (24–48 vs. 51–73 μm), the distance from the anterior end to the excretory pore (102–120 vs. 130–157 μm), the distance from the anterior end to the nerve ring (58–80 vs. 81–108 μm), the tail length (16–32 vs. 32–45 μm), D% (99–107 vs. 102–120 µm), GP1 at spicule level (vs. more anterior), the shape of the spicule manubrium (quadrangular vs. oblongate) and gubernaculum (more than half of the spicule length vs. similar length). Compared to males of H. egyptii, differences lie in the c ratio (24–35 vs. 19.5). When compared to males of H. georgiana, differences lie in the position of the excretory pore (at bulb or intestine level vs. posterior to the basal bulb only), spicules with rectangular manubrium with strongly refringent walls (vs rectangular with scarcely refringent walls) and gubernaculum (more than a half of the spicule length vs. a half of the spicule length). Compared to males of H. ruandica, differences include the shape of the spicule manubrium (well developed, quadrangular and with strongly refringent walls vs. poorly developed, triangular and not refringent), the shape of the gubernaculum manubrium (hook-like vs. straight) and gubernaculum (more than a half of the spicule length vs. a half). Compared to males of H. zacatecana, differences include the shape of the spicule manubrium (quadrangular with strongly refringent walls vs. rounded and not refringent), bursa with GP1-GP2 distance shorter (less than the corresponding body diameter vs. slightly longer), GP2–GP3 slightly separated (vs. very closed), spicule manubrium (with angular anterior end vs. with rounded anterior end), the shape of the gubernaculum manubrium (hook-like vs. slightly curved) and gubernaculum more than a half of the spicule length (vs. shorter). Lastly, differences from males of H. hambletoni include the distance from the anterior end to the nerve ring (58–80 vs. 80–90 μm). With respect to the males of all of the other species, H. casmirica n. sp. has a different spicule morphology (manubrium with thick and refringent walls and lacking a dorsal hump vs. thin walls and a small dorsal hump) and gubernaculum with a hook-like manubrium (vs. straight).
The hermaphroditic females of H. casmirica n. sp. can be distinguished from those of H. bacteriophora based on several characteristics, including the distance from the anterior end to the nerve ring (77–100 vs. 121–130 μm), and a larger V% (46–57 vs. 41–47). The hermaphroditic females of the new species can be differentiated from those of H. beicherriana by the distance from the anterior end to the nerve ring (77–100 vs. 135–243 μm), and a smaller anal body diameter (36–56 vs. 51–92 µm). Additionally, hermaphroditic females of H. casmirica n. sp. differ from those of H. egyptii by the distance from the anterior end to the nerve ring (77–100 vs. 101–147 μm); from those of H. georgiana by the distance from the anterior end to the excretory pore (180–211 vs. 200–277 μm) and the distance from the anterior end to the nerve ring (77–100 vs. 143–217 μm); from those of H. ruandica in tail shape (conoid vs. conoid-elongate) and size (longer vs. short), visible uteri (vs. not well visible), the a ratio (19–27 vs. 12–16), and c ratio (56–84 vs. 34–51); from those of H. zacatecana by shorter length (0.28–0.42 vs. 0.44–0.62 mm), the distance from the anterior end to the nerve ring (77–100 vs. 96–169 μm), visible oviducts and uteri (vs. not well visible), and shorter neck length (154–176 vs. 174–231 µm).
Amphimictic females of H. casmirica n. sp. can be differentiated from those of H. bacteriophora by their shorter rectum (slightly longer than the anal body diameter vs. about three times longer in the type population), smaller phasmids (vs. inconspicuous), shorter length (0.13–2.0 vs. 0.32–0.39 mm), smaller maximum body diameter (73–150 vs. 160–220 μm), the distance from the anterior end to the excretory pore (135–157 vs. 174–214 μm), and demanian ratios. Compared to H. beicherriana, amphimictic females of H. casmirica n. sp. have a shorter tail (conoid vs. conoid-elongate), with an acute tip (vs. finely rounded tip), differ in their phasmids (very small vs. inconspicuous), and have a smaller anal body diameter (22–30 vs. 35–81 μm). In comparison to H. egyptii, amphimictic females of H. casmirica n. sp. have a shorter tail (conoid vs. conoid-elongate), longer distance from the anterior end to the excretory pore (135–157 vs. 69–106 μm), and longer neck length (132–156 vs. 106–125 μm). Additionally, amphimictic females of H. casmirica n. sp. differ from those of H. georgiana by having smaller phasmids (vs. inconspicuous), and from those of H. ruandica by having a longer neck (132–156 vs. 107–132 μm), different a ratio (14–15 vs. 15–20), and smaller phasmids (vs. inconspicuous). Finally, compared to amphimictic females of H. zacatecana, those of the new species have a smaller maximum body diameter (73–150 vs. 160–228 μm), different b ratio (10–13 vs. 16–21), c ratio (16–31 vs. 31–63), smaller phasmids (vs. inconspicuous), and smaller anal body diameter (22–30 vs. 31–41 μm). Summaries of the similarities and differences between males, hermaphroditic females, and amphimictic females of H. casmirica n. sp. and other Heterorhabditis species are presented in Tables 3, 4, 5, respectively.
Heterorhabditis casmirica n. sp. is a highly pathogenic nematode species that can be easily raised on G. mellonella larvae at a temperature ranging from 18 to 24 °C. The life cycle of this new species is comparable to that of other Heterorhabditis species. When G. mellonella larvae are exposed to 50–100 IJs, they die within 36–48 h and appear bright reddish after 48–72 h. First- and second-generation adults of H. casmirica n. sp. can be found in the insect cadavers 5–6 and 7–9 days after infection, respectively. The pre-infective juveniles left the host body, matured for a few days, and then migrated to the water traps after 15–21 days.
Type host and locality
The specific host for H. casmirica n. sp. is currently unknown as these nematodes were isolated from soil samples using the insect baiting technique [24, 77, 78]. Heterorhabditis casmirica n. sp. populations were collected from soil samples in the union territory of Jammu and Kashmir, located in the northwest region of India, and specifically in the Himalayan Pir Panjal region.
The type material for H. casmirica n. sp. (holotype male, 15 hermaphroditic female paratypes, 15 male paratypes, 15 amphimictic female paratypes and 19 J3, all belonging to the HM population) were deposited in the National Nematode Collection of India, Indian Agricultural Research Institute, New Delhi. Nematode cultures are maintained at the Sher-e-Kashmir University of Agricultural Sciences and Technology of Kashmir, India.
The specific name “casmirica” is derived from the Kashmir division (Casmiria in Latin), the geographical region where the nematodes used to describe the new species were collected.
No viable offspring were observed when H. casmirica n. sp. nematodes of the HM strain were allowed to interact with Indian populations of H. bacteriophora, H. indica, and H. baujardi. However, fertile progenies were observed when six different populations of H. casmirica n. sp. nematodes were allowed to interact, indicating that these populations are conspecific but reproductively isolated from closely related species, including H. bacteriophora, H. indica, and H. baujardi. Fertile progeny was also observed when all the nematode strains self-fertilized.
Nematode molecular characterization
The six populations of H. casmirica n. sp. were molecularly characterized based on the sequences of various genetic regions, including the ITS region of the rRNA (NCBI accession numbers OQ517936–OQ517941), the D2–D3 expansion segments of 28S rRNA (NCBI accession numbers OQ517947–OQ517952), mitochondrial 12S rRNA (NCBI accession numbers OQ517975-OQ517980), and MT-COI (NCBI accession numbers OQ517969–OQ517974). The ITS region of H. casmirica n. sp. is 771 bp in length, with ITS1 comprising 389 bp, 5.8S comprising 154 bp, and ITS2 comprising 228 bp. The MT-COI region flanked by primers HCF and HCR of H. casmirica n. sp. shows sequence similarity scores ranging from 75 to 94% with other Heterorhabditis species, and differs in 17–57 nucleotide positions (Table 6). Considering this genetic region, H. casmirica n. sp. is closely related to H. bacteriophora, H. ruandica, and H. zacatecana (Table 6). Heterorhabditis bacteriophora and H. ruandica both share 94% similarity with H. casmirica n. sp. and differ in 17 nucleotide positions. Heterorhabditis zacatecana shares 93% similarity with H. casmirica n. sp., and differs in 21 nucleotide positions. Fewer differences between H. casmirica n. sp. and its more closely related species were observed in the rRNA gene sequences. When compared with H. casmirica n. sp., H. bacteriophora and H. zacatecana both share 99.7% similarity and differ in two nucleotide positions, while H. ruandica shares 99.5% similarity and differs in four nucleotide positions in the ITS rRNA sequences flanked by primers TW81 and AB28 (Additional file 1: Table S2). All these three species share 100% similarity in the D2–D3 rRNA sequences flanked by primers D2A and D3B (Additional file 1: Table S3). Currently, very few mitochondrial 12S rRNA gene sequences are publicly available for molecular comparisons and phylogenetic analysis. Nevertheless, the sequences obtained in this study were deposited in the NCBI database for future taxonomic studies.
Nematode phylogenetic reconstructions
Phylogenetic analyses based on different genetic markers show that H. casmirica n. sp. belongs to the “bacteriophora” clade, which is currently composed of H. bacteriophora, H. beicherriana, H. georgiana, H. ruandica, and H. zacatecana (Figs. 7, 8, 9). MT-COI is particularly useful for the differentiation of all of these closely related species, and clearly shows that H. casmirica n. sp. and H. bacteriophora, its more closely related species, form two independent subclusters (Fig. 7). However, sequences of the ITS and D2–D3 regions of the rRNA gene, although allowing for the differentiation of certain species (Figs. 7, 8), provide lower phylogenetic resolving power than the MT-COI gene, as reported by Dhakal et al.  and Machado et al. . Hence, MT-COI is particularly useful for the molecular discrimination of closely related species of the genus Heterorhabditis.
Morphological and molecular relationships between H. casmirica n. sp. and specimens of H. bacteriophora present in India
At the morphological level, H. casmirica n. sp. differs from previously reported Indian isolates of H. bacteriophora [22, 30] (Additional file 1: Table S1). In particular, we observed that the males differ in spicule manubrium with strongly refringent walls (vs with scarcely refringent walls), gubernaculum more than a half of the spicule length (vs. shorter) and GP1 at manubrium level (vs. more anterior in the type population). The amphimictic females differ in smaller phasmids (vs. inconspicuous). The IJs differ in the distance from the anterior end to the nerve ring (79–94 vs. 48–74 µm), presence of bacterial sac (invisible vs. visible in the ventricular portion of the intestine), and size of phasmids (very small at posterior part of tail vs. inconspicuous) (Additional file 1: Table S1).
At the molecular level, H. casmirica n. sp. differs in 17 nucleotide positions in the MT-COI gene from several H. bacteriophora isolates from India, such as DH7, DH8, CH21, P5 and P6. On average, H. casmirica n. sp. shares 94% similarity with these isolates. In addition, the Indian populations of H. bacteriophora share 99.7% similarity with H. casmirica n. sp., and differ in two nucleotide positions in the ITS rRNA gene. Lastly, these two species do not differ in the sequences of the D2–D3 rRNA gene. Notably, the Indian populations DH7, DH8, CH21, P5 and P6 share 100% similarity with the type population of H. bacteriophora across all the gene markers used, and hence corroborate the conclusions of previous studies [22, 30]. The phylogenetic study further confirms the distinctiveness of the Indian populations of H. bacteriophora from H. casmirica n. sp. and establishes their similitude with the type population of H. bacteriophora (Figs. 7, 8, 9).
Phylogenetic reconstructions based on core genome sequences and sequence comparisons show that the bacterial symbionts isolated from H. casmirica n. sp. are very similar and belong to the subspecies Photorhabdus laumondii subsp. clarkei (Fig. 10). The digital DNA–DNA hybridization (dDDH) scores between BOJ47T, the type strain of the species P. laumondii subsp. clarkei, and strains HH4, HPH, and HP1, isolated from H. casmirica n. sp. HH4, HPH and HP1, are 94.3%, which is above the 70 and 79% thresholds that delimit prokaryotic species and subspecies, and confirms that they are conspecific .
A side note on the nomenclature of Heterorhabditis marelatus
The term “marelatus” was created by combining the Latin words “mare” meaning sea and “latus” meaning side in an attempt to translate the type locality “seaside” into Latin . Hence, marelatus was formed as a noun, not as an adjective. Sudhaus  changed the specific epithet of the species Heterorhabditis marelatus to “marelata.” This change was perhaps motivated by the fact that the genus noun Heterorhabditis is feminine and that, in Latin, the specific epithet should agree in gender with the genus. However, nouns in Latin do not vary according to gender, and therefore we propose that the correct term is “marelatus.” Hence, we propose that the original species nomenclature, Heterorhabditis marelatus, should be maintained.
Six populations of Heterorhabditis nematodes were identified in the present study that exhibited clear distinctions in their morphology, morphometric and molecular characteristics, as well as reproductive isolation and phylogenetic separation from all known Heterorhabditis species. We propose the name Heterorhabditis casmirica n. sp. for this new species, which is the second new Heterorhabditis entomopathogenic nematode species reported from the Indian subcontinent. Our results highlight the importance of using both classical taxonomy and molecular markers (MT-COI, ITS, small subunit, and large subunit) to accurately describe new Heterorhabditis species and their bacterial symbionts. The discovery of H. casmirica n. sp. and its associated bacterial symbiont expands our understanding of the biodiversity and distribution of these biocontrol agents and underscores their potential in the development of new biocontrol strategies against insect pests.
Availability of data and materials
Our sequences were deposited in the GenBank database under the accession numbers given in Additional file 1: Tables S3 and S4. Data supporting the conclusions of this article are included within the article. The datasets used and/or analyzed during the current study are available from the corresponding author upon reasonable request.
Internal transcribed spacer
- MT-COI :
Mitochondrially encoded cytochrome C oxidase subunit I
National Center for Biotechnology Information
Scanning electron microscopy
Clarke DJ. Photorhabdus: a tale of contrasting interactions. Microbiology. 2020;166:335–48.
Kajol Y, Bhat AH, Chaubey AK. Biochemical and molecular characterization of Photorhabdus akhurstii associated with Heterorhabditis indica from Meerut, India. Pak J Nematol. 2020;38:15–24.
Machado RAR, Bhat AH, Castaneda-Alvarez C, Půža V, San-Blas E. Photorhabddus aballayi sp. nov. and P. luminescens subsp. venezuelensis subsp. nov., isolated from Heterorhabditis amazonensis entomopathogenic nematodes. Int J Syst Evol Microbiol. 2023. https://doi.org/10.1099/ijsem.0.005872.
Bhat AH, Machado RAR, Abolafia J, Askary TH, Půža V, Ruiz-Cuenca AN, et al. Multigene sequence based and phenotypic characterization reveals the occurrence of a novel entomopathogenic nematode species, Steinernema anantnagense n. sp. J Nematol. 2023;55:e2023–31. https://doi.org/10.2478/jofnem-2023-0029.
Lefoulon E, McMullen JG, Stock SP. Transcriptomic analysis of Steinernema nematodes highlights metabolic costs associated to Xenorhabdus endosymbiont association and rearing conditions. Front Physiol. 2022;13:821845. https://doi.org/10.3389/fphys.2022.821845.
Ciche TA, Ensign JC. For the insect pathogen Photorhabdus luminescens, which end of a nematode is out? Appl Environ Microbiol. 2003;69:1890–7.
Hallem EA, Rengarajan M, Ciche TA, Sternberg PW. Nematodes, bacteria, and flies: a tripartite model for nematode parasitism. Curr Biol. 2007;17:898–904.
Bai X, Adams BJ, Ciche TA, Clifton S, Gaugler R, Kim K, et al. A lover and a fighter: the genome sequence of an entomopathogenic nematode Heterorhabditis bacteriophora. PLoS ONE. 2013;8:e69618. https://doi.org/10.1371/journal.pone.0069618.
RH Ffrench-Constant, Dowling A, Waterfield NR. Insecticidal toxins from Photorhabdus bacteria and their potential use in agriculture. Toxicon. 2007;49:436–51.
Rodou A, Ankrah DO, Stathopoulos C. Toxins and secretion systems of Photorhabdus luminescens. Toxins. 2020;2:1250–64.
Adams BJ, Fodor A, Koppenhofer HS, Stackebrandt E, Stock SP, Klein MG. Biodiversity and systematics of nematode-bacterium entomopathogens. Biol Control. 2006;37:32–49.
Mitani DK, Kaya HK, Goodrich-Blair H. Comparative study of the entomopathogenic nematode, Steinernema carpocapsae, reared on mutant and wild-type Xenorhabdus nematophila. Biol Control. 2004;29:382–91.
Ciche TA, Kim K, Kaufmann-Daszczuk B, Nguyen KCQ, Hall DH. Cell invasion and matricide during Photorhabdus luminescens transmission by Heterorhabditis bacteriophora nematodes. Appl Environ Microbiol. 2008;74:2275–87.
Ebssa L, Borgemeister C, Poehling HM. Effectiveness of different species/strains of entomopathogenic nematodes for control of western flower thrips (Frankliniella occidentalis) at various concentrations, host densities, and temperatures. Biol Control. 2004;29:145–54.
Bhat AH, Chaubey AK, Askary TH. Global distribution of entomopathogenic nematode, Steinernema and Heterorhabditis. Egypt J Biol Pest Control. 2020;30:31.
Hunt DJ, Subbotin SA. Taxonomy and systematics. In: Hunt DJ, Nguyen KB, editors. Advances in entomopathogenic nematode taxonomy and phylogeny. Nematol Monogr Perspect, vol. 12. Brill; 2016. p. 13–58.
Machado RAR, Bhat AH, Abolafia J, Muller A, Bruno P, Fallet P, et al. Multi-locus phylogenetic analyses uncover species boundaries and reveal the occurrence of two new entomopathogenic nematode species, Heterorhabditis ruandica n. sp. and Heterorhabditis zacatecana n. sp. J Nematol. 2021;53:e2021–89.
Abd-Elgawad MMM, Ameen HH. Heterorhabditis egyptii n. sp. (Rhabditida: Heterorhabditidae) from Egypt. Egypt J Agric Res. 2005;2:855–70.
Pereira C. Rhabditis hambletoni n. sp. nema apparentemente semiparasito da “broca do algodoeiro” (Gasterocercodes brasiliensis). Arch Inst Biol. 1937;8:215–30.
Poinar GO, Karunakar GK, David H. Heterorhabditis indicus n. sp. (Rhabditida: Nematoda) from India: separation of Heterorhabditis spp. by infective juveniles. Fund Appl Nematol. 1992;15:467–72.
Bhat AH, Chaubey AK, Shokoohi E, Machado RAR. Molecular and phenotypic characterization of Heterorhabditis indica (Nematoda: Rhabditida) nematodes isolated during a survey of agricultural soils in western Uttar Pradesh. India Acta Parasitol. 2021;66:236–52.
Bhat AH, Askary TH, Ahmad MJ, Suman, Aasha, Chaubey AK. Description of Heterorhabditis bacteriophora (Nematoda: Heterorhabditidae) isolated from hilly areas of Kashmir Valley. Egypt J Biol Pest Control. 2019;29:96.
Vanlalhlimpuia, Lalramliana, Lalramnghaki HC, Vanramliana. Morphological and molecular characterization of entomopathogenic nematode, Heterorhabditis baujardi (Rhabditida, Heterorhabditidae) from Mizoram, north eastern India. J Parasit Dis. 2018;42:341–9.
White GF. A method for obtaining infective nematode larvae from cultures. Science. 1927;66:302–3.
Leonar AL, Nimkingrat P, Aryal S, Martinez JG, Bhat AH, Sumaya NH. Natural association of the entomopathogenic nematode Heterorhabditis indica (Rhabditida: Heterorhabditidae) from the Philippines with the non-symbiotic bacterium Ochrobactrum anthropi (Proteobacteria: Brucellaceae). Egypt J Biol Pest Control. 2022;32:83.
Bhat AH, Istkhar, Chaubey AK, Půža V, San-Blas E. First report and comparative study of Steinernema surkhetense (Rhabditida: Steinernematidae) from subcontinental India. J Nematol. 2017;49:92–102.
Loulou A, Guerfali MM, Muller A, Bhat AH, Abolafia J, Machado RAR, et al. Potential of Oscheius tipulae nematodes as biological control agents against Ceratitis capitate. PLoS ONE. 2022;17:e0269106.
Bhat AH, Chaubey AK, Shokoohi E, Mashela PW. Study of Steinernema hermaphroditum (Nematoda, Rhabditida), from the West Uttar Pradesh. India Acta Parasitol. 2019;64:720–37.
Grisse AT. Rediscription ou modification de quelques techniques utilisees dans letude des nematodes phytoparasitaries. Meddelingen Rijksfauculteit landbouwweteschappen Bull. 1969;34:351–6.
Rana A, Bhat AH, Shokoohi E, Machado RAR. Morphological and molecular characterization Heterorhabditis bacteriophora nematodes isolated from Indian agricultural soils and their biocontrol potential. Zootaxa. 2020;4878:77–102.
Abolafia J. Extracción y procesado de nematodos de muestras de suelos de cuevas yotros hábitats. Monogr Biopg. 2022;16:6–17.
Siddiqi MR. Studies on Discolaimus spp. (Nematoda: Dorylaimidae) from India. J Zool Syst Evol Res. 1964;2:174–84.
Bharti L, Bhat AH, Chaubey AK, Abolafia J. Morphological and molecular characterisation of Merlinius brevidens (Allen, 1955) Siddiqi, 1970 (Nematoda: Rhabditida: Merlinidae) from India. J Nat Hist. 2020;54:1477–98.
Bhat AH, Chaubey AK, Hartmann J, Půža V. Notes on the morphology, bionomics, distribution and efficacy of Steinernema siamkayai (Rhabditida: Steinernematidae) from Western Uttar Pradesh. India Nematol. 2021;23:817–36.
Abolafia J. A low cost technique to manufacture a container to process meiofauna for scanning electron microscopy. Microsc Res Tech. 2015;78:771–6.
de Man JG. Die einheimischen, frei in der reinen Erde und im süssen Wasser lebende Nematoden. Tijdschrift Nederlandsche Dierkundige Vereeniging. 1880;5:1–104.
De Ley P, van de Velde MC, Mounport D, Baujard P, Coomans A. Ultrastructure of the stoma in Cephalobidae, Panagrolaimidae and Rhabditidae, with a proposal for a revised stoma terminology in Rhabditida (Nematoda). Nematologica. 1995;41:153–82.
Abolafia J, Peña-Santiago R. On the identity of Chiloplacus magnus Rashid & Heyns, 1990 and C. insularis Orselli & Vinciguerra, 2002 (Rhabditida: Cephalobidae), two confusable species. Nematology. 2017;19:1017–34.
Bird AF, Bird J. The structure of nematodes, vol. 2. New York: Academic Press; 1991. p. 317.
Baldwin JG, Perry RN. Nematode morphology, sensory structure and function. In: Chen ZX, Dickson DW, Chen SY, editors. Nematology: advances and perspectives. Nematode morphology, physiology and ecology. Wallingford: CAB International; 2004. p. 197–201.
Dix I, Burnell AM, Griffin CT, Joyce SA, Nugent MJ, Downes MJ. The identification of biological species in the genus Heterorhabditis (Nematoda: Heterorhabditidae) by cross-breeding second-generation amphimictic adults. Parasitology 1992;104:509–18.
Suman B, Bhat AH, Aasha R, Chaubey AK, Abolofia J. Morphological and molecular characterization of Distolabrellus veechi (Rhabditida: Mesorhabditidae) from India. J Nematol. 2020;22:439–52.
Vrain TC, Wakarchuk DA, Lévesque AC, Hamilton RI. Intraspecific rDNA restriction fragment length polymorphism in the Xiphinema americanum group. Fund Appl Nematol. 1992;15:563–73.
Nadler SA, Bolotin E, Stock SP. Phylogenetic relationships of Steinernema Travassos, 1927 (Nematoda: Cephalobina: Steinernematidae) based on nuclear, mitochondrial and morphological data. Syst Parasitol. 2006;63:159–79.
Kuwata R, Yoshiga T, Yoshida M, Kondo E. Phylogenetic relationships of Japanese Heterorhabditis nematodes and their symbiotic Photorhabdus bacteria. Jpn J Nematol. 2007;37:39–50.
Bhat AH, Ameni L, Abolafia J, Machado RAR, Kallel S, Comparative morphological and molecular analyses of Acrobeloides bodenheimeri and A. tricornis Cobb. (Rhabditida, Cephalobidae) from Tunisia. Nematology. 1924;2023:207–26.
Dhakal M, Nguyen KB, Hunt DJ, Ehlers RU, Spiridonov SE, Subbotin SA. Molecular identification, phylogeny and phylogeography of the entomopathogenic nematodes of the genus Heterorhabditis Poinar, 1976: a multigene approach. Nematology. 2020;23:451–66.
Kimura MA. Simple method for estimating evolutionary rates of base substitutions through comparative studies of nucleotide sequences. J Mol Evol. 1980;16:111–20.
Hasegawa M, Kishino H, Yano T. Dating of the human-ape splitting by a molecular clock of mitochondrial DNA. J Mol Evol. 1985;22:160–74.
Nei M, Kumar S. Molecular evolution and phylogenetics. Oxford University Press; 2000. p. 352.
Tamura K, Stecher G, Kumar S. MEGA11: molecular evolutionary genetics analysis version 11. Mol Biol Evol. 2021;38:3022–7.
Edgar RC. MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Res. 2004;32:1792–7.
Chevenet F, Brun C, Bañuls AL, Jacq B, Christen R. TreeDyn: towards dynamic graphics and annotations for analyses of trees. BMC Bioinform. 2006;7:1–9.
Letunic I, Bork P. Interactive tree of life (iTOL) v3: An online tool for the display and annotation of phylogenetic and other trees. Nucleic Acids Res. 2016;44:W242–5.
Machado RAR, Ameni L, Bhat AH, Mastore M, Terrettaz C, Brivio MF, et al. Acinetobacter nematophilus sp. nov., Alcaligenes nematophilus sp. nov., Enterobacter nematophilus sp. nov., and Kaistia nematophila sp. nov., isolated from soil-borne nematodes and proposal for the elevation of Alcaligenes faecalis subsp. faecalis, Alcaligenes faecalis subsp. parafaecalis, and Alcaligenes faecalis subsp. phenolicus to the species level. Taxonomy. 2023;3:148–68.
Machado RAR, Bhat AH, Fallet P, Turlings TCJ, Kajuga J, Yan X, et al. Xenorhabdus bovienii subsp. africana subsp. nov., isolated from Steinernema africanum entomopathogenic nematodes. Int J Syst Evol Microbiol. 2023;73:1–9.
Bolger AM, Lohse M, Usadel B. Trimmomatic: a flexible trimmer for Illumina sequence data. Bioinform. 2014;30:2114–20.
Bankevich A, Nurk S, Antipov D, Gurevich AA, Dvorkin M, Kulikov AS, et al. SPAdes: a new genome assembly algorithm and its applications to single-cell sequencing. J Comput Biol. 2012;19:455–77.
Walker BJ, Abeel T, Shea T, Priest M, Abouelliel A, Sakthikumar S, et al. Pilon: an integrated tool for comprehensive microbial variant detection and genome assembly improvement. PLoS ONE. 2014;9:e112963.
Page AJ, Cummins CA, Hunt M, Wong VK, Reuter S, Holden MTG, et al. Roary: rapid large-scale prokaryote pan genome analysis. Bioinform. 2015;31:3691–3.
Price MN, Dehal PS, Arkin AP. FastTree: computing large minimum evolution trees with profiles instead of a distance matrix. Mol Biol Evol. 2009;26:1641–50.
Poinar GO Jr. Description and biology of a new insect parasitic rhabitoid, Heterorhabditis bacteriophora n. gen. n. sp. (Rhabditida; Heterorhabditidae n. family). Nematologica. 1976;21:463–70.
Li XY, Qi-Zhi L, Nermut J, Půža V, Mráček Z. Heterorhabditis beicherriana n. sp. (Nematoda: Heterorhabditidae), a new entomopathogenic nematode from the Shunyi district of Beijing, China. Zootaxa. 2012;3569:25–40.
Nguyen KB, Shapiro-Ilan DI, Mbata GN. Heterorhabditis georgiana n. sp. (Rhabditida: Heterorhabditidae) from Georgia, USA. Nematology. 2008;10:433–48.
Andaló V, Nguyen KB, Moino A. Heterorhabditis amazonensis n. sp. (Rhabditida: Heterorhabditidae) from Amazonas, Brazil. Nematology. 2006;8:853–67.
Edgington S, Buddie AG, Moore D, France A, Merino L, Hunt DJ. Heterorhabditis atacamensis n. sp. (Nematoda: Heterorhabditidae), a new entomopathogenic nematode from the Atacama Desert, Chile. J Helminthol. 2011;85:381–94.
Phan LK, Subbotin SA, Nguyen CN, Moens M. Heterorhabditis baujardi sp. n. (Rhabditida: Heterorhabditidae) from Vietnam with morphometric data for H. indica populations. Nematology. 2003;5:367–82.
Stock SP, Griffin CT, Burnell AM. Morphological characterisation of three isolates of Heterorhabditis Poinar, 1976 from the ‘Irish group’ (Nematoda: Rhabditida: Heterorhabditidae) and additional evidence supporting their recognition as a distinct species, H. downesi n. sp. Syst Parasitol. 2002;51:95–106.
Nguyen KB, Gozel U, Koppenhöfer HS, Adams BJ. Heterorhabditis floridensis n. sp. (Rhabditida: Heterorhabditidae) from Florida. Zootaxa. 2006;1177:1–19.
Liu J, Berry RE. Heterorhabditis marelatus n. sp. (Rhabditida: Heterorhabditidae) from Oregon. J Invertebr Pathol. 1996;67:48–54.
Poinar GO Jr, Jackson T, Klein M. Heterorhabditis megidis sp. n. (Heterorhabditidae: Rhabditidia), parasitic in Japanese beetle, Popillia japonica (Scarabidae: Coleoptera), in Ohio. Proc Helminthol Soc Wash. 1987;53:53–9.
Nguyen KB, Shapiro-Ilan DI, Stuart RJ, Mccoy CW, James RR, Adams BJ. Heterorhabditis mexicana n. sp. (Rhabditida: Heterorhabditidae) from Tamaulipas, Mexico, and morphological studies of the bursa of Heterorhabditis spp. Nematology. 2004;6:231–44.
Malan AP, Knoetze R, Tiedt L. Heterorhabditis noenieputensis n. sp. (Rhabditida: Heterorhabditidae), a new entomopathogenic nematode from South Africa. J Helminthol. 2014;88:139–51.
Malan AP, Nguyen KB, De Waal JY, Tiedt L. Heterorhabditis safricana n. sp. (Rhabditida: Heterorhabditidae), a new entomopathogenic nematode from South Africa. Nematology. 2008;10:381–96.
Shamseldean MM, Abou El-Sooud AB, Abd-Elgawad MMM, Saleh MM. Identification of a new Heterorhabditis species from Egypt, Heterorhabditis taysearae n. sp. (Rhabditida: Heterorhabditidae). Egyp J Biol Cont. 1996;6:129–38.
Poinar GO. Taxonomy and biology of Steinernematidae and Heterorhabditidae. In: Gaugler R, Kaya HK, editors. Entomopathogenic nematodes in biological control. Boca Raton: CRC Press; 1990. p. 23–61.
Bedding RA, Akhurst RJ. A simple technique for the detection of insect parasitic rhabditid nematodes in soil. Nematologica. 1975;21:109–10.
Askary TH, Bhat AH, Machado RAR, Ahmad MJ, Abd-Elgawad MMM, Khan AA, et al. Virulence and reproductive potential of Indian entomopathogenic nematodes against the larvae of the rice meal moth. Arch Phytopathol Plant Prot. 2023;55:2237–49.
Machado RAR, Bhat AH, Castaneda-Alvarez C, Askary TH, Půža V, Pagès S, et al. Xenorhabdus aichiensis sp. nov., X. anantnagensis sp. nov., and X. yunnanensis sp. nov., isolated from Steinernema entomopathogenic nematodes. Curr Microbiol. 2023;80:300.
Sudhaus W. Phylogenetic systematisation and catalogue of paraphyletic Rhabditidae (Secernentea, Nematoda). J Nematode Morphol Syst. 2011;14:113–78.
The authors thank the Institute of Biology of the University of Neuchâtel, Switzerland, and the Swiss National Science Foundation for their support.
The work of AHB was supported by a Swiss Government Excellence Scholarship (grant no. 2021.0463 to AHB). The work of RARM is supported by the Swiss National Science Foundation (grant no. 186094 to RARM). ANRC is a postdoctoral researcher at the University of Jaén, and has a postdoctoral grant for requalification from the Spanish University System 2021–2023 (modality ‘Margarita Salas’), financed by European Union-Next Generation funding through the Spanish Ministry of Universities. JA thanks the University of Jaén, Spain, for the financial support provided by the Research Support Plan POAIUJA 2021/2022: EI_RNM02_2021. This research was also funded by Researchers Supporting Project number (RSP2023R364), King Saud University, Riyadh, Saudi Arabia.
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Comparative morphometrics of infective juveniles and adult generations of Heterorhabditis casmirica n. sp. with type populations of Heterorhabditis bacteriophora and Indian strains. All data, with the exception of ratios and percentages, are given in micrometers and as mean (range). Table S2. Pairwise distances in base pairs of the ITS rRNA regions between species of Heterorhabditis and Heterorhabditis casmirica n. sp. Data for H. casmirica n. sp. are in italic. Table S3. Pairwise distances in base pairs of the D2–D3 rRNA regions between species of Heterorhabditis and Heterorhabditis casmirica n. sp. Data for H. casmirica n. sp. are in italic. Table S4. National Center for Biotechnology Information (NCBI) accession numbers of the nucleotide sequences used for the phylogenetic analyses in this study; the sequences newly generated in this study are in italic. Table S5. NCBI accession numbers of the genomic sequences of different Photorhabdus species used in this study; the sequences newly generated in this study are in italic.
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Bhat, A.H., Machado, R.A.R., Abolafia, J. et al. Taxonomic and molecular characterization of a new entomopathogenic nematode species, Heterorhabditis casmirica n. sp., and whole genome sequencing of its associated bacterial symbiont. Parasites Vectors 16, 383 (2023). https://doi.org/10.1186/s13071-023-05990-z