STEINERNEMA ADAMSI N. SP. (RHABDITIDA: STEINERNEMATIDAE), A NEW ENTOMOPATHOGENIC NEMATODE FROM THAILAND

Entomopathogenic nematodes (EPNs) are a group of nematodes that specifically infect and parasitize insects and are distributed worldwide (Adams et al., 2006; Dillman et al., 2012). Broadly, most described species of EPNs are classified into 2 families: Steinernematidae and Heterorhabditidae, and they have a symbiotic relationship with bacteria from the genera Xenorhabdus and Photorhabdus, respectively (Stock and Blair, 2008). These nematodes are effective in controlling and killing insects; hence they have great potential in biological control and integrated pest management. Some species of EPNs are commercially produced and are readily available for use (Shapiro-Ilan et al., 2023).

Pest control in commercial agriculture is largely based on chemical pesticides, which are detrimental to the environment and human health (Rani et al., 2021). The growing awareness of these issues has sparked a rise in the popularity of biological control—including classical, augmentation, and conservation methods (Stuart et al., 2006, 2008). The success of EPNs in controlling some important insect pests has sparked research into various aspects of their biology, allowing researchers to identify effective strains of EPNs that can be used as biological control agents (Parkman and Smart, 1996; Nguyen and Duncan, 2002; Stock et al., 2018). Although EPNs have great potential in biological control, understanding is limited as to their diversity, behavior, and factors affecting their distribution (Stuart et al., 2006). One hundred species of Steinernematidae have been recovered and described from soils around the world, and the number of newly described species is increasing (Koppenhöfer et al., 2020; Lis et al., 2021; Tian et al., 2022). According to phylogenetic analyses, nematodes within Steinernematidae are divided into 12 multispecies clades: Glaseri, Karii, Longicaudatum, Khoisanae, Costaricense, Cameroonense, Feltiae, Kushidai, Monticolum, Carpocapsae, Bicornutum, and Affine, and 9 of these clades can be combined into 3 superclades (Spiridonov and Subbotin, 2016).

Discovering new EPN species within Steinernematidae and using them for agricultural pest management may result in a sustainable solution because few nematodes within Steinernematidae seem to have strong host specificity, making the newly identified species of Steinernema a potential source of effective biological control (Lis et al., 2021). A new species of Steinernema was discovered during an EPN survey of agricultural fields in Thailand. Here we describe the morphological, morphometric, and molecular characterization of the newly discovered species, Steinernema adamsi n. sp., from Thailand.

The nematode was first recovered from a soil sample from the rhizosphere of a longan tree (Dimocarpus sp.) in the Mueangjee Sub-District, Mueang Lamphun District (18°28ʹ05.1″N, 98°58ʹ17.7″E), Lamphun Province, Thailand, during an EPN survey between October 2017 and October 2019. Approximately 500 g of collected soil was placed in a plastic bag and transported to the laboratory at the Department of Microbiology and Parasitology, Faculty of Medical Science, Naresuan University, Phitsanulok Province, Thailand. To isolate nematodes, 5 waxworm (Galleria mellonella) larvae were used as bait as previously described (Bedding and Akhurst, 1975). The dead G. mellonella were washed, and infective juveniles (IJs) were collected using a modified White trap (White, 1927; Kaya and Stock, 1997). The new isolate of Steinernema was noted as Steinernema eALN11.5_TH and sent to the Department of Nematology, University of California Riverside (UCR) for further characterization. Newly obtained IJs were washed using M9 buffer (6 g Na₂HPO₄, 3 g KH₂PO₄, and 5 g NaCl dissolved in 900 ml distilled deionized [DI] water and autoclaved) and transferred to a lawn of its symbiotic bacteria on nematode growth media (NGM) plates (20 g agar, 3 g NaCl, 10 ml Uracil [2 g/L], 2.5 g Peptone, 975 ml DI water), autoclaved at 121 C at 15 psi for 15 min. After autoclaving, 1 ml cholesterol (5 mg/ml in ethanol), 1 ml 1 M MgSO₄, 1 ml 1 M CaCl₂, and 25 ml 1 M KPO₄ (pH 6.0) were added, and plates were incubated at 27 C. Nematodes used for morphological and morphometric studies were maintained on an NGM medium inoculated with the symbiotic bacteria and were relocated to fresh plates whenever necessary.

For light microscopy, infective juveniles (IJs) were collected from the infected cadavers using modified White traps. Males and females were also collected from the NGM plates. All nematodes were collected in M9 buffer and prepared for fixation in a hot water bath at 60 C for 2 min. An equal volume of double-strength formalin-glycerol solution (4:1) was heated with the nematode solution, mixed, and incubated at room temperature for 2 days. Formalin-preserved nematodes were processed to anhydrous glycerol using modified Seinhorst’s rapid method to glycerol (Seinhorst, 1959). The specimens were mounted on glass slides for morphometric measurement using Volocity image analysis software (Quorum Technologies, Puslinch, Ontario, Canada). The images of analyzed specimens were recorded on a Nikon Eclipse E600 (Nikon, Melville, New York) and Keyence (BZ-X800 Series) (Keyence Corporation, Osaka, Japan). Using Adobe Photoshop, line drawings of all life stages were created by tracing over images captured with the Keyenece and Nikon Eclipse E600 microscope.

Approximately 10 freshly emerged IJs were used for DNA extraction. The surface sterilized IJs were pipetted into a 0.2 ml microcentrifuge polymerase chain reaction (PCR) tube mixed with 9 μl of 10 mM Tris, 1 mm EDTA, 0.5 μl of 2% Triton X, and 0.5 μl of Proteinase K (20 mg/ml, New England Biolabs, Ipswich, Massachusetts). The nematodes’ cuticle was disrupted by freezing and thawing the sample 3 times using liquid nitrogen, and then the nematode was incubated overnight at −20 C. The next day the frozen lysate was incubated at 56 C for 1 hr followed by 95 C for 10 min for DNA extraction. PCR reaction was performed with a total volume of 50 μl, which included 4 μl of genomic DNA as the template, 25 μl of EconoTaq PLUS 2X Master Mix (Lucigen, LGC Biosearch Technologies, Madison, Wisconsin), 2.5 µl each of forward and reverse primers at a concentration of 10 µM, and 16 µl of nuclease-free water. The primer 18S: 5-TTGATTACGTCCCTGCCCTTT-3 (forward) and 26R: 5-TTTCACTCGCCGTTACTAAGG-3 (reverse) was used to amplify the ITS1–5.8S–ITS2 region of rDNA gene (Vrain et al., 1992). Similarly, the 28S rDNA gene was amplified using D2F: 5-CCTTAGTAACGGCGAGTGAAA-3 (forward) and 536: 5-CAGCTATCCTGAGGAAAC-3 (reverse) primers (Nguyen et al., 2006). Similar conditions were used for the PCR amplification of both loci: initial denaturation at 94 C for 5 min, then 35 cycles at 94 C for 1 min, annealing at 57 C for 90 sec, extension at 72 C for 2 min, and a final extension step at 72 C for 10 min. PCR results were visualized on a 1% agarose gel stained with 0.0003% ethidium bromide and a 1-kb plus DNA ladder (New England Biolabs). PCR products were purified using the QIAquick® PCR Purification Kit (Qiagen, Germantown, Maryland) following the manufacturer’s protocol. Purified PCR products were Sanger sequenced from the forward and reverse strands at the UCR Core Instrumentation Facility according to the manufacturer’s protocol. Chromatogram sequence ambiguities of both forward and reverse sequences of each locus were visually checked and assembled using SeqManII software (DNASTAR, Madison, Wisconsin). Closely related species were identified in the National Center for Biotechnology Information (NCBI) database using the Basic Local Alignment Search Tool (BLAST). The sequences generated as part of this investigation have been deposited in NCBI GenBank under the accession numbers OR050811 for ITS sequences and OR050810 for D2-D3 rDNA sequences.

The ITS and 28S sequences of S. adamsi n. sp. and their respective best hits from NCBI (Table I) were aligned using Clustal W in MEGA 11 with the default parameters, and some alignment inconsistencies were corrected manually (Tamura et al., 2021). The aligned files were used to infer the phylogenetic relationships using the BEAST v. 1.10.4 program (Drummond and Rambaut, 2007) which used Bayesian analyses using the GTR + I + G model of DNA evolution as analyzed by Spiridonov and Subbotin (2016). The BEAST package uses a family of Markov chain Monte Carlo (MCMC) algorithms for Bayesian phylogenetic inference. To construct the concatenated phylogenetic tree of ITS and 28S, all aligned sequences of the isolates that are represented in both ITS and 28S phylogenetic trees were concatenated with a Bayesian analysis using the GTR + I + G model of DNA evolution as analyzed by Spiridonov and Subbotin (2016). The characteristics of the BEAST file were determined using BEAUti, which included the best substitution model from a previously determined model, clock type strict, MCMC chain length of 10,000,000, and all other parameters were at the default setting. The resulting tree was visualized using TreeAnnotator v. 1.10.4, and a burn-in of 10,000 states was specified. The phylogenetic trees were visualized using Figtree v. 1.4.4 (http://tree.bio.ed.ac.uk/software/figtree/).

To examine the reproductive compatibility of S. adamsi n. sp. with other close relative species within the genus Steinernema a cross-hybridization assay was conducted in NGM plates. Two EPN species Steinernema glaseri and Steinernema diaprepesi were used to test the reproductive compatibility. There were 10 replicates per cross; in the first 5 plates, individual females of test species were crossed with 2 males of S. adamsi n. sp.; in the remaining 5 plates, 2 males of test species were crossed with 1 female of S. adamsi n. sp. The experiment was conducted twice with controls consisting of 2 male and 1 female nematodes of the same species.

To extract the symbiotic bacteria associated with S. adamsi n. sp., G. mellonella larvae were infected with IJs. After 48 hr of infection, the insect cadaver was surface sterilized by dipping it into 75% ethanol for 1 minute and placed in a sterile Petri dish to dry. With sterile forceps, the third segment from the head of G. mellonella was opened, and its hemolymph was excessed using a sterile loop. The loop was then streaked on nutrient agar plates supplemented with bromothymol blue and triphenyl-2,3,5-tetrazolium chloride (NBTA; Sigma-Aldrich, St. Louis, Missouri). Normally colonies of Xenorhabdus on NBTA are blue or brown, but we have observed that the symbionts of S. adamsi n. sp. are greenish. The green colonies of the Xenorhabdus bacteria were observed after 5 days of incubation in a dark condition at 28 C. After 5 days a single colony of the Xenorhabdus bacteria was transferred to 5 ml of Luria-Bertani (LB) broth and incubated at 28 C overnight with shaking at 180 rpm. To identify the symbiotic bacteria, genomic DNA was extracted from bacterial pellets using the Proteinase K method as described above. The PCR amplification of the 16S rRNA gene used EconoTaq PLUS 2X Master Mix (Lucigen). The primers used for PCR reaction are F8: 5-CAGGCATCCAGACTTTGATYMTGGCTCAG-3 (forward) and R1512: 5-GTGAAGCTTACGGYTAGCTTGTTACGACTT-3 (reverse) as previously described (Dreyer et al., 2018). The PCR conditions were the same as above except for the annealing temperature, which was set at 55 C. The PCR product was gel purified and Sanger sequenced as previously described. A partial 16S rDNA gene sequence was compared with existing gene sequences in the NCBI database using BlastN (Agarwala et al., 2017). Based on the Blast result, different Xenorhabdus species were used to construct the phylogeny. The 16S sequences of Morganella morganii (AJ301681) were used as an outgroup. All the sequences were aligned using Clustal W in MEGA 11 with the default parameters, and some alignment inconsistencies were corrected manually (Tamura et al., 2021). The program IQ-Tree (Nguyen et al., 2015) was used to select the best-fitting model for the 16S data set using the “find best model” function and to construct a maximum-likelihood (ML) phylogenetic tree, utilizing ultrafast bootstrap branch support with 1,000 replicates.

The cryopreservation for Steinernema adamsi n. sp. was optimized by modifying a trehalose-DMSO freezing protocol for C. elegans (Poullet and Braendle, 2015). IJs freshly emerged from wax worms were placed on the NGM plates seeded with symbionts of S. adamsi n. sp. The NGM plates were incubated at 28 C until the bacteria on the NGM plates were completely depleted. At this stage, the plates are full of the mixed stage nematodes with most of the eggs freshly hatched, and females are full of J1 stage nematodes that are internally hatched. Nematodes from the several plates were collected by washing nematodes using M9 buffer and concentrated using centrifugation for 1 min at ∼1,000 g at room temperature in a 15 ml tube. Once the nematodes were concentrated the pellet was incubated in 5 ml of trehalose-DMSO solution for more than 30 min (1 L of trehalose-DMSO solution is prepared with 30.2 gm D-Trehalose anhydrous [Thermo Fisher Scientific, Waltham, Massachusetts]), 35.4 ml dimethylsulfoxide [DMSO, Fisher Scientific], volume adjusted to 1 L using M9, buffer and is filter sterilized). The nematodes were transferred into cryotubes to slowly reduce the temperature, and cryotubes were placed inside Styrofoam boxes and placed in a −80 C freezer overnight. After incubation cryotubes were taken out of the Styrofoam box and transferred to a cardboard box in a −80 C freezer for long-term storage.

• Order Rhabditida Chitwood, 1933 Family Steinernematidae Filipjev, 1934 Genus Steinernema; Travassos, 1927 Steinernema adamsi n. sp.

Heat-killed and relaxed females are C-shaped (Figs. 1, 2), 628–2,803 µm long (Table II). Head round and cuticle smooth under light microscopy. An excretory pore opening is present anterior to the nerve ring. Stoma reduced (cheilo-, gymno- and stegostom vestigial), short and wide funnel shaped, inconspicuous sclerotized walls. The nerve ring surrounds the isthmus just anterior to the basal bulb. The esophagus is muscular with a cylindrical procorpus, followed by a slightly swollen non-valvated metacorpus, isthmus, and basal bulb. Gonads are didelphic-amphidelphic. Vulva is located at the mid-portion of the body. The vagina is short and connected to paired uteri. Vulva a transverse slit varying from slightly protruding to significantly protruding from the body surface, with a thick flap. Females deposit the eggs internally; subsequently, juveniles hatch inside the female and bore their way out. Tail blunt, conoid. Post-anal lips non-protruding or slightly protruding.

Heat-killed specimens have J-shaped slender body and are ventrally curved posteriorly (Figs. 3, 4). Cuticle of the male is smooth, inconspicuous lateral field and phasmids under light microscopy. The head is truncated to somewhat round, and it is continuous with the body. Stoma like that of the female. Esophagus muscular; procorpus cylindrical; metacorpus slightly swollen, non-valvate; isthmus distinct Nerve ring just anterior to the basal bulb. Excretory pores at the middle of the esophagus. The reproductive system (testis) is monorchic and ventrally reflexed. The spicules average 65.85 µm long and are moderately curved and golden brown with a sharp tip (Table II). Gubernaculum fusiform, approximately about one-half of the length of spicules. Tail conoid and non-mucronate, and bursa absent.

Body straight or slightly curved, tapering gradually towards the anterior end from the base of esophagus and towards the distal end when infective juveniles are heat killed and relaxed (Figs. 3, 4). The mouth and the anus are closed, and the esophagus and intestine are collapsed. Excretory pores are located approximately around the mid-corpus region. The nerve ring is located at isthmus level slightly above basal bulb. The lateral field patterns and lateral lines were not observed in light microscopy. The intestine lumen is narrow and is filled with fat globules. Rectum is long, straight, and anus is distinct. The hyaline occupies c. 46% of the tail length (Table II). The tail is conoid with pointed terminus.

The rhizosphere of the longan tree (Dimocarpus sp.) in Lamphun Province, Thailand (18°28′05.1″N, 98°58′17.7″E).

The natural host of this novel species is unknown.

Six females (allotype and paratypes), 6 males (holotype and paratypes), and 6 infective juveniles (IJs) were deposited in the nematode collection at the University of California–Davis, Davis, California. The accession number for the holotype male and allotype female is UCDNC 5260, for the paratype male is UCDNC 5261, for the paratype female is UCDNC 5262, and for the infective juvenile is UCDNC 5263.

ITS rDNA genes (OR050811) and D2/D3 domain of the 28S (OR050810) are deposited in GenBank.

urn:lsid:zoobank.org:act:B9B5622C-D2F9-4DA8-8249-1BD1336BBB8D.

This species is named in honor of Dr. Byron J. Adams, acknowledging his extensive contributions to the study of the ecology and biology of entomopathogenic nematodes and our understanding of species.

Cross-hybridization assays between males and females of S. adamsi with S. glaseri and S. diaprepesi yielded no progeny. However, offspring were produced in all self-crossed species.

Sequences obtained from the ITS rDNA genes (OR050811) and D2/D3 domain of the 28S (OR050810) are deposited in GenBank. Based on the BLAST analysis in NCBI of ITS sequences of S. adamsi, we selected 81 different Steinernema species. In addition to the 82 steinernematids (81 plus S. adamsi), 3 outgroup species were included to construct phylogenetic trees. The phylogenetic tree inferred from the Bayesian analysis of ITS datasets is given in Figure 5. Based on the ITS phylogeny all the Steinernema nematodes were differentiated into 11 clades: ‘Glaseri’ (11 species), ‘Karii’ (7), ‘Longicaudatum’ (6), ‘Khoisanae’ (4), ‘Costaricense’ (2), ‘Feltiae’ (16), ‘Kushidai’ (2), ‘Monticolum’ (5), ‘Carpocapsae’ (13), ‘Bicornutum’ (6), ‘Affine’ (7), and 3 monospecies clades for Steinernema unicornum, Steinernema neocurtillae, and Steinernema rarum. The phylogenetic tree inferred from the Bayesian analysis of 28S data sets is represented in Figure 6. All the Steinernema species were divided into 11 clades similar to those of the ITS phylogeny except for 1 additional Cameroonense clade (3 species) instead of the Costaricense clade. The placement of the new species was consistent with the ITS phylogeny in the Longicaudatum clade with the posterior possibility (77%) (Fig. 6). For the concatenated phylogeny, we used species represented in both ITS and D2D3 phylogenies. Bayesian analysis of the concatenated dataset of 48 species yielded a similar topology to that of the ITS and D2D3 phylogenies, placing S. adamsi firmly in Longicaudatum clade, with a very high posterior probability (100%) (Fig. 7).

The efficacy and survival of the frozen nematode stocks were tested on NGM plates 2 wk and 2 mo after the cryopreservation at −80 C. We were able to recover mixed-stage nematodes from both tubes, and these nematodes reproduced stably downstream.

The sequence obtained from the 16S rDNA gene with accession number OR047834 is deposited in GenBank. The BLAST analysis of 16S rDNA gene sequences revealed that the symbiotic bacterium is 99.7% similar to Xenorhabdus griffiniae. Based on the BLAST result, 25 different Xenorhabdus species were used to construct the phylogeny. The IQ-Tree program selected TIM2+F+I+G4 as the best-fitting model to construct a maximum-likelihood (ML) phylogenetic tree. Subsequently, a maximum likelihood phylogenetic tree for Xenorhabdus spp. isolated from S. adamsi n. sp. was generated based on a partial 16S rDNA sequence (1,286 bp), in combination with Xenorhabdus strains from GenBank. Morganella morganii was used as an outgroup. The bootstrap values were computed based on 1,000 replicates, and the numbers displayed on the branches represent the support values for Maximum Likelihood clades above the 50% threshold (Fig. 8).

Based on molecular, morphological, and morphometric traits, S. adamsi is a member of the ‘Longicaudatum-clade.’ Nematodes of this group are characterized by having third-stage IJs between 832 mm to 1,055 mm long. Steinernema adamsi, a member of the ‘Longicaudatum-clade,’ presents several traits common to this group. Morphologically, the IJs of S. adamsi can be recognized by having a medium body length of 879.66 (range: 774.69–956.96) µm, compared to other members of the same clade, Steinernema lamjungense 832 (690–950) µm being the smallest and Steinernema guangdongense 1,055 (987–1,145) µm being the largest. A similar pattern was observed for the body diameter with 34.96 (31.58–39.85) µm, with other members S. lamjungense 29 (23–31) µm being the smallest and S. guangdongense 42 (30–48) µm as the largest. Within the Longicaudatum clade, the new species shares some of the hallmark morphological traits. However, the new species can be differentiated from closely related taxa by several morphometric and morphological differences between the IJs and males (Tables III, IV). Based on the ITS phylogeny, the novel species is closely related to S. guangdongense. However, morphologically the third-stage juveniles of new species are much shorter (average 879.66 µm vs. 1,055 µm). Based on the 28S phylogeny, the new species is placed together with Steinernema hermaphroditum. Morphologically, S. hermaphroditum IJs are larger compared to the new species (average 1,043 µm vs. 879.66 µm). Other morphological traits such as execratory pores of S. adamsi are more anteriorly located than that of S. guangdongense and posteriorly located compared to that of S. hermaphroditum (average 69.84 µm vs. average 80 µm vs. average: 65 µm). The closely related species also differ in the values of ‘c’ (ratio of total body length to the tail length) with an average of 12.08 in S. adamsi, 11.6 in S. guangdongense, and 10.6 in S. hermaphroditum. Similarly, other morphometric values for IJs also differed between closely related species such as ‘D%’ (the proportion of the length from the anterior end to the excretory pore to the length from the anterior end to the esophagus; 67.87 in S. adamsi vs. 59 S. guangdongense and 50 in S. hermaphroditum) and ‘E%’ (the proportion of the length from the anterior end to the excretory pore to the tail length; 96.24 in S. adamsi vs. 88 S. guangdongense and 85 in S. hermaphroditum). Additionally, IJs of the novel species differ from S. guangdongense, S. hermaphroditum, S. longicaudum, and S. lamjungense by having a shorter tail (average: 72.91 μm). The new species can be recognized further by the morphological characteristics of males. The spicules of S. adamsi are shorter, i.e., an average of 61.53 µm vs. an average of 68 µm in S. hermaphroditum and an average of 86 µm in S. guangdongense. Similarly, gubernaculum length also varied between 3 closely relative species (37.01µm in S. adamsi vs. 64 µm in S. guangdongense. vs. 48 µm in S. hermaphroditum), the values of the ‘SW%’ (the proportion of spicule length to the anal body diameter) (average: 126.79 in S. adamsi vs. 150 in S. hermaphroditum and 175 in S. guangdongense), and ‘D%’ (the proportion of the length from the anterior end to the excretory pore to the length from the anterior end to the esophagus) were different 68.23 in S. adamsi vs. 48 in S. hermaphroditum vs. 70 in S. guangdongense. Steinernema adamsi can be distinguished from other related species by morphometrical characteristics listed in Tables III and IV.

In this paper we focus more on the molecular characterization of the nematodes than morphological and morphometric characterization. The addition of detailed morphological features may provide more support for the classification of steinernematid species into different clades; however, support for the distinction between 2 closely related nematodes inferred from the molecular data does not significantly differ from morphological and morphometric characteristics (Spiridonov et al., 2004). Some morphological features are shared between nematodes of different clades; for example, most nematodes within the clades Bicornutum, Glaseri, Khoisane, and Longicaudatum have a lateral field with 8 ridges of equal height and width, which might create confusion during the species differentiation between these congeners (Spiridonov and Subbotin, 2016). Another morphological feature that is used to differentiate males of steinernematids is the number and distribution of the genital papillae. This morphological feature varies even between specimens of the same species, whereas, most of the time, steinernematids share a uniform number of papillae, i.e., 11 pairs of papillae, and 1 midventral precloacal papilla. Thus, the number of papillae (22 + 1) is a shared characteristic of most steinernematids, providing little help in differentiating species between different clades (Spiridonov and Subbotin, 2016).

In this study we identified the native symbiont of S. adamsi. Based on the 16S sequence results; the bacteria are identified as Xenorhabdus spp. highly similar to Xenorhabdus griffiniae. Bacterium X. griffiniae is also found in a symbiotic relationship with other species, including S. hermaphroditum and Steinernema khoisanae (Mothupi et al., 2015; Cao et al., 2022). Despite having high similarity to the species X. griffiniae, Xenorhabdus spp. isolated from S. adamsi grown on NGM have a greenish hue, which is not true of other X. griffiniae isolates. In this study, we did not investigate this further, but it seems to be a novel characteristic of this strain of bacteria that warrants further investigation.

Entomopathogenic nematodes parasitize insects and are used in biological control programs against insect pests in agriculture. The use of chemical pesticides for insect control has negative impacts on both human health and the environment. Using EPNs in pest management selectively targets insects, reducing harmful effects on human health, non-target creatures, and the environment (Piedra-Buena et al., 2015). Some species of EPNs have a broad host range such as Steinernema carpocapsae, which can infect approximately 200 insects across 10 orders, and some species have a specific host range such as Steinernema scapterisci, which is adapted to infect insects within the order Orthoptera (Koppenhöfer et al., 2020). However, the host range of the majority of EPNs is largely unknown (Peters, 1996; Shapiro-Ilan et al., 2017). In laboratory assays, nematodes may have a wider than normal host range that does not translate to the field. However, species that are isolated from the natural environment are well adapted to those environmental conditions and are effective against existing pests (Campos-Herrera et al., 2016). The host range of S. adamsi has not been explored yet, so future studies should focus on studying the host range of this novel species to identify the potential host to improve the efficacy of EPNs in biological control. Most of the EPNs that are isolated from the environment are reared in the waxworm Galleria mellonella, as the waxworm is highly susceptible to EPNs, is commercially available, and is easy to maintain. However, it has been reported that continuous lab rearing results in the trait deterioration for many valuable traits of EPNs such as infectivity, virulence, longevity, reproductive capacity, heat tolerance, and sex ratio (Gaugler and Campbell, 1991; Hopper et al., 1993; Shapiro et al., 1996; Bai et al., 2005; Bilgrami et al., 2006; Chaston et al., 2011; Stuart and Gaugler, 2011). Though deterioration of biocontrol traits can occur in a laboratory or commercial conditions due to repeated culturing, strains can be stabilized through cryopreservation or by establishing single or multiple purebred homozygous lines (Bai et al., 2005; Anbesse et al., 2013). In this study we were able to successfully modify the cryopreservation technique for novel species and conserve their inherent pathogenic trait.

To summarize, the results obtained from molecular analyses using ITS rDNA, D2D3 of the 28S rDNA gene sequences, and morphological and morphometric analyses provide evidence supporting the designation of S. adamsi as a new species of Steinernema based on the morphological, phylogenetic, and evolutionary species concept (Adams, 1998). This species possesses distinctive characteristics and is well suited to the environmental conditions of Thailand, where it was isolated. However, additional research is necessary to assess the host range of this newly identified species and enhance the efficiency of EPNs in biological control.

We are grateful to members of the Vitta lab who helped isolate and rear this nematode. This research was supported by the United States Department of Agriculture, National Institute of Food and Agriculture (USDA NIFA) Multistate project W5185. This work was partially supported by the Royal Golden Jubilee Ph.D. Program (PHD/0086/2561), through the National Research Council of Thailand (formerly the Thailand Research Fund).