Steinernema Africanum n. Sp. (Rhabditida, Steinernematidae), a New Entomopathogenic Nematode Species Isolated in the Republic of Rwanda

Abstract Alternatives to hazardous insecticides are urgently needed for an environmentally friendly and effective management of insect pests. One such option is the use of entomopathogenic nematodes (EPN). To increase the availability of EPN with potential for biocontrol, we surveyed agricultural soils in the Republic of Rwanda and collected two Steinernema isolates. Initial molecular characterization showed that they represent a new species, for which we propose the name S. africanum n. sp. To describe this new species, we reconstructed phylogenetic relationships, calculated sequence similarity scores, characterized the nematodes at the morphological level, conducted crossing experiments, and isolated and characterized their symbiotic bacteria. At the molecular level, S. africanum n. sp. is closely related to S. litorale and S. weiseri. At the morphological level, S. africanum n. sp. differs from closely related species by the position of the nerve ring and also because the stoma and pharynx region is longer. The first-generation males have ventrally curved spicules with lanceolate manubrium and fusiform gubernaculum and the second-generation males have rounded manubrium and anteriorly hook-like gubernaculum. Steinernema africanum n. sp. does not mate or produce fertile progeny with any of the closely related species.

The genus Steinernema Travassos, 1927 is one of two major genera of entomopathogenic nematodes (EPNs) used to control insect pests in agriculture. The members of this genus infest and kill numerous insects aided by their symbiotic, entomopathogenic bacteria of the genus Xenorhabdus. Together, they constitute a highly valuable pest management tool in sustainable and ecofriendly agriculture (Smart, 1995).
An important step for the use of EPNs in agriculture is the proper description and characterization of the nematode isolates with promising biocontrol traits. In the case of Steinernema, there are hundreds of isolates in different laboratories around the globe, which have been assigned to one of the more than 100 Steinernema species described so far . Many isolates still await being assigned to formal taxonomic studies, and it is very likely that they also represent new, undescribed species.

Nematode morphological and morphometrical characterization, light and scanning electron microscopy
First-and second-generation adult nematodes were obtained by dissecting infested G. mellonella cadavers in Ringer's solution after 5 d to 6 d and 8 d to 9 d of post infestation, respectively. Infective juveniles (IJs) were collected after their emergence from G. mellonella larvae in White traps (White, 1927). Nematodes were killed with water at 60°C, fixed in 4% formalin solution (4 mL formaldehyde, 1 mL glycerol, and 95 mL ddH 2 O) and transferred to anhydrous glycerin by the Seinhorst method (Seinhorst, 1959). Nematodes were then mounted on permanent glass slides with a thicker layer of paraffin wax to prevent the flattening of the nematodes (Grisse, 1969). Morphological measurements were taken using the Olympus BX51 software built into the ZEISS Axio Lab. A1 light microscope (Carl Zeiss Microscopy GmbH, Jena, Germany). Fifteen specimens of S. africanum n. sp. RW14-M-C2b-1 at each developmental stage were measured. To obtain light microscopy (LM) and scanning electron microscopy (SEM) photographs, specimens were processed following techniques described in detail by Abolafia (2022). Briefly, the nematodes, fixed in 4% formalin solution, were processed to anhydrous glycerin with Siddiqi's method using lactophenol-glycerin solutions (Siddiqi, 1964). Then, the nematodes were permanently mounted on glass microscope slides using the glycerin-paraffin method (Maeseneer and d'Herde, 1963;Siddiqi, 1964). Light microscopy photographs were taken using a Nikon Eclipse 80i microscope (Olympus, Tokyo, Japan) equipped with differential interference contrast optics (DIC) and a Nikon Digital Sight DS-U1 camera. For scanning electron microscopy, nematodes preserved in glycerin were taken from the permanent microscope slides by removing the cover glass, re-hydrated in distilled water, dehydrated in a graded ethanol-acetone series, critical point dried with liquid CO 2 , 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) (Abolafia, 2015). Light microscopy micrographs, obtained at different levels for each structure, were processed and combined using AdobeÒ PhotoshopÒCS (Microsoft Corporation, Redmond, WA). Morphological characters of closely related species were taken from the original publications (Hunt and Nguyen, 2016).

Self-crossing and cross-hybridization experiments
Self-crossing and cross-hybridization experiments were conducted as described by Kaya and Stock (1997) with some minor modifications (Kaya and Stock, 1997). Briefly, drops of hemolymph obtained from surface-sterilized G. mellonella larvae were placed in sterile Petri dishes (35 mm x 10 mm). A few micrograms of phenylthiourea were added to hemolymph drops to prevent melanization. Then 40 to 60 surface-sterilized juvenile nematodes (IJs) were added to the hemolymph drops. Nematodes were surface sterilized by immersing them in 0.1% NaOCl, and then washed thrice with autoclaved double distilled water. Then, Petri dishes were wrapped in moistened paper tissue and kept in plastic bags at 25°C. Petri plates were observed daily until IJs developed into adults. Then, male and female adults were separated by observing them under a light microscope. For self-crossing experiments, three males and three females of the same species were transferred to fresh hemolymph drops as described above. For cross-hybridization experiments, three males and three females of different species were transferred to fresh hemolymph drops as described above. Females without males were also included to confirm their virginity. Petri plates were observed daily to determine the production of offspring. For each crossing type, 10 independent Petri plates were included. Experiments were conducted twice under the same conditions. The following species were included in these experiments: Steinernema africanum n. sp. RW14-M-C2b-1 and RW14-M-C2a-3, S. feltiae Jakub, S. ichnusae Sardinia, S. litorale Aichi, and S. weiseri 1025(Yoshida, 2004Tarasco et al., 2008;Půža et al., 2021).

Nematode molecular characterization and phylogenetic relationships
Genomic DNA from about 20 females was extracted using the genomic DNA isolation kit from QIAamp DNA Mini Kit (Qiagen, Valencia, CA) following the manufacturer's instructions. The following genes/ genomic regions were amplified by polymerase chain reaction (PCR): the D2-D3 expansion segments of the 28S rRNA, the ITS region of the rRNA, the mitochondrial 12S rRNA, and the cytochrome oxidase subunit I (COI). To amplify the ITS rRNA, the following primers were used: 18S (5´-TTGATTACGTCCCTGCCC TTT-3´) and 26S (5´-TTTCACTCGCCGTTACTAAGG-3´) (Joyce et al. 1994). To amplify the D2-D3 region, the following primers were used: D2F (5´-CCTTAGTAAC GGCGAGTGAAA-3´) and 536 (5´-CAGCTATCCTGA GGAAAC-3´) (Subbotin et al., 2006). To amplify the 12S mitochondrial rRNA gene, primers 505F: 5´-GTTCCAG AATAATCGGCTAGAC-3´ and 506R: 5´-TCTACTTTACT ACAACTTACTCCCC-3´ were used (Nadler et al., 2006). Primers LCO-1490 (5´-GGTCAACAAATCATAAA GATATTGG-3´) and HCO-2198 (5´-TAAACTTCAGGGT GACCAAAAAATCA-3´) were used to amplify the COI (Folmer et al., 1994). PCR reactions consisted of 12.5 μL of DreamTaq Green PCR Master Mix (Thermo Scientific, Waltham, MA USA), 0.5 μL of each forward and reverse primers at 10 μM, 1 μL of genomic DNA, and 10.5 μL of nuclease free distilled water. The PCR reactions were performed using a thermocycler with the following settings. For ITS and D2-D3: 1 cycle of 5 min at 94°C followed by 40 cycles of 30 sec at 94°C, 30 sec at 50°C, 1 min 30 sec at 72°C, and by a single final elongation step at 72°C for 10 min. For the 12S gene, the PCR protocol included initial denaturation at 94°C for 3 min, followed by 30 cycles of 94°C for 30 sec, 50°C for 30 sec, and 72°C for 45 sec, followed by a final extension at 72°C for 15 min. For the COI gene, the PCR program was as follows: 1 cycle of 94°C for 2 min, followed by 37 cycles of 94°C for 30 sec, 51°C for 45 sec, 72°C for 2 min, and a final extension at 72°C for 12 min. PCR was followed by electrophoresis (45 min, 100 V) of 10 μL of PCR products in a 1% TBA (Trisboric acid-EDTA) buffered agarose gel stained with SYBR Safe DNA Gel Stain (Invitrogen, Carlsbad, CA). PCR products were purified using QIAquick PCR Purification Kit (Qiagen, Valencia, CA) and sequenced using reverse and forward primers by Sanger sequencing (Microsynth AG, Balgach, Switzerland). Obtained sequences were manually curated and trimmed and deposited in the National Center for Biotechnology Information (NCBI) under the accession numbers given on the phylogenetic trees. To obtain genomic sequences of nematodes that belong to all the validly described species of the "feltiae-clade," we searched the database of the NCBI using the Basic Local Alignment Search Tool (Altschul et al. 1990). The resulting sequences were used to reconstruct phylogenetic relationships by the maximum likelihood method based on the following nucleotide substitution models: Hasegawa-Kishino-Yano model (HKY + G) (ITS), General Time Reversible model (GTR + G + I) (COI), Kimura 2-parameter (K2 + G + I) (D2-D3), and Tamura 3-parameter model (T92) (12S) (Kimura, 1980;Hasegawa et al., 1985;Tamura, 1992;Nei and Kumar, 2000). To select the best substitution models, best-fit nucleotide substitution model analyses were carried out in MEGA 7 (Kumar et al., 2016). Sequences were aligned with MUSCLE (v3.8.31) (Edgar, 2004). The trees with the highest log likelihood are shown. The percentage of trees in which the associated taxa clustered together is shown next to the branches. Initial tree(s) for the heuristic search were obtained automatically by applying Neighbor-Join and BioNJ algorithms to a matrix of pairwise distances estimated using the maximum composite likelihood (MCL) approach, and then selecting the topology with superior log likelihood value. In some cases, a discrete Gamma distribution (+G) was used to model evolutionary rate differences among sites and the rate variation model allowed for some sites to be evolutionarily (+I). The trees are drawn to scale, with branch lengths measured in the number of substitutions per site. Graphical representation and edition of the phylogenetic trees were performed with Interactive Tree of Life (v3.5.1) (Chevenet et al., 2006;Letunic and Bork, 2016).

Symbiotic relationships
The entomopathogenic Xenorhabdus bacteria associated with S. africanum n. sp. RW14-M-C2b-1 nematodes were isolated as described (Machado et al., 2018(Machado et al., , 2019. To establish their taxonomic identities, we reconstructed phylogenetic relationships based on whole genome sequences of the isolated bacteria and all the different species of the genus Xenorhabdus. Genomic sequences were obtained as described (Machado et al., 2021b(Machado et al., , 2021c. Genome sequences were deposited in the National Centre for Biotechnology Information. Accession numbers are listed in Table S1 in Supplementary Material. Phylogenetic relationships were reconstructed based on the assembled genomes and the genome sequences of all validly published species of the genus with publicly available genome sequences as described (Machado et al., 2021a). Whole genome sequence similarities were calculated by the digital DNA-DNA hybridization (dDDH) method using the recommended formula 2 of the genome-togenome distance calculator (GGDC) web service of the Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH (DSMZ) (Auch et al., 2010a(Auch et al., , 2010bMeier-Kolthoff et al., 2013, 2014.

Life cycle
The life cycle of S. africanum n. sp. was studied by infesting wax moth larvae (G. mellonella) with either 50 or 150 S. africanum n. sp. RW14-M-C2b-1 IJs per larva (n = 30). Larvae were individually placed in Petri plates lined with a sheet of moist filter paper and incubated at 24°C. Upon infection, a few cadavers were dissected daily to collect and observe the number of nematodes at each developmental stage.

Results and Discussion
Two populations of Steinernema nematodes, RW14-M-C2b-1 and RW14-M-C2a-3, were isolated from agricultural soils in the Republic of Rwanda (Yan et al., 2016). Initial molecular characterization showed that they are identical, belong to the "feltiae-clade," are closely related to S. feltiae, S. citrae, S. litorale, S. nguyeni, and S. weiseri, and represent a new species, which we named S. africanum n. sp. To describe this new species, we compared it with other closely related species at the molecular and morphological level, and conducted cross-hybridization and self-crossing experiments. As both populations are identical at the molecular level, we selected RW14-M-C2b-1 for detailed morphological characterization.

S. weiseri
Mrácek et al. posterior to lateral labial papillae. Stoma shallow, funnelshaped, short and wide, with inconspicuous sclerotized walls; cheilostom short with small rhabdia; gymnostom scarcely developed with minute rhabdia stegostom robust with funnel-shaped lumen and walls with minute rhabdia. Deirids inconspicuous. Pharynx muscular with a cylindrical procorpus, a slightly swollen and nonvalvate metacorpus, narrower isthmus and basal bulb spheroid with reduced valves. Nerve ring (NR) usually located about mid-isthmus level or on the anterior part of the basal bulb. Secretory-EP opening circular, located posterior to NR, close to isthmus-basal bulb junction. Cardia prominent, conoid. Intestine tubular without differentiations. Reproductive system monorchic, ventrally reflexed. Spicules paired, symmetrical, ventrally curved with manubrium rhomboidal, calamus narrower and lamina ventrad curved at anterior part, bearing two longitudinal ribs, and ending in a blunt terminus, with scarcely developed velum not reaching the spicule tip, without rostrum or retinaculum. Gubernaculum fusiform with elongate tip, about one-half of the length of spicules. Tail conoid with rounded terminus bearing a fine mucron. Bursa absent. There are 23 GP (11 pairs and one single) arranged as follows: five pairs subventral precloacal, one pair lateral precloacal, one single mid-ventral papilla, two pairs sub-ventral ad-cloacal, one pair subdorsal post-cloacal, and two pairs of terminal papillae. Phasmids terminal, located between the last pair of GP.

Second-generation male
General morphology similar to that of first-generation males, but smaller in size and slenderer. Tail with mucron robust, dorsally curved. Spicules ventrally curved, with manubrium rounded, calamus slightly narrower than manubrium, and lamina ventrally curved at anterior part, lanceolate posterior part with finely rounded tip, reduced ventral velum, and two longitudinal lateral ribs. Gubernaculum slenderer than that of first-generation male, with manubrium ventrad bent, corpus robust, and narrow and slender terminus. Genital papillae and phasmids with arrangement similar to that in first-generation male.

First-generation female
Body C-shaped when heat-relaxed and fixed. Cuticle with transversal incisures marked, appearing poorly visible annuli. Lateral fields not observed. Deirids inconspicuous, difficult to observe even under SEM. Labial region rounded, continuous with the adjacent part of body. Stoma and pharynx region similar to males. Excretory pore located at level of the

Second-generation female
Similar to first-generation female but smaller. Tail conoid, longer than the first-generation female, lacking mucron. Phasmid located at the posterior part of the tail, at 58% to 59% of tail length.

Third-stage infective juvenile
Body straight or slightly curved when heat-killed, tapering gradually from the base of pharynx to the anterior end and from anus to the distal end. Cuticle with transverse incisures, appearing well-developed annuli. Lateral fields begin as a single line close to the anterior end, increasing to eight ridges, posteriorly gradually reduced to four (anus level) and two (phasmid level). Lip region slightly narrower than the adjacent part of body, with six lips, the lateral ones smaller with six reduced labial and four prominent cephalic papillae. Amphidial apertures pore-like. Stoma reduced, with small cheilostom and elongate gymno-stegostom. Pharynx reduced with narrow corpus, slightly narrower isthmus, and pyriform basal bulb with reduced valves. Nerve ring surrounding the isthmus. Excretory pore located at metacorpus level. Hemizonid present, between NR and pharynx base. Cardia conoid. Deirids inconspicuous.
Intestine bears a bacterial sac at its anterior part. Rectum long, almost straight, with very short cuticular part and elongate cellular part. Anus distinct. Genital primordium located equatorial. Tail conoid, tapering gradually with pointed terminus; cellular part longer than hyaline part, which comprises 28% to 39% of tail length; cellular-hyaline junction irregular. Phasmids located at 34% to 43% of tail length.

Life cycle
Steinernema africanum n. sp. readily infests and develops in G. mellonella larvae. However, the development of S. africanum is unusually slow at

Nematode molecular characterization and phylogenetic relationships
Phylogenetic reconstructions based on the nucleotide sequences of the D2-D3 expansion segments of the 28S rRNA, the ITS region of the rRNA, the mitochondrial 12S rRNA, and the COI show that S. africanum n. sp. belongs to the "feltiae-clade" (Fig. 8). Based on sequence similarities, S. africanum n. sp. is closely related to S. citrae, S. ichnusae, S. litorale, S. nguyeni, and S. weiseri (Figs. S1 and S2 in Supplementary Material). These species share between  Table S1 in Supplementary Material. 92.4% and 95.9% and differ in 24 to 58 nucleotides with S. africanum n. sp. in the ITS sequences flanked by primers 18S and 26S (Fig. S1 in Supplementary Material). Less sequence similarities were observed between S. africanum n. sp. and all the other species of the "feltiae-clade," supporting its novel taxonomic status (Figs. S1 and S2 in Supplementary Material).

Symbiotic relationships
Phylogenetic reconstructions based on whole genome sequences show that the bacterial symbiont isolated from S. africanum n. sp. RW14-M-C2b-1, named here XENO-1, is closely related to X. bovienii T228 T (Fig. 9). The dDDH value between X. bovienii T228 T and XENO-1 is 71.2%, suggesting that the symbiont of S. africanum n. sp. represents a novel subspecies within the X. bovienii species (Fig. S3 in Supplementary Material). This subspecies will be formally described elsewhere.