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

Steinernema africanum n. sp. RW14-M-C2b-1 and RW14-M-C2a-3 nematodes were isolated from soils of a banana, pumpkin, and sorghum intercrop in a valley of the Republic of Rwanda (GPS coordinates: 1°28´11.1"S 29°41´36.2"E; 1,865 m. s. n. m.) (Yan et al., 2016). Nematode isolation was achieved by baiting mixed soil samples with Galleria mellonella (Lepidoptera: Pyralidae) larvae, and placing the infected larvae in White traps (White, 1927).

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 ddH₂O) 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 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) (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 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, 2004; Tarasco et al., 2008; Půža et al., 2021).

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 (Tris– boric 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).

The entomopathogenic Xenorhabdus bacteria associated with S. africanum n. sp. RW14-M-C2b-1 nematodes were isolated as described (Machado et al., 2018, 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, 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-to-genome distance calculator (GGDC) web service of the Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH (DSMZ) (Auch et al., 2010a, 2010b; Meier-Kolthoff et al., 2013, 2014).

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.

(Figures 1–7 and Tables 1–4)

Figure 1

Body slender, ventrally curved posteriorly, C- or J-shaped when heat-killed. Cuticle with transversal incisures scarcely marked, with annuli appearing slightly visible. Lateral fields and phasmids inconspicuous under LM. Lip region truncate to slightly round, continuous with body. Six lips amalgamated, with one acute labial papilla and one low cephalic papilla each, except lateral lips. Amphidial apertures small, located at lateral lips posterior to lateral labial papillae. Stoma shallow, funnel-shaped, 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 non-valvate 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 sub-ventral 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.

Figure 2

Figure 3

Figure 4

Figure 5

Figure 6

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.

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 metacorpus-isthmus junction. Nerve ring surrounding the isthmus. Reproductive system didelphic, amphidelphic. Ovaries reflexed in dorsal position; oviducts well developed with glandular spermatheca, and uteri tubular with numerous uterine eggs; vagina short, with muscular walls; vulva protruding, in the form of transverse slit located slightly post-equatorial with lips slightly protruding, asymmetrical, with small epiptygma. Rectum 0.6 to 1.1 times the BD, with three rectal glands. Tail conoid, shorter than body anal diameter, with terminus bearing a fine mucron. Phasmids located at anterior part of tail, at 23% to 34% of tail length.

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.

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.

Steinernema africanum n. sp. readily infests and develops in G. mellonella larvae. However, the development of S. africanum is unusually slow at 24°C compared to several other Steinernema species including S. feltiae, S. weiseri, S. ischunanense, S. litorale, S. surkhetense, S. hermaphroditum, S. akhursti, S. cholashanense, and S. xueshanense. We typically observe that G. mellonella larvae infested with 50 to 150 IJs of the above-mentioned species die within 2 d to 3 d, but insects take 5 d to 6 d to die when infested with S. africanum. Nematode adults of the first and second generations are found in insect cadavers within 5 d to 6 d and 8 d to 9 d, respectively, when infested by the above-mentioned species, while they are found after 8 d to 9 d and 11 d to 12 d, respectively, when infested with S. africanum. Pre-IJs emerge from insect cadavers after 12 d to 15 d upon infestation by the above-mentioned species, but only after 18 d to 21 d when infested by S. africanum.

The type hosts are unknown as the nematodes of this genus can be hosted by different insect species (Yan et al., 2016; Kajuga et al., 2018; Fallet et al., 2022) and were isolated from mixed soil samples by the Galleria baiting technique (White, 1927; Bedding and Akhurst, 1975). Briefly, S. africanum n. sp. RW14-M-C2b-1 and RW14-M-C2a-3 nematodes were isolated, using the “Galleria baiting” method, from soil samples collected in a banana, pumpkin, and sorghum intercrop in a valley in the Republic of Rwanda (GPS coordinates: 1°28’11.1”S 29°41’36.2”E; 1,865 m. s.

n. m.) (Yan et al., 2016). Cultures of this species are maintained in the Institute of Biology, University of Neuchatel (Switzerland), in the Rwanda Agriculture and Animal Resource Development Board (Rubona, Rwanda), and in CABI Swiss laboratories in Hungary.

RW14-M-C2b-1 nematodes are the type material for S. africanum n. sp. Three slides of each stage, including first-generation adults (males and females), second-generation adults (males and females), and IJs, were deposited in the Nematology Collection of the Aquaculture Research Unit of the University of Limpopo, South Africa with the accession numbers ULRS-N1 to ULRS-N15. Additional specimens were deposited at the nematode collection of the Department of Animal Biology, Plant Biology and Ecology of the University of Jaén, Spain with the following accession numbers: RWA004-01 to RWA004-20 and RWA005-01 to RWA005-15. Slides will be made available upon reasonable request. Nematode cultures are maintained in the Institute of Biology, University of Neuchatel, Switzerland and in the Rwanda Agriculture and Animal Resource Development Board, Rubona, Rwanda.

The specific name refers to the continent where the species was isolated.

No progeny was observed when S. africanum n. sp. (isolates RW14-M-C2b-1 or RW14-M-C2a-3 nematodes) were allowed to interact with specimens of S. feltiae, S. ichnusae, S. litorale, or S. weiseri. No progeny was observed in the single-female control plates. When S. africanum n. sp. (isolates RW14-M-C2b-1 or RW14-M-C2a-3 nematodes) were allowed to interact, fertile progeny was observed. Fertile progeny was also observed when all nematode strains were self-fertilized. Hence, S. africanum n. sp. (RW14-M-C2b-1 or RW14-M-C2a-3) are conspecific and reproductively isolated from closely related species such as S. feltiae, S. ichnusae, S. litorale, and S. weiseri.

Steinernema africanum n. sp. adults have short stoma with rounded cheilorhabdia, pharynx robust with rounded basal bulb; males monorchid with ventrally curved spicules having lanceolate manubrium in the first generation and rounded manubrium in the second generation, gubernaculum fusiform in the first generation and anteriorly hook-like in the second generation, tail conoid and slightly ventrally curved with fine mucron in the first generation (34–46 mm, c = 25–34, c´ = 0.9–1.1, mucron = 4.0–5.3 mm) and with more robust mucron in the second generation (28– 46 mm, c = 26–36, c´ = 0.7–1.1, mucron = 5–10 mm); females didelphic–amphidelphic with shorter conoid tail bearing a fine mucron in the first generation (35–55 mm, c = 51–104, c´ = 0.7–1.0, mucron =3.7– 5.0 mm) and longer conoid, tail-lacking mucron in the second generation (40–60 mm, c = 17–51, c´ = 1.2– 2.3); and IJs with short body (0.69–0.80 mm), poorly developed pharynx (132–153 mm), H% (28–39), D% (79–105), and E% (135–290), lateral fields with eight longitudinal wings, and tail conoid-elongate (52– 72 mm, c = 10–15, c´ = 2.9–4.2).

Based on morphological and morphometric traits, S. africanum n. sp. belongs to the “feltiae-clade.” Nematodes of this group are characterized by having third-stage IJs between 700 mm and 1,000 mm long. The lateral fields of IJs in the “feltiae-clade” are characterized by eight ridges arranged evenly in the mid-body region. Steinernema africanum n. sp., a member of the “feltiaeclade,” presents several traits common to this group. Specifically, the IJs have a large body size (690–802 mm), and they have eight ridges in the mid-body region of the lateral field. Several of the morphometric traits of the IJs overlap with those of other species in the “feltiae-clade” (Tables 2 and 3).

Steinernema africanum n. sp. IJs and first-generation adults are morphologically similar to S. citrae, S. feltiae, S. ichnusae, S. litorale, S. nguyeni, S. weiseri, S. jollieti, and S. puntauvense. The IJs of S. africanum n. sp. can be distinguished from the IJs of these latest species and other species of the “feltiae-clade” because the position of the NR is more posterior in S. africanum n. sp. and the stoma and pharynx regions are longer. In addition, S. africanum n. sp. IJs differ from S. citrae IJs in hyaline region occupying posterior (28%–39% vs 37%–50%) of tail length, hemizonid (present vs not observed), and cardia (conoid vs inconspicuous). Steinernema africanum n. sp., S. feltiae, and S. ichnusae IJs differ in tail length (52–72 μm vs 81–8 μm vs 76–89 μm), c´ (2.9–4.2 vs 4.5–5.1 vs 4.2–5.1), D% (34–46 vs 44–50 vs 42–49), hyaline region comprising (28–39% vs 37–51% vs 44–50%) of tail length, and E% (79–129 vs 67–81 vs 68–83). Steinernema africanum n. sp. and S. litorale IJs differ in body length (690–802 μm vs 834–988 μm), tail length (52–72 μm vs 72– 91 μm), b ratio (4.3–6.3 vs 6.7–7.9), and location of hemizonid (between NR and pharynx base vs basal bulb). Steinernema africanum n. sp. and S. nguyeni IJs differ in b ratio (4.3–6.3 vs 6.2–7.4), neck length (NL) (123–167 μm vs 101–121 μm), and hyaline region comprising (28–39% vs 20–31%) of tail length. Steinernema africanum n. sp. and S. weiseri IJs differ in deirids (inconspicuous vs located in center of lateral fields at level of pharyngeal bulb or slightly posterior), location of hemizonid (between NR and pharynx base vs at level of basal bulb or immediately posterior), hyaline region comprising (28–39% vs 18–24%) of tail length, and D% (34–46 vs 44–55). Steinernema africanum n. sp. and S. jollieti IJs differ in the number of structures of the lateral field at mid-body (eight vs six), D% (34–46 vs 46–50), and hyaline region comprising (28–39% vs 46–60%) of tail length. Steinernema africanum n. sp. and S. puntauvense IJs differ in hemizonid (present vs not observed), hyaline region comprising (28–39% vs 52–55%) of tail length, BD (25–32 μm vs 31–38 μm), distance from anterior region to EP (54–68 μm vs 20–30 μm), distance from anterior region to NR (87–132 μm vs 46–69 μm), a ratio (23–30 vs 17–23), and D% (79–129 vs 35–56) (Table 2).

First-generation males of S. africanum n. sp. differ from the males of S. citrae in lateral field (inconspicuous vs present in mid-body, with one narrow ridge) and tail length (34–46 μm vs 17–31 mm); from the males of S. feltiae in body length (0.98– 1.40 mm vs 1.41–1.81 mm), the position of the EP (69–109 μm vs 110–126 μm), NL (132–147 μm vs 164–180 μm), and SW% (144–197 vs 99–130); from the males of S. ichnusae in the presence of prominent mucron vs absent, spicule manubrium rhomboidal vs always oblongate, body size (0.98– 1.40mm vs 1.15–1.49 mm), and a ratio (9–12 vs 20–29); from the males of S. litorale in shape of spicule manubrium (rhomboidal vs oval to somewhat angular, rectangular), c (25–34 vs 33–56), D% (52–74 vs 34– 56), and GS% (49–68 vs 62–81); from the males of S. nguyeni in presence of amphidial apertures (small vs inconspicuous), lateral field inconspicuous vs with one ridge, the position of the EP (69–109 μm vs 47–71 μm), tail length (34–46 μm vs 18–25 μm), c (25–34 vs 38–53), and D% (52–74 vs 38–57); from the males of S. weiseri in the presence of mucron vs absent, spicule manubrium rhomboidal vs distinctly elongated, tail length (34–46 μm vs 19–32 μm), gubernaculum length (GL) (32–49 μm vs 46–57 μm), c (25–34 vs 36–64), and GS% (49–68 vs 70–85); from the males of S. jollieti in the presence of prominent mucron vs absent, shape of spicule manubrium (rhomboidal vs elongated), shape of gubernaculum (fusiform vs boat-shaped), deirids inconspicuous vs located posterior to level of pharyngo-intestinal junction, and a ratio (9–12 vs 12–19); from the males of S. jollieti in the distance of anterior end NR (79–104 μm vs 104– 128 μm), presence of spicule calomus (narrower vs inconspicuous), lamina with rostrum or retinaculum (present vs absent), and velum (scarcely developed velum not reaching the spicule tip vs extending from rostrum to spicule terminus) (Table 3).

Figure 7

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 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).

Figure 8

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.

Figure 9