Morphological and molecular characterization of Butlerius butleri Goodey, 1929 (Nematoda: Diplogastridae) from South Africa: First report

Soil samples were collected from two home garden compost heaps (garden 1/population 1: 26°42′25.3″S 27°06′25.9″E; garden 2/population 2: 26°41′17.4″S 27°06′05.6″E) in Potchefstroom, North-West Province South Africa. Following, the samples were transported in cooler boxes to the North-West University and stored at 6 to 8°C until further processing. Nematodes were extracted from soil samples using the adapted decanting and sieving followed by the sugar centrifugal-flotation method (Marais et al., 2017). The extracted nematodes were fixed in a heated 4% formaldehyde plus 1% propionic acid (FPG) solution, dehydrated in a glycerin solution following De Grisse (1969) and mounted in glycerin on glass microscope slides.

Measurements and drawings of the mounted specimens were done with a Zeiss Axio Imager A2 equipped with an Axiocam ERc5s camera and iPad with the Labscope imaging application as a drawing tube. All measurements and identifications were done at 1,000 × magnification. Curved structures were measured along the median line. All measurements in the descriptions are given in micrometers (µm) and in the form: mean ± standard deviation (range). All specimens were deposited in the National Collection of Nematodes (NCN), Biosystematics, Agricultural Research Council – Plant Health and Protection (ARC-PHP), Pretoria.

One specimen from each population was transferred into an Eppendorf tube containing 15 µl of ddH₂O and DNA was extracted from each population using the chelex method (Musapa et al. 2013) modified by Rashidifard et al. (2019) as follow: 20 μl chelex-100 (5% w/v) and 5 μl proteinase K (20 mg ml⁻¹) were added to each tube containing the nematodes, the tubes were vortexed (15 sec) and centrifuged at 8,000 rpm for 10 sec. Finally, the tubes were incubated at 56°C for 2 hr followed by incubation at 95°C for 10 min before they were stored at −20°C. The following DNA markers were used to amplify the small and large sub units (SSU and LSU) genes, respectively: SSU F04 (5′-GCTTGTCTCAAAGATTAAGCC-3′), SSU R26 (5′-CATTCTTGGCAAATGCTTTCG-3′) (Blaxter et al., 1998), and D2A (5′-ACAAGTACCGTGAGGGAAAGTTG-3′), D3B (5′-TCGGAAGGAACCAGCTACTA-3′) (Subbotin et al., 2006). Following DNA extraction, the polymerase chain reaction (PCR) was carried out using an Eppendorf Mastercycler gradient thermal cycler (Eppendorf, Hamburg, Germany); more details are provided in Table 1. The amplification tube contained 12.5 μl master mix (Promega Corporation, USA), 1 μl of each of the primers (i.e. forward and reverse), 5 μl DNA, and 5.5 μl ddH₂O.

Four microliters of PCR product were loaded on a 1% agarose gel to check the DNA quality. The DNA was stained using GelRed and then visualized under a UV transilluminator. The PCR product was stored at −20°C before sequencing by inqaba biotec™, South Africa (www.inqaba-southafrica.co.za).

Selection of the appropriate taxa for SSU phylogenetic analysis was done according to Shokoohi et al. (2015). For the LSU analysis the available sequences for Diplogastridae, as well as the outgroups, were obtained from NCBI GenBank. The sequences of selected taxa for each gene were aligned using the MUSCLE tool (Edgar, 2004) implemented in Geneious Prime® 2019.2.1 (https://www.geneious.com). Post-editing for each alignment was done using Gblock program (version 0.91b) (http://phylogeny.lirmm.fr/phylo_cgi/one_task.cgi?task_type=gblocks) with all three less stringent parameters. The jModelTest program 2.1.10 (Darriba et al., 2012) was used to identify the best nucleotide substitution model. General time reversible model with a Gamma distribution (GTR + G) was selected as the most appropriate model for SSU and D2-D3 LSU analyses. Bayesian inference (BI) was performed using MrBayes v3.1.2 (Ronquist and Huelsenbeck, 2003) Geneious Prime® 2019.2.1 and the chains were running for 2 × 10⁶ generations for both the SSU and LSU datasets. After discarding a 25% burn-in sample to estimate the posterior probabilities of the phylogenetic trees, the Markov Chain Monte Carlo (MCMC) algorithm was used (Larget and Simon, 1999) to estimate the posterior probabilities (PI) with the 50% majority rule. One population of Odontopharynx longicaudata De Man, 1912 was selected as outgroup for each dataset.

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Measurements: Table 2.

Female (n = 10): Body 1,166 to 1,423 µm long, slender, tapering posterior to anus. Cuticle with evenly spaced punctations. Lateral field marked by three lines. Labial region continuous with body contour, lips low, fused each with a setae 8 to 9 µm long. Stoma 23 to 28 µm long, 9 to 13 µm wide. Cheilostom barrel shaped, 11 to 15 µm long, cheilorhabdions anteriorly arched inwards. Gymnostom 6 to 9 µm long, connected to cheilostom by hyaline ligament. Stegostom anisomorphc, dorsal metastegostom bearing a prominent 7 to 10 µm long dorsal tooth, subventrally a smaller 4 to 6 µm sickle-shaped tooth. Dorsal pharyngeal gland opening on dorsal tooth. Pharynx diplogasteroid, 241 to 269 µm long. Corpus (procorpus + metacorpus) muscular, cylindrical, 133 to 147 µm long. Lumen of corpus heavily sclerotized. Metacorpus swollen, distinguished from procorpus by slightly more sclerotized lumen. Nerve ring situated at isthmus. Excretory pore situated posterior to nerve ring. Postcorpus (isthmus + basal bulb) 107 to 126 µm long, gradually expanding posteriorly. Cardia prominent, 8 to 12 µm long. Reproductive system didelphic, amphidelphic, both branches equal in length. Ovaries reflexed, oocytes arranged in single row except for multiple rows in growth zones. Spermatheca not well demarcated, uterus muscular, inner wall sclerotized dorsally. Four large glands opening into proximal part of uterus. Vagina narrow, 10 to 16 µm long, one-third of the corresponding body width. Vulva situated at 44 to 47% of body length, pore-like opening, not protruding. Rectum 28 to 37 µm long. Tail 225 to 325 µm long, filiform (broken in many specimens). Phasmid prominent, 28 to 31 µm posterior to anus.

Male (n = 12): similar to female, slightly smaller in size (1,082-1,399 µm). Reproductive system monorchid, testis 392 to 703 µm long, reflexed. Prominent sphincter present in mid-region of vas deferens. Spicules 36 to 43 µm long, slender, and curved, capitulum anteriorly flattened, circular margins, lamina narrow, ventrally curved. Gubernaculum 23 to 31 µm long, proximal part narrow, curved, handle-like, distal part rounded with a sleeve, distal tip bearing small, laterally directed, hook-like processes. Nine pairs of genital papillae, three pre-cloacal and six post-cloacal, formula: v1, v2, v3d/v4, ad, ph, v5, 6, 7, pd. The v5, 6, 7 clusters greatly separated, left subventral group at level of phasmid, right subventral group at level of posterior dorsal papilla. Anterior cloacal lip large, lobe-like, protruding posteriorly, bearing single prominent papilla, posterior lip bearing single pair of smaller papillae. Tail similar to female, 242 to 356 µm long, filiform.

The Nucleotide BLAST (Blastn) search based on the partial SSU sequences (MN710517 and MN710518) showed > 99% similarity to a B. butleri from Iran (accession no. KP453998). Blastn search using the partial LSU sequences of our populations (MN710521 and MN71522) revealed 92% similarity to a Diplogastrellus sp. from Germany (accession no. KJ877248).

Bayesian topology based on partial SSU sequences confirmed the well-supported sister relationship (PB: 0.99) of South African populations (MN710517 and MN710518) with another population of B. butleri from Iran (KP453998) (Fig. 4). Moreover, these populations formed a highly supported group (BP: 0.93) with two unidentified populations of Diplogastrellus Paramonov, 1952 (KJ77205 and AB597239), two Pseudodiplogasteroides Körner, 1954 populations (AB597237 and AB597238) and an unidentified population of Butlerius (KJ877204). Bayesian inference tree using the D2-D3 LSU sequences showed that two newly sequenced populations of B. butleri are closely related and clustered (BP: 0.84) with two populations of Pseudodiplogasteroides (AB597248 and AB597249) and an unidentified population of Rhabditidoides Rahm, 1928 (AB597251) (Fig. 5).

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