First molecular observation on Mylonchulus hawaiiensis from South Africa

Samples from Magoebaskloof mountain (GPS coordinates: 23°51′23.0″S 29°57′26.7″E), in Limpopo Province, South Africa, were collected in March 2022 (Fig. 1). Nematode extraction was achieved using the Whitehead and Hemming tray method (Whitehead & Hemming, 1965; Shokoohi, 2022). Extracted individuals were fixed with a hot 4 % formaldehyde solution (except those specimens used for molecular analyses), transferred to anhydrous glycerin utilizing the method of De Grisse (1969), and mounted on permanent glass slides. The glass slide with a paraffin ring in the middle was used to fix the nematodes permanently. Then, the ring was covered by a coverslip, heated up, and left to get solid. The specimens for molecular analyses were freshly taken and transferred to the PCR tube. The specimen processing for molecular study, morphometrics, and microscopic morphological studies was done at the Aquaculture Research Unit (ARU) of the University of Limpopo. No stain was used for morphological observations.

Fig. 1.

Measurements of specimens mounted on permanent slides were taken, and de Man’s (1881) indices were calculated. Drawings were made using a camera attached to a Zeiss microscope (Axio Lab, A series; Germany) at the Aquaculture Research Unit, University of Limpopo. Micrographs Pictures were taken under a Nikon Eclipse 80i light microscope provided with differential interference contrast optics (DIC) and a Nikon Digital Sight DS-U1 camera (Nikon, Tokyo, Japan). Micrographs were edited using Adobe® Photoshop® CS.

To evaluate the morphological variations between the populations of M. hawaiiensis, a principal component analyses (PCA) with different morphological traits were conducted. PCA analyses were carried out in XLSTAT (Addinsoft, 2007). Various morphometric features were obtained from fixed nematodes, and the available literature, including an average of body length, a, b, c, c′, V, lip region width, buccal cavity length, buccal cavity width, dorsal tooth apex, and tail length were included in the PCA analyses.

Before their analysis, the measures were normalized by the Log₁₀ as used for morphometric data provided by Nattero et al. (2017) using XLSTAT software (Addinsoft, 2007). The scores values were determined for each species based on each of the principal components, and the scores for the first two components were used to form a two-dimensional plot (F1 and F2) of each isolate based on the eigenvalues given by the software XLSTAT.

DNA extraction was done using the Chelex method (Shokoohi et al., 2023). Three specimens of M. hawaiiensis were hand-picked with a fine-tip needle and transferred to a 1.5 ml Eppendorf tube containing 10 μl double distilled water. The nematodes in the tube were crushed with the tip of a fine needle and vortexed. Thirty microliters of 5 % Chelex® 50 and 2 μL of proteinase K were mixed into the microcentrifuge tube containing the crushed nematodes. The microcentrifuge tube with the nematode lysate was incubated at 56 °C for two hours and then set at 95 °C for 10 minutes to deactivate the proteinase K and finally spun for 2 min at 16000 rpm (Shokoohi, 2022). The supernatant was extracted from the tube and stored at −20 °C. Following this step, the forward and reverse primers, SSU R26 (5′-CATTCTTGGCAAATGCTTTCG–3′); 18s (5′-TTGATTACGTCCCTGCCCTTT–3′) and 26s (5′-TTTCACTCGCCGTTACTAAGG–3′) (Vrain et al., 1992; Blaxter et al., 1998), were used in the PCR reactions for partial amplification of the 18S, and 28S rDNA regions, respectively. PCR was conducted with eight μl of the DNA template, 12.5 μl of 2X PCR Master Mix Red (NEB, UK), one μl of each primer (10 pmol μl⁻¹), and ddH₂O for a final volume of 30 μl. The amplification was processed using an Eppendorf master cycler gradient (Bio-Rad, USA), with the following program: initial denaturation for 3 min at 94 °C, 37 cycles of denaturation for 45 s at 94°C; 54 °C, and 56 °C annealing temperatures for 18S and ITS rDNA, respectively; extension for 45 s to 1 min at 72 °C, and finally an extension step of 6 min at 72 °C followed by a temperature on hold at four 4 °C. After DNA amplification, 4 μl of product from each tube was loaded on a 1 % agarose gel in TBE buffer (40 mM Tris, 40 mM boric acid, and one mM EDTA) for evaluation of the DNA bands. The bands were stained with safe view classic (Applied Biological Materials Inc. (abm), Richmond, Canada) and visualized and photographed on a UV transilluminator. The amplicons of each gene were stored at −20 °C. Finally, Inqaba Biotech (South Africa) purified the PCR products for sequencing. Also, as outgroups, Bathyodontus mirus Andrássy, 1956 (FJ969116), and B. cylindricus Fielding, 1950 (AY552964) were used for 18S rDNA trees. Besides, Mermis nigrescens Dujardin, 1842 (KF886021) was used for ITS rDNA outgroup. The ribosomal DNA sequences were analyzed and edited with BioEdit (Hall, 1999) and aligned using CLUSTAL W (Thompson et al., 1994). Phylogenetic trees were generated using the Bayesian inference method as implemented in the program MrBayes 3.1.2 (Ronquist & Huelsenbeck, 2003). The GTR+I+G model was selected for the 18S, and ITS rDNA trees using jModeltest 2.1.10 (Guindon & Gascuel, 2003; Darriba et al., 2012). The selected model was then initiated with a random starting tree and ran with the Markov chain Monte Carlo (MCMC) for 10⁶ generations. The new partial 18S rDNA (OR035485), and ITS rDNA (OR035486) were deposited in GenBank.

To understand the relationship between the different populations of M. hawaiiensis, a principal component analysis was performed using morphometric features of the females (Fig. 3). The analyzed morphological characters allowed a clear separation between the populations of M. hawaiiensis. An accumulated variability of 49.44 % was observed in female-based PCA, specifically, 27.26 % in the F1 and 22.18 % in the F2. Body length (r = 0.661) and b (r = 0.783), and tail length (r = 0.842) displayed a significant coefficient correlation with F1 (Fig. 3; Table 3).

Fig. 3.

The result indicated that South African M. hawaiiensi grouped close to Hawaiian population in the PCA. In contrast, the result also showed a variation among the populations of M. hawaiiensis. In addition, the present population of M. hawaiiensis stands separate from the previous population of the same species reported from South Africa (Fig. 3; Table 4).

A molecular study of nblast of 18S rDNA region of M. hawaiiensis indicated a 100 % similarity between the South African population and the Japanese population of M. hawaiiensis (AB361438; AB361439; AB361440; AB361442). However, the first time sequenced ITS rDNA of M. hawaiiensis worldwide, and therefore, its comparison with the same species or another species belonging to Mylonchulus is not possible. The phylogenetic analysis of 18S rDNA (Fig. 4) showed a close relation of M. hawaiensis with 0.97 posterior probability. The phylogenetic analysis of ITS rDNA (Fig. 5) placed South African M. hawaiiensis (OR035486) close to Prionchulus oleksandri Winiszewska and Susulovsky, 2003 with 1.00 posterior probability. In addition, network analysis (Fig. 6) based on Median Neighbor Joining showed a variation within M. hawaiiensis. The result indicated that South African M. hawaiiensis is closer to the Japanese population of the same species.

Fig. 4.

Fig. 5.

Fig. 6.