First Reports and Morphological and Molecular Characterization of Pratylenchus delattrei and Quinisulcius capitatus Associated with Chickpea in Ethiopia

Soil and root samples were collected from chickpea growing areas in Minjar, Adea’a, and Mesekan districts during the 2021 main growing season, located in central and southern parts of Ethiopia. Details regarding sample locality, altitude, GPS coordinators, and GenBank accession numbers are summarized in Table 1. From each sampling locality, 20 soil cores were taken in a zig-zag pattern from within the top 30 cm using a 3 cm diameter tube from the chickpea rhizosphere, mixed to obtain a 500 g soil sample. For each sample, 80 chickpea roots were collected and put in labelled plastic bags. Subsequently, both soil and root samples were taken to the Plant Disease Diagnostics laboratory at Jimma University and stored at 4°C until nematode extraction (Barker et al., 1969). The nematodes were extracted from aliquots of 100 ml of soil and 10 g of roots by the modified Baermann tray method described by Hooper et al. (2005).

Morphological and morphometric data were recorded from both temporary and permanent slides. In order to link molecular data with morphological vouchers of individual nematodes, live nematodes were heat relaxed by quickly passing them over a flame and examined, photographed, and measured using an Olympus BX51 DIC Microscope (Olympus Optical, Tokyo, Japan), equipped with an HD Ultra camera. Subsequently, each specimen was recovered from the temporary slide for genomic DNA extraction. For permanent slides, the nematode suspensions were concentrated in a drop of water in an embryo glass dish, with a few drops of fixative (4% formalin, 1% glycerol (in water) in it. The nematodes were immediately heated in a microwave (700 watts) for about 4 sec and left at room temperature for 1 h at 4°C for 24 h. This was followed by gradually transferring to anhydrous glycerin, ready to be mounted on glass slides as described by Seinhorst (1959). Specimens for scanning electron microscopy (SEM) were fixed in Trump's fixative, washed in 0.1 M-phosphate buffer (pH = 7.5), dehydrated in a graded series of ethanol solutions, critical point dried with liquid CO₂ and mounted on stubs with carbon tabs (double conductive tapes), coated with 25 nm gold, and photographed with a JSM-840 EM (JEOL) at 12 kV (Singh et al., 2021).

After making morphological vouchers, nematodes were recovered from temporary slides, washed with distilled water, cut into 2–3 pieces, and transferred to a PCR tube containing 20 μL of worm lysis buffer (WLB) (50 mM KCl;10 mM Tris pH 8.3; 2.5 mM MgCl2; 0.45% NP-40 (Tergitol Sigma); 0.45% Tween-20). Then, the samples were incubated at −20°C for 10 min, followed by adding 1 μL proteinase K (1.2 mg/ml) and incubation for 1 h at 65°C and 10 min at 95°C and centrifugation for 1 min at 14,000 rpm. Finally, the samples were stored at −20°C until used for the PCR, as previously described by Singh et al. (2019) and Nguyen et al. (2019). A DNA template of 3 μL was transferred to an Eppendorf tube containing 23.5 μL master mix containing 10 μL of PCR water, 12.5 μL Dream tag, and 0.5 μL of each primer (Derycke et al., 2010) and PCR amplification was performed using a Bio-Rad T100™ thermocycler. PCR amplifications of the D2-D3 region of 28S-rDNA were performed using the forward primer D2A (5′-ACA AGT ACC GTG AGG GAA AGT TG-3′), and reverse primer D3B (5′-TCG GAA GGA ACC AGC TAC TA-3′) (Subbotin et al., 2006). For ITS rDNA, the forward primer Vrain2F (5′-CTT TGT ACA CAC CGC CCG TCG CT-3′), and reverse primer Vrain2R (5′-TTT CAC TCG CCG TTA CTA AGG GAA TC-3′), were used following the protocol of Vrain et al. (1992) with the touch-down thermal profiles described by Singh et al. (2019). For the amplification of the cytochrome oxidase subunit 1 (COI) gene of mitochondrial DNA, the primer JB3 (5′-TTT TTT GGG CAT CCT GAA GTC TAT-3′) and JB4.5 (5′-CCT ATT CTT AAA ACA TAA TGA AAA TG-3′) and the primer JB3Prat (5′-TTT TTT GGG CAT CCT GAA GTC TAT-3′) and JB4Prat (5′-CCT ATT CTT AAA ACA TAA TGA AAA TG-3′) were used following the protocol of Bowles et al. (1992) with the thermal profile described in the study of Singh et al. (2019). All the PCR products were checked by gel electrophoresis stained with GelRed (Biotium) and visualized under UV light illumination. The successful PCR reactions were purified and sent to Macrogen (https://dna.macrogen.com, Europe) for sequencing. Consensus sequences were assembled in forward and reverse directions using Geneious 2022.1 (Biomatters; http://www.geneious.com) and deposited in the NCBI GenBank (Table 1).

Resulting sequences were compared with other relevant sequences available in the GenBank. Multiple alignments of the different DNA sequences were made using MUSCLE with default parameters, followed by manual trimming of the poorly aligned ends using Geneious 2022.1. Phylogenetic trees were created by using MrBayes 3.2.6, adding Geneious with the GTR + I + G model. The Markov chains for generating phylogenetic trees were set at 1 × 106 generations, four runs, 20% burn-in and sub-sampling frequency of 500 generations (Huelsenbeck and Ronquist, 2001).

See Table 2.

Vermiform and slightly curved ventrally after heat-killing and fixation. Labial region continuous from the rest of the body and lip region with three annuli. Under SEM (Figs. 1 C,D), en face view showing an oval oral aperture surrounded by six inner labial sensilla, submedian segments fused to oral disc, corresponding to head pattern group 2 according to Corbett and Clark (1983). Stylet was well developed (16–18 μm long) with anteriorly directed rounded knobs. The areolation was only visible at tail level and lateral field with four incisures, with the outer two being entirely crenate, and the inner lines being finely striated. Rounded to oval-shaped metacorpus with short isthmus, pharyngeal gland overlapped ventrally. Excretory pore ws located slightly above pharyngo-intestinal junction. There was a vulva, a transverse slit in ventral view, and well developed post-vulval uterine sac. The tail had (27–30) annuli, subcylindrical, and with rounded to conical, smooth terminus.

Vouchers (two females) are available in the UGent Nematode Collection (slide UGnem-314) of the Nematology Research Unit, Department of Biology, Ghent University, Ghent, Belgium.

Pratylenchus delattrei Four sequences of the D2-D3 28S rDNA (OP646167-OP646170; 571-619 bp; 1–3 bp differences), two ITS rDNA sequences (OP646172-OP646171; 731bp; 17 bp differences) and two COI sequences (OP730535-OP7330534; 421 bp; with 100% identical) were generated for P. delattrei from Minjar and Adea’a districts (Table 1; Figs. 3 A–C). Based on the D2–D3 sequences, isolates formed the highest supported clade with P. delattrei sequences from Cape Verde (KY677820) and two sequences from Iran (JX261949 and JX261948), which are 99.6–100% identical. For ITS, the P. delattrei sequences formed a maximally supported clade with three unidentified Pratylenchus sp. sequences from India (MN100134, MN100135 and MH375058) with 94–97% similarity (Fig. 3B). Finally, two identical COI sequences have been generated for the first time for P. delattrei, and these sequences were in a poorly supported sister relationship (0.68 PP) with P. parazeae (Fig. 3C).

Male nematodes were not found. As first reported on chickpea and from Ethiopia, this species was recovered from the Minjar and Adea’a districts in the central parts of Ethiopia, both in the rhizosphere and the roots (Table 1). It has been also reported in other African countries, including Madagascar (cotton), Sudan (sugarcane), and Cape Verde (tomato), and from several Asian countries: South Korea, Pakistan, Oman, Iran (on tomato and eggplant, date palm, pigeon pea and peanut, and medicinal plants) (Luc, 1958; Sharma et al., 1992; Jothi et al., 2004; Mani et al., 2005; Castillo & Vovlas, 2007; MajdTaheri et al., 2013; Flis et al., 2018). The studied female morphology and morphometrics are in agreement with the original description (Luc, 1958), and other descriptions of P. delattrei from Iran and Cape Verde (MajdTaheri et al., 2013; Flis et al., 2018). The characteristics also agree without variation with the matrix code for the tabular key of Castillo and Vovlas (2007): A2 (three labial annuli), B1 (male absent), C3 (stylet length 16–17 μm), D1 (shape of spermatheca absent or reduced), E2 (V ratio = 75–79.9%), F3 (PUS: 20–24.9 μm), G3 (conoid tail shape), H1 (smooth tail tip), I1 (<30 μm pharyngeal overlapping length), J1 (four lateral field lines), K1 (smooth bands of lateral field structures), and a subcylindrical tail shape with a conical to rounded tail tip. The Ethiopian P. delattrei has a slightly longer tail compared to populations from Iran and Cape Verde (27.5–30.1 vs. 21–29 μm); however, the tail length was not included in the original description. In the phylogenetic tree of the ITS region, the Ethiopian P. delattrei sequences formed a maximally supported clade with three unidentified Pratylenchus species from India; these may therefore also represent P. delattrei based on the relatively limited molecular variability (26–49 bp difference). This study links for the first time ITS sequences to P. delattrei.

(Fig. 2)

Figure 2:

See Table 3.

The studied specimens are morphologically and morphometrically similar to the original description (Allen, 1955) and subsequent descriptions by Mekete et al. (2008), Munawar et al. (2021), and Iqbal et al. (2021), except for the slightly longer stylet compared to populations from coffee in Ethiopia (17.8–19.7 vs. 15–18 μm) and longer tail compared to the Canadian population (41.0–50.0 vs. 31.3–40.4 μm) (Table 3). All of the specimens have five incisures in the lateral field and also conoid, enlarged and striated terminus shape (Fig. 2), agreeing with genus Quinisulcius, which sets it apart from Tylenchorhynchus (5 vs. 3–4) according to the key of Hunt et al. (2012). As in the current study, males have rarely been found (Hopper, 1959; Siddiqi, 1971; Knobloch and Laughlin, 1973; Maqbool, 1982; Mekete et al., 2008; Geraert, 2011). However, Iqbal et al. (2021), described Q. capitatus male specimens from apple, tomato, maize, potato, cabbage, and onion in Pakistan. Quinisulcius capitatus is known to parasitize over 27 plants across all continents (North America, Central and South America, temperate parts of Europe, Africa, Asia, Australia, and New Zealand) (Munawar et al., 2021). In Africa, this species has been reported in Ethiopia from coffee (Mekete et al., 2008), soybean in South Africa (Mbatyoti et al., 2020), and tomato and carrot in Benin (Baimey et al., 2009). The Ethiopian Q. capitatus specimens formed a well-supported clade with the Pakistan populations in our D2–D3 of 28S and ITS rDNA, however, the tree topology to the Canadian Q. capitatus population was not resolved for both gene regions (Figs. 4A,B). This suggests that the Canadian populations may have been mislabelled.

Figure 4: