Rotylenchus wimbii n. sp. (Nematoda: Hoplolaimidae) associated with finger millet in Kenya

A soil sample mixed with some roots was collected using a shovel from 15 to 25 cm soil depth in a zig-zag pattern from a finger millet field [Eleusine coracana (L.) Gaertn.] in Kipkaren Estate, Eldoret, Kenya in February, 2019. The GPS coordinates of the field location are 00°30.395’ N; 035°14.825’E. Nematodes were extracted from 100 ml of soil by using modified Baermann’s method (Whitehead and Hemming, 1965). The extracted nematodes were stored at 4°C during the course of analysis.

Host roots were stained using acid fuchsin (Byrd et al., 1983). For this, clean roots were bleached in 2.5% NaOCl for 5 min, followed by rinsing the roots in running tap water, and boiling them in 30 ml distilled water with 1 ml of stock staining solution (0.35 g acid fuchsin, 25 ml acetic acid, and 75 ml distilled water) in a microwave for 30 sec. After cooling, excess fuchsin was drained and roots were washed with running tap water and de-stained by immersing them in 70% acidified glycerol, and finally observed under a stereo microscope for presence of stained PPN.

For morphological studies, live nematodes were heat-relaxed by quickly passing over a flame in a drop of water on a glass slide until nematode movement stopped and examined, photographed, and measured using an Olympus BX51 DIC Microscope (Olympus Optical, Tokyo, Japan), equipped with an Olympus C5060Wz camera as described in Singh et al. (2018). After recording morphological information, each specimen was recovered from the slide and its genomic DNA was extracted. For fixing, the nematode suspension obtained after extraction was concentrated in a drop of water in a glass embryo dish, followed by addition of a few drops of Trump’s fixative [2% paraformaldehyde, 2.5% glutaraldehyde in 0.1 M Sorenson buffer (Sodium phosphate buffer at pH = 7.5)] into it. The nematodes were then immediately heated in a microwave (700 watts) for about 4 sec and left for 1 hr at room temperature and finally at 4°C for 24 hr. This was followed by gradually transferring the nematodes to anhydrous glycerin, ready to be mounted on glass slides as described in Singh et al. (2018). For SEM, specimens fixed in Trump’s fixative were washed in 0.1 M phosphate buffer (pH = 7.5) and dehydrated in a graded series of ethanol solutions, critical-point-dried with liquid CO₂, 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., 2018).

After morphological analysis, heat-relaxed nematodes were recovered from temporary slides and each individual nematode was cut into pieces in distilled water using a blade and the pieces were transferred to a PCR tube containing 20 µl of worm lysis buffer [50 mM KCl, 10 mM Tris at pH = 8.3, 2.5 mM MgCl₂, 0.45% NP 40 (Tergitol Sigma), 0.45% Tween 20]. The PCR tubes were incubated at −20°C (10 min) followed by addition of 1 µl proteinase K (1.2 mg/ml), incubation at 65°C (1 hr) and 95°C (10 min), and finally centrifuging the mixture at 14,000 rpm for 1 min (Singh et al., 2018). PCR amplifications of the partial ITS, 18S, and D2-D3 expansion segment of 28S of rDNA were done using the primer pairs, Vrain2F: 5´-CTTTGTACACACCGCCCGTCGCT-3´/Vrain2R: 5´-TTTCACTCGCCGTTACTAAGGGAATC-3´ (Vrain et al., 1992), SSU18A: 5´-AAAGATTAAGCCATGCATG-3´/SSU26R: 5´-CATTCTTGGCAAATGCTTTCG-3´ (Mayer et al., 2007) and D2A: 5´-ACAAGTACCGTGAGGGAAAGTTG-3´/D3B: 5´-TCCTCGGAAGGAACCAGCTACTA-3´ (Nunn, 1992), respectively, using thermal profiles described in Singh et al. (2018, 2019). For amplification of the COI region of mtDNA, the primer pair, JB3: 5´-TTTTTTGGGCATCCTGAGGTTTAT-3´/JB4.5: 5´-TAAAGAAAGAACATAATGAAAATG-3´ was used according to Bowles et al. (1992). The PCR products were enzymatically cleaned as mentioned in Singh et al. (2020) and contigs were made from the newly produced forward and backward sequences using Geneious Prime 2020.0.5 (https://www.geneious.com) and deposited in GenBank.

The phylogenetic relationships of the new species with other related species were analyzed based on the partial sequences of ITS, 18S, 28S, and COI. Phylogenetic programs implemented in Geneious Prime 2020.0.5 were used. The obtained consensus contigs were subjected to BLAST search to check for closely related species on GenBank and all the collected sequences for each gene fragment were aligned using MUSCLE alignment of Geneious Prime 2020.0.5 using default parameters, followed by manually trimming of the poorly aligned ends. The best nucleotide substitution model of each gene alignment was determined by jModelTest 2.1.10 and using the selected models, phylogenetic trees were created using Bayesian phylogenetic analyses (MrBayes 3.2.6), which was run under 1 × 10⁶ generations (4 runs) and Markov chains sampled at every 100 generations, and 20% of the converged runs regarded as burn-in (Huelsenbeck and Ronquist, 2001). To check for additional distinctiveness of our sequences in phylogenetic trees, the species delimitation plugin of Geneious Prime 2020.0.5 (Masters et al., 2011) was used to calculate Rosenberg’s PAB, which tests the probability for reciprocal monophyly of sequence clusters (Rosenberg, 2007).

Rotylenchus wimbii n. sp.

Figures 1 and 2, Tables 1 and 2.

Figure 1:

Figure 2:

Rotylenchus wimbii n. sp. is characterized by a moderate body size of 0.6 to 0.8 mm, a hemispherical, continuous lip region with four annuli, basal lip annule with 3 to 4 irregular blocks, absence of longitudinal cuticular striations at anterior region, lateral field with four lines forming three equal bands which are areolated around pharynx level, a robust stylet of less than 30 µm of which 45 to 53% is cone and with rounded to sometimes indented knobs, didelphic-amphidelphic reproductive system, vulva without distinct epiptygma, indistinct to empty spermatheca, very short truncated tail with 5 to 9 annuli, and pore like phasmids located at 7 to 17 annuli anterior to anus. According to Castillo and Vovlas (2005), the matrix code of this species is A4, B1, C1, D4, E1, F2, G3, H (commonly truncated), I2, J2, K1.

This new species is morphologically closest to Rotylenchus abnormecaudatus Van den Berg and Heyns, 1974 and Rotylenchus brevicaudatus Colbran, 1962. The females of these three species have similar body sizes, lip region of more or less hemispherical to rounded and very slightly offset to continuous with 4 to 5 annuli, stylet length below 30 µm, lateral field areolation only at pharynx level, absence of longitudinal cuticular striations, vulva position of 52 to 62% of body length from anterior end, phasmids positioned at 7 to 17 annuli anterior to anus, and short tail with only 5 to 11 tail annuli. The new species differs from R. abnormecaudatus in having 4 vs 4 to 5 lip annuli, a slightly longer cone part of stylet (45-53% vs 42-45%), and often truncated vs irregularly rounded tail terminus. Rotylenchus abnormecaudatus has also been reported with a peculiar and irregular arrangement of annuli and lateral field ending in the posterior part of tail which is not seen in other species including R. wimbii n. sp. While males of R. wimbii n. sp. and R. abnormecaudatus are not known and subsequently, no sperm is observed in the female spermathecae, males of R. brevicaudatus have been reported and the female spermatheca are often filled with sperm. Rotylenchus wimbii n. sp. further differs from R. brevicaudatus by the presence of 2 to 3 longitudinal striations on the basal lip annule vs 6 to 12 such longitudinal striations as reported by Sher (1965) from the study of six face views of the 20 female topotypes supplied by R.C. Colbran, and commonly truncated vs hemispherical tail terminus. Information on the number of longitudinal striations on basal lip annule for R. abnormecaudatus is not available.

The new species is also comparable to Rotylenchus cypriensis Antoniou, 1981, Rotylenchus mabelei Van den Berg and De Waele, 1989, and Rotylenchus unisexus Sher, 1965, all of which are species without males and reported to occur in South Africa and Kenya. The new species can be separated from R. cypriensis in continuous vs well offset lip region and absence vs presence of a ventral mucron at the tail tip. It can be differentiated from R. mabelei by a slightly smaller body (0.6-0.8 vs 0.9-1.1 mm), a lesser number of tail annuli (5-7 vs 14-18), and the position of phasmids with respect to anus (7-17 anterior vs 3 anterior to 5 posterior of anus). Finally, our new species can be differentiated from R. unisexus by the longitudinal striae on basal lip annule (3-4 vs 16-18 striae), tail annuli number (5-9 vs 9-13), and phasmid position (7-17 annuli anterior vs 6-7 annuli anterior of anus). An additional comparison of the new species with female morphological characters and morphometrics of all the Rotylenchus species that have been reported from Africa till date is provided in Table 2.

The species epithet refers to its host. Wimbi originates from Swahili and is used as a common name for finger millet in Eastern Africa.

The new species was found parasitizing the host plant Eleusine coracana (L.) Gaertn. (Planta: Poaceae), commonly known as finger millet, Kipkaren Estate, Eldoret, Kenya. The host status was also confirmed by a positive greenhouse test at the National Plant Protection Organization, Wageningen, The Netherlands. The GPS coordinates of the type location are 00°30.395’ N; 035°14.825’E.

Female holotype (Slide: WT 3796) and seven female paratypes in two slides (Slides: WT 3797 and WT 3798) are deposited at the National Plant Protection Organization, Wageningen Nematode Collection (WaNeCo), the Netherlands. One slide containing 10 female paratypes is deposited at Ghent University Museum, Zoology Collections, Belgium. An additional slide containing eight female paratypes is also deposited at UGent Nematode Collection (Slide: UGnem-300) of Nematology Research Unit of Ghent University, Belgium.

Several stained spiral-shaped nematodes were observed in the host roots which are Rotylenchus. As no other Rotylenchus species or other spiral PPN such as Scutellonema and Helicotylenchus was detected from the corresponding soil sample, the PPN observed in the roots are most likely R. wimbii n. sp. confirming that this novel species can parasitize finger millet.