A New Root-Knot Nematode species, Meloidogyne karsseni n. sp. (Nematoda: Meloidogynidae), From Mexico and a Taxonomic Update on M. paranaensis From Guatemala

The new species was found to be associated with sweet pepper in 2019 in Mexico and has been maintained in the greenhouse of the NPPO in potted tomato (Solanum lycopersicum L.) plants (NPPO collection number: F5919). The M. paranaensis population was received by the NPPO from Guatemala in 1995 as a RKN-infested tomato root sample. This population has also since been in the greenhouse culture at the NPPO in potted tomato plants (NPPO collection number: C7729).

The J2 and males of both species were extracted from about 100 ml of soil from their respective culture pots by the centrifugation method (van Bezooijen, 2006). Obese females were directly picked out from the root galls of the host plants using fine forceps, needles, and glass pipettes. The nematodes were stored at 4°C during the study.

Both temporary mounts of heat-killed and permanent mounts of chemically fixed nematode specimens were used for morphological characterization. For preparation of a temporary mount of a J2 or male, a Cryo-Pro label (VWR International) was cut into two halves and stuck at the center of a glass slide, leaving a small parallel gap between them. A single specimen was transferred in a drop of distilled water onto the glass slide in the center of the gap, then the slide was passed intermittently over a flame until nematode movements stopped. The specimen was covered with a glass coverslip and examined, photographed, and measured using an Olympus BX50 DIC Microscope (Olympus Optical, Tokyo, Japan), equipped with a UCMOS 5MP camera. After recording morphological data, individual specimens were recovered from the slide by adding a few drops of water from one end of the gap and collecting the nematodes that were flushed out from the other end of the gap (Singh et al., 2021). Obese female specimens were individually teased out from galled roots in water in a petri dish using two fine needles and pipetted into a new embryo glass dish. The recovered specimens were subsequently used to extract genomic DNA, as described in this manuscript. For making permanent mounts, live nematodes were killed and fixed using warm 4% formaldehyde (about 60°C) and left in the fixative for 10 days at 4°C. The fixed nematodes were gradually transferred to glycerin (Seinhorst, 1959). The fixed specimens were mounted in glycerin on glass slides, covered with glass coverslips and studied using the camera-equipped compound microscope. Drawings were made with the aid of a drawing tube attached to the microscope and further improved using Photoshop CS6 (Adobe, San Jose, CA).

For scanning electron microscopy (SEM), specimens fixed in Trump's fixative (2% paraformaldehyde, 2.5% glutaraldehyde in 0.1 M Sorenson buffer [sodium phosphate buffer at pH = 7.3]) 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 of gold, and photographed with a JSM-840 EM (JEOL, Tokyo, Japan) operating at 12 kV (Singh et al., 2020).

Individual nematodes (J2 and females) were used to extract genomic DNA. The cuticle of each nematode was first punctured with the help of a metallic pin used as a worm-picking tool, and transferred to a polymerase chain reaction (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 Belgium], and 0.45% Tween 20). The PCR tubes were then frozen at −20°C (10 min), then 1 μl proteinase K (1.2 mg/ml) was added. The mixture was incubated at 65°C (1 h) and 95°C (10 min), and finally the lysate was centrifuged at 14,000 × g for 1 min (Singh et al., 2020).

The D2–D3 of 28S, partial 18S and ITS of rDNA were amplified by PCR using the primer pairs D2A: 5′-ACA AGT ACC GTG AGG GAA AGT TG-3′/D3B: 5′-TCC TCG GAA GGA ACC AGC TAC TA-3′ (Nunn, 1992), SSU18A: 5′-AAA GAT TAA GCC ATG CAT G-3′/SSU26R: 5′-CAT TCT TGG CAA ATG CTT TCG-3′ (Mayer et al., 2007) and Vrain2F: 5′-CTT TGT ACA CAC CGC CCG TCG CT-3′/Vrain2R: 5′-TTT CACT CGC CGT TAC TAA GGG AAT C-3′ (Vrain et al., 1992) respectively, and following the thermal profiles described in Singh et al. (2019). For amplification of the partial sequence of the mitochondrial cox1 gene, the primer pair JB3: 5′-TTT TTT GGG CAT CCT GAG GTT TAT-3′/JB4.5: 5′-TAA AGA AAG AAC ATA ATG AAA ATG-3′ (Bowles et al., 1992) was used with thermal profile from Etongwe et al. (2020). The PCR products were enzymatically cleaned using alkaline phosphatase (1 U/ml) and exonuclease I (20 U/ml) for 15 min at 37°C followed by 15 min at 80°C (Singh et al., 2020), and sequenced at Macrogen (Seoul, South Korea, https://dna.macrogen.com). The contigs were made from the newly produced forward and backward sequences using Geneious Prime 2020.0.5 (Dotmatics, Boston, MA, https://www.geneious.com) and were deposited in GenBank.

The isozyme characterizations were based on esterase (Est; EC 3.1.1.1) and malate dehydrogenase (Mdh; EC 1.1.1.37) staining of individual young egg-laying females (Karssen et al., 1995).

The phylogenetic relationships of the two species with other RKN species were analyzed based on the D2–D3, ITS, 18S, and cox1 sequences. 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 in GenBank. All of the collected sequences for each gene fragment were aligned using MUSCLE alignment in Geneious Prime 2020.0.5 using default parameters, followed by manual trimming of the poorly aligned ends. Bayesian phylogenetic analysis (MrBayes 3.2.6) was carried out using the GTR + I + G nucleotide substitution model, analyses were run under 1 × 10⁶ generations (4 runs), with Markov chains being sampled every 200 generations. 20% of the converged runs were regarded as burn-in (Huelsenbeck and Ronquist, 2001).

Figures 1–3, Table 1.

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Figure 2:

Figure 3:

The species is named in honor of the eminent nematologist, Prof. Dr. Gerrit Karssen, for his valuable and exemplary contributions as a nematode taxonomist, especially in the RKN group.

Sweet pepper (Capsicum annuum L.).

Guanojuato state in the city of San Jose Iturbide, Mexico.

Holotype female and paratypes of females, J2 and males have been deposited at the Wageningen Nematode Collection (WaNeCo), Wageningen, the Netherlands. Additionally, paratypes of all life stages have also been deposited at Ghent University Zoological Museum and UGent Nematode Collection of the Nematology Research Unit of Ghent University, Belgium (slide: UGMD_104440).

Meloidogyne karsseni n. sp. is characterized by J2 of 0.3 to 0.4 mm long with conical-rounded continuous head, post-labial annule without transverse striation, thin stylet of 11 to 12 μm, DGO at 5.2 to 6.0 μm from the knobs, hemizonid just above SE-pore, tail tapering to a finely rounded terminus and hyaline part bearing one or two very weak constrictions; globular females have rounded to oval perineal patterns with coarse striations, especially on the lateral sides around anus, and low dorsal arch with fine striations; males are without striations on the post-labial annule, with 20 to 24 μm long robust stylet and DGO at 2.4 to 2.9 μm from the knobs.

This species can be morphologically separated from all the known 20 species of the tropical RKN species complex, and especially from the five most common and also morphologically close RKN species found in North, Central, and South America – M. arabicida López & Salazar, 1989; M. izalcoensis Carneiro, Almeida, Gomes & Hernandez, 2005; M. lopezi Humphreys-Pereira, Flores-Chaves, Gomez, Salazar, Gomez-Alpízar & Elling, 2014; M. konaensis Eisenback, Bernard & Schmitt, 1994; and M. paranaensis (Subbotin et al., 2021).

Meloidogyne karsseni n. sp. is morphologically close to M. arabicida in that they have similar perineal patterns, stylet length in all stages, spicule length, ‘a’ and ‘c’ ratios in J2, tail, and tail hyaline lengths. However, it can be differentiated from M. arabicida based on the DGO lengths in males (2.4 to 2.9 vs. 3 to 5 μm from knobs) and in the J2 (5.2 to 6.0 vs. 2.0 to 4.7 μm), and absence vs. presence of transverse striations on the post-labial annule of both male and J2.

Meloidogyne karsseni n. sp. and M. izalcoensis are similar in J2, female and male stylet lengths; the absence of transverse striation on post-labial annule of both J2 and males; and ‘a’ and ‘c’ ratios, tail, and tail hyaline lengths of J2. However, the new species can be separated from M. izalcoensis based on the lengths of the DGO from knobs in females (2.9 to 3.8 vs 4.5 to 6.0 μm), males (2.4 to 2.9 vs. 4.0 to 7.0 μm), and the J2 (5.2 to 6.0 vs. 3.0 to 4.0 μm), and female perineal patterns (coarse striations around anus with low dorsal arch containing fine striations vs. coarse striations around the vulval lips and with a comparatively higher dorsal arch).

The new species and M. lopezi are similar in the J2 and male stylet length; male DGO length from knobs; absence of transverse striation on the post-labial annule of both the J2 and males; the ‘a’ and ‘c’ ratios; tail and hyaline lengths; and tail shape of the J2. However, M. karsseni n. sp. differs from M. lopezi by having a shorter female stylet (11.8 to 15.6 vs. 15 to 23 μm), more posterior DGO position in the J2 (5.2 to 6.0 vs. 2.5 to 3.5 μm), and female perineal pattern (lower dorsal arc with fine striations vs. comparatively higher dorsal arch with coarser striations).

Meloidogyne karsseni n. sp. and M. konaensis are similar in male stylet lengths; DGO position in J2; and lack of transverse striation on the post-labial annule in J2 and males. However, they differ in the stylet lengths of females (11.8 to 15.6 vs. 14.0 to 20.0 μm) and J2 (10.8 to 12.4 vs. 12.6 to 15.0 μm); DGO lengths in females (2.9 to 3.8 vs. 3.4 to 7.0 μm) and males (2.4 to 2.9 vs. 5 to 9 μm); absence vs. presence of numerous large projections on stylet shaft of male; and the lip region of the J2 (conical-rounded vs. more truncated).

Meloidogyne karsseni n. sp. is similar to M. paranaensis in J2 and male stylet lengths, J2 tail and hyaline lengths, and the ‘a’ and ‘c’ ratios. However, M. karsseni n. sp. differs from M. paranaensis by the absence vs. presence of transverse striation on the post-labial annule of both males and the J2; shorter female stylet length (11.8 to 15.6 vs. 15.0 to 17.5 μm); DGO length in the females (2.9 to 3.8 vs. 3 to 6 μm), males (2.4 to 2.9 vs 3.0 to 5.5 μm), and J2 (5.2 to 6.0 vs 2.0 to 4.5 μm); and the female perineal pattern (low dorsal arch vs. higher dorsal arch).

Lastly, the new species can be easily separated from the globally distributed tropical RKN species – M. arenaria (Neal, 1889) Chitwood, 1949, M. incognita (Kofoid & White, 1919) Chitwood, 1949, and M. javanica (Treub, 1885) Chitwood, 1949 – based on the combination of the following features: absence of transverse striation on post-labial annule and longer DGO length (>5 μm) in the J2; perineal patterns with coarse striations, especially on the lateral sides of anus; and very low dorsal arch.

Figures 4–6, Table 2.

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Figure 5:

Figure 6:

The current population is morphologically similar to the original description of M. paranaensis (Carneiro et al., 1996) and also to the populations from Brazil and Guatemala (Santos et al., 2020). However, in these compared populations, a distinct sclerotization of the duct near the SE-pore of male was not mentioned, whereas such a sclerotization was prominent in all the males of our population. Therefore, this characteristic should be re-examined in the type specimens, and also in the Guatemalan population of M. paranaensis mentioned above, to ensure that it is a characteristic that separates it from other very similar species, such as M. incognita. Furthermore, the esterase phenotype of the herein-studied population was identical to that of the Guatemalan M. paranaensis population (see details under molecular characterization) (Santos et al., 2020). The exact location where the currently studied sample was collected in Guatemala remains unknown. Permanent mount of females, J2, and males have been deposited at the Wageningen Nematode Collection (WaNeCo), Wageningen, the Netherlands, and also at the UGent Nematode Collection of the Nematology Research Unit of Ghent University, Belgium.

Three identical sequences, each of the D2–D3 expansion segment of 28S (OQ772268–OQ772270; 735–740 bp), partial 18S (OQ772271–OQ772273; 860bp) and partial ITS (OQ772265–OQ772267; 709–713 bp) of rDNA, and five identical sequences of cox1 of mtDNA (OQ773540–OQ773544; 375–415 bp) were generated for M. karsseni n. sp. The D2–D3 sequences were found to be closest to an unidentified Meloidogyne sp. (JX465613; 99.6% identity; 3 out of 716 bp difference); the 18S sequences were closest to M. incognita (JX100421; 99.8% identity; 2 out of 854 bp difference); the ITS sequences were closest to M. luci Carneiro, Correa, Almeida, Gomes, Mohammad Deimi, Castagnone-Sereno and Karssen, 2014 (LN626964; 98.4% identity; 10 out of 641 bp difference); and the cox1 sequences was identical to M. incognita sequence (ON276646; 100% identity).

Similarly, for M. paranaensis, three D2–D3 of 28S (OQ753107–OQ753109; 630–730 bp long; 98.3–98.6% similarity; 14–15 bp differences), two identical partial 18S (OQ750410–OQ750411; 850 bp) and one partial ITS (OQ753116; 707 bp) sequences of rDNA, and three identical cox1 (OQ750575–OQ750577; 360–410 bp) of mtDNA were generated. The D2–D3 sequences were found to be closest to M. incognita sequence (MT193445; 99.6% identity; 4 out of 630 bp difference); the 18S sequences were closest to M. arabicida sequence (AY942625; 99.7% identity; 2 out of 850 bp difference); the ITS sequences were closest to M. incognita sequence (KP901064; 97.2% identity; 25 out of 709 bp difference); and the cox1 sequences were also closest to M. incognita sequence (ON276646; 99.5%; 2 out of 414 bp differences). Interestingly, upon comparison of the D2–D3 and the ITS sequences with the available corresponding sequences of M. paranaensis from Guatemala (Santos et al., 2020), our sequences were found to be relatively dissimilar to KY911103 (97.2–98.6% identity, 9 to 24 bp differences) and KY911109 (92.3% identity, 58 out of 730 bp difference).

In Santos et al. (2020), the 28S and the ITS sequences from the Brazilian and the Guatemalan M. paranaensis populations showed remarkable intraspecific sequence variations, up to 2%, and 8%, respectively. In the current study, a similar level of variation in the 28S (up to 3%) and the ITS sequences (7.3%) were also observed between our population and the Guatemalan population (Santos et al., 2020), indicating that these gene regions may not be reliable for the identification of M. paranaensis.

Based on the D2–D3 tree (770 bp-long alignment) (Fig. 7), both M. karsseni n. sp. and M. paranaensis were placed together in an unresolved clade (PP = 0.6) consisting of only tropical and subtropical RKN species (also known as Tropical RKN complex) (Álvarez-Ortega et al., 2019). Similarly, in the ITS tree (1000 bp-long alignment) (Fig. 8), both species appear in the tropical and subtropical RKN species clade (PP = 1), but the phylogenetic position of M. karsseni is only slightly resolved, i.e., it is sister to 16 species (PP = 0.79) (see trees for the respective related species). As 18S and cox1 tropical and subtropical RKN sequences were also highly conserved, and thus yielded similar tree topologies with an unresolved clade of tropical and subtropical RKN species, they are not shown here.

Figure 7:

Figure 8:

The isozyme codes (Fig. 9) of esterase phenotypes and malate dehydrogenase phenotypes of M. karsseni n. sp. are, respectively, F1-I1-S1 and N1 types. For M. paranaensis, they are F1–S1 (= P2a) and N1, respectively. The isozyme codes for the new species have not previously been seen, whereas those of M. paranaensis were identical to the corresponding codes of M. paranaensis from Guatemala (Santos et al., 2020).

Figure 9: