First report of Meloidogyne hapla on kiwifruit in South Africa

Worldwide, the kiwi (Actinidia spp.) fruit trees are increasingly being cultivated in temperate regions. The genus Actinidia comprises more than 70 species (Peng et al., 2019), with A. deliciosa being the most popular across the world. At Magoebaskloof, Limpopo Province, South Africa, kiwifruits are produced in subtropical regions much closer to the tropical regions in the Southern hemisphere. The location has a high altitude that confers temperate climatic conditions, which are suitable for the production of kiwifruits. Subsequently, the kiwifruit is becoming increasingly important outside of temperate regions in South Africa.

During November 2019, root samples were collected from roots of kiwifruit trees in the Magoebaskloof area (S: 23°52′43′′; E: 29°56′13′′) (Fig. 1). Roots were washed, cut into pieces and mature female specimens were removed using a scalpel, using a Zeiss stereomicroscope. The specimens were fixed with a hot 4% formaldehyde solution and transferred to anhydrous glycerin using De Grisse’s (1969) method. The characteristics perineal patterns of the second-stage juvenile (J2) were used to identify the test Meloidogyne species (Perry et al., 2009).

Figure 1:

The molecular characterization followed the methods described in Álvarez-Ortega et al. (2019). The ribosomal ITS and LSU sequences were analyzed and aligned using the program BioEdit (Hall, 1999), aligned using CLUSTAL W (Thompson et al., 1994). The length of each alignment was 946 and 1186 bp for ITS rDNA and 28S rDNA, respectively. Bayesian inference was used to reconstruct the phylogeny, with Bayesian trees generated using the Bayesian inference method as implemented in the program MrBayes 3.1.2 (Ronquist and Huelsenbeck, 2003). The GTR + I + G model was selected using jModeltest 2.1.10 (Guindon and Gascuel, 2003; Darriba et al., 2012). Analysis using the GTR + I + G model was initiated with a random starting tree and ran with the Markov chain Monte Carlo (MCMC) for 10⁶ generations for ITS and 28S rDNA. The trees were visualized with the TreeView program. The original partial ITS rDNA and 28S (D2-D3 expansion) sequence of M. hapla were deposited in GenBank under the accession numbers MT256109 and MT258534, respectively. The morphological and molecular analyses confirmed that the species was M. hapla.

Morphometric mean, standard deviation and range values of M. hapla J2 were (n = 10): L = 337 ± 15.5 (322-353) μm; a = 30.8 ± 2.4 (28-32); b = 3.7 (n = 1); c = 8.5 ± 1.0 (7.4-9.5); stylet length = 12.6 ± 0.6 (11.9-13.0) μm; center of the median bulb to anterior end = 48.8 ± 2.6 (46-51) μm; excretory pore to anterior end = 67 ± 1.0 (66-68) μm; length body to the middle of genital primordium = 208.3 ± 8.6 (199-216) µm, hyaline part of tail length = 11.2 ± 1.3 (10.0-12.6) μm and tail length = 39.6 ± 3.1 (37-43) μm. The J2 had the smooth and spherical head, with a tail tapering to a blunt or rounded terminus. This species already has been studied from tropical areas of Africa (Whitehead, 1969), India (Waliullah, 2005), Chile (Carneiro et al., 2007), Brazil (Somavilla et al., 2011), Italy (D’Errico and Giacometti, 2012), and Turkey (Akyazi et al., 2017). Configuration of perineal patterns of females, their morphologies and morphometrics of M. hapla J2 were similar to those reported previously for isolates of this nematode species from tropical areas of Africa (Whitehead, 1969). However, the stylets of J2 from South Africa were slightly longer than those of the studied by Whitehead (1969). In comparison with Turkish population of M. hapla, they differ in body length (322-353 vs 340-440 µm), excretory pore to anterior end (66-68 vs 60.7-82.4 µm), hyaline part of the tail (10-12.6 vs 12-18.5 µm), and tail length (37-43 vs 50.2-54.8 µm). Although, our population of M. hapla showed no significant differences with those second-stage juveniles of M. hapla studied by Handoo et al. (2005).

The sequence lengths flanked by the forward primer TW81 [5′-GTTTCCGTAGGT GAACCTGC-3′] and AB28 [5′-ATATGCTTAAGTTCA GCGGGT-3′] (Joyce et al., 1994); D2A (5″-ACAAGTACCGTGAGGGAAAGTTG-3″) and the reverse primer D3B (5″-TCGGAAGGAACCAGCTACTA-3″) (De Ley et al., 1999) of the ITS rDNA and 28S region of M. hapla isolate 505 and 702 base pairs long, respectively. The nBlast test of ITS rDNA showed that the test population had only one base pair, which was different to those of M. hapla from South Korea (MK188473), Japan (LC030357), and Taiwan (KJ572385), all with 99% similarity. Despite high similarity (99%) with M. hapla populations, our sequence of M. hapla showed the lowest similarity, 85% with M. incognita (KJ739707) and M. javanica (KJ739709), and 79% with M. enterolobii (KM046989) using ITS rDNA marker. The nBlast of 28S rDNA showed four bp differences with 98% similarity with the Chinese (MN752204; KJ755183) and Ethiopian population (KP410845). Despite high similarity (98%) with M. hapla populations, our sequence of M. hapla showed the lowest similarity, 89% with M. incognita (JX100425), M. javanica (JX100426), and M. enterolobii (KJ146862) using 28rDNA marker. Therefore, molecular result confirmed our populations as M. hapla.

The phylogenetic analysis using ITS and 28S rDNA, placed the South African M. hapla population in a clade together with other M. hapla populations (Figs. 2, 3). The molecular characterization of several species of M. hapla suggested that they formed a monophyletic group. Findings in the current study were in agreement with the phylogenies of Meloidogyne species studied using 18S rDNA, ITS, 28S rDNA and COII of mtDNA (De Ley et al., 2002; Tigano et al., 2005; Tao et al., 2017). Two permanent microscope slides containing the perennial patterns and female and J2 of M. hapla were deposited in the Nematology collection of the University of Limpopo, South Africa. According to literature, this is the first record of M. hapla from kiwifruits in South Africa. Besides, ITS and 28S rDNA information of this species are being reported for the first time. M. hapla was associated with kiwifruits in Limpopo Province and therefore, host-status studies are necessary to find out the severity of this root-knot nematode.

Figure 2:

Figure 3: