Introduction

Global withdrawal of the highly effective synthetic chemical fumigant nematicides, which had been relied upon for over a century in the management of plant-parasitic nematodes (PPN), has had severe economic ramifications in crop production systems (Caboni et al. 2015). Parasitism by root-knot nematodes (RKN), Meloidogyne species is considered one of the main biotic factors responsible for reduced productivity in various agricultural crops (Mhatre et al. 2019). RKN results in up to 30% yield decline by direct infestation and indirect losses owing to predisposing or breakdown of resistance to other root diseases, such as bacterial wilt, attributing to quantity and quality losses (Muimba-Kankolongo 2018). Meloidogyne genus is a worldwide economically significant pest, comprising over 100 species, (Karuri et al. 2017) including approximately 22 described from Africa (Onkendi et al. 2014) widely distributed on leguminous and flowering plants.

Two Meloidogyne species, M. incognita and M. javanica, have been declared economically important to roughly 4000 host plants, including field crops, ornamentals, medicinal, aromatics plants, and even weeds (Jones et al. 2013; Onkendi et al. 2014). Second-stage juveniles (J2) penetrate roots to establish a feeding site, called giant cell, usually within the pericycle and vascular tissues and form root galls soon after their infection (Mashela et al. 2015). In nematode-susceptible hosts, infection by Meloidogyne species induces the formation of severe root galls, stunted growth, decreased water uptake, imbalances of essential nutrient elements, low evapotranspiration and increased root exudation of amino acids, which reduces soil pH (Saikia et al. 2013).

Up-to-date cultural management procedures are insufficient to fully manage RKN (Trudgill and Blok 2001), even more so with continued restrictions on synthetic chemical use (Desaeger et al. 2017). As a result, it has become critical to develop additional PPN management strategies that are environmentally friendly. Currently, there are numerous studies conducted on the subject all around the world (Baum et al. 2015; Brito et al. 2020; Damasceno et al. 2015; Gupta et al. 2017; Hussain et al. 2018; Laquale et al. 2015; Seo et al. 2019), including screening nematode-resistant genotypes (Chiamolera et al. 2018; Da Silva-Mattos et al. 2019; Hajihassani et al. 2019). These studies are proving to be beneficial, providing additional insights that can lead to increased profits for farmers.

Nematode-resistant hosts may exhibit pre- or post-penetration resistance (Thurau et al. 2010). Pre-penetration nematode resistance is the form of resistance that occurs prior to nematodes coming into contact with the root systems (Ferraz and Brown 2002). This form of resistance prevents penetration of nematode J2 and is characterized by pre-existing morphological factors or the production of root exudates that either attract or repel J2 (Trudgill 2003). Root penetration by RKN has also been attributed to the lack of metabolites required for host identification, repellent host exudates, or the existence of a physical barrier over which the nematode cannot pass (Lee et al. 2017).

In post-penetration nematode resistance J2 are allowed to penetrate the root systems (Desmedt et al. 2020), with passive chemicals previously called elicitors, activated to form the phytoalexins, that have nematicidal properties (Desmedt et al. 2020, 2022). Some of the phytoalexins induce hypersensitive response (HR), that appear as necrotic spots, where cells around the nematode wither (Huysmans et al. 2017), thereby preventing feeding, development of J2 and reproduction. According to Lopez-Gomez and Verdejo-Lucas (2017), post-penetration incompatibility in resistant crops is associated with failure of giant cells to develop further into root galls. Rootlet interference and small underdeveloped root galls are also characteristics of post-penetration nematode resistance (Benková and Bielach 2010). In sedentary RKN, this type of resistance is further subdivided into early and late resistance, wherein early resistance that occurs during migration or early site establishment, and late resistance that occurs after the establishment of a feeding site (Fuller et al. 2007).

Between the two mechanisms of resistance, only post-penetration nematode resistance can be introgressed (Thurau et al. 2010), dictating the need to establish the mechanism of nematode resistance in any nematode resistant plant species in order for it to serve as a candidate of introgression. Among the available alternative techniques to methyl bromide, plant resistance is one of the most investigated techniques in PPN management (Onkendi et al. 2014). Most crops lack resistant genotypes to Meloidogyne species as observed in four commercial genera of Cucumis, Citrullus, Cucurbita and Lagenaria within the Cucurbitaceae family (Liu et al. 2016; Thies et al. 2016; Verdejo-Lucas and Talavera 2019; Singh and Patel 2015). Cucumis africanus is highly resistant to Meloidogyne species (Pofu et al. 2012); however, the mechanism of nematode resistance in this crop has not been established.

Therefore, the objective of this study was to determine the mechanism of nematode resistance in C. africanus to M. incognita and M. javanica.

Materials and methods

Experimental procedures

Two separate experiments were conducted under greenhouse conditions at North-West University, South Africa. Greenhouse temperature were set at 25 ± 2 °C, with temperatures and humidity controlled using thermostatically activated fans and wet-wall at opposite ends. Seeds of C. africanus were sown in seedling trays filled with pasteurized (300 ℃ for 1 hour) fine sand and raised for 6 weeks. Uniform seedlings were transplanted into 250 ml polystyrene cups, filled with 200 ml pasteurized fine sand and placed at 10-cm inter- and intra-row spacing. In each experiment, the treatments comprised of 15 harvesting times, experimentally done in a randomized complete block design (RCBD), with five replications. Isolates of M. incognita and M. javanica were each raised on nematode-susceptible tomato (Solanum lycopersicum) cv. ‘Floradade’ seedlings and roots collected for egg masses when needed. Egg masses were hand-picked using a tooth pick and hatched in distilled water for 72 h (Powers et al. 1991). A day after transplanting, Cucumis seedlings were each inoculated by dispensing approximately 100 J2 of M. incognita or M. javanica using a 20 ml plastic syringe into 5-cm-deep furrow around the seedling stem and covered with growing medium. Harvesting was done every other day, for a period of 30 days starting from 2-days after inoculation. Seedlings were fertilized once with Super Phosphate (Efekto Care, Bryanston, South Africa) and NPK (2:3:2) and irrigated with 30 ml tap water every other day.

Data collection

At each harvest, seedling roots were severed and the shoots discarded. Roots were rinsed in tap water to remove soil particles, blotted dry using paper towel and stained (Bybd et al. 1983). The whole root system was soaked in 1.5% NaOCl solution for four minutes to remove any associated microbe, rinsed in tap water, followed by a 15 min immersion in tap water to remove excess NaOCl. Root samples were each stained by covering with 30 ml tap water mixed with 1 ml acid fuchsin and boiled for 30 s. The solution was cooled to room temperature and roots distained by putting in acidified glycerin with a few drops of 5 N HCl, which were heated to boiling, followed by cooling to room temperature. Root samples were each placed in a petri dish and closed with the top lid for assessment under the stereomicroscope at 45 × magnification for necrotic spots, rootlet interference, giant cells and root gall number.

Data analysis

Prior to analysis of variance (ANOVA), all data were transformed through log10(x + 1) to normalize the variances. Data were subjected to ANOVA through the 2008 SAS software. The mean sum of squares were partitioned to provide the contribution of sources of variation in the total treatment variation (TTV) of variables (Gomez and Gomez 1984). Treatment means were separated using Waller-Duncan Multiple Range test at 5% level of probability. Unless stated otherwise, all treatment effects were discussed at 5% level of probability.

Results

Harvesting times were highly significant (P ≤ 0.01) on necrotic spot number, rootlet interference number and root gall number contributing 59, 64 and 50% in TTV of the respective variables, but were not significant on giant cell number (Table 1). In the first 18 days after inoculation, necrotic spots, rootlet interference and root galls were not noticeable. The first necrotic spots and root galls were observed after 20 days, whereas rootlets after 22 days (Table 2).

Table 1 Total treatment variation (TTV) on necrotic spot, giant cell number, rootlet interference and root gall number in Cucumis africanus seedlings infected by Meloidogyne incognita and Meloidogyne javanica under greenhouse conditions at 30 days after inoculation (n = 75). Where DF is degrees of freedom and MS is mean sum of squares
Full size table
Table 2 Mean separation for necrotic spot, giant cell number, rootlet interference and root gall number in Cucumis africanus infected by Meloidogyne incognita and Meloidogyne javanica under greenhouse conditions at 30 days after inoculation (n = 75)
Full size table

Harvesting time had highly significant effects on necrotic spot number, giant cell number, rootlet interference number and root gall number, contributing 55, 71, 63 and 59% in TTV of the respective variables (Table 1). HR was noticeable 26 days after inoculation, giant cells and root galls after 18 days, whereas, rootlets were observed after 16 days (Table 2).

Discussion

According to Chitambar and Raski (1984), M. incognita is more pathogenic and becomes aggressive with time in comparison with M. javanica, which could explain why necrotic spots for C. africanus—M. javanica relations were observed 6 days after C. africanus—M. incognita relations. HR is known to be a common response to RKN infection in resistant crops (Chitambar and Raski 1984; Das et al. 2008; Freire et al. 2010; Lee et al. 2021), resulting in cell death and prevention of nematode feeding site formation and nematode development, leading to subsequent nematode death (Postnikova et al. 2015). HR in nematode-infected cells representatives of hyperactive responses in nematode resistant plants (Mashela et al. 2016). According to Nicholson and Hammerschmidt (2003), HR indicates the presence of phenols that play a role in plant defense. When Abifarin et al. (2019) investigated phytochemical and antioxidant activities of C. africanus, they found that the plant has phenolic compounds in fruits, leaves and roots.

The presence of such phytochemicals could be responsible for pathogen-associated molecular patterns (PAMP) in C. africanus. PAMP result in incompatible nematode-host interactions that triggers the up-regulation of a network of host genes and corresponding proteins involved in an innate response known as pathogen-triggered immunity (PTI) (Melillo et al. 2006). Coffee (Coffea canephora cv. ‘Apoata’), resistant to M. exigua exhibited HR, which further inhibited formation of feeding site (Vieira et al. 2013). Moon et al. (2010) also observed necrotic spots in resistant C. annuum cultivars exposed to M. incognita. Two resistant alyce clover (Alysicarpus ovalifolium hybrids, FL-1 and FL-3), showed attributes of HR to M. arenaria (Powers et al. 1991). Marini et al. (2016) observed similar results for resistant oats (Avena sativa cv. ‘IPR Afrodite’) when exposed to M. incognita at 15 days after inoculation.

Delay in giant cells response had been previously reported in nematode resistant trials using molecular approaches (Escobar and Fenoll 2021). The giant cells appeared as deeper stained spots with multiple nuclei that failed to develop beyond the zygote-like size. In nematode-susceptible plant species, giant cells are formed as multinucleate structures formed when the feeding cell and those around it responds to RKN infection by undergoing repeated mitosis without cytokinesis (Huang et al. 2003). The successful establishment of giant cells is essential for nematode development. Meloidogyne species evolved strategies that enable them to induce giant cell formation on thousands of plant species by manipulating important factors of plant cell development (Caillaud et al. 2008). This group of notorious pathogens secrete chemical compounds called gene products through the sub-ventral and dorsal gland cells during migration and sedentary phases, respectively (Gheysen and Fenoll 2002; Tripathi et al. 2015). The release of gene products is important during RKN migration and feeding site establishment because it enables nematode growth to subsequent stages (Curtis 2008; Siddique et al. 2022). Anti-plant gene, on the other hand, is a strategy by host plants when plant genes that respond to nematode feeding and secretions to allow for successful partnerships between PPN and plants are silenced (Mashela et al. 2016). Thus, the phytotoxic chemical compounds that destroy the feeding structures, giant cells, are upregulated.

Marini et al. (2016) also noticed that M. incognita gradually initiated giant cells that failed to develop into root galls in resistant roots of A. sativa 18 days after inoculation. Similarly, Wehner et al. (1991) observed small, poorly formed giant cells in resistant cucumber (C. sativus) and African horned cucumber (C. metuliferus) exposed to M. hapla. Observation of the under-developed giant cells also agreed with observations in resistant soybean (Glycine max cvs. ‘Jackson’ and ‘PI 200,538’) exposed to M. arenaria (Pedrosa et al. 1996) and in resistant cayenne pepper (Capsicum annuum cvs. ‘02G132’ and ‘03G53’) (Moon et al. 2010). Pedrosa et al. (1996) indicated that resistance to M. arenaria was expressed in G. max as small, poorly formed giant cells. In all the cited examples, the cultivars had post-penetration nematode resistance. The giant cell serves as a source of nutrients for the developing nematode (Bartlem et al. 2014). The post-penetration compatibility in susceptible crops is usually associated with optimal development of giant cells that form a large multinucleate structure which, however, fail to develop in nematode resistant crops (Ortiz 2011).

Compensatory rootlet growth was observed originating adjacent to the under-developed giant cells. Mechanisms behind compensatory rootlet development in RKN-infected resistant hosts have not been investigated. However, considering the tactics RKN implement throughout migratory phases, it is possible that resistant hosts develop lateral rootlets to supplement for roots that can no longer transport nutrients and water from soil to aboveground parts. After root penetration, RKN seek for a suitable feeding site and position themselves strategically in the vascular bundles. While host feeder roots absorb nutrients and water, which move in the plant's vascular system to aboveground plant parts, these nutrients are channeled into RKN throughout their development (Bartlem et al. 2014). Villordon and Clark (2018) noted an increase in lateral root growth on sweet potato resistant (cv. ‘Bayou Belle’) compared to susceptible variety (cv. ‘Beauregard’) hosts. The observations supported those in nematode-resistant G. max that was exposed to M. javanica (Doyle and Lambert 2003) and on nematode-resistant Trifolium repens that was exposed to M. trifoliophila (Mercer et al. 2004).

Generally, in nematode-susceptible plant species, when Meloidogyne J2 develop through J3, J4 and adult female stages, the adjacent root cells bulge to form a root gall. Of 39 cultivars of C. annuum screened for nematode resistance, six (‘02G132’, ‘03G62’, ‘04G8’, ‘99G198’, ‘03G53’ and ‘CM334’) were resistant to M. incognita, with few undeveloped root galls (Moon et al. 2010). Pedrosa et al. (1996) and Herman et al. (1991) noticed fewer J2 advancing to subsequent stages of Meloidogyne species.

In a host-parasitic interaction study, tomato host reactions to Meloidogyne species parasitism were initiated during the first 12 h after infection (Siddique et al. 2014). However, in the two Cucumis species against the Meloidogyne species in the current study, there was no evidence of rapid host reactions. Findings by Ramatsitsi and Dube (2020) explained and supported the findings in the current study wherein there were no detectable nematode juveniles in roots at 30 days after inoculation even though they were observed earlier after inoculation. At 30 days after inoculation, Marini et al. (2016) also found a decrease in nematode numbers inside the roots of a resistant A. sativa cv. ‘IPR Afrodite’ that was exposed to M. incognita. At the onset of feeding, the nematode becomes sedentary, going through three molts before becoming a mature adult female, with males migrating out of the plant without playing any role in reproduction (Caillaud et al. 2008).

The results showed similar mechanisms of resistance in the roots of nematode resistant C. africanus for both M. incognita and M. javanica. The discovery of post-penetration nematode resistance to Meloidogyne species would very certainly increase the use of C. africanus in plant breeding and crop rotation systems, hence extending the applications and economic relevance of C. africanus. For future research, efforts could be made to investigate whether C. africanus is predisposed to other soil-borne pathogens through puncture wounds from penetration of the nematodes.