Front. Plant Sci. Frontiers in Plant Science Front. Plant Sci. 1664-462X Frontiers Media S.A. 10.3389/fpls.2022.1093657 Plant Science Review RETRACTED: Meloidogyne enterolobii risk to agriculture, its present status and future prospective for management Sikandar Aatika Jia Luming Wu Haiyan Yang Shanshan * Guangxi Key Laboratory of Agro-Environment and Agric-Products safety, College of Agriculture, Guangxi University, Nanning, China

Edited by: Raja Asad Ali Khan, Hainan University, China

Reviewed by: Isabel Abrantes, University of Coimbra, Portugal

Huan Peng, Institute of Plant Protection (CAAS), China

*Correspondence: Shanshan Yang, yangshanshan12@126.com

This article was submitted to Plant Pathogen Interactions, a section of the journal Frontiers in Plant Science

 24 01 2023 2022 13 1093657 09 11 2022 05 12 2022 Copyright © 2023 Sikandar, Jia, Wu and Yang 2023 Sikandar, Jia, Wu and Yang

This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

Meloidogyne enterolobii, commonly known as guava root-knot nematode, poses risk due to its widespread distribution and extensive host range. This species is recognized as the most virulent root-knot nematode (RKN) species because it can emerge and breed in plants that have resistance to other tropical RKNs. They cause chlorosis, stunting, and yield reductions in host plants by producing many root galls. It is extremely challenging for farmers to diagnose due to the symptoms’ resemblance to nutritional inadequacies. This pathogen has recently been considered a significant worldwide threat to agricultural production. It is particularly challenging to diagnose a M. enterolobii due to the similarities between this species and other RKN species. Identified using traditional morphological and molecular techniques, which is a crucial first in integrated management. Chemical control, biological control, the adoption of resistant cultivars, and cultural control have all been developed and effectively utilized to combat root-knot nematodes in the past. The object of this study was to get about the geographical distribution, host plants, symptoms, identification, and control techniques of M. enterolobii and recommend future initiatives to progress its management.

 RKN virulent resistant root galls integrated disease management National Natural Science Foundation of China10.13039/501100001809 Introduction

Nematodes are one of the most abundant organisms on the planet (Hoogen et al., 2019; Sikandar et al., 2021a) and is a major component of soil (Hailu and Hailu, 2020). Plant-parasitic nematodes (PPNs) pose a significant threat to agriculture, causing an estimated yearly output loss of more than $157 billion globally (Youssef et al., 2013). The root-knot nematodes (RKN), are considered one of the most pathogenic PPN (Sikandar et al., 2019). These parasites are economically significant and one of the most destructive pests of vegetables and other crops (Tileubayeva et al., 2021). Root-knot nematodes are obligate endoparasites that live in the roots of more than 3,000 different plant species (Sikandar et al., 2020a). They are found worldwide, and their population multiplies when conditions are favorable (Feyisa, 2022).

Meloidogyne enterolobii, known as guava root-knot nematode, poses a risk to agriculture because of its worldwide distribution and diverse host range (Dareus et al., 2021). This species is recognized as being among the most virulent RKNspecies due to its ability to emerge and breed in host plants having resistance against major tropical RKN (Koutsovoulos et al., 2020). M. enterolobii was previously identified as M. incognita in 1983 in the Chinese pacara earpod tree (Enterolobium contortisiliquum) (Yang and Eisenback, 1983). In 1988, it was represented as a novel species found in Puerto Rico, identified as Meloidogyne mayaguensis (Rammah and Hirschmann, 1988). However, in 2004 it was reclassified as Meloidogyne enterolobii based on morphological and molecular evidence (Xu et al., 2004). This nematode had caused tremendous harm in the Psidium guajava (guava trees) in South America, that’s why it commonly called “guava root-knot nematode” (Palomares-Rius et al., 2021). M. enterolobii may cause more than 65% of the losses alone, which is significantly greater than any other RKNs species (Castagnone-Sereno, 2012). The growers still may not recognize that crops infecteduntil the harvest occurs and then notice a high number of galls on roots (Philbrick et al., 2020). Because of the similarities between M. enterolobii and other RKN species, diagnosing an infestation of M. enterolobii is very difficult (Min et al., 2012).

Synthetic chemicals have been used to control nematodes, but they are very poisonous and hazardous to the environment (Sikandar et al., 2021b). Most nematicide compounds, including ethylene dibromide (EDB), dibromochloropropane, and methyl bromide have been withdrawn from the market because several are carcinogenic (Onkendi et al., 2014). Bio-control, crop rotation, cultural practices, and plant resistance are now the main research areas for researchers attempting to address this challenging problem (Sikandar et al., 2020b). Compared to chemicals, bio-control is safer and more environmentally friendly because it has no residual effect (Köhl et al., 2019).

Thus, we present an overview of M. enterolobii research from all over the world. Moreover, we focused on how this accomplishment can help with M. enterolobii control. This review also includes species details as well as some recommendations for additional research on this lethal pathogen.

 Geographical distribution and host plants

Meloidogyne enterolobii nematode has been documented globally and is primarily found in tropical and subtropical areas (Silva and Santos, 2017). However, it was discovered in China and has now been recorded in Africa, Asia, Amercia (North and South) and Europe ( Table 1 ).

Geographic distribution of Meloidogyne enterolobii.

Continent Country Reference
Africa Cote d’Ivoire Fargette (1987)
Togo Fargette (1987)
Burkina Faso Fargette et al. (1994)
Senegal Diop (1994)
South Africa Willers (1997)
Malawi Trudgill et al. (2000)
Congo Onkendi et al. (2014)
Nigeria Kolombia et al. (2016)
Kenya Chitambo et al. (2016)
Niger Assoumana et al. (2017)
Benin Affokpon et al. (2017)
Mozambique Kisitu et al. (2017)
America (North) Guadeloupe Rammah and Hirschmann (1988)
Puerto Rico Rammah and Hirschmann (1988)
Cuba Decker and Rodriguez Fuentes (1989)
Guatemala Carneiro et al. (2000)
Martinique Carneiro et al. (2000)
United State of America Brito et al. (2004)
Costa Rica Humphreys et al. (2012)
America (South) Trinidad and Tobago Trudgill et al. (2000)
Brazil Carneiro et al. (2001)
Venezuela Lugo et al. (2005)
Asia China Yang and Eisenback (1983)
Vietnam Iwahori et al. (2009)
Thailand Jindapunnapat (2012)
India Poornima et al. (2016)
Europe Switzerland Kiewnick et al. (2008)
Portugal Santos et al. (2019)

It is a polyphagous RKN with various plant host species ( Table 2 ). Only a few number of fruit and vegetable species (Allium fistulosum, A. sativum, Anacardium occidentale, Annona cherimola, Arachis hypogaea, Averrhoa carambola, Brassica oleracea, Citrus aurantium, Citrus limonia, Citrus paradise, Citrus reticulate, C. reticulate, C. sunki, C. trifoliate, C. volkameriana, Cocos nucifera, Euterpe oleracea, Fragaria ananassa, Mangifera indica, Olea europaea, Passiflora spp., Persea americana and Zea mays) have been documented to be poor hosts for M. enterolobii (Freitas et al., 2017; EPPO-Datasheet, 2020).

Meloidogyne enterolobii host plants reported worldwide.

Scientific name Common name Family References
Abelmoschus esculentus Okra Malvaceae Vinícius-Marin et al. (2017)
Acalypha australis Copperleaf Euphorbiaceae Jia et al. (2022)
Acanthospermum australe Spiny-bur Asteraceae Bellé et al. (2019)
Ajuga reptans Bugleweed Lamiaceae Brito et al. (2010)
Allium porrum Garden leek Amaryllidaceae Rosa et al. (2015)
Amaranthus deflexus Perennial pigweed Amaranthaceae Bellé et al. (2019)
A. hybridus Pigweed Rich et al. (2009)
A. spinosus Spiny amaranth Rich et al. (2009)
A. tricolor Edible amaranth Bellé et al. (2019)
Ananas comosus Pineapple Bromeliaceae Silva and Oliveira (2010)
Angelonia angustifolia Summer Snapdragon Plantaginaceae Kaur et al. (2006)
Apium graveolens Celery Apiaceae Rodriguez et al. (2003)
Artocarpus heterophyllus Jackfruit Moraceae Brito et al. (2015)
Beta vulgaris Sugar beet Chenopodiaceae Moens et al. (2009)
Bidens pilosa Hairy beggar ticks Asteraceae Bellé et al. (2019)
Brassica oleracea Mustard Brassicaceae Bitencourt and Silva (2010)
Brugmansia suaveolens White angel trumpet Solanaceae Brito et al. (2010)
Buddleja davidii Butterfly bush Scrophulariaceae Brito et al. (2010)
Byrsonima cydoniifolia Nanche Malpighiaceae Paes et al. (2012)
Cactus sp. Cactus Cactaceae Silva et al. (2016)
Callistemon citrinus Lemon bottlebrush Myrtaceae Brito et al. (2010)
C. rigidus Stiff bottlebrush Marques et al. (2012)
C. viminalis Weeping bottlebrush Brito et al. (2010)
Cannabis sativa Hemp Cannabaceae Ren et al. (2021)
Canavalia ensiformis Jack bean Fabaceae Moens et al. (2009)
Capsicum annuum Bell pepper Solanaceae Assoumana et al. (2017)
Capsicum baccatum Orchid pepper Solanaceae Pinheiro et al. (2014)
Capsicum chinense Habanero pepper Solanaceae Melo et al. (2011)
Capsicum frutescens Bird pepper Solanaceae Silva et al. (2016)
Carica papaya Papaya Caricaceae Freitas et al. (2017)
Caryopteris clandonensis Bluebeard Lamiaceae Brito et al. (2010)
Cassia occidentalis Coffee Senna Caesalpiniacea Mendes and Dickson (2016)
Cereus fernambucensis Cactus Cactaceae Souza et al. (2006)
Chamaesyce hypericifolia Graceful spurge Euphorbiaceae Quénéhervé et al. (2011)
Citrullus lanatus Watermelon Cucurbitaceae Ramírez-Suárez et al. (2014)
Coffea arabica Coffee Rubiaceae Muniz et al. (2008)
Commelina benghalensis Wandering Jew Commelinaceae Bellé et al. (2019)
Coriandrum sativum Coriander Apiaceae Bitencourt and Silva (2010)
Cucumis melo Sweet melon Cucurbitaceae Freitas et al. (2017)
C. sativus Cucumber Bitencourt and Silva (2010)
Cucurbita moschata Pumpkin Cucurbitaceae Marques et al. (2012)
Daucus carota Carrot Apiaceae Wang et al. (2014)
Dioscorea rotundata Yam Dioscoreaceae Kolombia et al. (2016)
Elaeocarpus decipiens Japanese blueberry Elaeocarpaceae Moore et al. (2020)
Emilia sonchifolia Lilac Tassel Flower Asteraceae Rich et al. (2009)
Enterolobium contortisiliquum Tamboril Fabaceae Yang and Eisenback (1983)
Erechtites hieraciifolius American burnweed Asteraceae Carneiro et al. (2006)
Euphorbia heterophylla Desert spurge Euphorbiaceae Silva and Krasuski (2012)
E. prostrata Ground spurge Rich et al. (2009)
E. punicea Jamaican poinsettia Han et al. (2012)
E. tirucalli Indian tree spurge Souza et al. (2006)
Fatoua villosa Mulberryweed Moraceae Brito et al. (2008)
Ficus carica Common fig Moraceae Freitas et al. (2014)
Galinsoga parviflora Potato weed Asteraceae Bellé et al. (2019)
Gardenia jasminoides Cape jasmine Rubiaceae Lu et al. (2019)
Glycine max Soybean Fabaceae Ye et al. (2013)
Gossypium hirsutum Cotton Malvaceae Ye et al. (2013)
Helianthus sp. Sunflower Asteraceae Rosa et al. (2015)
Hibiscus grandiflorus Swamp rose mallow Malvaceae Brito et al. (2010)
Hydrocotyle bonariensis Largeleaf pennywort Araliaceae Souza et al. (2006)
Hypericum sp. St. John’s wort Hypericaceae Silva et al. (2016)
Inga edulis Ice-cream bean Fabaceae Silva et al. (2016)
Ipomoea batatas Sweet potato Convolvulaceae Melo et al. (2011)
Ixora chinensis Chinese Ixora Rubiaceae Wu et al. (2022a)
Jatropha urens Bull nettle Euphorbiaceae EPPO-Datasheet (2020)
Lactuca sativa Garden lettuce Asteraceae Correia et al. (2015)
Lagerstroemia indica Crape myrtle Lythraceae Brito et al. (2010)
Lampranthus sp. Ice plant Aizoaceae EPPO-Datasheet (2020)
Lantana camara Shrub lantana Verbenaceae Silva et al. (2016)
L. montevidensis Weeping lantana Brito et al. (2010)
Leonotis nepetifolia Christmas candlestick Lamiaceae Quénéhervé et al. (2011)
Leonurus sibiricus Siberian motherwort Lamiaceae Bellé et al. (2019)
Ligustrum sp. Privet Oleaceae Brito et al. (2010)
Luffa cylindrica Sponge gourd Cucurbitaceae Marques et al. (2012)
Lycopersicon peruvianum Peruvian tomato Solanaceae Melo et al. (2011)
L. hirsutum Hairy tomato Melo et al. (2011)
Malpighia emarginata Acerola cherry Malpighiaceae (Humphreys et al., 2012)
M. glabra Barbados cherry Silva et al. (2016)
Manihot esculenta Cassava Euphorbiaceae Rosa et al. (2014)
Maranta arundinacea Arrow root Marantaceae Zhuo et al. (2010)
Melaleuca linearis Pine-leave bottlebrush Myrtaceae EPPO-Datasheet (2020)
M. viminalis Weeping Bottlebrush EPPO-Datasheet (2020)
Merremia aegyptia Hairy woodrose Convolvulaceae Cunha e Castro (2019)
Mesona chinensis Chinese mesona Lamiaceae Wu et al. (2022a)
Morinda citrifolia Indian mulberry Rubiaceae Brito et al. (2010)
Morus alba White mulberry Moraceae Sun et al. (2019)
M. celtidifolia Texas mulberry Soares et al. (2018)
M. nigra Black mulberry Paes-Takahashi et al. (2015)
Musa spp. Banana Musaceae Freitas et al. (2014)
Myrica cerifera Wax myrtle Myricaceae Brito et al. (2010)
Nicandra physaloides Apple of Peru Solanaceae Bellé et al. (2019)
Nicotiana tabacum Tobacco Solanaceae Moens et al. (2009)
Ocimum basilicum Thai basil Lamiaceae Gu et al. (2021)
Oeceoclades maculata Monk orchid Orchidaceae Carneiro et al. (2006)
Ormosia hosiei Horse-eye bean Fabaceae Wu et al. (2022b)
Panicum sp. Panic Grass Poaceae Brito et al. (2008)
Passiflora mucronata Passion Flower Passifloraceae Rich et al. (2009)
Paulownia elongata Empress tree Paulowniaceae EPPO-Datasheet (2020)
Pentas lanceolata Star Cluster Rubiaceae Brito et al. (2010)
Phaseolus vulgaris Common bean Fabaceae Moens et al. (2009)
Physalis angulata Cutleaf ground cherry Solanaceae Marques et al. (2012)
P. peruviana Cape gooseberry Castagnone-Sereno (2012)
Psidium guajava Guava Myrtaceae Almeida et al. (2010)
P. guineense Brazilian guava Marques et al. (2012)
P. myrtoides Guava Marques et al. (2012)
Punica granatum Pomegranate Punicaceae Silva and Krasuski (2012)
Raphanus sativus Radish Brassicaceae Rosa et al. (2014)
Rhynchelytrum repens Natal grass Poaceae Bellé et al. (2019)
Rosa sp. Rose Rosaceae EPPO-Datasheet (2020)
Rotheca myricoides Butterfly bush Lamiaceae EPPO-Datasheet (2020)
Saccharum sp. Sugarcane Poaceae Marques et al. (2012)
Salix sp. Willow Salicaceae Brito et al. (2010)
Salvia leucantha Mexican bush sage Lamiaceae Levin et al. (2005)
Senna alata Candle bush Fabaceae Souza et al. (2006)
S. occidentalis Coffee senna Souza et al. (2006)
Sida rhombifolia Arrow leaf sida Malvaceae Bellé et al. (2019)
Solandra maxima Hawaiian lily Solanaceae Brito et al. (2010)
Solanum americanum American black nightshade Solanaceae Pinheiro et al. (2019)
S. gilo Scarlet eggplant Marques et al. (2012)
S. lycopersicum Tomato Silva et al. (2020)
S. melongena Aubergine Ghule et al. (2020)
S. pseudocapsicum Jerusalem cherry Groth et al. (2017)
S. quitoense Naranjilla Ghule et al. (2020)
S. scabrum Garden huckleberry EPPO-Datasheet (2020)
S. sisymbriifolium Sticky nightshade Bellé et al. (2019)
S. paniculatum Jurubeba Silva and Santos (2017)
S. pseudocapsicum Jerusalem cherry Groth et al. (2017)
S. tuberosum Potato Kan et al. (2008)
Stenocereus queretaroensis Pitaya Cactaceae Ramírez-Suárez et al. (2016)
Syngonium sp. Arrowhead plant Araceae EPPO-Datasheet (2020)
Syzygium aromaticum Clove Myrtaceae EPPO-Datasheet (2020)
Talinum patens Jewels of Opar Talinaceae Groth et al. (2017)
T. fruticosum Ceylon spinach EPPO-Datasheet (2020)
T. triangulare Philippine spinach Souza et al. (2006)
Tecoma capensis Cape honeysuckle Bignoniaceae Moens et al. (2009)
Tibouchina elegans Glory bush Melastomataceae Moens et al. (2009)
Ulmus parvifolia Chinese elm Ulmaceae Quénéhervé et al. (2011)
Washingtonia sp. Washington fanpalm Arecaceae Brito et al. (2010)
Vigna unguiculata Cowpea Fabaceae Guimarães et al. (2003)
Zingiber officinale Ginger Zingiberaceae Xiao et al. (2018)
Zizyphus jujuba Chinese date Rhamnaceae Long et al. (2014)
Symptoms

Plants infected with M. enterolobii have reduced growth, life span, and resistance against several abiotic stresses (Dareus et al., 2021). Generally, M. enterolobii effects may include reduced yield quality and quantity (Abd-Elgawad, 2021). Above-ground symptoms include leaf yellowing, wilting, and stunted growth while below-ground symptoms, such as root galls, can be considerable in size and quantity (Jia et al., 2022). Plants infected by M. enterolobii are more vulnerable to secondary plant infections, such as Fusarium solani parasitizing guava after infestation (Gomes et al., 2014).

 Identification of <italic>Meloidogyne enterolobii</italic> Morphology

Species of Meloidogyne have been identified based on adults’ morphology, along with an examination of the perineal patterns, which are structures of cuticle folds around the anus and vulva in adult females (Archidona-Yuste et al., 2018). Such detection techniques need tremendous experience and expertise, and negligence in using them may result in misdiagnosis (Bogale et al., 2020). The perineal patterns’ characteristics effectively distinguish M. enterolobii from other species of Meloidogyne (Ydinli and Mennan, 2016). M. enterolobii perineal patterns are often oval, with a round and high dorsal arch, large phasmids, a round tail tip part that lacks striae, and sometimes weak lateral lines present (Hunt and Handoo, 2009). Furthermore, perineal patterns within the species might differ between individuals, making diagnosis difficult (Karssen and Van Aelst, 2001). Moreover, M. enterolobii and M. incognita can exhibit eerily alike perineal patterns (Iwahori et al., 2009; Cunha et al., 2018), which is why M. enterolobii was initially believed to be M. incognita based upon the perineal investigation. Female RKN may be distinguished by their stylet, neck length, body form, and perineal pattern (Subbotin et al., 2021). Body morphometrics can be used to identify males and second-stage juveniles (J2) (Nyaku et al., 2018). Most RKN species have overlapping characteristics and measurements, making species identification challenging (Maleita et al., 2018).

 Isozyme analysis

Isozyme analysis is a biochemically standard diagnostic procedure that involves staining and observing malate dehydrogenase (Mdh), esterase, and cellulose acetate isozyme profiles after separation and migration through the electrophoresis (Siddiquee et al., 2010). The inter-species diversity produces a lot of isozymes, which have the same catalytic roles but differing chemical characteristics, like mobility in electrophoresis (Simonsen, 2012). The distinct pattern of one Mdh band and two distinct esterase bands in M. enterolobii distinguishes it from other species (Palomares-Rius et al., 2021). This approach successfully differentiated young adult females into species, while not being applicable for J2s (Castillo and Castagnone-Sereno, 2020). Additionally, this is extremely sensitive and carried out using only one adult female’s isolated protein (Birithia et al., 2012). Even though isozyme investigation was commonly used for identification of Meloidogyne (Nisa et al., 2022), more than single polymorphic enzyme was required to authenticate the identification of specific isolates because the presence or absence of an enzyme signal could varywithin and between samples (Cunha et al., 2018).

 Species specific polymerase chain reaction assay

This method has been designed and employed to distinguish the RKN species (Bhat et al., 2022). M. enterolobii was identified using a sequence characterized amplified region (SCAR) primer pair, such as MK7F/MK7R (GATCAGAGGCGGGCGCATTGCGA/CGAACTCGCTCGAACTCGAC) (Tigano et al., 2010). The IGS2 primers MeF/MeR (AACTTTTGTGAAAGTGCCGCTG/TCAGTTCAGGCAGGATCAACC) were substantially specific than MK7F/MK7R primers (Villar-Luna et al., 2016). TW81F/AB28R internal transcribed spacer (ITS) region primers were employed to diagnose M. enterolobii (Suresh et al., 2019). The multiplex PCR was intended to diagnose M. javanica, M. enterolobii, and M. incognita by DNA obtained directly from a single gall at different life cycle stages (Hu et al., 2011). A quantitative real-time PCR (qPCR) technique that measures the quantity of nucleic acid presence was developed for the precise detection, identification, and possibly quantification of M. enterolobii in both host roots and soil (Sapkota et al., 2016). In M. enterolobii, a unique satellite DNA family called pMmPet was found, providing species-specific PCR, dot blot, and southern blot analysis identification (Braun-Kiewnick et al., 2016). It was discovered that the satellite repetition was highly abundant and persistent across various populations of M. enterolobii, enabling single-individual identification and rendering it an efficient screening tool (Philbrick et al., 2020).

 Loop-mediated isothermal amplification

This approach has been designed to amplify DNA with selectivity, sensibility, accuracy, and quickly in isothermal conditions (Cai et al., 2018). Moreover, LAMP could amplify DNA in 1 hour in isothermal conditions using two or three sets of primers (Chen et al., 2011). A simple screening technique designed and employed in the field to detect M. enterolobii, M. arenaria, M. hapla, M. javanica, and M. incognita is recognized as the LAMP assay (Niu et al., 2012). Using a single-tube assay method based on the PCR melting curve methodology, the novel post-PCR analysis approach known as high-resolution melting curve analysis (HRMC) may distinguish between different DNA sequences according to their length, composition, and GC content (Holterman et al., 2012). Various tropical Meloidogyne species might be distinguished using HRMC analysis (Palomares-Rius et al., 2021). M. enterolobii isolates displayed distinct melting peak trends, having 1 or 2 peaks with varying centered heights at various melting temperatures, indicating a risk of employing a fragment that generated multiple amplicons of different lengths inside the same species (Chen et al., 2022). Moreover, examining novel single copy genes and regions in multiplex HRMC tests may be efficient in distinguishing M. enterolobii from other RKN species (Chen et al., 2022). Single nucleotide polymorphisms (SNPs) analysis may be an effective and reasonable method for diagnosing M. enterolobii (Holterman et al., 2012). The phylogenetic genetic relationships of the M. javanica, M. enterolobii, and M. incognita populations in South Africa were successfully investigated, and 34 SNPs that effectively distinguished these Meloidogyne species were discovered by using the genotyping-by-sequencing (GBS) technique (Rashidifard, 2019). Koutsovoulos et al. (2020) reported the genomes of M. hapla, M. incognita, and M. enterolobii. Because mitotic parthenogenesis is also a mode of reproduction in M. enterolobii, there has been little genetic variability found inside it (Humphreys-Pereira and Elling, 2015). The M. enterolobii isolates from various hosts and regions were tested using DNA markers, which revealed that they were genetically homogenous (Schwarz et al., 2020).

 Life cycle

M. enterolobii’s life cycle ( Figure 1 )is similar to other RKN species (Castillo and Castagnone-Sereno, 2020). Mature females lay their eggs in a gelatinous matrix (Kole, 2020). This matrix holds the eggs together, which protects them from severe climatic conditions (Mwesige, 2013). The nematode develops into a first-stage juvenile (J1), then molts into J2, and then hatchs from the egg (Velloso et al., 2022). Hatching can be affected by moisture, temperature, and the pH of the soil (Velloso et al., 2022). Second-stage juveniles travel toward the new host and penetrate the root system (Rashidifard et al., 2021). These nematodes travel to the vascular cylinder, and make massive feeding sites by causing physical damage with the stylets and releasing cellulolytic and proteolytic enzymes (Pulavarty et al., 2021). Giant cells form on the feeding sites, resulting in the characteristic galls observed on infected root systems (Nguyen, 2016). Giant cells are multinucleated, larger cells that normally develop in plant vascular tissues, and nourish nematodes by redistributing the metabolites of plants (Sreekavya et al., 2019). The J2 further molt three times, transitioning to the third-stage (J3) and fourth-stage (J4) until becoming sexual adults (Jagdale et al., 2021). Due to a malfunctioning stylet, the J3 and J4 stage nematodes cannot feed (Rashidifard, 2019). Vermiform male M. enterolobii worms emerge from the root system of the host plant (Castillo and Castagnone-Sereno, 2020). Furthermore, various Meloidogyne species only develop males in non-favorable circumstances, like extremely hot soil and inadequate moisture content (Giné et al., 2021). The Meloidogyne species have a 30-35 days life cycle in ideal circumstances, and every female may produce 500-1000 eggs in a gelatinous matrix (Feyisa, 2022). Koutsovoulos et al. (2020) demonstrated that M. enterolobii can also reproduce by obligate mitotic parthenogenesis, which occurs when the nucleus splits into two daughter nuclei that share similar genetic information as their parents. Meanwhile males can arise from genetically predisposed females under harsh environmental circumstances (Philbrick et al., 2020).

Life cycle of Meloidogyne enterolobii.

 Disease incidence condition

The incidence of the disease and yield losses caused by root-knot nematodes are frequently undetermined because their foliar signs are identical to those of other biotic diseases and abiotic stresses, such as stunted growth and yellow leaves (Liang et al., 2020). M. enterolobii is an extremely pathogenic species that causes extensive root galling as compared to other Meloidogyne species. It is also a very effective parasitic species with a high infestation rate on the host plant’s roots. Tomato yield declined by up to 65% in a microplot experiment (Cetintas et al., 2008). In just two greenhouses in Switzerland, output losses of up to 50% and substantial stunting of cucumber and tomato rootstocks were observed (Kiewnick et al., 2008). Infected Mulberry (Morus spp.) plants developed many galls on their roots, which are characteristic indications of root-knot nematode (M. enterolobii) infection, and the disease incidence was 100% (Sun et al., 2019). M. enterolobii reduced guava production in Brazil by 70% in 7 years, resulting in a US$61 million economic loss, that’s why cultivation may become unprofitable in highly infested areas with M. enterolobii (Carneiro et al., 2007).

 Integrated disease management strategies

It includes the combined application of several disease management strategies in order to reduce disease prevalence and severity while also reducing the pathogenic population below the devastating economic threshold (Forghani and Hajihassani, 2020). While integrated disease management (IDM) is a cost-effective and environmentally friendly strategy, it might be difficult to control the disease when a severe M. enterolobii infection has developed (Schwarz et al., 2020). M. enterolobii management is difficult because of its diverse host range and rapid reproduction cycles (Castagnone-Sereno, 2012). Therefore, developing successful strategies and incorporating them into disease management programs might effectively prevent disease outbreaks, lower disease severity, and boost agricultural output (Desaeger et al., 2020). They can be managed using various methods, such as chemical control, biological control, the adoption of resistant cultivars, and cultural control (Abd-Elgawad, 2022). The researchers usually use a single management strategy at a time to control this virulent nematode, so there is an urgent need to design a study in which different management strategies are applied at a time and also focus on inventing new management strategies. Additionally, a reliable and accurate diagnostic technique for M. enterolobii investigation might promote agricultural productivity and improve preventative activities to protect epidemiological research and crop management strategies internationally.

Chemical control

The application of chemical nematicides has controlled Meloidogyne species, although most of these substances are being banned due to safety concerns and hazards (Abd-Elgawad, 2021). Non-fumigants and fumigants are two major chemical nematicides used to regulate M. enterolobii (Castillo and Castagnone-Sereno, 2020). Non-fumigant nematicides are often prepared as liquids or grains form that can be properly mixed in water (Morris, 2015). Ethoprop, fluopyram, terbufos, fluensulfone, and oxamyl are some popular non-fumigant nematicides that are often used to manage Meloidogyne species (Desaeger et al., 2020). While the fumigants are typically composed of gases or liquids, this enables them to be rapidly evaporated and circulate in air holes between soil particles (Stejskal et al., 2021). The fumigants 1,3-dichloropropene, metam sodium, and metam potassium are commonly used to control M. enterolobii (Talavera-Rubia and Verdejo-Lucas, 2021). While fumigants are effective in controlling Meloidogyne species, these are generally costly and vulnerable to heightened legal scrutiny (Nyczepir and Thomas, 2009). Moreover, nematicides are classified as contact or systemic based on whether they directly kill nematodes in the soil or are first absorbed by plants (Lahm et al., 2017). Such chemical nematicides are incredibly hazardous as their residues can be detected in the food chain (Abd-Elgawad, 2016). Nematicide mode of action refers to the lethal action of nematicides on important life processes within nematode (Oka, 2020). Broad-spectrum fumigant nematicides, enter the nematode’s body wall directly and do not need to be eaten to be effective (Desaeger et al., 2020). Once they enter the nematode’s body cavity, they affect various internal organs when these organs are drenched in body fluids containing the nematicide (Desaeger et al., 2020). However, they are characterized biocidal compounds because they effect on fungus, bacteria, seeds, and other organisms in the soil and can pose environmental disruption and phyto-toxicity (Ebone et al., 2019; Oka, 2020). Nonfumigants can also directly enter nematodes’ body walls (Ebone et al., 2019).

 Biological control

Biological control with microbial antagonists (bacteria and fungi) has generated tremendous attention as a safe alternate and potential method of controlling plant-parasitic nematodes for ecological balance and safety (Riascos-Ortiz et al., 2022). Bacillus firmus, B. firmus, B. amyloliquefaciens, B. subtilis, B. urkholderia spp., Microbacterium spp., Paenibacillus spp., Pseudomonas spp., Serratia spp., Sinorhizobium spp., and Streptomyces spp. have exhibited nematicidal action against eggs, juveniles, and adults of Meloidogyne species (Aioub et al., 2022). Paenibacillus alvei increased the mortality of juveniles and decreased the hatching of M. enterolobii (Bakengesa, 2016). Microbacterium maritypicum and Sinorhizobium fredii have been shown to restrain nematode development and promote systemic resistance (Zhao et al., 2019). Plant-parasitic nematodes M. enterolobii are suppressed by plant growth promoting bacteria (PGPB) via several processes depending on microorganisms’ ability to compete successfully for ecological niches, colonize plant surfaces, and release nematicidal and antimicrobial chemicals (hydrolytic enzymes, toxins, antibiotics, siderophores, etc.) (Bakengesa, 2016; Gamalero and Glick, 2020). Bacteria and their metabolites have an impact on both the plant and microbial communities (Burkett-Cadena et al., 2008). Antibiosis, parasitism, or competition for resources or infection sites can all have a direct antagonistic effect (Migunova and Sasanelli, 2021). Bacteria can indirectly boost host defensive systems, resulting in induced systemic resistance (ISR) (Yu et al., 2022). Acremonium, Arthrobotrys, Chaetomium, Monacrosporium, Paecilomyces, Pochonia, Purpureocillium, and Trichoderma are fungi that are antagonistic and trap nematodes with sticky mycelia (Moliszewska et al., 2022). Endophytic fungi like Paecilomyces and Trichoderma can capture and destroy Meloidogyne species in the soil or root systems and restrain their development (Kassam et al., 2022). Similarly, Purpureocillium lilacinum and Pochonia chlamydosporia display the most significant effects and are suitable for biocontrol of M. enterolobii (Flores Francisco et al., 2021). To control M. enterolobii, additional study is required on the efficiency and broad-spectrum action, improving growth conditions, and sustainability of beneficial antagonistic bacteria or fungi for their marketing and use in IDM. Arbuscular mycorrhizal fungi (AMF) form a mutualistic symbiotic relationship with plants. As a result, they alter root structure, increasing plant tolerance, altering rhizosphere interactions, limiting plant-parasitic nematode feeding and space in the root, and inducing systemic resistance (ISR) (Vishwakarma et al., 2022). As microbiome research expands, the discovery of beneficial microbial agents for M. enterolobii for field application will be critical in the coming years (Galileya Medison et al., 2021). It is also crucial to consider how beneficial microbes interact with plant roots and symbiotic connections to better understand the various mechanisms behind their activities against M. enterolobii (Mohamed et al., 2022). According to research on its direct effects on plant-parasitic nematodes and its numerous benefits, AMF may be utilized as a biocontrol agent in suppressing M. enterolobii and improving nutrient absorption for improved crop productivity and quality (Forghani and Hajihassani, 2020). Fungi are recognized as a biocontrol agent through various mechanisms of action, including antibiosis, mycoparasitism, competition with pathogens, stimulation of plant growth, improved plant tolerance to abiotic stressors, and activation of pathogen defenses (Hermosa et al., 2012). The major direct contact mechanisms are competition and the formation of lytic enzymes and/or secondary metabolites (antibiosis) (Poveda, 2020). In order to colonize plant tissues, endophytic fungi must at least partially inhibit the plant defenses that allow them to produce induced systematic resistance (ISR) and systematic acquired resistance (SAR) against the invasion of pests and/or diseases (Busby et al., 2016). The strictly direct mechanisms of mycorrhizal fungi against nematodes are not yet adequately described, as they typically act through the plant host, altering root morphology by increasing root growth and branching, increasing water uptake and nutrients, making plants competitive with other plants for nutrients and space, or changing rhizosphere interactions (Schouteden et al., 2015). Nematodes can be directly attacked, killed, rendered immobile, or repelled by endophytic fungi. They can also be rendered unable to locate their hosts, have an effect on the development of nurse cells, compete in resource competition, or combine several of these tactics (Schouten, 2016).

 Resistance

Numerous research projects are being conducted worldwide to improve plant resistance to RKN (Padilla-Hurtado et al., 2022). The most cost-effective and environmentally friendly way to eradicate RKNs is to plant resistant cultivars (Ayala-Doñas et al., 2020). Meloidogyne species resistance is conferred by at least ten plant-resistance genes (Mi-1, Mi-2, Mi-3, Mi-4, Mi-5, Mi-6, Mi-7, Mi-8, Mi-9, and Mi-HT) (Rezk et al., 2021). Only five of them (Mi-1, Mi-3, Mi-5, Mi-9, and Mi-HT) have now had their genes mapped (El-Sappah et al., 2019). However, M. enterolobii is more pathogenic than other Meloidogyne species in crop genotypes with multiple sources of resistance genes (Collett et al., 2021). For instance, M. enterolobii thrives in crop genotypes resistant to other Meloidogyne species, such as resistant Capsicum annuum (N gene, Tabasco gene), Vigna unguiculata (Rk gene), Glycine max (Mir1 gene), Gossypium hirsutum, Ipomoea batatas, Solanum lycopersicum (Mi-1 gene), and Solanum tuberosum (Mh gene) (Schwarz, 2019).

Currently, researchers have concentrated on finding alternative sources of genetic resistance against M. enterolobii because this species has the potential to reproduce on a variety of crops that have resistance genes against other nematodes species (Castillo and Castagnone-Sereno, 2020). The exploration of new sources of tolerance or resistance against M. enterolobii has required a tremendous amount of research. In the previous research, Silva et al. (2019) reported that three varieties of wild and commercial tomatoes (Solanum pimpinellifolium “CGO 7650”, and S. lycopersicum “CNPH 1246 and Yoshimatsu”) exhibited resistance against M. enterolobii. Pinheiro et al. (2020) studied thirty-seven pepper genotypes to identify their resistance against three root-knot nematode species (M. incognita, M. javanica, and M. enterolobii). Only two genotypes (CNPH 6144 and CNPH 30118) were resistant against M. enterolobii.

Moreover, translationally controlled tumor protein (TCTP) was initially discovered in mice (Yenofsky et al., 1982). A new M. enterolobii TCTP (MeTCTP) effector exhibited the potential to increase parasitism, most likely by reducing programmed cell death in the host (Zhuo et al., 2017). The silencing of the MeTCTP effector reduced the reproduction and parasitic ability of M. enterolobii, indicating the nematode effector gene as a target for host-generated RNAi to establish disease resistance (Zhuo et al., 2017). Furthermore, new bioinformatics tools and genome sequence data have both become available for efficient dsRNA construction and stacking dsRNA sequences to target several genes for management of nematodes (Banerjee et al., 2017). The identification and functional studies of nematode-effector targets utilizing RNAi technology could carry substantial potential to enhance resistance in plants to M. enterolobii.

 Cultural control

Cultural control is an old and cost-effective approach to manage nematodes, such as crop rotation with non-host crops or resistant cultivars (Molendijk and Sikora, 2021). Crop rotation to non-host crops suppresses M. enterolobii populations because it cannot reproduce without a suitable host (Niere and Karuri, 2018). Nematode populations can be reduced by rotating hosts for at least a year (McSorley, 2011). As a result, crops should rotate to non-hosts for at least three years (Seid et al., 2015). While crop rotation is impeded because of M. enterolobii’s vast variety of hosts (Groover, 2017). The rotation crops of garlic (Allium sativum), grapefruit (Citrus paradise), maize (Zea mays), peanut (Arachis hypogaea), sour orange (C. aurantium), and wheat can be used because they have been known to be poor hosts of M. enterolobii (Rodriguez et al., 2003). Additional cultural practices such as steaming, flooding, and soil solarization could be applied (Schwarz, 2019; Schwarz et al., 2020). A key prophylactic tactic is weed control, as many of them can act as M. enterolobii’s hosts (Bellé et al., 2019). Nematodes may spread rapidly through agricultural tools, water, and plant matter; thus, sterilization prevents the nematodes from spreading to unaffected fields (Philbrick et al., 2020). To promote the effective management of M. enterolobii, a more specific study on cultural control measures like soil amendments, crop rotational strategies, and tillage is required.

 Conclusion and perspectives

In this review, we particularly emphasized the advancements achieved by numerous researchers in biology, identification and control of M. enterolobii. The new outbreak of the extremely pathogenic and destructive nematode M. enterolobii threatens agriculture worldwide. Biological control with microbial antagonists (bacteria and fungi) has generated tremendous attention as a safe alternate and potential method of controlling M. enterolobii for ecological balance and safety. Extensive investments are required in fundamental research aimed at identifying species and understanding parasitism mechanisms, evolution, and genetic diversity at a deep level to control this emerging RKN. Therefore, it is more vital than ever to create accurate and reliable identifying genetic markers, specifically for proper identification and to restrict the emergence of this pathogenic RKN. Both traditional methods and modern technologies must be considered to maintain food security. Currently, researchers have also concentrated on finding alternative sources of genetic resistance against M. enterolobii because this species has the potential to reproduce on a variety of crops that have resistance genes against other nematodes species. The fresh insights on existing and forthcoming concerns, underpinned by only a better knowledge of the relationship between the host and M. enterolobii, may increase the potential for inventing new management strategies. Controlling such an economically destructive nematode in agricultural production systems must involve broad research alliances and bring multidisciplinary researchers studying M. enterolobii.

 Author contributions

AS, LJ and HW discussed and conceived ideas. AS gathered the literature and wrote the manuscript. HW and SY helped to revise the manuscript. All authors have read, edited, and approved it for publication.

 Funding

We gratefully acknowledge the financial support of the National Natural Science Foundation of China (32160627, 32202245), the Guangxi Natural Science Foundation (2020GXNSFDA297003) and Guangxi Innovation Team of National Modern Agricultural Technology System (nycytxgxcxtd-10-04) for this research.

 Conflict of interest

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

 Publisher’s note

All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article, or claim that may be made by its manufacturer, is not guaranteed or endorsed by the publisher.

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