Abstract
Nematodes and fungi are soil inhabitants. Both are essential for maintaining the stability of food-web and facilitation of the nutrient cycle. Interaction between nematodes and fungi is possible in multiple ways. Here, we supply a platform for nematophagous (nematode destroying) fungi (NF), their mode of action, and their importance in agricultural ecosystems. They are potentially important for sustainable agriculture and play a major role in integrated pest management programs. Nematophagous fungi belong to a broad taxonomic group, such as Ascomycota, Oomycota, Basidiomycota, and distinct groups of fungi. Nematophagous fungi are broadly distributed in terrestrial and aquatic ecosystems that contain high densities of nematodes. Depending on the mechanism that affects nematode, NF can be divided into four types. Here, we described the classification, taxonomy, occurrence, distribution and ecology, types of nematophagous fungi, and potential mechanisms of NF in the control of plant-parasitic nematodes.
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Keywords
- Plant-parasitic nematodes
- Nematophagous
- Biological control
- Arthrobotrys oligospora
- Egg parasites
- Second-stage juveniles
12.1 Introduction
In this materialistic world, increasing food production for feeding the ever-growing population is the major world demand, while earning more profit from agriculture is the grower’s demand, who faces an occasional setback due to the serious outbreak of the diseases and keeps looking for an effective method to save his crop from destructive pathogens. Among them, one of the major limiting factors is plant-parasitic nematodes (PPNs) that continuously affected agricultural production by and large. Further, the nature of crops, varieties, nematode species, the population of primary inocula, and environmental factors influence the losses to a great extent. It is estimated that 12.3% of global annual losses of major crops are due to phytoparasitic nematodes (Sasser 1989). Such type of losses become intolerable for poor and developing countries including India. Plant-parasitic nematodes cause 18–25% losses in vegetables, 20–25% in pulses, 18–23% in oilseed crops, and 15–18% in cereals crops (Indian Economy 2004).
Management of plant-parasitic nematodes is largely dependent on the use of toxic pesticides, the majority of which are soil fumigants. Farmers are using toxic pesticides intensively for the last few decades in order to reduce such a magnitude of losses to sustained crop production. However, the residual effects of these pesticides on nontarget soil flora and fauna are of great concern. Further, long-term residual effects are responsible for eroding biodiversity, increasing resistance and resurgence in the pathogen, and causing pollution that poses health hazards to humans, animals, and the environment. Present circumstances of environmental awareness evoke urgent need to search for and establish compatible alternatives to these hazardous agrochemicals. The persistence of pesticides in soils, deterrents to ecosystems, environmental contamination, detrimental impact on human health, deterrents to ecosystems, and the creation of resistant pathogenic strains are all consequences of heavy pesticide use. In order to reduce the use of pesticides, researchers have intensified resistance breeding programs along with transgenic plants to control the losses caused by these notorious pathogens. However, several constraints limit the scope of the resistance breeding program, i.e., unavailability of suitable donor parents having a high degree of resistance, detection of the source of resistance, and transferring desirable traits into a cultivar using a resistance breeding program is a great challenge. Biological control of plant-parasitic nematodes offers a promising alternative to pesticides, which had attained lots of attention over the years. Under these circumstances, using microbe-based strategies for the control of plant-parasitic nematodes has been reported to be an environmentally friendly, safe, and residue-free approach (Singh et al. 2012a, b, 2013, 2019a, b). Several biological control agents of microbial origin have been evaluated and used to control the plant-parasitic nematodes in many crops. Among them, Trichoderma asperellum, T. harzianum, T. virens, Bacillus subtilis, B. licheniformis, Pseudomonas fluorescens, Purpureocillium lilacinus, Arthrobotrys oligospora, Pochonia chlamydosporia, Dactylaria spp., Monacrosporium spp., Drechslerella dactyloides, Syncephalastrum racemosum, Hirsutella spp., and Duddingtonia spp. were noted worthy and are used to manage nematodes worldwide (Singh 2013; Singh et al. 2013, 2017, 2019b; Wang et al. 2014; Huang et al. 2014; Gupta et al. 2015a, b). Predacious fungi are an important part of the soil’s biodiversity. Interactions between predatory fungi and parasitic nematodes are widespread and dynamic in the soil. Several reviews and research publications have demonstrated the ability of nematophagous fungi (NF or NPF) to suppress plant-parasitic nematodes; however, they are scattered. Nematophagous (nematode-eating) fungi are found in both terrestrial and aquatic habitats and are diverse in nature (Pramer 1964; Nordbring-Hertz et al. 2006). More than 200 fungal species, which can develop specific trapping devices, belong to the NF group such as adhesive knobs, constricting rings, and adhesive networks to capture nematodes juveniles, eggs, and adults and then use various strategies to extract nutrients from their nematode prey (Jansson and Lopez-Llorca 2001; Nordbring-Hertz et al. 2006; Yang et al. 2007; Schmidt et al. 2007). Most of the nematode-trapping fungi can function as both saprophytes and parasites (Pramer 1964; Nordbring-Hertz et al. 2006). Nematode-trapping fungi develop sophisticated hyphal structures, such as hyphal knobs, hyphal branches or rings, and hyphal nets, by adhesion or mechanical capture (Nordbring-Hertz et al. 2006; Singh 2007; Singh et al. 2012b). They have an essential role in maintaining nematode population density via natural settings. Many egg-parasitic and trap-forming fungi may exist in soil saprophytically while endoparasites are obligate parasites. Considering the importance of the problem and the potentiality shown by previous workers, the present study was undertaken with the objectives to give an overview of the biology, ecology, and potential application of nematophagous fungi to sustain crop production in changing climatic scenarios.
12.2 Historical Background
The word “predacious” comes from the Latin word preada, which means “to grab out all valuables and things of a victim after they have been killed.” Predacious fungi are distributed in all types of soils. These fungi are more significant in decomposing plant waste, and the organic matter supplied to the soil increases the number of predacious fungi. Arthrobotrys oligospora was first described as a common inhabitant of organic plant debris by Fresenius (1852). Woronin (1870) reported that the conidia of A. oligospora germinated on the old manure and some of the hyphae-produced net-like bails, although he did not know the functions of such bails. Sorokin (1876) created the genus Catenaria with the type species C. anguillulae. He found C. anguillulae parasitizing eelworms in a vessel, which were eventually killed. He also described that round zoospores were liberated from the sporangia through a discharge tube. Zopf (1888) was the first to record the predacious behavior of A. oligospora. Further, his studies show that the cuticle of the captured nematode is penetrated, and the fungus grew within the nematode body and consumed it by its hyphae. Thus, Zopf established the predatory relationship of a fungus on nematodes. Drechsler (1937) established a base for studies on predacious fungi responsible for capturing and killing nematodes. He described that the predacious fungi produced different trapping devices for the predation of nematodes. Such capturing devices include adhesive hyphae, adhesive branches, adhesive nets, and adhesive knots, whereas the nonadhesive organs include non-constricting rings and constricting rings. Some predaceous fungi produce sticky knobs that capture nematodes (Drechsler 1937; Barron 1977).
12.3 Occurrence and Distribution
Predacious fungi can be found as saprophytes in the soil or on decaying plant materials where they live saprophytically and/or feeding on plant-parasitic nematodes. The efficacy of capturing predacious fungi may be influenced by the nature of the soil and environmental conditions. The presence of nematodes in the soil and organic matter is necessary in maintaining the biodiversity of soil and increasing the population of predacious fungi. Most of the predatory species belong to either the Zoopagales or Moniliales, while endo-parasites are found in the lower fungi such as Chytridiales, Saprolegniales, Peronosporales, Lagenidiales, Mucorales, Entomophthorales, and in higher fungi Deutromycetes.
12.4 Classification
Fungi represent the fifth kingdom in the living organisms (Kendrick 2001). NPF are found in all lower and higher groups of fungi, such as Basidiomycetes, Ascomycetes, and Deuteromycetes in higher fungi and Chytridiomycetes, Oomycetes, and Zygomycetes in lower fungi. In distinct taxonomic groups of fungi, the habit of nematophagous fungi evolved separately. Barron (1992) reported that the habit of nematophagous fungi evolved from lignolytic (characterized by a unique ability to depolymerize and mineralize lignin) and cellulolytic fungi (hydrolyzing or having the capacity to hydrolyze cellulose) for adaptation to overcome nutrient competition in the soil.
In tandem with entomopathogenic species of Verticillium, which were relocated to the genus Lecanicillium based on both morphological and molecular features, egg-parasitic fungi previously placed within the genus Verticillium were recently shifted to the new genus Pochonia (Zare and Gams 2001; Zare et al. 2001). Cordyceps contains the teleomorphs of the Pochonia species. P. chlamydosporia and P. rubescens are the most well-known egg parasites, and other taxa reported to parasitize nematode eggs include Paecilomyces lilacinus and Lecanicillium lecanii. Based on the molecular evidence, Scholler et al. (1999) proposed the following classification: Arthrobotrys (adhesive three-dimensional networks), Dactylellina (stalked adhesive knobs or non-constricting rings), Drechslerella (constricting rings), and Gamsylella (non-constricting rings) (having adhesive branches and unstalked knobs).
The taxonomy and phylogeny of endoparasitic fungi are far less well-understood (Fig. 12.1). Some, such as the zoosporic Catenaria anguillulae, are classified as Chytridiomycetes, while others are classified as Haptocillium (previously Verticillium), Harposporium, or Drechmeria. Harposporium spp. teleomorphs have lately been shifted to Podocrella from Atricordyceps (Chaverri et al. 2005). Pleurotus is a category of toxin-producing fungi that contains species such as the oyster mushroom Pleurotus ostreatus. Luo et al. (2004) reported that Coprinus comatus was recently discovered to have similar abilities, suggesting that nematophagy is more widespread among Basidiomycetes than previously thought.
12.5 Ecology
Nematophagous or nematode-trapping fungi (NF or NPF) are found in the soil. They are mostly found in the topsoil, meadows, leaf litter, mangroves, and some shallow aquatic areas. NF employ adhesive knobs, adhesive hyphal strands, and nets made of hyphal threads, hyphal loops, and non-constricting loops that capture nematodes. When the nematode is bridled, the NF hyphae enter the cuticle and eat the nematode’s internal tissues (Zhang et al. 2014).
Arthrobotrys oligospora, a species of Arthrobotrys, is one of the most well-studied nematode-trapping fungi (Nordbring-Hertz et al. 2006). Strains of A. oligospora have been discovered in different soil conditions (Pfister and Liftik 2018; Money 1998). By creating intricate three-dimensional networks, A. oligospora enters the parasitic stage in the presence of nematodes to capture them. Nematode trapping triggers a chain of actions that include nematode adhesion, penetration, and immobilization (Nordbring-Hertz 2004; Nordbring-Hertz et al. 2006). The fungus’ strong ability to capture nematodes makes it a promising candidate for controlling plant-parasitic worms. To catch nematodes mechanically, A. oligospora forms three-dimensional adhesive nets. The fungus actively seeks out its prey by creating chemical signals or olfactory cues that are similar to those used by worms to find food and mate (Yu’e et al. 2005; Zhang et al. 2015; Hsueh et al. 2017). Some nematophagous fungi produce toxins that render nematodes immobile. The hypha of the shaggy ink cap (Coprinus comatus) attacks the nematode Panagrellus redivivus as a spiny ball structure, which immobilizes and breaks the nematode cuticle, following which the hypha pierces the skin and digests the contents (Luo et al. 2007). The spores of the most endoparasitic fungi are attracted to and concentrated in the mouth region of soil nematodes. The hyphae proliferate throughout the nematode after penetration of the cuticle and absorption of nematode tissues. Conidia are contacted by the nematode in other fungal species and are infected in a similar fashion. Harposporium anguillulae having sickle-shaped conidia are consumed by the nematodes and lodge themselves in the esophagus or gut, where they destroy the tissues (Aschner and Kohn 1958).
The hypha flattens itself against the egg in egg-parasitic species, and the presence of appressoria indicates that infection is about to occur or has already occurred. After piercing the egg and devouring the developing juvenile worm, the hypha produces conidiophores and moves on to nearby eggs (Money 1998).
12.6 Plant-Parasitic Nematodes have an Impact on Agriculture
Plant-parasitic nematodes are a serious constraint in agricultural crop production. These nematodes have been discovered in over 4100 different species (Decraemer and Hunt 2006). Crop loss is projected to cost between US$118 and 80 billion per year (Sasser and Freckman 1987; Nicol et al. 2011). The most economically important nematode species accounts for 15% of all identified nematode species. They directly target the plant roots of major crops, preventing nutrient uptake and water, resulting in decreased agronomic performance, overall yield, and quality of the crop. Surprisingly, just a small percentage of the nearly 4000 reported plant-parasitic nematodes cause major agricultural losses. In a survey, the principal genera of phytoparasitic nematodes identified to cause crop losses in the United States were Meloidogyne, Heterodera, Hoplolaimus, Rotylenchulus, Xiphinema, and Pratylenchus (Koenning et al. 1999).
12.7 Types of Nematophagous Fungi
Nematophagous fungi can be divided into four major groups (Fig. 12.2) depending on their mode of attacking nematodes (Jansson and Lopez-Llorca 2001):
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1.
Nematode-trapping fungi (previously sometimes called predacious or predatory fungi)
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2.
Endoparasitic fungi
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3.
Egg- and female-parasitic fungi
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4.
Toxin-producing fungi
The nematode-trapping fungi use hyphal trapping devices of various shapes and sizes to capture nematodes, such as sticky/adhesive three-dimensional nets, adhesive knobs, and nonadhesive constricting rings. Some “nematode-trappers” capture nematodes by an adhesive substance formed on their hyphae without any visible traps, e.g., Stylopage spp. Endoparasitic fungi use their conidia or zoospores to infect the nematodes. The propagules of fungi adhere to the cuticle of the nematode, and then spore contents are injected into them or spores are swallowed by the host. Most of them are obligate parasites and the entire vegetative stages of their life live inside the infected nematodes. The egg- and female-parasitic fungi are facultative parasites. They infect nematode females and their eggs, using appressoria or zoospores. Being facultative parasites, they grow on nematodes and parasitize the sedentary stages such as eggs. The toxin-producing fungi produce toxic compounds that can immobilize nematodes, prior to penetration by hyphae through the cuticle of the nematode. Parasitism of nematodes results in complete prey or egg digestion in all four nematophagous fungal groups, an action that provides the fungus with nutrients and energy for continuous growth.
12.8 Mechanism and Mode of Action of Nematophagous Fungi to Control Plant-Parasitic Nematode
Biological control of phytonematodes is described as a reduction in populations of nematodes caused by actions of living organisms other than those found naturally in the host plant, or by introduction of antagonist organisms into the environment (Kim 2015). More than 200 taxonomically distinct fungi have demonstrated the ability to kill live nematodes in all the stages of development such as juveniles, adults, and eggs (Nordbring-Hertz et al. 2006). Two types of barriers to fungus invasion are created by the morphology of nematodes. The eggshell is the first barrier, made up of three layers in root-knot and cyst nematodes: the outer vitelline (mostly proteins), the inner lipoprotein layer, and the chitin layer, and the cuticle is the second barrier. The parasitism, poisonous chemicals, and enzyme methods used by nematophagous fungi to infect nematodes can be separated into three categories (Fig. 12.3).
Different nematophagous fungi infect nematodes and their eggs in a similar, general way. Infection of nematode eggs by Pochonia rubescens, as well as the zoospores of Catenaria anguillulae, which infect vermiform worms, demonstrate this. P. rubescens begins penetrating nematode eggs by contacting the egg with its hyphae and then forming an appressorium. The appressorium forms an extracellular matrix (ECM) or adhesive, which is disclosed by lectin Concanavalin A labeling. The fungus uses both mechanical and enzymatic components to enter the worm eggshell from the appressorium. Because the nematode eggshell is mostly made up of chitin and proteins (Bird and Bird 1991), chitinases and proteases are vital during the penetration of eggshell (Lopez-Llorca 1990a, b; Tikhonov et al. 2002). Eggshells are degraded as a result of proteolytic action.
12.8.1 Chemotaxis and Adhesion (Host Recognition, Host Specificity, and Infection)
The recognition phase of nematode begins, which includes chemotaxis of the host toward fungal traps, hyphae, or zoospore chemotaxis toward the host’s natural apertures (Jansson and Nordbring-Hertz 1979; Jansson and Thiman 1992). It is unknown which chemicals are involved in the chemotactic events (Jansson and Friman 1999; Bordallo et al. 2002). After contact with a nematode, the adhesive on A. oligospora traps changes from amorphous to fibrillar, in contrast to the adhesive on D. coniospora conidia, which always appears fibrillar (Jansson and Nordbring-Hertz 1988). Lopez-Llorca et al. (2002) suggested that the adhesion on the appressoria of P. rubescens and P. chlamydosporia can be identified using the lectin Concanavalin A, indicating that it is a glycoprotein having glucose/mannose moieties. A. oligospora’s Gal-NAc-specific lectin and D. coniospora’s sialic acid-specific lectin have both been implicated in worm recognition (Nordbring-Hertz and Mattiasson 1979; Jansson and Nordbring-Hertz 1984). Infection events eventually trigger a signaling cascade that is required for nematode prey penetration and colonization (Tunlid et al. 1992). An extracellular substance is generated after contact, which keeps the fungus attached to the nematode surface. Proteins or carbohydrates are typically found in the adhesives of nematophagous fungi (Tunlid et al. 1991a, b). Carbohydrates on nematode surfaces are engaged in the lectin-binding recognition stage, but they also appear to play a role in nematode chemotaxis (Zuckerman and Jansson 1984; Jansson 1987). Major nematode sensory organs, such as inner labial papillae and amphids, are positioned around their mouth in the labial and cephalic region (Ward et al. 1975). Zuckerman (1983) and Zuckerman and Jansson (1984) proposed that carbohydrates play a role in nematode chemoreception. Lectins (Concanavalin A binds with mannose/glucose residues, and Limulin binds with sialic acid) could block the chemoreceptors, leading bacterial-feeding nematodes to lose their chemotactic behavior to microbial exudates (Jeyaprakash et al. 1985). Further, nematode chemotaxis was reduced when enzymes (mannosidase, sialidase) obliterated the terminal carbohydrates (Jansson et al. 1984), demonstrating the importance of carbohydrate moiety in nematode chemotaxis. An endoparasitic nematophagous fungus, D. coniospora, uses conidia to infect nematodes that cling to the host’s chemosensory organs (Jansson and Nordbring-Hertz 1983). Both Limulin treatment of nematodes and sialic acid treatment of spores reduced conidial adhesion, implying that a sialic acid-like carbohydrate is involved (Jansson and Nordbring-Hertz 1984). Furthermore, it is evaluated that nematode adherent with spores lost their capacity to respond chemotactically to all the attracting sources, including hyphae, conidia, or bacteria, implying a link between chemotaxis and adhesion via carbohydrates on the surface of nematode (Jansson and Nordbring-Hertz 1983). D. coniospora conidia stick to Meloidogyne spp. chemosensory organs, but they do not penetrate and cannot infect the worms.
12.8.2 Differentiation and Signaling
When recognizing the surface of the host, or even synthetic surfaces, most harmful fungi distinguish appressoria. Appressoria formation in plant pathogenic fungi infecting leaves has been examined in depth (Lee et al. 2003; Basse and Steinberg 2004). St. Leger (1993) proposed a signaling hypothesis for the insect pathogen Metarhizium anisopliae during appressorium production, based in part on knowledge of plant pathogenic fungi. Appressoria on their hosts are differentiated by nematophagous fungi, particularly egg parasites (Lopez-Llorca and Claugher 1990). The signaling pathways that lead to nematode infection by nematophagous fungus are poorly understood. Using expressed sequence tag (EST) techniques, it was recently demonstrated that genes involved in the creation of infection structures and fungal morphogenesis were expressed during trap formation in the nematophagous fungus Dactylellina haptotyla (syn. Monacrosporium haptotylum) (Ahren et al. 2005). As a response to chemical and tactile inputs, fungi-infecting vermiform nematodes differentiate multiple trapping organs. The three ring cells that make up the trapping mechanism are inflated by the constricting ring traps. When a nematode comes into contact with the inner ring wall, an unknown mechanism causes the nematode to inflate and close, which takes around 0.1 s.
12.8.3 Nematodes Cuticle and Eggshell Penetration by NPF
Nematophagous fungi penetrate the worm cuticle or eggshell after a solid adhesion to the host surface. Both enzymatic and physical mechanisms appear to be used by the nematophagous fungus to penetrate host surfaces, as in many other cases of fungal penetration. Because the nematode cuticle is mostly made up of proteins (Bird and Bird 1991), proteolytic enzymes are necessary for penetration of the nematode cuticle. The PII serine protease of A. oligospora has been characterized, sequenced, and cloned (Ahman et al. 1996). The protein presence, such as nematode cuticles, increases PII expression (Ahman et al. 1996). The subtilisin PII has a molecular mass of 32 kDa and belongs to the subtilisin family (Fig. 12.4).
Another serine protease (Aoz1) was recently identified from A. oligospora, having 38 kDa molecular mass and 97% similarity with PII (Zhao et al. 2004). Other fungi have been isolated and characterized, including Arthrobotrys microscaphoides (Mlx) (Wang et al. 2006a, b) and Arthrobortys shizishanna (Ds1) (Wang et al. 2006a, b), both of which show significant similarities to the A. oligospora serine (Wang et al. 2006a, b).
Protein and chitin are structured in a microfibrillar and amorphous form in nematode eggshells (Clarke et al. 1967). As a result, extracellular enzymes that degrade such polymers were sought. Lopez-Llorca (1990a, b) identified, purified, and characterized P32, a 32 kDa serine protease from the egg parasite P. rubescens for the first time. P32 suppression by polyclonal antibodies and chemicals reduced the penetration and egg infection, despite pathogenesis being a complex process involving many variables (Lopez-Llorca et al. 2002). An extracellular protease (VcP1) is produced by P. chlamydosporia, which is linked to P32 and similar entomopathogenic fungal enzymes (Segers et al. 1994). Eggs treated with VcP1 enzyme were more easily infected than non-treated eggs, implying that the enzyme plays a role in eggshell penetration by fungi that feed on eggs. Recently, a serine protease (Ver112) from Lecanicillium psalliotae was isolated and described, exhibiting approximately 40% homology with Arthrobotrys proteases (PII and Aoz1) and have 60% homology with egg-parasitic serine proteases (Yang et al. 2005a, b). Non-nematophagous fungi such as Clonostachys rosea and Trichoderma harzianum are additional sources of nematicidal serine proteases (Suarez et al. 2004; Li et al. 2006). Huang et al. (2014) reported that Pochonia rubescens and Pochonia chlamydosporia both have chitinolytic enzymes that have been discovered. A 43 kDa endochitinase (CHI43) was one of those responsible for the majority of the activity (Tikhonov et al. 2002). Damage to eggshells was more widespread when treating G. pallida eggs with both P32 and CHI43, which indicated that the two enzymes work together to destroy eggshells (Tikhonov et al. 2002). A hydrolytic enzyme chitosanase from the egg-parasitic fungus P. lilacinus was recently identified and described (Chen et al. 2005).
12.9 Potential Application
For many years, nematophagous fungi have been tested for biological control of plant-parasitic nematodes, but the strategy had limited success due to the lack of understanding of these species’ ecology (Stirling 1991) (Table 12.1).
The colonization of plant roots by endophytes is a significant element. By induced resistance or by the production of secondary metabolites, NF may protect plants from several fungal diseases and plant-parasitic nematodes. Plant growth can also be boosted by nematophagous fungi participating in the nutrient intake or modifying plant growth regulators. As a result, endophytic colonization must be taken into consideration while looking for nematophagous fungi as biocontrol agents. Combining numerous forms of nematophagous fungi, such as egg-parasitic and nematode-trapping fungi, that kill nematodes at different phases of their lives, could be a key requirement. When choosing the right fungi for biological control of plant-parasitic nematodes, interactions with other soil fungi, including plant-parasitic and biocontrol agents, are also crucial factors to consider. Larriba et al. (2015) demonstrated the egg-parasitic fungus Pochonia chlamydosporia having potential for biological control of plant-parasitic nematodes. They act as an endophyte in both monocot and dicot plants and have shown plant growth promotion in a variety of crops. Nematophagous fungus Pochonia chlamydosporia promotes growth of barley (Hordeum vulgare) plants by endophytic colonization of roots and provides defense against stresses. Escudero and Lopez-Llorca (2012) stated that endophytic colonization of tomato roots by P. chlamydosporia is important for plant growth and may influence root-knot nematode management.
12.10 Future Prospects
Plant-parasitic nematodes cause major yield and monetary losses in agriculture all over the world. The utilization of nematophagous fungi as endophytes is a potential technique for the biocontrol of nematodes in the soil. Larriba et al. (2015) observed that at the molecular level, plants colonized endophytically having growth-promoting effect by P. chlamydosporia, paving the way for more research into the fungus’ ability to mitigate the negative effects of biotic and abiotic factors on plant crops. Bioproducts formulated with these NF have various advantages over chemical nematicides for more sustainable agriculture, including ease of application, environmental safety, little impact on soil biota, and no residues in harvested products. However, when producing a commercial bionematicidal product, there are various aspects to be kept in mind as it is a living system. As a result, new technology such as real-time quantitative PCR is used to quantify and track the biocontrol agent after its application into the soil. Biocontrol agents can be genetically modified to have their efficacy increased by increasing the expression of genes implicated in nematicidal activity or pathogenicity (Zhang et al. 2020a, b). To improve aggression and virulence against nematodes, expression of heat shock factors, UV protectants, immunological modulators, destroying enzymes of cuticle, and genetic modification techniques can be used. Several studies established the efficacy of applying a combination of treatments to manage plant-parasitic nematode populations under diverse conditions, including various cultural techniques (such as soil amendment and soil solarization), biological agents, and chemical nematicides (Zhang et al. 2014). Finally, the unpredictability of nematode antagonists against PPN in field circumstances, as well as their limited efficacy, are key barriers to using biocontrol agents to manage plant-parasitic nematodes. The intrinsic mechanisms governing ecosystem stability in field circumstances may be one of the causes for the disparities between the results of laboratory trials and field trials. Understanding interactions between nematodes and nematophagous fungi in native niches aids in the development of better applications for long-term crop protection approaches. The effects of combining various partners, such as NF, plant-pathogen mycoparasites, and plant growth-promoting microorganisms, could provide useful information for the development of biocontrol agents to reduce the impact of nematode and fungal pathogens on agriculturally important crops (Luns et al. 2018; Baron et al. 2020).
12.11 Conclusion
Nematophagous fungi are common soil organisms that may infect, attack, and consume nematodes at any stage of their development, including adults, juveniles, and eggs. To infect their nematode hosts, they use trapping organs, spores, and appressoria. In addition to infecting nematodes, nematophagous fungi can infect other fungi as mycoparasites and colonizing plant roots endophytically. Because of their various capacities, nematophagous fungi, in particular, may be a promising candidate for the biological management of plant root diseases. The use of nematophagous fungi as a substitute for synthetic chemicals used in the production of nematicides is fascinating. Obtaining bionematicides efficiently is a goal and a prerequisite for all agricultural researchers seeking sustainability in the system. Depending on the pathogenicity factor, some information is still lacking. Some NF enzymes such as serine proteases, chitinases, and toxins function as virulence factors and are especially interesting in the parasitic worm infection process. Some NF strains’ success implies that they have different host preferences. Finally, we suggest that NF is a potential alternative to synthetic pesticides in the management of plant-parasitic nematodes, and that they may be more effective in making agriculture sustainable by replacing hazardous chemicals and mitigating the effects of their residues on the environment.
References
Ahman J, Ek B, Rask L, Tunlid A (1996) Sequence analysis and regulation of a gene encoding a cuticle-degrading serine protease from the nematophagous fungus Arthrobotrys oligospora. Microbiology 142:1605–1616
Ahren D, Tholander M, Fekete C, Rajashekar B, Friman E, Johansson T et al (2005) Comparison of gene expression in trap cells and vegetative hyphae of the nematophagous fungus Monacrosporium haptotylum. Microbiology 151:789–803
Aschner M, Kohn S (1958) The biology of Harposporium anguillulae. Microbiology 19(1):182–189
Ashraf MS, Khan TA (2010) Integrated approach for the management of Meloidogyne javanica on eggplant using oil cakes and biocontrol agents. Arch Phytopathol Plant Protect 43(6):609–614
Baheti BL, Dodwadiya M, Bhati SS (2017) Eco-friendly management of maize cyst nematode, Heterodera zeae on sweet corn (Zea mays L. saccharata). J Entomol Zool Stud 5(6):989–993
Baron CN, Souza-Pollo A, Rigobelo E (2020) Purpureocillium lilacinum and Metaehizium marquandii as plant growth-promoting fungi. Peer Rev J 8:e9005
Barron GL (1977) The nematode destroying fungi. Canadian Biological Publication Ltd., Guelph, p 140
Barron GL (1992) Lignolytic and cellulolytic fungi as predators and parasites. In: Carroll GC, Wicklow DT (eds) The fungal community, its organization and role in the ecosystems. Marcel Dekker, New York, pp 311–326
Basse CW, Steinberg G (2004) Ustilago maydis, model system for analysis of the molecular basis of fungal pathogenicity. Mol Plant Pathol 5(2):83–92
Bird AF, Bird J (1991) The structure of nematodes, 2nd edn. Academic, New York
Bordallo JJ, Lopez-Llorca LV, Jansson HB, Salinas J, Persmark L, Asensio L (2002) Colonization of plant roots by egg-parasitic and nematode-trapping fungi. New Phytol 154(2):491–499
Brahma U, Borah A (2016) Management of Meloidogyne incognita on pea with bioagents and organic amendment. Indian J Nematol 46(1):58–61
Chaverri P, Samuels GJ, Hodge KT (2005) The genus Podocrella and its nematode-killing anamorph Harposporium. Mycologia 97(2):433–443
Chen YY, Cheng CY, Huang TL, Li YK (2005) Chitosanase from Paecilomyces lilacinus with binding affinity for specific chitooligosaccharides. Biotechnol Appl Biochem 41(2):145–150
Clarke AJ, Cox PM, Shepherd AM (1967) The chemical composition of the egg shells of the potato cyst-nematode, Heterodera rostochiensis woll. Biochem J 104(3):1056–1060
Decraemer W, Hunt DJ (2006) Structure and classification. In: Perry RN, Moens M (eds) Plant nematology. CABI Publishing, Wallingford, pp 3–32
Devi TS, Mahanta B, Borah A (2016) Comparative efficacy of Glomus fasciculatum, Trichoderma harzianum, carbofuran and carbendazim in management of Meloidogyne incognita and Rhizoctonia solani disease complex on brinjal. Indian J Nematol 46:161–164
Dhawan SC, Singh S (2009) Compatibility of Pochonia chlamydosporia with nematicide and neem cake against root-knot nematode, meloidogyne incognita infesting okra. Indian J Nematol 39(1):85–89
Drechsler C (1937) Some hyphomycetes that prey on free living terricolous nematodes. Mycologia 29:447–552
Escudero N, Lopez-Llorca LV (2012) Effects on plant growth and root-knot nematode infection of an endophytic GFP transformant of the nematophagous fungus Pochonia chlamydosporia. Symbiosis 57(1):33–42
Feyisa B, Lencho A, Selvaraj T, Getaneh G (2015) Evaluation of some botanicals and Trichoderma harzianum for the management of tomato root-knot Nematode (Meloidogyne incognita) (Kofoid and White) Chitwood. Adv Crop Sci Technol 4:201
Fresenius G (1852) Beitrage zur mykologie. Heft 1-2:1–80
Gogoi D, Mahanta B (2013) Comparative efficacy of Glomus fasciculatum, Trichoderma harzianum, carbofuran and carbendazim in management of Meloidogyne incognita and Rhizoctonia solani disease complex on French bean. Ann Plant Prot Sci 21(1):72–175
Gupta R, Tiwari S, Saikia SK, Shukla V, Singh R, Singh SP, Kumar PV, Pandey R (2015a) Exploitation of microbes for enhancing bacoside content and reduction of M. incognita infestation in B. monnieri L. Protoplasma 252(1):53–61
Gupta R, Saikia SK, Pandey R (2015b) Bioconsortia augments antioxidant and yield in Matricaria recutita L. against M. incognita (Kofoid and White) Chitwood infestation. Proc Natl Acad Sci India Sect B Biol Sci 87(2):335–342
Hastuti LDS, Faull J (2018) Wheat bran soil inoculant of sumateran nematode-trapping fungi as biocontrol agents of the root-knot nematode Meloidogyne incognita on deli tobacco (nicotiana tabaccum l) cv. deli 4. IOP Conf Ser 130(1):012009)
Hsueh YP, Gronquist MR, Schwarz EM, Nath RD, Lee CH, Gharib S, Schroeder FC, Sternberg PW (2017) Nematophagous fungus Arthrobotrys oligospora mimics olfactory cues of sex and food to lure its nematode prey. elife 6:e20023
Huang WK, Sun JH, Cui JK, Wang GF, Kong LA, Peng H, Chen SL, Peng DL (2014) Efficacy evaluation of fungus Syncephalastrum racemosum and nematicide avermectin against the RKN M incognita on cucumber. PLoS One 9(2):e89717
Indian Economy (2004) Special issue updated with economic survey 2004–05 and budget 2005. Published by Pearson Education; 15th Floor World Trade Tower, C01, Sector 16, Noida, Uttar Pradesh
Jansson H-B (1987) Receptors and recognition in nematodes. In: Veech J, Dickson D (eds) Vistas on nematology. Society of Nematologists, Hyattsville, pp 153–158
Jansson HB, Friman E (1999) Infection-related surface proteins on conidia of the nematophagous fungus Drechmeria coniospora. Mycol Res 103(2):249–256
Jansson HB, Lopez-Llorca LV (2001) Biology of nematophagous fungi. In: Misra JK, Horn BW (eds) Mycology: trichomycetes, other fungal groups and mushrooms. Science Publishers, Enfield, pp 145–173
Jansson HB, Nordbring-Hertz B (1979) Attraction of nematodes to living mycelium of nematophagous fungi. Microbiology 112(1):89–93
Jansson H-B, Nordbring-Hertz B (1983) The endoparasitic fungus Meria coniospora infects nematodes specifically at the chemosensory organs. J Gen Microbiol 129:1121–1126
Jansson H-B, Nordbring-Hertz B (1984) Involvement of sialic acid in nematode chemotaxis and infection by an endoparasitic nematophagous fungus. J Gen Microbiol 130:39–43
Jansson HB, Nordbring-Hertz B (1988) Infection events in the fungus-nematode system. Dis Nematodes 2:59–72
Jansson HB, Thiman L (1992) A preliminary study of chemotaxis of zoospores of the nematode-parasitic fungus Catenaria anguillulae. Mycologia 84(1):109–112
Jansson H-B, Jeyaprakash A, Damon RA, Zuckerman BM (1984) Caenorhabditis elegans and Panagrellus redivivus: enzyme-mediated modification of chemotaxis. Exp Parasitol 58:270–277
Jeyaprakash A, Jansson HB, Marban-Mendoza N, Zuckerman BM (1985) Caenorhabditis elegans: lectin-mediated modification of chemotaxis. Experimental Parasitol 59(1):90–97
Kendrick B (2001) The fifth kingdom, 3rd edn. Focus Publishing, Newburyport
Kim YH (2015) Predatory nematodes as biocontrol agents of phytonematodes. In: Askary TK, Martinelly PRP (eds) Biocontrol agents of phytonematodes. CABI, Wallingford, pp 393–420
Koenning SR, Overstreet C, Noling JW, Donald PA, Becker JO, Fortnum BA (1999) Survey of crop losses in response to phytoparasitic nematodes in the United States for 1994. J Nematol 31(4):587–618
Larriba E, Jaime MD, Nislow C, Martín-Nieto J, Lopez-Llorca LV (2015) Endophytic colonization of barley (Hordeum vulgare) roots by the nematophagous fungus Pochonia chlamydosporia reveals plant growth promotion and a general defense and stress transcriptomic response. J Plant Res 128:665–678
Lee N, D’Souza CA, Kronstad JW (2003) Of smuts, blasts, mildews, and blights: cAMP signaling in phytopathogenic fungi. Annu Rev Phytopathol 41:399–427
Li J, Yang J, Huang X, Zhang KQ (2006) Purification and characterization of an extracellular serine protease from Clonostachys rosea and its potential as a pathogenic factor. Process Biochem 41:925–929
Lopez-Llorca LV (1990a) Purification and properties of extracellular proteases produced by the nematophagous fungus Verticillium suchlasporium. Can J Microbiol 36:8
Lopez-Llorca LV (1990b) Purification and properties of extracellular proteases produced by the nematophagous fungus Verticillium suchlasporium. Can J Microbiol 36:530–537
Lopez-Llorca LV, Claugher D (1990) Appressoria of the nematophagous fungus Verticillium suchlasporium. Micron Microsc Acta 21:125–130
Lopez-Llorca LV, Olivares-Bernabeu C, Salinas J, Jansson HB, Kolattukudy PE (2002) Prepenetration events in fungal parasitism of nematode eggs. Mycol Res 106:499–506
Luns FD, Assis RCL, Silva LPC, Ferraz CM, Braga FR, Araujo JVD (2018) Coadministration of nematophagous fungi for biocontrol over nematodes in bovine in the South-Eastern Brazil. Biomed Res Int 2018:2934674
Luo H, Mo M, Huang X, Li X, Zhang K (2004) Coprinus comatus: a basidiomycete fungus forms novel spiny structures and infects nematodes. Mycologia 96:1218–1225
Luo H, Liu Y, Fang L, Li X, Tang N, Zhang K (2007) Coprinus comatus damages nematode cuticles mechanically with spiny balls and produces potent toxins to immobilize nematodes. Appl Environ Microbiol 73(12):3916–3923
Mei X, Wang X, Li G (2021) Pathogenicity and volatile nematicidal metabolites from Duddingtonia flagrans against meloidogyne incognita. Microorganisms 9(11):2268
Money NP (1998) Mechanics of invasive fungal growth and the significance of turgor in plant infection. In: Molecular genetics of host-specific toxins in plant disease. Kluwer Academic Publishers, Dordrecht, pp 261–271
Muthulakshmi M, Kumar S, Subramanian S, Anita B (2012) Compatibility of Pochonia chlamydosporia with other biocontrol agents and carbofuran. J Biopest 5:243–245
Narasimhamurthy HB, Ravindra H, Sehgal M (2017) Management of rice root knot nematode, Meloidogyne graminicola. Int J Pure Appl Biosci 5:268–276
Nicol J, Turner D, Coyne L, den Nijs L, Hockland S, Maafi Z (2011) Current nematode threats to world agriculture. In: Jones J, Gheysen G, Fenoll C (eds) Genomics and molecular genetics of plant-nematode interactions. Springer, Berlin, pp 21–43
Nordbring-Hertz B (2004) Morphogenesis in the nematode-trapping fungus Arthrobotrys oligospora - an extensive plasticity of infection structures. Mycologist 18(3):125–133
Nordbring-Hertz B, Mattiasson B (1979) Action of a nematode-trapping fungus shows lectin mediated host-microorganism interaction. Nature 281:477–479
Nordbring-Hertz B, Jansson HB, Tunlid A (2006) Nematophagous fungi. In: Encyclopedia of life sciences. Wiley, Hoboken, pp 1–11
Parihar K, Rehman B, Ganai MA, Asif M, Siddiqui MA (2015) Role of oil cakes and Pochonia chlamydosporia for the management of Meloidogyne javanica attacking Solanum melongena L. J Plant Pathol Microbiol 1:1–5
Pfister DH, Liftik ME (2018) Two Arthrobotrys anamorphs from Orbilia auricolor. Mycologia 87(5):684–688
Pramer D (1964) Nematode-trapping fungi. Science 144:382–388
Raveendra HR, Krishna MR, Mahesh KR (2011) Management of root knot nematode Meloidogyne incognita by using oil cake, bioagent, trap crop, chemicals and their combination. Int J Sci Nat 2:519–523
Sasser JN (1989) Plant parasitic nematodes: the farmer’s hidden enemy. North Carolina State University, Raleigh, p 115
Sasser JN, Freckman DW (1987) A world perspective on nematology: the role of the society. In: Veech JA, Dickson DW (eds) Vistas on nematology. Society of Nematologists Inc., Hyattsville, pp 7–14
Schmidt AR, Dorfelt H, Perrichot V (2007) Carnivorous fungi from Cretaceous Amber. Science 318:1743
Scholler M, Hagedorn G, Rubner A (1999) A reevaluation of predatory orbiliaceous fungi. II. A new generic concept. Sydowia 51:89–113
Segers R, Butt TM, Kerry BR, Peberdy F (1994) The nematophagous fungus Verticillium chlamydosporium Goddard produces a chymoelastase-like protease which hydrolyses host nematode proteins in situ. Microbiology 140:2715–2723
Shamalie BVT, Fonseka RM, Rajapaksha RGAS (2011) Effect of Trichoderma viride and carbofuran (Curator®) on management of root-knot nematodes and growth parameters of Gotukola (Centella asiatica L.). Trop Agric Res 23:61–69
Singh UB (2007) Occurrence, characterization and performance of some predacious fungi. M.Sc. Thesis. Hindu University, Varanasi, India
Singh S (2013) Integrated approach for the management of the root-knot nematode, Meloidogyne incognita, on eggplant under field conditions. Nematology 15:747–757
Singh UB, Sahu A, Sahu N, Singh RK, Renu RP, Singh DP, Sharma BK, Manna MC (2012a) Co-inoculation of Dactylaria brochopaga and Monacrosporium eudermatum affects disease dynamics and biochemical responses in tomato (Lycopersicon esculentum Mill.) to enhance bioprotection against Meloidogyne incognita. Crop Prot 35:102–109
Singh UB, Sahu A, Singh RK, Singh DP, Meena Kamlesh K, Srivastava JS, Renu Manna MC (2012b) Evaluation of biocontrol potential of Arthrobotrys oligospora against Meloidogyne graminicola and Rhizoctonia solani in Rice (Oryza sativa L.). Biol Control 60:262–270
Singh UB, Sahu A, Sahu N, Singh BP, Singh RK, Renu S, Jaiswal RK, Sharma BK, Singh HB, Manna MC, Subba Rao A, Prasad RS (2013) Can endophytic Arthrobotrys oligospora modulate accumulation of defence related biomolecules and induced systemic resistance in tomato (Lycopersicon esculentum Mill.) against root knot disease caused by Meloidogyne incognita. Appl Soil Ecol 63:45–56
Singh UB, Singh S, Malviya D, Chaurasia R, Mohd I, Rai JP, Sharma AK (2017) Harnessing biocontrol potential of Trichoderma harzianum for control Meloidogyne incognita in tomato. Indian Phytopathol 70:331–335
Singh UB, Malviya D, Singh S, Kumar M, Sahu PK, Singh HV, Kumar S, Roy M, Imran M, Rai JP, Sharma AK (2019a) Trichoderma harzianum and methyl jasmonate-induced resistance to Bipolaris sorokiniana through enhanced phenylpropanoid activities in bread wheat (Triticum aestivum L.). Front Microbiol 10:1697
Singh UB, Singh S, Khan W, Malviya D, Sahu PK, Chaurasia R, Sharma SK, Saxena AK (2019b) Drechslerella dactyloides and Dactylaria brochopaga mediated induction of defense related mediator molecules in tomato plants pre-challenged with Meloidogyne incognita. India Phytopathol 72:309–320
Soliman MS, El-Deriny MM, Ibrahim DSS, Zakaria H, Ahmed Y (2021) Suppression of root-knot nematode Meloidogyne incognita on tomato plants using the nematode trapping fungus Arthrobotrys oligospora Fresenius. J Appl Microbiol 131(5):2402–2415
Sorokin N (1876) Note sur les vegetaux parasites des Anguillulae. Ann Sci Nat Bot Ser 6(4):62–71
St. Leger RJ (1993) Biology and mechanism of insect-cuticle invasion by Deuteromycete fungal pathogens. In: Beckage NE, Thompson SN, Federici BA (eds) Parasites and pathogens of insects: pathogens, vol 2. Academic, San Diego, pp 211–229
Stirling GR (1991) Biological control of plant parasitic nematodes: progress, problems and prospects. CAB International, Wallingford
Suarez B, Rey M, Castillo P, Monte E, Llobell A (2004) Isolation and characterization of PRA1, a trypsin-like protease from the biocontrol agent Trichoderma harzianum CECT 2413 displaying nematicidal activity. Appl Microbiol Biotechnol 65:46–55
Tikhonov VE, Lopez-Llorca LV, Salinas J, Jansson HB (2002) Purification and characterization of chitinases from the nematophagous fungi Verticillium chlamydosporium and V. suchlasporium. Fungal Genet Biol 35(1):67–78
Tunlid A, Johansson T, Nordbring-Hertz B (1991a) Surface polymers of the nematode-trapping fungus Arthrobotrys oligospora. J Gen Microbiol 137:1231–1240
Tunlid A, Nivens DE, Jansson HB, White DC (1991b) Infrared monitoring of the adhesion of Catenaria anguillulae zoospores to solid surfaces. Exp Mycol 15:206–214
Tunlid A, Jansson H-B, Nordbring-Hertz B (1992) Fungal attachment to nematodes. Mycol Res 96:401–412
Wang M, Yang J, Zhang KQ (2006a) Characterization of an extracellular protease and its cDNA from the nematode-trapping fungus Monacrosporium microscaphoides. Can J Microbiol 52:130–139
Wang RB, Yang JK, Lin C, Zhang Y, Zhang KQ (2006b) Purification and characterization of an extracellular serine protease from the nematode-trapping fungus Dactylella shizishanna. Lett Appl Microbiol 42:589–594
Wang X, Li GH, Zou CG, Ji XL, Liu T, Zhao PJ, Liang LM, Xu JP, An ZQ, Zheng X, Qin YK (2014) Bacteria can mobilize nematode-trapping fungi to kill nematodes. Nat Commun 5(1):1–9
Ward S, Thomson N, White JG, Brenner S (1975) Electron microscopical reconstruction of the anterior sensory anatomy of the nematode Caenorhabditis elegans. J Comp Neurol 160:313–337
Woronin MS (1870) Sphaeria lemaneae, Sordaria coprophila, Arthrobotrys oligospora. Nat Ges 7:325
Yang J, Huang X, Tian B, Sun H, Duan J, Wu W, Zhang M (2005a) Characterization of an extracellular serine protease gene from the nematophagous fungus Lecanicillium psalliotae. Biotechnol Lett 27:1329–2334
Yang J, Huang X, Tian B, Wang M, Niu Q, Zhang K (2005b) Isolation and characterization of a serine protease from the nematophagous fungus Lecanicillium psalliotae, displaying nematicidal activity. Biotechnol Lett 27:1123–1128
Yang JK, Tian BY, Liang LM, Zhang KQ (2007) Extracellular enzymes and the pathogenesis of nematophagous fungi. Appl Microbiol Biotechnol 75:21–31
Yu’e H, Mo M, Su H, Zhang K (2005) Ecology of aquatic nematode-trapping hyphomycetes in southwestern China. Aquat Microb Ecol 40:175–181
Zare R, Gams W (2001) A revision of Verticillium section Prostrata. IV. The genera Lecanicillium and Simplicillium gen. nov. Nova Hedwigia 73:1–50
Zare R, Gams W, Evans HC (2001) A revision of Verticillium section Prostrata. V. The genus Pochonia, with notes on Rotiferophthora. Nova Hedwigia 73:51–86
Zhang Y, Zhang K-Q, Hyde KD (2014) The ecology of nematophagous fungi in natural environments. Nematode-trapping fungi. Springer, Dordrecht, pp 211–229
Zhang Z, Zhang X, Jhao J, Zhang X, Liang W (2015) Tillage and rotation effects on community composition and metabolic footprints of soil nematodes in a black soil. Eur J Soil Biol 66:40–48
Zhang Y, Li S, Li H, Wang R, Zhang KQ, Xu J (2020a) Fungi-nematode interactions: diversity, ecology, and biocontrol prospects in agriculture. J Fungi 6:1–24
Zhang FA, Liu S-R, Zhou X-J et al (2020b) Fusarium xiangyunensis (Nectriaceae) a remarkable new species of the Nematophagous fungi from Yunnan, China. Phytotaxa 450:273–284
Zhao M, Mo M, Zhang Z (2004) Characterization of a serine protease and its full-length cDNA from the nematode-trapping fungus Arthrobotrys oligospora. Mycologia 96:16–22
Zopf W (1888) Zur. Kenntnis der infections-Krankheiten niederer Tiere. Nova Acta Leop Acad Nat Halle 52:7
Zuckerman BM (1983) Hypothesis and possibilities of intervention in nematode chemoreceptors. J Nematol 15:173–182
Zuckerman BM, Jansson H-B (1984) Nematode chemotaxis and mechanisms of host/prey recognition. Annu Rev Phytopathol 22:95–113
Acknowledgments
The authors sincerely thank the Director of ICAR-NBAIM, Mau, for providing scientific and technical support during preparation of the manuscript. The authors gratefully acknowledge the Science and Engineering Research Board, Department of Science and Technology, Government of India, for providing financial support for the study.
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The authors declare that they have no known competing financial interest or personal relationship that could have appeared to influence the content reported in this manuscript. The authors declare no conflict of interest.
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This research was supported by Science and Engineering Research Board, Department of Science and Technology, New Delhi (India).
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Yadav, B. et al. (2023). Nematophagous Fungi: Biology, Ecology and Potential Application. In: Singh, U.B., Kumar, R., Singh, H.B. (eds) Detection, Diagnosis and Management of Soil-borne Phytopathogens. Springer, Singapore. https://doi.org/10.1007/978-981-19-8307-8_12
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