Abstract
Plant parasitic nematodes cause economic damage to food crops throughout the world. Many harmful chemical pesticides have been withdrawn due to environmental toxicity and health effects resulting in the development of non-chemical alternatives for nematode pest management. Biological control based on nematophagous fungi has marked remarkable success in various parts of the world. The widely used fungi include Purpureocillium lilacinum, Pochonia chlamydosporia, and Trichoderma harzianum, Arthrobotrys, Hirsutella, etc. for the management of root knot, cyst, and other groups of plant parasitic nematodes. Improvisation in quality of the product, mass production of fungi, and improvement of delivery systems will increase the usage of fungal products. The molecular mechanisms of the interaction between nematode and fungal species can be understood with the application of molecular biology tools, which further helps to develop new screening procedures for nematophagous fungi to control plant parasitic nematodes. However, the use of nematophagous fungi along with other practices in integrated nematode management systems can ensure sustainable control.
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10.1 Introduction
Plant parasitic nematodes are considered to be the major constraint on plant productivity, and they have been found to attack almost every part of the plant including roots, stems, leaves, fruits, and seeds. They are also responsible for invasion of secondary pests and pathogens by causing wounds in the host roots (Caboni et al. 2016). Global annual yield losses from plant parasitic nematodes have been estimated at 12.6% ($215.77 billion) for top 20 life sustaining crops (Abd-Elgawad and Askary 2015). Among different groups of plant parasitic nematodes, root knot (Meloidogyne spp.) and cyst nematodes (Heterodera and Globodera spp.) are economically important by forming complex feeding structures like giant cells and syncytia in the roots of their hosts which act as a nutrient sink to the nematode. Apart from these, root-lesion nematodes (Pratylenchus spp.), burrowing nematode (Radopholus similis), stem nematode (Ditylenchus dipsaci), reniform nematode (Rotylenchulus reniformis), and several other plant parasitic nematodes causing economic damage to the crops (Jones et al. 2013).
Many traditional chemical pesticides such as fumigants, organophosphates, and carbamates have been used for the management of plant parasitic nematodes (Kour et al. 2020). However, the usage of harmful chemical pesticides cause environmental toxicity, health effects and further withdrawal of some pesticides has led to the development and adoption of non-chemical alternatives for the management of plant parasitic nematodes (Davies and Curtis 2011; Kumar et al. 2018). In addition, these chemicals do not affect the development of nematode eggs in soil, since the egg shell of nematode acts as a barrier which makes them resistant to chemical pesticides (Baron et al. 2019). Crop rotation and use of resistant cultivars have been used as the management strategies for plant parasitic nematodes. However, these practices are limited by the size of growing area, wide host range of nematode species, and high genetic diversity among or within nematode populations (Berlitz et al. 2014; Li et al. 2015).
Biological control , an alternative for the management of plant parasitic nematodes. The usage of biocontrol agents is increasing these days, due to their reduced environmental toxicity, target specificity, and safety to non-target organisms (Kumar and Singh 2015; Jiang et al. 2017). These biocontrol agents include fungi, bacteria, viruses, nematodes, and other invertebrates which have an antagonistic activity against plant parasitic nematodes. The market for biological nematicides has risen by nearly 20% from 2009 to the present, according to the Markets and Markets report, with most of the development in field crops. Nematicide sales amounted to just over $1 billion in 2014 and are anticipated to expand at a compound annual growth rate (CAGR) of 2.7% over the next 10 years to $1.3 billion over 20 years (Jiang et al. 2017).
Currently several biocontrol agents such as fungi and bacteria have been widely used as commercial products for the management of plant parasitic nematodes. Fungal antagonists have been considered as the most promising biocontrol agents and attention is being paid to the exploitation of fungi to control plant parasitic nematodes (Stirling 2011; Moosavi and Zare 2012; Lamovsek et al. 2013). Among different fungal biocontrol agents, the most widely studied species for nematode management include Pochonia chlamydosporia, Purpureocillium lilacinum, Arthrobotrys oligospora, Trichoderma spp. and Verticillium spp. (Rastegari et al. 2020a, b; Yadav et al. 2020a, b). This chapter reviews different categories of nematophagous fungi, mode of action, and knowledge of the interactions between nematodes and nematode trapping fungi and commercialization of different fungal products for the management of plant parasitic nematodes.
10.2 Nematophagous Fungi
This group of fungi has been widely used as biocontrol agents of plant parasitic nematodes due to their antagonistic, parasitic, and predatory action against different species. There are approximately 700 fungal species reported to be able to attack different life stages of nematodes (juveniles, adults, and eggs) (Li et al. 2015; Jiang et al. 2017). Over the decades, several researchers have worked on fundamental studies and their potential as biocontrol agents against different plant parasitic nematodes. Based on their infection mechanism, they are subdivided into four categories viz., nematode trapping fungi, endoparasitic fungi, egg and female parasitic fungi, and toxin producing fungi (Dackman et al. 1992; Nordbring-Hertz et al. 2006) (Table 10.1).
10.2.1 Nematode Trapping Fungi
These are soil borne fungi that have the ability to capture and kill nematodes by using trapping structures, including adhesive hyphae, adhesive knobs, constricting rings and non-constricting rings, and adhesive networks (Nordbring-Hertz et al. 2011; Jiang et al. 2017). The type of trapping structure formed mainly depends on the fungal species or strains of fungal species and both biotic and abiotic environmental factors (Nordbring-Hertz et al. 2006). Based on phylogenetic analysis results using ribosomal RNAs (rRNAs) and protein coding genes, these trapping devices could serve as indicators for generic delimitation among these fungi (Ahren et al. 1998; Scholler et al. 1999; Li et al. 2005). There are about 380 species of nematode trapping fungi have been reported from different regions of the world (Zhang et al. 2011). They are usually non-host specific and can infect all soil dwelling nematodes. All the known nematode trapping fungi belong to the order Orbiliales of the phylum Ascomycota (Zhang and Hyde 2014). These fungi can live both saprophytically on organic matter and as predatory by capturing tiny animals. However, they can shift from saprophytic to predatory lifestyle in the presence of nematode prey by the production of trapping structures (Li et al. 2011; Yang et al. 2012; Zhang and Hyde 2014). The application of these fungi for nematode suppression was accompanied by addition of large amounts of organic matter and in most cases the levels of nematode control were unpredictable (Stirling 1991).
10.2.1.1 Adhesive Hyphae
These are mostly produced by lower fungi belongs to the genera Stylopage and Cystopage under phylum Zygomycota (Table 10.1). They cannot produce complex devices for capturing nematodes due to the absence of septa (Barron 1977). However, septate fungi such as Arthrobotrys botryospora, Dactylaria psychrophila, and A. superba can capture nematodes with adhesive hyphae under certain conditions (Chen and Dickson 2004). The infection begins with the capture organ being adhered by nematode. Within few minutes, the nematodes added to a fungal colony were captured and penetrated within 1 h by the trap forming cell hyphae. After 6 h, the whole nematode body was filled with hyphae. The dead nematode did not induce traps but developed a large number of thin hyphae around the nematode.
10.2.1.2 Adhesive Knobs
These are very effective trapping structures and nematodes are often attacked at more than one infection site (Barron 1977). These knobs are covered by a thin layer of adhesive materials and the infection process involves one knob and several associated hyphal elements. These adhesive knobs produce an infection peg and infection bulb. From this bulb, the infection hyphae proliferate within the nematode and digest the inner contents. The adhesive knobs are found in Ascomycota and Basidiomycota (Table 10.1). This group of fungi is unique in forming clamp connections on the secondary hyphae (Chen and Dickson 2004). The genomic studies of adhesive knob forming species, Monacrosporium haptotylum indicated the enrichment of small secreted proteins (SSPs) that were highly and differentially expressed during the interaction with nematode hosts (Meerupati et al. 2013).
10.2.1.3 Adhesive Networks
This is the most common type of nematode trapping structure observed in Arthrobotrys spp. (Table 10.1). The trap may consist of a single ring or a three-dimensional network. A. oligospora, a model organism for understanding the interaction between fungi and nematodes, forms a three-dimensional adhesive network during infection process. The genomic and proteomic analyses of this fungus provide insights into nematode trap formation including G-protein coupled receptors, adhesive proteins, cell division cycle, peroxisome related proteins, and proteins involved in energy supplementation (Yang et al. 2011).
10.2.1.4 Constricting Rings
Among different trapping structures , constricting rings are probably the only ones that actually capture the nematodes and considered to be the lineal strategy after which other trapping structures evolved (Yang et al. 2007a; Liu et al. 2012). In general, trapping fungi that form constricting rings are more abundant in soils rich in organic matter and influenced by the population density of nematodes (Linford 1937; Gray 1985). However, the trapping mechanism is completely different in contrast to others. When a nematode passing into the ring, it stimulates the inner face of the ring such that the three cells comprising the ring inflate centripetally to about three times and get trapped within 0.1 s. The pressure exerted by nematode on the ring during this process causes the activation of G-protein coupled receptors which leads to an increase in cytoplasmic Ca2+, and activation of calmodulin that opens the water channels, and thereby inflates the ring trapping the nematode (Chen et al. 2001). The genome sequence of Drechslerella stenobrocha constricting ring forming fungus revealed that the downregulation of saprophytic enzyme genes and the upregulation of infection related genes during the capture of nematodes indicated a transition between saprophagous and predatory life strategies. This study also indicated that protein kinase C (PKC) signal pathway and Zn (2)-C6 type transcription factors were responsible for trap formation in D. stenobrocha (Liu et al. 2014). Following the capture an infection peg is produced by one of the three ring cells and a small globose infection bulb develops after the penetration of infection peg into the body cavity of the nematodes. Trophic hyphae quickly digest and absorb the internal contents of nematode leaving only the cuticle structures .
10.2.1.5 Non-constricting Rings
Non-constricting rings are generally produced by erect lateral branches produced from septate hyphae. When a nematode thrusts its head inside the non-constricting ring, it gets wedged and held by friction. During this, the ring often breaks at the point of weakness near the stalk apex and this process may be repeated until several rings are wedged around the host. Following capture, the fungi penetrate and consume the inner contents of the host (Zaki 1994). Dactylaria candida and D. lysipaga usually generally form non-constricting rings during infection process (Table 10.1).
10.2.2 Endoparasitic Fungi
These are a group of fungi that infect nematodes mainly by conidia (Drechmeria coniospora), ingestive spores (Harposporium spp.), adhesive spores (Nematoctonus concurrens) or zoospores (Catenaria anguillulae) (Table 10.1). Their spores either adhere to the nematode cuticle or swallowed by the host and then germinate inside the nematode body which finally result in the death of nematode. In D. coniospora surface proteins and chymotrypsin-like proteases were involved in the infection process (Lopez-Llorca et al. 2008; Jansson and Friman 1999). Although some of these like Nematophthora gynophila could parasitize a large number of nematodes viz., Heterodera avenae, H. trifolii, H. schachtii, H. goettengiana, H. cruciferae, etc., and their bioefficacy was dependent on adequate soil moisture and on the relative number of nematodes and endozoic units present in the soil. Besides, their obligate nature posed a problem for mass production (Kerry 1987).
10.2.3 Egg and Female Parasitic Fungi
The research on egg and female parasitic fungi has been initiated in the 1990s. These fungi use either appressoria (Purpureocillium spp., Pochonia spp.,) or zoospores (Nematophthora gynophila) to kill different life stages (Lopez-Llorca et al. 2008) (Table 10.1). The relationship between nematodes and these fungi was variable, but as some of the isolates were highly virulent, they were extensively used and are still available as formulations in the market, especially for sedentary endoparasites. However, it is important to ascertain the quality parameters, namely colony forming units (CFU), presence of contaminants of these formulations before application in the field for control of nematode population. Among different egg parasitic fungi, P. chlamydosporia and P. lilacinum are widely used as biocontrol agents.
10.2.3.1 Pochonia chlamydosporia
P. chlamydosporia is considered as potential biocontrol agent of endoparasitic nematodes due to their ability to colonize the rhizosphere of plants and cultivars, to produce chlamydospores in vitro and to infect eggs of endoparasitic nematodes such as root knot (Meloidogyne spp.) and cyst nematodes (Heterodera avenae, H. glycines, H. schachtii) (Manzanilla-Lopez et al. 2013; Dallemole-Giaretta et al. 2015). In addition, this fungus has shown activity against false root-knot nematode, Nacobbus aberrans, burrowing nematode, Radopholus similis, citrus nematode, Tylenchulus semipenetrans, and reniform nematode, Rotylenchulus spp. (Manzanilla-Lopez et al. 2013; Abd-Elgawad and Askary 2018). The specificity of this fungus towards sedentary endoparasitic nematodes is associated with the recognition of both quantitative and qualitative changes in root exudates patterns during nematode infection (Wang and Bergeson 1974). Crops such as beans, cabbage, crotalaria, kale, pigeon pea, potato, pumpkin, and tomato are considered as good hosts (200 CFU/cm2 of root) for colonization of this fungus in the rhizosphere. Whereas, chilli, sweet potato, cowpea, rye, tobacco, and cotton are considered as moderate hosts (100–200 CFU/cm2 of root) and poor hosts (<100 CFU/cm2 of root) include aubergine, okra, soybean, sorghum, and wheat (de Leij 1992; Bourne et al. 1996).
P. chlamydosporia infects nematode eggs through the development of appressoria at the hyphal tip and the presence of mucilaginous material between appressoria and surface of the egg shell appear to assist in egg shell penetration (Lopez-Llorca et al. 2002). A post infection bulb leads to the development of a mycelium within the egg and destruction of internal contents (Segers 1996; Kerry and Hirsch 2011). The fungus produce extracellular enzymes such as Serine proteases (VCP1) and chitinases which are involved in degradation of egg shells (Segers 1996). SCP1, a serine carboxypeptidase from P. chlamydosporia was immunolocalized in M. javanica eggs infected by fungus (Escudero et al. 2016).
The ability to produce chlamydospores in vitro without the addition of other energy sources lead to the consideration of this fungus as biocontrol agent. Hence application of chlamydospores at the rate of 5000 spores/g soil in aqueous suspension is recommended to test the efficacy of fungus but it may vary according to the strain and target nematode (de Leij et al. 1992; Stirling and Smith 1998; Bourne and Kerry 2000; Kerry and Hidalgo-Diaz 2004). Application of organic amendments may increase fungal abundance in soil and fungus remains in saprophytic phase but not necessarily increase in nematode parasitic activity (Atkins et al. 2003). Gene expression studies showed differences among several genes involved in cellular signals, transport or DNA repair, with a distinct cluster of genes commonly expressed during transition from saprophytic to parasitic phase of this fungus (Rosso et al. 2011). Niu (2017) reviewed that the secondary metabolites released from P. chlamydosporia and other species such as resorcylic acid lactone, pyranones, alkaloid, phenolics, etc. may act as virulence factors to plant and animal-parasitic nematodes parasitized by the fungus .
10.2.3.2 Purpureocillium lilacinum (=Paecilomyces lilacinus)
P. lilacinum is a soil inhabiting fungus reported from different parts of the world and widely used as potential biocontrol agent against plant parasitic nematodes (Vasanthi and Kumaraswamy 1999; Brand et al. 2010). This fungus infects eggs, egg masses, females, and cysts of many plant parasitic nematodes such as Meloidogyne spp., Globodera spp., Heterodera spp., Tylenchulus sp., and Nacobbus sp. The fungus first colonizes gelatinous matrix of different nematodes and penetrate the eggs with the help of appressoria or hyphae. In addition, the fungus produce several extracellular enzymes such as serine proteases, chitinases depending on the recognition of the host surface hydrophobicity and these were purified and characterized from different strains (Bonants et al. 1995; Lopez-Llorca et al. 2002; Khan et al. 2003; Huang et al. 2004; Khan et al. 2004). Enzymes produced by P. lilacinum strain 251 resulted in reduction of egg hatching in M. javanica (Khan et al. 2004). Similarly, the secondary metabolites produced by the fungi (Australian isolate), such as leucinostatins, have significant effect on the colonization of M. javanica eggs (Park et al. 2004).
In addition to control of plant parasitic nematodes, species of this fungus produce several metabolites which promote plant growth and defensive substances against biotic and abiotic stresses (Khan et al. 2012). Further, this fungus promote plant growth by performing Phosphorus solubilization in soil (Rinu and Pandey 2011; Lima-Rivera et al. 2016).
10.2.4 Toxin Producing Fungi
This group of fungi produces toxic metabolites which act as nematicidal toxins before the penetration of hyphae into nematode body through cuticle. The toxic effects on nematodes include reduced egg hatching, immobility, mortality, etc. The productions of toxins also help in preventing the consumption of fungal colonies by fungivorous nematodes and other invertebrates (de Freitas Soares et al. 2018). So far more than 200 compounds have been identified from nearly 280 fungal species (Li et al. 2007a; Li and Zhang 2014). The commonly included genera under this category are Nematoctonus, Pleurotus, and Penicillium, etc. (de Freitas Soares et al. 2018). These toxic metabolites include peptides, terpenoids, alkaloids, quinines, sterols, etc. (Li and Zhang 2014). Table 10.3 shows a list of toxic metabolites identified in different fungi and their effect on different nematode species . The discovery of these toxic metabolites further lead to the development of these chemicals as biocontrol agents against plant parasitic nematodes (Guo et al. 2012) (Table 10.2).
10.2.5 Endophytic Fungi
The approach of using endophytic fungi as biocontrol agents of plant parasitic nematodes has gained attention in recent years to rectify the difficulties involved in establishing an introduced organism in the rhizosphere environment. Endophytes have the benefit that they occur in the same ecological niche as endoparasitic nematodes but are not subject to competition from microorganisms in the soil and rhizosphere (Stirling 2011; Yadav et al. 2019a, b; 2020c). The most research work has focused on the strains of Fusarium oxysporum that reduce the infection and reproduction of burrowing nematode, Radopholus similis (Athman et al. 2007), root-knot nematode, M. incognita (Hallmann and Sikora 1994; Dababat and Sikora 2007), lesion nematode, Pratylenchus goodeyi (Waweru et al. 2014), spiral nematode, Helicotylenchus multicinctus (Waweru et al. 2014).
Trichoderma spp. are endophytic and mycoparasitic fungi that have been described as biocontrol agent against plant parasitic nematodes (Zhang et al. 2014; Li et al. 2015; Dandurand and Knudsen 2016). Trichoderma parasitizes nematode eggs by the secretion of chitinolytic enzymes encoded by two genes chi 18-5 and chi 18-12 (Szabó et al. 2012). In addition, trypsin like protease (PRA1) and serine protease (SprT) were also observed against Meloidogyne juveniles during infection by the fungi (Suarez et al. 2004; Chen et al. 2009). Further, Szabó et al. 2013 reported that comparative analysis of protease expression profiles in T. harzianum revealed 13 peptidase encoding genes, suggesting that these genes might play an important role in infection process by the fungi. In addition to direct antagonism, nematophagous activity has also observed on eggs of M. incognita, M. javanica, M. arenaria, and M. exigua (Windham et al. 1989; Eapen et al. 2005; Sharon et al. 2007; Spiegel et al. 2007; Ferreira et al. 2008).
The future market for fungal products may be improved by commercializing several virulent strains for nematode pests. Zhao et al. (2013) reported that the culture filtrates of new fungal species, Simplicillium chinense (strain-Snef5) showed potential effect against soybean cyst nematode, Heterodera glycines, root-knot nematode M. incognita, white tip nematode Aphelenchoides besseyi and Caenorhabditis elegans. Whereas, Trichoderma sp. KAV1 and C. rosea KAV2 showed 100% mortality to second stage juveniles of M. incognita and M. javanica under in vitro conditions (Migunova et al. 2018). Further, the fungal isolate, Mortierella globalpina proved pathogenic to M. chitwoodi in vitro using trapping structures and subsequently reduced root galls in vivo on the roots of tomato (Solanum lycopersicum var. Rutgers) (DiLegge et al. 2019). Recently, Du et al. (2019) reported inhibition rate of 84.61%. 78.91% and 84.25% and 79.48% for adult females, juveniles, egg mass , and gall index of M. incognita under greenhouse experimental conditions at a concentration of 3 × 108 cfu mL-1 Phanerochaete chrysosporium (strain B-22) in tomato.
10.3 Nematode-Fungal Interaction
The infection process of nematophagous fungi to nematodes involves a variety of virulence factors which have been studied by various techniques such as electron microscopy, bioassays, and several biochemical, physiological, immunological, and molecular techniques (Thorn and Barron 1984; Murray and Wharton 1990; Singh and Yadav 2020). In general, the interaction of fungi with nematode species involves five different stages, viz., recognition, attraction, adhesion, penetration, and digestion.
10.3.1 Recognition
The mechanism of host recognition by fungi is not completely understood. However, few reports suggested that lectin, a carbohydrate binding protein plays an important role in nematode–fungal interaction (Borrebaeck et al. 1984; Rosenzweig and Ackroyd 1983; Nordbring-Hertz and Mattiasson 1979; Nordbring-Hertz and Chet 1986). For example, the interaction between A. oligospora and nematode was mediated by GalNAc-(N-acetyl-d-galactosamine) specific lectin which binds to carbohydrate on the nematode surface (Nordbring-Hertz and Mattiasson 1979). Whereas, Hsueh et al. (2013) reported that ascarosides, a group of glycolipids constitutively secreted by soil dwelling nematodes could trigger the trap formation in A. oligospora. This type of ascaroside induced morphogenesis is conserved in several closely related species of nematophagous fungi and occurs under nutrient stress conditions (Jiang et al. 2017).
10.3.2 Attraction
Nematodes are attracted by the culture filtrates and living mycelia of several nematophagous fungi (Li et al. 2015). The volatile compounds such as monoterpenes (α-pinene and β-pinene) and a terpenoid (camphor) produced by an endoparasitic fungus Esteya vermicola hypothesized to be involved in the interaction of pinewood nematode, Bursaphelenchus xylophilus to E. vermicola. Fungi which have more parasitic ability, i.e., endoparasitic fungi are more effective in attracting nematodes than saprophytic fungi and it was tested in soil microcosms (Tunlid et al. 1992; Dijksterhuis et al. 1994; Nordbring-Hertz et al. 2006). In the same way, the volatile compounds produced by host plant roots could also play a role in the interaction of nematode and fungi (Zhao et al. 2007).
10.3.3 Adhesion
The adhesion of nematodes to the spores and trapping structures is an essential requirement in the infection process. The presence of extracellular fibrillar layer with residues of neutral sugars, uronic acid and proteins on the surface of adhesive traps, spores and appresoria mediates the adhesion of fungi to the nematode cuticle surface (Tunlid et al. 1991; Whipps and Lumsden 2001; Su et al. 2015).
10.3.4 Penetration and Digestion
During this stage the nematophagous fungi penetrate the host by mechanical pressure and the activity of several extracellular hydrolytic enzymes that can degrade the polysaccharides and proteins of the nematode cuticle and egg shells. These extracellular enzymes include proteases, collagenases, and chitinases that have identified in different nematode trapping fungi, which act as key factors in the penetration process. After penetration, the nematode is digested by the fungus.
10.4 Extracellular Enzymes
Extracellular hydrolytic enzymes such as proteases, chitinases, and collagenases produced by nematophagous fungi play an important role in nematode cuticle penetration and host cell digestion (Åhman et al. 2002; Huang et al. 2004; Morton et al. 2004). Among these, proteases produced more rapidly in higher concentrations by nematophagous fungi than collagenases and chitinases. So far, more than 20 serine proteases have been detected, characterized, and cloned from different nematode trapping and egg parasititc fungi by several researchers (Tunlid et al. 1994; Yang et al. 2007c, 2008). Lopez-Llorca (1990) first isolated serine protease P32 from Pochonia rubens (Verticillium suchlasporium). With the availability of genomic data, the number of genes encoding serine proteases have been identified in different nematophagous fungi (Table 10.3).
Chitinases are the enzymes usually produced by nematophagous fungi to penetrate the nematode eggshell during infection (Gortari and Hours 2008). The first chitinase (Chi43) was purified from P. chlamydosporia and P. suchlasporia (Tikhonov et al. 2002). So far, 20 chitinases have been purified or cloned from various nematophagous fungi (Li et al. 2015). Table 10.4 showed a list of chitinases either purified or cloned from different nematophagous fungi and their activity on different species of nematode eggs under in vitro conditions.
Collagenases are another group of enzymes which are suspected to play a role in nematode infection. Initially Schenck et al. (1980) reported that eight nematophagous fungi could secrete collagenases. Later Tosi et al. (2002) reported that Arthrobotrys genus could produce collagenases.
10.5 Commercialization
The development of a biocontrol agent needs several steps aimed at isolation in pure culture and screening by different bioassay tests under in vitro, in vivo, and ex vivo conditions (Montesinos 2003). For commercialization of any product, the bioagent must be produced in a large scale, formulated by means of biocompatible additives to improve the storage capacity of the product. Further, quality control, registration of the particular formulated product and implementation are required (Ravensberg 2011).
In case of nematophagous fungi, various species have been tested for their efficacy in control of plant parasitic nematodes. However, only a few species have been commercialized for large scale multiplication and field application. Table 10.5 demonstrates a list of commercial products of nematophagous fungi produced by various companies under different trade names. Among these, the formulations of P. chlamydosporia, P. lilacinum, and A. robusta, A. irregularis have been widely used for nematode management in vegetables and fruit crops.
Although, nematophagous fungi provides several advantages over traditional products, the Commercialization of these bioproducts lags far behind due to inconsistent performance, quality control issues, limited shelf life of product, slow rate of kill, lack of field persistence of some formulations, difficulties in scale-up production, expensive and time consuming registration process and marketable issues, etc. (Moosavi and Askary 2015; Venkatesan and Pattar 2017). There is a need to focus on improving the formulation and manufacturing technologies that reduce costs and enhance shelf life of the commercial product.
The development and success of a biocontrol agent for plant parasitic nematodes require a better knowledge on the biology and ecology of the nematophagous fungi and the nematode, its host cultivar, method and time of application, and the various biotic and abiotic factors regulating the efficacy of biocontrol agent, the root diffusates differ markedly between plant species and cultivar which influences the proliferation of fungi (Tunlid and Ahrén 2001; Morton et al. 2004; Davies 2005). In addition, several soil microbes or their metabolites compete with the introduced bioagents for scarce energy sources. These can significantly affect the efficacy of the agent even when added to soil in a pre-colonized substrate. For example, egg masses of Meloidogyne spp. harbored 122 bacteria and 19 fungi, 23% and 74% of which, respectively, were antagonistic to P. chlamydosporium (Kok and Papert 2001). Bacillus sp. strain H6 isolated from a fungistatic soil produced iturin like compounds from that caused swelling in the conidia and germ tube of nematophagous fungi (Li et al. 2007a). Such sensitivity of a biocontrol agent to antagonism by an isolate of another microbe varies with the isolate (Montfort et al. 2006).
10.6 Conclusion and Future Prospects
Over the past 50 years, the number of scientists involved in research on the biocontrol of nematodes has increased significantly. Although several biocontrol agents for nematodes have been reported, only few organisms were developed as commercial bioagents. Surveys and empirical tests are being replaced by quantitative experimentation and basic research at genomic levels is being undertaken. Such information is essential for a realistic appraisal of the impact of microbial agents on nematode pests and for monitoring the spread and survival of the releases organisms. Our research efforts need to be directed towards identifying the factors governing their proliferation in soil and to remove the constraints wherever feasible. With the application of molecular biology, the molecular mechanisms of the interaction between nematode and fungal species can be understood, which further helps to develop new screening procedures of nematophagous fungi to control plant parasitic nematodes.
References
Abd-Elgawad MMM, Askary TH (2015) Impact of phytonematodes on agriculture economy. In: Askary TH, Martinelli PRP (eds) Biocontrol agents of phytonematodes. CAB International, Wallingford, pp 3–49
Abd-Elgawad MMM, Askary TH (2018) Fungal and bacterial nematicides in integrated nematode management strategies. Egypt J Biol Pest Co 28:74
Åhman J, Johanson T, Olsson M, Punt PJ, van den Hondel CAMJJ, Tunlid AS (2002) Improving the pathogenicity of a nematodetrapping fungus by genetic engineering of a subtilisin with nematotoxic activity. Appl Environ Microbiol 689:3408–3415
Ahren D, Ursing BM, Tunlid A (1998) Phylogeny of nematode-trapping fungi based on 18S rDNA sequences. FEMS Microbiol Lett 158:179–184
Athman SY, Dubois T, Coyne D, Gold CS, Labuschagne N, Viljoen A (2007) Effect of endophytic Fusarium oxysporum on root penetration and reproduction of Radopholus similis in tissue culture derived banana (Musa sp.) plants. Nematol 9:599–607
Atkins SD, Hidalgo-Diaz L, Kalisz H, Mauchline TH, Hirsch PR, Kerry BR (2003) Development of a new management strategy for the control of root-knot nematodes (Meloidogyne spp.) in organic vegetable production. Pest Manag Sci 59:183–189
Baron NC, Rigobelo EC, Zied DC (2019) Filamentous fungi in biological control: current status and future perspectives. Chil J Agr Res 79(2):307–315
Barron GL (1977) The nematode-destroying fungi. Lancaster Press, Lancaster, PA
Berlitz DL, Knaak N, Cassal MC, Fiuza LM (2014) Bacillus and biopesticides in control of phytonematodes. In: Sahayaraj K (ed) Basic and applied aspects of biopesticides. Springer, New Delhi, pp 3–16
Bonants PJM, Fitters PFL, Thijs H, den Belder E, Waalwijk C, Henfling JWDM (1995) A basic serine protease from Paecilomyces lilacinus with biological activity against Meloidogyne hapla eggs. Microbiol 141:775–784
Borrebaeck CA, Mattiasson B, Nordbring-Hertz B (1984) Isolation and partial characterization of a carbohydrate-binding protein from a nematode-trapping fungus. J Bacteriol 159:53–56
Bourne JM, Kerry BR (2000) Observations on the survival and competitive ability of the nematophagous fungus Verticillium chlamydosporium in soil. Int J Nematol 10:9–18
Bourne JM, Kerry BR, de Leij FAAM (1996) The importance of the host plant on the interaction between root-knot nematodes (Meloidogyne spp.) and the nematophagous fungus Verticillium chlamydosporium Goddard. Biocontr Sci Technol 6:539–548
Braga FR, Araújo JV, Soares FEF, Geniêr HLA, Queiroz JH (2012) An extracellular serine protease of an isolate of Duddingtonia flagrans nematophagous fungus. Biocontrol Sci Tech 22:1131–1142
Brand D, Soccol CR, Sabu A, Roussos S (2010) Production of fungal biological control agents through solid state fermentation: a case study on Paecilomyces lilacinus against root-knot nematodes. Micol Aplicada Int 22:31–48
Caboni P, Aissani N, Demurtas M, Ntalli N, Onnis V (2016) Nematicidal activity of acetophenones and chalcones against Meloidogyne incognita, and structure–activity considerations. Pest Manag Sci 72:125–130
Chen SY, Dickson DW (2004) Biological control of nematodes by fungal antagonists. In: Chen ZX, Chen SY, Dickson DW (eds) Nematology advances and perspectives, Vol-II Nematode management and utilization. CAB International, Wallingford, pp 979–1025
Chen TH, Hsu CS, Tsai PJ, Ho YF, Lin NS (2001) Heterotrimeric G-protein and signal transduction in the nematode-trapping fungus Arthrobotrys dactyloides. Planta 212:858–863
Chen LL, Liu LJ, Shi M, Song XY, Zheng CY, Chen XL, Zhang YZ (2009) Characterization and gene cloning of a novel serine protease with nematicidal activity from Trichoderma pseudokoningii SMF2. FEMS Microbiol Lett 299(2):135–142
Ciancio A (1995) Observations on the nematicidal properties of some mycotoxins. Fundam Appl Nematol 18(5):451–454
Dababat AEA, Sikora RA (2007) Induced resistance by the mutualistic endophyte, Fusarium oxysporum strain 162, toward Meloidogyne incognita on tomato. Biocontrol Sci Tech 17(9):969–975
Dackman C, Jansson HB, Nordbring-Hertz B (1992) Nematophagous fungi and their activities in soil. In: Stotzky G, Bollag JM (eds) Soil biochemistry. Marcel Dekker, New York, pp 95–103
Dallemole-Giaretta R, Grassi de Freitas L, Lopes EA, da Silva MCS, Kasuya MCM, Ferraz S (2015) Pochonia chlamydosporia promotes the growth of tomato and lettuce plants. Acta Scientiarum 37(4):417–423
Dandurand LM, Knudsen GR (2016) Effect of the trap crop Solanum sisymbriifolium and two biocontrol fungi on reproduction of the potato cyst nematode, Globodera pallida. Ann Appl Biol 169(2):180–189
Davies KG (2005) Interactions between nematodes and microorganisms: bridging ecological and molecular approaches. Adv Appl Microbiol 57:53–78
Davies KG, Curtis RH (2011) Cuticle surface coat of plant-parasitic nematodes. Annu Rev Phytopathol 49:135–156
de Freitas Soares FE, Sufiate BL, de Queiroz JH (2018) Nematophagous fungi: far beyond the endoparasite, predator and ovicidal groups. Agric Natu Resour 52(1):1–8
de Leij FAAM (1992) Significance of ecology in the development of Verticillium chlamydosporium as a biological control agent against root-knot nematodes (Meloidogyne spp.). Ph.D. thesis, University of Wageningen, The Netherlands
de Leij FAAM, Davies KG, Kerry BR (1992) The use of Verticillium chlamydosporium and Pasteuria penetrans alone and in combination to control Meloidogyne incognita on tomato plants. Fund Appl Nematol 15:235–242
Dijksterhuis J, Veenhuis M, Harder W, Nordbring-Hertz B (1994) Nematophagous fungi: physiological aspects and structure—function relationships. Adv Microb Physiol 36:111–143
DiLegge MJ, Manter DK, Vivanco JM (2019) A novel approach to determine generalist nematophagous microbes reveals Mortierella globalpina as a new biocontrol agent against Meloidogyne spp. nematodes. Sci Rep 9(1):7521
Du B, Xu Y, Dong H, Yan L (2019) Phanerochaete chrysosporium strain B-22, a parasitic fungus infecting Meloidogyne incognita. PLoS One 15(1):e0216688
Eapen SJ, Beena B, Ramana KV (2005) Tropical soil microflora of spice-based cropping systems as potential antagonists of root-knot nematodes. J Invertebr Pathol 88:218–225
Escudero N, Ferreira SR, Lopez-Moya F, Naranjo-Ortiz MA, MarinOrtiz AI, Thornton CR, Lopez-Llorca LV (2016) Chitosan enhances parasitism of Meloidogyne javanica eggs by the nematophagous fungus Pochonia chlamydosporia. Fungal Biol 120:572–585
Ferreira PA, Ferraz S, Lopes EA, Freitas LG (2008) Parasitismo de ovos de Meloidogyne exigua por fungos nematófagos e estudo da compatibilidade entre os isolados fúngicos. Revista Trópica: Ciências Agrárias e Biológicas 2:15–21. [in Portuguese]
Gan Z, Yang J, Tao N, Liang L, Mi Q, Li J, Zhang K-Q (2007a) Cloning of the gene Lecanicillium psalliotae chitinase Lpchi1 and identification of its potential role in the biocontrol of root-knot nematode Meloidogyne incognita. Appl Microbiol Biotechnol 76(6):1309–1317. https://doi.org/10.1007/s00253-007-1111-9
Gan Z, Yang J, Tao N, Yu Z, Zhang K-Q (2007b) Cloning and expression analysis of a chitinase gene Crchi1 from the mycoparasitic fungus Clonostachys rosea (syn. Gliocladium roseum). J Microbiol 45(5):422–430
Gortari MC, Hours RA (2008) Fungal chitinases and their biological role in the antagonism onto nematode eggs. A review. Mycol Progress 7:221–238. https://doi.org/10.1007/s11557-008-0571-3
Gray N (1985) Ecology of nematophagous fungi: effect of soil moisture, organic matter, pH and nematode density on distribution. Soil Bio Biochem 17:499–507
Guo J, Zhu C, Zhang C, Chu Y, Wang Y et al (2012) Thermolides, potent nematocidal PKS-NRPS hybrid metabolites from thermophilic fungus Talaromyces thermophilus. J Am Chem Soc 134:20306–20309
Hallmann J, Sikora RA (1994) Influence of Fusarium oxysporum, a mutualistic fungal endophyte, on Meloidogyne incognita infection of tomato. J Plant Dis Prot 101(5):475–481
Hsueh YP, Mahanti P, Schroeder FC, Sternberg PW (2013) Nematode trapping fungi eavesdrop on nematode pheromones. Curr Biol 23:83–86. https://doi.org/10.1016/j.cub.2012.11.035
Huang XW, Zhao NH, Zhang KQ (2004) Extracellular enzymes serving as virulence factors in nematophagous fungi involved in infection of the host. Res Microbiol 115:811–816
Jansson HB, Friman E (1999) Infection-related surface proteins on conidia of the nematophagous fungus Drechmeria coniospora. Mycol Res 103(2):249–256
Jiang X, Xiang M, Liu X (2017) Nematode-trapping fungi. Microbiol Spectr 5(1):28128072. https://doi.org/10.1128/microbiolspec.FUNK-0022-2016
Jones JT, Haegeman A, Danchin EG, Gaur HS, Helder J, Jones MG, Kikuchi T, Manzanilla-López R, Palomares-Rius JE, Wesemael WM, Perry RN (2013) Top 10 plant-parasitic nematodes in molecular plant pathology. Mol Plant Pathol 14(9):946–961
Kerry BR (1987) Biological control. In: Brown RH, Kerry BR (eds) Principles and practice of nematode control in crops. Academic Press, Sydney, pp 233–263
Kerry BR, Hidalgo-Diaz L (2004) Application of Pochonia chlamydosporia in the integrated control of root-knot nematodes on organically grown vegetable crops in Cuba. Multitr Inter Integ Contr 27:123–126
Kerry BR, Hirsch PR (2011) Ecology of Pochonia chlamydosporia in the rhizosphere at the population, whole organisms and molecular scale. In: Davies KG, Spiegel Y (eds) Biological control of plant-parasitic nematodes: building coherence between microbial ecology and molecular mechanisms. Progress in biological control. Springer, Dordrecht, pp 71–182
Khan A, Williams K, Molloy MP, Nevalainen H (2003) Purification and characterization of a serine protease and chitinases from Paecilomyces lilacinus and detection of chitinase activity on 2D gels. Prot Expres Purif 32:210–220
Khan A, Williams KL, Nevalainen HK (2004) Effects of Paecilomyces lilacinus protease and chitinase on the eggshell structures and hatching of Meloidogyne javanica juveniles. Biol Control 31(3):346–352
Khan AL, Hamayun M, Kang SM, Kim YH, Jung HY, Lee JH et al (2012) Endophytic fungal association via gibberellins and indole acetic acid can improve plant growth under abiotic stress: an example of Paecilomyces formosus LHL10. BMC Microbiol 12:14
Kok CH, Papert A (2001) Microflora of Meloidogyne egg masses: species composition, population density and effect on the biocontrol agent Verticillium chlamydosporium (Goddard). Nematol 3(8):729–734
Kour D, Rana KL, Yadav AN, Yadav N, Kumar M, Kumar V, Vyas P, Dhaliwal HS, Saxena AK (2020) Microbial biofertilizers: bioresources and eco-friendly technologies for agricultural and environmental sustainability. Biocatal Agric Biotechnol 23:101487. https://doi.org/10.1016/j.bcab.2019.101487
Kumar S, Singh A (2015) Biopesticides: present status and the future prospects. J Fertil Pestic 6:e129
Kumar KK, Sridhar J, Murali-Baskaran RK, Senthil-Nathan S, Kaushal P, Dara SK, Arthurs S (2018) Microbial biopesticides for insect pest management in India: current status and future prospects. J Invertebr Pathol 165:74–81
Kwok OCH, Plattner R, Weisleder D et al (1992) A nematicidal toxin from Pleurotus ostreatus NRRL 3526. J Chem Ecol 18:127
Lamovsek J, Urek G, Trdan S (2013) Biological control of rootknot nematodes (Meloidogyne spp.): microbes against pests. Acta Agric Slov 101(2):263–275
Li G, Zhang K-Q (2014) Nematode-toxic fungi and their nematicidal metabolites. In: Zhang K-Q, Hyde KD (eds) Nematode-trapping fungi. Springer, Dordrecht, pp 313–375
Li Y, Hyde K, Jeewon R, Cai L, Vijaykrishna D, Zhang K (2005) Phylogenetics and evolution of nematode-trapping fungi (Orbiliales) estimated from nuclear and protein coding genes. Mycologia 97:1034–1046
Li J, Yang JK, Huang XW, 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
Li G, Zhang K, Xu J, Dong J, Liu Y (2007a) Nematicidal substances from fungi. Recent Pat Biotechnol 1:212–233
Li G, Wang X, Zheng L, Li L, Huang R, Zhang K (2007b) Nematicidal metabolites from the fungus Pleurotus ferulae Lenzi. Ann Microbiol 57:527–529
Li L, Ma M, Liu Y, Zhou J, Qu Q, Lu K, Fu D, Zhang K (2011) Induction of trap formation in nematode-trapping fungi by a bacterium. FEMS Microbiol Lett 322:157–165
Li J, Zou C, Xu J, Ji X, Niu X, Yang J, Huang X, Zhang K-Q (2015) Molecular mechanisms of nematode-nematophagous microbe interactions: basis for biological control of plant-parasitic nematodes. Annu Rev Phytopathol 53:67–95
Lima-Rivera DL, Lopez-Lima D, Desgarennes D, Velazquez-Rodriguez AS, Carrion G (2016) Phosphate solubilization by fungi with nematicidal potential. J Soil Sci Plant Nutr 16:507–524
Linford MB (1937) Stimulated activity of natural enemies of nematodes. Science 85:123–124
Liu K, Tian J, Xiang M, Liu X (2012) How carnivorous fungi use three-celled constricting rings to trap nematodes. Prot Cell 3(5):325–328
Liu K, Zhang W, Lai Y, Xiang M, Wang X, Zhang X, Liu X (2014) Drechslerella stenobrocha genome illustrates the mechanism of constricting rings and the origin of nematode predation in fungi. BMC Genom 15:114
Lopez-Llorca LV (1990) Purification and properties of extracellular proteases produced by the nematophagous fungus Verticillium suchlasporium. Can J Microbiol 36:530–537
Lopez-Llorca LV, Olivares-Bernabeu C, Salinas J, Jansson HB, Kolattukudy PE (2002) Pre-penetration events in fungal parasitism of nematodes eggs. Mycol Res 106:499–506
Lopez-Llorca LV, Maciá-Vicente JG, Jansson HB (2008) Mode of action and interactions of nematophagous fungi. In: Ciancio A, Mukerji KG (eds) Integrated management and biocontrol of vegetable and grain crops nematodes. Integrated management of plant pests and diseases, vol 2. Springer, Dordrecht, pp 51–76
Lu Z-X, Laroche A, Huang HC (2005) Isolation and characterization of chitinases from Verticillium lecanii. Can J Microbiol 51:1045–1055
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 73:3916–3923
Manzanilla-Lopez RH, Esteves I, Finetti-Sialer MM, Hirsch PR, Ward E, Devonshire J, Hidalgo-Diaz L (2013) Pochonia chlamydosporia: advances and challenges to improve its performance as a biological control agent of sedentary endo-parasitic nematodes. J Nematol 45(1):1–7
Meerupati T, Andersson K-M, Friman E, Kumar D, Tunlid A et al (2013) Genomic mechanisms accounting for the adaptation to parasitism in nematode trapping fungi. PLoS Genet 9(11):e1003909
Mi Q, Yang J, Ye F, Gan Z, Wu C, Niu X, Zou C, Zhang KQ (2010) Cloning and overexpression of Pochonia chlamydosporia chitinase gene pcchi44, a potential virulence factor in infection against nematodes. Process Biochem 45(5):810–814
Migunova V, Sasanelli N, Kurakov A (2018) Effect of microscopic fungi on larval mortality of the root-knot nematodes Meloidogyne incognita and Meloidogyne javanica. Biol Integr Contr Plant Pathog 133:27–31
Montesinos E (2003) Development, registration and commercialization of microbial pesticides for plant protection. Int Microbiol 6(4):245–252
Montfort WS, Kirkpatrick TL, Long DL, Rideout S (2006) Efficacy of a novel nematicidal seed treatment against Meloidogyne incognita on cotton. J Nematol 38:245–249
Moosavi MR, Askary TH (2015) Nematophagous Fungi: commercialization. In: Askary TH, Martinelli PRP (eds) Biocontrol agents of phytonematodes. CAB International, Wallingford, pp 187–202
Moosavi MR, Zare R (2012) Fungi as biological control agents of plant-parasitic nematodes. In: Mérillon J, Ramawat K (eds) Plant defence: biological control. Progress in biological control, vol 12. Springer, Dordrecht, pp 67–107
Morton O, Hirsch P, Kerry B (2004) Infection of plant-parasitic nematodes by nematophagous fungi–a review of the application of molecular biology to understand infection processes and to improve biological control. Nematol 6(2):161–170
Murray DS, Wharton DA (1990) Capture and penetration processes of the free-living juveniles of (Nematoda) by the nematophagous fungus, Arthrobotrys oligospora. Parasitol 101(1):93–100
Nguyen V-N, Oh I-J, Kim Y-J, Kim K-Y, Kim Y-C, Park R-D (2009) Purification and characterization of chitinases from Paecilomyces variotii DG-3 parasitizing on Meloidogyne incognita eggs. J Ind Microbiol Biotechnol 36:195–203
Niu XM (2017) Secondary metabolites from Pochonia chlamydosporia and other species of Pochonia. In: Manzanilla-López R, Lopez-Llorca L (eds) Perspectives in sustainable nematode management through Pochonia chlamydosporia applications for root and rhizosphere health. Springer, Cham, pp 131–168
Nordbring-Hertz B, Chet I (1986) Fungal lectins and agglutinins. In: Mirelman D (ed) Microbial lectins and agglutinins: properties and biological activity. Wiley, New York, pp 393–408
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. Wiley, Hoboken, NJ
Nordbring-Hertz B, Jansson H-B, Tunlid A (2011) Nematophagous fungi. Wiley, Chichester, pp 1–13
Park JO, Hargreaves JR, McConville EJ, Stirling GR, Ghisalberti EL (2004) Production of leucinostatins and nematicidal activity of Australian isolates of Paecilomyces lilacinus (Thom) Samson. Lett Appl Microbiol 38:271–276
Rastegari AA, Yadav AN, Awasthi AA, Yadav N (2020a) Trends of microbial biotechnology for sustainable agriculture and biomedicine systems: diversity and functional perspectives. Elsevier, Cambridge
Rastegari AA, Yadav AN, Awasthi AA, Yadav N (2020b) Trends of microbial biotechnology for sustainable agriculture and biomedicine systems: perspectives for human health. Elsevier, Cambridge
Ravensberg WJ (2011) A roadmap to the successful development and commercialization of microbial pest control products for control of arthropods, vol 10. Springer, New York
Rinu K, Pandey A (2011) Slow and steady phosphate solubilization by a psychrotolerant strain of Paecilomyces hepiali (MTCC 9621). World J Microbiol Biotechnol 27:1055–1062
Rosenzweig WD, Ackroyd D (1983) Binding characteristics of lectins involved in the trapping of nematodes by fungi. Appl Environ Microbiol 46:1093–1096
Rosso LC, Finetti-Sialer MM, Hirsch PR, Ciancio A, Kerry BR, Clark IM (2011) Transcriptome analysis shows differential gene expression in the saprotrophic to parasitic transition of Pochonia chlamydosporia. Appl Microbiol Biotechnol 90(6):1981–1994
Schenck S, Chase T, Rosenzweig W, Pramer D (1980) Collagenase production by nematode-trapping fungi. Appl Environ Microbiol 40:567–570
Scholler M, Hagedorn G, Rubner A (1999) A reevaluation of predatory orbiliaceous fungi II A new generic concept. Sydowia 51:89–113
Segers R (1996) The nematophagous fungus Verticillium chlamydosporium: aspects of pathogenicity. PhD thesis, University of Nottingham, UK
Segers R, Butt TM, Kerry BR, Peberdy JF (1994) The nematophagous fungus Verticillium chlamydosporium produces a chymoelastaselike protease which hydrolyses host nematode proteins in situ. Microbiol 140:2715–2723
Sharon E, Chet I, Viterbo A, Bar-Eyal M, Nagan H, Samuels GJ, Spiegel Y (2007) Parasitism of Trichoderma on Meloidogyne javanica and role of the gelatinous matrix. Eur J Plant Pathol 118:247–258
Singh J, Yadav AN (2020) Natural bioactive products in sustainable agriculture. Springer, Singapore
Soares FE, Braga FR, Araújo JV, dos Santos LW, Mozer LR, Queiróz JH (2012) In vitro activity of a serine protease from Monacrosporium thaumasium fungus against first-stage larvae of Angiostrongylus vasorum. Parasitol Res 110:2423–2437
Spiegel Y, Sharon E, Bar-Eyal M, Maghodia A, Vanachter A, Van Assche A et al (2007) Evaluation and mode of action of Trichoderma isolates as biocontrol agents against plant-parasitic nematodes. IOBC WPRS Bulletin 30(6/2):129
Stadler M, Mayer A, Anke H, Sterner O (1994a) Fatty acids and other compounds with nematicidal activity from cultures of basidiomycetes. Planta Med 60:128e132
Stadler M, Sheldrick WS, Dasenbrock J, Steglich W, Anke H (1994b) Antibiotics from the nematode-trapping basidiomycete Nematoctonus robustus. Nat Prod Lett 4:209e216
Steyn PS (1970) The isolation, structure and absolute configuration of secalonic acid D, the toxic metabolite of Penicillium oxalicum. Tetrahedron Lett 26(1):51–57
Stirling GR (1991) Biological control of plant parasitic nematodes. CAB International, Wallingford, p 282
Stirling GR (2011) Biological control of plant parasitic nematodes: an ecological perspective, a review of progress and opportunities for further research. In: Davies KG, Spiegel Y (eds) Biological control of plant parasitic nematodes: building coherence between microbial ecology and molecular mechanisms. Springer, Dordrecht, pp 1–38
Stirling GR, Smith L (1998) Field tests of formulated products containing either Verticillium chlamydosporium or Arthrobotrys dactyloides for biological control of root-knot nematodes. Biol Contr 11:231–239
Su H, Zhao Y, Zhou J, Feng H, Jiang D, Zhang KQ, Yang J (2015) Trapping devices of nematode-trapping fungi: formation, evolution, and genomic perspectives. Biol Rev Camb Philos Soc 31:371–378
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(1):46–55
Szabó M, Csepregi K, Gálber M, Virányi F, Fekete C (2012) Control plant-parasitic nematodes with Trichoderma species and nematode-trapping fungi: the role of chi18-5 and chi18-12 genes in nematode egg-parasitism. Biol Control 63(2):121–128
Szabó M, Urbán P, Virányi F, Kredics L, Fekete C (2013) Comparative gene expression profiles of Trichoderma harzianum proteases during in vitro nematode egg-parasitism. Biol Control 67(3):337–343
Thorn RG, Barron GL (1984) Carnivorous mushrooms. Science 224(4644):76–78
Tikhonov VE, Lopez-Llorca LV, Salinas J, Jansson H-B (2002) Purification and characterization of chitinases from the nematophagous fungi Verticillium chlamydosporium and V.suchlasporium. Fungal Genet Biol 35:67–78
Tosi S, Annovazzi L, Tosi I, Iadarola P, Caretta G (2002) Collagenase production in an Antarctic strain of Arthrobotrys tortor Jarowaja. Mycopathologia 153:157–162
Tunlid A, Ahrén D (2001) Application of genomics to the improvement of nematode-trapping. In: Vurro M, Gressel J, Butt T et al (eds) Enhancing biocontrol agents and handling risks. IOS Press, Amsterdam, pp 193–200
Tunlid A, Johansson T, Nordbring-Hertz B (1991) Surface polymers of the nematode-trapping fungus Arthrobotrys oligospora. J Gen Microbiol 137:1231–1240
Tunlid A, Jansson H-B, Nordbring-Hertz B (1992) Fungal attachment to nematodes. Mycol Res 96:401–412
Tunlid A, Rosen S, Ek B, Rask L (1994) Purification and characterization of an extracellular serine protease from the nematode-trapping fungus Arthrobotrys oligospora. Microbiology 140:1687–1695
Vasanthi D, Kumaraswamy K (1999) Efficacy of vermicompost to improve soil fertility and rice yield. J Ind Soc Soil Sci 47(2):268–272
Venkatesan T, Pattar S (2017) Intellectual property guidelines and commercialization of biocontrol agents/biopesticides technologies. In: Varshney R, Gupta A, Patil J (eds) Training manual on “identification, mass production and utilization of parasitoids, predators and entomopathogens for sustainable insect pest management”. ICAR-NBAIR, Bengaluru, pp 115–126
Wang ELH, Bergeson GB (1974) Biochemical changes in root exudates and xylem sap of tomato plants infected with Meloidogyne incognita. J Nematol 6:194–202
Wang M, Yang JK, 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 KQ (2006b) Purification and characterization of an extracellular serine protease from the nematodetrapping fungus Dactylella shizishanna. Lett Appl Microbiol 42:589–594
Wang B, Wu WP, Liu XZ (2007) Purification and characterization of a neutral serine protease with nematicidal activity from Hirsutella rhossiliensis. Mycopathologia 163:169–176
Wang B, Liu X, Wu WP, Liu X, Li S (2009) Purification, characterization, and gene cloning of an alkaline serine protease from a highly virulent strain of the nematode-endoparasitic fungus Hirsutella rhossiliensis. Microbiol Res 164:665–673
Wang X, Guan T, Zhang L, Li H (2015) Cloning of a serine protease gene from the nematophagous fungus Esteya vermicola and expressed activity of the recombinant enzyme against Bursaphelenchus xylophilus. Nematology 17(9):1071–1080
Waweru B, Turoop L, Kahangi E, Coyne D, Dubois T (2014) Non-pathogenic Fusarium oxysporum endophytes provide field control of nematodes, improving yield of banana Musa sp. Biol Control 74:82–88
Whipps JM, Lumsden RD (2001) Comercial use of fungi as plant disease biological control agents: status and prospects. In: Butt TM, Jackson CW, Magan N (eds) Fungi as biocontrol agents progress, problems and potential. CA National, New York, pp 9–22
Windham GL, Windham MT, Williams WP (1989) Effects of Trichoderma spp. on maize growth and Meloidogyne arenaria reproduction. Plant Dis 73:493–495
Yadav AN, Singh S, Mishra S, Gupta A (2019a) Recent advancement in white biotechnology through fungi. Volume 2: perspective for value-added products and environments. Springer, Cham
Yadav AN, Singh S, Mishra S, Gupta A (2019b) Recent advancement in white biotechnology through fungi. Volume 3: perspective for sustainable environments. Springer, Cham
Yadav AN, Rastegari AA, Yadav N, Kour D (2020a) Advances in plant microbiome and sustainable agriculture: diversity and biotechnological applications. Springer, Singapore
Yadav AN, Rastegari AA, Yadav N, Kour D (2020b) Advances in plant microbiome and sustainable agriculture: functional annotation and future challenges. Springer, Singapore
Yadav AN, Singh J, Rastegari AA, Yadav N (2020c) Plant microbiomes for sustainable agriculture. Springer, Cham
Yang JK, Huang XW, Tian BY, Wang M, Niu QH, Zhang KQ (2005) Isolation and characterization of a serine protease from the nematophagous fungus, Lecanicillium psalliotae, displaying nematicidal activity. Biotechnol Lett 27:1123–1128
Yang Y, Yang E, An Z, Liu X (2007a) Evolution of nematode-trapping cells of predatory fungi of the Orbiliaceae based on evidence from rRNA-encoding DNA and multiprotein sequences. Proc Natl Acad Sci 104:8379–8384
Yang JK, Li J, Liang LM, Tian BY, Zhang Y, Chen CM, Zhang KQ (2007b) Cloning and characterization of an extracellular serine protease from the nematode-trapping fungus Arthrobotrys conoides. Arch Microbiol 188:167–174
Yang JK, Liang LM, Zhang Y, Li J, Zhang L, Ye FP, Gan ZW, Zhang KQ (2007c) Purification and cloning of a novel serine protease from the nematode-trapping fungus Dactylellina varietas and its potential roles in infection against nematodes. Appl Microbiol Biotechnol 75:557–565
Yang JK, Ye FP, Mi QL, Tang SQ, Li J, Zhang KQ (2008) Purification and cloning of an extracellular serine protease from the nematodetrapping fungus Monacrosporium cystosporium. J Microbiol Biotechnol 18:852–858
Yang J, Wang L, Ji X, Feng Y, Li X, Zou C, Xu J, Ren Y, Mi Q, Wu J et al (2011) Genomic and proteomic analyses of the fungus Arthrobotrys oligospora provide insights into nematode-trap formation. PLoS Pathog 7:e1002179
Yang E, Xu L, Yang Y, Zhang X, Xiang M, Wang C, An Z, Liu X (2012) Origin and evolution of carnivorism in the Ascomycota (fungi). Proc Natl Acad Sci USA 109:10960–10965
Zaki FA (1994) Mode of action of nematode destroying organisms. In: Bhatti DS, Walia RK (eds) Nematode pest management in crops. CBS Publishers, Delhi, pp 239–256
Zhang K-Q, Hyde KD (2014) Nematode-trapping fungi. Springer, Dordrecht
Zhang YJ, Liu XZ, Wang M (2008) Cloning, expression, and characterization of two novel cuticle-degrading serine proteases from the entomopathogenic fungus Cordyceps sinensis. Res Microbiol 159:462–469
Zhang Y, Yu Z-F, Xu J, Zhang K-Q (2011) Divergence and dispersal of the nematode-trapping fungus Arthrobotrys oligospora from China. Environ Microbiol Rep 3:763–773
Zhang S, Gan Y, Xu B, Xue Y (2014) The parasitic and lethal effects of Trichoderma longibrachiatum against Heterodera avenae. Biol Control 72:1–8
Zhao ML, Mo MH, Zhang KQ (2004) Characterization of a neutral serine protease and its full-length cDNA from the nematodetrapping fungus Arthrobotrys oligospora. Mycologia 96:16–22
Zhao ML, Huang JS, Mo MH, Zhang KQ (2005) A potential virulence factor involved in fungal pathogenicity: serine-like protease activity of nematophagous fungus Clonostachys rosea. Fungal Divers 19:217–234
Zhao LL, Wei W, Kang L, Sun JH (2007) Chemotaxis of the pinewood nematode, Bursaphelenchus xylophilus, to volatiles associated with host pine, Pinus masso-niana, and its vector Monochamus alter-natus. J Chem Ecol 33:1207–1216
Zhao D, Liu B, Li LY, Zhu XF, Wang YY, Wang JQ, Duan YX, Chen LJ (2013) Simplicillium chinense: a biological control agent against plant parasitic nematodes. Biocontrol Sci Tech 23:980–986
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Kumar, K.K. (2020). Fungi: A Bio-resource for the Control of Plant Parasitic Nematodes. In: Yadav, A., Mishra, S., Kour, D., Yadav, N., Kumar, A. (eds) Agriculturally Important Fungi for Sustainable Agriculture. Fungal Biology. Springer, Cham. https://doi.org/10.1007/978-3-030-48474-3_10
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