InTechOpen uses cookies to offer you the best online experience. By continuing to use our site, you agree to our Privacy Policy.

Biochemistry, Genetics and Molecular Biology » "Nematology - Concepts, Diagnosis and Control", book edited by Mohammad Manjur Shah and Mohammad Mahamood, ISBN 978-953-51-3416-9, Print ISBN 978-953-51-3415-2, Published: August 16, 2017 under CC BY 3.0 license. © The Author(s).

Chapter 8

Harnessing Useful Rhizosphere Microorganisms for Nematode Control

By Seloame Tatu Nyaku, Antoine Affokpon, Agyemang Danquah and Francis Collison Brentu
DOI: 10.5772/intechopen.69164

Article top


Model for the modulation of host immunity in ectomycorrhizal (EMF) and arbuscular mycorrhizal (AMF) fungi. (A) Root exudates recruit symbiotic mycorrhizal fungi and prime them for the interaction. Host plants initially recognize ectomycorrhizal (EMF) and arbuscular mycorrhizal (AMF) fungi as potential invaders; pattern recognition receptors (PRR) in the host perceive microbe‐associated molecular patterns (MAMPs) and a signaling cascade is initiated that results in MAMP‐triggered immunity (MTI). (B) The establishment of the symbiotic program in plant cells, which is activated upon perception of the mycorrhizal Myc factors, counteracts MTI with mechanisms yet to be defined. Molecules secreted in the apoplastic or peri‐arbuscular space (PAS) may act as either apoplastic or cytoplasmic effectors to suppress the MTI response or promote the symbiotic program. The AMF Glomus intraradices secretes the SP7 effector which is translocated into the plant cytosol; a nuclear localization signal (NLS) targets SP7 to the nucleus, where it interacts with the defense‐related transcription factor ERF19 to block the ERF19‐mediated transcriptional program [85].
Figure 1. Model for the modulation of host immunity in ectomycorrhizal (EMF) and arbuscular mycorrhizal (AMF) fungi. (A) Root exudates recruit symbiotic mycorrhizal fungi and prime them for the interaction. Host plants initially recognize ectomycorrhizal (EMF) and arbuscular mycorrhizal (AMF) fungi as potential invaders; pattern recognition receptors (PRR) in the host perceive microbe‐associated molecular patterns (MAMPs) and a signaling cascade is initiated that results in MAMP‐triggered immunity (MTI). (B) The establishment of the symbiotic program in plant cells, which is activated upon perception of the mycorrhizal Myc factors, counteracts MTI with mechanisms yet to be defined. Molecules secreted in the apoplastic or peri‐arbuscular space (PAS) may act as either apoplastic or cytoplasmic effectors to suppress the MTI response or promote the symbiotic program. The AMF Glomus intraradices secretes the SP7 effector which is translocated into the plant cytosol; a nuclear localization signal (NLS) targets SP7 to the nucleus, where it interacts with the defense‐related transcription factor ERF19 to block the ERF19‐mediated transcriptional program [85].
Signal exchange during symbiosis. (A) An asymbiotic cell constitutively releases root exudates, including strigolactones. The root cell monitors the concentration of minerals and microbial organisms in the soil and transduces the respective signals. Integration of the signals occurs at the cellular and organismic levels and includes cell‐to‐cell communication. (B) A root‐hair cell primed for interaction with rhizobia or AM fungi, respectively. Plant roots release flavonoids and strigolactones that prime the rhizobia and AM fungi. Nod and Myc factors act as signals from the symbionts to plant root cells that activate calcium spiking via the Sym pathway (boxed). The potential differential activation of CaMK/Cyclops leads to differential induction of nodulation‐specific transcription factors (NSP1, NSP2, and ERN) and unknown mycorrhizal‐specific transcription factors. Rhizobial and mycorrhizal infection require the common Sym pathway but also exhibit recognition and signaling independent of this pathway. The path for fungal infection and the IT is predicted by the PiT and the PPA, respectively, indicating directed signaling to neighboring cells. Nodule organogenesis is induced in inner cortical cells after nod‐factor perception by epidermal cells. This requires cytokinin signaling and is associated with changes in auxin levels [89].
Figure 2. Signal exchange during symbiosis. (A) An asymbiotic cell constitutively releases root exudates, including strigolactones. The root cell monitors the concentration of minerals and microbial organisms in the soil and transduces the respective signals. Integration of the signals occurs at the cellular and organismic levels and includes cell‐to‐cell communication. (B) A root‐hair cell primed for interaction with rhizobia or AM fungi, respectively. Plant roots release flavonoids and strigolactones that prime the rhizobia and AM fungi. Nod and Myc factors act as signals from the symbionts to plant root cells that activate calcium spiking via the Sym pathway (boxed). The potential differential activation of CaMK/Cyclops leads to differential induction of nodulation‐specific transcription factors (NSP1, NSP2, and ERN) and unknown mycorrhizal‐specific transcription factors. Rhizobial and mycorrhizal infection require the common Sym pathway but also exhibit recognition and signaling independent of this pathway. The path for fungal infection and the IT is predicted by the PiT and the PPA, respectively, indicating directed signaling to neighboring cells. Nodule organogenesis is induced in inner cortical cells after nod‐factor perception by epidermal cells. This requires cytokinin signaling and is associated with changes in auxin levels [89].

Harnessing Useful Rhizosphere Microorganisms for Nematode Control

Seloame Tatu Nyaku1, 2, Antoine Affokpon3, Agyemang Danquah1, 2 and Francis Collison Brentu4
Show details


Nematodes are very diverse and parasitize various plants including vegetables, and their management is of concern. Biological control of nematodes provides an environmentally friendly management option and there are various micro‐soil‐borne organisms which can be considered for this purpose. The primary goal of this chapter is to provide a review on the progress made so far, in application of biological control agents in nematode management in vegetables, cereals, and root and tuber crops. This chapter will be divided into five (5) sections: (1) herbivore‐induced plant volatiles, (2) root exudates and nematode control, (3) inhibitory metabolites in bacteria for nematode management, (4) fungi and symbiotic reprogramming in host cells, and (5) fungi antagonists of nematodes.

Keywords: arbuscular mycorrhizal fungi (AMF), biocontrol, volatile organic compounds (VOCs)

1. Introduction

Plant‐parasitic nematodes (PPNs) represent serious threat to the world economy and are responsible for great losses in production systems worldwide [1]. In monetary terms, world agricultural economy losses are approximately $215.8 billion annually, because of 12.6% crop loss inflicted on top 20 life‐sustaining crops by PPN based on 2010–2013 production figures and prices. These figures do not cover all crops throughout the world especially crops produced in the developing countries which will probably exceed these estimates if combined. Therefore, nematode management is a major constraint in food security efforts worldwide. However, PPNs are difficult to control compared to other pests because nematodes mostly inhabit the soil, and usually attack the underground parts of the plants [2]. Although chemical nematicides are effective, easy to apply, and show rapid effects, the growing dissatisfaction with chemical nematicides due to environmental and health issues has created redirections in the type and choice of applicable nematicides [3]. In view of these challenges posed by traditional nematicides, for the past 20 years the search for novel, environmentally friendly alternatives with which to manage PPN populations has therefore become increasingly important. The role of different beneficial microorganisms in the soil ranks high as environmentally friendly biological alternatives to synthetic nematicides [3].

Volatile compounds are emitted both by eukaryotes and by prokaryotes; these volatile organic compounds (VOCs) are lipophilic, with a molecular mass of about 300 Da or less, and a vapor pressure of 0.01 kPa. These chemicals evaporate easily and are produced through diffusion; however, other mechanisms (passive or active) for their emission and transmission exist [4]. Three chemical groups can be associated with the volatile compounds (terpenoids, phenylpropanoids, and fatty acid derivatives). Volatile compound penetration and movement in soils is greatly influenced by the mineral type, soil texture, and particle design [5]. The rhizosphere has within it various microorganisms because of its conducive environment; furthermore, about 20% of carbon can be released by roots [6]. Root exudates are made up of various chemical compounds, among these are amino acids and amides, organic acids, sugars, phenols, polysaccharides, secondary metabolites, and proteins [7]. Volatile metabolites effused in the soil could have an impact on the organism within the soil community. Mycorrhizal and non‐mycorrhizal plants also release distinct root exudates which contain organic acids and sugars [8].

Plant‐parasitic nematodes move toward their host and this phenomenon is important in agriculture [9]. Carbon dioxide is a root volatile with specific roles in luring plant‐parasitic nematodes, for example, to their hosts Meloidogyne incognita [10], Caenorhabditis elegans [11], and Ditylenchus dipsaci [12]. In a previous study, a tracking system linked to a computer was implored to monitor the responses of second‐stage juveniles of M. incognita exposed to carbon dioxide [10]. Results revealed a positive correlation among carbon dioxide concentration increase and nematode locomotion rate. Higher carbon dioxide concentrations (>10%) resulted in a reduction of nematode movement. In a second experiment, the movement of nematodes was monitored on a gradient, maintaining the carbon dioxide concentration constant. Thresholds were maintained either above or below 0.01% CO2/cm. The migration rate under optimal CO2concentrations was 0.7 cm/h. Plants secrete chemicals, for example, benzaldehyde, thymol, limonene, neral, geranial, and carvacrol which are needed for defense against other pathogens in the soil [1318]. These chemicals may have within them nematicidal properties.

2. Herbivore‐induced plant volatiles

Herbivore‐induced plant volatiles (HIPVs) are generated after a herbivore feeds on its host roots and their roles to attract nematodes and other predators are still been explored [1921]. Lima bean (Phaseolus lunatus) releases volatiles after the feeding activities of spider mites (Tetranychus urticae); this volatile attracts Phytoseiulus persimilis which is a predatory mite [22]. Among the compounds present in the oral secretions of herbivores are volicitin and fatty acid amides, which stimulate volatile release in plants [23, 24].

The roles herbivores play in relation to nematode parasitism on plants have been investigated [25, 26]. Signals released from plant roots, which are also parasitized by insects, influence the actions of entomopathogenic nematodes (EPNs) [27, 28]. Feeding mechanisms of herbivores stimulate the release of EPN‐attracting volatiles, especially in annual grasses [29]. A hybrid root stock “Swingle citrumelo” lures EPNs (Steinernema diaprepesi) toward its roots after parasitism by larval Diaprepes abbreviatus root weevils; this is because of the production of subterranean volatiles (terpenoid) [30]. The citrus nematode Tylenchulus semipenetrans is a devastating pest of citrus causing damage to about 8–12% of citrus species; however, higher infection rates (53–89%) have been observed on citrus in Florida [31]. This nematode life cycle has the second‐stage juvenile (J2) as the most infective stage. These nematodes are attracted to citrus roots that have been parasitized by weevil larvae (D. abbreviatus) compared to non‐parasitized plants [26]. In their experiment, the response of four entomopathogenic nematodes (S. diaprepesi, S. carpocapsae, S. riobrave, and Heterorhabditis indica) and a plant‐parasitic nematode (T. semipenetrans) to D. abbreviatus parasitism on citrus root stocks (Poncirus trifoliata, S. citrumelo‐(C. paradisi × P. trifoliata), and Citrus aurantium) was investigated. Results revealed high nematode numbers that moved toward S.citrumelo weevil‐infested roots, compared to the non‐infested ones in spite of the foraging strategy implored by the nematode‐foraging strategy and its trophic status. Further, parasitism or non‐parasitism of D. abbreviatus on the citrus parent line P. trifoliata did not influence the attraction level of nematodes, because the nematode responses to the root stock were similar. Production of the volatile, pregeijerene was released after feeding activity by D. abbreviatus only within the root zone and absent in the upper portions of shoots. Feeding activity by the adult beetle (D. abbreviatus) on the shoots did not stimulate the production of pregeijerene; however, limonene was released. Within the P. trifoliata roots, pregeijerene was released; however, the feeding activity of D. abbreviatus had no influence in its production.

Maize root volatiles can be associated with the ability of entomopathogenic nematodes in controlling the western corn rootworm. The roots of maize release the volatile (E)‐β‐caryophyllene (EβC) after parasitism by the larvae of Diabrotica virgifera virgifera. This chemical, which is a sesquiterpene, serves as an attractant to some species of entomopathogenic nematodes [29, 32, 33]. The volatile (E)‐β‐caryophyllene (EβC) was investigated on the EPN H. bacteriophora, H. megidis, and S. feltiae against D. v. virgifera larvae in southern Hungary. The maize variety that released (E)‐β‐caryophyllene (EβC) was protected from H. megidis and S. feltiae.

The roots of cotton (Gossypium herbaceum) also emit terpenoid volatiles after the feeding activity of the larvae of the chrysomelid beetle D. balteata [25]. This sesquiterpenoid aristolene may be a useful volatile for attraction of the nematode H. megidis.

3. Root exudates and nematode control

Plant root exudates and their impact on root‐knot nematode egg hatchability are an important development for nematode management. The chemicals within root exudates may either attract or repel nematodes to their host roots. There is experimental evidence to show the influence of root exudates on nematode egg hatch [3436]. There are specific signals which are generated from exudates of roots; these enable nematodes to be attracted to their hosts. Known compounds that attract second‐stage juveniles to host roots include tannic acids, flavonoids, glycoside, fatty acids, and volatile organic molecules [37, 38]. Semiochemicals, for example, small lipophilic molecules produced from root exudates of tomato and rice, enable stylet movement into host cells [39].

Root exudates have within them organic acids and sugars which are generated from mycorrhizal and non‐mycorrhizal plants [8]. Flavonoids [40], phenolic compounds [41], amino acids [42], and the plant hormone strigolactone [43] are also constituents of root exudates. Root exudates released by mycorrhizal plants have the potential of attracting Pseudomonas fluorescens [44] and the fungus Trichoderma spp. [45], both organisms poses nematicidal properties for biocontrol of nematodes [46, 47]. Tomato plants, which formed symbiosis with Funneliformis mosseae, had low juvenile numbers of M. incognita compared to control plots [48].

In a recent study, the impact of tomato root exudates on M. incognita was investigated. These exudates were obtained from the root stocks Baliya (highly resistant, HR), RS2 (moderately resistant, MR), and L‐402 (highly susceptible, T). These had varying impacts on M. incognita egg hatch and the movements of the second‐stage juveniles (J2) [49]. The various root exudates obtained from the tomato root stocks (HR, MR, and T strains) decreased M. incognita egg hatchability; furthermore, populations of J2 decreased with the highest mortality rate associated with exudates from the HR plants. There was a much higher repelling rate from the HR genotypes to M. incognita J2 compared to the other genotypes. However, exudates from the susceptible genotype (T) attracted the juveniles. The root exudates are made up of varying constituents from the different AMF species [50]. Microbial diversity occurring within soils is positively influenced by root exudates [51], and AMF in soils may also produce high facultative anaerobic bacteria, for example, Streptomyces species, and actinomycetes [5254].

4. Soil bacteria and nematode control

Nematodes in soil are subject to infections by bacteria and fungi. This creates the possibility of using soil bacteria to control PPN [5557]. An effective natural enemy of nematodes is nematophagous bacteria which are ubiquitous with wide host ranges. These organisms have been isolated from soil, plant tissues, cysts, and eggs of nematodes. They directly suppress the activities of nematodes through the production of antibiotics, toxins, as well as enzymes; they also compete for nutrients and space through parasitizing, and therefore provide systemic resistance for plant growth. Their activities promote plant growth though facilitating rhizosphere colonization and enhanced microbial antagonism. Antagonism may be direct, which might result from physical contact, or indirect, which includes activities that do not involve sensing or targeting the PPN. Nematophagous bacteria may be grouped into parasitic and non‐parasitic bacteria, opportunistic parasitic bacteria, rhizobacteria, Cry protein‐forming bacteria, endophytic bacteria, and symbiotic bacteria based on their mode of parasitism [58].

Biocontrol agents, for example, Agrobacterium, Alcaligenes, Bacillus, Clostridium, Desulfovibrio, Pseudomonas, Serratia, Streptomyces, and Pasteuria penetrans have potentials for nematode control, have shown great potential for the biological control of nematodes [59, 60]. Nematophagous bacteria affect nematodes by the following modes of action: parasitizing; producing toxins, antibiotics, or enzymes; interfering with nematode‐plant‐host recognition; competing for nutrients; inducing systemic resistance of plants; and promoting plant health [58].

Among microorganisms occurring in soil, only few have been identified as biocontrol agents for phytonematodes, and some species of fungi and bacteria are the most common parasites of nematodes [57]. Some bacteria are potent antagonists of phytonematodes, and currently some have been developed into commercial bionematicides which are being used to control on the field mainly in advanced countries [61] (Table 1). These nematophagous bacteria can be categorized into two groups based on their mechanisms of infection: (i) bacteria that are pathogenic to nematodes or nematode diseases producing bacteria and (ii) bacteria whose secretions or metabolic products are harmful to nematodes or the nematode toxin‐producing bacteria. The genus Pasteuria are endospore forming which are parasites of nematodes and water fleas [62, 63]. The control of most economically important genera of phytonematodes using nematophagous bacteria has been associated with this genus—Pasteuria. The other group includes strains of Agrobacterium radiobacter, Azotobacter chroococcum, Bacillus spp., Clostridium spp., and Streptomyces spp.

Product nameMicrobial originCompany or institutionCountryNematode targetReferences
EconemPasteuria usgae (or P. penetrans)Bayer Crop ScienceMultinationalSting (or root knot)[76]
Avid 0.15EC (orAbamectinBacillus thuringensisSyngenta Group companyMultinationalRoot-knot and other nematodes[190]
Bionem-WP, BioSafe-WP, and Chancellor-WPB. armusAgro GreenMultinationalRoot-knot and other nematodes including[190]
Nortica VOTIVO PONCHO/VOTIVOB. armusBayer CropScienceMultinationalHeterodera avenae[76]
Deny Blue circleBurkholderia capaciaStine Microbial Wisconsin ProductsUSAMeloidogyne incognita[191]
Biostart®Bacillus subtilisBio-CatUSARoot knot nematodes[192]
BiostartL™B. laterosporus, B. ncheniformis (mixture)Rhcon-Vltova
NemixBacillus subtilis, B. ncheniformisAgriLife/Chr. HansenBrazil[192]
NemalessSerrata marcescensAgriculltural Research CentreGiza, EgyptRoot-knot and other phytonematode[193]
SHEATHGUARD (or Sudozone)Pseudomonas, P. fluorescensAgri Life (Ind Limited or Agri Land Biotech)Hyderabad,IndiaNematode such as root-knot,cyst and Citrus nematode
Xlan MileBacillus cereusXlnYlZhong kai Agro-Chemical Industry Co., ltdChinaMeloidogyne spp. on vegetables[194]
Pathway Consortia®Bacillus spp. Trichoderma spp., P. flurescens, Streptomyces spp.Pathway HoldingsUSAPhytonematodes[1]
MicronemaBacillus sp., Pseudomonas sp., Rhizobacterium sp., Rhizobium sp.Agricultural Research CentreGiza, EgyptRoot-knot and other phytonematodes[195]

Table 1.

Commercial products of bacteria for phytonematode control.

CAB International 2015. Biocontrol Agents of Phytonematodes (eds T.H. Askary and P.R.P. Martinelli)

Actinobacteria are a group of soil bacteria of importance as biocontrol agents with nematicidal properties [6467]. The diversity and biocontrol ability of nematicidal actinobacteria have been investigated [67]. In their study, 200 soil samples were obtained from 20 provinces within China. Results revealed 4000 actinobacteria, and these isolates 533 (13.3%) and 488 (12.2%) have some nematicidal activities on the nematodes Panagrellus redivivus and Bursaphelenchus xylophilus, respectively. Actinobacteria are generally Gram positive bacteria, and have G+C content of >55%. There has been over 70% of bioactive compounds released by these microorganisms with their usage in agriculture and pharmaceutical industry. These organisms release lytic enzymes, and secondary metabolites. One group of metabolites are avermectins which are produced by S. avermitilis [68]. Avermectins are useful for nematode control [69]. A previous screen of 502 actinobacteria showed 15 of these with nematicidal impact on P. redivivus, a free‐living nematode [65].

Streptomyces isolate (CR‐43) from Costa Rica had inhibitory impacts on C. elegans after a laboratory experimentation [69]. Other studies conducted in the greenhouse showed CR‐43 with the potential of reducing root galls on tomato inoculated with M. incognita. Furthermore, field studies in Puerto Rico revealed pepper and tomato plants that received CR‐43 as treatments having the least gall numbers compared to controls. In an in vitro investigation, Streptomyces sp. (CMU‐MH021), which is an actinomycete isolated from nematode‐infested soils in Thailand, showed the release of secondary metabolites which prevented M. incognita egg hatch, and also a decrease in juvenile numbers [70]. The nematicidal properties of various culture filtrates were explored. The modified basal (MB) medium gave the highest activity against M. incognita. The broth microdilution technique was applied for understanding the nematicidal activity of fervenulin. Inhibitory concentrations for both egg hatch (30 μg/ml) and M. incognita juvenile mortality (120 μg/ml) were noted. An evaluation of both in vitro and in vivo nematicidal potential of extracts from S. hydrogenans strain DH16 against M. incognita prevented egg hatch (>95%) and a high mortality rate (95%) of juveniles after 96 h [71].

Furthermore, two compounds [10‐(2,2‐dimethyl‐cyclohexyl)‐6,9‐dihydroxy‐4,9‐dimethyl‐dec‐2‐enoic acid methyl ester] purified from the streptomycete were evaluated for their efficacy against M. incognita. The juvenile nematode mortality varied with the concentration rates with high mortality observed at high concentrations, for example, a concentration of 100 μg/ml caused 95% mortality after 96 h.

The marine bacteria B. firmus strain YBf‐10 shows its efficacy as a biocontrol agent on M. incognita (eggs and juveniles) through a systemic action [72]. The application of this strain through drenching of tomato plants inoculated with M. incognita produced plants with reduced galls and egg masses, and nematode numbers in soil samples.

Pasteuria, which is an endospore‐forming bacteria with various species within this genus, may be implored as biocontrol agents and there are four nematode antagonists within this genus. Among these, P. penetrans, P. thornei, P. nishizawae, and P. usgae are parasites on root‐knot nematodes, lesion nematodes [73], and Belonolaimus spp. [74]. Commercialization of Pasteuria products for nematode control is, however, limited by two factors: (i) a narrow host range [75] and (ii) growth in vitro is slow and production is tedious [76]. In vitro production of Pasteuria spp. was initiated after Pasteuria. Bioscience Alachua (Florida, USA) filed a patent in 2004, for the production of the product EconemTM, a product which is target‐specific and has been designed to control sting nematodes (Belonolaimus spp.) in turf.

5. Fungi and symbiotic reprogramming in host cells

Arbuscular mycorrhiza fungi (AMF) are in the phylum Glomeromycota [77]; these fungi form symbiotic associations with plant roots and provide phosphorus, nitrogen, and water to plants [78]. Another advantage derived from this association is tolerance to biotic and abiotic stresses by host plants [79, 80]. Native strains of AMF are used as bio‐fertilizers for enhanced plant growth, including root and tuber crops and for nematode management [81, 82]. The AMF releases signal that are transmitted systemically and these are to target non‐infected parts of roots [83, 84].Within the soil microbes with beneficial properties, for example, AMF are recognized by plants as invaders leading to the triggering of an immune response (Figure 1A) [85], and this signaling is associated with microbe‐associated molecular patterns (MAMPs), which further induce MAMP‐triggered immunity (MTI) [86, 87]. Second, there symbiotic activities within cells can be activated through mycorrhizal Myc factors if perceived (Figure 1B). The SP7 effector within the AMF Glomus intraradices is a characteristic defense signal in the fungi [88], and its expression occurs in host roots [85].


Figure 1.

Model for the modulation of host immunity in ectomycorrhizal (EMF) and arbuscular mycorrhizal (AMF) fungi. (A) Root exudates recruit symbiotic mycorrhizal fungi and prime them for the interaction. Host plants initially recognize ectomycorrhizal (EMF) and arbuscular mycorrhizal (AMF) fungi as potential invaders; pattern recognition receptors (PRR) in the host perceive microbe‐associated molecular patterns (MAMPs) and a signaling cascade is initiated that results in MAMP‐triggered immunity (MTI). (B) The establishment of the symbiotic program in plant cells, which is activated upon perception of the mycorrhizal Myc factors, counteracts MTI with mechanisms yet to be defined. Molecules secreted in the apoplastic or peri‐arbuscular space (PAS) may act as either apoplastic or cytoplasmic effectors to suppress the MTI response or promote the symbiotic program. The AMF Glomus intraradices secretes the SP7 effector which is translocated into the plant cytosol; a nuclear localization signal (NLS) targets SP7 to the nucleus, where it interacts with the defense‐related transcription factor ERF19 to block the ERF19‐mediated transcriptional program [85].

Plant cells with roots undergo reprogramming activities for successful establishment of symbiosis with symbionts (e.g., arbuscular mycorrhizal (AM) and root‐nodule (RN) symbiosis) [89] (Figure 2B). However, this reprogramming phenomenon is absent in an asymbiotic root cell (Figure 1B). Within the soil, roots of plants continuously produce and release root exudates and strigolactines as observed in an asymbiotic root cell. Signals are transmitted to the nucleus through transcription factors, gene expression occurs, and there is cell‐to‐cell communication. There are also plant receptors within the root cells that detect mineral concentration in soils. In a root cell that either interacts with AM or RN fungi, there is release of both flavonoids and strigolactones, two factors (Nod and Myc) are released from the symbionts and these turn on the calcium spiking. Within the RN symbiosis, flavonoids from the plant root turn on the Nod transcription factor, and enables bacteria to produce lipochitooligosaccharide nod factors. These Nod factors stimulate root‐nodule development, which are needed by rhizobia. Strigolactones further stimulate AM fungi and hyphal branching occurs [90]. The root cortex is usually colonized by AM fungi and produces substantial hyphae (arbuscules). During the development of the arbuscle, it becomes enveloped within the peri‐arbuscular membrane (PAM), and essential proteins are moved to the plant cell within the PAM [91]. Jasmonic acid (JA) and methyl jasmonate (MeJA) can stimulate the expression of Nod genes [92] and release of Nod factors [93], in rhizobia after their application exogenously.


Figure 2.

Signal exchange during symbiosis. (A) An asymbiotic cell constitutively releases root exudates, including strigolactones. The root cell monitors the concentration of minerals and microbial organisms in the soil and transduces the respective signals. Integration of the signals occurs at the cellular and organismic levels and includes cell‐to‐cell communication. (B) A root‐hair cell primed for interaction with rhizobia or AM fungi, respectively. Plant roots release flavonoids and strigolactones that prime the rhizobia and AM fungi. Nod and Myc factors act as signals from the symbionts to plant root cells that activate calcium spiking via the Sym pathway (boxed). The potential differential activation of CaMK/Cyclops leads to differential induction of nodulation‐specific transcription factors (NSP1, NSP2, and ERN) and unknown mycorrhizal‐specific transcription factors. Rhizobial and mycorrhizal infection require the common Sym pathway but also exhibit recognition and signaling independent of this pathway. The path for fungal infection and the IT is predicted by the PiT and the PPA, respectively, indicating directed signaling to neighboring cells. Nodule organogenesis is induced in inner cortical cells after nod‐factor perception by epidermal cells. This requires cytokinin signaling and is associated with changes in auxin levels [89].

6. Fungi antagonists of nematodes

Biological control, defined as the management of plant diseases and pests by means of other living organisms, mainly concerns the exploitation of microbial agents [94]. Under natural conditions, biocontrol agents that are associated with plant‐parasitic nematodes usually exist [95]. These organisms act through parasitism, predation, antagonism, or competition [96], but their successful activity depends on a number of parameters, including soil environmental factors [97]. Many beneficial organisms were found to attack plant‐parasitic nematodes but most research has been focused on bacteria and fungi [94, 98]. Although few biological agents had been until recently adopted for nematode control with successful use, the current progress in studies of biological control has gradually led to the development of commercial biocontrol products with proven efficacy against plant‐parasitic nematodes. Studies on fungal antagonists of nematodes have been started since 1874 with the first observations of Harposporium anguillulae, by Lohde.

7. Types of nematode‐antagonistic fungi and their mode of action

Species of several fungal genera have been reported to have biological activity against plant‐parasitic nematodes [58]. Hallmann et al. [98] classified these beneficial fungi into nematophagous fungi, saprophagous fungi, and endophytic fungi.

7.1. Nematophagous fungi

Nematophagous fungi are the largest and the most studied group of the fungi involved in the biological control against plant‐parasitic nematodes. Among nematophagous fungi, which have been tested for their efficacy in controlling nematodes, some are obligate parasites (e.g., Nematophthora gynophila), others are facultative or opportunistic parasites (e.g., Pochonia chlamydosporia) [98].

Obligate parasites require a residual population of nematodes for their survival. Infection is initiated when fungal spores penetrate the host nematode either through the gastrointestinal tract after being ingested or directly after adhering to the cuticle [98]. Among the obligate fungal parasites, Hirsutella spp. and Drechmeria coniospora have shown to be interesting in terms of their biology, mode of action, and nematode control potential. Infection of these fungi is initiated by the adhesion of small conidia to the nematode cuticle. However, obligate parasites are difficult to grow in culture.

The facultative parasites are able to switch between saprophytic state in soil and rhizosphere into parasites that infect nematodes, depending on environmental circumstances. Nematode infection occurs either by way of adhesives spores or by trapping structures, or through an appressorium [94]. Depending on their mode of action, nematophagous fungi can attack nematodes during all stages of their life cycle.

In addition to the fungi described above, some form a mycelium able to capture plant‐parasitic nematodes. They are called predacious fungi or nematode‐trapping fungi and act through different trapping structures including fungal hyphae covered with adhesive secretions (e.g., Stylopage spp.), adhesive branches (e.g., Monacrosporium cionopagum), adhesive spores (Meristacrum spp.), or adhesive knobs (Arthrobotrys spp., Nematoctonus spp.) [99, 100]. These fungi also produce nematicidal compounds such as linoleic acid (e.g., A. oligospora) or pleurotin (e.g., N. robustus) [101].

7.2. Saprophagous fungi

Among the saprophagous fungi present in the bulk soil, some have been reported to be antagonistic toward plant‐parasitic nematodes. This group was represented by the genus Trichoderma, a ubiquitous soil fungus that also colonizes the root surface and cortex [98]. Trichoderma spp. was first reported to be parasite of other fungi [102], before being identified as an antagonist of plant‐parasitic nematodes [103, 104]. A number of Trichoderma species, for example, T.asperellum, T. hamatum, harzianum, and T. viride, were reported to infect eggs and juveniles of root‐knot nematodes [105, 106]. Several possible mechanisms including the production of antifungal metabolites, competition for space and nutrients, mycoparasitism, plant growth promotion, and induction of the defense responses in plants have been suggested as mechanisms for their biocontrol activity [107, 108]. Other saprophagous fungi with antagonistic activity against plant‐parasitic nematodes include species of the genus Gliocladium, Acremonium, and Cylindrocarpon [109111].

7.3. Endophytic fungi

Endophytic fungi have been considered as important fungi in the biological control of plant‐parasitic nematodes. The implication of endophytic fungi in root‐knot nematode reduction was first demonstrated with arbuscular mycorrhizal fungi on vegetables [112].

AMFs are obligate fungi, which form symbiotic associations with numerous plant species, with the primary function of improving plant nutrient uptake [113]. Arbuscular mycorrhizal fungi are obligate plant symbionts. According to Harley and Smith [114], AMFs establish with their host plant an interdependent mutualistic relationship (symbiosis) where the host plant receives mineral nutrients, while the fungus obtains photosynthesis‐derived carbon compounds from the plant [115]. Three major types of mycorrhizal associations—ectomycorrhiza, endomycorrhiza, and ectomycorrhizal—endomycorrhizal intermediate type—have been distinguished [116]. Their endophytic nature enables associated (infected) plants to overcome biotic [117] and abiotic stresses [118]. Potential modes of actions developed by AMF during the protective activity against plant pathogens reviewed by Whipps [119] include (1) the direct competition or inhibition, (2) enhanced or altered plant growth, morphology, and nutrition, (3) biochemical changes associated with plant defense mechanisms and induced resistance, and (4) development of an antagonistic microbiota. Other studies have recently reported the ability of AMF to induce systemic resistance against plant‐parasitic nematodes in the root system [120].

Another important endophytic fungus in nematode control but with saprophytic nature is the non‐pathogenic Fusarium species, Fusarium oxysporum. Reduction of nematode penetration into the host plant root and induction of systemic resistance to plants have been considered as the main mechanisms by which F. oxyporum reduced nematode parasitism [121123].

8. Potential of antagonistic fungi in nematode control

A large number of fungi have been tested for their potential as biological control agents of plant‐parasitic nematodes. Until recently, few had been adopted for nematode control with successful use [98]. However, the current progress in studies of biological control has gradually led to the development of commercial biocontrol products with proven efficacy against plant‐parasitic nematodes. In this section, most fungal studies will be discussed.

8.1. P. chlamydosporia

Species of Pochonia are widely distributed in agricultural soils and infect eggs of plant‐parasitic nematodes, snails, and slugs [96].

Within the genus Pochonia, P. chlamydosporia appears the most effective in infecting nematode eggs [124]. P. chlamydosporia includes two subspecies P. chlamydosporia var. chlamydosporia and P. chlamydosporia var. catenulatum [125] which are considered non‐pathogenic to plants, higher animals, and humans [126]. This species is one of the major facultative antagonistic fungi that can parasitize egg and female stages of root‐knot nematodes and female cyst nematodes [96, 127, 128]. Parasitism of this fungus is based on appressorial formation developed from undifferentiated hyphae, which allows the colonization of the egg surface and penetration through both mechanical and enzymatic actions [129]. Observations during the infection process have shown that the penetration of the eggshell occurs from both the appressorium and the lateral branch of the mycelium, and leads to the disintegration and the dissolution of three layers composing the eggshell: the vitelline layer, chitin layer, and lipoprotein layer [130, 131]. The infection process is affected by the nematode host [130], suggesting that fungal growth, development, and penetration of the eggshell may be influenced by signals from the eggs [132]. Different enzymes, in particular proteases and chitinases, are important for the infection processes, and VCP1 proteases being the most known proteases with enzymatic activity against the nematode eggshells [94, 130].

The efficacy of P. chlamydosporia has been reported to be affected by three key factors: the fungal density in the rhizosphere, the rate of egg development in the egg masses, and the size of the galls in which the female nematodes develop [133]. P. chlamydosporia is found to be more abundant in the rhizosphere and on nematode‐infected roots, and parasitism may promote the long‐term survival of the fungus in soil [96]. However, the extent of colonization depends on the fungus isolate and the plant species [134, 135]. Although isolates of P. chlamydosporia differ significantly in their ability to parasitize the eggs of different nematode species, they have shown little host specificity [136].

Formulations based on P. chlamydosporia have been developed and are currently being commercialized (e.g., KlamiC®based on P. chlamydosporia var. catenulata RES 392 from Cuba) [98, 137].

8.2. Trichoderma spp

Species of Trichoderma are ubiquitous soil‐borne fungi that can colonize the root surface as well as the cortex [138, 139]. Several species of Trichoderma have been considered for biocontrol of plant‐parasitic nematodes [104]. Some species were found to be associated with eggs of root‐knot nematodes in vegetable fields [106].

Against nematodes, Trichoderma spp. can provide excellent control and are viewed as strong contenders for development as biocontrol agents [104]. In various studies, species of Trichoderma were reported to show antagonistic activity against eggs and juveniles of root‐knot nematodes in in vitro conditions [105] and to infect nematode egg masses and reduce juvenile populations in non‐sterilized field soil [140]. Trichoderma spp. were shown to efficiently control root‐knot nematodes when they were applied before planting [104, 141]. Methods suggested for their application include seed treatment, dry formulation, or soil drench [98]. However, isolates of the same species of Trichoderma can differ markedly in their rhizosphere competence, biocontrol potential toward nematodes, and plant growth promotion [141].

Different mechanisms have been suggested as mechanisms developed by Trichoderma against nematodes. The first observable interaction between Trichoderma spp. and its host is expressed by direct growth of the mycoparasite hyphae initiated by a chemotropic reaction toward the host [105]. The hyphae, upon contact, coil around and penetrate the host. This process involves the release of lytic enzymes by Trichoderma spp. [142], which serves to partially degrade the host cell wall. Lytic enzymes such as chitinases, glucanases, and proteases, seem to be particularly important in the mycoparasitic process. Induction of defense responses in plants by Trichoderma spp. was also observed through increased peroxidase and chitinase activities following fungal inoculation and a strengthening of the epidermal and cortical cell walls as the deposition of newly formed barriers [143]. These authors also reported increased enzyme activities in the leaves, suggesting a systemic defense response to the presence of Trichoderma in the rhizosphere. When monitoring fungus‐nematode interactions, Sharon et al. [105] observed that in pre‐inoculated soil, the fungus colonizing the roots interacts with the penetrating juveniles and colonizes their penetration sites, indicating also a competition for spaces. Trichoderma‐based products are commercially available and used to control plant‐parasitic nematodes on different crops. Successful examples include BioNem® [144] and T‐22™ Planter Box [145].

9. Arbuscular mycorrhizal fungi

A number of studies have demonstrated the contribution of arbuscular mycorrhizal fungi in improving soil structure [146], plant mineral uptake, and plant growth [113, 147, 148] enhancing plant tolerance to pollution with toxic metals [149, 150], resistance to drought stress [151], and reducing the effect of plant diseases [117, 152154]. AMFs have also been reported to protect host plants against plant‐parasitic nematodes [81, 98, 155]. The interaction between AMF‐colonized plants and plant‐parasitic nematodes has been reviewed by several authors [98, 156, 157]. AMFs have also been shown to suppress the effect of damage [112, 158], although some studies have shown no effects against these pests [159, 160]. However, the efficacy of AMF against nematodes may be influenced by a number of factors including prevailing environmental conditions [161], cultivar [159], nutrient status of the field [162], and the timing of application [163]. Existing knowledge suggests the application of the fungi in the nursery or to introduce suitable mycorrhizal crops into the rotation pattern for efficient pest control [98]. Pre‐inoculation of seedlings with AMF, for example, has resulted in high levels of root colonization, followed by a significant reduction of nematode infection [164]. However, recent studies showed that the level of reduction of RKN was not necessarily dependent on high‐root mycorrhization, while the interaction between crop cultivar‐AMF strains is also important [165]. Furthermore, direct inoculation of AMF inoculum into the transplanting hole prior to planting may provide plant protection against root‐knot nematodes, indicating possible use of AMF for seed‐growing crops [165]. Some studies on the combination of AMF with other antagonists have provided promising clues for their successful integration into nematode control strategies [166, 167]. Different formulations based on AMF strains (e.g., F. mosseae‐ and G. dussii‐based products from BIORIZE© in Dijon, France) were commercially developed for use in crop protection against plant‐parasitic nematodes [81, 165].

9.1. Paecilomyces lilacinus

Paecilomyces lilacinus (Thom) Samson seems to be most frequent in warmer regions, although it has been reported in different parts of the world and from various habitats [126, 168]. Investigations on the biocontrol activity of the fungus toward plant‐parasitic nematodes started after Jatala et al. [169] discovered infection of eggs and females of M. incognita and eggs of Globodera pallida. Both mechanical and enzymatic activities may be involved in the host penetration. P. lilacinus first colonizes the gelatinous matrix of Meloidgyne, Tylenchulus, and Naccobus, and cysts of cyst nematodes, develops a mycelium network, then engulfs and penetrates the nematode eggs through an appressorium or simple hyphae [126, 169]. Following penetration, the fungus grows on the early embryonic development, depletes all nutrients in the eggs, breaks the cuticle of the infected egg and infects other eggs. Although P. lilacinus is considered as egg‐pathogenic fungus, Holland et al. [170] observed in in vitro experiment infection of third‐ and fourth‐stage juveniles and adult females of M. javanica.

P. lilacinus is among the most widely studied microorganisms used for the management of plant‐parasitic nematodes. Its success in controlling plant‐parasitic nematodes has led to the development of commercial products such as MeloCon® WG by Bayer in Germany and PAECILO® by AgriLife in India [171].

9.2. Fusarium oxysporum

The interest in the non‐pathogenic Fusarium oxysporum for nematode control is stimulated after several isolates were reported to reduce the banana root rotting caused by Pratylenchus goodeyi [172]. This endophytic fungus was reported as the most abundant endophytes of banana (Musa spp.), for example, in Uganda [173, 174]. In various studies, the strain F. oxysporum FO162 has shown the ability to reduce penetration of damage caused by plant‐parasitic nematodes on tomato and banana [175178]. Dababat and Sikora [123] reported that plants colonized by F. oxyporum were less attractive or exuded substances that were repellent toward nematodes. The endophytic fungus can infect nematodes at any stages and reduce significantly the plant damage [121, 179]. Recent studies indicate that the non‐pathogenic F. oxysporum is a successful biocontrol agent for plant‐parasitic nematodes with positive effect on the plant growth [180].

9.3. Arthrobotrys spp

Arthrobotrys species are trapping fungi which immobilize nematodes [189] using different trap structures [181]. The species A. oligospora was the first recognized nematode‐trapping fungus [182]. A. conoides and A. oligospora makes three‐dimensional adhesive network to trap soil‐inhabiting nematodes [94, 183]. A. candida usually forms non‐constructing rings [184] but Al kader [181] reported a formation of adhesive hyphae capturing nematodes and then trophic hyphae within nematodes’ body to digest nematode contents. A. brochopaga forms ring traps that constrict around the body of a nematode passing through them [185]. The presence of the nematode is important in the initiation of the trapping structures [186]. Nematode species did not affect the type of trap structure but most probably the quantity of these traps. Santos et al. [187] reported substantial variability in virulence among isolates of the same species. Host recognition and adhesion by the fungus were the first steps in the infection of the host nematode. This recognition has been attributed to a molecular interaction of certain proteins on the fungal surface with sugar molecules on the nematode cuticle [183]. Substantial variability in virulence among isolates of the same species was observed [187]. Nordbring‐Hertz et al. [188] reported that Aphelenchusavenae can avoid to be captured by the fungi structures, especially for the young nematode.

10. Conclusions

Beneficial microbial inocula can be applied for large‐scale field management of nematodes which will result in increased yields. However, further research into the various biocontrol measures used by organisms is necessary, and this can be achieved through genomic approaches; this will enhance understanding of the various complex mechanisms used by these organisms on nematodes. Strains of these organisms may be effective in their local occurrences, and therefore countrywide surveys of soils will enable location‐specific strains to be isolated and characterized. These local strains once characterized can be produced in large quantities and distributed to farmers for applications in their fields.


1 - Abd‐Elgawad MM, Askary TH. Impact of phytonematodes on agriculture economy. In: Askary TH, Martinelli PRP, editors. Biocontrol Agents of Phytonematodes. Wallingford, UK: CAB International; 2015. pp. 3-49
2 - Stirling GR. Biological Control of Plant Parasitic Nematode: Progress, Problems and Prospects. Wallington, UK: CAB International; 1991. p. 282
3 - Abd‐Elgawad MM. Biological control agents of plant‐parasitic nematodes. Egyptian Journal of Biological Pest Control. 2016;26:423-429
4 - Effmert U, Buss D, Rohrbeck D, Piechulla B. Localization of the synthesis and emission of scent compounds within the Xower. In: Dudareva N, Pichersky E, editors. Floral Scents. London: CRC Press Taylor and Francis Group; 2006. pp 105-124
5 - Aochi YO, Farmer WJ. Impact of soil microstructure on the molecular transport dynamics of 1, 2‐dichlorethane. Geoderma. 2005;127:137-153
6 - Barber DA, Martin JK. The release of organic substances by cereal roots into soil. New Phytology. 1976;76:69-80
7 - Roshchina VV, Roshchina VD. The Excretory Function of Higher Plants. Berlin: Springer; 1993. p. 314. doi: 10.1007/978‐3‐642‐78130‐8
8 - Hage‐Ahmed K, Moyses A, Voglgruber A, Hadacek F, Steinkellner S. Alterations in root exudation of intercropped tomato mediated by the arbuscular mycorrhizal fungus Glomus mosseae and the soilborne pathogen Fusarium oxysporum f. sp. lycopersici. Journal of Phytopathology. 2013;161:763-773. doi:10.1111/jph.12130
9 - Lewis EE, Campbell JF, Griffin C, Kaya HK, Peters A. Behavioral ecology of entomopathogenic nematodes. Biological Control. 2006;38:66-79
10 - Pline M, Dusenbery DB. Responses of plant‐parasitic nematode Meloidogyne incognita to carbon dioxide determined by video camera‐computer tracking. Journal of Chemical Ecology. 1987;13(4):873-888. doi: 10.1007/BF01020167 PMID: 24302053
11 - Dusenbery DB. Theoretical range over which bacteria and nematodes could use carbon dioxide to locate plant roots. Journal of Chemical Ecology. 1987;13:1617-1624
12 - Klinger J. Die Orientierung von Ditylenchus dipsaci in gemessenen künstlichen und biologischen CO2‐Gradienten. Nematologica. 1963;9:185-199
13 - Bauske EM, Rodríguez‐Kábana R, Estaun V, Kloepper JW, Robertson DG, Weaver CF, King PS. Management of Meloidogyne incognita on cotton by use of botanical aromatic compounds. Nematropica. 1994;24:143-150
14 - Oka Y, Nacar S, Putievsky E, Ravid U, Yaniv Z, Spiegel Y. Nematicidal activity of essential oils and their components against the root‐knot nematode. Phytopathology. 2000;90:710-715
15 - Rohlof J. Volatiles from rhizomes of Rhodiola rosea L. Phytochemistry. 2002;59:655-661
16 - Kokalis‐Burelle N, Martinez‐Ochoa N, Rodríguez‐Kabana R, Kloepper JW. Development of multi‐component transplant mixes for suppression of Meloidogyne incognita on tomato (Lycopersicum esculentum). Journal of Nematology. 2002;34:362-369
17 - Bertoli A, Pistelli L, Morelli I, Fraternale D, Giamperi L, Ricci D. Volatile constituents of different parts (roots, stems, and leaves) of Smyrnium olusatrum L. Flavour Fragrance Journal. 2004;19:522-525
18 - Weissteiner S, Schütz S. Are different volatile pattern influencing host plant choice of belowground living insects. Mitteilungen der Deutschen Gesellschaft für Allgemeine und Angewandte Entomologie. 2006;15:51-55
19 - Turlings TCJ, Tumlinson JH, Lewis WJ. Exploitation of herbivore‐induced plant odors by host‐seeking parasitic wasps. Science. 1990;250:1251-1253
20 - Heil M. Indirect defence via tritrophic interactions. New Phytologist. 2008;178:41-61
21 - Dicke M, Baldwin IT. The evolutionary context for herbivore‐induced plant volatiles: Beyond the ‘cry for help’. Trends in Plant Science. 2010;15:167-175
22 - Dicke M, Sabelis MW. How plants obtain predatory mites as bodyguards. Netherlands Journal of Zoology. 1988;38:148-165
23 - Alborn HT, Jones TH, Stenhagen GS, Tumlinson JH. Identification and synthesis of volicitin and related components from beet armyworm oral secretions. Journal of Chemical Ecology. 2000;26:203-220
24 - Turlings TCJ, Alborn HT, Loughrin JH, Tumlinson JH. Volicitin, an elicitor of maize volatiles in the oral secretion of Spodoptera exigua: Its isolation and bio‐activity. Journal of Chemical Ecology. 2000;26:189-202
25 - Rasmann S, Turlings TCJ. First insights into specificity of belowground tritrophic interactions. Oikos. 2008;117:362-369
26 - Ali JG, Alborn HT, Stelinski LL. Constitutive and induced subterranean plant volatiles attract both entomopathogenic and plant‐parasitic nematodes. Journal of Ecology. 2011;99:26-35
27 - van Tol RWHM, van der Sommen ATC, BoV MIC, van Bezooijen J, Sabelis MW, Smits PH. Plants protect their roots by alerting the enemies of grubs. Ecological Letters. 2001;4:292-294
28 - Neveu N, Grandgirard J, Nenon JP, Cortesero AM. Systemic release of herbivore‐induced plant volatiles by turnips infested by concealed root‐feeding larvae Delia radicum L. Journal of Chemical Ecology. 2002;28:1717-1732
29 - Rasman S, Köllner TG, Degenhardt J, Hiltpold I, Toepfer S, Kuhlmann U, Gershenzon J, Turlings TCJ. Recruitment of entomopathogenic nematodes by insect‐damaged maize roots. Nature. 2005;434:732-737
30 - Ali JG, Alborn HT, Stelinski LL. Subterranean herbivore‐induced volatiles released by citrus roots attract entomopathogenic nematodes. Journal of Chemical Ecology. 2010;36:361-368
31 - Duncan LW, Ferguson JJ, Dunn RA, Noling JW. Application of Taylor’s power law to sample statistics of Tylenchulus semipenetrans in Florida citrus. Supplement to the Journal of Nematology (Annals of Applied Nematology). 1989;21:707-711
32 - Hiltpold I, Toepfer S, Kuhlmann U, Turlings TCJ. How maize root volatiles affect the efficacy of entomopathogenic nematodes in controlling the western corn rootworm. Chemoecology. 2010;20:155-162
33 - Degenhardt J, Hiltpold I, Kollner TG, Frey M, Gierl A, Gershenzon J, Hibbard BE, Ellersieck MR, Turlings TCJ. Restoring a maize root signal that attracts insect‐killing nematodes to control a major pest. In: Proceedings the National Academy of Sciences United States of America. 2009;106:13213-13218
34 - Perry RN, Clarke AJ. Hatching mechanisms of nematodes. Parasitology. 1981;83(02):435-449
35 - Curtis RH, Robinson AF, Perry RN. Hatch and Host Location of Root‐Knot Nematodes. Wallington, UK: CAB International; 2009. pp. 139-162. 10.1079/9781845934927.0139
36 - Pudasaini MP, Viaene N, Moens M. Hatching of the root‐lesion nematode, Pratylenchus penetrans, under the influence of temperature and host. Nematology. 2008;10(1):47-54
37 - Chitwood DJ. Phytochemical based strategies for nematode control. Annual Review of Phytopathology. 2002;40(1):221-249
38 - Rasmann S, Hiltpold I, Ali J. The Role of Root-Produced Volatile Secondary Metabolites in Mediating Soil Interactions, Advances in Selected Plant Physiology Aspects, Montanaro, G, editor. InTech Open Access Publisher; 2012 DOI: 10.5772/34304. Available from:
39 - Dutta TK, Powers SJ, Gaur HS, Birkett M, Curtis RH. Effect of small lipophilic molecules in tomato and rice root exudates on the behaviour of Meloidogyne incognita and M. graminicola. Nematology. 2012;14(3):309-320
40 - Steinkellner S, LendzemoV, Langer I, Schweiger P, Khaosaad T, Toussaint J‐P, et al. Flavonoids and strigolactones in root exudates as signals in symbiotic and pathogenic plant‐fungus interactions. Molecules. 2007;12:1290-1306. doi: 10.3390/12071290
41 - McArthur DA, Knowles NR. Resistance responses of potato vesicular‐arbuscular mycorrhizal fungi under varying abiotic phosphorus levels. Plant Physiology. 1992;100:341-351. doi: 10.1104/pp.100.1.341
42 - Harrier LA, Watson CA. The potential role of arbuscular mycorrhizal (AM) fungi in the bio‐protection of plants against soil‐borne pathogens in organic and/or other sustainable farming systems. Pest Management Science. 2004;60:149-157. doi: 10.1002/ps.820
43 - López‐Ráez JA, Charnikhova T, Fernández I, Bouwmeester H, Pozo MJ. Arbuscular mycorrhizal symbiosis decreases strigolactone production in tomato. Journal of Plant Physiology. 2011;168:294-297. doi: 10.1016/j.jplph.2010. 08.011
44 - Sood GS. Chemotactic response of plant‐growth‐promoting bacteria towards roots of vesicular‐arbuscular mycorrhizal tomato plants. FEMS Microbiology Ecology. 2003;45:219-227. doi: 10.1016/S0168‐6496(03)00155‐7
45 - Druzhinina IS, Seidl‐Seiboth V, Herrera‐Estrella A, Horwitz BA, Kenerley CM, Monte E, et al. Trichoderma: The genomics of opportunistic success. Nature Reviews Microbiology. 2011;9:749-759. doi: 10.1038/nrmicro2637
46 - Dong LQ, Zhang KQ. Microbial control of plant‐parasitic nematodes: A five‐party interaction. Plant Soil. 2006;288:31-45. doi: 10.1007/s11104‐006‐9009‐3
47 - Sikora RA, Pocasangre L, FeldeZum A, Niere B, Vu TT, Dababat AA. Mutualistic endophytic fungi and in planta suppressiveness to plant‐parasitic nematodes. Biological Control. 2008;46:15-23. doi: 10.1016/j.biocontrol.2008.02.011
48 - Vos C, Claerhout S, Mkandawire R, Panis B, de Waele D, Elsen A. Arbuscular mycorrhizal fungi reduce root‐knot nematode penetration through altered root exudation of their host. Plant Soil. 2012b;354:335-345. 10.1007/s11104‐011‐1070‐x
49 - Yang G, Zhou B, Zhang X, Zhang Z, Wu Y, Zhang Y, et al. Effects of tomato root exudates on Meloidogyne incognita. PLoS One. 2016;11(4):e0154675. doi: 10.1371/journal.pone.0154675
50 - Kobra N, Jalil K, Youbert G. Effects of three Glomus species as biocontrol agents against Verticillium‐induced wilt in cotton. Journal of Plant Protection Research. 2009;49:185-189. 10.2478/v10045‐009‐0027‐z
51 - Lioussanne L. The role of the arbuscular mycorrhiza‐associated rhizobacteria in the biocontrol of soilborne phyto‐pathogens. Spanish Journal of Agricultural Research. 2010;8:3-5. doi: 10.5424/sjar/201008S1‐5301
52 - Scheublin TR, Sanders IR, Keel C, van der Meer JR. Characterisation of microbial communities colonising the hyphal surfaces of arbuscular mycorrhizal fungi. ISMEJ. 2010;4:752-763. doi: 10.1038/ismej. 2010.5
53 - Miransari M. Interactions between arbuscular mycorrhizal fungi and soil bacteria. Applied Microbiology Biotechnology. 2011;89:917-930. doi: 10.1007/s00253‐010‐3004‐6
54 - Nuccio EE, Hodge A, Pett‐Ridge J, Herman DJ, Weber PK, Firestone MK. An arbuscular mycorrhizal fungus significantly modifies the soil bacterial community and nitrogen cycling during litter decomposition. Environmental Microbiology. 2013;15:1870-1881. doi: 10.1111/1462‐2920.12081
55 - Mankau R. Biological control of nematodes pests by natural enemies. Annual Review of Phytopathology. 1980;18:415-440
56 - Jatala P. Biological control of plant‐parasitic nematodes. Annual Review of Phytopathology. 1986;24:453-489
57 - Tranier M, Pognant-Gros J, De la Cruz Quiroz R, González C, Mateille T, Roussos S. Commercial Biological Control Agents Targeted Against Plant-Parasitic Root-knot Nematodes. Brazilian Archives of Biology and Technology. 2014;57:831-841.
58 - Siddiqui ZA, Mahmood I. Role of bacteria in the management of plant‐parasitic nematodes: A review. Bioresource Technology. 1999;69:167-179
59 - Emmert EAB, Handelsman J. Biocontrol of plant disease: A (Gram) positive perspective. FEMS Microbiology Letters. 1999;171:1-9
60 - Meyer SLF. United States Department of Agriculture—Agricultural Research Service research programs on microbes for management of plant‐parasitic nematodes. Pest Management Science. 2003;59:665-670
61 - Abd‐Elgawad MMM, Vagelas IK. Nematophagous Bacteria: Field Application and Commercialization. In: Askary TH, Martinelli PRP, editors. Biocontrol Agents of Phytonematodes. UK: CAB International; 2015. pp. 276-309. 10.1079/9781780643755.0276
62 - Sayre RM, Starr MP. Pasteuria penetrans (ex Thorne 1940) non. rev. comb. n. sp. n. a mycelial and endospore forming bacterium parasite in plant parasitic nematodes. Proceedings of the Helminthological Society of Washington. 1985;52:149-165
63 - Bekal S, Borneman J, Springer MS, Giblin‐Davis RM, Becker JO. Phenotypic and molecular analysis of a Pasteuria strain parasitic to the sting nematode. Journal of Nematology. 2001;33:110-115
64 - Kun XC, Jun LX, Qin XJ, Lei G, Qun DC, He MM, Qin ZK, Xiang YF, Huang FD. Phylogenetic analysis of the nematicidal actinobacteria from agricultural soil of China. African Journal of Microbiology Research. 2011;5(16):2316-2324
65 - Mishra SK, Keller JE, Miller JR, Heisey RM, Nair MG, Putnam AR. Insecticidal and nematicidal properties of microbial metabolites. Indian Journal of Microbiology. 1987;2:267-276
66 - Dicklow MB, Acosta N, Zuckerman BM. A novel Streptomyces species for controlling plant‐parasitic nematodes. Journal of Chemical Ecology. 1993;19:159-173
67 - Nour SM, Lawrence JR, Zhu H, Swerhone GDW, Welsh M, Welacky TW, Topp E. Bacteria associated with cysts of the soybean cyst nematode (Heterodera glycines). Applied Environmental Microbiology. 2003;69:607-615
68 - Burg RW, Miller BM, Baker EE, Birnbaum J, Currie SA, Hartman R, Kong YL, Richard L, Monaghan RL, Olsonm G, Putter I, Tunac JB, Wwllick H, Stapley EO, Oiwa R, Omura S. Avermectins, new family of potent anthelmintic agents: Producing organism and fermentation. Antimicrobial Agents and Chemotherapy. 1979;15:361-367
69 - Sun MH, Li G, Shi YX, Li BJ, Liu XZ. Fungi and actinomycetes associated with Meloidogyne spp. eggs and females in China and their biocontrol potential. Journal of Invertebrate Pathology. 2006;93:22-28
70 - Ruanpanun P, Laatsch H, Tangchitsomkid N, Lumyong S. Nematicidal activity of fervenulin isolated from a nematicidal actinomycete, Streptomyces sp. CMU‐MH021, on Meloidogyne incognita. World Journal of Microbiology and Biotechnology. 2011;27(6):1373-1380. doi: 10.1007/s11274‐010‐0588‐z
71 - Kaur T, Jasrotia S, Ohri P, Manhas RK. Evaluation of in vitro and in vivo nematicidal potential of a multifunctional streptomycete, Streptomyces hydrogenans strain DH16 against Meloidogyne incognita. Microbiological Research. 2016;192:247-252. doi: 10.1016/j.micres.2016.07.009
72 - Xiong J. Zhou Q, Luo H, Xia L, Li L, Sun M, Yu Z. Systemic nematicidal activity and biocontrol efficacy of Bacillus firmus against the root‐knot nematode Meloidogyne incognita. World Journal of Microbiology and Biotechnology. 2015;31:661. doi: 10.1007/s11274‐015‐1820‐7
73 - Sayre M, Starr MP. Bacterial diseases and antagonism of nematode. In: Poinar Jr, GO, Jannso HB, editors. Diseases of Nematodes. Boca Raton, FL: CRC Press; 1988. pp. 69-101
74 - Giblin‐Davis RM, Williams DS, Bekal S, Dickson DW, Brito JA, Becker JO, Preston JF. Candidatus Pasteuria usage sp. nov., an obligate endoparasite of the phytoparasitic nematode Belonlaimus longicaudatus. International Journal of Systematic and Evolutionary Microbiology. 2003;53:197-200
75 - Mohan S, Mauchline TH, Rowe J, Hirsch PR, Davies KG. Pasteuria endospores from Heterodera cajani (Nematoda: Heteroderidae) exhibit inverted attachment and altered germination in cross‐infection studies with Globodera pallida (Nematoda: Heteroderidae). FEMS Microbiology Ecology. 2012;79:675-684
76 - Wilson MJ, Jackson TA. Progress in the commercialisation of bionematicides. BioControl. 2013;58:715-722
77 - Schüßler AH, Gehrig H, Schwarzott D, Walker C. Analysis of partial Glomales SSU rRNA gene sequences: Implications for primer design and phylogeny. Mycological Research. 2001;105:5-15. doi: 10.1017/S0953756200003725
78 - Smith SE, Read DJ. Mineral nutrition, toxic element accumulation and water relations of arbuscular mycorrhizal plants. In: Smith SE, Read DJ, editors. Mycorrhizal Symbiosis. 3rd ed. London: Academic Press; 2008. pp. 145-148
79 - Porcel R, Aroca R, Ruiz‐Lozano JM. Salinity stress alleviation using arbuscular mycorrhizal fungi. A review. Agronomy for Sustainable Development. 2011;32:181-200. doi: 10.1007/s13593‐011‐0029‐x
80 - Augé RM, Toler HD, Saxton AM. Arbuscular mycorrhizal symbiosis alters stomatal conductance of host plants more under drought than under amply watered conditions: A meta‐analysis. Mycorrhiza. 2015;25:13-24. doi: 10.1007/s00572‐014‐0585‐4
81 - Tchabi A, Hountondji FCC, Ogunsola B, Lawouin L, Coyne D, Wiemken A, Oehl F. The influence of arbuscular mycorrhizal fungi inoculation on micro‐propagated hybrid yam (Dioscorea spp.) growth and root‐knot nematode (Meloidogyne spp.) suppression. International Journal of Current Microbiology and Applied Sciences. 2016;5(10):267-281
82 - Ceballos I, Ruiz M, Fernandez C, Pena R, Rodriguez A, Sanders IR. The in vitro mass‐produced model mycorrhizal fungus, Rhizophagus irregularis, significantly increases yields of the globally important food security crop Cassava. PLoS One. 2013;8:e70633. doi: 10.1371/journal.pone.0070633
83 - Khaosaad T, García‐Garrido JM, Steinkellner S, Vierheilig H. Take‐all disease is systemically reduced in roots of mycorrhizal barley plants. Soil Biology and Biochemistry. 2007;39:727-734. 10.1016/j.soilbio.2006.09.014
84 - Castellanos‐Morales V, Keiser C, Cárdenas‐Navarro R, Grausgruber H, Glauninger J, García‐Garrido JM, et al. The bioprotective effect of AM root colonization against the soil‐borne fungal pathogen Gaeumannomyces graminis var. tritici in barley depends on the barley variety. Soil Biology and Biochemistry. 2011;43:831-834. 10.1016/j.soilbio.2010.12.020
85 - Zamioudis C, Pieterse CMJ. Modulation of host immunity by beneficial microbes. Molecular Plant Microbe Interactions. 2012;25:139-150. 10.1094/MPMI‐06‐11‐0179
86 - Boller T, Felix G. A renaissance of elicitors: Perception of microbe‐associated molecular patterns and danger signals by pattern recognition receptors. Annual Review of Plant Biology. 2009;60:379-406
87 - Jones JDG, Dangl JL. The plant immune system. Nature. 2006;444:323-329. 10.1038/nature05286
88 - Kloppholz S, Kuhn H, Requena N. A secreted fungal effector of Glomus intraradices promotes symbiotic biotrophy. Current Biology. 2011;21:1204-1209
89 - Oldroyd GED, Harrison MJ, Paszkowski U. Reprogramming plant cells for endosymbiosis. Science. 2009;324(5928):753-754. doi: 10.1126/science.1171644
90 - Besserer A, Becard G, Jauneau A, Roux C, Sejalon‐Delmas N. GR24, a synthetic analog of strigolactones, stimulates the mitosis and growth of the arbuscular mycorrhizal fungus Gigaspora rosea by boosting its energy metabolism. Plant Physiology. 2008;148:402413. doi: 10.1104/pp.108.121400
91 - Parniske M. Arbuscular mycorrhiza: The mother of plant root endosymbioses. Nature Reviews Microbiology. 2008;6:763775
92 - Rosas S, Soria R, Correa N, Abdala G. Jasmonic acid stimulates the expression of nod‐genes in rhizobium. Plant Molecular Biology. 1998;38:1161-1168
93 - Mabood F, Souleimanov A, Khan W, Smith DL. Jasmonates induce Nod factor production by Bradyrhizobium japonicum. Plant Physiology and Biochemistry. 2006;44:759-765. doi: 10.1038/nrmicro1987
94 - Viaene N, Coyne DL, Kerry BR. Biological and cultural management. In: Perry RN, Moens M, editors. Plant Nematology. Wallingford, UK: CAB International; 2006. pp. 346-369
95 - Yu O, Coosemans J. Fungi associated with cysts of Globodera rostochiensis, G‐pallida, and Heterodera schachtii; and egg masses and females of Meloidogyne hapla in Belgium. Phytoprotection. 1998;79:63-69. DOI: 10.7202/706135ar
96 - Kerry BR. Rhizosphere interactions and the exploitation of microbial agents for the biological control of plant‐parasitic nematodes. Annual Review of Phytopathology. 2000;38:423-441. DOI: 10.1146/annurev.phyto.38.1.423
97 - Whitehead AG. Plant Nematode Control. Wallingford, UK: CAB International; 1998. p. 384
98 - Hallmann J, Davies KG, Sikora R. Biological control using microbial pathogens, endophytes and antagonists. In: Perry RN, Moens M, Starr JL, editors. Root‐Knot Nematodes. Wallingford, UK: CAB International; 2009. pp. 380-411
99 - Kerry BR, Jaffee BA. Fungi as biological control agents for plant parasitic nematodes. In: Wicklow DT, Soderstrom BE, editors. The Mycota: A Comprehensive Treatise on Fungi as Experimental Systems for Basic and Applied Research, Volume 4 Environmental and Microbial Relationships. Berlin: Springer; 1997. pp. 203-218
100 - Lopez‐Llorca LV, Macià‐Vicente JG, Jansson H‐BJ. Mode of action and interactions of nematophagous fungi. In: Ciancio A, Mukerji KG, editors. Integrated Management and Biocontrol of Vegetable and Grain Crops Nematode. Dordrecht: Springer; 2008. pp. 51-76
101 - Anke H, Stadler M, Mayer A, Sterner O. Secondary metabolites with nematicidal and antimicrobial activity from nematophagous fungi and Ascomycetes. Canadian Journal of Botany. 1995;73:932-939. DOI: 10.1139/b95‐341
102 - Chet I, Harman GE, Baker R. Trichoderma hamatum: Its hyphal interactions with Rhizoctonia solani and Pythium spp. Microbial Ecology. 1981;7:29-38. DOI: 10.1007/BF02010476
103 - Windham GL, Windham MT, Williams WP. Effects of Trichoderma spp. on maize growth and Meloidogyne arenaria reproduction. Plant Disease. 1989;73:493-495. DOI: 10.1094/PD‐73‐0493
104 - Spiegel Y, Chet I: Evaluation of Trichoderma spp. as a biocontrol agent against soilborne fungi and plant‐parasitic nematodes in Israel. Integrated Pest Management Review. 1998;3:1-7. DOI: 10.1023/A:1009625831128
105 - Sharon E, Chet I, Viterbo A, Bar‐Eyal M, Nagan H, Samuels GJ, Spiegel Y. Parasitism of Trichoderma on Meloidogyne javanica and role of the gelatinous matrix. European Journal of Plant Pathology. 2007;118:247-258. DOI: 10.1007/s10658‐007‐9140‐x
106 - Affokpon A, Coyne DL, De Proft M, Coosemans J. In vitro growth characterization and biocontrol potential of naturally occurring nematophagous fungi recovered from root‐knot nematode infested vegetable fields in Benin. International Journal of Pest Management. 2015;61:273-283. DOI: 10.1080/09670874.2015.1043971
107 - Chet I, Inbar J, Hadar Y. Fungal antagonists and mycoparasitism. In: Wicklow DT, Soderstrom BE, editors. The Mycota. Volume IV: Environmental and Microbial Relationships. Heidelberg: Springer‐Verlag; 1997. pp. 165-184
108 - Howell CR. Mechanisms employed by Trichoderma species in the biological control of plant diseases: The history and evolution of current concepts. Plant Disease. 2003;87:4-10. DOI: 10.1094/PDIS.2003.87.1.4
109 - Rodríguez‐Kábana R, Morgan‐Jones G, Godroy G, Gintis BO. Effectiveness of species of Gliocladium, Paecilomyces and Verticillium for control of Meloidogyne arenaria in field soil. Nematropica. 1984;14:155-170
110 - Freitas LG, Ferraz S, Muchovey JJ. Effectiveness of different isolates of Paecilomyces lilacinus and an isolate of Cylindorcarpon destructans on the control of Meloidogyne javanica. Nematropica. 1995;25:109-115
111 - Goswami J, Pandey RK, Tewari JP, Goswami BK. Management of root knot nematode on tomato through application of fungal antagonists, Acremonium strictum and Trichoderma harzianum. Journal of Environmental Science and Health. 2008;43:237-240. DOI: 10.1080/03601230701771164
112 - Sikora RA, Schönbeck F. Effect of vesicular‐arbuscular mycorrhizae, Endogone mosseae on the population dynamics of the root‐knot nematodes Meloidogyne incognita and M. hapla. In: Proceedings of the 8th International Congress of Plant Protection; 21-27 August 1975; Moscow. pp. 158-166
113 - Jeffries P, Gianinazzi S, Perotto S, Turnau K, Barea JM. The contribution of arbuscular mycorrhizal fungi in sustainable maintenance of plant health and soil fertility. Biology and Fertility of Soils. 2003;37:1-16. DOI: 10.1007/s00374‐002‐0546‐5
114 - Harley JL, Smith SE. Mycorrhizal Symbiosis. London: Academic Press; 1983. p. 483
115 - Bonfante‐Fasolo P. Anatomy and morphology VA mycorrhizae. In: Powell CL, Bagyaraj DJ, editors. VA Mycorrhiza. Boca Raton: CRC Press; 1984. pp. 5-32
116 - Peyronel B, Fassi B, Fontana A, Trappe JM. Terminology of mycorrhizae. Mycologia. 1969;61:410-441
117 - Azcón‐Aguilar C, Barea JM. Arbuscular mycorrhizas and biological control of soil‐borne plant pathogens—An overview of the mechanisms involved. Mycorrhiza. 1996;6:457-464. DOI:10.1007/s005720050147
118 - Ruiz‐Lozano JM. Arbuscular mycorrhizal symbiosis and alleviation of osmotic stress: New perspectives for molecular studies. Mycorrhiza. 2003;13:309-317. DOI: 10.1007/s00572‐003‐0237‐6
119 - Whipps JM. Prospects and limitations for mycorrhizals in biocontrol of root pathogens. Canadian Journal of Botany. 2004;82:1198-1227. DOI: 10.1139/b04‐082
120 - Elsen A, Gervacio D, Swennen R, De Waele D. AMF‐induced biocontrol against plant parasitic nematodes in Musa sp.: A systemic effect. Mycorrhiza. 2008;18:251-256. DOI: 10.1007/s00572‐008‐0173‐6
121 - Hallmann J, Sikora RA. Influence of F. oxysporum, a mutualistic fungal endophyte on M.  incognita on tomato. Journal of Plant Diseases and Protection. 1994a;101:475-481
122 - Dababat AA, Sikora RA. Induced resistance by the mutualistic endophyte, Fusarium oxysporum strain 162, toward Meloidogyne incognita on tomato. Biocontrol Science and Technology. 2007a;17:969-975. DOI:10.1080/09583150701582057
123 - Dababat AA, Sikora RA. Influence of the mutualistic endophyte Fusarium oxysporum 162 on Meloidogyne incognita attraction and invasion. Nematology. 2007b;9:771-776. DOI: 10.1163/156854107782331225
124 - Kerry BR. Exploitation of the nematophagous fungal Verticillium chlamydosporium Goddard for the biological control of root knot nematodes (Meloidogyne spp.). In: Butt MT, Jackson CW, Magan N, editors. Fungi as Biocontrol Agents‐Progress, Problems and Potential. Wallingford, UK: CAB International; 2001. pp. 155-166
125 - Zare R, Gams W. A revision of Verticillium sect. Prostrata. III. Generic classification. Nova Hedwigia. 2001;72:329-337
126 - Chen S, Dickson DW. Biological control of nematodes by fungal antagonists. In: Chen ZX, Chen SY, Dickson DW, editors. Nematology Advances and Perspectives. Volume II Nematode Management and Utilization. Wallingford, UK: CAB International; 2004. pp. 977-1039
127 - Kerry BR, Bourne JM. The importance of rhizosphere interactions in the biological control of plant parasitic nematodes—a case study using Verticillium chlamydosporium. Pesticide Science. 1996;47:69-75. DOI: 10.1002/(SICI)1096‐9063(199605)47:1<69::AID‐PS386>3.0.CO;2‐6
128 - Jalali AAH, Segers R, Coosemans J. Biocontrol of Heterodera schachtii using combinations of the sterile fungus, StFCh1‐1, Embellisia chlamydospora and Verticillium chlamydosporium. Nematologica. 1998;44:345-355. DOI: 10.1163/005525998X00025
129 - Lysek H, Sterba J. Colonization of Ascaris lumbricoides eggs by the fungus Verticillium chlamydosporium Goddard. Folia Parasitologica. 1991;8:255-259
130 - Segers R, Butt TM, Kerry BR, Beckett A, Peberdy JF. The role of the proteinase VCP1 produced by the nematophagous Verticillium chlamydpsporium in the infection process of nematode eggs. Mycological Research. 1996;100:421-428
131 - Morton CO, Hirsch AM, Kerry BR. 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. Nematology. 2004;6:161-170. DOI: 10.1163/1568541041218004
132 - Dackman C. Fungal parasites of the potato cyst nematode Globodera rostochiensis: Isolation and reinfection. Journal of Nematology. 1990;22:594-597
133 - Kerry BR, De Leij AAM. Key factors in the development of fungal agents for the control of cyst and root‐knot nematodes. In: Tjamos EC, Papavizas GC, Cook RJ, editors. Biological Control of Plant Diseases. New York, NY: Plenum Press; 1992. pp. 139-144
134 - De Leij FAAM, Kerry BR. The nematophagous fungus Verticillium chlamydosporium Goddard, as a potential biological control agent for Meloidogyne arenaria (Neal) Chitwood. Revue de Nématologie. 1991;14:157-164
135 - Bourne JM, Kerry BR, De Leij FAAM. The importance of the host plant in the interaction between root‐knot nematodes (Meloidogyne spp.) and the nematophagous fungus Verticillium chlamydosporium Goddard. Biocontrol Science and Technology. 1996;6:539-548. DOI: 10.1080/09583159631172
136 - Kerry BR, Bourne JM. A Manual for Research on Verticillium chlamydosporium, a Potential Biological Control Agent for Root‐Knot Nematodes. Darmstadt: Druckform GmbH; 2002. p. 84
137 - De Oca NM, Arévalos J, Nuñez A, Riverón Y, Villoch A, Hidalgo‐Díaz L. KlamiC: Experiencia técnica‐productiva. Revista de Protección Vegetal. 2009;24:62-65
138 - Esposito E, Da Silva M. Systematics and environmental application of the genus Trichoderma. Critical Reviews in Microbiology. 1998;24:89-98. DOI: 10.1080/10408419891294190
139 - Harman GE, Howell CR, Viterbo A, Chet I, Lorito M. Trichoderma species—Opportunistic, avirulent plant symbionts. Nature Reviews, Microbiology. 2004;2:43-56. DOI: 10.1038/nrmicro797
140 - Kyalo G, Affokpon A, Coosemans J, Coyne DL. Biological control effects of Pochonia chlamydosporia and Trichoderma isolates from Benin (West‐Africa) on root‐knot nematodes. Communications in Agricultural and Applied Biological Sciences. 2007;72:219-223
141 - Affokpon A, Coyne DL, Htay CC, Dossou Agbèdè R, Lawouin L, Coosemans J. Biocontrol potential of native Trichoderma isolates against root‐knot nematodes in West African vegetable production systems. Soil Biology and Biochemistry. 2011a;43:600-608. DOI: 10.1016/j.soilbio.2010.11.029
142 - Kullnig‐Gradinger CM, Szakacs G, Kubicek CP. Phylogeny and evolution of the fungal genus Trichoderma‐a multigene approach. Mycological Research. 2002;106:757-767. DOI: 10.1017/S0953756202006172
143 - Yedidia I, Benhamou N, Chet I. Induction of defense response in cucumber plants (Cucumis sativus L.) by the biocontrol agent Trichoderma harzianum. Applied Environmental Microbiology. 1999;65:1061-1070
144 - Sikora RA, Oka Y, Sharon, E, Hok CJ, Keren‐Zur M. Achievements and research requirements for the integration of biocontrol into farming systems. Nematology. 2000;2:737-738. DOI: 10.1163/156854100509592
145 - Bennett AJ, Mead A, Whipps JM. Performance of carrot and onion seed primed with beneficial microorganisms in glasshouse and field trials. Biological Control. 2009;51:417-426. DOI: 10.1016/j.biocontrol.2009.08.001
146 - Rilling MC, Wright SF, Eviner VT. The role of arbuscular mycorrhizal fungi and glomalin in soil aggregation: Comparing effects of five plant species. Plant and Soil. 2002;238:325-333. DOI: 10.1023/A:1014483303813
147 - Tchabi A, Burger S, Coyne D, Hountondji F, Lawouin L, Wiemken A, Oehl F. Promiscuous arbuscular mycorrhizal symbiosis of yam (Dioscorea spp.), a key staple crop in West Africa. Mycorrhiza. 2009;19:375-392. DOI: 10.1007/s00572‐009‐0241‐6
148 - Tchabi A, Coyne D, Hountondji F, Lawouin L, Wiemken A, Oehl F. Efficacy of indigenous arbuscular mycorrhizal fungi for promoting white yam (Dioscorea rotundata) growth in West Africa. Applied Soil Ecology. 2010;45:92-100. DOI: 10.1016/j.apsoil.2010.03.001
149 - Davies JFT, Puryear JD, Newton RJ, Egilla JN, Saraiva Grossi SJA. Mycorrhizal fungi enhance accumulation and tolerance of chromium in sunflower (Helianthus annuus). Journal of Plant Physiology. 2001;158:777-786. DOI: 10.1078/0176‐1617‐00311
150 - Chen BD, Roos P, Borggaard OK, Zhu Y‐G, Jakobsen I. Mycorrhiza and root hairs in barley enhance acquisition of phosphorus and uranium from phosphate rock but mycorrhiza decreases root to shoot uranium transfer. New Phytologists. 2005;165:591-598. DOI: 10.1111/j.1469‐8137.2004.01244.x
151 - Al‐Karaki GN. Benefit, cost and water‐use efficiency of arbuscular mycorrhizal during wheat grown under drought stress. Mycorrhiza. 1998;8:41-45. DOI: 10.1007/s005720050209
152 - Jaizme‐Vega MC, Tenoury P, Pinochet J, Jaumot M. Interactions between the root‐knot nematode Meloidogyne incognita and Glomus mosseae in banana. Plant and Soil. 1997;196:27-35. DOI:10.1023/A:1004236310644
153 - Declerck, S, Risèd JM, Rufyikiri G, Delvaux B. Effects of arbuscular mychorrhizal fungi on severity of root rot of bananas caused by Cylindrocladium spathiphylli. Plant Pathology. 2002;51:109-115. DOI: 10.1046/j.0032‐0862.2001.656.x
154 - Gange AC, Brown VK, Aplin D. Multitrophic links between arbuscular mycorrhizal fungi and insect parasitoids. Ecology Letters. 2003;6:1051-1055. DOI: 10.1046/j.1461‐0248.2003.00540.x
155 - Marro N, Lax P, CabelloM, Doucet ME, Becerra AG. Use of the arbuscular mycorrhizal fungus Glomus intraradices as biological control agent of the nematode Nacobbus aberrans Parasitizing Tomato. Brazilian Archives of Biology and Technology. 2014;57:668-674. DOI: 10.1590/S1516‐8913201402200.
156 - Mohanty KC, Sahoo NK. Prospects of mycorrhizae as potential nematode antagonist. In: Trivedi PC, editor. Advances in Nematology. India: Scientific Publishers; 2003. p. 317
157 - Hol WHG, Cook R. An overview of arbuscular mycorrhizal fungi‐nematode interactions. Basic and Applied Ecology. 2005;6:489-503. DOI: 10.1016/j.baae.2005.04.001
158 - Castillo P, Nico AI, Azcón‐Aguilar C, Del Río Rincón C, Calvet C, Jiménez‐Díaz RM. Protection of olive planting stocks against parasitism of root‐knot nematodes by arbuscular mycorrhizal fungi. Plant Pathology. 2006;55:705-713. DOI: 10.1111/j.1365‐3059.2006.01400.x
159 - Masadeh B, Von Alten H, Grunewaldt‐Stoecker G, Sikora RA. Biocontrol of root‐knot nematodes using the arbuscular mycorrhizal fungus Glomus intraradices and the antagonist Trichoderma viride in two tomato cultivars differing in their suitability as hosts for the nematodes. Journal Plant Disease and Protection. 2004;111:322-333
160 - Ryan NA, Deliopoulos T, Jones P, Haydock PPJ. Effects of a mixed‐isolate mycorrhizal inoculum on the potato‐potato cyst nematode interaction. Annals of Applied Biology 2003;143:111-119. DOI: 10.1111/j.1744‐7348.2003.tb00275.x
161 - Schwob I, Ducher M, Coudret A. Effects of climatic factors on native arbuscular mycorrhizae and Meloidogyne exigua in a Brazilian rubber tree (Hevea brasiliensis) plantation. Plant Pathology. 1999;48:19-25. DOI: 10.1046/j.1365‐3059.1999.00300.x
162 - Waceke JW, Waudo SW, Sikora R. Effect of inorganic phosphatic fertilizers on the efficacy of an arbuscular mycorrhiza fungus against a root‐knot nematode on pyrethrum. International Journal of Pest Management. 2002;48:307-313. DOI: 10.1080/09670870210149862
163 - De La Peña E, Echeverría SR, Van Der Putten HH, Freitas H, Moens M. Mechanism of control of root‐feeding nematodes by mycorrhizal fungi in the dune grass Ammophila arenaria. New Phytologists. 2006;169:829-840. DOI: 10.1111/j.1469‐8137.2005.01602.x
164 - Zhang L, Zhang J, Christie P, Li X. Pre‐inoculation with arbuscular mycorrhizal fungi suppresses root knot nematode (Meloidogyne incognita) on cucumber (Cucumis sativus). Biology and Fertility of Soils. 2008;45:205-212. DOI: 10.1007/s00374‐008‐0329‐8
165 - Affokpon A, Coyne DL, Lawouin L, Tossou C, Dossou Agbèdè R, Coosemans J. Effectiveness of native West African arbuscular mycorrhizal fungi in protecting vegetable crops against root‐knot nematodes. Biology and Fertility of Soils. 2011b;47:207-217. DOI 10.1007/s00374‐010‐0525‐1
166 - Diedhiou PM, Hallmann J, Oerke EC, Dehne HW. Effects of arbuscular mycorrhizal fungi and a non‐pathogenic Fusarium oxysporum on Meloidogyne incognita infestation of tomato. Mycorrhiza. 2003;13:199-204. DOI 10.1007/s00572‐002‐0215‐4
167 - Rumbos C, Reimann S, Kiewnick S, Sikora RA. Interactions of Paecilomyces lilacinus strain 251 with the mycorrhizal fungus Glomus intraradices: Implications for Meloidogyne incognita control on tomato. Biocontrol Science and Technology. 2006;16:981-986. DOI: 10.1080/09583150600937667
168 - Domsch KH, Gams W, Anderson T‐H. Compendium of Soil Fungi. London: Academic Press; 1980. p. 406
169 - Jatala P, Kaltenback R, Bocangel M. Biological control of Meloidogyne acrita and Globodera pallida on potatoes. Journal of Nematology. 1979;11:303
170 - Holland RJ, Williams KL, Khan A. Infection of Meloidogyne javanica by Paecilomyces lilacinus. Nematology. 1999;1:131-139. DOI: 10.1163/156854199508090
171 - Kiewnick S, Sikora RA. Biological control of the root‐knot nematode Meloidogyne incognita by Paecilomyces lilacinus strain 251. Biological Control. 2006;38:179-187. DOI: 10.1016/j.biocontrol.2005.12.006
172 - Speijer PR. Interrelationships between Pratylenchus goodeyi Sher & Allen and strains of nonpathogenic Fusarium oxysporum Schl. emd. Snyd. & Hans. in roots of two banana cultivars [thesis]. Bonn: University of Bonn; 1993
173 - Schuster RP, Sikora RA, Amin N. Potential of endophytic fungi for the biological control of plantlet parasitic nematodes. Mededelingen van de Faculteit Landbouwwetenschappen Rijksuniversiteit Gent. 1995;60:1047-1052
174 - Griesbach M. Occurrence of mutualistic fungal endophytes in bananas (Musa spp.) and their potential as biocontrol agents of banana weevil Cosmopolites sordidus (Germar) in Uganda [thesis]. Bonn: University of Bonn; 2000
175 - Hallmann J, Sikora RA. Occurrence of plant parasitic nematodes and nonpathogenic species of Fusarium in tomato plants in Kenya and their role as mutualistic synergists for biological control of root knot nematodes. International Journal of Pest Management. 1994b;40:321-325. DOI: 10.1080/09670879409371907
176 - Pocasangre LE. Biological enhancement of banana tissue culture plantlets with endophytic fungi for the control of the burrowing nematode Radopholus similis and the Panama disease (Fusarium oxysporum f.sp. cubense) [thesis]. Bonn: University of Bonn; 2000
177 - Niere B. Significance of non‐pathogenic isolates of Fusarium oxysporum Schlecht.: Fries for the biological control of the burrowing nematode Radopholus similis (Cobb) Thorne on tissue cultured banana [thesis]. Bonn: University of Bonn; 2001
178 - Vu TT, Hauschild R, Sikora RA. Fusarium oxysporum endophytes induced systemic resistance against Radopholus similis on banana. Nematology. 2006;8:847-852. DOI: 10.1163/156854106779799259
179 - Mennan S, Aksoy HM, Ecevit O. Antagonistic effect of Fusarium oxysporum on Heterodera cruciferae. Journal of Phytopathology. 2005;153:221-225. DOI: 10.1111/j.1439‐0434.2005.00957.x.
180 - Waweru BW, Turoop L, Kahangi E, Coyne D, Dubois T. Non‐pathogenic Fusarium oxysporum endophytes provide field control of nematodes, improving yield of banana (Musa sp.). Biological Control. 2014;74:82-88. DOI: 10.1016/j.biocontrol.2014.04.002
181 - Al kader MAA. In vitro studies on nematode interactions with their antagonistic fungi in the rhizosphere of various plants [thesis]. Freiburg im Breisgau: Universität Freiburg im Breisgau; 2008
182 - Zopf W. Zur kenntnis der infektions‐krankheiten niederer thiere und pflanzen. Nova Acta Academiae Caesareae Leopoldino‐Carolinae Germanicae Naturae Curiosorum. 1888;52:314-376
183 - Niu X‐M, Zhang K‐Q. Arthrobotrys oligospora: A model organism for understanding the interaction between fungi and nematodes. Mycology. 2011;2:59-78. DOI: 10.1080/21501203.2011.562559
184 - Saikawa M, Takahashi A. Nonconstricting‐ring formation in two species of nematode‐capturing hyphomycetes. Mycoscience. 2002;43:417-419. DOI: 10.1007/s102670200061
185 - Ferris H, Castro CE, Caswell EP, Jaffee BA, Roberts PA, Westerdahl BB, Williamson VM. Biological approaches to the management of plant‐parasitic nematodes. In: Madden JP, editor. Beyond Pesticides: Biological Approaches to Pest Management in California. CA: University of California; 1992. pp. 68-101
186 - Nordbring‐Hertz B, Jansson H‐B, Tunlid A. Nematophagous fungi. Encyclopedia of Life Science. 2002;12:681-690. DOI: 10.1038/npg.els.0004293
187 - Santos MA, Ferraz S, Muchovej JJ. Evaluation of 20 species of fungi from Brazil for biocontrol of Meloidogyne incognita race 3. Nematropica. 1992;22:183-192
188 - Nordbring‐Hertz B, Jansson H‐B, Friman E, Persson Y, Dackman C, Hard T, Poloczek E, Feldmann R. Nematophagous Fungi. Film no. V, 1851. Göttingen: Institut für den Wissenschaftlichen Film; 1995
189 - Martin SB. Nematode control. Available at: (accessed 05 may, 2017).
190 - Wang B, Liu XZ. Mass production and formulation of nematode‐antagonistic microbes. In: Liu XZ, Zhang KQ, Li TF, editors. Biological Control of Plant‐Parasitic Nematodes (in Chinese, with English abstract). Beijing, China: China Science and Technology Press; 2004, pp. 285-297.
191 - Meyer SLF, Roberts DP, Chitwood DJ, Carta LK, Lumsden RD, Mao W. Application of Burkholderia cepacia and Trichoderma virens, alone and in combinations, against Meloidogyne incognita on bell pepper. Nematropica. 2001;31:75-86.
192 - Raddy HM, Fouad AFA, Montasser SA, Abdel‐Lateef MF, El‐Samadisy AM. Efficacy of six nematicides and six commercial bioproducts against root‐knot nematode, Meloidogyne incognita on tomato. Journal of Applied Sciences Research. 2013;9:4410-4417.
193 - Abd‐Elgawad MMM, Aboul‐Eid HZ. Effects of oxamyl, insect nematodes and Serratia marcescens on a polyspecific nematode community and yield of tomato. Egyptian Journal of Agronematology. 2001;5:79-89.
194 - Wei LH, Xue QY, Wei BQ, Wang YM, Li SM, Chen LF, Guo JH. Screening of antagonisticbacterial strains against Meloidogyne incognita using protease activity. Biocontrol Science and Technology. 2010;20:739-750.
195 - Abd‐Elgawad MMM, Mohamed MMM, El‐Gamal NGS. Development of safe chemicaland biological formulations for control of nematodes in cucumber. Egyptian Pharmaceutical Journal. 2008;7:41-50.