Submitted Successfully!
To reward your contribution, here is a gift for you: A free trial for our video production service.
Thank you for your contribution! You can also upload a video entry or images related to this topic.
Version Summary Created by Modification Content Size Created at Operation
1
Mati Ur Rahman
 -- 2177 2023-11-08 13:03:28 |
2 format correct
Wendy Huang
 Meta information modification 2177 2023-11-09 03:03:26 |

Video Upload Options

Do you have a full video?
Send video materials Upload full video

Confirm

Are you sure to Delete?
Yes No
Cite
If you have any further questions, please contact Encyclopedia Editorial Office.
Rahman, M.U.; Chen, P.; Zhang, X.; Fan, B. Nematophagous Fungi. Encyclopedia. Available online: https://encyclopedia.pub/entry/51293 (accessed on 03 May 2024).
Rahman MU, Chen P, Zhang X, Fan B. Nematophagous Fungi. Encyclopedia. Available at: https://encyclopedia.pub/entry/51293. Accessed May 03, 2024.
Rahman, Mati Ur, Peng Chen, Xiuyu Zhang, Ben Fan. "Nematophagous Fungi" Encyclopedia, https://encyclopedia.pub/entry/51293 (accessed May 03, 2024).
Rahman, M.U., Chen, P., Zhang, X., & Fan, B. (2023, November 08). Nematophagous Fungi. In Encyclopedia. https://encyclopedia.pub/entry/51293
Rahman, Mati Ur, et al. "Nematophagous Fungi." Encyclopedia. Web. 08 November, 2023.
Nematophagous Fungi
Edit
This entry is adapted from the peer-reviewed paper 10.3390/agronomy13112685

Plant-parasitic nematodes significantly threaten agriculture and forestry, causing various diseases. They cause annual losses of up to 178 billion dollars worldwide due to their parasitism. Nematophagous fungi (NF) are valuable in controlling or reducing parasitic nematode diseases by killing nematodes through predatory behavior.

nematophagous fungi nematodes predatory fungi egg- and female-parasitic fungi endoparasitic fungi toxins

1. Introduction

Nematodes are highly ubiquitous animal groups found throughout the planet. Despite being over 28,000 known species, they are challenging to differentiate. Out of all these species, 16,000 are recognized as parasitic towards animals, insects, or plants [1]. Over 4000 species of plant-parasitic nematodes (PPNs) have been identified [2] which severely threaten agricultural and horticultural crops [3]. These harmful pathogens are hideous crop parasites that cause global annual losses between USD 78 and 125 billion [4][5]. PPNs can actively move in the soil region adjacent to the plant roots, known as the rhizosphere, as well as on the aerial parts of the plant, or within the plant itself. They feed and breed on living organisms, with some nematodes feeding on plant parts such as flowers, leaves, stems, and seeds [6]. However, most of these worms feed on underground plant parts like roots, bulbs, and tubers, causing substantial plant damage ranging from mild harm to destruction [5].
NF are capable of capturing, killing, and digesting nematodes [7]. They reside externally and internally within the host organism, exploiting it for sustenance. These fungi use specific traps to ensnare prey, hyphae tips to parasitize females and eggs, and conidia to adhere while generating toxins to attack nematodes. Based on these strategies, NF are traditionally classified into four groups (Figure 1): (1) The group of fungi that prey on nematodes using specialized traps; (2) some fungi are egg-parasitic and invade nematode eggs or females through their hyphal tips, (3) while others are endoparasitic and use their spores. (4) Additionally, toxin-producing fungi immobilize nematodes before invading them [8][9].
Agronomy 13 02685 g002
Figure 1. Mode of action of nematophagous fungi.

2. Predatory Fungi

Predatory fungi employ hyphal structures in nematodes. Studies on nematode-trapping fungi have drawn much attention to their diverse and intricate catching structures [10]. The traps generated by the fungi’s mycelium adversely affect the nematode’s cuticle. The mycelium proliferates within the nematode’s body, resulting in the formation of a penetration peg, whose growth over time causes the hyphae to cover the outer surface of the colonized nematode [11]. The fungus A. oligospora has a unique mechanism for penetrating the cuticle of nematodes via penetrating tubes, and its impact on Meloidogyne javanica in tomato cultivars has been empirically demonstrated [12]. Predatory structures are vital for the life and activity of trapping fungi. Compared to regular hyphae, adhesive traps have a longer lifespan [13][14]. These specialized traps are utilized by over two hundred fungal species (found within the Zygomycota, Basidiomycota, and Ascomycota phyla) to capture free-living nematodes in soil [15]. NF play a vital role in maintaining the population of nematodes through natural methods, such as parasitism, trapping, and poisoning [16][17]. Basidiomycota trapping fungi use spores and adhesive knobs to capture nematodes [18][19][20]. The Orbiliaceae family comprises over 80% of nematode-trapping fungi within the Ascomycota Phylum, where constricting rings, adhesive networks, adhesive branches, adhesive knobs, and non-constricting rings are all used to ensnare nematodes (Figure 1) [21][22]. The study of nematode-trapping fungi in Zygomycota has faced obstacles due to inadequate isolation and culture techniques, despite the growing interest in it [23].

2.1. Adhesive Branches

Adhesive branches, also known as adhesive columns, have a simpler morphological structure than other capture organs (Figure 1). These vertical branches consist of one to three cells that merge via anastomosis, forming adhesive hoops or networks with two-dimensional structures resembling crochet or lines. The nematode is easily captured upon contact with the branch due to its complete coverage by a delicate adhesive layer. Due to the proximity of adhesive branches, nematodes frequently become stuck to more sticky hyphae upon contact and struggle to detach themselves. The species commonly found in temperate soils with developed adhesive branches is Dactylella cionopaga [24]. These branches serve as typical trapping mechanisms for Monacrosporium cionopagum and M. gephyrophagum [25]. For example, M. cionopagum produces adhesive branches that trap and immobilize the sugar beet cyst nematode Heterodera schachtii [26]. Likewise, Gamsylella gephyropaga produces adhesive branches to trap nematodes [27][28].

2.2. Adhesive Hyphal Network

The adhesive network, widely distributed in fungi, is comprised of an upright lateral branch emerging from a vegetative hypha (Figure 1), extending approximately 20–25 μm from the parent hypha [29], and is characterized by a longer lifespan in comparison to typical hyphae [13]. These adhesive nets are constructed using intricate three-dimensional networks. A. oligospora, with a global distribution, is the most frequently observed species in this specific trapping structure [30]. Adhesive nets form from vegetative hyphae by curving a solitary lateral branch and can combine with parental hyphae. Adhesive nets are regarded as an evolutionary progression from adhesive branches. More lateral hyphae are generated from the parental hyphae, or a loop is formed to generate additional loops once a complex of interconnected loops that extend away from the potential hyphae in all logical directions is established. Nematodes are attracted to the network’s surface, which is coated with a thin layer of adhesives [31].

2.3. Adhesive Knobs

Adhesive knobs are specialized cells with a small layer of adhesive covering them (Figure 1). When a nematode becomes ensnared, the contact area between it and the spherical knob is limited, allowing it to resist and free itself. However, upon coming into contact with a flattened, sticky pad, the fungus takes control and traps the nematode. This significantly increases the adhering surface, resulting in a secure binding of the captured nematode, followed by fungal penetration that involves both enzymatic and physical mechanisms. For example, the fungus synthesis of collagenase aids in penetrating the nematode’s cuticle, while the dense sticky pad provides strength and rigidity, allowing the piercing hyphae to move toward the cuticle [24]. Once a spherical infection bulb has formed, assimilative hyphae emerge to consume the internal contents of the nematode [32][33]. Dactylellina arcuata, D. asthenopaga, D. leptosphora, D. copepodii, and D. ellipsospora use adhesive knobs to capture nematodes [34][35][36].

2.4. Constricting Rings

Constricting rings are hyphal branches with a circular arrangement typically composed of three cells (Figure 2). These structures are highly sophisticated and actively capture prey. A nematode entering the cavity triggers the rapid expansion of the three surrounding cells, resulting in a threefold increase in their volume. This process effectively seals the orifice and confines the nematode inside the cavity. Subsequently, the hyphae penetrate and assimilate the nematode [37]. Twelve species of hyphomycetes have been discovered to form constricting rings of varying internal diameters ranging from 20 to 40 μm [24]. Constricting rings are distinct from other mechanisms because they encircle nematodes upon contact with the inner edge of the trap, resulting in closure. The nematode is strangled by the expansion of an internally located cell wall of a constricting ring, causing inward swelling through the outer cell wall of these rings. Rapid water intake causes an increase in the volume of the cells forming the ring [38]. The traps of D. brochopaga mutants are considerably larger, almost eight times more than conventional traps. It has been observed that the cells in these traps release fluid droplets, reducing their volume. The humidity of the surrounding environment can be adjusted to facilitate the entry or exit of atmospheric water into or out of these cells. Additionally, a correlation has been established between the ambient humidity level and the frequency of ring closures [39]. The source of the water supply has been investigated, and evidence suggests that it mainly originates primarily from stalk cells or mycelium [40]. Furthermore, once the stalk cell closed, there was no noticeable movement of internal components, suggesting that water from the surrounding environment may have been retained [41]. This is supported by the observation that rings can continue to spread even after being detached from the original stalk on which they first appeared [39][42]. This idea is plausible as live nematodes are typically surrounded by a thin layer of water, which could serve as a sufficient source of moisture for the process of ring closure. Additionally, the ring closure process in D. brochopaga can be chemically induced, in addition to physical methods like touch, elevated temperature, or electrical stimulation [43]. When exposed to solutions containing methanol, ethanol, propanol, or butanol, or to chlorobutanol vapor, this fungus’s traps expanded in 10 to 15 s. In contrast, it is important to note that benzene, ether, and chloroform did not have any detectable effect, suggesting that unidentified variables drive this significant phenomenon [44].

2.5. Non-Constricting Rings

Non-constricting rings are a type of three-celled rings that grow on a short supportive stalk originating from prostrate septate hyphae (Figure 2D). The nematodes display passive behavioral responses during the predation process. As noted, the attachment point between the supporting stalk and the ring was weakened. As the nematode attempted to free itself, the ring often detached, indicating that the fungus may have facilitated its escape by allowing it to carry the non-constricting ring tightly wrapped around its body. This seems to be a favored method for achieving widespread dissemination in the soil [45]. For example, fungi, such as Dactylaria candida and D. lysipaga, which produce non-constricting rings, often create adhesive knobs [34][46] and capture nematodes using non-constricting rings [35]. A similar pattern was observed in Dactylellina daliensis via non-constricting rings [47].
Agronomy 13 02685 g003
Figure 2. Different morphological structures used by predatory fungi for capturing nematodes. (A) Adhesive branches (adhesive column [48]), (B) adhesive hyphae network [49], (C) adhesive knob [50], (D) constricting ring, and (E) non-constricting ring [51].

3. Egg- and Female-Parasitic Fungi

Research on egg- and female-parasitic fungi has been in progress since the 1990s. These fungi employ appressoria (Purpureocillium spp. and Pochonia spp.), zoospores (Nematophthora gynophila), lateral mycelial branches, and penetration pegs to parasitize eggs, females, and other growth stages of the PPNs [52]. An in vitro assessment was conducted to evaluate the parasitism of 10 isolates of P. chlamydosporia on Globodera pallida eggs; the levels of observed pathogenicity ranged from 34% to 49%. The event of impulsive hatching occurs when P. chlamydosporia isolates aggressively parasitize immature eggs as opposed to those containing second-stage juveniles [53]. Additionally, the use of wild-type Beauveria bassiana 08F04 and transformant G10 resulted in a substantial decrease in the cereal cyst nematode (female) population in the roots [54]. In a greenhouse study, it was found that the presence of the arbuscular mycorrhizal fungus (AMF) Glomus etunicatum reduced the population of H. glycines female nematodes by 28.21% in root systems compared to untreated roots. This finding suggests that G. etunicatum may play a role in promoting the ability of host plants to tolerate the presence of the soybean cyst nematode (SCN) [55]. Figure 3 displays fungal species that parasitize the egg and female.
Agronomy 13 02685 g004
Figure 3. The nematode egg- and female-parasitic fungi and their infection modes.

4. Endoparasitic Fungi

Endoparasitic fungi are a category of nematophagous fungi which infect nematodes by producing spores. These spores can either be internalized by the nematodes through ingestion, leading to infection, or attach to the nematode epidermis, initiating the infection [8][56][57]. Endoparasites utilize spores, such as conidia and zoospores, for infection, which may attach to the nematode cuticle or be ingested [58]. It has been found that endoparasitic fungi can reduce the number of root-knot nematodes that create galls on tomatoes and alfalfa in greenhouse experiments [8]. These fungi exhibit varying degrees of diversity, with studies indicating differences in their production of conidia per infected nematode. D. coniospora fungi produce a significant amount of conidia, with up to 10,000 per hyphal material, while H. rhossoliensis yields 100–1000 conidia per infected nematode. Conidia germinate immediately, and assimilative hyphae infiltrate and absorb the entire contents of the nematode body, enabling the fungus to penetrate the host’s outer layer [59]. D. coniospora is an aggressive endoparasitic fungus that targets nematodes. The endoparasitic fungus Drechmeria coniospora YMF1.01759 strain exhibited excellent nematode-infecting ability. The study revealed that it hindered nematodes from hatching their eggs, infected them with spores, and produced active metabolites that killed them [60].

5. Toxin Production

Some nematophagous fungi produce toxins that kill nematodes and impact plant defense and resistance mechanisms against parasitic nematodes [61][62][63]. Toxin-producing fungi originate from various orders and families. The fungus assaults nematodes via the secretion of inhibitory metabolites without physically interacting and immobilized them [52][64]. After immobilization, the hyphae penetrate the nematode cuticle. Culture filtrates of these fungi contain strong enzymatic (proteolytic and chitinolytic) activities, low-molecular-weight metabolites, and specific non-volatile oil components that cause larval death or inhibit egg hatching [65]. The metabolites secreted by the fungi alter the composition of nematode eggs and prevent embryonic development, rendering them unable to hatch due to their varying shapes and sizes. Similarly, fungi produce toxic chemicals, other than enzymes, that immobilize nematodes and later consume them [66]. Basidiomycetes are the predominant fungi that produce toxins. Recent research on Basidiomycetous fungi (Coprinus comatus and Stropharia rugosoannulata) has revealed that the action mechanisms of these toxins against nematodes are varied and multifaceted [67]. Among Basidiomycetes, numerous Pleurotus species produce toxins with nematotoxic activity [66][68]. For example, P. ostreatus produces trans-2-decenoic acid, a compound obtained from linoleic acid that is detrimental to nematodes, insects, and other fungi [69]. Basidiomycetes are not the only fungi that generate these kinds of toxins; some fungi also produce toxins that are harmful to nematodes, but these are not nematophagous [11]. These compounds exhibit diverse chemical properties, including simple fatty acids or other organic acids such as lactones, pyrones, anthraquinones, benzoquinones, alkaloids, furans, peptaibiotics, and cyclodepsipeptides.

References

  1. Hugot, J.-P.; Baujard, P.; Morand, S. Biodiversity in helminths and nematodes as a field of study: An overview. Nematology 2001, 3, 199–208.
  2. Jones, J.T.; Haegeman, A.; Danchin, E.G.; Gaur, H.S.; Helder, J.; Jones, M.G.; Kikuchi, T.; Manzanilla-López, R.; Palomares-Rius, J.E.; Wesemael, W.M. Top 10 plant-parasitic nematodes in molecular plant pathology. Mol. Plant Pathol. 2013, 14, 946–961.
  3. Elling, A.A. Major emerging problems with minor meloidogyne species. Phytopathology 2013, 103, 1092–1102.
  4. Batish, D.R.; Singh, H.P.; Kohli, R.K.; Kaur, S. Eucalyptus essential oil as a natural pesticide. For. Ecol. Manag. 2008, 256, 2166–2174.
  5. Chitwood, D.J. Research on plant-parasitic nematode biology conducted by the United States Department of Agriculture–Agricultural Research Service. Pest Manag. Sci. Former. Pestic. Sci. 2003, 59, 748–753.
  6. Palomares-Rius, J.E.; Escobar, C.; Cabrera, J.; Vovlas, A.; Castillo, P. Anatomical alterations in plant tissues induced by plant-parasitic nematodes. Front. Plant Sci. 2017, 8, 1987.
  7. Nordbring-Hertz, B.; Jansson, H.-B.; Tunlid, A. Nematophagous Fungi. In eLS; Wiley Online Library: Hoboken, NJ, USA, 2011.
  8. Liu, X.; Xiang, M.; Che, Y. The living strategy of nematophagous fungi. Mycoscience 2009, 50, 20–25.
  9. Abd-Elgawad, M.M.; Askary, T.H. Fungal and bacterial nematicides in integrated nematode management strategies. Egypt. J. Biol. Pest Control 2018, 28, 74.
  10. Corda, A.K.J. Pracht-Flora Europaeischer, Schimmelbildungen; Bei G. Fleischer: Leipzig, Germany, 1839.
  11. de Freitas Soares, F.E.; Sufiate, B.L.; de Queiroz, J.H. Nematophagous fungi: Far beyond the endoparasite, predator and ovicidal groups. Agric. Nat. Resour. 2018, 52, 1–8.
  12. Mostafanezhad, H.; Sahebani, N.; Nourinejhad Zarghani, S. Induction of resistance in tomato against root-knot nematode Meloidogyne javanica with salicylic acid. J. Crop Prot. 2014, 3, 499–508.
  13. Veenhuis, M.; Nordbring-Hertz, B.; Harder, W. An electron-microscopical analysis of capture and initial stages of penetration of nematodes byArthrobotrys oligospora. Antonie Van Leeuwenhoek 1985, 51, 385–398.
  14. Bedekovic, T.; Brand, A.C. Microfabrication and its use in investigating fungal biology. Mol. Microbiol. 2022, 117, 569–577.
  15. Li, T.; Zhang, K.; Liu, X. Taxonomy of Nematophagous Fungi; China Science Publishing & Media Ltd: Beijing, China, 2000.
  16. Jaffee, B.; Tedford, E.; Muldoon, A. Tests for density-dependent parasitism of nematodes by nematode-trapping and endoparasitic fungi. Biol. Control 1993, 3, 329–336.
  17. Linford, M.; Yap, F.; OLIVEIRA, J.M. Reduction of soil populations of the root-knot nematode during decomposition of organic matter. Soil Sci. 1938, 45, 127–142.
  18. Poloczek, E.; Webster, J. Conidial traps in Nematoctonus (nematophagous Basidiomycetes). Nova Hedwig. 1994, 59, 201–205.
  19. Dürschner-Pelz, U. Traps of Nematoctonus leiosporus—An unusual feature of an endoparasitic nematophagous fungus. Trans. Br. Mycol. Soc. 1987, 88, 129–130.
  20. Thorn, R.G.; Moncalvo, J.-M.; Reddy, C.; Vilgalys, R. Phylogenetic analyses and the distribution of nematophagy support a monophyletic Pleurotaceae within the polyphyletic pleurotoid-lentinoid fungi. Mycologia 2000, 92, 241–252.
  21. Hyde, K.D.; Swe, A.; Zhang, K.-Q. Nematode-trapping fungi. In Nematode-Trapping Fungi; Springer: Berlin/Heidelberg, Germany, 2014; pp. 1–12.
  22. Swe, A.; Li, J.; Zhang, K.; Pointing, S.; Jeewon, R.; Hyde, K. Nematode-trapping fungi. Curr. Res. Environ. Appl. Mycol. 2011, 1, 1–26.
  23. Saikawa, M. Ultrastructural studies on zygomycotan fungi in the Zoopagaceae and Cochlonemataceae. Mycoscience 2011, 52, 83–90.
  24. Poinar, G.O.; Jansson, H.-B. Diseases of Nematodes; CRC Press: Boca Raton, FL, USA, 1988; Volume 1.
  25. Saxena, G. Biological control of root-knot and cyst nematodes using nematophagous fungi. In Root Biology; Springer: Berlin/Heidelberg, Germany, 2018; pp. 221–237.
  26. Andersson, K.-M.; Kumar, D.; Bentzer, J.; Friman, E.; Ahrén, D.; Tunlid, A. Interspecific and host-related gene expression patterns in nematode-trapping fungi. Bmc Genom. 2014, 15, 968.
  27. Jaffee, B.; Strong, D. Strong bottom-up and weak top-down effects in soil: Nematode-parasitized insects and nematode-trapping fungi. Soil Biol. Biochem. 2005, 37, 1011–1021.
  28. Zhang, Y.; Zhang, K.-Q.; Hyde, K. The ecology of nematophagous fungi in natural environments. In Nematode-Trapping Fungi; Springer: Berlin/Heidelberg, Germany, 2014; pp. 211–229.
  29. Nordbring-Hertz, B.; Friman, E.; Veenhuis, M. Hyphal fusion during initial stages of trap formation in Arthrobotrys oligospora. Antonie Van Leeuwenhoek 1989, 55, 237–244.
  30. Niu, X.-M.; Zhang, K.-Q. Arthrobotrys oligospora: A model organism for understanding the interaction between fungi and nematodes. Mycology 2011, 2, 59–78.
  31. Pala Martinelli, P.R.; dos Santos, J.M. Scanning electron microscopy of nematophagous fungi associated Tylenchulus semipenetrans and Pratylenchus jaehni. Biosci. J. 2010, 26, 809–816.
  32. Barron, G.L. The Nematode-Destroying Fungi; Canadian Biological Publications Ltd.: Guelph, ON, Canada, 1977.
  33. Gray, N. Fungi attacking vermiform nematodes. Dis. Nematodes 1988, 2, 3–38.
  34. Li, Y.; Hyde, K.D.; Jeewon, R.; Cai, L.; Vijaykrishna, D.; Zhang, K. Phylogenetics and evolution of nematode-trapping fungi (Orbiliales) estimated from nuclear and protein coding genes. Mycologia 2005, 97, 1034–1046.
  35. Hai-Yan, W.; Dong-Geun, K.; Xun-Bo, Z. First report of an unrecorded nematode-trapping fungus species Dactylellina candidum in Korea. Afr. J. Microbiol. Res. 2012, 6, 203–205.
  36. Xie, H.; Aminuzzaman, F.; Xu, L.; Lai, Y.; Li, F.; Liu, X. Trap induction and trapping in eight nematode-trapping fungi (Orbiliaceae) as affected by juvenile stage of Caenorhabditis elegans. Mycopathologia 2010, 169, 467–473.
  37. Liu, K.; Tian, J.; Xiang, M.; Liu, X. How carnivorous fungi use three-celled constricting rings to trap nematodes. Protein Cell 2012, 3, 325–328.
  38. Heintz, C.; Pramer, D. Ultrastructure of nematode-trapping fungi. J. Bacteriol. 1972, 110, 1163–1170.
  39. Insell, J.; Zachariah, K. The mechanism of the ring trap of the predacious hyphomycete Dactylella brochopaga Drechsler. Protoplasma 1978, 95, 175–191.
  40. Rudek, W.T. The constriction of the trapping rings in Dactylaria brochopaga. Mycopathologia 1975, 55, 193–197.
  41. Barron, G. Nematophagous fungi: A new Harposporium producing aerial arthroconidia. Can. J. Bot. 1979, 57, 886–889.
  42. Barron, G. Host range studies for Haptoglossa and a new species, Haptoglossa intermedia. Can. J. Bot. 1989, 67, 1645–1648.
  43. Higgins, M.; Pramer, D. Fungal morphogenesis: Ring formation and closure by Arthrobotrys dactyloides. Science 1967, 155, 345–346.
  44. Zachariah, K. Chemical induction of trap closure in Dactylella brochopaga. Protoplasma 1989, 148, 87–93.
  45. Askary, T.H.; Martinelli, P.R.P. Biocontrol Agents of Phytonematodes; CABI: Egham, UK, 2015.
  46. Drechsler, C. Some hyphomycetes that prey on free-living terricolous nematodes. Mycologia 1937, 29, 447–552.
  47. Su, H.; Hao, Y.e.; Yang, X.; Yu, Z.; Deng, J.; Mo, M. A new species of Dactylellina producing adhesive knobs and non-constricting rings to capture nematodes. Mycotaxon 2008, 105, 313.
  48. Jiang, X.; Xiang, M.; Liu, X. Nematode-Trapping Fungi. Microbiol. Spectr. 2017, 5, 5-1.
  49. Nordbring-Hertz, B. Morphogenesis in the nematode-trapping fungus Arthrobotrys oligospora—An extensive plasticity of infection structures. Mycologist 2004, 18, 125–133.
  50. Li, J.; Zou, C.; Xu, J.; Ji, X.; Niu, X.; Yang, J.; Huang, X.; Zhang, K.-Q. Molecular mechanisms of nematode-nematophagous microbe interactions: Basis for biological control of plant-parasitic nematodes. Annu. Rev. Phytopathol. 2015, 53, 67–95.
  51. Liu, K.; Zhang, W.; Lai, Y.; Xiang, M.; Wang, X.; Zhang, X.; Liu, X. Drechslerella stenobrocha genome illustrates the mechanism of constricting rings and the origin of nematode predation in fungi. BMC Genom. 2014, 15, 114.
  52. Lopez-Llorca, L.; Maciá-Vicente, J.; Jansson, H.-B. Mode of action and interactions of nematophagous fungi. In Integrated Management and Biocontrol of Vegetable and Grain Crops Nematodes; Springer: Berlin/Heidelberg, Germany, 2008; pp. 51–76.
  53. Dos Santos, M.C.V.; Horta, J.; Moura, L.; Pires, D.V.; Conceicao, I.; Abrantes, I.; Costa, S.R. An integrative approach for the selection of Pochonia chlamydosporia isolates for biocontrol of potato cyst and root knot nematodes. Phytopathol. Mediterr. 2019, 58, 187–199.
  54. Zhang, J.; Fu, B.; Lin, Q.; Riley, I.T.; Ding, S.; Chen, L.; Cui, J.; Yang, L.; Li, H. Colonization of Beauveria bassiana 08F04 in root-zone soil and its biocontrol of cereal cyst nematode (Heterodera filipjevi). PLoS ONE 2020, 15, e0232770.
  55. Benedetti, T.; Antoniolli, Z.I.; Sordi, E.; Carvalho, I.R.; Bortoluzzi, E.C. Use of the Glomus etunicatum as biocontrol agent of the soybean cyst nematode. Res. Soc. Dev. 2021, 10, e7310615132.
  56. Dijksterhuis, J.; Harder, W.; Wyss, U.; Veenhuis, M. Colonization and digestion of nematodes by the endoparasitic nematophagous fungus Drechmeria coniospora. Mycol. Res. 1991, 95, 873–878.
  57. Tunlid, A.; Jansson, H.-B.; Nordbring-Hertz, B. Fungal attachment to nematodes. Mycol. Res. 1992, 96, 401–412.
  58. Braga, F.R.; de Araújo, J.V. Nematophagous fungi for biological control of gastrointestinal nematodes in domestic animals. Appl. Microbiol. Biotechnol. 2014, 98, 71–82.
  59. Nordbring-Hertz, B.; Jansson, H.; Tunlid, A. Encyclopedia of Life Sciences; Wiley: Chichester, UK, 2006.
  60. Wan, J.; Dai, Z.; Zhang, K.; Li, G.; Zhao, P. Pathogenicity and metabolites of endoparasitic nematophagous fungus Drechmeria coniospora YMF1.01759 against nematodes. Microorganisms 2021, 9, 1735.
  61. Sarker, M.S.; Mohiuddin, K.; Al-Ani, L.K.T.; Hassan, M.N.; Akter, R.; Hossain, M.S.; Khand, M. Effect of bio-nematicide and bau-biofungicide against root-knot (Meloidogyne spp.) of soybean. Malays. J. Sustain. Agric. 2020, 4, 44.
  62. Comans-Pérez, R.J.; Sánchez, J.E.; Al-Ani, L.K.T.; González-Cortázar, M.; Castañeda-Ramírez, G.S.; Mendoza-de Gives, P.; Sánchez-García, A.D.; Millán-Orozco, J.; Aguilar-Marcelino, L. Biological control of sheep nematode Haemonchus contortus using edible mushrooms. Biol. Control 2021, 152, 104420.
  63. Girardi, N.S.; Sosa, A.L.; Etcheverry, M.G.; Passone, M.A. In vitro characterization bioassays of the nematophagous fungus Purpureocillium lilacinum: Evaluation on growth, extracellular enzymes, mycotoxins and survival in the surrounding agroecosystem of tomato. Fungal Biol. 2022, 126, 300–307.
  64. Liu, X.; Li, S. Fungal secondary metabolites in biological control of crop pests. In Handbook of Industrial Mycology; CRC Press: Boca Raton, FL, USA, 2004; pp. 742–767.
  65. Westphal, A.; Becker, J. Components of soil suppressiveness against Heterodera schachtii. Soil Biol. Biochem. 2001, 33, 9–16.
  66. Satou, T.; Kaneko, K.; Li, W.; Koike, K. The toxin produced by Pleurotus ostreatus reduces the head size of nematodes. Biol. Pharm. Bull. 2008, 31, 574–576.
  67. Luo, H.; Li, X.; Li, G.; Pan, Y.; Zhang, K. Acanthocytes of Stropharia rugosoannulata function as a nematode-attacking device. Appl. Environ. Microbiol. 2006, 72, 2982–2987.
  68. Nordbring-Hertz, B.; Neumeister, H.; Sjollema, K.; Veenhuis, M. A conidial trap-forming mutant of Arthrobotrys oligospora. Mycol. Res. 1995, 99, 1395–1398.
  69. Kwok, O.; Plattner, R.; Weisleder, D.; Wicklow, D. A nematicidal toxin from Pleurotus ostreatus NRRL 3526. J. Chem. Ecol. 1992, 18, 127–136.
More
© Text is available under the terms and conditions of the Creative Commons Attribution (CC BY) license; additional terms may apply. By using this site, you agree to the Terms and Conditions and Privacy Policy.
Upload a video for this entry
Information
Subjects: Others
Contributors MDPI registered users' name will be linked to their SciProfiles pages. To register with us, please refer to https://encyclopedia.pub/register :
Mati Ur Rahman
,
Peng Chen
,
Xiuyu Zhang
,
Ben Fan
View Times: 289
Revisions: 2 times (View History)
Update Date: 09 Nov 2023
Table of Contents
    1000/1000
    Hot Most Recent
    Feedback