Thielaviopsis basicola

Thielaviopsis basicola is a plant-pathogenic fungus in the division Ascomycota. It is a soil-borne fungus that causes black root rot.[1] It has a wide host range consisting of gerbera, kalanchoe, pansy, petunia, poinsettia, primula, snapdragon, sweet pea, verbena, and viola.[2] After T. basicola infects the host some of the symptoms consist of “stunting of foliage and root systems, blackened area on roots, yellowing of leaves between the veins or along the margins, and branch dieback. The yellowing of leaves means the plant cannot do photosynthesis and the blackened tissue means it is dead. And some of the signs include dark brown, multi-celled spores form in the infected roots. The individual cells appear to snap apart. Light colored spores are formed in a long tapering cell and extruded in chains”.[1] If the hypocotyl along with the roots become infected it can lead to “black, rough, longitudinal cracks.”.[2]

Thielaviopsis basicola
Thielaviopsis basicola 1.JPG
Microscopic view of Thielaviopsis chlamydospores (black) and endoconidia (hyaline)
Scientific classification edit
Kingdom: Fungi
Division: Ascomycota
Class: Sordariomycetes
Order: Microascales
Family: Ceratocystidaceae
Genus: Thielaviopsis
Species:
T. basicola
Binomial name
Thielaviopsis basicola
(Berk. & Broome) Ferraris (1912)
Synonyms

Chalara elegans Nag Raj & W.B. Kendr. (1975)
Torula basicola Berk. & Broome (1850)
Trichocladium basicola (Berk. & Broome) J.W. Carmich. (1980)

PathogenesisEdit

Thielaviopsis basicola is a soilborne fungus that belongs to the Ascomycota division of the "true fungi" and is a hemibiotrophic parasite.[3] Fungi belonging to Ascomycota are known to produce asexual and sexual spores, however, a sexual stage has yet to be observed and validated in the Thielaviopsis basicola life cycle, which classifies this species as a Deuteromycete or an imperfect fungus.[4] During the asexual reproductive cycle of Thielaviopsis basicola, two types of asexual spores are borne from the hyphae including endoconidia and chlamydospores.[4] Endoconidia are a distinctive type of conidium in that they develop within a hollow cavity inside a hyphal tube and are ejected from the end of this tube to disperse.[5] Both of the aforementioned spores must first undergo physical dissemination in order to begin locating an infection court on a new, viable host. Aside from the normal translocation of spores within the soil environment, vectors such as shore flies have been observed carrying and aerially transmitting Thielaviopsis basicola spores, a phenomenon uncharacteristic of soilborne fungal pathogens.[6] Upon landing on an infected plant, the shore flies feed on the infected tissue and ingest spores along with the plant material, only to excrete the hitchhiking spores in their frass, which ultimately lands on healthy plant tissue continuing the disease cycle.[6] However, it is important to note that this association between vector and soilborne fungi has only been observed in commercial agricultural settings in which artificially controlled environments (i.e. greenhouses) promote conditions that deviate from the natural world.[6]


Following dispersal (via vector-insect, cultural practice, or other translocation means within the soil matrix), the spores will detect an infection site on the host plant (usually root hairs) and germinate in response to the stimuli produced by the root exudates, some of which include sugars, lecithins, and unsaturated triglycerides.[7] Germ tubes emerge from the spores and directly penetrate into the cells of the root hairs (typically the single-cell epidermal layer) via penetration hyphae.[7] The living host plant will typically respond with the development of cell appositions called papillae, which attempt to block the pathogen from penetrating the cell wall and subsequently parasitizing the host's cells.[8] However, most of these early defense mechanisms prove unsuccessful, hence the significance and prevalence of the disease around the world. Advancing, the vegetative hyphal cells differentiate into feeding structures that resemble haustoria, which absorb nutrients biotrophically from the host cells.[9] Once the pathogen has breached the cell wall of the epidermal root cell, it proceeds to release effector compounds that disrupt the host's systemic defense mechanisms.[10] Systemic acquired resistance (SAR) is employed by the host to actively address localized infection and initiate defense signaling cascades throughout the plant. For example, the SAR NPR1 (AtNPR1) gene is of special importance and acts to suppress the infection faculties of Thielaviopsis basicola, effectively imparting resistance to some host plants.[10] Furthermore, research suggests that the NPR1 gene, when over-expressed in transgenic plants, aids in the expression of other defense-related genes such as PR1, effectively improving resistance to infection by Thielaviopsis basicola.[10] NPR1 and its associated benefits for enhancing disease resistance have been recognized as possible tools to use when equipping economically indispensable crops with transgenic resistance to disease.[10]


Once penetration and the establishment of biotrophic feeding structures are successful, the pathogen progresses into the root tissue leaving distinctive black/brown lesions in its wake (lesion coloration can be attributed to thick-walled chlamydospore clusters); it continues proliferating until eventually entering its necrotrophic stage.[4] Hemibiotrophs, like Thielaviopsis basicola, transition from a biotrophic stage to a necrotrophic stage by way of a coordinated effort between different pathogenesis genes that secrete effector proteins capable of manipulating their host's defense system.[11] Research suggests that during biotrophy, certain types of effectors from the pathogen are expressed over others and vice versa during the necrotrophic stage.[11] Once the biotrophic stage is no longer preferred by the pathogen, it will initiate this complicated genetic transition and commence the necrotrophic stage. In order to digest and metabolize nutritive compounds from a necrotic host plant, Thielaviopsis basicola secretes enzymes such as xylanase and other hemicellulases, which breakdown cell tissues making them available to the fungus.[12] During this stage, the pathogen also produces its asexual spores in the lesions to reproduce and disseminate more propagules for continued survival in the soil.[4] In addition to its normal infection process, studies have shown that Thielaviopsis basicola and it's pathogenesis are synergistically linked to a fortuitous coinfection process involving Meloidogyne incognita nematodes when the two are present in the same soil.[13] It has been observed that the infection of host tissues by Meloidogyne incognita facilitates the infection of Thielaviopsis basicola into the root and vascular tissues, effectively allowing the fungal pathogen to optimize infection even when environmental conditions are suboptimal.[13]

ImportanceEdit

Thielaviopsis basicola was discovered in the mid-1800s and has remained an important plant pathogen affecting ornamental and agricultural plants in over 31 countries around the world.[4] The pathogen is known to stunt or delay maturity in the species it parasitizes, which, coupled with environmental limitations, can lead to severe economic losses.[14] It has been observed that black root rot can delay plant maturity for up to a month and result in over a 40% yield reduction in the affected crop.[14] One crop that is affected by Thielaviopsis basicola and that is of significant economic importance is cotton. In the United States alone, between the years 1995 and 2005, the total annual loss in revenue due to diseases in the cotton crop was $897 million.[15] Thielaviopsis basicola was a significant contributor to that economic loss. In other parts of the world, such as in major cotton producer Australia, Thielaviopsis basicola has a very severe economic impact as well. In Australia, the disease was initially observed in sweet peas in the 1930s.[7] However, black root rot spread to a range of cultivated hosts, especially into Australian cotton production. In fact, surveys taken in 2010 and 2011 of Australian agriculture statistics reported black root rot to be present in 93% of farms and 83% of fields studied.[7] Of the fields affected, yield losses have reached 1.5 bales per acre.[7] The national average of cotton production per hectare in Australia is about 10 bales, so a loss of 1.5 bales per acre (or roughly 3 bales per hectare) to black root rot adds up to a significant loss.[16] In addition to cotton, carrot, lupin, cabbage, clover, and tobacco are all crops cultivated in many different countries that suffer from black root rot.[17] Some important ornamental crops affected by black root rot include: Begonia sp., poinsettia, African daisy, pansy, marigold, and petunia; the list is quite extensive.[18] However, cultural practices have led to the eradication of this disease in many ornamental crops, including poinsettia. During the 1950s and 1960s, poinsettia production was ravaged by black root rot disease.[19] Despite faltering, once the use of soil mixes was traded for soilless alternatives throughout the floriculture industry, black root rot was no longer a threat to poinsettias.[19] Thielaviopsis basicola (black root rot) has been and will remain a significant threat to crops grown globally in both agricultural and horticultural systems.

Disease cycleEdit

Thielaviopsis basicola is a soil inhabiting disease. The pathogen typically colonizes root tissue in the first two to eight weeks of crop growth. This causes cortical cell death which gives a brown to blackened appearance in the roots. The death of root cells also reduces development of new tissue in both roots and shoots. Once the fungus has successfully infected, it grows vegetatively via hyphae and produces the two types of spores.[20] In this particular situation, state means imperfect form of the fungi. The “chalara state produces endospores (conidia) and the Thielavopsis produces aleuriospores (chlamydospores). Chlamydospores survive in soil for many years”.[21] During wet and cool soil the spores will germinate. It is most “severe from 55° to 61°F, while only a trace of disease develops at 86°F. Alkaline soil favors the disease, which can be prevented at pH 4.8 and greatly reduced at pH 5.5 or below”.[20] The fungus can “spread via vectors including- fungus gnats and shore flies, from infected roots to healthy roots if they come into contact with each other and when spores (conidia) are splashed from pot to pot when watered”.[22]

ManagementEdit

Cultural Practices and Mechanical MeasuresEdit

The first and foremost strategy for controlling T. basicola at the first sign of disease should be cultural control including- “maintaining a soil pH below 5.6, removing and destroying all diseased plants, using soil-less media, sterilizing equipment, keeping work areas clean, and controlling fungus gnats and shore flies. Fungus gnats and shore flies can be vectors; therefore, controlling these pests can lead to minimizing the spread of the fungus”.[21] In addition, “crop rotation is recommended for management of black root rot. Soil fumigants, such as chloropicrin, can be helpful for controlling seedbed and sterol inhibitors”.[23] Furthermore, “to avoid contamination of plants and potting media, greenhouse floors and walkways should be lightly misted with water to cut down on airborne dust transmission of T. basicola during cleaning operations”.[24] At the end of the “growing season, doing a thorough clean-up of the greenhouse can be beneficial because it reduces the possibility of the fungus surviving as a resistant chlamydospores on the soil floor and in wooden benches".[22]

Disease ResistanceEdit

Disease resistance can be naturally coded in the genome of the host itself and induced via natural or artificial means, artificially introduced via a number of transgenic or breeding measures, and/or mutually associated with beneficial microbes found within soil ecosystems. Most, if not all, vascular plants utilize a system of defense, which consists of PAMP-triggered immunity (PTI) and effector-triggered immunity (ETI).[25] Following localized infection and the influx of associated pathogen stimulants, the aforementioned immune system responses trigger systemic acquired resistance (SAR), which sets off a cascade of defense signaling throughout the plant to initiate defense strategies at distal locations targeted to attack any recognized foreign pathogens. However, even with these innate lines of defense, the pathogen often prevails. This calls for selective breeding, genetic manipulation, or other novel biological control methods. Assessing varieties/cultivars for disease resistance and breeding for selected resistance traits is an important management method utilized by growers and breeders in the fight against Thielaviopsis basicola.[26] Commercially available resistant species of plants, include select varieties of Japanese holly (among other species of holly) and woody plants such as boxwood and barberry.[26][27] However, in some important crops like cotton, no commercially viable cultivars have been bred with sufficient resistance against black root rot.[7] Interestingly, in Australia, researchers have identified diploid cotton species displaying marked resistance against black root rot, yet cross-breeding these traits into viable commercial crops has proven to be difficult.[7] Similarly, researchers in Poland have uncovered innate disease resistance in the germplasm of a wild-type relative of Nicotiana tabacum called Nicotiana glauca.[17] Moreover, disease resistance genes derived from Nicotiana debneyi (a relative of the previously mentioned tobacco species) have successfully been incorporated into tobacco varieties displaying resilience to multiple races of Thielaviopsis basicola.[17] That being said, selective variety breeding is not the only source of resistance to black root rot in modern plant pathology. Transgenic methods of disease management offer promising new avenues scientists can take to aid in adapting plants to increasingly virulent pathogens. One such mechanism includes the manipulation of the expression of the NPR1 gene in the host plant defense genome sequences.[25] By over-expressing NPR1 genes transgenically in host plants such as cotton, scientists were able to increase the induction of PR genes like PR1 and LIPOXYGENASE1, which led to enhanced resistance by improving yield and limiting stunting.[25] In addition to genetic tools, inventive plant pathologists are exploring other novel methods of control, which include beneficial microbes and biological control agents (BCAs), among many others. Symbiotic associations between arbuscular mycorrhizal fungi and plant roots are well-documented, yet scientists studying host-plant defense have discovered this association may be more arcane than previously thought. Some researchers suggest this association extends to the realm of disease resistance and defense.[28] This phenomenon was analyzed in research conducted by German scientists who studied the transcript expression of defense related genes in Petunia hybrida when they were exposed to Thielaviopsis basicola and also colonized by arbuscular mycorrhizal fungal networks in their rhizosphere.[28] They found that the arbuscular mycorrhiza (AM) symbiosis functioned as a first line of defense by antagonizing the pathogenic fungi before it could ever induce a defense response in the host itself.[28] Thus, it is not inconceivable that control measures involving biotic compliments, such as AM, may be used in the future to control for disease presence in agricultural fields without the use of deleterious chemicals and/or genetic meddling.

Infected plantsEdit

See:

ReferencesEdit

  1. ^ a b "Black Root Rot (Thielaviopsis) (Plant Diseases)". Plant Diseases (Penn State Extension). Retrieved 2016-12-08.
  2. ^ a b "UC IPM: UC Management Guidelines for Thielaviopsis Root Rot on Floriculture and Ornamental Nurseries". ipm.ucanr.edu. Retrieved 2016-12-08.
  3. ^ Coumans, Joëlle V. F., Moens, Pierre D. J., Polijak, Anne, Al-Jaaidi, Samiya, Pereg, Lily and Raferty, Mark J. 2010. Plant-extract-induced changes in the proteome of the soil-borne pathogenic fungus Thielaviopsis basicola. Proteomics 10:1573-1591. doi: https://doi.org/10.1002/pmic.200900301
  4. ^ a b c d e Nel, W. J., Duong, T. A., de Beer, Z. W. and Wingfield, M. J. 2019. Black root rot: a long known but little understood disease. Plant Pathology. doi: 10.1111/ppa.13011
  5. ^ Delvecchio, V. G., Corbaz, R., and Turian, G. 1969. An Ultrastructural Study of the Hyphae, Endoconidia and Chlamydospores of Thielaviopsis basicola. Journal of General Microbiology 58:23-27. doi: 10.1099/00221287-58-1-23
  6. ^ a b c Stanghellini, M.E., Rasmussen, S. L., and Kim, D. H. 1999. Aerial transmission of Thielaviopsis basicola, a pathogen of corn-salad, by adult shore flies. Phytopathology 89:476-479. doi: https://doi.org/10.1094/PHYTO.1999.89.6.476
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  9. ^ Hood, M. E. and Shew, H. D. 1997. Reassessment of the Role of Saprophytic Activity in the Ecology of Thielaviopsis basicola. Phytopathology 87:1214-1219. doi: https://doi.org/10.1094/PHYTO.1997.87.12.1214
  10. ^ a b c d Kumar, Vinod, Joshi, Sameer G., Bell, Alois A. and Rathore, Keerti S. 2012. Enhanced resistance against Thielaviopsis basicola in transgenic cotton plants expressing Arabidopsis NPR1 gene. Transgenic Research 22:359-368. doi: 10.1007/s11248-012-9652-9
  11. ^ a b Lee, S.J., and Rose, J. K. 2010. Mediation of the transition from biotrophy to necrotrophy in hemibiotrophic plant pathogens by secreted effector proteins. Plant Signaling & Behavior 5(6):769-772. https://doi.org/10/4161/psb.5.6.11778
  12. ^ Ghosh, V. K. and Deb, J. K. 1988. Production and characterization of xylanase from Thielaviopsis basicola. Applied Microbiology and Biotechnology 29:44-47. doi: https://doi.org/10.1007/BF00258349
  13. ^ a b Walker, N. R., Kirkpatrick, T. L. and Rothrock, C. S. 1999. Effect of Temperature on and Histopathology of the Interaction Between Meloidogyne incognita and Thielaviopsis basicola on Cotton. Phytopathology 89:613-617. doi: https://doi.org/10.1094/PHYTO.1999.89.8.613
  14. ^ a b Holman, Sharna. 2016. Black root rot: The research roundup. https://www.cottoninfo.com.au/sites/default/files/documents/BRR%20update%20%28long%29%20v2%20-%20Oct%202016.pdf
  15. ^ Niu, Chen, Lister, Harriet E., Nguyen, Bay, Wheeler, Terry A. and Wright, Robert J. 2008. Resistance to Thielaviopsis basicola in the cultivated A genome of cotton. Theoretical and Applied Genetics 117:1313-1323. doi: 10.1007/s00122-008-0865-5
  16. ^ Farrell, Roger. 2018.  Australia: Cotton and Products Annual. USDA Foreign Agricultural Service: Global Agricultural Information Network. https://apps.fas.usda.gov/newgainapi/api/report/downloadreportbyfilename?filename=Cotton%20and%20Products%20Annual_Canberra_Australia_3-28-2018.pdf
  17. ^ a b c Trojak-Goluch, Anna and Berbec, Apoloniusz. 2005. Potential of Nicotiana glauca (Grah.) as a source of resistance to black root rot Thielaviopsis basicola (Ber. and Broome) Ferr. in tobacco improvement. Plant Breeding 124:507-510. doi: 10.1111/j.1439-0523.2005.01135.x
  18. ^ Greenhouse Plants, Ornamental-Black Root Rot. Pacific Northwest Pest Management Handbooks. Retrieved October 18, 2020, from https://pnwhandbooks.org/plantdisease/host-disease/greenhouse-plants-ornamental-black-root-rot
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  20. ^ a b Mondal, A. H.; Nehl, D. B.; Allen, S. J. (2005). "Acibenzolar-S-methyl induces systemic resistance in cotton against black root rot caused by Thielaviopsis basicola". Australasian Plant Pathology. 34 (4): 499–507. doi:10.1071/AP05089. ISSN 0815-3191. S2CID 37007553.
  21. ^ a b Pscheidt, J.W. "Black Root Rot: Thielaviopsis basicola" (PDF). Black Root Rot: Thielaviopsis basicola. Cornell University.
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  25. ^ a b c Silva, Katchen Julliany P., Mahna, Nasser, Mou, Zhonglin, and Folta, Kevin M. 2018.  NPR1 as a transgenic crop protection strategy in horticultural species. Horticulture Research 5:15. doi: 10.1038/s41438-018-0026-1
  26. ^ a b Lambe, R.C., and Ridings, W. H. 1979. Black Root Rot of Japanese Holly. Plant Pathology Circular. No. 204. https://www.fdacs.gov/content/download/11211/file/pp204.pdf
  27. ^ Hansen, Mary Ann. Black Root Rot of Japanese Holly. Virginia Cooperative Extension publication 450-606. https://vtechworks.lib.vt.edu/bitstream/handle/10919/48796/450-606_pdf.pdf?sequence=1&isAllowed=y
  28. ^ a b c Hayek, Soukayna, Gianinazzi-Pearson, Vivienne, Gianinazzi, Silvio, and Franken, Philipp. 2014. Elucidating mechanisms of mycorrhiza-induced resistance against Thielaviopsis basicola via targeted transcript analysis of Petunia hybrida genes. Physiological and Molecular Plant Pathology 88:67-76. doi: https://doi.org/10.1016/j.pmpp.2014.09.003