Cryphonectria parasitica (blight of chestnut)
- Taxonomic Tree
- Notes on Taxonomy and Nomenclature
- Distribution Table
- Hosts/Species Affected
- Host Plants and Other Plants Affected
- Growth Stages
- List of Symptoms/Signs
- Biology and Ecology
- Notes on Natural Enemies
- Seedborne Aspects
- Plant Trade
- Detection and Inspection
- Similarities to Other Species/Conditions
- Prevention and Control
- Distribution Maps
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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- Cryphonectria parasitica (Murrill) M.E. Barr
Preferred Common Name
- blight of chestnut
Other Scientific Names
- Diaporthe parasitica Murrill
- Endothia parasitica (Murrill) P.J. Anderson & H.W. Anderson
International Common Names
- English: blight: chestnut; blight: oak; canker of chestnut; canker: chestnut; canker: oak; chestnut blight; chestnut canker; swollen butt: scarlet oak
- Spanish: chancro de la corteza; chancro del castano
- French: chancre de l'écorce du chataignier; chancre du chataignier; maladie chancreuse du chataignier
Local Common Names
- Germany: Kastaniensterben; Krebs: Eiche; Krebs: Kastanie; Rindenkrebs: Kastanie
- ENDOPA (Cryphonectria parasitica)
Taxonomic TreeTop of page
- Domain: Eukaryota
- Kingdom: Fungi
- Phylum: Ascomycota
- Subphylum: Pezizomycotina
- Class: Sordariomycetes
- Subclass: Sordariomycetidae
- Order: Diaporthales
- Family: Valsaceae
- Genus: Cryphonectria
- Species: Cryphonectria parasitica
Notes on Taxonomy and NomenclatureTop of page The causal agent of chestnut blight, was referred to initially in 1906 as Diaporthe parasitica and then in 1912 as Endothia parasitica (Shear et al., 1917) until Barr (1978) placed the fungus in the genus Cryphonectria. Barr placed the genera Cryphonectria and Endothia in different families, Valsaceae and Gnomoniaceae, respectively, based on the arrangement of perithecia, either valsoid or diatrypoid, in stromatic tissues (see Morphology and Similarities to other species/conditions). The extensive early work of Shear et al. (1917) emphasized perithecial arrangement in C. parasitica in terms of the number of stromal layers in which perithecia were found: a Valsa-like (valsoid) arrangement was described early by Heald (1926). Roane (1986) retained the fungus in Endothia because of the lack of clarity in defining the old terms of stromatic arrangement used by Barr and the variability in orientation of perithecia in some species. The two genera are closely related (Walker et al., 1985), but also differ in the shape and septation of ascospores (see Similarities to other species/conditions). These characters aid in identification of C. parasitica, but were not used by Barr to place either of the two genera in the two familes indicated above. The type of fungal tissue (prosenchyma versus pseudoparenchyma) present in stromatic tissues is emphasized by some as being important, either qualitatively or quantitatively, in distinguishing Endothia from Cryphonectria, but all descriptions do not agree on the characterization, absence or presence, or relative abundance of each tissue type in stromata of C. parasitica (Shear et al., 1917; Kobayashi, 1970; Barr, 1978; Sivanesan and Holliday, 1981; Roane, 1986; Micales and Stipes, 1987; Hanlin, 1990). Both Cryphonectria and Endothia have Endothiella conidial states, but this name is seldom used for the chestnut blight fungus as both asexual and sexual states are typically present in the same stroma.
DescriptionTop of page On blight-susceptible American chestnut stems, conspicuous, yellow-orange or orange-brown stromata, 0.75-3 mm in diameter by 0.5-2.5 mm high, are typically abundant, randomly distributed, and sometimes contiguous on smooth-bark cankers. On rough-bark cankers of blight-resistant chestnuts, or on oaks, stromata may be infrequent or inconspicuous. They may be found only in cracks and crevices, solitary or in small or large linear groups that may be confluent. Stromata are typically slightly to moderately erumpant through the bark, are composed of both fungal and host cells, and contain conidiomata and ascomata.
The conidiomata are the first structures to form in the stroma and are surrounded by a loose growth of hyphae or prosenchyma. When the stroma first breaks through the bark periderm, stromatic cells become shorter and thicker, densely crowded together, and appear as a pseudoparenchymous tissue. This tissue layer covers the exposed surface of the stroma and the necks of the typically later-formed perithecia. Conidiomata locules are often large, convoluted, and 100 to 300 µm in diameter. Conidiophores are branched, bearing differentiated conidiogenous cells that are cylindrical, tapering at the apex, sometimes with a collarette. Conidia are hyaline, aseptate, oblong to cylindrical, 3 to 5 x 1-2 µm, and during moist periods are expelled in mucilaginous spore tendrils that are yellowish when young and coral red when old. The tendrils readily wash away during rains. Some stromata contain conidiomata and no ascomata; this is especially common for hypovirulent strains of C. parasitica, which form very small stromata on blight-susceptible chestnuts.
The ascoma is a subglobose perithecium, 300-400 µm in diameter, which often develops in the stroma below the conidiomata, but at times is formed well up in the stroma at the level with or above the conidiomata. The perithecia are white to brown, with black necks. Perithecia commonly have a valsoid arrangement, with the oblique and central long (600 or more µm) periphysate necks converging and erumpent through the stromatic disc. Some stromata have an upright arrangement of perithecia. Stromata may contain one to 60 perithecia (usually 15 to 30), which show on the surface of the stroma as a number of raised papillae or a number of black ostiolate necks. Asci (30-60 x 7-9 µm) are unitunicate, ellipsoid to subclavate with a refractive apical ring, evanescent at the base, then free at maturity, 8-spored. Ascospores (7-12 x 3-5.5 µm) are hyaline, two-celled with a median septum, ellipsoid or ovoid, with a gelatinous envelope.For further information see Shear et al. (1917); Heald (1926); Barr (1978); Sivanesan and Holliday (1981); Roane (1986); Micales and Stipes (1987); Hanlin (1990).
DistributionTop of page C. parasitica is widespread throughout the eastern USA, in China and Japan, where it is native, and in many countries of Europe that have significant Castanea populations.
See also CABI/EPPO (1998, No. 194).
Distribution TableTop of page
The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.
|Continent/Country/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
|Azerbaijan||Present||Aghayeva and Harrington, 2008; EPPO, 2014; CABI/EPPO, 2015|
|China||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Anhui||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Guangdong||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Guangxi||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Guizhou||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Hebei||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Henan||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Hubei||Present||Li GuoYuan, 2002; CABI/EPPO, 2015|
|-Jiangsu||Restricted distribution||EPPO, 2014; CABI/EPPO, 2015|
|-Jiangxi||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Liaoning||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Shaanxi||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Shandong||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Yunnan||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Zhejiang||Present||EPPO, 2014; CABI/EPPO, 2015|
|Georgia (Republic of)||Present||EPPO, 2014; CABI/EPPO, 2015|
|India||Restricted distribution||EPPO, 2014; CABI/EPPO, 2015|
|-Uttar Pradesh||Restricted distribution||EPPO, 2014; CABI/EPPO, 2015|
|Iran||Restricted distribution||EPPO, 2014; CABI/EPPO, 2015|
|Japan||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Honshu||Present||EPPO, 2014; CABI/EPPO, 2015|
|Korea, DPR||Present||EPPO, 2014; CABI/EPPO, 2015|
|Korea, Republic of||Present||EPPO, 2014; CABI/EPPO, 2015|
|Taiwan||Present||EPPO, 2014; CABI/EPPO, 2015|
|Turkey||Restricted distribution||1979||EPPO, 2014; CABI/EPPO, 2015|
|Tunisia||Present, few occurrences||EPPO, 2014; CABI/EPPO, 2015|
|Canada||Restricted distribution||EPPO, 2014; CABI/EPPO, 2015|
|-British Columbia||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Ontario||Present||EPPO, 2014; CABI/EPPO, 2015|
|USA||Restricted distribution||EPPO, 2014; CABI/EPPO, 2015|
|-California||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Connecticut||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Florida||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Georgia||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Indiana||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Iowa||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Kentucky||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Louisiana||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Maryland||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Massachusetts||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Michigan||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Minnesota||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Mississippi||Present||EPPO, 2014; CABI/EPPO, 2015|
|-New Hampshire||Present||CABI/EPPO, 2015|
|-New Jersey||Present||EPPO, 2014; CABI/EPPO, 2015|
|-New York||Present||EPPO, 2014; CABI/EPPO, 2015|
|-North Carolina||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Oregon||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Pennsylvania||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Tennessee||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Virginia||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Washington||Present||EPPO, 2014; CABI/EPPO, 2015|
|-West Virginia||Present||EPPO, 2014; CABI/EPPO, 2015|
|Austria||Restricted distribution||****||EPPO, 2014; CABI/EPPO, 2015|
|Belgium||Present||EPPO, 2014; CABI/EPPO, 2015|
|Bosnia-Hercegovina||Present||EPPO, 2014; CABI/EPPO, 2015|
|Bulgaria||Restricted distribution||EPPO, 2014; CABI/EPPO, 2015|
|Croatia||Widespread||Halembek, 1991; EPPO, 2014; CABI/EPPO, 2015|
|Czech Republic||Eradicated||Haltofová, 2006; EPPO, 2014; CABI/EPPO, 2015|
|France||Restricted distribution||EPPO, 2014; CABI/EPPO, 2015|
|-France (mainland)||Restricted distribution||CABI/EPPO, 2015|
|Germany||Present, few occurrences||198*||EPPO, 2014; CABI/EPPO, 2015|
|Greece||Restricted distribution||EPPO, 2014; CABI/EPPO, 2015; Tziros et al., 2015|
|-Crete||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Greece (mainland)||Restricted distribution||CABI/EPPO, 2015|
|Hungary||Restricted distribution||1970||Tarcali et al., 2008; EPPO, 2014; CABI/EPPO, 2015|
|Italy||Present||1938||EPPO, 2014; CABI/EPPO, 2015|
|-Italy (mainland)||Present||CABI/EPPO, 2015|
|-Sardinia||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Sicily||Present||Pennisi et al., 2005; CABI/EPPO, 2015|
|Macedonia||Present||EPPO, 2014; CABI/EPPO, 2015|
|Netherlands||Transient: actionable, under eradication||NPPO of the Netherlands, 2013; EPPO, 2014; CABI/EPPO, 2015|
|Poland||Eradicated||EPPO, 2014; CABI/EPPO, 2015|
|Portugal||Widespread||EPPO, 2014; CABI/EPPO, 2015|
|-Azores||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Madeira||Present||EPPO, 2014; CABI/EPPO, 2015|
|-Portugal (mainland)||Widespread||CABI/EPPO, 2015|
|Romania||Restricted distribution||Bolen et al., 1995; EPPO, 2014; CABI/EPPO, 2015|
|Russian Federation||Restricted distribution||EPPO, 2014; CABI/EPPO, 2015|
|-Southern Russia||Restricted distribution||EPPO, 2014; CABI/EPPO, 2015|
|Serbia||Widespread||EPPO, 2014; CABI/EPPO, 2015|
|Slovakia||Restricted distribution||1976||Bolvanský et al., 2014; EPPO, 2014; CABI/EPPO, 2015|
|Slovenia||Restricted distribution||Halembek, 1991; Tarcali et al., 2008; EPPO, 2014; CABI/EPPO, 2015|
|Spain||Restricted distribution||Bascón et al., 2014; EPPO, 2014; CABI/EPPO, 2015|
|-Spain (mainland)||Restricted distribution||CABI/EPPO, 2015|
|Switzerland||Widespread||1947||EPPO, 2014; CABI/EPPO, 2015|
|UK||Transient: actionable, under eradication||IPPC, 2012; Hunter et al., 2013; EPPO, 2014; CABI/EPPO, 2015|
|-England and Wales||Transient: actionable, under eradication||EPPO, 2014; CABI/EPPO, 2015|
|Ukraine||Present||EPPO, 2014; CABI/EPPO, 2015|
|Australia||Present, few occurrences||EPPO, 2014; CABI/EPPO, 2015|
|-Victoria||Present, few occurrences||EPPO, 2012; EPPO, 2014; CABI/EPPO, 2015|
Hosts/Species AffectedTop of page C. parasitica is most important as a pathogen of Castanea and Quercus species though it may be found as saprophyte or a weak pathogen on other species. The most important susceptible species are C. dentata (American chestnut), and C. sativa (European chestnut), although the latter is considered less susceptible than Ameican chestnut. The important Asian species, C. mollissima (Chinese chestnut) and C. crenata (Japanese chestnut) are blight resistant but can develop severe disease; C. seguinii and C. henryi, from China, are hosts and C. pumila, from eastern USA, and other chinquapins are suscepstible. Oaks such as Q. virginiana (live oak) and Q. stellata (post oak) are the only oaks in North American to be seriously affected by C. parasitica and some trees may be killed; Q. coccinea (scarlet oak) is commonly infected by C. parasitica (Roane et al., 1986; Nash and Stambaugh, 1989; Torsello et al., 1994). C. parasitica has been reported from Quercus in Slovakia (Juhasova, 1991), on Q. petraea (durmast oak) in Switzerland (Bissegger and Heiniger, 1991), and on a range of trees including Ostrya carpinifolia, Q. ilex (evergree oak), Q. pubescens (downy oak) and Alnus cordata (Italian alder) in Italy (Turchetti et al., 1991), although the symptoms were mild. Eucalyptus is also a host as are Castanopsis chrysophylla, Q. rubra, Malus, Acer , Fagus, Rhus , Carpinus, Carya, and Liriodendron species. Some of these hosts are documented from artificial inoculations (Shear et al.,1917) and have not been found to be commonly infected in nature; some hosts are based on artificial inoculations of dead (girdled) trees (Baird, 1991).
Outside the USA, it has been reported from Quercus in Slovakia (Juhasova, 1991) on Q. petraea in Switzerland (Bissegger and Heiniger, 1991), and on a range of trees including Ostrya carpinifolia, Quercus ilex, Q. pubescens and Alnus cordata in Italy (Turchetti et al., 1991) although disease symptoms were mild.
Host Plants and Other Plants AffectedTop of page
|Alnus cordata (Italian alder)||Betulaceae||Wild host|
|Castanea dentata (American chestnut)||Fagaceae||Main|
|Castanea sativa (chestnut)||Fagaceae||Main|
|Malus domestica (apple)||Rosaceae||Wild host|
|Quercus coccinea (scarlet oak)||Fagaceae||Unknown|
|Quercus frainetto (Hungarian oak)||Fagaceae||Other|
|Quercus ilex (holm oak)||Fagaceae||Unknown|
|Quercus petraea (durmast oak)||Fagaceae||Unknown|
|Quercus rubra (northern red oak)||Fagaceae||Unknown|
|Quercus stellata (Post oak)||Fagaceae||Unknown|
Growth StagesTop of page Vegetative growing stage
SymptomsTop of page
The first evidence of infection on blight-susceptible chestnut trees may be a small, flat, orange-brown area on the smooth bark tissues of the main stem or branches. These small lesions may be associated with a small, shade-killed branch. The lesions develop into sunken cankers as buff-colored mycelial fans develop in the bark, at one or more bark depths in the phloem to the vascular cambium. The margin of the cankers may slight slightly and the bark may crack. Small, yellowish to orange stromata containing conidiomata break through the bark and become larger and more numerous as the canker grows. Distinctive yellow tendrils (cirrhi) of conidia extrude from the stroma in wet weather. In the later months of the first year, papillae and/or the black necks of the perithecia become apparent on the stromata. Stromata may reach densities of 50 or more per cm² and over 1000 per canker on American chestnuts. Cankers or stromata on European chestnuts may contain no, few, or many perithecia. Soon after, depending on the diameter of the stem, the canker expands around the circumference of the stem and the vascular cambium is girdled and killed on susceptible chestnuts. Wilting and death of the foliage above the branch or stem canker follows. On older and rougher bark with rhytidome, or on blight-resistant chestnuts or oaks, cankers may not be as obvious, and stromata may be infrequent. Blight-resistant trees infected with virulent strains may have superficial and swollen cankers with thick rhytidome, due to the wound periderm formation, or callused and swollen cankers, which develop after a small area of the vascular cambium has been killed. Intermediate cankers may be formed also with irregularly swollen and sunken areas within a canker. Killing of the vascular cambium and stem death can occur in normally blight-resistant Chinese chestnuts after severe spring frosts or at high altitude, but it is uncommon. Blight-susceptible chestnuts infected with hypovirulent strains may exhibit the same canker types exhibited by blight-resistant chestnuts and oaks infected with virulent strains. In the case of the former, only stromata with conidiomata may be formed by the hypovirulent strains. For further information, see Shear et al. (1917); Heald (1926); Sivanesan and Holliday (1981); Roane et al. (1986); Heineger and Rigling (1994); Guérin et al. (2000); Hogan and Griffin (2002).
List of Symptoms/SignsTop of page
|Leaves / wilting|
|Stems / canker on woody stem|
|Stems / dieback|
|Stems / gummosis or resinosis|
|Whole plant / plant dead; dieback|
Biology and EcologyTop of page Wounds in the bark, of mechanical or biological origin are the main infection court of C. parasitica and cankers are often associated with a dead branch stub on the stem. The primary source of virulent inoculum is ascospores, which are produced in abundance on blight-susceptible chestnut trees, and, to a lesser extent, on blight-resistant chestnut trees and oaks. Ascospores are disseminated in air currents and have been detected in large numbers as far as 90-120 m from a perithecial source; they were expelled every day for 168 days. Ascospores are expelled from perithecia in stroma on cankers following a warm rain, primarily in the spring and early summer. The optimum temperature range for expulsion is 20-27°C. The amount of sexual reproduction in a C. parasitica population is critical to the genetic diversity in that population and the number of vegetative compatibilty types of the fungus, which affects the spread of hypoviruses involved in blight control (see Control). The smaller conidia are considered less infective than ascospores and are primarily water and insect-borne. Tendrils of conidia in cirrhi, formed in wet weather on a stroma, may contain 100 million conidia (Heald, 1926). Propagules, likely conidia or mycelial fragments, have been associated with insects in Coleoptera and Diptera (Russin et al., 1984;Pakaluk and Anagnostakis, 1997) mites (Wendt et al., 1983; Nannelli et al., 1998), and slugs (Turchetti and Chelazzi, 1984). Large numbers of conidia have been recovered from birds (Heald and Studhalter, 1914).
The fungus colonizes bark tissues by the growth of mycelial fans that may be found in several layers of the bark; host cells are killed in advance of colonization. Resistant hosts, such as Chinese chestnut, infected with virlulent strains, and susceptible chestnut hosts, infected with hypovirulent (reduced virulence) strains, have fewer mycelial fans. Hypovirulent strains of C. parasitica may naturally be present in cankers (see Control) and are frequently infected with a dsRNA virus (hypovirus). Most chestnut cankers resulting from colonization by virulent strains are sunken, which kills the stem or branch and all foliage above the canker. Resistant hosts, infected with virulent strains, or susceptible chestnuts, infected with hypovirulent strains, may have non-killing, swollen, superficial cankers or callused and swollen cankers. Stromata containing conidiomata and/or perithecia are abundant on sunken cankers but often infrequent on swollen cankers. Sunken cankers on main stems can kill the entire tree above the ground, often within 1or 2 years, but stump sprouts commonly develop after stem death unless sunlight is too low for shoot growth or browse damage is high (Sivanesan and Holliday, 1981; Roane et al., 1986; Griffin et al., 1991).
Notes on Natural EnemiesTop of page Naturally occurring dsRNA viruses (hypoviruses) which confer hypovirulence have been extensively studied (see Control).
Soil or bark microbes can be natural antagonists.
Seedborne AspectsTop of page Incidence
C. parasitica is not well known as a seedborne pathogen. Around 14% of the nuts harvested from a planting of American chestnuts at Hamden, Connecticut, USA, in which chestnut blight was prevalent were infected with C. parasitica. Signs of infection appeared after storage at 4°C followed by incubation at 18-25°C, but the infections apparently were initiated while the nuts were on the tree. Infections were confined to the shell and appeared not to affect seed germination or seedling growth (Jaynes and DePalma, 1984).
Effect on Seed Quality
The only study available suggests that seed viability is not affected by this fungus, as only the outer tissues are affected (Jaynes and DePalma, 1984).
The fungus can be transmitted through seed contamination (Jaynes and DePalma, 1984) but infection of seedlings was not demonstrated.
Seed Health Tests
No tests have been developed.
Plant TradeTop of page
|Plant parts liable to carry the pest in trade/transport||Pest stages||Borne internally||Borne externally||Visibility of pest or symptoms|
|Bark||fruiting bodies; hyphae|
|Leaves||hyphae||Yes||Pest or symptoms usually invisible|
|Stems (above ground)/Shoots/Trunks/Branches||fruiting bodies; hyphae||Yes||Yes||Pest or symptoms usually visible to the naked eye|
|True seeds (inc. grain)||hyphae||Yes||Yes||Pest or symptoms usually invisible|
|Plant parts not known to carry the pest in trade/transport|
|Fruits (inc. pods)|
|Growing medium accompanying plants|
ImpactTop of page The American chestnut was the most important tree species in the former oak-chestnut forest and a dominant species in the mixed mesophytic forest of eastern North America. It was very important economically for timber, edible nuts, and tannins. Within 40 years following discovery in 1904 of the chestnut blight disease in New York City, almost all canopy or 3.5 billion American chestnut trees were killed by C. parasitica in these regions. The area affected ranged from Maine in the north to Alabama in the south to southeastern Michigan, Indiana, and Ontario in the west. The American chestnut survives presently as mostly non-flowering, small understory trees on which C. parasitica is endemic. In Europe, the chestnut blight disease was discovered in 1938, and has been less destructive there, killing fewer trees than in North America. The lower level of disease may result in part from a higher level of blight resistance in the European chestnut; also, the recovery of chestnut stands and coppice stems in Italy, southern Switzerland, and surrounding countries has been associated with the natural occurrence of hypovirulent strains of C. parasitica (Anagnostakis, 1982; Heineger and Rigling, 1994; Roane et al., 1986). Although Chinese chestnut is considered to be highly blight resistant, variation in resistance has been found among Chinese chestnut cultivars and wild chestnut trees, and C. parasitica presently is considered to be the most important pathogen affecting the genus Castanea in China (Qin et al., 2002). The pathogen is also sometimes destructive in Japan on the relatively blight-resistant Japanese chestnut (Uchida, 1977). On oaks, the pathogen has been an important pathogen on live oak and post oak in the USA (Roane et al., 1986).
UsesTop of page Enzymes from C. parasitica are widely used as coagulants in cheese production, as substitutes for animal rennets (International Dairy Foundation, 1996; Krause et al., 1996; Perennou, 1997).
DiagnosisTop of page Diagnosis of C. parasitica on chestnuts should rely on the presence of small to large cankers on the stems or branches, yellow-orange or orange-brown stromata with characteristic ascospores in perithecia (see Morphology) and the presence of buff-colored mycelial fans in bark tissues. Sometimes only stromata with conidiomata may be present. Death of the crown should not be relied upon alone as root disease, associated with Phytophthora and Pythium species, can kill trees or seedlings, especially on heavy soils. However, death of a stem or branch with foliage blight above a canker is useful in diagnosis. On oaks, the presence of cankers with mycelial fans and ellipsoid ascospores in perithecia can be used to diagnose C. parasitica. On blight-susceptible chestnuts, ascospores may be present as early as the last months of the first year of disease, and mycelial fans will be present in small cankers. Molecular probes, reproductive morphology, and virulence characters can be used to distinguish C. parasitica from related species (see Similarities to other conditions). Virulent C. parasitica strains are readily isolated from diseased tissues with conventional surface disinfestation and tissue plating on acidified, fungal-isolation media, such as acidified, potato-dextrose agar. Colonies should be yellow-orange and radially symetric. Hyovirulent C. parasitica strains, infected with dsRNA hypoviruses, often have an abnormal colony appearance or atypical yellow-orange pigmentation and asexual sporulation. For European hypovirulent strains, colonies typically have a predominantly white colony appearance; single-conidium isolation can be used to recover the normal, yellow-orange pigmented, hypovirus-free colony type. For further information, see Sivanesan and Holliday (1981) and Roane et al. (1986).
Detection and InspectionTop of page Most smooth-bark cankers on stems or branches of Castanea species are conspicuous, especially if yellow-orange stromata are present. Rainfall or damp weather enhances the detection of cankers from a distance due to increased brilliance of the stromatal colors. The presence of stromata is common on cankers a few months or more old. These cankers are also typically sunken with mycelial fans in the infected tissues; after several months the necrotic bark may slough away, exposing the wood. Foliage blight symptoms, or flagging, may develop on the branch or stem above the canker within 1 or 2 years after cankers are apparent. On stems or branches with rough bark , or on blight-resistant chestnut or oak species, cankers and stromata may not be conspicuous. A knife or axe may be required to dissect and reveal the necrosis in the bark. There is often a slight to moderate swelling or callus formation on the stems and stromata may only be found in bark cracks. Use of a magnifying lens facilitates observation of stromata in bark cracks. On Q. coccinea (scarlet oak), C. parasitica infection is usually indicated by the pronounced swelling of the base of the stem, giving rise to the name, 'swollen butt disease'. Careful examination may or may not reveal stromata in bark crevices. Dissection of bark tissues, as indicated above, should reveal the presence of necrosis and mycelial fans characteristic of C. parasitica. For chestnuts, the base of the stem is also the most common location of cankers, but branch cankers and flagging may be more obvious. For further information see Sivanesan and Holliday (1981) and Roane et al. (1986).
Similarities to Other Species/ConditionsTop of page The Cryphonectria species most likely to be encountered on the same hosts as C. parasitica is C. radicalis. Unlike C. parasitica, this uncommon species is only weakly pathogenic on chestnut, and can be distinguished from C. parasitica by the significantly smaller ascospores, by nuclear and mitochondrial probes, and by the inability to cross with C. parasitica (Hoegger et al., 2002; Shear et al., 1917). Other related Cryphonectria species (C. gyrosa and C. havanensis) are not pathogenic, nor are they reported on chestnut and, except for Japan, are not usually found in regions where C. parasitica is present (Roane et al., 1986). Also, these two similar species have more narrowly ellipsoid ascospores than C. parasitica (Barr, 1987). Endothia gyrosa, in the family Gnomoniaceae, occurs on oak and other hosts of C. parasitica, and has a diatrypoid or upright configuration of perithecia in stromata; C. parasitica, in the family Valsaceae, commonly has a valsoid configuration of perithecia. In addition, E. gyrosa has allantoid, one-celled ascospores (Barr, 1987; Roane et al., 1986), whereas C. parasitica has ellipsoid or ovoid two-celled ascospores. The buff-colored mycelial fans of C. parasitica in bark tissues are not formed by any other Cryphonectria species and furnish the most reliable field character for distinguishing it from related species.
Prevention and ControlTop of page Research on the control of C. parasitica centres around breeding for blight resistance using the high levels of blight resistance found in Asian chestnut species; breeding using the lower levels of blight resistance in some European and large, surviving American chestnuts; the use of hypovirulent strains of C. parasitica that are infected with dsRNA hypoviruses; forest management practices; and application of soil and microbe compresses to cankers. After early efforts to hybridize Chinese and Japanese chestnut with American chestnut did not produce blight-resistant trees with forest-tree form, the backcross method of breeding was suggested and adopted (Burnham, 1981). With this approach, blight-resistance genes from Chinese chestnut are transferred to American chestnut by backcrossing the most resistant Chinese x American hybrids to American chestnut three or more times. Then, the most blight-resistant progeny are intercrossed to obtain the level of blight resistance of Chinese chestnut with the forest-tree form of American chestnut. Good progress with the aid of molecular mapping is being made in this approach (Anagnostakis, 1992; Kubisiak et al., 1997; Hebard, 2002). Selected trees of European chestnut (Bazzigher, 1981) and some large, surviving American chestnut trees (Griffin et al., 1983) have been shown to have useful levels of blight resistance. The American chestnuts have been intercrossed to increase the level of blight resistance (Griffin, 2000).
Hypovirulent strains of C. parasitica, infected with Cryphonectria hypovirus 1 (CHV1), have spread naturally on European chestnut in Italy, southern Switzerland, and surrounding countries (Alleman et al., 1999). The spread of CHV1 and the hypovirulent strain have been associated with natural chestnut blight control on European chestnut, even though C. parasitica strains in different vegetative compatibility types potentially limit hypovirus transmission (Heiniger and Rigling, 1994). In addition, artificial inoculation of cankers with hypovirulent strains has been used successfully to control blight on European chestnut infected with a few vegetative compatiblity types of C. parasitica (Grente, 1981;Turchetti and Maresi, 1988). In contrast, cork-borer hole inoculations around cankers with mixtures of CHV1-infected C. parasitica and other hypovirulent strains of American origin have not generally resulted in blight control on blight-susceptible American chestnuts. Vegetative incompatibility among the many strains of C. parasitica present on American chestnut has been identified as an important barrier to transmission of hypoviruses and conversion of virulent strains to hypovirulence (Anagnostakis and Day, 1979; Liu and Milgoom, 1996). In addition, the abundance of virulent inoculum in American chestnut stands, the high blight susceptibility of American chestnut, and stressful site factors may result in rapid tree death and insufficient time for CHV1 and other hypoviruses to spread (Griffin, 2000). Inoculation of CHV1-infected C. parasitica strains around cankers on grafted American chestnut trees, derived from large survivors, has resulted in a high level and a long period (20 years) of blight control; spread of CHV1 into a large number of C. parasitica vegetative compatibility types on these trees has occurred (Hogan and Griffin, 2002). In this integrated management approach, low levels of blight resistance in the chestnut grafts, spread of CHV1, favorable spatial patterns of hypovirulent strains and vegetative compatibility types on the trees, and favorable forest management factors (low attitude, mesic site with control of competing hardwoods) have been associated with blight control.
Application of soil, compost, or sphagnum peat compresses, or antagonistic microbes (Roane et al., 1986; Tattar et al., 1996) to individual cankers has resulted in some blight control, but is limited to accessible parts of a tree, such as the lower stem or branches. Each canker must be treated separately. Soil compresses should be applied early in canker development to be effective. Graft unions can be protected from C. parasitica using soil from the base of the tree in a similar manner. Chemical control has not been effective except for the protection of graft unions with fungicides (Turchetti et al., 1981; Canciani et al., 1995).
ReferencesTop of page
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