Ceratocystis fimbriata (Ceratocystis blight)
- Summary of Invasiveness
- Taxonomic Tree
- Notes on Taxonomy and Nomenclature
- Distribution Table
- History of Introduction and Spread
- Risk of Introduction
- Hosts/Species Affected
- Host Plants and Other Plants Affected
- Growth Stages
- List of Symptoms/Signs
- Biology and Ecology
- Means of Movement and Dispersal
- Pathway Vectors
- Plant Trade
- Wood Packaging
- Impact Summary
- Environmental Impact
- Impact: Biodiversity
- Social Impact
- 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
- Ceratocystis fimbriata Ellis & Halst.
Preferred Common Name
- Ceratocystis blight
Other Scientific Names
- Ceratostomella fimbriata (Ellis & Halst.) J.A. Elliott
- Endoconidiophora fimbriata (Ellis & Halst.) R.W. Davidson
- Ophiostoma fimbriatum (Ellis & Halst.) Nannf.
- Rostrella coffeae Zimm.
- Sphaeronaema fimbriata (Ellis & Halst.) Sacc.
International Common Names
- English: black cane rot of Syngonium; black canker of aspen; black rot of sunn hemp; black rot of sweet potato; black rot of taro; blight of mango; cacao wilt; canker of coffee; canker stain of plane tree; Ceratocystis canker; Ceratocystis wilt; Ceratostomella wilt; mallet canker; mallot wound canker; mango blight; mango wilt; mouldy rot of rubber; sweet potato black rot; target canker of aspen; wilt disease of cocoa
- Spanish: llaga macana de cacao; mal de choroni de cacao; mal de machete; mal del machete de cacao; marchitez de chupones: cacao; muerte subita de citrus; necrosis del tronco del cacao; secamiento de los citricos
- French: chancre colore du platane; fletrissement du cacaoyer; pourriture des saignees de l'hevea; tache chancreuse
Local Common Names
- Brazil: seca da mangueira
- Germany: Schwarzfäule: Süsskartoffel; Welke: Kaffee; Welke: Kakao
- CERAFI (Ceratocystis fimbriata)
Summary of InvasivenessTop of page The literature suggests that introduced populations of C. fimbriata have thus far remained restricted to particular cultivated hosts. This is probably due to the relatively narrow host range of the introduced strains (Walker et al., 1988; Johnson et al., 2002; Baker et al., 2003). However, there has been little attempt to see if the introduced populations have spread to other hosts. Some forms of the fungus, such as those occurring in Brazil, have relatively broad host ranges (Baker et al., 2003) and may potentially be more invasive if introduced to new ecosystems. The inefficiency of dispersal by insects may also limit the invasiveness of C. fimbriata.
Taxonomic TreeTop of page
- Domain: Eukaryota
- Kingdom: Fungi
- Phylum: Ascomycota
- Subphylum: Pezizomycotina
- Class: Sordariomycetes
- Subclass: Hypocreomycetidae
- Order: Microascales
- Family: Ceratocystidaceae
- Genus: Ceratocystis
- Species: Ceratocystis fimbriata
Notes on Taxonomy and NomenclatureTop of page Ceratocystis fimbriata, the type species of the genus, was originally described on sweet potato (Ipomoea batatas) in 1890 (Halsted, 1890). Saccardo (1892) transferred the species to Sphaeronaema, Elliott (1923) transferred it to Ceratostomella, Melin and Nannfeldt (1934) transferred it to Ophiostoma, and Davidson (1935) transferred it to Endoconidiophora. Placement in Ceratocystis has been accepted since 1950 (Bakshi, 1950).
A fungus attacking Coffea in Indonesia was described as Rostrella coffea (Zimmerman, 1900), and this species was later synonymized with C. fimbriata (Pontis, 1951), although no careful comparisons have been made. Walter et al. (1952) designated the pathogen attacking Platanus as a separate form on the basis of its purported host specificity; see separate datasheet on C. fimbriata f. platani. Another form, occurring on Acacia mearnsii and species of Protea in South Africa, is now considered a separate species, C. albofundus (Wingfield et al., 1996); it is probably native to southern Africa (Roux et al., 2000). C. variospora, found on Quercus and described by Davidson (1944), is similar to C. fimbriata (Hunt, 1956). Although Upadhyay (1981) considered C. variospora a synonym of C. fimbriata, it is probably a separate species. It is becoming increasingly apparent that C. fimbriata is a complex of many species, each with a unique host range and geographic distribution.
DescriptionTop of page C. fimbriata grows readily on most agar media. Mycelium is hyaline at first, later turning dark greenish-brown. Within a few days there are usually abundant conidiophores that produce chains of hyaline conidia, sometimes called endoconidia, characteristic of the anamorph genus Chalara. However, Chalara species are anamorphs of discomycetes, and the genus Thielaviopsis is now used for anamorphs of Ceratocystis species (Paulin et al., 2002). Endoconidia are cylindrical and may vary in size from 11 to 16 mm long by 4 to 5 mm wide (all measurements are from Hunt, 1956). Specialized conidiophores give rise to thick-walled, pigmented aleurioconidia (sometimes called chlamydospores), probably a survival spore. Aleurioconidia are typically 9-16 mm long and 6-13 mm wide, borne singly or in short chains. Endoconidia may also darken and become thick walled chlamydospores, thus resembling aleurioconidia. Endoconidia, chlamydospores formed from endoconidia, and aleurioconidia may be produced on and within the substratum.
The teleomorph of the fungus is well adapted to insect dispersal. The fungus has two mating types, and MAT-1 isolates can only produce perithecia when paired with MAT-2 isolates. However, MAT-2 isolates are self-fertile due to uni-directional mating type switching (Harrington and McNew, 1997; Witthuhn et al., 2000). Most field isolates are MAT-2 and self-fertile, producing many fruiting bodies (ascomata) on the surface of the host or in culture, often within one week. Ascomata are dark brown to black and globose, 130-200 µm diameter with a long, thin neck up to 800 µm long, through which the ascospores are exuded. The opening at the tip of the neck has 8 to 15 ostiolar hyphae ranging in length from 50 to 90 µm. Ascospores are small, hyaline and hat-shaped, 4.5-8 µm long by 2.5-5.5 µm wide, and accumulate in a sticky matrix at the tip of the ascomatal neck, where they appear as a cream to pink ball or coil.
DistributionTop of page In addition to the published reports, the following specimens are held in the US National Fungus Collections: Mexico (BPI 596218 and 595433), St Vincent and Grenadines (BPI 596219), Massachusetts and Rhode Island, USA (BPI 595868 and 595867, respectively); and there is an accession from Suriname in the American Type Culture Collection (ATTC 14503). Confirmed isolates of C. fimbriata have also been collected from Iowa (on Carya cordiformis), Missouri (on Platanus occidentalis) and Wisconsin, USA (on C. cordiformis) (TC Harrington, Iowa State University, USA, unpublished data).
Several older reports of C. fimbriata (cited in CMI, 1983) may be erroneous but have been included in the listed distribution. The fungus has been reported as a saprobe on Hevea in Uganda (Snowden, 1926), and two reports have suggested it as a pathogen on Hevea in the Congo Democratic Republic (Ringoet, 1923; Anon., 1948). Unverified voucher specimens from Fagus and Larix in the UK are cited in CMI (1983), but Larix is a very unlikely host, and there are no confirmed reports of the fungus from the UK. The report of the fungus on Theobroma in the Philippines (Eloja and Gandia, 1963) was only a tentative identification.
Several unnamed forms of C. fimbriata appear to be indigenous to North and South America or Asia but have been introduced elsewhere. Different hosts are attacked in different regions, and even in regions where the fungus is common, not all potential hosts are attacked. For example, mango wilt is known only in Brazil, although Mangifera is grown in other areas where C. fimbriata is common on other plants. The Theobroma form is restricted to Central America and northern and eastern South America, while Coffea forms apparently occur only in Central America and northern South America and, perhaps, a few locations in South-East Asia (Zimmerman, 1900).
Because of the numerous cryptic species in the C. fimbriata complex and the history of human-mediated movement of host-specialized strains around the world (Baker et al., 2003), it is difficult to know which of the reports of C. fimbriata in specific countries are of native populations of C. fimbriata or of exotic populations. For some cases where there is clear evidence that the pathogen was introduced, such as on the ornamental cultivars of Syngonium (Walker et al., 1988), it appears that the fungus has been restricted to cultivated plants in nurseries or greenhouses. Otherwise, the introduced strains are considered to be invasive populations.
Note: IMI Herbarium, various dates, in the distribution table indicates records for which specimens are held in the herbarium at the International Mycological Institute (now CABI Bioscience). The herbarium also contains specimens from Honduras and Australia.
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.Last updated: 23 Apr 2020
|Continent/Country/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
|Congo, Democratic Republic of the||Present, Localized||EPPO (2020)|
|Congo, Republic of the||Present||Introduced||2000||Invasive||Roux et al. (2000)|
|Côte d'Ivoire||Present, Localized||DAVET (1962); EPPO (2020)|
|Ghana||Absent, Unconfirmed presence record(s)||EPPO (2020); Herb IMI (Undated)|
|Seychelles||Present, Localized||EPPO (2020)|
|South Africa||Present, Localized||Roux et al. (2000); EPPO (2020)|
|Uganda||Present, Localized||Introduced||2001||Invasive||Roux et al. (2001); EPPO (2020)|
|Brunei||Present, Localized||EPPO (2020)|
|Cambodia||Present, Localized||EPPO (2020)|
|China||Present, Localized||Native||SY (1956); EPPO (2020)|
|-Fujian||Present||Hu FangPing et al. (1999)|
|-Sichuan||Present||Xu et al. (2011)|
|-Yunnan||Present||Huang et al. (2008); Li et al. (2014); Li et al. (2014a); Li et al. (2016)|
|India||Present, Localized||Kaushik and Toky (1992); EPPO (2020); CABI (Undated); Herb IMI (Undated)|
|-Andhra Pradesh||Present||Somasekhara (2006)|
|-Maharashtra||Present||Somasekhara and Wali (2000)|
|Indonesia||Present, Localized||Wright (1925); Tayler and Stephens (1929); EPPO (2020)|
|-Java||Present||Zimmerman (1900); South and Sharples (1925); Leefmans (1934)|
|-Sumatra||Present||Tayler and Stephens (1929)|
|Japan||Present, Localized||Asuyama (1938); Shimizu (1939); Okamoto (1940); EPPO (2020)|
|-Kyushu||Present||Native||Kato et al. (1982); Kajitani and Kudo (1993)|
|Malaysia||Present, Localized||South and Sharples (1925); Beeley (1929); EPPO (2020); Herb IMI (Undated)|
|Myanmar||Present, Localized||Turner and Myint (1980); EPPO (2020)|
|North Korea||Present, Localized||EPPO (2020)|
|Oman||Present||Wyk et al. (2005)|
|Pakistan||Present||Fateh et al. (2006); CABI (Undated)|
|Philippines||Present, Localized||EPPO (2020)|
|South Korea||Present, Localized||EPPO (2020)|
|Taiwan||Present, Localized||Lee ChunYee and Kuo KerChung (1997); EPPO (2020)|
|Thailand||Present, Localized||EPPO (2020)|
|Vietnam||Present, Localized||EPPO (2020)|
|France||Present, Localized||Introduced||1974||Invasive||Vigouroux (1986); Grosclaude et al. (1991); EPPO (2020); CABI (Undated);|
|Italy||Present, Localized||Introduced||Invasive||Panconesi (1981); Panconesi (1999); EPPO (2020)||First reported: 1940s|
|Poland||Present||Introduced||1977||Przybył (1980); Przybyl (1986); CABI (Undated)|
|Portugal||Present, Localized||EPPO (2014); EPPO (2020)|
|-Azores||Present||Introduced||Bensaude (1927); EPPO (2020)|
|Switzerland||Present||Introduced||1986||Invasive||Matasci and Gessler (1997); CABI (Undated)|
|United Kingdom||Absent, Unconfirmed presence record(s)||EPPO (2020); Herb IMI (Undated)|
|Canada||Present, Localized||EPPO (2020)|
|-British Columbia||Present||Native||Lowe (1969); Hinds (1985)|
|-Quebec||Present||Native||Vujanovic et al. (1999); CABI (Undated)|
|Costa Rica||Present, Localized||Native||Martin (1949); ECHANDI and SEGALL (1956); Siller (1958); EPPO (2020)|
|Cuba||Present, Localized||Native||Rodriguez T. and Alfonso H. (1978); Herrera Isla and Grillo Ravelo (1989); Martín Triana and Alvárez Díaz (1989); EPPO (2020)|
|Dominican Republic||Present, Localized||Schieber (1969); EPPO (2020)|
|El Salvador||Present, Localized||EPPO (2020)|
|Grenada||Present, Localized||EPPO (2020)|
|Guatemala||Present, Localized||Native||SZKOLNIK (1951); SCHIEBER and SOSA (1960); Tejada (1983); EPPO (2020)|
|Haiti||Present, Localized||Barker (1926); EPPO (2020)|
|Honduras||Absent, Unconfirmed presence record(s)||Herb IMI (Undated)|
|Jamaica||Present, Localized||Native||LEATHER (1966); EPPO (2020); Herb IMI (Undated)|
|Mexico||Present, Localized||Native||MARTIN (1947); EPPO (2020)|
|Nicaragua||Present, Localized||EPPO (2020)|
|Panama||Present, Localized||EPPO (2020)|
|Puerto Rico||Present, Localized||EPPO (2020)|
|Saint Lucia||Present, Localized||EPPO (2020)|
|Saint Vincent and the Grenadines||Present, Localized||EPPO (2020); CABI (Undated)|
|Trinidad and Tobago||Present, Localized||Briant (1932); Baker (1936); Leach (1946); Baker and Dale (1951); Iton (1959); EPPO (2020); Herb IMI (Undated)|
|United States||Present, Widespread||Native||EPPO (2020); Herb IMI (Undated)|
|-Alaska||Present||Native||Hinds and Laurent (1978)|
|-Arkansas||Present||Native||McCracken and Burkhardt (1977)|
|-California||Present||Native||Davis (1953); DeVay et al. (1968); Hinds (1985); Perry and McCain (1988)|
|-Delaware||Present||Native||MOOK (1940); Walter (1946)|
|-District of Columbia||Present||Native||WALTER et al. (1952)|
|-Florida||Present||Native||Alfieri et al. (1994)|
|-Hawaii||Present||Native||Uchida and Aragaki (1979); EPPO (2020)|
|-Louisiana||Present||Native||WEBSTER and BUTLER (1967); Baker et al. (2003)|
|-Massachusetts||Present||CABI (Undated)||Original citation: Herb. BPI|
|-Minnesota||Present||Native||WOOD and FRENCH (1963); Hinds and Anderson (1970)|
|-New Jersey||Present||Native||DODGE (1940); Walter (1946)|
|-New Mexico||Present||Native||HINDS (1972)|
|-New York||Present||Native||Walter (1946)|
|-North Carolina||Present||Native||Walter (1946)|
|-North Dakota||Present||Native||Hinds (1985)|
|-Pennsylvania||Present||Native||Jackson and Sleeth (1935); DODGE (1940); Walter (1946); WEBSTER and BUTLER (1967)|
|-Rhode Island||Present||Native||CABI (Undated)||Original citation: Herb. BPI|
|-South Dakota||Present||Native||Hinds (1985)|
|-Tennessee||Present||Native||MOOK (1940); Walter (1946)|
|-Virginia||Present||Native||Walter (1946); WEBSTER and BUTLER (1967)|
|-West Virginia||Present||Native||Walter (1946)|
|American Samoa||Present, Localized||EPPO (2020)|
|Australia||Present||CABI (Undated a)||Present based on regional distribution.|
|-New South Wales||Present, Localized||Introduced||Walker et al. (1988)|
|-Queensland||Present, Localized||Introduced||Walker et al. (1988)|
|-South Australia||Present, Localized||Introduced||Vogelzang and Scott (1990)|
|-Victoria||Present, Localized||Introduced||Walker et al. (1988)|
|Fiji||Present, Localized||Graham (1965); Firman (1972); Walker et al. (1988); EPPO (2020)|
|New Zealand||Present, Localized||SLADE (1960); Baker et al. (2003); EPPO (2020)|
|Papua New Guinea||Present, Localized||MANN (1953); Walker et al. (1988); Baker et al. (2003); EPPO (2020); Herb IMI (Undated)|
|Samoa||Present, Localized||Walker et al. (1988); EPPO (2020)|
|Solomon Islands||Present, Localized||EPPO (2020)|
|Brazil||Present, Localized||Native||EPPO (2020); Herb IMI (Undated)|
|-Bahia||Present, Localized||Native||Pereira and Santos (1986); Bezerra (1997); Firmino et al. (2013)|
|-Goias||Present||Melo Filho et al. (2002)|
|-Mato Grosso||Present||Firmino et al. (2012)|
|-Minas Gerais||Present||Native||MÜLLER (1937); CHARDON et al. (1940); Melo et al. (2016)|
|-Para||Present||Native||Deslandes (1944); Albuquerque et al. (1972); Muchovej et al. (1978)|
|-Pernambuco||Present, Localized||Native||Batista (1947); Batista (1960); Upadhyay (1981)|
|-Piaui||Present||Viana and Silva (2001)|
|-Rio de Janeiro||Present||Native||Baker et al. (2003); Carvalho and Carmo (2003)|
|-Rio Grande do Sul||Present||Santos and Ferreira (2003)|
|-Rondonia||Present||Native||Bastos and Evans (1978)|
|-Sao Paulo||Present, Widespread||Native||Arruda (1940); Oliveira (1966); Valarini and Tokeshi (1980); Silveira et al. (1985); Ribeiro et al. (1987); Firmino et al. (2015)|
|Colombia||Present, Widespread||Native||Garces (1944); PONTIS and VIDELA] (1951); Arbelaez (1957); Mourichon (1994); Borja et al. (1995); Marin et al. (2003); EPPO (2020); Herb IMI (Undated)|
|Ecuador||Present, Localized||Native||Rorer (1918); Desrosiers and Diaz (1956); Desrosiers (1957); Chalmers (1969); EPPO (2020)|
|Guyana||Present, Localized||Bisessar (1965); EPPO (2020)|
|Peru||Present, Localized||Rada (1939); Krug and Quartey-Papafio (1964); Soberanis et al. (1999); EPPO (2020)|
|Suriname||Present, Localized||Native||Baker et al. (2003); EPPO (2020)|
|Uruguay||Present||Barnes et al. (2003)|
|Venezuela||Present, Widespread||Native||PONTIS and VIDELA] (1951); Malaguti (1952); MALAGUTI (1952a); Reyes (1988); EPPO (2020); Herb IMI (Undated)|
History of Introduction and SpreadTop of page The Populus form is most abundant in North America, but it has also appeared in Poland and perhaps India, most probably from recent introductions. Cuttings of various Populus species and hybrids were brought into Poland from North America in the 1970s, and C. fimbriata may have been introduced to Poland in these cuttings. Cuttings of P. balsamifera have been shown to harbour the fungus in nurseries in Quebec, Canada (Vujanovic et al., 1999). The disease was severe in experimental plantings in Poland (Gremmen and de Kam, 1977; Przybyl, 1980, 1986); however, the disease appears to have lessened in importance in recent years and may no longer be present.
The pathogen on Platanus species, f. platani, is believed to be specialized to that genus and was probably introduced to Naples, Italy, during World War II on colonized crating material or dunnage from the USA (Panconesi, 1981, 1999; Santini and Capretti, 2000; Baker et al., 2003). The pathogen has spread throughout northern Italy (Pancosi 1981, 1999) to Switzerland in 1986 (Matasci and Gessler, 1997) and to southern France (Ferrari and Pichenot, 1974, 1976, 1979; Vigouroux, 1986; Grosclaude et al., 1991b).
The cacao form of the pathogen may have been introduced to the state of Bahia in Brazil on infected cuttings of Theobroma cacao (Harrington, 2000; Baker et al., 2003). The recent reports of the eucalyptus form of the pathogen in Uganda and the Congo may also be due to introductions on cuttings from Brazil (Roux et al., 2000, 2001a; Baker et al., 2003).
The Syngonium form of the pathogen has been dispersed on cuttings of this plant and has been reported in greenhouses in California, Florida, Hawaii and Australia (Davis, 1953; Uchida and Aragaki, 1979; Walker et al., 1988; Alfieri et al., 1994).
The Ipomoea form of the fungus has probably been spread to many locations on storage roots. For example, the report of C. fimbriata in the Azores (Bensaude, 1927) was on experimental plantings of Ipomoea germplasm imported from the Caribbean. The Ipomoea form is apparently native to Latin America and/or the Caribbean (Baker et al., 2003).
Risk of IntroductionTop of page As most forms of C. fimbriata are easily transmitted in cuttings, unrestricted movement of cuttings or other propagative material is potentially dangerous. It is likely that the fungus has been spread to new countries or regions on cuttings of Populus, Theobroma, Eucalyptus and Syngonium and on storage roots of Ipomoea. Circumstantial evidence points to packing materials as the source of the plane tree pathogen in southern Europe, and the fungus is known to survive for up to 5 years in wood, probably in the form of aleurioconidia. C. fimbriata is listed as among the highest risk pathogens that could be imported into the USA on eucalyptus logs and chips from South America (Kliejunas et al., 2001). The Platanus form (C. fimbriata f. platani) is listed as an EPPO A2 quarantine pest (OEPP/EPPO, 1986).
Hosts/Species AffectedTop of page A wide variety of annual and perennial plants are attacked by C. fimbriata. There are several apparently host-specialized strains that are sometimes called 'types', 'races' or 'forms' (Wellman, 1972; Harrington, 2000; Baker et al., 2003), and many of these may prove to be distinct species. Webster and Butler (1967a) considered such types as members of a single, highly variable species. However, isolates from some hosts and some regions are genetically unique (Santini and Capretti, 2000; Barnes et al., 2001; Johnson et al., 2002; Baker et al., 2003; Marin et al., 2003). Harrington (2000) proposed that the cryptic species within the C. fimbriata complex fall into three broad geographic clades: the North American, the Latin American and the Asian clades. Both rDNA and alloenzyme analyses support these three major clades (Harrington, 2000; Johnson et al., 2002; Baker et al., 2003).
Cross-inoculation studies have established the host-specificity of some of these types. For example, isolates from Mangifera (Ribeiro and Coral, 1968), Ipomoea, Platanus, Gmelina, Coffea, Xanthosoma, Eucalyptus (Baker et al., 2003), Crotalaria, Cajanus and Acacia (Coral et al., 1984) did not infect Theobroma. Isolates from Ipomoea and Colocasia were host-specific when inoculated to these two hosts (Mizukami, 1951), as were isolates from Hevea and Ipomoea (Olson and Martin, 1949), and Coffea and Ipomoea (Pontis, 1951). Isolates from Coffea, Prunus, Theobroma, Quercus and Colocasia failed to infect Ipomoea (Kojima and Uritani, 1976). Isolates from Platanus, Prunus (almond and apricot), Mangifera, Xanthosoma, Gmelina, Eucalyptus and Theobroma were not pathogenic to Ipomoea, and isolates from Ipomoea, Prunus (almond and apricot), Platanus, Coffea, Mangifera, Xanthosoma, Gmelina, Eucalyptus and Theobroma were not pathogenic to Platanus (Crone, 1963; Baker et al., 2003). Costa Rican isolates from Theobroma, Coffea and Xanthosoma were specialized to their respective hosts (Baker et al., 2003). Among Brazilian isolates from various hosts, only a Gmelina isolate could infect Gmelina (Baker et al., 2003). A Syngonium isolate from Australia infected various cultivars of Syngonium, other Araceae and Crotolaria, but not Platanus, Prunus spp., or Ipomoea (Vogelzang and Scott, 1990). Each host-specific type of C. fimbriata appears to have a distinct geographic distribution, although the total number of types and the geographic and host boundaries of each of them have not been fully determined.
Several recorded host plants for C. fimbriata are not included in the listing because they have not been confirmed. Some of these are probably erroneous reports, including the reports of C. fimbriata on soyabean (Glycine max), tobacco (Nicotiana species), potato (Solanum tuberosum), chestnut (Castanea sativa), cucumber (Cucumis sativa), kidney bean (Phaseolus vulgaris), coconut (Cocos nucifera), pineapple (Ananas comosus) and yam (Dioscorea species). There is also considerable confusion over the scientific and common names of edible members of the Araceae (for example, Xanthosoma, Colocasia and Alocasia), and it is not always clear which of these genera are referred to in the various reports.
Laboratory experiments have demonstrated C. fimbriata infection of Caladium, Dieffenbachia (Vogelzang and Scott, 1990) and several wild Ipomoea species (Clark and Watson, 1983) that have not been recorded as hosts in nature.
Host Plants and Other Plants AffectedTop of page
|Acacia decurrens (green wattle)||Fabaceae||Main|
|Acacia mangium (brown salwood)||Fabaceae||Main|
|Acacia mearnsii (black wattle)||Fabaceae||Main|
|Acrocarpus fraxinifolius (shingle tree)||Fabaceae||Other|
|Alocasia macrorrhizos (giant taro)||Araceae||Other|
|Annona squamosa (sugar apple)||Annonaceae||Main|
|Arracacia xanthorrhiza (arracacha)||Apiaceae||Other|
|Cajanus cajan (pigeon pea)||Fabaceae||Main|
|Carya cordiformis (bitternut hickory)||Juglandaceae||Main|
|Cassia javanica (pink shower)||Fabaceae||Main|
|Citrus aurantium (sour orange)||Rutaceae||Main|
|Coffea arabica (arabica coffee)||Rubiaceae||Main|
|Coffea canephora (robusta coffee)||Rubiaceae||Main|
|Colocasia esculenta (taro)||Araceae||Main|
|Crotalaria juncea (sunn hemp)||Fabaceae||Main|
|Daucus carota (carrot)||Apiaceae||Main|
|Eriobotrya japonica (loquat)||Rosaceae||Other|
|Ficus carica (common fig)||Moraceae||Main|
|Gmelina arborea (candahar)||Lamiaceae||Main|
|Hevea brasiliensis (rubber)||Euphorbiaceae||Main|
|Ipomoea batatas (sweet potato)||Convolvulaceae||Main|
|Lactuca sativa (lettuce)||Asteraceae||Main|
|Lactuca sativa (lettuce)||Asteraceae||Other|
|Mangifera indica (mango)||Anacardiaceae||Main|
|Manihot esculenta (cassava)||Euphorbiaceae||Main|
|Passiflora edulis (passionfruit)||Passifloraceae||Other|
|Pimenta dioica (allspice)||Myrtaceae||Main|
|Prunus (stone fruit)||Rosaceae||Main|
|Prunus armeniaca (apricot)||Rosaceae||Main|
|Prunus dulcis (almond)||Rosaceae||Main|
|Punica granatum (pomegranate)||Punicaceae||Main|
|Spathodea campanulata (African tulip tree)||Bignoniaceae||Main|
|Syngonium podophyllum (arrowhead vine)||Araceae||Main|
|Tectona grandis (teak)||Lamiaceae||Other|
|Theobroma cacao (cocoa)||Malvaceae||Main|
|Theobroma grandiflorum (cupuassu)||Malvaceae||Other|
Growth StagesTop of page Flowering stage, Fruiting stage, Post-harvest, Seedling stage, Vegetative growing stage
SymptomsTop of page C. fimbriata is primarily a xylem pathogen. On trees (Theobroma, Mangifera, Prunus, etc.), infection typically occurs through fresh wounds (Giraldo, 1957; Viégas, 1960; Moller et al., 1969), although root infections are also common (Ribeiro et al., 1986; Rossetto and Ribeiro, 1990; Laia et al., 2000). Mycelium and spores enter wounds and move through the xylem in water-conducting cells and into ray parenchyma cells. The fungus causes dark reddish-brown to purple to deep-brown or black staining in the xylem. This staining may extend several metres from the roots, up the trunk of the tree, and into branches. When affected branches or trunks are cut in cross-section, the staining along the rays gives a distinctive wedge-shaped or starburst-like pattern (Sinclair et al., 1987). On the surface of the trunk or branches, cankers may develop over areas of xylem discoloration, and the cankers may exude gum. Branch and trunk cankers are particularly common on Populus, Prunus, Platanus (Sinclair et al., 1987) and Eucalyptus (Laia et al., 2000), though wilting may also occur in the absence of canker development. Wilted leaves typically become dry and curled rather suddenly but remain attached to the tree for several weeks. On Platanus, individual leaves of affected branches often show interveinal chlorosis and necrosis, perhaps associated with fungal-produced phytotoxins (Ake et al., 1992; Alami et al., 1998; Pazzagli et al., 1999).
Infection of many trees (Theobroma, Mangifera, Punica and others) is often accompanied by secondary attack by various ambrosia beetles (such as Xyleborus and Hypocryphalus species), which bore into the xylem of the diseased trunk and produce copious amounts of frass (wood particles mixed with faeces) (Iton, 1959, 1960; Rossetto and de Medeiros, 1967; Somasekhara, 1999). Frass may cling to the gallery entrance holes in long strands or accumulate on the bark or at the base of the tree. Aleurioconidia may be present in such frass and may be an important source of inoculum. Frass containing C. fimbriata may be dispersed by wind or rainsplash.
On rubber trees (Hevea brasiliensis), C. fimbriata attacks the tapping panel, causing a pale-grey mould on the surface of the panel and dark discoloration in the wood under the surface (Martin, 1949; Silveira et al., 1994).
On herbaceous plants (Colocasia, Ipomoea, etc.), C. fimbriata may attack through wounded roots or stems, causing a root rot or seedling rot, or it can travel through the xylem, causing rapid wilting of the plant and extensive dark discoloration of the vascular system. It may also occur as a black, sunken rot on the surface of storage roots or corms of Ipomoea and Araceae such as Colocasia and Xanthosoma, either before or after harvest (Clark and Moyer, 1988).
The fungus has also been reported as a superficial pathogen of harvested cocoa pods, causing soft, brown, rotted lesions (Malaguti, 1958), especially during rainy periods (Siller, 1958). However, a related fungus, Ceratocystis paradoxa, is more common on rotten cocoa pods, most probably as a secondary invader (Thorold, 1975).
List of Symptoms/SignsTop of page
|Fruit / lesions: black or brown|
|Fruit / lesions: on pods|
|Leaves / abnormal colours|
|Leaves / necrotic areas|
|Leaves / wilting|
|Leaves / yellowed or dead|
|Roots / cortex with lesions|
|Stems / canker on woody stem|
|Stems / dieback|
|Stems / discoloration|
|Stems / discoloration of bark|
|Stems / gummosis or resinosis|
|Stems / internal discoloration|
|Stems / internal feeding|
|Stems / mould growth on lesion|
|Stems / mycelium present|
|Stems / ooze|
|Stems / visible frass|
|Stems / wilt|
|Whole plant / frass visible|
|Whole plant / plant dead; dieback|
|Whole plant / seedling blight|
|Whole plant / wilt|
Biology and EcologyTop of page Although outcrossing is possible, most isolates are self-fertile due to unidirectional mating type switching (Webster and Butler, 1967a, b; Harrington and McNew, 1997; Witthuhn et al., 2000). Fruiting bodies (perithecia) are produced from the mycelium in culture in about a week. The fungus may be dispersed as fragments of mycelium, conidia, aleurioconidia or ascospores. Aleurioconidia are probably the most common survival units because they are thick-walled and durable, and they probably facilitate survival in soil (Accordi, 1989) and in insect frass (Iton, 1960). The fungus may survive in wood fragments in river water (Grosclaude et al., 1991a) and in the soil (Accordi, 1989) for at least 3 months in the winter. C. fimbriata produces a strong fruity odour that varies with the medium. This has been assumed to be an adaptation for dispersal by insects, which are attracted to diseased plants and can become covered with sticky spores if the fungus is sporulating (see Means of Movement and Dispersal).
Wounds, either natural or from human activities, are important infection courts for all members of the genus Ceratocystis, including C. fimbriata. Inoculum may reach an open wound by being blown in the wind in insect frass (Iton, 1960) or by being carried by insects that visit the wound. Nitidulid beetles that feed on fungi and plant sap may be important vectors (Moller and DeVay, 1968b). Cultivation practices such as pruning may also provide infection courts (Teviotdale and Harper, 1991).
C. fimbriata usually grows best at temperatures from 18 to 28°C and is able to produce ascospores within a week. The fungus probably survives adverse conditions as mycelium within the plant host, or as aleurioconidia in the soil or in plant hosts or debris. The disease in Theobroma has been thought to be most severe during periods of abiotic stresses, particularly drought stress (Spence, 1958), or excessive rain (Malaguti, 1952a). On Ipomoea, attack by C. fimbriata may be enhanced by boron deficiency in the soil (Hu et al., 1999).
Means of Movement and DispersalTop of page Natural Dispersal
The fungus spreads readily between adjacent Platanus trees via root grafts (Accordi, 1986). It may also infect Platanus trees through wounds in the roots (Vigouroux and Stojadinovic, 1990). Mangifera trees may be infected through the roots from soilborne inoculum (Rossetto and Ribeiro, 1990), and root crops such as Ipomoea are commonly infected through wounds made by insects and rodents (Clark and Moyer, 1988). Ascopores are probably spread naturally by insects and are not likely to be airborne. Airborne dispersal of conidia is also not likely, except in insect frass. Rainsplash dispersal of conidia has not been documented.
Many Ceratocystis species produce fruiting bodies and fruity aromas that are believed to be adaptations for dispersal by insects, and C. fimbriata is frequently associated with insects. On Populus (Hinds, 1972b) and Prunus (Moller and DeVay, 1968b), circumstantial evidence suggests that fungal-feeding nitidulid beetles acquire the fungus and visit fresh wounds on susceptible trees. Also, spores of C. fimbriata may be carried upon the bodies of ambrosia beetles (Iton, 1966), and the spores can survive passage through an insect gut (Iton, 1960, 1966; Crone, 1963).
Ambrosia beetles (especially Xyleborus and Hypocryphalus species) are attracted to diseased plants (such as Theobroma, Mangifera and Eucalyptus) and produce large amounts of fine wood shavings (frass) when creating breeding galleries in the trunk and branches (Goitia and Rosales, 2001). These wood shavings and faecal material are pushed outside the tree as the galleries are excavated, and the frass contains spores and fragments of mycelium that may be blown in the wind (Iton, 1960).
No instances of spread of C. fimbriata on or with seed have been reported. However, one specimen in the US National Fungus Collections (BPI 596218) of an Erythrina seed pod has many fruiting bodies of C. fimbriata, suggesting that seedborne spread is possible.
Pruning wounds are common entry points for C. fimbriata, and the fungus can be carried on machetes or pruning tools (Walter, 1946, 1952; Teviotdale and Harper, 1991). Platanus street trees may become infected through pruning wounds, and the fungus may be spread on pruning tools or in wound dressings (Walter, 1946). Indeed, proper sanitation and disinfecting tools played a major role in stopping the epidemic on plane trees in urban areas of the eastern USA in the 1920s to 1940s (Walter, 1952). Infected wood and sawdust may harbour viable spores for at least 5 years (Grosclaude et al., 1995). On Theobroma, wounds made by harvesting pods, removing stem sprouts or weeding may become infected (Malaguti, 1958), and the fungus also infects pruning wounds and wounds made in harvesting almond fruit (Teviotdale and Harper, 1991).
As there may be extensive mycelial growth within a plant before symptoms appear, propagative cuttings may be an effective method of dispersal. Apparently healthy propagative cuttings of Populus were found to be infested with C. fimbriata (Vujanovic et al., 1999). BPI specimen 595645, of propagative material from Costa Rica intercepted in Miami, Florida, USA, contains several Manihot cuttings with abundant perithecia at the nodes. Infected Syngonium cuttings were the apparent means of spread of the Syngonium form of the fungus throughout the greenhouse industry (Walker et al., 1988). The fungus has also been found in both symptomatic and apparently healthy Eucalyptus cuttings in a Brazilian Eucalyptus plantation (CJ Baker, Iowa State University, USA, personal observation). Cuttings, roots and corms are used to propagate many other common hosts of C. fimbriata, including Theobroma, Ipomoea and Colocasia, and this may facilitate long-distance transport of the fungus. The Ipomoea form of the fungus, which is probably native to Latin America, is probably spread on storage roots (Bensaude, 1927; Baker et al., 2003).
Movement in Trade
It is apparent that several host-specialized forms of the fungus have been introduced into many regions. Propagative materials, especially cuttings, are a likely source. Packaging material and dunnage are also likely means of dispersal of the fungus. The Platanus form may have been introduced on packing material to Europe from North America during World War II (Panconesi, 1981, 1999) and has caused substantial damage to ornamental Platanus in southern Europe. This form can survive in Platanus wood taken from diseased trees (Grosclaude et al., 1995), which may be an efficient means of introducing the pathogen to new locations.
Pathway VectorsTop of page
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; spores||Yes||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Bulbs/Tubers/Corms/Rhizomes||fruiting bodies; hyphae; spores||Yes||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Fruits (inc. pods)||fruiting bodies; hyphae; spores||Yes||Pest or symptoms usually visible to the naked eye|
|Growing medium accompanying plants||fruiting bodies; hyphae; spores||Yes||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Leaves||fruiting bodies; hyphae; spores||Yes||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Roots||fruiting bodies; hyphae; spores||Yes||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Seedlings/Micropropagated plants||fruiting bodies; hyphae; spores||Yes||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Stems (above ground)/Shoots/Trunks/Branches||fruiting bodies; hyphae; spores||Yes||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Wood||fruiting bodies; hyphae; spores||Yes||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Plant parts not known to carry the pest in trade/transport|
|True seeds (inc. grain)|
Wood PackagingTop of page
|Wood Packaging liable to carry the pest in trade/transport||Timber type||Used as packing|
|Solid wood packing material with bark||Platanus spp.||Yes|
|Solid wood packing material without bark||Platanus spp.||Yes|
|Wood Packaging not known to carry the pest in trade/transport|
|Loose wood packing material|
|Processed or treated wood|
Impact SummaryTop of page
|Fisheries / aquaculture||None|
ImpactTop of page Diseases caused by C. fimbriata can be of high local importance, and there is a history of sporadic epidemics. The disease in Theobroma has been of major importance in Costa Rica (Echandi and Segall, 1956), Trinidad and Tobago (Iton, 1959), Ecuador (Desrosiers, 1957), parts of Colombia (Arbelaez, 1957) and Venezuela (Reyes, 1988), and most recently in Bahia, Brazil (Bezerra, 1997). In Theobroma plantations, the fungus has killed as many as 50% of the trees in some locations (Idrobo, 1958). The disease in Coffea is particularly important in Colombia (Pontis, 1951), where citrus is another major economic host (Borja et al., 1995). The disease in Mangifera in São Paulo, Brazil is of major importance (Oliveira, 1966; Ribeiro and Coral, 1968; Rossetto et al., 1969; Yamashiro and Myazaki, 1985; Rossetto and Ribeiro, 1990; Ribeiro et al., 1995). The fungus has also decimated certain clones of Eucalyptus in plantations in Brazil, and recent reports of the disease in Eucalyptus in the Congo and Uganda have indicated serious levels of mortality (Roux et al., 2000, 2001a). Almonds in California, USA, particularly in older orchards, have been seriously affected by the disease, especially after the initial introduction of mechanical shakers, which severely wounded the trees and led to more infections (DeVay et al., 1968). Platanus plantings in Italy, France and Switzerland are also seriously affected, and over 10% of the London plane trees in southern Switzerland have been killed since the early 1980s (Matasci and Gessler, 1997). More than 87% of plane trees (Platanus acerifolia) were lost during the period 1926-1949 in the community of Gloucester, New Jersey, the earliest recognized epidemic on plane tree in the USA (Walter et al., 1952). By 1952, they had estimated losses in excess of $1,000,000 (in 1952 dollars) in the north-east. Loss from Ceratocystis wilt on Punica in the Bijapur district of India from 1995 to 1998 was estimated at 7.5% of the crop (Somasekhara, 1999). Although damage from the Ipomoea form is now less severe in south-eastern USA than previously, mostly due to the use of resistant varieties and sanitary measures, it remains an important constraint to Ipomoea production in China and Japan (Clark and Moyer, 1988).
Environmental ImpactTop of page C. fimbriata is probably a natural component of many forest ecosystems in the Americas and Asia. On native tree hosts it primarily colonizes wounds but does not move throughout the tree or kill the host. Most mortality of woody hosts appears to be on exotic tree species or native trees in plantations or used as street trees, perhaps because of wounding and movement of the pathogen on tools. The plane tree pathogen, for instance, has been devastating on street trees but is rare in natural forests with little human activity (Walter et al., 1952). Even where the fungus has been introduced, the damage is primarily to planted species. Thus, the impact in natural environments has been minimal. However, some plantation species have been abandoned in some regions, such as Gmelina arborea in Pará state in Brazil and Platanus in the south-eastern USA.
Impact: BiodiversityTop of page There has been no clear impact of C. fimbriata on biodiversity.
Social ImpactTop of page Platanus species, especially P. acerifolia (London plane) is a very common street tree in many regions of the world, especially in the eastern USA and southern Europe. The loss of plane trees in Italy and southern France due to C. fimbriata has been dramatic, thus seriously reducing the aesthetics of urban areas. Earlier epidemics in urban areas of the eastern USA also had severe impact, though sanitation practices have greatly reduced the impact of the disease since the 1940s (Walter et al., 1952).
DiagnosisTop of page
Pure cultures of C. fimbriata may be obtained by placing chips of discoloured wood from the base of an infected tree or diseased vegetative plant parts in a moist chamber or plating them out on nutrient agar. When the fungus is present, conidia appear in 1-3 days and perithecia in 5-10 days. The presence of fast-growing contaminants, such as Fusarium and Penicillium, may necessitate the use of baits. The Platanus form may be baited from wood, soil or water samples with healthy Platanus twigs stripped of their bark (Grosclaude et al., 1988). All forms of the fungus may be baited from infected plant material by placing a small piece of colonized plant material between two slices of fresh carrot in high humidity for 4-10 days (Moller and DeVay, 1968a). Carrot slices may also be used to bait the fungus from soil (Laia et al., 2000), although carrot is not completely species-specific, allowing the growth of C. moniliformis, Thielaviopsis basicola (Yarwood, 1946), Fusarium spp. and some bacteria. The fungus can also be isolated from the frass of ambrosia beetles (Xyleborus and Hypocryphalus species) in Mangifera, Theobroma and Eucalyptus by using the carrot slice technique.
Molecular or serological diagnostic techniques have not been developed, but there are DNA sequences of ITS-rDNA and other genes unique to C. fimbriata and these could be developed for diagnosis.
A diagnostic protocol is detailed in OEPP (2003).
Detection and InspectionTop of page Disease caused by C. fimbriata may be visible on cuttings or other plant material as dark discoloration of the xylem, although symptomless cuttings may still be infected. Ascomata may also occasionally be produced on the surface of stem cuttings, particularly at the nodes. On Ipomoea storage roots and Araceae corms, the fungus may appear as a dry, black rot, usually with perithecia and ascospores. Incubation of colonized plant parts in a humid environment will usually result in ascomata production in only a few days. Unless perithecia are present on the infected plant, a pure culture of the fungus is usually required for reliable identification.
Similarities to Other Species/ConditionsTop of page C. fimbriata is usually recognized by its distinctive fruiting bodies, which are somewhat similar to those produced by other species of Ceratocystis and species of Ophiostoma. Ophiostoma species, in contrast to Ceratocystis, do not produce the endoconidial or aleurioconidial states. C. fimbriata has sometimes been confused with Ceratocystis paradoxa, a pathogen of mostly monocotyledonous plants. Both C. paradoxa and C. fimbriata may produce a pod rot of cocoa, although C. fimbriata can be distinguished by its hat-shaped ascospores (Hunt, 1956). Most forms of C. paradoxa are heterothallic, and isolates of this species usually do not produce perithecia unless paired with isolates of the opposite mating type.
On Theobroma trees, C. fimbriata may be confused with Ceratocystis moniliformis, which is weakly pathogenic, usually causing only partial wilting or wilting of only a few branches (Barba and Hansen, 1962). In the laboratory, C. moniliformis grows much more quickly on nutrient agar than does C. fimbriata, and when viewed under a compound microscope, the perithecial bases of C. moniliformis have characteristic spine-like ornamentations (Hunt, 1956). Also, C. moniliformis does not produce aleurioconidia. However, C. moniliformis produces hat-shaped ascospores similar to those of C. fimbriata.
Ceratocystis albofundus is morpholocially very similar to C. fimbriata but can be distinguished by its hyaline perithecial bases (Wingfield et al., 1996). Thus far, C. albofundus has only been reported from Africa (Roux et al., 2001b).
Infection by many other wilt-type fungi and species of Botryosphaeria may cause xylem discoloration in trees, and it is necessary to isolate C. fimbriata from the discoloured xylem or canker in order to confirm it as the causal agent.
Prevention and ControlTop of page
Due to the variable regulations around (de)registration of pesticides, your national list of registered pesticides or relevant authority should be consulted to determine which products are legally allowed for use in your country when considering chemical control. Pesticides should always be used in a lawful manner, consistent with the product's label.Host-Plant Resistance
Host-plant resistance has been used successfully with Mangifera (Ribeiro et al., 1984, 1986, 1995; Rossetto et al., 1997), Theobroma (Desrosiers, 1956; Delgado and Echandi, 1965; Gardella et al., 1982; Ocampo et al., 1982; Simmonds, 1994), Ipomoea (Martin, 1954), Coffea (Castilla, 1982) and Crotalaria (Ribeiro et al., 1977). Species and varieties of citrus also vary in susceptibility to Colombian strains of the fungus (Paez-Redondo and Castano-Zapata, 2001).
Cultural Control and Sanitary Measures
Sanitation is also effective for disease control. For example, disinfecting machetes and pruning tools between plants may help control the disease in Platanus (Walter, 1946; Walter et al., 1952) and Prunus (Teviotdale and Harper, 1991). Heat treatment of Ipomoea roots used in propagation has been suggested (Daines et al., 1962).
Fungicides are used with some success to treat tapping panels of Hevea (Chee, 1970) and in Ipomoea fields (Martin, 1971) or as post-harvest dips of Ipomoea roots (Daines, 1971; Yang et al., 2000). Fungicides injected into the stems of Platanus species may provide some protection (Causin et al., 1995; Minervini et al., 2001). Fungicides are also used to control the disease in Ficus (Hirota et al., 1984).
No biological control methods currently exist.
ReferencesTop of page
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