Ceratocystis fimbriata

The following information was compiled for the CABI Crop Protection Compendium, CABI Publishing. Updated 2001 by C.J. Baker and T. C. Harrington and by T. Harrington in 2004. This information was gathered in part through funding by the National Science Foundation (DEB-9870675 and DEB-0128104).

  • Taxonomy and Nomenclature
  • Host Range
  • Geographical Distribution
  • Biology and Ecology
  • Movement and Dispersal
  • Symptoms
  • Morphology
  • Detection Methods and Diagnostics
  • Similarities to Other Conditions
  • Impact
  • Control
  • Phytosanitary Risk
  • Pictures
  • References
  •  

    Taxonomy and Nomenclature

    PREFERRED SCIENTIFIC NAME: Ceratocystis fimbriata Ellis and Halsted

    CLASS: Pyrenomycetes

    ORDER: Microascales

    FAMILY: Ceratocystiaceae

     NON-PREFERRED SCIENTIFIC NAMES  TAXONOMIC AUTHORITY
     Ceratostomella fimbriata  (Ellis and Halsted) Elliott
     Endoconidiophora fimbriata  (Ellis and Halsted) Davidson
     Ophiostoma fimbriatum  (Ellis and Halsted) Nannf.
     Sphaeronema fimbriata  (Ellis and Halsted) Sacc.
     Rostrella coffea  Zimmerman
     Ceratocystis fimbriata f. platani  (Ellis and Halsted) Walter

    Common names of diseases

    English : Ceratocystis blight , blight of mango , wilt disease of cocoa , cacao wilt , mouldy rot of rubber , black rot of sweet potato , canker of coffee , Ceratostomella wilt, mango blight , black cane rot of Syngonium , black canker of aspen , black rot of sunn hemp , black rot of taro , canker stain of plane tree , Ceratocystis canker , Ceratocystis wilt , mallet canker , mango wilt , sweet potato black rot , target canker of aspen ,
    Spanish : llaga macana de cacao , mal de choroni de cacao , mal del machete de cacao , muerte subita de citrus , secamiento de los citricos , mal de machete , necrosis del tronco del cacao , marchitez de chupones: cacao ,
    French : chancre colore du platane , tache chancreuse , fletrissement du cacaoyer , pourriture des saignees de l'hevea ,
    Brazil : seca da mangueira ,

    Notes on taxonomy and nomenclature
    Ceratocystis fimbriata, the type species of the genus, was originally described on Ipomoea batatus (sweet potato) 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, C. fimbriata f. platani, based on its purported host specificity. 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 likely 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.

     HOST PLANTS AFFECTED BY C. FIMBRIATA
     Theobroma cacao (cacao)
     Mangifera indica (mango)
     Ipomoea batatas (sweet potato)
     Platanus acerifolia (London plane)
     Platanus orientalis (Oriental plane)
     Platanus occidentalis (American sycamore)
     Platanus racemosa (California sycamore)
     Coffea spp. (coffee)
     Eucalyptus spp.
     Citrus spp.
     Citrus aurantium (sour orange)
     Prunus dulcis (almond)
     Prunus armeniaca (apricot)
     Crotalaria juncea (sunn hemp)
     Hevea brasiliense (rubber)
     Gmelina arborea (melina, snapdragon tree)
     Pimenta officinalis (pimento)
     Punica granatum (pomegranate)
     Colocasia esculenta (taro, dasheen, inhame)
     Xanthosoma sp. (dasheen, malanga, cocoyam, yautia)
     Syngonium spp.
     Ficus carica (fig)
    Cajanus cajan (pigeon pea)
     Populus spp.
    Carya cordiformis (bitternut hickory)
      Carya ovata (shagbark hickory)
    Spathodea campanulata (African tulip tree)
     Cassia renigera
      Acacia decurrens
     Acacia mearnsii
     Alocasia sp. (giant taro)
     Annona
     Erythrina sp.
      Manihot esculenta (cassava)
     Herrania sp.

    Notes on host range
    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 allozyme 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 (Xanthosoma, Colocasia and Alocasia for example), 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.

    Geographical Distribution

     COUNTRY/REGION

     HOSTS AFFECTED

     REFERENCES
     ASIA    
     CHINA  Ipomoea, Punica, Colocasia  Sy, 1956, BPI specimens
     Fujian  Ipomoea  Hu et al., 1999
     Yunnan  Punica  Huang et al., 2003
     INDIA  Populus, Hevea Kaushik and Toky, 1992; Ramakrishnan and Radhakrishna, 1963
    Karnataka  Punica  Somasekhara, 1999
     Maharashtra  Punica  Somasekhara and Wali, 2000
     INDONESIA  Hevea Wright, 1925
    Java  Hevea, Coffea  Leefmans, 1934; South and Sharples, 1925; Zimmermann, 1900
     Kalimantan  Hevea  Tayler and Stephens, 1929
     Sumatra  Hevea  Tayler and Stephens, 1929
     JAPAN  Ipomoea, Colocasia  Asuyama, 1938; Okamoto, 1940; Shimizu, 1939
     Kyushu  Ficus, Ipomoea  Kajitani and Kudo, 1993; Kato et al., 1982;
     MALAYSIA  Hevea  Beeley, 1929; South and Sharples, 1925
     MYANMAR (BURMA)  Hevea  Turner and Myint, 1980
     TAIWAN  Crotalaria  Lee and Kuo, 1997
     AFRICA    
     CONGO  Eucalyptus  Roux et al., 2000
     COTE D'IVOIRE  Crotalaria  Davet, 1962
     KENYA Ipomoea Kihurani et al., 2000
     UGANDA  Eucalyptus  Roux et al., 2001
     SOUTH AFRICA  Acacia  Roux et al., 2000
     NORTH AND CENTRAL AMERICA    
     CANADA    
    British Columbia  Populus  Lowe, 1969, Hinds, 1985
     Manitoba  Populus  Zalasky, 1965
     Quebec  Populus  Vujanovic et al., 1999
     Saskatchewan  Populus  Zalasky, 1965
     Yukon Territory  Populus  Hinds, 1985
     COSTA RICA  Theobroma, Herrania, Coffea Baker et al., 2003; Echandi and Segall, 1956; Martin, 1949; Siller, 1958
     CUBA  Spathodea, Colocasia, Citrus  Isla and Ravelo 1989; Rodriguez and Alfonso 1978; Triana and Diaz, 1989;
     DOMINICAN REPUBLIC  Theobroma  Schieber, 1969
     GUATEMALA  Theobroma, Coffea, Hevea  Schieber and Sosa, 1960; Szkolnik, 1951; Tejada, 1983
     HAITI  Ipomoea  Barker, 1926
     JAMAICA  Pimenta  Leather, 1966
     MEXICO  Hevea, Erythrina  Martin, 1947; BPI specimens 596218, 595433
     ST VINCENT & GRENADINES  Ipomoea  BPI specimen 596219
     TRINIDAD & TOBAGO  Ipomoea, Theobroma  Baker, 1936; Baker and Dale, 1951; Briant, 1932; Iton, 1959; Leach, 1946
     UNITED STATES    
     Alaska  Populus  Hinds and Laurent, 1978
     Arkansas  Platanus  McCracken and Burkhardt, 1977
     Arizona  Populus  Hinds, 1972
     California  Syngonium, Platanus, Prunus  Davis, 1953; DeVay et al. 1968; Perry and McCain, 1988; Teviotdale and Harper 1991
     Colorado  Populus  Hinds, 1972
     Delaware  Platanus  Mook, 1940; Walter, 1946
     District of Columbia  Platanus  Walter et al., 1952
     Florida  Syngonium, Alocasia, Colocasia  Alfieri et al., 1994
     Hawaii  Syngonium, Colocasia  Uchida and Aragaki, 1979
     Idahao  Populus  Hinds, 1985
     Kentucky  Platanus  Mook, 1940
     Louisiana  Ipomoea  Baker et al., 2003; Webster and Butler, 1967
     Maryland  Platanus  Dodge, 1940
     Massachusetts  Ipomoea  BPI specimen 595868
     Minnesota   Populus  Hinds and Anderson, 1970; Wood and French, 1962
     Mississippi  Platanus  Walter, 1946
     Montana  Populus  Hinds, 1985
     Nevada  Populus  Hinds, 1985
     New Jersey  Platanus  Dodge, 1940; Walter, 1946
     New Mexico  Populus  Hinds, 1972
     New York  Platanus  Walter, 1946
     North Carolina  Platanus, Ipomoea  Baker et al., 2003; Walter, 1946
     North Dakota  Populus  Hinds, 1985
     Oregon  Populus  Hinds, 1985
     Pennsylvania  Platanus  Dodge, 1940; Jackson and Sleeth, 1935; Walter, 1946; Webster and Butler, 1967
     Rhode Island  Ipomoea  BPI specimen 595867
     Tennessee  Platanus  Mook, 1940; Walter, 1946
     Utah  Populus  Hinds, 1972
     Virginia  Platanus  Walter, 1946; Webster and Butler, 1967
     West Virginia  Platanus  Walter, 1946
     Wyoming  Populus  HInds, 1972
     SOUTH AMERICA    
     BRAZIL    
     Bahia  Theobroma, Hevea, Eucalyptus  Baker et al., 2003; Bezerra, 1997; Ferreira et al., 1999; Laia et al., 1999; Pereira and Santos, 1986
     Distrito Federal  Crotolaria  Melo-Filho et al., 2002
      Minas Gerais  Crotolaria  Chardon et al., 1940; Muller, 1937
     Pará Hevea, Gmelina, Acacia Albuquerque et al., 1972; Deslandes, 1944; Muchovej et al., 1978
     Pernambuco  Crotalaria, Mangifera, Coffea  Batista, 1947, 1960; Upadhyay, 1981
     Rio de Janeiro  Mangifera, Annona, Daucus  Cavalho and Carmo, 2003; Baker et al., 2003
     Rio Grande do Sul  Acacia  Santo and Ferreira, 2003
     Rondônia  Theobroma  Bastos and Evans, 1978
     São Paulo  Cassia, Crotolaria, Ficus, Hevea, Mangifera, Acacia  Arruda, 1940; Galli, 1958; Oliveira, 1966; Ribeiro et al., 1987; Ribeiro et al., 1988; Silveira et al. 1985; Valarini and Tokeshi, 1980
     COLOMBIA  Theobroma, Coffea, Citrus  Arbelaez, 1957; Borja et al., 1995; Garces, 1944; Marin et al., 2003; Mourichon, 1994; Pontis, 1951
     ECUADOR  Theobroma  Chalmers, 1969; Desrosiers, 1957; Desrosiers and Diaz, 1956; Rorer, 1918
     GUYANA  Theobroma  Bisessar, 1965
     PERU  Theobroma, Ipomoea  Krug and Quartey-Papafio, 1964; Rada, 1939; Soberanis et al., 1999
     SURINAME  Coffea  Baker et al., 2003
    URAGUAY  Eucalyptus  Barnes et al., 2003
     VENEZUELA  Coffea, Theobroma  Malaguti, 1952a, 1952b; Pontis, 1951; Reyes, 1988
     EUROPE    
     AZORES Ipomoea   Bensaude, 1927
     FRANCE  Platanus  Ferrari and Pechenot, 1979, 1974, 1976; Grosclaude et al., 1991; Vigouroux, 1986
     ITALY  Platanus  Pancohesi, 1981, 1999
     POLAND  Populus  Gremmen and deKam, 1976; Przybyl, 1980, 1986
     SWITZERLAND  Platanus  Matasci and Gessler1997
     OCEANIA    
     AUSTRALIA    
     New South Wales  Syngonium  Walker et al., 1988
     Queensland  Syngonium  Walker et al., 1988
    Victoria    Syngonium  Walker et al., 1988
     FIJI  Xanthosoma  Firman, 1972; Graham, 1965; Walker et al., 1988
     NEW ZEALAND  Ipomoea  Baker et al., 2003; Slade, 1960
     PAPUA NEW GUINEA  Ipomoea, Hevea  Baker et al., 2003; Mann, 1953
     WESTERN SAMOA  Colocasia  Walker et al., 1988

    Geographical distribution--further information

    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 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. Thus, many of the above reports have a question mark in the column designating exotic or native. For some of the 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 only cultivated plants in nurseries or greenhouses. Otherwise, the introduced strains are considered to be invasive populations.

    HISTORY OF INTRODUCTION / SPREAD

    The Populus form is most abundant in North America, but it has also appeared in Poland and perhaps India, most likely 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 Quebec nurseries (Vujanovic et al., 1999). The disease was severe in experimental plantings in Poland (Gremmen & 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 Pechenot, 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, 2001; 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 & Aragaki, 1979; Walker et al., 1988; Alfieri et al., 1994).

    The Ipomoea form of the fungus has likely 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).

    Biology and Ecology

    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 dispersal

    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 airborne. Airborne disperal of conidia is also not likely, except in insect frass. Rainsplash dispersal of conidia has not been documented.

    Vector Transmission

    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).

    Seedborne Spread

    No instances of its spread 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.

    Agricultural Practices

    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-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).


    Because there may be extensive mycelial growth within a plant before symptoms appear, propagative cuttings may be an effective method of dispersal. Healthy-appearing 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, 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 likely 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.

    Plant parts liable to carry the pest in trade/transport:
    - Bark: Spores, hyphae, fruit bodies; borne internally; borne externally; invisible to naked eye.
    - Bulbs/tubers/corms/rhizomes: Spores, hyphae, fruit bodies; borne internally; borne externally; invisible.
    - Fruits (inc. pods): Spores, hyphae, fruit bodies; borne externally; invisible to naked eye.
    - Growing medium accompanying plants: Spores, hyphae, fruit bodies; borne internally; borne externally; invisible.
    - Leaves: Spores, hyphae, fruit bodies; borne internally; borne externally; invisible to naked eye.
    - Seedlings/micropropagated plants: Spores, hyphae; borne internally; invisible.
    - Roots: Spores, hyphae, fruit bodies; borne internally; borne externally; invisible.
    - Stems (above ground)/shoots/trunks/branches: Spores, hyphae, fruit bodies; borne internally; borne externally; invisible.
    - Wood: Spores, hyphae, fruit bodies; borne internally; borne externally; invisible to naked eye.

    Plant parts not known to carry the pest in trade/transport:
    - Flowers/inflorescences/cones/calyx.

    Transport pathways for long distance movement:
    - Containers and packing: Wood used in packaging or dunnage.(Panconesi 1981; Grosclaude et al. 1995)
    - Soil, gravel, water, etc.: River water.(Grosclaude et al. 1991a)


    Symptoms-Description
    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 with 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, C. paradoxa, is more common on rotten cocoa pods, most likely as a secondary invader (Thorold, 1975).

    Descriptors: Whole plant: plant dead; dieback; seedling blight; frass visible; wilt. Leaves: necrotic areas; abnormal colours; wilting; yellowed or dead. Stems: discoloration of bark; canker on woody stem; gummosis or resinosis; dieback; mould growth on lesion; internal discoloration; internal feeding; visible frass; wilt; ooze; mycelium present; discoloration. Roots: cortex with lesions. Fruits/pods: lesions: black or brown; lesions: on pods.

    Click here to see pictures of symptoms caused by C. fimbriata

    Morphology

    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; Witthun 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.

    Click here to see pictures of C. fimbriata morphology

    Detection and inspection methods

    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.

    Diagnosis

    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.

    Similarities to other species or conditions

    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 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. fimbrata.


    Ceratocystis albofundus is morpholocially very similar to C. fimbriata but can be distinguished by its lightly pigmented perithecial bases (Wingfield et al., 1996). Thus far, C. albofundus has only been reported from Africa (Roux et al., 2001).

    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.


    Impact

    Economic Impact
    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, 2001). 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 the 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 Northeast. 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 southeastern 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 impact

    Ceratocystis fimbriata is likely 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 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 southeastern USA.

    Social impact
    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, southern France and Italy 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 greatly reduced the impact of the disease since the 1940s (Walter et al., 1952).


    Control

    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 (Castillo, 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, 1952) and Prunus (Teviotdale and Harper, 1991). Heat treatment of Ipomoea roots used in propagation has been suggested (Daines et al., 1962).


    Chemical Control
    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).


    Phytosanitary risk

    Because most forms of the species 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 up to 5 years in wood, probably in the form of aleurioconidia. Ceratocystis fimbriata is listed as among the highest risk pathogens that could be imported into the United States 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).

    References

    Back to top

    To Tom Harrington's Ceratocystis page