Invasive Species Compendium

Detailed coverage of invasive species threatening livelihoods and the environment worldwide

Datasheet

Claviceps africana
(ergot)

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Datasheet

Claviceps africana (ergot)

Summary

  • Last modified
  • 27 September 2018
  • Datasheet Type(s)
  • Invasive Species
  • Pest
  • Preferred Scientific Name
  • Claviceps africana
  • Preferred Common Name
  • ergot
  • Taxonomic Tree
  • Domain: Eukaryota
  •   Kingdom: Fungi
  •     Phylum: Ascomycota
  •       Subphylum: Pezizomycotina
  •         Class: Sordariomycetes
  • Summary of Invasiveness
  • C. africana is the ergot pathogen of Sorghum bicolor, now found in most sorghum-producing areas of the world. It is primarily a problem for hybrid seed production, but epidemics on local varieties have occ...

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Pictures

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PictureTitleCaptionCopyright
Sphacelia of C. africana appear in advance of spore-bearing honeydew.
TitleSphacelia
CaptionSphacelia of C. africana appear in advance of spore-bearing honeydew.
CopyrightD.E. Frederickson, INTSORMIL, Zimbabwe
Sphacelia of C. africana appear in advance of spore-bearing honeydew.
SphaceliaSphacelia of C. africana appear in advance of spore-bearing honeydew.D.E. Frederickson, INTSORMIL, Zimbabwe
Profuse honeydew exudation and secondary conidiation often renders infected panicles spectacularly white. Wherever honeydew dries a stiff crust is formed.
TitleHoneydew
CaptionProfuse honeydew exudation and secondary conidiation often renders infected panicles spectacularly white. Wherever honeydew dries a stiff crust is formed.
CopyrightD.E. Frederickson, INTSORMIL, Zimbabwe
Profuse honeydew exudation and secondary conidiation often renders infected panicles spectacularly white. Wherever honeydew dries a stiff crust is formed.
HoneydewProfuse honeydew exudation and secondary conidiation often renders infected panicles spectacularly white. Wherever honeydew dries a stiff crust is formed.D.E. Frederickson, INTSORMIL, Zimbabwe
Honeydew oozing from sorghum florets infected with C. africana. Note the white droplet surfaces, indicative of secondary conidiation.
TitleHoneydew
CaptionHoneydew oozing from sorghum florets infected with C. africana. Note the white droplet surfaces, indicative of secondary conidiation.
CopyrightD.E. Frederickson, INTSORMIL, Zimbabwe
Honeydew oozing from sorghum florets infected with C. africana. Note the white droplet surfaces, indicative of secondary conidiation.
HoneydewHoneydew oozing from sorghum florets infected with C. africana. Note the white droplet surfaces, indicative of secondary conidiation.D.E. Frederickson, INTSORMIL, Zimbabwe
Parasitic bodies of C. africana with variable amounts of sclerotial tissue forming towards the base.
TitleParasitic bodies
CaptionParasitic bodies of C. africana with variable amounts of sclerotial tissue forming towards the base.
CopyrightD.E. Frederickson, INTSORMIL, Zimbabwe
Parasitic bodies of C. africana with variable amounts of sclerotial tissue forming towards the base.
Parasitic bodiesParasitic bodies of C. africana with variable amounts of sclerotial tissue forming towards the base.D.E. Frederickson, INTSORMIL, Zimbabwe
Parasitic bodies of C. africana: Parasitic bodies of C. africana from Texas, USA. The orange-brown portion of each is composed of sclerotial tissues whilst the upper part is residual sphacelial tissue.
TitleParasitic bodies
CaptionParasitic bodies of C. africana: Parasitic bodies of C. africana from Texas, USA. The orange-brown portion of each is composed of sclerotial tissues whilst the upper part is residual sphacelial tissue.
CopyrightD.E. Frederickson, INTSORMIL, Bulawayo, Zimbabwe
Parasitic bodies of C. africana: Parasitic bodies of C. africana from Texas, USA. The orange-brown portion of each is composed of sclerotial tissues whilst the upper part is residual sphacelial tissue.
Parasitic bodiesParasitic bodies of C. africana: Parasitic bodies of C. africana from Texas, USA. The orange-brown portion of each is composed of sclerotial tissues whilst the upper part is residual sphacelial tissue.D.E. Frederickson, INTSORMIL, Bulawayo, Zimbabwe
Teleomorph of C. africana, the purple-pigmented stipe and capitulum are diagnostic of the species.
TitleTeleomorph
CaptionTeleomorph of C. africana, the purple-pigmented stipe and capitulum are diagnostic of the species.
CopyrightMycological Research, 95:1101-1107
Teleomorph of C. africana, the purple-pigmented stipe and capitulum are diagnostic of the species.
TeleomorphTeleomorph of C. africana, the purple-pigmented stipe and capitulum are diagnostic of the species.Mycological Research, 95:1101-1107
Macroconidia (C) of C. africana, many of which have germinated to produce a hyphal process (H) and secondary conidium (SC). (N) indicates a nucleus.
TitleMacroconidia
CaptionMacroconidia (C) of C. africana, many of which have germinated to produce a hyphal process (H) and secondary conidium (SC). (N) indicates a nucleus.
CopyrightD.E. Frederickson, INTSORMIL, Zimbabwe
Macroconidia (C) of C. africana, many of which have germinated to produce a hyphal process (H) and secondary conidium (SC). (N) indicates a nucleus.
MacroconidiaMacroconidia (C) of C. africana, many of which have germinated to produce a hyphal process (H) and secondary conidium (SC). (N) indicates a nucleus.D.E. Frederickson, INTSORMIL, Zimbabwe

Identity

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Preferred Scientific Name

  • Claviceps africana Frederickson, Mantle & De Milliano 1991

Preferred Common Name

  • ergot

Other Scientific Names

  • Sphacelia sorghi McRae

International Common Names

  • English: sorghum ergot; sugary disease

Local Common Names

  • India: Asali

EPPO code

  • CLAVAF (Claviceps africana)

Summary of Invasiveness

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C. africana is the ergot pathogen of Sorghum bicolor, now found in most sorghum-producing areas of the world. It is primarily a problem for hybrid seed production, but epidemics on local varieties have occurred (Pazoutová and Frederickson, 2005). Large numbers of secondary conidia produced on infected panicles become airborne and are presumed to be the means by which the fungus has spread rapidly across continents in recent years. The fungus may also be carried in the form of sclerotia and/or sphacelia among harvested seed, and this may be the means of spread between continents, but the seed lots can be cleaned or treated with fungicides. Alternative hosts are predominantly wild and weedy Sorghum spp., but some wild grasses can become infected; any of these might provide a reservoir between planting seasons or a bridge between regions.

Taxonomic Tree

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  • Domain: Eukaryota
  •     Kingdom: Fungi
  •         Phylum: Ascomycota
  •             Subphylum: Pezizomycotina
  •                 Class: Sordariomycetes
  •                     Subclass: Hypocreomycetidae
  •                         Order: Hypocreales
  •                             Family: Clavicipitaceae
  •                                 Genus: Claviceps
  •                                     Species: Claviceps africana

Notes on Taxonomy and Nomenclature

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When sorghum ergot was first found in Kenya in 1924 (specimen in the IMI Herbarium, c/o CABI Bioscience, Egham, UK; accession number 93464; IMI Herbarium, undated), the imperfect stage of a pathogen in India had already been described (McRae, 1917). As a consequence, the African pathogen became known by the same name, Sphacelia sorghi. In 1976, Kulkarni et al. described the perfect stage of an Indian pathogen as Claviceps sorghi with the similar result that ergot in Africa was then sometimes additionally referred to as C. sorghi. However, C. sorghi does not appear to exist in Africa. Any records of C. sorghi in Africa date from the period 1976-1991, when the existence of two distinct species was unknown. With the generation of the perfect stage of a Zimbabwean pathogen in 1991 (Frederickson et al., 1991), it became clear that the African pathogen was a new species entirely, which was named C. africana.

Description

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Sphacelia white, subglobose, 4-6 x 2-3 mm, forming two types of spore: macroconidia oblong, hyaline, with polar vacuoles and a slight central constriction, 9-17 x 5-8 µm; microconidia spherical, 2-3 µm diameter. Germination of the macroconidium produces the pyriform secondary conidium, 8-14 x 4.0-6.5 µm on the tip of the sterigma-like process (Frederickson et al., 1989; 1991; Pazoutová et al., 2004). The secondary conidia are capable of two or three iterations, as well as of forming vegetative mycelium on a suitable medium (Pazoutová et al., 2004).

The structure popularly called the sclerotium is actually comprised of both sphacelial and sclerotial tissues and is similar in size and shape to the initial sphacelium. The true sclerotial tissues are proximal, spherical to oval in shape and, in contrast to the sphacelium, are firm (even after soaking in water), hydrophobic and contain the alkaloid dihydroergosine. The sclerotial cortex is orange-brown, but may appear superficially pink, orange or red due to adherent floral membranes. Germination of the sclerotium gives rise to up to six stromata each with a purple stipe, 8-15 x 0.3-0.6 mm long, and purple capitulum (0.5-1.3 mm diameter). Ascomata (perithecia), ovate-pyriform, 123-226 x 86-135 µm, asci cylindrical 140 x 3-4 µm: ascospores eight, filiform, septate, 45 x ca. 1 µm (Frederickson et al., 1991).

Distribution

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In 1991, ergot in Zimbabwe was described as a new species, C. africana, quite distinct from Claviceps sorghi (Frederickson et al., 1991). Ergot on sorghum worldwide had been assumed to be C. sorghi prior to this. Earlier reports of C. sorghi in Africa are erroneous and are considered to refer to C. africana.

Bandyopadhyay (ICRISAT, India, personal communication, 1998) recorded the presence of C. africana in Burkina Faso.

When, in 1998, C. africana was confirmed to be present in India (Bogo and Mantle, 1999; Pazoutová et al., 2000) and it was realised that since the late 1980s all ergot samples from India had probably been of C. africana, it was hypothesized that C. africana may have replaced or marginalized C. sorghi (DE Frederickson, INTSORMIL, Box 776, Bulawayo, Zimbabwe, personal communication, 1999). The earlier report of sorghum ergot in India of Kulkarni et al. (1976) describes both the elongate sclerotia of C. sorghi and the small, subglobose sclerotia of C. africana indicating that C. africana was present in 1976. Bandyopadhyay (ICRISAT, India, personal communication, 1998) reported C. africana in Maharashtra, Karnataka, Andhra Pradesh, Gujarat, Tamil Nadu and Uttar Pradesh states in India. Confirmation of the presence and distribution of C. sorghi, and distribution of C. africana, were the objectives of a survey in India (Bandyopadhyay, ICRISAT, India, 2000).

Ergot of sorghum was recorded in Thailand in 1983 (Boon-Long, 1992) and, in 1991, Frederickson et al. (1991) identified the pathogen as C. africana; the first clear record of the pathogen outside Africa. In 1994, C. africana was identified in southern Japan (Mantle and Hassan, 1994). In 1995, the widespread appearance of C. africana outside Africa began in Brazil (Reis et al., 1996) and soon followed in other parts of South and Central America (Bandyopadhyay et al., 1998) and, in 1996, in Queensland, Australia (Ryley et al., 1996). By March 1997, the fungus crossed the Rio Grande valley between Mexico and the USA into Texas (Isakeit et al., 1998) and outbreaks in other states quickly followed (Odvody et al., 1998; Zummo et al., 1998).

Pazoutová et al. (2000) used DNA-based marker methods to compare C. africana isolates from across the globe. RAPD banding patterns of pathogens from the USA, Mexico, Puerto Rico, Bolivia, India and Australia were evaluated with nearly 100 primers. The American isolates and three isolates from South Africa were identical, suggesting Africa as the origin of the American clones. C. africana isolates from Australia and India separated into another group. Thus two strains of C. africana were recognized, designated East (India/Australia) and West (Africa/Americas).

The distribution map includes records based on specimens of C. africana from the herb. IMI (c/o CABI UK, Egham -- IMI Herbarium, undated). Whereas most major sorghum-producing countries are represented in reports of C. africana, no data were found concerning its presence or absence in other countries where sorghum is a significant crop, including China and Russia.

Distribution Table

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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/RegionDistributionLast ReportedOriginFirst ReportedInvasiveReferenceNotes

Asia

IndiaRestricted distributionIntroduced Invasive Bogo and Mantle, 1999; CABI/EPPO, 2006; EPPO, 2014
-Andhra PradeshPresentIntroduced Invasive Bogo and Mantle, 1999; CABI/EPPO, 2006; EPPO, 2014
-GujaratPresentIntroduced Invasive CABI/EPPO, 2006; EPPO, 2014
-KarnatakaPresentIntroduced Invasive Venkateshwaran et al., 2006
-Madhya PradeshPresentIntroduced Invasive CABI/EPPO, 2006; EPPO, 2014
-RajasthanPresentIntroduced Invasive Venkateshwaran et al., 2006
-Tamil NaduPresentIntroduced Invasive CABI/EPPO, 2006; EPPO, 2014
-Uttar PradeshPresentIntroduced Invasive Venkateshwaran et al., 2006
JapanPresentIntroduced Invasive Tsukiboshi and, 1999; CABI/EPPO, 2006; EPPO, 2014
-HonshuPresentIntroduced Invasive CABI/EPPO, 2006; EPPO, 2014
-KyushuPresentIntroduced Invasive Mantle and Hassan, 1994; Tsukiboshi and, 1999; CABI/EPPO, 2006; EPPO, 2014
ThailandPresentIntroduced Invasive Frederickson et al., 1991; CABI/EPPO, 2006; EPPO, 2014
YemenPresentIntroduced1980s Invasive CABI/EPPO, 2006; EPPO, 2014

Africa

AngolaPresentIntroduced1980sDe Milliano et al., 1991; Bandyopadhyay et al., 1998; CABI/EPPO, 2006; EPPO, 2014
BotswanaPresentIMI Herbarium, undated; Molefe, 1975; CABI/EPPO, 2006; EPPO, 2014
Burkina FasoPresentBandyopadhyay et al., 1998
BurundiPresentIntroduced1980sBandyopadhyay et al., 1998; CABI/EPPO, 2006; EPPO, 2014
EthiopiaPresentIMI Herbarium, undated; Hulluka and Gebrekidan, 1980; Hulluka, 1982; Frederickson et al., 1994; CABI/EPPO, 2006; EPPO, 2014
GhanaPresentIntroduced1980sIMI Herbarium, undated; CABI/EPPO, 2006; EPPO, 2014
KenyaPresentIntroduced1980sIMI Herbarium, undated; CABI/EPPO, 2006; EPPO, 2014
LesothoPresentIntroduced1980sDe Milliano et al., 1991; CABI/EPPO, 2006; EPPO, 2014
MalawiPresentIntroduced1980sDe Milliano et al., 1991; CABI/EPPO, 2006; EPPO, 2014
MozambiquePresentIntroduced1980sPlumb-Dhindsa and Mondjane, 1984; De Milliano et al., 1991; CABI/EPPO, 2006; EPPO, 2014
NigeriaPresentFutrell and Webster, 1965; Futrell and Webster, 1966; Mower et al., 1973; CABI/EPPO, 2006; EPPO, 2014
RwandaPresentFrederickson et al., 1994; CABI/EPPO, 2006; EPPO, 2014
SenegalPresentCABI/EPPO, 2006; EPPO, 2014
South AfricaPresentDoidge et al., 1953; De Milliano et al., 1991; McLaren, 1994; CABI/EPPO, 2006; EPPO, 2014
SudanPresentPazoutová and Frederickson, 2005
SwazilandPresentDe Milliano et al., 1991; CABI/EPPO, 2006; EPPO, 2014
TanzaniaPresentIntroduced1980sWallace and Wallace, 1949; De Milliano et al., 1991; Mansuetus, 1995; CABI/EPPO, 2006; EPPO, 2014
UgandaPresentIntroduced1980sIMI Herbarium, undated; CABI/EPPO, 2006; EPPO, 2014
ZambiaPresentIMI Herbarium, undated; Angus, 1965; Frederickson, 1990; De Milliano et al., 1991; CABI/EPPO, 2006; EPPO, 2014
ZimbabwePresentIMI Herbarium, undated; Frederickson, 1990; Frederickson et al., 1991; Frederickson et al., 1993; Frederickson et al., 1994; Mantle and Hassan, 1994; CABI/EPPO, 2006; EPPO, 2014Samples obtained from Matopos, Bulawayo, Hunters Road, Kwe Kwe, Harare, Mazoe, Gwebi and Shamva

North America

MexicoPresentIntroduced1997Torres and Montez, 1997; Bandyopadhyay et al., 1998; Valasquez-Valle et al., 1998; CABI/EPPO, 2006; EPPO, 2014
USARestricted distributionIntroduced1997Alderman et al., 2004; CABI/EPPO, 2006; EPPO, 2014
-ArkansasRestricted distributionIntroduced2000Ross et al., 2002
-FloridaPresentIntroducedMiller, 1997; Bandyopadhyay et al., 1998; CABI/EPPO, 2006; EPPO, 2014
-GeorgiaPresentIntroduced1997Bandyopadhyay et al., 1998; Odvody et al., 1998; Alderman et al., 2004; CABI/EPPO, 2006; EPPO, 2014
-KansasPresentIntroduced1997Bandyopadhyay et al., 1998; Claflin, 1998; Alderman et al., 2004; CABI/EPPO, 2006; EPPO, 2014
-MississippiPresentIntroduced1998Bandyopadhyay et al., 1998; Zummo et al., 1998; Alderman et al., 2004; CABI/EPPO, 2006; EPPO, 2014
-NebraskaPresentIntroduced1997Bandyopadhyay et al., 1998; Odvody et al., 1998; CABI/EPPO, 2006; EPPO, 2014
-OklahomaPresentIntroducedCABI/EPPO, 2006; EPPO, 2014
-TexasPresentIntroduced1997Bandyopadhyay et al., 1998; Isakeit et al., 1998; Alderman et al., 2004; EPPO, 2014

Central America and Caribbean

Dominican RepublicPresentIntroduced1997Bandyopadhyay et al., 1998; CABI/EPPO, 2006; EPPO, 2014
HaitiPresentIntroduced1997CABI/EPPO, 2006; EPPO, 2014
HondurasPresentIntroduced1996Bandyopadhyay et al., 1998; CABI/EPPO, 2006; EPPO, 2014
JamaicaPresentIntroduced1997Bandyopadhyay et al., 1998; CABI/EPPO, 2006; EPPO, 2014
Netherlands AntillesPresentEPPO, 2014
Puerto RicoPresentIntroduced1997Bandyopadhyay et al., 1998; CABI/EPPO, 2006; EPPO, 2014

South America

ArgentinaPresentIntroduced1996Bandyopadhyay et al., 1998; CABI/EPPO, 2006; EPPO, 2014
BoliviaPresentIntroduced1996Bandyopadhyay et al., 1998; CABI/EPPO, 2006; EPPO, 2014
BrazilPresentIntroduced1995De Almeida Pinto et al., 1997; Bandyopadhyay et al., 1998; CABI/EPPO, 2006; EPPO, 2014
-GoiasPresentIntroduced1995Reis et al., 1996; CABI/EPPO, 2006; EPPO, 2014
-Minas GeraisPresentIntroduced1995Reis et al., 1996; CABI/EPPO, 2006; EPPO, 2014
-Santa CatarinaPresentIntroducedBogo and Boff, 1997; CABI/EPPO, 2006; EPPO, 2014
-Sao PauloPresentIntroduced1995Reis et al., 1996; CABI/EPPO, 2006; EPPO, 2014
ColombiaPresentIntroduced1996Varon de Agudelo et al., 1996; Bandyopadhyay et al., 1998; CABI/EPPO, 2006; EPPO, 2014
ParaguayPresentIntroduced1996Bandyopadhyay et al., 1998; CABI/EPPO, 2006; EPPO, 2014
UruguayPresentIntroduced1996Bandyopadhyay et al., 1998; CABI/EPPO, 2006; EPPO, 2014
VenezuelaPresentIntroduced1996Bandyopadhyay et al., 1998; CABI/EPPO, 2006; EPPO, 2014

Oceania

AustraliaRestricted distributionIntroduced1996CABI/EPPO, 2006; EPPO, 2014
-New South WalesPresentIntroduced1996CABI/EPPO, 2006; EPPO, 2014
-QueenslandPresentIntroduced1996Ryley et al., 1996; Bandyopadhyay et al., 1998; CABI/EPPO, 2006; EPPO, 2014

Risk of Introduction

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Risk Criteria Category

Economic Importance - moderate
Distribution - widespread in all major sorghum-growing countries
Seedborne Incidence - moderate
Seed Transmitted - not recorded
Seed Treatment - yes

Overall Risk - low to moderate

(Caution needed to avoid the introduction of 'East strains' to America and Africa.)

Phytosanitary Risk

Recent experiences in the USA and Australia prove that quarantine has limited impact on preventing ergot introductions and spread of C. africana in the face of windborne spread (Bandyopadhyay et al., 1998). Airborne secondary conidia are by far the most important means of disease transmission over a contiguous land mass (Frederickson et al., 1989; 1993; Bandyopadhyay et al., 1998). In Texas, USA, and Mexico, ergot is annually observed on Johnson grass (Sorghum halepense) and sorghum regrowth in winter fields. Even if only a proportion of inoculum survives to summer, and conidia survive several months (Claflin, 1998) there will be enough airborne inoculum to initiate new season's infections within and across international borders. Quarantine of seed is thus futile for controlling the spread of C. africana once the disease is present in a region. Ergot is now present in all the major sorghum-growing countries of the world. However, the existence of two strains of C. africana (Pazoutová et al., 2000), one (east strain) confined to India/Australia and the other (west strain) to Africa/America suggests caution in the exchange of seed between these areas. Toxigenic effects of the ingestion of sclerotia of the east strain have been reported in cattle, pigs and poultry (Bandyopadhyay et al., 1998). It may, therefore, be important to maintain the exclusion of the east strain from the Americas and Africa to protect the animal feed industry. See also 'Biology and Ecology'.

Habitat List

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CategoryHabitatPresenceStatus
Terrestrial-managed
Cultivated / agricultural land Present, no further details Harmful (pest or invasive)
Managed grasslands (grazing systems) Present, no further details Harmful (pest or invasive)
Terrestrial-natural/semi-natural
Natural grasslands Present, no further details Harmful (pest or invasive)

Hosts/Species Affected

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Pennisetum glaucum (pearl millet) has been artificially infected by C. africana (Frederickson and Mantle, 1996), but is not expected to be an important secondary host in nature. Reports of other grass hosts in the literature are conflicting and it is recommended that alternative hosts are investigated by cross-inoculation tests following Koch's postulates (Frederickson, 1999). In general, only sorghum species appear to be infected naturally. Artificial inoculations of a number of grass species with 106 conidia, resulted in ergot infection of only Sorghum arundinaceum, Sorghum halepense, Sorghum versicolor, Sorghum virgatum [Sorghum bicolor subsp. verticilliflorum], and P. glaucum (Muthusubramanian et al., 2005). However, in Mexico, S. halepense, Cenchrus echinatus and Panicum maximum [Megathyrsus maximus] were shown to be hosts for local isolates by inoculation from cultivated sorghum to grass and back to sorghum (Montes-Belmont et al., 2002a).

Individual florets of the inflorescence are affected by sphacelia/sclerotia and honeydew; the panicle, seeds, leaves and stalks are affected (but not infected) by honeydew.

Host Plants and Other Plants Affected

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Plant nameFamilyContext
SorghumPoaceaeWild host
Sorghum bicolor (sorghum)PoaceaeMain
Sorghum halepense (Johnson grass)PoaceaeOther

Growth Stages

Top of page Flowering stage

Symptoms

Top of page Individual ovaries between the glumes of some or all sorghum florets are replaced by a soft, white, subglobose-shaped growth of mycelium (sphacelium) from which sticky, liquid droplets of spore-bearing honeydew (thin to viscous, orange-brown or superficially white) may exude. Under conditions of high relative humidity, the copious honeydew is of low viscosity and the surface white. The surfaces of the panicle, seed, leaves, stalk and soil also become smeared by the dripping honeydew and appear conspicuously white. A white, powdery crust forms wherever such honeydew dries. For more information, see Frederickson et al. (1989; 1991).

When the honeydew and sphacelia are colonized by the hyperparasite, Cerebella andropogonis, black, spherical, convoluted growths are seen at floret tips (Bandyopadhyay et al., 1998). Upon dissection, a discoloured sphacelium of reduced size is found underneath. Other moulds may also grow on the honeydew.

List of Symptoms/Signs

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SignLife StagesType
Inflorescence / honeydew or sooty mould
Leaves / honeydew or sooty mould
Stems / honeydew or sooty mould

Biology and Ecology

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Most details of the life cycle focus on the macroconidia because the secondary conidia, although of greater epidemiological significance (Frederickson et al., 1989; 1993) are difficult to isolate. The path of infection of C. africana on sorghum is similar to that of C. sorghi, although the timing of infection events differs. Macroconidia germinate on the stigmatic hairs of unpollinated ovaries after 12-16 h, the germ tube penetrates the stigmatic surface and grows down the transmission tracts of the style into the top of the ovary. Germination occurs in the range 14-32°C, optimum 20°C (Frederickson, 1990). Hyphae grow down the inner ovary wall to the base, occupying the tissues adjacent to the rachilla, which provides the necessary nutrients to support the massive proliferation of hyphae. By day 4 the ovule has been invaded and hyphae simultaneously emerge onto the ovary surface. All but the very apex of the ovary is colonized. The incubation period before disease symptoms appear (formation of the sphacelium) is 6-12 days (Frederickson, 1990); the optimum temperature for disease development is 20°C (McLaren and Wehner, 1990).

Spores, in honeydew, are exuded about a day after the sphacelia become obvious (c.f. C. sorghi where honeydew appears long before the sphacelia are visible). Under conditions of high relative humidity, macroconidia at the honeydew surface germinate iteratively to produce aerially-supported secondary conidia, rendering the honeydew white in appearance (Frederickson et al., 1989; 1993; Bandyopadhyay et al., 1990). Secondary conidiation is a prominent feature of C. africana epiphytotics across the world. Secondary conidia are wind-borne spores. In field trials in Zimbabwe, C. africana spread rapidly through replicated plots of male-sterile sorghum A-lines from a group of centrally situated, inoculated plants. Prominent secondary conidiation by the pathogen on the surface of exuded honeydew provided airborne spores, which were trapped in a Burkard continuous spore trap and showed diurnal peaks in air close to the primary inoculum source. From the rate and pattern of disease spread it was concluded that secondary conidia of C. africana were the principal disease agents within the experimental area, and that ergot spread by windborne secondary conidia has significant epidemiological and economic implications for sorghum hybrid breeding in southern Africa (Frederickson et al., 1993). Secondary conidia were responsible for explosive disease development and, by inference, for disease up to 300 m away (Frederickson et al., 1989; 1993). Although they are present throughout the day in Zimbabwe, there is a sharp rise in the incidence of secondary conidia at nightfall (Frederickson et al., 1993). Hyaline secondary conidia are too fragile to survive the negative effects of high temperatures, relative humidity and ultraviolet light to account for intercontinental disease transmission, but are almost certainly responsible for disease spread over moderate distances across contiguous land masses, for example, in the Americas (DE Frederickson, INTSORMIL, Box 776, Bulawayo, Zimbabwe, personal communication, 1999). Autoinfection and many secondary cycles are possible in a season, even from a small initial inoculum. Inoculum may thus rapidly build up and spread to cause high disease incidence and severity. Chakraborty and Ryley (2008) found that new infections could occur even 1 day after sources of inoculum had been removed from plots, apparently due to persistently airborne secondary conidia.

Ergot pathogens only infect and colonize unfertilized ovaries. There is, therefore, a strong correlation between unpollinated flowers and ergot infection (Bandyopadhyay et al., 1998). Rapid anthesis lowers the chance of infection and any factor prolonging the period from floret opening to fertilization will promote ergot infection (Futrell and Webster, 1965; Puranik et al., 1973). For each day of delay in fertilization after inoculation, Musabyimana et al. (1995) recorded an 8.3% increase in ergot severity.

At the end of the cropping season the pathogen exists in the form of sphacelia and sclerotia and either may enable survival. Sclerotia are not separate entities, but form from within sphacelia and carry a considerable residual sphacelial portion. Sphacelia, however, may have no sclerotial tissues to fully-developed sclerotial tissues associated with them (Frederickson et al., 1999). Bandyopadhay et al. (1990) observed that lower humidity and higher temperatures favoured the production of sclerotia. Use of the term sphacelia/sclerotia is recommended (Alderman et al., 1999; Frederickson et al., 1999) to help eliminate the expectation of sclerotia appearing as discrete structures from sphacelia.

Macroconidia dry-stored in sphacelia/sclerotia for 7-12 months in Nigeria, Botswana and Zimbabwe (Futrell and Webster, 1966; Mower et al., 1973; Frederickson et al., 1991; 1993) have been shown experimentally to initiate infections with low severity following inoculation. In Kansas, USA, conidia applied to rubber, leather and metal were able to survive from November until temperatures rose in April in an unheated, outdoor storeroom (Claflin, 1998). Conidia showed poor survival on paper. However, in normal seed store conditions, conidial viability would decline to zero after about 4 months (Odvody et al., 1999). Low temperatures and low moisture content, which conditions prevail in the 'winter' or 'off-season' in Zimbabwe, promote the survival of conidia in sphacelia through winter (Odvody et al., 1999). Honeydew on seed is one potential source of infection, which can be eliminated by seed treatment with captan (Dahlberg et al., 1999). Although there are deep recesses in sphacelia, the efficacy of chemical seed treatment, which is essentially superficial, at reducing germination of conidia diffusing from deep within has remarkable impact (Odvody et al., 1999). Tests by Prom et al. (2005) in Texas, USA showed that macroconidia on panicles in the field could survive in an infective condition long enough to serve as inoculum the following year.

Sclerotia have only been experimentally germinated with difficulty to produce the teleomorph (Frederickson et al., 1991) and they may be less important than the conidia for survival. Frederickson et al. (1991) finally generated the teleomorph by incubating sclerotia in pots of soil outdoors under natural growing season conditions in Zimbabwe. Initial germination took 4 weeks under diurnal fluctuations of 19-28°C. Although well-developed sclerotia have been observed in the USA (Texas), Mexico and Puerto Rico, they have not been germinated beyond the initial eruption of a globose, papillate structure through the rind despite extensive efforts (DE Frederickson, INTSORMIL, Box 776, Bulawayo, Zimbabwe, unpublished data). Sclerotia may be important as vehicles of conidia in the attached sphacelia. For example, macroconidia may ooze from the sphacelial portion as the moisture content rises following contact with the soil, resulting in secondary conidiation (Bandyopadhyay et al., 1991). This, however, requires experimental verification.

The sclerotial tissues of C. africana produce alkaloids. The alkaloid content of ergot bodies varies between 0.02 and 0.98% wt/wt (Mantle, 1968; Mantle and Waight, 1968; Frederickson 1990; Frederickson et al., 1991), largely depending upon the proportion of sclerotial tissue present. Dihydroergosine accounts for nearly 90% of the alkaloid (Frederickson, 1990) with trace amounts of the biosynthetic intermediates chanoclavine, festuclavine, pyroclavine and dihydroelymoclavine present. The presence of dihydroergosine in sclerotia is an important diagnostic criterion for C. africana (Frederickson et al., 1991; Mantle and Hassan, 1994; Reis at al., 1996).

The question of toxicity inevitably arises by analogy to the potent ergotamine alkaloids of Claviceps purpurea and agroclavine of Clavicepsfusiformis (Youngken, 1947; Shone et al., 1959; Loveless, 1967). Potential toxicity of C. africana alkaloids was investigated by Mantle (1968) using an African pathogen. Diets incorporating up to 50% sclerotia exhibited no toxigenicity to mice. However, in 1997, piggeries in Australia reported feed refusal and loss of milk production in sows, resulting in piglet starvation, when sorghum feed contained 1-20% sclerotia naturally (Blaney et al., 2000). A decline in milk production was similarly reported for dairy cows. Effects could be mimicked by incorporation of 1-5% sclerotia into pig feed, where 5% sclerotia resulted in feed refusal, weight loss and reduction in blood prolactin concentration. Similarly 2.5-5.0% sclerotia in poultry diets resulted in respiratory difficulties, diarrhoea and death. Investigations (Bailey et al., 1999) in the USA, reiterated those of Mantle: there were no adverse effects, other than reduced feed efficiency, when chickens were fed up to 10% C. africana sclerotia for 3 weeks. These conflicting results reflect the distribution of the two strains of C. africana: the 'west' strain in Africa and the Americas and the 'east' strain in Australia and India (Pazoutová et al., 2000). As both strains produce dihydroergosine, and intermediates, in sclerotia, the origin of this toxigenic difference is unclear. Porter et al. (1998) stated that the toxin fusaric acid, produced by Fusarium species commonly associated with honeydew, co-occurred with all samples of ergot alkaloids analysed. Whether Fusarium toxins could be responsible for clinical symptoms or could produce such effects in synergy with rather low levels of ergot alkaloids needs further investigation (Porter, 2000).

Alternative hosts are known and have a proven role in inoculum survival in some parts of the world. In the Americas and Australia, C. africana infections on Johnson grass (Sorghum halepense) and other species of sorghum are common (Ryley et al., 1996; Odvody et al., 1998) and experimental cross-inoculation onto cultivated sorghum and back has been successful. Ergot was first recorded in Texas, USA, in 1997 (Isakeit et al., 1998) and mild winter weather allowed the pathogen to persist in an active form throughout the southern part of the state until the spring of 1998. The summer of 1998 in Texas, however, was unusually hot and dry and ergot was not observed on sorghum during the growing season. Ergot was only seen throughout the state on sorghum forages and regrowth, and on Johnson grass, with the resumption of cool temperatures and rains the following winter. C. africana has repeatedly survived the winter as active, conidial inoculum in Texas and Mexico in following years (Odvody et al., 2002). In Africa, the involvement of alternative hosts in the life cycle of C. africana has not been demonstrated, but merits further investigation. There are often conflicting reports in the literature. In Zimbabwe, it is postulated that a wild sorghum species, or other grass, may play a vital role in bridging the gap between the end of the winter, when surviving conidial inoculum is very limited, and the flowering of cultivated sorghums a few months later (Frederickson and Odvody, 2002). C. africana inoculum was collected from Hyparrhenia rufa in Zimbabwe (Frederickson, INTSORMIL, Zimbabwe; Pazoutova et al., 2004), but it has not yet been determined if H. rufa is an alternative host (see also Loveless, 1964). Lineages of C. africana in Hyparrhenia spp. recently were shown to differ genetically, and their macroconidia differed in size, from those affecting sorghum (Pazoutová and Frederickson, 2005). Pearl millet (Pennisetum glaucum) suffered a low severity of infection following artificial inoculation with C. africana under high inoculum pressure in Zimbabwe (Frederickson and Mantle, 1996) and India (Muthusubramanian et al., 2005). However, it is not expected to be a significant source of inoculum under natural conditions.

Insects are known to carry ergot conidia non-specifically on their bodies after feeding on honeydew (Prom et al., 2003). Sucking bugs that feed on the ovaries and developing seeds of Sorghum bicolor and S. halepense can also transport infective conidia (Prom, 2005). Prom and Lopez (2004) found that corn earworm moths (Helioverpa zea) could ingest conidia and excrete them in a viable and infective condition for more than 24 hr, and thus are conceivably capable of transporting the pathogen from Mexico to Texas and delivering it while ovipositing on susceptible flowering panicles.

Climate

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ClimateStatusDescriptionRemark
Am - Tropical monsoon climate Preferred Tropical monsoon climate ( < 60mm precipitation driest month but > (100 - [total annual precipitation(mm}/25]))
As - Tropical savanna climate with dry summer Preferred < 60mm precipitation driest month (in summer) and < (100 - [total annual precipitation{mm}/25])
Aw - Tropical wet and dry savanna climate Preferred < 60mm precipitation driest month (in winter) and < (100 - [total annual precipitation{mm}/25])
B - Dry (arid and semi-arid) Preferred < 860mm precipitation annually
BS - Steppe climate Preferred > 430mm and < 860mm annual precipitation
C - Temperate/Mesothermal climate Preferred Average temp. of coldest month > 0°C and < 18°C, mean warmest month > 10°C
Cf - Warm temperate climate, wet all year Preferred Warm average temp. > 10°C, Cold average temp. > 0°C, wet all year
Cw - Warm temperate climate with dry winter Tolerated Warm temperate climate with dry winter (Warm average temp. > 10°C, Cold average temp. > 0°C, dry winters)

Seedborne Aspects

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Incidence

Seed lots may be contaminated with sphacelia, the highly convoluted, cream-white to grey cushions of mycelia that usually become visible on the host 6-12 days after infection (Frederickson, 1990; Bandyopadhyay et al., 1990). Sclerotial tissues may also be present. Sclerotia have been induced to germinate experimentally only once and the production of ascosporic inoculum is thought to be infrequent in nature. However, sclerotial tissue always carries residual sphacelial tissue with it and so sphacelia/sclerotia in seed are a cause for concern as either tissue type may be a source of inoculum. Honeydew can also contaminate seeds and the spore concentration on seed from a naturally-infected field in Africa was 55,000,000 spores/g seed (McLaren, 1993). However, where C. africana is endemic, the likelihood of ergot infection resulting from the importation of contaminated seed into an area is very slim compared to infection arising from the dissemination of airborne inoculum (secondary conidia) from relatively local sources (Odvody et al., 2002).

Bhuiyan et al. (2002) showed that macroconidia of C. africana survived in dried honeydew on soil for 13-14 weeks in a glasshouse at ambient temperatures, but for less than half that time on seed stored in a shadehouse over summer. However, those on seeds stored at 4°C survived for over a year (58-62 weeks). During summer, conidia on ergot-infected panicles buried in soil, or on the soil surface, survived for 7.5-12 weeks, whereas over winter the survival times were 4 weeks and 19-27 weeks, respectively. Macroconidia on infected panicles held above the soil surface survived for >38 weeks (8 calendar months) over winter, suggesting that they may play a role in the perennial appearance of C. africana in Australia. Prom et al. (2005) obtained similar results for survival, as well as retention of infectivity, on panicles on or above the soil surface in Texas, USA.

The effects of temperature and relative humidity on infection of Sorghum bicolor male sterile line AQL 33 by C. africana Australian isolate 10765 was studied under controlled conditions (Tonapi et al., 2002). Secondary conidia produced from macroconidia were brushed onto 100 stigmas per flowering panicle and placed at either 10, 20, 25, 30, 35 or 40°C (60% RH, 10 h daylight/14 h night). Relative humidities were increased to 100% for 3 days in half the treatments by sealing panicles into plastic bags. The optimum conditions for ergot infection were 20°C and 100% RH. The optimum temperature for pollen germination and viability was 25°C, with maximum pollen production at 60% RH occurring at 35°C and at 100% RH occurring at 30°C.

Komolong et al. (2003) demonstrated a possible histological basis for partial resistance in male-sterile sorghum lines. Using chitin-specific fluorescin-isothiocyanate-conjugated wheat germ agglutin and callose-specific aniline blue, this study investigated the process of sorghum ovary colonization by C. africana. Conidia germinated within 24 h after inoculation (a.i.); the pathogen was established in the ovary by 79 h a.i., and at least half of the ovary was converted into sphacelial tissue by 120 h a.i. Changes in fungal cell wall chitin content and strategic callose deposition in the host tissue were associated with penetration and invasion of the ovary. The rate of ovary colonization differed in three male-sterile lines that also differed in ergot susceptibility.

Effect on Seed Quality

Honeydew exudate of C. africana on seeds reduced germination by up to 60% (McLaren, 1992a) and severe moulding occurred owing to saprophytic colonization of the exudate. As more honeydew was removed from seeds by washing, so mesocotyl discoloration and post-emergence damping-off were reduced (McLaren, 1993). Honeydew exudates contain toxic or inhibitory substances to sorghum plumule and root growth (McLaren, 1993). The proportion of small seeds in infected plants was 6% greater than in uninfected plants in a study in the State of Guanajuato, Mexico (Hernández-Martínez et al., 2006).

Pathogen Transmission

Studies in Africa and Mexico indicate that sorghum seed from C. africana-infected plants are coated with sugary exudates in which the fungal spores are borne (McLaren, 1993; Valasquez-Valle et al., 1998), but the role of these seeds, or seed-sphacelia/sclerotia admixtures, in the spread of ergot has not yet been determined. Sorghum is not susceptible at sowing (seed), but only much later at flowering, so a flowering, alternative host would have to be present at sowing to permit infection. Even if secondary conidia were produced by iterative germination of macroconidia, the spores, being planted with seed in the ground, would have no inoculum potential. The likelihood of ergot infection resulting from the importation of contaminated seed into an area is very slim compared to infection arising from the dissemination of airborne inoculum (secondary conidia) from relatively local sources (Odvody et al., 2002).

The rapid spread of the disease across vast continental areas in the Americas and Australia is consistent only with wind-borne spread of conidia (Bandyopadhyay et al., 1998). Relationships between weather conditions and rate of spread of C. africana have been studied in Mexico (Velasquez-Valle et al., 2001), Texas, USA (Workneh and Rush, 2002) and Australia (Wang et al., 2000). However, wind-borne dissemination of the pathogen is unlikely to have been responsible for the initial introduction of ergot disease to these countries. Identity of genotypes and evidence from historical phytosanitary samples support transmission of the pathogen across the Atlantic Ocean as sclerotia in imported seed (Pazoutová and Frederickson, 2005).

Seed Treatment

The contamination of C. africana honeydew on infected seeds could be reduced by seed washing, but large volumes of water (4 L/15g seed) were required to reduce the concentration to levels, which would not affect germination and seedling development (McLaren, 1993). In Texas commercial seed production, air-blowing, screen-cutting (sieving) and use of the gravity table removes almost all sclerotia/sphacelia from seed (DE Frederickson, INTSORMIL, Box 776, Bulawayo, Zimbabwe, personal communication, 2001). The flotation test, in which seed is stirred into 10% salt solution with the expectation that all floating bodies are ergot, is very unreliable (Alderman et al., 1999; Frederickson et al., 1999). In practice, ergot bodies float only if there is a large proportion of sclerotial tissue present. Other foreign bodies float, and seed may float if it is cracked or has glumes attached. Conversely, all sphacelia sink.

Coating of seed with captan renders conidia in honeydew deposits inviable and prevents germination of conidia from any remaining sphacelial tissues (Dahlberg et al., 1999; Odvody et al., 2002). To assess the inhibitory effect of captan towards conidia from sphacelia/sclerotia in seed admixtures, fresh sphacelia were mixed with sorghum seeds. Captan was applied at 94 g a.i/100 kg seed . Sphacelia were retrieved and intact sphacelia, or the central core tissues resulting from trimming, were plated onto Kirchoff's agar. Secondary conidiation per sphacelium or core was evaluated on a 0-3 scale; germination incidence (percentage of total number with any associated secondary conidiation) was also calculated. Secondary conidiation per sphacelium was significantly inhibited in the captan-treated sphacelia, with the mean just below the experimental threshold of detection. Captan treatment also significantly reduced the proportion of sphacelia with germination (Frederickson and Odvody, 2003). Seeds treated with thiram had 31% less fungal colonies than non-treated seeds in a Mexican study (Cisneros-López and Mendoza-Onofre, 2010).

Seed Health Tests

Visual examination (Alderman et al., 1999; Frederickson et al., 1999).

Seed lots are examined by at least a x10 magnification for the presence of sphacelia: cream-white to grey cushions of fungal growth. If present at the base of sphacelia, sclerotial tissues are red-brown and oval-spherical in shape. Host tissues (lemma, palea, glumes) may be attached. (c.f. Sclerotia of the Indian Claviceps sorghi are thin, cylindrical and elongate; sclerotia of the Japanese C. sorghicola are conical, elongate and purple-black).

Pathway Causes

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CauseNotesLong DistanceLocalReferences
Seed tradepresumed means of reaching Americas Yes Bandyopadhyay et al., 1998; Pazoutová and Frederickson, 2005

Pathway Vectors

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Plant Trade

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Plant parts liable to carry the pest in trade/transportPest stagesBorne internallyBorne externallyVisibility of pest or symptoms
Flowers/Inflorescences/Cones/Calyx hyphae; sclerotia; spores Yes Pest or symptoms usually visible to the naked eye
Leaves spores Yes Pest or symptoms usually visible to the naked eye
Stems (above ground)/Shoots/Trunks/Branches spores Yes Pest or symptoms usually visible to the naked eye
True seeds (inc. grain) hyphae; sclerotia; 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
Bark
Bulbs/Tubers/Corms/Rhizomes
Fruits (inc. pods)
Growing medium accompanying plants
Roots
Seedlings/Micropropagated plants
Wood

Impact Summary

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CategoryImpact
Economic/livelihood Negative

Impact

Top of page Ergot disease is primarily an economic problem in F1 hybrid seed production. It is particularly severe in male-sterile lines (A-lines) when either nonsynchronous flowering of A-line and restorer lines (R-lines) or adverse environmental conditions result in lack of viable pollen and delayed seed set (Bandyopadhyay et al., 1998). Losses of 10-80% have been reported in hybrid seed production fields in India and regular annual losses of 12-25% recorded in Zimbabwe (Frederickson and Leuschner, 1997; Bandyopadhyay et al., 1998). It has been estimated that ergot will cost the Australian seed industry A$4 annually (Bandyopadhyay et al., 1998) and in the USA, annual production cost increases due to ergot are projected at $5 million (Anon., 1997).

Risk and Impact Factors

Top of page Invasiveness
  • Invasive in its native range
  • Proved invasive outside its native range
  • Abundant in its native range
  • Highly mobile locally
  • Fast growing
  • Has high reproductive potential
  • Reproduces asexually
Impact outcomes
  • Host damage
  • Negatively impacts agriculture
  • Negatively impacts livelihoods
Impact mechanisms
  • Pathogenic
Likelihood of entry/control
  • Highly likely to be transported internationally accidentally
  • Difficult to identify/detect as a commodity contaminant
  • Difficult/costly to control

Diagnosis

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C. africana can be distinguished from the other sorghum ergot pathogens by the detection of the alkaloid dihydroergosine in sclerotial tissues (Frederickson et al., 1991; Mantle and Hassan, 1994) or by comparison of the nucleotide sequences of ITS1 and part of 5.8S rDNA (Pazoutová et al., 2000). The east and west strains of C. africana could be distinguished by the RAPD banding patterns produced with seven primers (Pazoutová et al., 2000; Pazoutová and Frederickson, 2005). The use of assays, with specially-designed PCR primers and probes, for the detection of C. africana in sorghum is detailed in Tooley et al. (2010).

Detection and Inspection

Top of page In the field, C. africana infection is usually obvious from the dripping of honeydew from infected florets and honeydew deposition on the panicle, leaves, stalk and soil. Often the panicle is spectacularly white (see Symptoms). The pathogen is less easily detected in seed batches and poses potential problems for seed exchange (see Seed Treatment and Seed Health Tests).

Similarities to Other Species/Conditions

Top of page Two other species of Claviceps, C. sorghi (Kulkarni et al., 1976) and C. sorghicola (Tsukiboshi et al., 1999) infect sorghum and overlap in distribution with C. africana. Records of C. sorghi in India go back to 1917 (McRae, 1917), but uncertainty about the present distribution of the pathogen originated a recent survey (Muthusubramanian et al., 2006; Tooley et al., 2006). C. sorghi may have been marginalized or eradicated by the more virulent C. africana. C. sorghicola is found only in central Japan (Tsukiboshi, 1999).

Both C. sorghi and C. sorghicola form elongate sphacelia and sclerotia, whereas those of C. africana are subglobose. Unfortunately, this difference will not be apparent to the untrained eye early in infection. Other diagnostic criteria are that C. sorghi honeydew appears before the sphacelia are visible so that infected florets show the honeydew ooze even though the fungus is still invisible (Frederickson and Mantle, 1988). The converse is true for C. africana, the bulky sphacelium causing the glumes to bulge open before honeydew is produced (Frederickson, 1990; Frederickson et al., 1991). Conidia of C. sorghi and C. sorghicola do not form secondary conidia in nature, so honeydew never appears superficially white.

In the absence of the teleomorph, which is difficult to obtain, the best diagnostic test for C. africana is the presence of the alkaloid dihydroergosine in sclerotial tissues (Frederickson et al., 1991; Mantle and Hassan, 1994; Reis et al., 1996). C. sorghicola and C. sorghi sclerotia produce trace amounts of caffeine (Bogo and Mantle, 2000; Bogo et al., 2003), (N.b. sphacelia do not synthesise alkaloid in any species). Pazoutová et al. (2000) used nucleotide sequence differences of ITS1 and 5.8S rDNA to distinguish between the species. Tooley et al. (2006) observed distinct AFLP locus patterns for C. sorghi and C. africana from India. For a full comparison of C. sorghi and C. africana morphology, see Frederickson et al. (1991) and Muthusubraniam et al. (2005). For details of C. sorghicola, see Tsukiboshi et al. (1999)

The sori of covered kernel smut (Sporisorium sorghi) and long smut (Tolyposporium ehrenbergii) are sometimes confused with C. africana sphacelia. However, in these fungi the sack-like sori comprise a smooth, cream to grey outer covering or peridium enclosing the powdery-black teliospores (Frederiksen, 1986; Hilu, 1986). The ergot sphacelia are devoid of an outer covering, being solid, spongy bodies with convolutions which are the microscopic spore-bearing cavities.

Prevention and Control

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Cultural Control and Sanitary Methods

Cultural control is not a reliable control technique, often depending on the capricious nature of the climate. In Zimbabwe, crops may escape ergot if early rains permit sowing in November so that flowering both avoids pollen sterility induced by cool nights and coincides with a mid-season dry spell in January or February (Frederickson and Leuschner, 1997). Sorghum is unaffected by ergot when seed multiplication is performed under irrigation in the dry season at Mazarabani. Similarly, early sowings of sorghum avoid ergot infection in India (Singh, 1964; Sangitrao et al., 1979; Anahosur and Patil, 1982) and central Mexico (Montes-Belmont et al., 2002b).

Field practices aimed at reducing the risk or severity of infection include the removal of infected panicles at harvest, 3-year crop rotations and deep ploughing of field residues. However, despite these measures, a serious epiphytotic occurs every 5-10 years in Zimbabwe (Frederickson and Leuschner, 1997). Increasing the ratio of pollen-producing rows to the male-sterile, female parent, or staggering the planting dates of the pollen donor rows helped reduce ergot by increasing the period when pollen was available (Frederickson and Leuschner, 1997), but only if the weather conditions were favourable for pollination. Cold nights 2-3 weeks before flowering and cool, wet weather at flowering and during the 5 days after flowering (McLaren and Wehner, 1990; 1992) have an overriding negative effect on all planting systems, promoting disease.

Host-Plant Resistance

There is currently no source of resistance to sorghum ergot for use in the field in A-lines. Resistant fertile sorghums have been reported (Tegegne et al., 1994; Musabyimana et al., 1995), but resistance has proved to be a function of cleistogamy, or fast and efficient pollination and fertilization (Bandyopadhyay, 1992; Frederickson et al., 1994) with no potential use in A-lines. In trying to evaluate resistance, simple comparisons of incidence data from genotypes from different localities, following natural infection or artificial inoculation, are meaningless (McLaren, 1992b) because susceptibility to ergot is extremely sensitive to environmental factors at flowering and a few weeks before (McLaren and Wehner, 1990; 1992; McLaren, 1997; Montes-Belmont et al., 2002b). Cool nights of <12°C at 2-3 weeks before anthesis result in pollen sterility and increased ergot severity. Therefore, tolerance of low, pre-flowering temperatures is important for disease avoidance (McLaren, 1997). Similarly, the mean maximum temperature 1-4 days after pollen shed affects incidence with no disease occurring at >28°C. Interactions between genotype, location and flowering date must be compared by regression analyses because flowering dates of even a day or two apart affect the severity of ergot (McLaren, 1992b; McLaren and Flett, 1998).

Careful screening and selection for floral characteristics that reduce disease severity may prove to be one useful strategy. In Puerto Rico, Dahlberg and Bandyopadhyay (USDA-ARS-TARS, Puerto Rico, personal communication, 1999) found a male-fertile accession with glumes, which tightly clasp the ovary, apparently conferring tolerance to high inoculum loads. This line also showed potential in a male sterile background. In the USA, many A-line sorghums have a protracted stigma receptivity period that confers high ergot susceptibility (Odvody, 1997) and disease reduction may possibly be achieved by decreasing the ergot susceptible period of the A-line stigma. Other advantageous modifications might include reducing the floret gaping period, selecting for more rapid post-fertilization changes in the A-line, breeding for cold temperature tolerance in R-line pollen production and during fertilization, and extending the pollen production period.

Chemical Control

Chemical control is barely cost effective, only feasible for controlling disease on the A-lines and unnecessary on the hybrids themselves. In South Africa, three ground applications of bitertanol or procymidone were effective at controlling C. africana in A-lines in the field when sprayed preventatively (McLaren, 1994). In Zimbabwe, a single spray of benomyl at heading or flowering controlled ergot, but was ineffective when applied to plants already showing disease symptoms (Frederickson and Leuschner, 1997). In Brazil, three to four ground sprays of propiconazole or tebuconazole (triazole fungicides) applied at 5- to 7-day intervals were successful at controlling ergot in the A-lines of seed production plots in the absence of rains (Ferreira et al., 1996; Odvody, 1997; Odvody et al., 1998). These chemicals, as well as azoxystrobin, were also the most efficacious of 14 fungicides tested in greenhouse and field trials in the USA (Prom and Isakeit, 2003). The fungicide must be precisely deposited on the stigmas, a requirement rendering aerial application ineffective so far. See also ‘Seed Treatments’.

Biological Control

Bhuiyan et al. (2003) found that some fungal isolates, including those in Trichoderma species-containing commercial products, could reduce or prevent disease in glasshouse trials, particularly when the biocontrol agent was applied several days before inoculation.

Development of IPM Strategies

The integration of several control practices would provide the greatest potential for control. Cultural control alone has little impact on disease. Chemical use in the field is only economically feasible in seed production plots and then only with marginal profit. Fungicide usage should be viewed as a short- term control measure pending the development of other strategies because over-dependence on chemicals may result in problems of fungicide tolerance by the pathogen in the future (Frederickson and Leuschner, 1997). Fungicide treatment of seed, however, is an effective and economic way to control potential seed-borne, conidial inoculum (Dahlberg et al., 1999; Odvody et al., 1999).

Disease forecasting by using weather variables to predict ergot severity is still in its infancy. In South Africa, a simple model based on multiple regression analysis of pre-flowering minimum temperature, mean daily maximum temperature and mean daily maximum relative humidity (McLaren and Flett, 1998) accurately predicted ergot potential in experimental plots. The model was not applicable to Australian conditions and Meinke and Ryley (1997) developed an alternative model, based on climatic conditions before and around flowering, which enabled the risk assessment of 10 locations. With a view towards more precise timing of fungicide application to breeding line crops, Ryley and Chakraborty (2008) then investigated patterns of conidium release in South Queensland. Release occurred 1-3 days following rainfall, and airborne conidia were detected during hours characterized by higher median temperature, windspeed and vapour pressure deficit, lower relative humidity, and leaf wetness values of 0%. In the USA, Workneh and Rush (2006) obtained “promising results” with a model, based on cumulative departures from maximum temperature and minimum relative humidity thresholds, for forecasting ergot in hybrid seed production fields of northern Texas.

Combining some form of host genetic resistance with disease prediction and limited fungicide use during high risk periods may be the best strategy for control in the future.

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India: International Crops Research Institute for the Semi-Arid Tropics (ICRISAT), Patancheru 502 324, Andhra Pradesh

USA: Sorghum Improvement Conference of North America, P.O. Box 530 Abernathy, Texas 79311

Contributors

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10/09/09 Updated by:

Systematic Mycology & Microbiology Laboratory, USDA-ARS, 10300 Baltimore Ave., Beltsville, MD 20705, USA

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