Tilletia indica (Karnal bunt of wheat)
- Summary of Invasiveness
- Taxonomic Tree
- Distribution Table
- Risk of Introduction
- Hosts/Species Affected
- Host Plants and Other Plants Affected
- Growth Stages
- List of Symptoms/Signs
- Biology and Ecology
- Means of Movement and Dispersal
- Seedborne Aspects
- Plant Trade
- Economic Impact
- Environmental Impact
- Social Impact
- Detection and Inspection
- Similarities to Other Species/Conditions
- Prevention and Control
- Distribution Maps
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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- Tilletia indica Mitra
Preferred Common Name
- Karnal bunt of wheat
Other Scientific Names
- Neovossia indica (Mitra) Mundk.
International Common Names
- English: Indian bunt of wheat; new bunt; partial bunt of wheat
- French: carie de Karnal
Local Common Names
- Germany: Indischer Weizenbrand
- NEOVIN (Tilletia indica)
Summary of InvasivenessTop of page
T. indica is the fungal pathogen causing Karnal bunt of wheat seeds. Its distribution is mainly limited to northwest India and adjoining countries, North America and South Africa; it is listed as a quarantine pest in Europe, Australia, South America and elsewhere. In North America it was confined to an area in northwest Mexico until 1996, when it was reported from Arizona. Since then, surveys have detected it in a few locations in southwest USA. It has also recently been found in germplasm imported into other continents.
Crop damage after a recent introduction of T. indica is minor. Infection is usually only in a few kernels in a spike making the disease much less conspicuous than other bunts where whole ears are infected. This makes Karnal bunt much more difficult to detect and can lead to a major infection of wheat (bread and durum) and triticale crops before it is fully realised. The fungus is almost impossible to eradicate from the soil and the teliospores remain viable for up to 5 years in infested soil. Yield losses can occasionally be high in susceptible varieties but the main effect is on seed quality. Cost implications can be large because of the quality loss. In addition, export of grain to wheat-growing countries that are presently free of T. indica would be restricted and expensive to control plus monitoring operations would need to be implemented to satisfy the importers’ phytosanitary requirements which is usually area freedom.
Taxonomic TreeTop of page
- Domain: Eukaryota
- Kingdom: Fungi
- Phylum: Basidiomycota
- Subphylum: Ustilaginomycotina
- Class: Ustilaginomycetes
- Subclass: Exobasidiomycetidae
- Order: Tilletiales
- Family: Tilletiaceae
- Genus: Tilletia
- Species: Tilletia indica
DescriptionTop of page
Teliospores are dark reddish to coppery, dull brown or dark brown, some spores typically black/opaque, globose to subglobose, occasionally with a mycelial fragment (apiculus) attached; 24-47 µm diameter (about twice that of Tilletia caries); exospore with thick, truncate, compact projections, 1.5-5 µm high, seen in median view. The opacity of the teliospores (Duran and Fischer, 1961) is one factor that differentiates T. indica from T. walkeri that is found on ryegrass (Lolium spp.) (Castlebury and Carris, 1999); other factors include surface ornamentation and size parameters. Sterile cells are intermingled with teliospores in the sori; very variable, globose, subglobose, frequently lacrymiform, yellowish-brown, 10-28 µm wide at their widest point to 48 µm in overall length, with a well developed stalk; walls laminated, up to 7 µm thick.
Primary sporidia usually 64-79 x 1.5-2 µm; secondary sporidia usually 12-13 x 2 µm.
For more information, see Duran and Fischer (1961), Khanna et al. (1968) and Waller and Mordue (1983).
DistributionTop of page
The first report of a new bunt disease in wheat came from the region of Faizalabad (Pakistan) in 1909. This was presumably Karnal bunt, which was first formally recorded in 1930 near the north Indian city of Karnal (Mitra, 1931). Within India the pathogen spread and can now be considered widespread in northern and central India (in regions where low winter temperatures and high humidity prevail, viz. Delhi, Uttar Pradesh, Haryana, Punjab, Himachal Pradesh, Rajasthan, Madhya Pradesh, Jammu and Kashmir, West Bengal and Gujarat) (Singh et al., 1985).
The first report of Karnal bunt from a non-Asian country came from Mexico in 1972, where the disease has been reported from localized areas (500,000 ha) within the state of Sonora (EPPO, 1991) and from the states of Sinaloa and Baja California Sur.
Reports in the literature of occurrences in Lebanon, Sweden, Syria and Turkey (Lambat et al., 1983) and Kenya (intercepted plant material, IMI Herbarium) have not been substantiated. These records were based on intercepted wheat consignments and have not been confirmed by the countries or by a screening survey of the International Center for Agricultural Research in the Dry Areas (ICARDA) on wheat germplasm from the Middle East (Diekmann, 1987). For more information, see Locke and Watson (1955), Warham (1986), Gill et al. (1993). Isolated outbreaks have been found in south-western USA since its first reported occurrence there in 1996. In 2000, Karnal bunt was reported in South Africa (Rong, 2000; Crous et al., 2001).
See also CABI/EPPO (1998, No. 246).
Distribution TableTop of page
The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.
|Continent/Country/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
|Afghanistan||Present||IMI, 1996; EPPO, 2014|
|India||Widespread||IMI, 1996; EPPO, 2014|
|-Bihar||Present||Singh et al., 1985; IMI, 1996; EPPO, 2014|
|-Delhi||Present||Singh et al., 1985; IMI, 1996; EPPO, 2014|
|-Gujarat||Present||IMI, 1996; EPPO, 2014|
|-Haryana||Present||Anil Gupta et al., 1990; IMI, 1996; EPPO, 2014|
|-Himachal Pradesh||Present||Sharma and Anil Sirdhi, 1994; IMI, 1996; EPPO, 2014|
|-Indian Punjab||Present||IMI, 1996; EPPO, 2014|
|-Jammu and Kashmir||Present||IMI, 1996; EPPO, 2014|
|-Madhya Pradesh||Present||Singh et al., 1985; IMI, 1996; EPPO, 2014|
|-Rajasthan||Present||Singh et al., 1985; IMI, 1996; EPPO, 2014|
|-Uttar Pradesh||Present||IMI, 1996; EPPO, 2014|
|-Uttarakhand||Present||Singh et al., 2002|
|-West Bengal||Present||Singh et al., 1985; IMI, 1996; EPPO, 2014|
|Iran||Restricted distribution||EPPO, 2014|
|Lebanon||Absent, invalid record||IMI, 1996; EPPO, 2014|
|Nepal||Present||IMI, 1996; EPPO, 2014|
|Pakistan||Restricted distribution||IMI, 1996; EPPO, 2014|
|Syria||Absent, invalid record||IMI, 1996; EPPO, 2014|
|Turkey||Absent, invalid record||IMI, 1996; EPPO, 2014|
|Kenya||Absent, intercepted only||IMI, 1996; EPPO, 2014|
|South Africa||Present, few occurrences||EPPO, 2014|
|Mexico||Restricted distribution||IMI, 1996; EPPO, 2014|
|USA||Present, few occurrences||1996||IMI, 1996; EPPO, 2014|
|-Alabama||Absent, invalid record||EPPO, 2014|
|-Arizona||Present, few occurrences||IMI, 1996; NAPPO, 2013; EPPO, 2014|
|-New Mexico||Eradicated||USDA-APHIS, 2009, personal communication; IMI, 1996; EPPO, 2014|
|-Tennessee||Absent, invalid record||EPPO, 2014|
|-Texas||Eradicated||IMI, 1996; EPPO, 2014|
|Brazil||Present, few occurrences||EPPO, 2014|
|-Rio Grande do Sul||Present, few occurrences||EPPO, 2014|
|Denmark||Absent, no pest record||DCA - Nationalt Center for Fødevarer og Jordbrug, Denmark, 2018|
|Hungary||Absent, confirmed by survey||EPPO, 2014|
|Netherlands||Absent, confirmed by survey||NPPO of the Netherlands, 2013; EPPO, 2014||Based on long-term annual surveys, 63 survey observations in 2012.|
|Poland||Absent, intercepted only||IMI, 1996; EPPO, 2014|
|Russian Federation||Absent, intercepted only||EPPO, 2014|
|Sweden||Absent, invalid record||IMI, 1996; EPPO, 2014|
Risk of IntroductionTop of page
RISK CRITERIA Category
ECONOMIC IMPORTANCE Moderate
SEEDBORNE INCIDENCE Low
SEED TRANSMITTED Not recorded
SEED TREATMENT Yes
T. indica has long been considered to be an important quarantine pest. Jones (2007a, b) has questioned whether the pathogen justifies this status but Sansford et al. (2008) have examined all the current evidence and consider T. indica does warrant quarantine pest status.
T. indica is regulated by the European Union (EU, 2000) and by other EPPO countries (Belarus, Russia, Turkey, Ukraine). It is considered to present a risk also in Africa, Asia, Near East and South America. Its distribution in North America remains very limited. Once introduced, the fungus would be almost impossible to eradicate because spores can remain viable in the soil for a considerable time. However, eradication has been achieved in New Mexico because of strict quarantine measures were imposed on affected fields. T. indica presents a hazard to both bread wheat and durum wheat in areas with favourable climatic conditions. According to Baker et al. (2000) and current Pest Risk Analysis (Sansford et al., 2006: http://karnalpublic.pestrisk.net/files/eu_karnalbunt_pra.pdf; Sansford et al., 2008), a huge area within the European and Mediterranean region which is devoted to the cultivation of wheat and other Poaceae would be at risk to infection by T. indica, especially in the temperate and cool areas of western and northern Europe. For further information on this assessment of risk associated with karnal bunt in the EU, see http://karnalpublic.pestrisk.net/deliverables/.
Australia, where wheat is grown in a semi-arid environment, has been deemed to be at risk (Murray and Brennan, 1998; Stansbury and McKirdy, 2000; Stansbury et al., 2002).
Hosts/Species AffectedTop of page
The main host of T. indica is wheat (Triticum spp.) (Aujla et al., 1986, 1987); durum wheat and triticale are less susceptible. Plants are infected within 2-3 weeks of heading.
In inoculation experiments Aegilops spp., Bromus spp., Lolium spp. and Oryzopsis showed varying degrees of susceptibility (Royer and Rytter, 1988).
Host Plants and Other Plants AffectedTop of page
Growth StagesTop of page Flowering stage, Fruiting stage, Post-harvest
SymptomsTop of page
Symptoms depend on climate and are manifested most clearly when cool/warm humid conditions prevail at heading. The fungus causes a reduction in the length of ears as well as in the number of spikelets of bunted ears. Infected plants may be dwarfed. In general, T. indica rarely infects more than a few spikelets per ear and then the affected grains are not swollen. Oblong or ovoid sori, 1-3 mm diameter, develop, containing dusty, brown to black spore masses. These characteristically smell of decaying fish (trimethylamine) as do those of T. tritici, T. foetida and T. contraversa (EPPO/CABI, 1996). Feeding studies have revealed no adverse health effects, but consumers can begin to taste and smell the fishiness when 3% or more of grain is affected.
The grain is partially destroyed, the attack starting at the hilum and running along the suture, leaving the endosperm intact and covered by the whole or partly ruptured seed coat. In the case of mild infection, only a black point just below the embryo towards the suture is apparent. In advanced attack, tissues along the suture and adjacent endosperm are replaced by spores. The glumes spread apart, exposing the infected grains, and both glumes and grains may fall to the ground.
For more information, see Holton (1949) and Duran and Fischer (1961).
List of Symptoms/SignsTop of page
|Inflorescence / black fungal spores|
|Inflorescence / twisting and distortion|
|Seeds / discolorations|
|Seeds / galls|
Biology and EcologyTop of page
T. indica survives in the soil. In certain areas a 2-year period free from wheat reduces, but does not eliminate, the disease. However, survival and spread of the fungus can occur by transport of infested and infected seed. Teliospores germinate at or near the soil surface in response to temperature and moisture, normally at temperatures between 20 and 25°C (Krishna and Singh, 1982). Teliospores produce a promycelium bearing many filiform primary sporidia. Primary sporidia give rise to allantoid or filiform secondary sporidia, or hyphae that can also produce secondary sporidia. Dhaliwal and Singh (1989) found that two types of secondary sporidia are produced: allantoid sporidia and filiform sporidia, of which only the allantoid type is thought to be able to infect and cause the disease. Allantoid secondary sporidia are ballistospores (i.e. forcibly discharged).
Primary and secondary sporidia are dispersed by wind or rainsplash to the wheat ears and act as the primary source of infection. Germ tubes arise from secondary sporidia and grow towards stomatal openings of the glume, lemma or palea, where they enter. The hyphae grow intercellularly within the glume, lemma, palea and possibly rachis, entering the base of the ovary from these tissues and leading to infection of the seed, which is normally limited to the pericarp (Goates, 1988). Spread of the pathogen then appears to take place systemically from primary infection sites to adjacent spikelet and florets (Singh and Dhaliwal, 1988). Another factor regarding floral infection is the finding that sporidia develop on the outer glumes of florets, indicating that repeated cycles of sporidial production in spikes provide secondary inoculum (Bains and Dhaliwal, 1989). Sporidia also develop on leaves and other plant parts. According to Dhaliwal, the fungus colonizes the surfaces of lower plant leaves to produce more secondary sporidia. These are splashed or blown to higher leaves. In this way the pathogen moves in steps up the plant to infect the spike.
The disease seems to be favoured by cool temperatures and high relative humidity at heading (Royer and Rytter, 1985). Temperature, rainfall and humidity factors have been utilized in models proposed to predict the development of Karnal bunt (Singh et al., 1990; Jhorar et al., 1992). Temperatures of 8-20°C and high humidity associated with light rain showers and cloudy weather are most favourable for infection of the ears. Environmental conditions are considered to play a decisive role in infection, with dry weather, high temperatures (20-25°C) and bright sunlight being unfavourable.
Seed- or soil-borne teliospores and their subsequent germination are believed to play only a starting role in Karnal bunt epidemics (Dhaliwal, 1989). According to Bains and Dhaliwal (1989), repeated cycles of sporidial production in the ears provide more inoculum than soil-borne teliospores of T. indica. Secondary sporidia were also able to germinate and multiply on surface-sterilized leaves and in sterile soil as well as on glumes and leaves of resistant wheats, thus providing a large inoculum for airborne infection (Dhaliwal, 1989). These secondary sporidia have been shown to be very durable and can remain dormant and then regenerate very rapidly under conditions conducive for the disease (Goates 2010).
For more information, see Mitra (1931, 1935, 1937), Mundkur (1943a, b), Warham (1986), Goates (1988), Tan et al. (2013) and USDA-APHIS (http://www.aphis.usda.gov/plant_health/plant_pest_info/kb/index.shtml).
Means of Movement and DispersalTop of page
Natural spread can be substantial as teliospores can be carried over long distances and at high altitudes by wind and man-made elements like movement of people, agricultural products and machinery (Warham, 1986). Teliospores can pass through the digestive tracts of animals undamaged (Smilanick et al., 1986), thus making it possible that the pathogen is distributed with farm manure. However, the main mode of international spread is on infected or contaminated wheat seeds. Marshall et al. (2003) suggest that, for movement from Mexico into USA, this may involve not only consignments, but small lots of wheat moving in automobiles, trucks and railway cars. Grain imports have also been targeted as possible avenues of introduction. Teliospores of the pathogen may also be transported in straw, i.e. dried up wheat plants with a few ears mixed up in it.
Seedborne AspectsTop of page
Although high numbers of wheat samples may have infested seed in years conducive to Karnal bunt, generally the numbers of infested seeds per sample are low.
T. indica is commonly found in wheat seeds in northern and central regions of India and Pakistan. A survey of 2144 seed lots in Pakistan showed seed infection by T. indica in 28% of samples in 1983-84 (Shamshad Begum and Mathur, 1989). In India in 1986, bunt spores were detected in 52% of 100 wheat seed samples (Kailash Agrawal et al., 1986). In a later survey in Pakistan (Bhutta et al., 1999) a total of 730 wheat seed samples were tested to assess the incidence of Karnal bunt using the dry inspection method from 1993/94 to 1996/97. High infection percentage (3%) of Karnal bunt in various seed lots was found in central Punjab and north-west areas of Pakistan. Southern parts of the country were found to be free from the disease from 1994/95 to 1996/97. The incidence of Karnal bunt showed a decreasing trend (up to 0.5%) at the country level. As Karnal bunt is known to be a quarantine disease, strict quarantine measures are needed to contain the spread of the disease. During 1989-1990 to 1996-1997, wheat grain samples were collected from markets in different districts of the Indian Punjab. No completely Karnal bunt free area was found in the state but some areas of low disease severity were observed in Bathinda (Singh et al., 1999).
In a survey in 1979-1981, infection levels as high as 17% infected grains were found in wheat seed lots in north-western Mexico (Royer and Rytter, 1985). The latter finding prompted the United States Department of Agriculture (USDA) to quarantine wheat and triticale produced in Mexico. Infestations of T. indica detected in wheat seed lots produced in south-western states of the USA in 1996 has led to the USDA placing these areas under quarantine.
For further discussion of recent and historical distribution records, see Gill et al. (1993) and Jones (2007, 2009).
Histological studies of the infection of wheat seeds indicate that T. indica is restricted to the pericarp (Cashion and Luttrell, 1988; Goates, 1988; Rashmi Aggarwal et al., 1994). Cashion and Luttrell (1988) showed that hyphae proliferate intercellularly in the space formed by disintegration of the middle layers of parenchymatous cells of the pericarp during normal development of the grain and prevent fusion of the remaining outer and inner layers of the pericarp with one another and with the seed coat. The hyphae form a compact, hymenium-like layer over all surfaces of the pericarp tissue surrounding this cavity and give rise to short, septate sporogenous hyphal branches that produce teliospores singly from their terminal cells. Growth of the fungus ruptures the connection between the pericarp tissue surrounding the vascular bundle in the bottom of the adaxial groove in the pericarp and the nucellar projection along the length of the developing seed. The consequence is atrophy of the seed through disruption of the normal flow of nutrients from the pericarp. The endosperm is shrunken to varying degrees and cartilaginous in appearance. The embryo with attached endosperm may be easily dissected out and is germinable before or after removal from the grain. In the most severe infections, the grain is reduced to a black membranous sack of teliospores and the shrivelled embryo is dead. For further information on infection, see Biology and Ecology.
The dormancy of freshly produced teliospores has been mentioned by many workers studying Karnal bunt. For example, teliospores collected from newly mature wheat spikes germinated poorly <10%), but increased to 40-60% over a 4-month period (Smilanick et al., 1986). Teliospores of T. indica in diseased grains of wheat germinated at a rate of 10.3% after storage at 10°C for 1 year (Dhiman and Bedi, 1988).
Another concern about the potential occurrence of T. indica relates to other hosts. Suspect teliospores of T. indica were found in south-eastern USA in USDA-APHIS surveys of wheat seed in 1996 (Cunfer and Castlebury, 1999). Between April and June 1997, ryegrass seed samples were collected from 190 wheat fields in 47 counties in Georgia and from 26 fields in 17 counties in Alabama and south-central Tennessee. In 1998, 70 samples were collected from 40 counties in the same regions of the three states. The teliospores from these samples were 23-45 µm in diameter (average approx. 33 µm) and ranged from light brown to dark reddish-brown. Teliospores had coarse, widely spaced cerebriform ridges on the surface and were surrounded by a gelatinous sheath. The ryegrass bunt was identified as the recently described species T. walkeri (Castlebury and Carris, 1999). In 1997, teliospores of T. walkeri were found in 13 samples from eight counties in central Georgia and from one field in Tennessee. In 1998, more teliospores and bunted seeds were found, possibly due to frequent rain in the region throughout the flowering period for ryegrass. In one wheat field in Morgan County, Georgia, approximately 50% of the ryegrass seed collected were partially bunted and a small percentage were completely bunted. Fields with teliospores were widely distributed and generally matched the locations where teliospores were found in APHIS wheat seed surveys in 1996-1998. No bunted wheat seeds or teliospores of T. indica were found in the survey.
Effect on Seed Quality
Grains of six wheat varieties infected with Karnal bunt were evaluated for number of teliospores and percentage reduction in 1000-grain weight in comparison with non-infected controls. The different grades of infection of the grains ranged from slight infection <10% of endosperm converted into spore mass) to very severe infection (>75% converted into spore mass). Reduction in 1000-grain weight was observed in all the varieties, ranging from 56.9 to 65.9% under severe infection conditions (Karwasra et al., 1991). T. indica infection increased the prolamine and decreased the albumin and globulin protein content of the seed. The decreased level of glutelins lowered the gluten quality in diseased compared with healthy grain (Singh and Bedi, 1985).
Various tests of the effect of T. indica infection on wheat seed quality indicated that it had very little effect on seed viability, irrespective of the age of the seed, whereas germination of infected seed appeared to depend upon the wheat cultivar and age of the seed. In contrast, there was a significant reduction in the vigour of seedlings arising from infected seed (Warham, 1990). Rattan and Aujla (1992a) showed that germination and shoot length, root length and fresh and dry weight of wheat seedlings were severely affected only when >75% of the grain was converted into a spore mass.
There is no direct evidence that T. indica can be transmitted from planted seeds to the plants grown from the seed. However, teliospores that heavily contaminate seeds do survive and germinate in the soil and are considered to be an important inoculum source of the pathogen (Bains and Dhaliwal, 1989).
Highest germination (55-60%) of teliospores from both Mexico and India occurred after 3-week incubation at 15-20°C in continuous light at pH 6-9.5. Germination was similar on soil, soil extract agar and water agar. The promycelia of teliospores germinating under 2 mm of soil or agar were incapable of reaching the surface; presumably they could not release the wheat floret-infecting sporidia which are necessary to complete the life cycle. Teliospore germination resumed unhindered after a 1- or 3-week interruption by freezing (-5°C) or desiccation. Teliospores collected from newly mature wheat spikes germinated poorly <10%), but increased to 40-60% over a 4 month period (Smilanick et al., 1986).
Germination of teliospores occurred on the soil surface over a temperature range of 10 to 25°C and at 5-40% soil moisture content (Rattan and Aujla, 1992b). Teliospores on heavily infested wheat seed germinated at a rate of 2.24% after burial in soil for 27 months (Rattan and Aujla, 1990). Haploid secondary sporidia are produced on germinating teliospores and forcibly discharged into the air. Sporidia can germinate over a fairly wide range of temperature and humidity conditions (Smilanick et al., 1989). They also germinate on glumes of wheat and infect the plants (Singh et al., 1988; Dhaliwal, 1989).
In a screen of 47 products, wheat seed treatments that reduced teliospore germination of T. indica appeared to be limited in the length of their activity. Most were effective for up to 6 months, but only a few for longer. Of those with a longer period of activity, triphenyltin hydroxide (use now discontinued in USA and elsewhere), methoxyethylmercury acetate and ethylmercury chloride were effective for up to 18 months. However, with the possible exception of the mercurial compounds, none was capable of killing T. indica teliospores when applied to infected seeds. It was concluded that no existing chemical seed treatment can ensure that wheat seed is not carrying viable T. indica teliospores (Warham et al., 1989).
In field trials in India, a single protective application of propiconazole controlled the incidence of T. indica on wheat by 71.4%; three sprays reduced disease by 100%. Triadimefon and carbendazim controlled the disease by 87 and 84%, respectively (Aujla et al., 1990). Carboxin + thiram, and chlorothalonil have been used as seed treatments in the USA and Mexico (B Goates, University of Idaho, USA, personal communication, 1998).
Exposing infected seeds to sonication has also been tested as a potential seed treatment. This killed the teliospores without affecting germination of the wheat seed (Satvinder Kaur, 1994). Feeding diseased seeds to animals showed that ingestion by chickens or passage through the intestinal tract of a cow reduced teliospore germination of these fungi, but did not prevent it (Smilanick et al., 1986).
Beniwal et al. (2000) reported that soaking seeds in water could help eradicate teliospores of T. indica without an adverse effect on wheat seed germination. Soaking at 35°C for 12 h was the most effective treatment.
Washing contaminated wheat seeds in 0.005% SDS, 0.001% Tween-20 or 0.05% Triton-X-100 removed 90-95% of teliospores on the seeds (Gupta et al., 2003). The detergents Triton-X-100 and SDS also increased germination rate.
Treatment of wheat seeds with Tilt (propiconazole, 25% EC) and Controll (hexaconazole, 5% EC) gave 94.7 and 88.0% control against karnal bunt; however, as a seed treatment Tilt is phytotoxic, resulting in stunting, curling and yellowing of emerging seedlings (Sharma and Sharma, 2004).
For organic farming, seed treatment with extracts of Canabis sativa, Eucalyptus globulus, Thuja sinensis and Datura stramonium are fully effective against T. indica in the field (Borgen, 2004).
Solar energy treatment can also significantly reduce karnal bunt incidence. Duhan and Beniwal (2004) soaked wheat seeds in water (1:1 w/v) in a galvanised tub that was tightly covered with a transparent polythene sheet. The tub was left out in the sun for six hours (08.00-16.00) during September in north-western India. The treatment gave 73.64% control of T. indica and is recommended by the authors for farmers in the north-western plains of India.
Seed Health Tests and Diagnosis
Visual examination (Shamshad Begum and Mathur, 1989)
Dry seeds are visually examined for the presence of bunted grains. Further examination and testing is required to determine the cause of the grain abnormality. Although Tilletia contraversa, T. tritici and T. foetida usually affect the entire seed, occasionally partial bunting occurs. As well, Puccinia striiformis can infect the testa and produce a small telium resembling a small Karnal bunt lesion but this contains two-celled teliospores. The physiological condition black point can also produce symptoms that could be mistaken for Karnal bunt unless examined more closely.
Wash test (Castro et al., 1994)
1. Samples of 1000 seeds are washed in 600 ml of sterile distilled water containing 0.001% Tween 20.
2. The wash water is filtered initially through a screen (60-µm diameter pore size) to remove large debris, and then through a membrane (12-µm pore size) on which the spores are collected.
3. The spores are washed from the membrane surface and pelleted in a centrifuge run for 30 s at 180 x g.
4. The pellet is resuspended in water and examined by light microscopy for teliospores characteristic of T. indica.
(Teliospores may also be mounted in Shear's mounting media, as modified by Graham (1959), before examination.)
Sodium hydroxide soak (Agarwal and Verma, 1983)
1. Wheat seed samples are soaked in 0.2% NaOH solution for 24 h at 20°C.
2. Seeds infected by T. indica can be identified by the black, diseased portions which contrast with the pale yellow healthy parts.
Wash test (size-selective sieving) and PCR (Peterson et al., 2000).
A seed wash of a 50 g grain sample was washed through 53 µm and 20 µm pore size nylon screens to remove unwanted debris and to concentrate and isolate teliospores. The material retained in the 20 µm screen was suspended for direct microscopic examination or plated on water agar for teliospore germination. DNA extracted from mycelia is used for identification by PCR utilizing one pair of T. indica-specific and one pair of T. walkeri-specific primers. The reliability of detection for both light microscopy and PCR are 100% at an infestation of five teliospores per 50 g sample. The proportion of teliospores recovered from grain samples artificially infested with T. indica was 0, 82, 88, 81 and 82%, respectively, at infestation levels of 0, 1, 2, 5 and 10 teliospores per 50 g wheat sample. Extraction efficiency was comparable to the centrifuge seed-wash method currently used by most seed health laboratories. Sample analysis using size-selective sieving was more than 83% faster than the standard centrifuge seed wash.
PCR–Differentiate and identify T. indica, T. horrida, T. controversa, T. tritici (McDonald et al., 1999)
A simple technique of conducting PCR on single ungerminated teliospores of Tilletia species using primers for either a portion of the nuclear ribosomal intergenic spacer region or a portion of the mitochondrial DNA. The success rate ranged from 100 to 10%. This technique avoids the difficulty and time delay in having to germinate teliospores.
This is a highly sensitive and specific molecular assay performed directly on ungerminated teliospores extracted from size-selective sieving of 50 g seed samples or on genetic material directly released from a single spore. The use of five dual-labelled, species-specific fluorescent probes and associated species-specific primers in a PCR mix enables the identification of T. indica and differentiation from bunt species of T. walkeri, T. horrida, T. ehrhartae and T. tritici that are commonly found in harvested grains. The assay was reported to have the sensitivity to detect three to five spores in grain washings from 50 g grain samples. Direct analysis of selected suspect spores leads to the elimination of the germination step in the confirmation for a small number of spores found in an incursion.
Seed immunoblot binding (Kumar et al., 1998)
The immuno probe generated from polyclonal antisera was used for the development of an immunoblot binding assay for detecting Karnal bunt infections in wheat seed samples. Further development of the anti-teliospore serum for diagnosis reported partial cross-reactivity with Puccinia recondita and T. horrida (Kesari et al., 2005).
Image Analysis (Chesmore et al., 2003; Deng et al., 2012)
Chesmore et al. (2003) developed a method of bleaching spores and then using image-processing software to identify T. indica and distinguishing it from similar-looking species T. walkeri and T. horrida. Image analysis is also advocated by Deng et al. (2012), who were able to identify T. indica with an accuracy of 82.9% by studying micrographs and employing a support vector machine (SVM).
Notes on methods
Visual examination of dry seeds will only detect bunted seeds showing symptoms. It is necessary to use a wash to detect teliospores that contaminate the seed lots when bunted seeds break during harvest (Shamshad Begum and Mathur, 1989).
Size-selective sieving (Peterson et al., 2000) is routinely used in surveys in the USA. It is much faster than other methods being used elsewhere for detecting Karnal bunt teliospores. When the wash technique was applied to seed lots to which known numbers of T. indica teliospores had been added, all samples containing as few as five teliospores per 1000 seeds gave a positive recovery of T. indica. Three replicate 50 g samples can detect a contamination level of 1 spore/50 g of grain, with a 99% confidence. This size-selective sieving wash test is also incorporated into the EU Recommended Diagnostic Protocol for T. indica (Inman et al., 2003). From samples to which a single teliospore had been added, the spore was recovered 25% of the time (Castro et al., 1994).
Detection and identification methods using PCR are based on DNA sequence data of the closely related bunt species and are more specific and sensitive than diagnostic platforms based on image analysis and immunotechniques. These PCR methods (McDonald et al., 1999; Tan et al., 2009) can be performed directly on un-germinated spores, and so reduced the time of a diagnosis by at least 2 weeks which is the time estimated for spore germination.
Plant TradeTop of page
|Plant parts liable to carry the pest in trade/transport||Pest stages||Borne internally||Borne externally||Visibility of pest or symptoms|
|True seeds (inc. grain)||spores||Yes||Yes||Pest or symptoms usually invisible|
|Plant parts not known to carry the pest in trade/transport|
|Fruits (inc. pods)|
|Stems (above ground)/Shoots/Trunks/Branches|
Economic ImpactTop of page
The disease appeared in the Punjab (India) around 1930. It was epidemic there in 1953-1954 (Agarwal et al., 1976). Until 1970, sporadic outbreaks occurred every 2-3 years in the Punjab, Haryana and Uttar Pradesh regions, with a disease incidence of 0.1-10% and annual yield losses of about 0.2% (Munjal, 1976). In 1974 and 1975, the disease was epidemic in other regions (Himachal Pradesh, Tarai areas of Uttar Pradesh and the Gurudaspur area of the Punjab) with 50% infection on the cultivar HD-2000. In 1976-1977, low levels of infection (up to 3%) were observed on cultivars HD-1553 and HD-1593 in Uttar Pradesh, Punjab, Haryana, Rajasthan and Madhya Pradesh. When infection is severe, yield, seed quality and germination are adversely affected. Food grain is unacceptable when infection exceeds 3%. The disease is controlled by the use of resistant cultivars in infested areas so that high levels of infection are seldom reached at present.
In Mexico, where Karnal bunt appears regularly, direct losses are not very significant and do not exceed 1%. However, indirect costs to the Mexican economy are more significant due to quarantine measures which have to be applied for grain exports (OEPP/EPPO, 1991a; Brennan et al., 1992). In addition, the presence of Karnal bunt in Mexico has created a need for considerable extra precautions in the dispatch of cereal germplasm material by the International Maize and Wheat Improvement Center (CIMMYT).
For further information, see Gill et al. (1993).
Environmental ImpactTop of page
T. indica mainly attacks an annual crop (wheat). It does not affect other species in the natural environment. Its economic impact on cereal growing is not such as to modify land use. Accordingly, the environmental impact of this pest is nil.
Social ImpactTop of page
Karnal bunt is not toxic to humans, but infection by T. indica can affect the appearance and smell of grain products. Bunted grain smells like rotting fish due to the production of trimethylamine.
DiagnosisTop of page
Chesmore et al. (2003) developed a method of bleaching spores and then using image-processing software to identify T. indica. This method of diagnosis is useful for quickly distinguishing between T. indica and similar-looking species T. waleri and T. horrida. A one-tube fluorescent assay developed by Tan et al. (2009) is also able to distinguish T. indica from a variety of related pathogens. Image analysis is also advocated by Deng et al. (2012), who were able to identify T. indica with an accuracy of 82.9% by studying micrographs and employing a support vector machine (SVM).
Details on diagnosis with PCR and western blotting are given in Dumalasová (2006). The use of a two-step PCR protocol using FRET probes, which allows for the detection and identification of T. indica from only a small number of spores, is detailed in Tan and Murray (2006). The development and use of specific primers for detecting T. indica are described in Thirumalaisamy et al. (2011).
Detection and InspectionTop of page
A quarantine procedure for testing seeds of Triticum spp. for T. indica has been described by EPPO (OEPP/EPPO, 1991b). An EU recommended diagnostic protocol for the detection and identification of T. indica has also been produced (Inman et al., 2003; EPPO, 2007). This protocol has been enhanced for increased sensitivity and specificity by the adoption of more advanced technology (Tan et al., 2010). An updated draft is available from the IPPC website (www.ippc.int).
Crops for seed should be inspected during the growing season, though not while the crop is still green. Field inspection at maturity prior to harvest could be useful, although field symptoms are often very slight and the disease can be difficult to discern even at maturity. Any bunted seeds detected during field inspections should be examined under the microscope for the characteristic teliospores of T. indica. For quarantine purposes, seed should be tested for the presence of the fungus by the washing test (see Seedborne Aspects).
Direct visual observation for Karnal bunt (dry seed inspection) is regarded as insufficient for quarantine purposes because low levels of infection might pass undetected (Agrawal et al., 1986) and even minimal seed infections can substantially contaminate healthy seed lots (Aujla et al., 1988).
Being a non-systemic pathogen, it generally produces not more than four or five bunted kernels in each spike. Detection in the field is very unlikely and the first year of an outbreak usually goes undetected. For instance, detection of T. indica in a seed test sample in Arizona, USA, in 1996 was traced to wheat harvested in 1993 suggesting the pathogen had been present since 1992 (Rust et al., 2005). It had taken at least 4 years for the pathogen to be detected.
Similarities to Other Species/ConditionsTop of page
There are other Tilletia species that can be confused with T. indica and are commonly found in harvested grain. These include T. walkeri (ryegrass bunt pathogen; Castlebury and Carris, 1999), T.horrida (rice bunt pathogen; Caris et al., 2006) and T. ehrhartae (veldt grass pathogen; Pascoe et al., 2005). Refer to the review article by Tan et al. (2013) for a comparative summary of the morphology of teliospores of these species.
Prevention and ControlTop of page
High nitrogen applications and excessive irrigation favour the disease (Warham, 1986). Crop rotation may help to control the pathogen, but its value is questionable because T. indica can survive for up to 4 years in the soil. To prevent the spread of T. indica into previously unaffected areas, the use of disease-free seed is essential. The movement of farm machinery and soil from contaminated fields may also be restricted.
Chemical seed treatments have proved to be ineffective in killing the teliospores of T. indica on seeds of wheat, with the exception of mercurial compounds (Warham et al., 1989) which are banned in most countries. Bleach, in combination with heat treatment, is effective (MR Bonde, USDA- ARS, Foreign Disease-Weed Science Research Unit, personal communication, 1998); see also Smilanick et al. (1989). Carboxin + thiram, and chlorothalonil have been used as seed treatments in the USA and Mexico (B Goates, University of Idaho, USA, personal communication, 1998).
Foliar sprays of fungicides may be used to control the airborne inoculum of primary and secondary sporidia. Propiconazole was shown to be effective against natural infection in India (Singh et al., 1989). In Pakistan, propiconazole and bitertanol reduced the disease by 79 and 67%, respectively (Chandhry and Khan, 1990).
Intensive research has been carried out to breed resistant cultivars of bread wheat. It has been reported (Warham, 1986) that all commercially available cultivars in India are susceptible to the pathogen. However, Gill et al. (1986a, b) reported two cultivars to be either tolerant or field resistant. Cultivars of bread wheat, durum wheat and triticale tested by Warham (1988) were all equally susceptible to inoculation at the boot stage. In Mexico, a few lines of bread wheat showed moderate resistance (Singh and Dhaliwal, 1989).
For Europe and Australia, seeds of host plants should come from a pest-free area. Grain should come from a pest-free area or from a pest-free place of production (with testing of the harvested grain). CIMMYT (1989) uses the following procedures for germplasm material sent to other continents: production in areas free from T. indica; propiconazole sprays of seed-production plots; treatment of seed batches in a sodium hypochlorite bath; seed treatment with carboxin, captan and chlorothalonil.
ReferencesTop of page
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Distribution MapsTop of page
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