Xanthomonas axonopodis pv. manihotis (cassava bacterial blight)
- Taxonomic Tree
- Notes on Taxonomy and Nomenclature
- Distribution Table
- Risk of Introduction
- Hosts/Species Affected
- Host Plants and Other Plants Affected
- Growth Stages
- List of Symptoms/Signs
- Biology and Ecology
- Natural enemies
- Seedborne Aspects
- Plant Trade
- Detection and Inspection
- Similarities to Other Species/Conditions
- Prevention and Control
- Distribution Maps
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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- Xanthomonas axonopodis pv. manihotis (Bondar) Vauterin et al., 1995
Preferred Common Name
- cassava bacterial blight
Other Scientific Names
- Aplanobacter robici (Bouriquet) Magrou 1953
- Bacillus manihotis Berthet & Bondar 1915
- Bacillus manihotus Bondar 1912
- Bacterium manihotis Drummond & Hipolito 1941
- Bacterium robici Bouriquet 1947
- Phytomonas manihotis (Berthet & Bondar) Viegas 1940
- Pseudomonas manihotis (Berthet & Bondar) Savulescu 1947
- Xanthomonas campestris pv. manihotis (Bondar 1915) Dye 1978
- Xanthomonas manihotis (Berthet & Bondar) Starr 1946
International Common Names
- Spanish: bacteriosis de la yuca
- French: bacteriose du manioc
Local Common Names
- Brazil: bacteriose; leiteira; mancha angular; murcha; queima; sapeco
- Congo Democratic Republic: maladie des cierges
- Germany: Bakterielle: Maniok Blattkrankheit
- Indonesia: bacterial dieback
- Madagascar: feu
- Venezuela: bacteriosis
- XANTMN (Xanthomonas axonopodis pv. manihotis)
Taxonomic TreeTop of page
- Domain: Bacteria
- Phylum: Proteobacteria
- Class: Gammaproteobacteria
- Order: Xanthomonadales
- Family: Xanthomonadaceae
- Genus: Xanthomonas
- Species: Xanthomonas axonopodis pv. manihotis
Notes on Taxonomy and NomenclatureTop of page Cassava bacterial blight was first described in Brazil by Bondar (1912) and the bacteria named Bacillus manihotus, but without bacteriological description. In a joint paper with Arthaud-Berthet in 1915, the name of the latter was omitted in the title, but the error was mentioned in a later issue of the journal (Bradbury, 1986). For this reason Bondar is often cited alone as original author.
In a study on the nutrition of phytopathogenic bacteria, Starr (1946) listed this bacterium as Xanthomonas manihotis. With the revision of the plant pathogenic bacteria and the adoption of the pathovar system, Dye (1988) renamed X. manihotis as X. campestris pv. manihotis. On the basis of their phenetic and genomic characteristics Vauterin et al. (1995) proposed the name X. axonopodis pv. manihotis, which is now used as reference name in the recent list of plant pathogenic bacteria (Young et al., 1996).
Bacterium robici, the name used by Bouriquet (1939) for a bacterium associated in Madagascar with leaf spots and blight on cassava, is most probably a synonym.
DescriptionTop of page Individual colonies of purified isolates streaked on nutrient agar plates become visible after 24 h incubation at 28°C. After 48 h the colonies measure about 1 mm in diameter. They are whitish-grey to cream, raised, convex, smooth, shiny, with entire edges, at first hyaline, then opaque and turbid, and of viscous consistency.
Growth on potato dextrose agar and tryptone soya agar is faster than on nutrient agar. Colonies on tetrazolium medium (Kelman, 1954) are 8 mm in diameter after 6 days and are round, smooth with bright red centres and a narrow edge. They resemble the colony type of weakly pathogenic mutants of Pseudomonas solanacearum [Ralstonia solanacearum] (Maraite and Meyer, 1975).
On glucose-yeast-saline (GYS) plates (glucose 5 g, yeast extract 5 g, ammonium dihydrogen phosphate 0.5 g, dipotassium hydrogen phosphate 1.5 g, magnesium sulphate heptahydrate 0.2 g, sodium chloride 5 g, agar 20 g, water 1 l) colonies are distinctly convex and shiny. For some strains, rough variants are easily detected on this medium (Maraite et al., 1981).
On preparations from young cultures on nutrient agar, the bacteria appear as Gram-negative rods of 1.3 x 0.4 µm (1.0-1.75 x 0.28-0.6 µm), mostly singly or in pairs. Motile cells in a 0.5% glucose solution have a single polar flagellum.
DistributionTop of page Cassava bacterial blight was first described in 1912 in Brazil (Bondar, 1912). During the 1970s it was found in most of the cassava-growing areas of Central and South America, the Caribbean, Africa and Asia (Bradbury, 1986). Some of the recent reports are due to the increased attention being paid to cassava as a major staple food crop. Indeed, detailed field surveys in several countries revealed the widespread occurrence of cassava bacterial blight soon after it was first detected, indicating that it had been established long before the epidemic development was noticed (Maraite and Meyer, 1975; Persley, 1979a).
In some areas, the recent introduction and rapid spread of cassava bacterial blight have been demonstrated (Kwaje, 1982; Joseph and Elango, 1991), justifying strict quarantine measures on exchange of germplasm. No relation between differences in aggressiveness and geographical origin has been demonstrated. Isolates from Africa could be differentiated from X. axonopodis pv. manihotis of other origins by bacteriophage typing (Persley, 1980), and also by their low amylase activity which contrasts with the variable amylase production observed for strains from South America (Maraite et al., 1981; Alves and Takatsu, 1984). Both ribotyping and restriction fragment length polymorphism (RFLP) analysis using several DNA probes indicate a clonal population structure of X. axonopodis pv. manihotis in Africa (Verdier et al., 1993). This suggests that the epidemic development in Africa originated from a limited number of introductions.
Cassava bacterial blight has not yet been reported from countries in the southern Pacific.
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|
|India||Restricted distribution||EPPO, 2014|
|-Kerala||Present||Bradbury, 1986; IMI, 1993; EPPO, 2014|
|-Tamil Nadu||Present||Nair et al., 1981|
|-Java||Absent, unreliable record||Booth and Lozano, 1978; Bradbury, 1986; IMI, 1993; EPPO, 2014|
|-Sulawesi||Present||Tominaga et al., 1978; IMI, 1993; EPPO, 2014|
|-Sumatra||Present||Booth and Lozano, 1978; Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Japan||Present||Kawano et al., 1983; Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Malaysia||Restricted distribution||EPPO, 2014|
|-Peninsular Malaysia||Present||IMI, 1993; EPPO, 2014|
|Philippines||Present||Dedal et al., 1980; Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Taiwan||Present||Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Thailand||Present||Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Benin||Present||Akle and Gnouhoué, 1979; Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Burkina Faso||Present||Wonni et al., 2015|
|Burundi||Present||Autrique and Perreaux, 1989|
|Cameroon||Present||Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Central African Republic||Present||Daniel et al., 1981; Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Congo||Present||Daniel et al., 1979; Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Congo Democratic Republic||Present||Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Côte d'Ivoire||Present||Notteghem et al., 1980; Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Ghana||Present||Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Kenya||Widespread||Onyango and Ramos, 1979|
|Madagascar||Present||Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Malawi||Present||Kranz and Hammat, 1979; Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Mali||Present||Kranz and Hammat, 1979; Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Mauritius||Present||Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Niger||Present||Kranz and Hammat, 1979; Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Nigeria||Present||Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Rwanda||Present||Bradbury, 1986; IMI, 1993; EPPO, 2014|
|South Africa||Present||Manicom et al., 1981; Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Sudan||Present||Kwaje, 1982; Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Tanzania||Present||Nyango, 1979; Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Togo||Present||Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Uganda||Present||Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Mexico||Present||IMI, 1993; EPPO, 2014|
Central America and Caribbean
|Cuba||Present||Bradbury, 1986; Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Dominican Republic||Present||EPPO, 2014|
|Panama||Present||Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Trinidad and Tobago||Present||Elango, 1991; IMI, 1993; EPPO, 2014|
|Argentina||Present||Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Brazil||Widespread||IMI, 1993; EPPO, 2014|
|-Amazonas||Widespread||Takatsu et al., 1979|
|-Parana||Present||Portz et al., 2006|
|-Santa Catarina||Present||EPPO, 2014|
|Colombia||Present||Bradbury, 1986; IMI, 1993; EPPO, 2014|
|French Guiana||Present||IMI, 1993; EPPO, 2014|
|Venezuela||Present||EPPO 1999b; Bradbury, 1986; IMI, 1993; EPPO, 2014|
|Fiji||Present||Frison and Feliu, 1991|
|Guam||Restricted distribution||Wall, 1991; EPPO, 2014|
|Micronesia, Federated states of||Present||EPPO, 2014|
Risk of IntroductionTop of page RISK CATEGORY
ECONOMIC IMPORTANCE High
DISTRIBUTION Worldwide except Europe
SEEDBORNE INCIDENCE Moderate
SEED TRANSMITTED Yes
SEED TREATMENT Yes
OVERALL RISK Moderate
Notes on Phytosanitary risk
Very soon after the detection of cassava bacterial blight in Nigeria, the Inter-African Phytosanitary Council (CPI) released a warning (Addoh, 1972) and Zambia introduced quarantine measures on the movement of cuttings (Logan, 1974).
Despite its widespread occurrence, there are still areas which are disease-free, including the southern Pacific. Quarantine measures in this area are particularly relevant. Quarantine methods have been adopted by the Philippine Root Crop Research and Training Center (Anon., 1979).
Because of the occurrence of differences in aggressiveness among X. axonopodis pv. manihotis isolates (Maraite et al., 1981; Alves and Takatsu, 1984) it is advisable to take measures limiting the spread of the aggressive strains.
HabitatTop of page In Africa, X. axonopodis pv. manihotis is more prevalent in the savannah and the forest-savannah transition zones than in the forest zones, or even the forest galleries within the savannahs (Maraite and Meyer, 1975; Persley, 1980).
Hosts/Species AffectedTop of page In addition to cassava (Manihot esculenta), this pathogen attacks the wild species Manihot aipi, M. glaziovii and M. palmata.
Symptoms similar to those seen on cassava were induced on Euphorbia pulcherrima (poinsettia) and Pedilanthus tithymaloides following artificial infection with X. axonopodis pv. manihotis (Dedal et al., 1980). Van den Mooter et al. (1987), however, observed no or only very restricted lesions on E. pulcherrima inoculated with a wide range of X. axonopodis pv. manihotis isolates.
Amaranthus species, Panicum fasciculatum, Sida species, Sorghum halepense and several species belonging to the Euphorbiaceae have been identified as possible alternative hosts in Venezuela (Marcano and Trujillo, 1982). Epiphytic survival was demonstrated on several plant species under high relative humidity, heavy rainfall and thick cloud cover, but not under dry conditions (Marcano and Trujillo, 1984). However, Ikotun (1981) concluded from studies in Nigeria and Colombia that X. axonopodis pv. manihotis does not survive naturally on alternative hosts in and around cassava plants. This author observed a hypersensitive reaction by inoculation of E. repanda.
Symptomless epiphytic development or survival on cassava flowers was found consistently by Daniel and Boher (1981).
Host Plants and Other Plants AffectedTop of page
Growth StagesTop of page Flowering stage, Vegetative growing stage
SymptomsTop of page Large variations in the predominance and severity of the various symptoms described here are observed according to the location, season, aggressiveness of the occurring bacterial strains and the cassava cultivars (Maraite, 1993).
Leaves show dark-green to blue, water-soaked, angular spots (1-4 mm in diameter), limited by veinlets and irregularly distributed on the lamina. With time they frequently extend and coalesce along the veins or the edges of the leaf; the central portion turns brown and the water-soaked part often becomes surrounded by a chlorotic halo. The lesions appear as translucent spots when viewed against the light. Under a magnifying glass, small droplets of exudate oozing from the central portion of the lesion are visible on the lower surface of the leaves. The droplets, which first glisten creamy-white and later yellow, are easily dissolved by rain or dew; on drying they form a thin scale. Under favourable conditions (young leaves, high soil and air humidities), water-soaked pinpoint spots develop, scattered around young angular spots. The surrounding part of the lamina turns light brown and within 2-3 days extensive areas of the leaflets become withered not only towards the tip or border of the leaflet, but also towards the base. The affected parts show light-brown and green zonations as if they had been superficially burnt. These necrotic areas are not translucent, no bacterial exudate is observed and bacteria are absent or present in only very limited amounts at the borders of the extending blight lesions. A severe attack leads to premature drying and shedding of the leaves.
Stem and Growing Point
Under conditions of high humidity, infection may spread through the vascular bundles from the leaflets to the petiole and twigs or stems, with the formation of black and dark-brown streaks as well as exudation drops along the pathway of progression. These parts may also become infected directly through wounds, which may be due to removal of leaves for consumption or insect punctures.
On the unlignified twig or stem, a dark-green to black water-soaked area develops around the infection point. Large gummy exudation drops appear some distance away from the infection point, in the axis of vascular bundles, and one or a small number of leaves, located on the same side, show a sudden loss of turgidity, followed by rapid wilting and shrivelling. Afterwards the base of the petiole collapses, but the dried leaves generally remain attached for some time. All leaves located above those showing the first symptoms wilt progressively. Finally the unlignified tip dies, appearing as a wick on the withered stem end, giving the 'candle' symptom, while new shoots grow out lower down the stem. As the infection progresses towards the base of the stem these shoots often also become wilted leading to plant dieback. In the infected shoots, xylem vessels are brownish.
Under the microscope the vessels appear obstructed by bacteria, tyloses and mucilaginous substances. Lytic pockets develop around the protoxylem. The spread of these pockets causes rupture of the xylem ring, development of lytic pockets in the phloem, and later rupture of the fibre ring in the cortical collenchyma. Externally the latter pockets become visible as dark-green water-soaked spots and small black streaks, corresponding to altered laticifers. These spots swell, rupture and extrude a sticky white-yellow gum. In fully lignified stems or branches only internal vascular browning is visible. Infection may spread more than 50 cm below any external visual symptom.
Only exceptionally does infection reach the roots in some very susceptible cultivars, whose swollen roots may show dry, rotted spots around the dead vascular strands (Lozano, 1986).
On the green capsules, water-soaked expanding spots can also be observed. Heavily infected seeds from such fruit may be deformed, with corrugation of the testa and necrotic areas on the cotyledons and endosperm.
List of Symptoms/SignsTop of page
|Growing point / rot|
|Leaves / abnormal colours|
|Leaves / abnormal leaf fall|
|Leaves / necrotic areas|
|Leaves / wilting|
|Leaves / yellowed or dead|
|Roots / necrotic streaks or lesions|
|Stems / dieback|
|Stems / discoloration of bark|
|Stems / gummosis or resinosis|
|Stems / internal discoloration|
|Stems / ooze|
|Stems / wilt|
Biology and EcologyTop of page Transmission
X. axonopodis pv. manihotis spreads to new areas in infected, symptomless cuttings and seed. Within the crop, spread is mostly by rain splash (Persley, 1979b) and epidemics build up during the rainy season. Contaminated tools are also an important means of spread, because planting material is prepared simultaneously with the harvest, and the pathogen may be spread to uninfected cuttings when planting stakes are prepared from apparently healthy mature stems harbouring the pathogen. Movement of man and animals through plantations, especially during or after rain, may also contribute to pathogen spread (Lozano, 1986).
The possibility of transmission of X. axonopodis pv. manihotis by seed has been demonstrated by Persley (1979a) by a leaf-infiltration technique, using as inoculum the supernatant from seeds soaked in sterile water at 30°C for 2-4 h. By immunofluorescence (IF) and enzyme-linked immunosorbent assay (ELISA) on enriched embryonic extracts, Elango and Lozano (1980) confirmed seed transmission, with embryo infection varying from 0-40% depending on the sampling period. Clear positive reactions were obtained with 100,000 cells/ml in IF and with 10,000 cells/ml in ELISA. In histological studies combined with IF, Daniel and Boher (1981) determined levels of pathogen populations on and within the seeds (testa, caruncle, endosperm, cotyledon and embryo) of up to 1 million bacteria per seed.
However, the distribution of infected vegetative planting material from diseased plantations has been the main means of disseminating X. axonopodis pv. manihotis over long distances into Africa and Asia. It is also the source of primary infections in newly established plantations. This is due mainly to the lack of visible symptoms on lignified stems and to the ability of the pathogen to survive in invaded tissue for a very long time (Lozano and Nolt, 1989).
Infection and Disease Development
Entry into the plants occurs through stomata or epidermal wounds. In central and western Africa, stem infections have often been found to originate from punctures by the bug Pseudotheraptus devastans, an occasional vector of the bacterium (Maraite and Meyer, 1975). A similar role has been found for Anoplecnemis madagascariensis (Bouriquet, 1939) and the grasshopper Zonocerus variegatus.
Infection requires 12 hours of 90-100% relative humidity with temperatures of 22-26°C (Lozano, 1986). The optimum temperature for disease development in growth chambers is around 30°C (Maraite and Perreaux, 1979). In South America, temperature is considered to be a major factor affecting the severity of cassava bacterial blight. In regions such as the Amazon, with average minimum and maximum temperatures above 20 and 30°C, respectively, cassava bacterial blight is not an important disease, despite the high rainfall (Takatsu et al., 1979). Fluctuating day/night temperatures exceeding 10°C during the rainy season were found to enhance disease severity (Lozano, 1986; Joseph and Elango, 1991), which may explain the low incidence in forest zones with buffered temperature variations. In Africa, cassava bacterial blight is also more prevalent in the savannah and the forest-savannah transition zones than in the forest zones, or even the forest galleries within the savannahs (Maraite and Meyer, 1975; Persley, 1980). As well as the temperature fluctuation effect, this may also be due to increased susceptibility of cassava on the leached savannah soil: potassium fertilization to increase the potassium content of the leaves tends to reduce disease severity and enhances yield (Odurukwe and Arene, 1980).
Bacterial multiplication in the intercellular spaces of parenchymatous leaf or stem tissues occurs together with cell-wall degradation, possibly due to production of pectinases and cellulases (Maraite and Weyns, 1979; Dianese, 1985; Boher et al., 1995), and leads to the formation of lytic pockets filled with mucilaginous substances, including bacterial extracellular polysaccharide (Ikotun, 1984). Either directly or after disruption, the lytic pockets provide the bacteria with access to the xylem which results in systemic spread throughout the plant. The appearance and extent of the blight symptoms are associated with an increased leaf concentration of the blight-inducing toxin 3-methylthiopropionic acid (Perreaux et al., 1986) and other phytotoxic carboxylic acids produced by the bacteria (Ewbank, 1992). These toxins result from the induction of transamination and decarboxylation catabolism of methionine (Ewbank and Maraite, 1990) or of other amino acids in the bacteria (Ewbank, 1992).
The bacteria remain viable for many months in stems and gum, renewing activity in wet periods. The survival rate of the bacteria is higher in dry than in moist conditions (Persley, 1979b) and better in cassava debris than in rhizosphere soil of cassava weeds (Thaveechai et al., 1993). Daniel and Boher (1985) found large numbers of the pathogen on symptomless cassava leaves in farm crops and cultivar collections in the Congo during the rainy season, when spread of the disease occurs. In the dry season, the numbers fell to undetectable levels, but the presence of X. axonopodis pv. manihotis a few weeks before new symptoms appeared at the beginning of the next season suggested that it could survive as an epiphyte.
Natural enemiesTop of page
Seedborne AspectsTop of page Incidence
Persley (1979a) detected X. axonopodis pv. manihotis in 20 of 50 botanical seed samples of cassava using a leaf-infiltration technique, using as inoculum the supernatant from seeds soaked in sterile water at 30°C for 2-4 hours. The threshold of detection of the bacterium with this technique was 100,000 cells/ml. Using immunofluorescence (IF) and ELISA on enriched embryonic extracts, Elango and Lozano (1980) demonstrated embryo infection varying from 0 to 40%; this depended on the sampling time. Clear positive reactions were obtained with 100,000 cells/ml in IF and with 10,000 cells/ml in ELISA. In histological studies combined with IF, Daniel and Boher (1981) determined levels of pathogen populations on and within the seeds (testa, caruncle, endosperm, cotyledon and embryo) of up to 1,000,000 bacteria/seed.
Effect on Seed Quality
Heavily infected seeds from infected fruit may be deformed, with necrotic spots on the cotyledons and endosperm (Lozano, 1986; Frison and Feliu, 1991). Very few of these seeds germinate (Lozano, 1986).
X. axonopodis pv. manihotis was recovered from seedlings grown from infected seeds that were pre-germinated in the laboratory then transplanted to sterile sand in the greenhouse (Elango and Lozano, 1980). Infected seeds are recognized as an important means of transmitting the disease (Lozano, 1986).
The distribution of infected vegetative planting material of cassava from diseased plantations has been the main means of disseminating X. axonopodis pv. manihotis over long distances into Africa and Asia. It is also the source of primary infections in newly established plantations. This is due mainly to the lack of visible symptoms on lignified stems and to the ability of the pathogen to survive in invaded tissue for a very long time (Lozano and Nolt, 1989).
Soaking of infested botanical seed in hot water at 60°C for 20 minutes followed by drying in shallow layers at 30°C overnight or at 50°C for 4 hours reduced the number of bacteria to less than the minimum detectable level without appreciably reducing germination (Persley, 1979a).
The FAO/IBPGR technical guidelines for the safe movement of cassava germplasm recommend visual inspection of the seeds, density selection, followed by treatment of the seeds by immersing them in water and heating in a microwave oven at full power until the water temperature reaches 73°C and then immediately removing the water. If a microwave oven is not available, a dry heat treatment for 2 weeks at 60°C is recommended (Frison and Feliu, 1991). A subsequent thiram dust treatment reduces seed reinfestation. Lozano and Nolt (1989) mentioned 77°C instead of 73°C for the microwave treatment.
Seed Health Tests
Leaf-infiltration technique (Persley, 1979a)
- Soak seeds in sterile water at 30°C for 2-4 h.
- Infiltrate the supernatant into the leaf of 1-month-old plants.
- Incubate at 30°C at high relative humidity for 1-2 weeks before looking for water-soaked angular spots.
Serology (Elango and Lozano, 1980)
The pathogen has been detected in seeds by ELISA and immunofluorescence techniques.
Dot-blot (Verdier and Mosquera, 1999)
A dot-blot assay was developed for the detection of X. axonopodis pv. manihotis, the causal agent of cassava bacterial blight. An 898 bp DNA fragment unique to X. axonopodis pv. manihotis strains was cloned and evaluated as a diagnostic DNA probe. The DNA probe (p898) hybridized with the total DNA of 362 X. axonopodis pv. manihotis strains tested but not with any of the 40 other Xanthomonas strains tested or with saprophytes associated with the cassava plant. A quick colony hybridization procedure was developed to identify X. axonopodis pv. manihotis strains. The probe also detected X. axonopodis pv. manihotis strains in crude extracts of leaf and stem lesions, and in cassava fruits and sexual seeds that were naturally infected. The overall sensitivity of the dot blot method for detecting X. axonopodis pv. manihotis in stem and leaf samples was about 103 colony-forming units per reaction. Because the dot-blot hybridization technique is sensitive and rapid, it can be easily used for culture indexing.
Ojeda and Verdier (2000) have developed a PCR assay to detect the pathogen in cassava true seeds. The technique is specific, sensitive, and rapid.
Selective medium for X. axonopodis pv. manihotis (Fessehaie et al., 1999).
The semi-selective agar medium, designated cefazolin trehalose agar (CTA) medium, contains (per litre) 3.0 g of K2HPO4, 1.0 g of NaH2PO4, 0.3 g of MgSO4.7H2O, 1.0 g of NH4Cl, 9.0 g of D(+)-trehalose, 1.0 (+)-glucose, 1.0 g of yeast extract, 0.025 g of cefazolin, 0.0012 g of lincomycin, 0.0025 g of phosphomycin, 0.25 g of cycloheximide and 14.0 g of agar. When X. axonopodis pv. manihotis occurs in high concentrations in diseased tissue, the standard yeast trehalose glucose agar medium supplemented with 250 µg of cycloheximide per ml appears to be satisfactory. The newly developed CTA medium should prove useful for control strategies to identify and remove infected planting material of cassava, as well as for basic ecological studies of the pathogen.
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|
|Bulbs/Tubers/Corms/Rhizomes||Yes||Pest or symptoms usually invisible|
|Flowers/Inflorescences/Cones/Calyx||Yes||Pest or symptoms usually visible to the naked eye|
|Fruits (inc. pods)||Yes||Pest or symptoms usually visible to the naked eye|
|Leaves||Yes||Pest or symptoms usually visible to the naked eye|
|Roots||Yes||Pest or symptoms usually visible to the naked eye|
|Seedlings/Micropropagated plants||Yes||Pest or symptoms usually invisible|
|Stems (above ground)/Shoots/Trunks/Branches||Yes||Yes||Pest or symptoms usually visible to the naked eye|
|True seeds (inc. grain)||Yes||Pest or symptoms usually invisible|
|Wood||Yes||Pest or symptoms usually visible to the naked eye|
|Plant parts not known to carry the pest in trade/transport|
|Growing medium accompanying plants|
ImpactTop of page Cassava bacterial blight is a major constraint on cassava cultivation and losses can be extremely severe after introduction of the pathogen, or possibly of more aggressive strains, in a region where highly susceptible cultivars are grown. In 1973, 1 year after the first report of cassava bacterial blight in Nigeria, estimated yield losses were 75% (Ezelio, 1977). Crop losses as high as 90-100% were observed in some parts of Uganda, 2 years after the disease was first recorded (Otim-Nape, 1980). In Zaire, the epidemics between 1971 and 1973 in the Kasaï and Bandundu provinces led to severe starvation, because of the importance of cassava roots and leaves as staple foods in these areas (Maraite and Meyer, 1975). In field experiments with X. axonopodis pv. manihotis-free and X. axonopodis pv. manihotis-infected stem cuttings, Otim-Nape (1983) observed a reduction in fresh tuber yield from 40.1 to 26.6 t/ha. By comparing the yield of susceptible clones to that of resistant ones under natural infection in Colombia during 1974-82, Umemura and Kawano (1983) observed an 18-92% yield reduction, depending on locality, planting time and degree of simultaneous infection by Elsinoë brasiliensis.
Under the selection pressure of severe cassava bacterial blight epidemics, the most susceptible clones are eliminated from the mixture generally planted by the smallholders, and overall severity decreases. Based on a CIAT survey of five major cassava-producing zones in Colombia, the estimated reduction in national production was, nevertheless, still 6.64% in 1973 (Lourido, 1974).
DiagnosisTop of page Cut small pieces of tissue from the edge of lesions and mount in a drop of water. Look for bacterial streaming under a microscope equipped with phase-contrast, dark-field or Abbe condenser with an iris diaphragm.
Crush a piece of infected tissue in a drop of sterile water, incubate for 30 min and streak a loopful onto nutrient agar plates. After 2-4 days' incubation at 28°C, analyse for the occurrence of typical whitish-grey and convex colonies. Subculture on GYS for confirmation of purity and store on yeast-dextrose-chalk-agar (YDCA) (Dye, 1962) slants or lyophilized. Rapid loss of viability is sometimes observed on peptone-containing media.
By Gram staining, X. axonopodis pv. manihotis (Gr-) can rapidly be distinguished from whitish saprophytic Bacillus species (Gr+). The Kovacs' oxidase test is most useful for distinguishing X. axonopodis pv. manihotis (negative or slow positive) from Ralstonia solanacearum (rapidly positive).
For further characteristics see Bradbury (1977) and Van den Mooter et al. (1987). Some strains show a week starch hydrolytic activity (Maraite et al., 1981).
Rapid and simple pathogenicity tests can be performed, preferably on young cassava shoots in field plots or in glasshouses, at temperatures between 25 and 35°C (Maraite and Perreaux, 1979). A sample is collected by passing a needle through a 3-day-old culture and using it to puncture the leaflets or the stem between the third and the fourth leaf from the top. High humidity after inoculation is useful but not essential. Between 1 and 4 weeks after inoculation, the leaf and stem are checked for the development of symptoms.
Seed Health Tests
No standardized method for seed testing has been published. The leaf-infiltration technique used by Persley (1979a) for demonstration of X. axonopodis pv. manihotis transmission by seed is described under Detection and Inspection Methods. This very simple technique has a sensitivity similar to immunofluorescence and only slightly lower than ELISA. The immunological techniques require special laboratory equipment and temperature-sensitive reagents.
Detection and InspectionTop of page Cassava bacterial blight infections are most conspicuous at the end of the rainy season and the beginning of the dry season. First observations should focus on the presence of abnormal leaf shedding and on drying of leaves still attached to the stem. Inspect the leaves for the presence of angular leaf spots and blighted areas. On the green part of the stem look for the presence of dark-brown to black streaks, for abnormal swellings of the bark and gummy exudates. Cut the outer part of the bark between wilted leaves and exudates and look for black stripes and lytic pockets in the phloem. After peeling the bark, slit the xylem and look for vascular browning.
Inspect the green capsules for the presence of dark-green water-soaked expanding spots.
For seed inspection, soak seeds in sterile water at 30°C for 2-4 h. Infiltrate the supernatant into the leaf of 1-month-old plants and incubate at 30°C at high relative humidity for 1-2 weeks before looking for water-soaked angular spots (Persley, 1979a).
For inspection of lignified stem cuttings remove 5 cm from the upper end and look for vascular browning.
Similarities to Other Species/ConditionsTop of page X. cassavae causes angular leaf spots very similar to those induced by X. axonopodis pv. manihotis. However, these spots do not evolve into blight areas, and produce very tiny bright yellow exudates (Maraite and Perreaux, 1979).
A bacterial soft rot of cassava roots linked to vascular browning in the stem and wilting of the leaves was reported by Schwarz from Indonesia (1926). The bacterium was identified as Pseudomonas solanacearum [Ralstonia solanacearum] on the basis of pathogenicity on tomatoes. Similar observations had been made in 1922 by Palm on Manihot glaziovii. Nishiyama et al. (1980) distinguished bacterial wilt caused by R. solanacearum, characterized by drooping, severe defoliation with remaining immature leaves at the top, and usually accompanied by affected roots, from bacterial dieback without root rot caused by X. axonopodis pv. manihotis. Bacterial wilt of cassava caused by R. solanacearum biovars 3 and 4 is confined to Lampung province, Sumatra, and parts of Java (Machmud, 1986).
Wilt symptoms and vascular browning in cassava can also be caused by Verticillium dahliae (Maraite and Meyer, 1976). However, lytic pockets and gummy exudates on the green stem are not induced by this fungus.
Prevention and ControlTop of page Introduction
Losses can be greatly reduced by a combination of measures taken within the perspective of IPM (Lozano, 1986).
In areas where cassava bacterial blight does not yet occur, great care must been taken in the introduction of germplasm. Vegetative propagated material must be introduced as meristem culture multiplied in vitro and certified disease-free. Botanical seed should originate from areas unfavourable for disease development, be heat-treated and planted in quarantine. Details can be found in the FAO/IBPGR technical guidelines for the safe movement of cassava germplasm (Frison and Feliu, 1991).
Cultural Control and Sanitary Methods
In areas where cassava bacterial blight is already widespread, disease incidence can be reduced by the use of clean planting material. Cuttings should be taken only from plantations that have been found to be free of the disease by inspections at the end of the rainy season. In cases of sporadic occurrence of the disease, great care must be taken in collecting cuttings only from healthy plants and from the most lignified portion of the stem, up to 1 m from the base, combined with visual inspection for the absence of vascular browning. Tools should be regularly disinfected using a bactericide.
Infected clones can be cleaned by rooting bacteria-free stem tips in conditions unfavourable for infection or by meristem cultures in vitro.
Crop rotation and fallowing proved very successful when the new crop was planted with uninfected cuttings. All infected plant debris and weeds on which epiphytic survival may occur should be removed and burned or incorporated into the soil. Rotation or fallowing should last at least one rainy season.
In some areas, planting towards the end of the rainy season instead of at the beginning will delay epidemic development during the growing period and so reduce yield loss. Intercropping cassava with maize or melon has been reported to reduce cassava bacterial blight significantly (Ene, 1977).
In potassium-deficient soils, increasing the potassium content of the leaves by fertilization tends to reduce disease severity (Odurukwe and Arene, 1980).
Soaking of infested botanical seed in hot water at 60°C for 20 min, followed by drying in shallow layers at 30°C overnight or at 50°C for 4 h, reduced the number of bacteria to less than the minimum detectable level without appreciably reducing germination (Persley, 1979a).
The FAO/IBPGR technical guidelines for the safe movement of cassava germplasm recommend visual inspection of the seeds, density selection, followed by treatment of the seeds by immersing them in water and heating in a microwave oven at full power until the water temperature reaches 73°C and then immediately pouring the water off. If a microwave oven is not available, a dry heat treatment for 2 weeks at 60°C is recommended (Frison and Feliu, 1991). A subsequent thiram dust treatment reduces seed re-infestation. Lozano and Nolt (1989) mentioned 77°C instead of 73°C for the microwave treatment.
Clear differences in host-plant resistance occur, especially with regard to stem infection and wilt; use of resistant genotypes is a major control strategy. Resistance to cassava bacterial blight appears to be due to several genes, mainly with additive effects but also to some extent with non-additive effects; resistance appears to be recessive to susceptibility (Hahn, 1979; Umemura and Kawano, 1983). A variation in aggressiveness, but no clear-cut pathogenic specialization, is observed among X. axonopodis pv. manihotis isolates from various countries, and also among those from a single country (Maraite et al., 1981; Alves and Takatsu, 1984). A strong genotype-environment interaction is often observed. Host-plant resistance is sustained by adequate fertilization.
Ninety-three varieties of M. esculenta were assessed by amplified fragment length polymorphisms (AFLPs) for genetic diversity and for resistance to X. axonopodis pv. manihotis. AFLP analysis was performed using two primer combinations, and a 79.2% level of polymorphism was found. The phenogram obtained showed between 74 and 96% genetic similarity among all cassava accessions analysed. The results demonstrate that resistance to X. axonopodis pv. manihotis is broadly distributed in cassava germplasm and that AFLP analysis is an effective and efficient means of providing quantitative estimates of genetic similarities among cassava accessions (Sanchez et al., 1999).
Foliar application of Pseudomonas fluorescens and P. putida has been shown significantly to reduce leaf infection by X. axonopodis pv. manihotis (Lozano, 1986). However, biological control has not yet gained practical acceptance.
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Distribution MapsTop of page
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