Xanthomonas axonopodis pv. phaseoli (bean 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. phaseoli (Smith 1897) Vauterin et al. 1995
Preferred Common Name
- bean blight
Other Scientific Names
- Bacillus phaseoli Smith 1897
- Bacterium phaseoli (Smith 1897) Smith 1905
- Bacterium phaseoli var. fuscans (Burkholder) Okabe 1933
- Phytomonas phaseoli (Smith 1897) Bergey et al. 1923
- Phytomonas phaseoli var. fuscans Burkholder 1930
- Pseudomonas phaseoli (Smith) Smith 1901
- Pseudomonas phaseoli var. fuscans (Burkholder) Stapp 1935
- Xanthomonas campestris pv. phaseoli (Smith 1897) Dye 1978
- Xanthomonas campestris pv. phaseoli var. fuscans (Burkholder 1930) Starr & Burkholder 194
- Xanthomonas campestris pv. phaseoli var. indica Uppel et al.
- Xanthomonas fuscans (Burkholder) Burkholder 1959
- Xanthomonas phaseoli (ex Smith 1897) Gabriel at al. 1989
- Xanthomonas phaseoli var. fuscans (Burkholder) Starr & Burkholder 1942
- Xanthomonas phaseoli var. indica Uppal, Patel & Nikam 1946
International Common Names
- English: bacterial blight of bean; bacterial leaf pustule; common bacterial blight; common blight; fuscous blight of bean
- Spanish: mancha bacterial; quema bacteriana de las judias; quemaz bacterial commún; tizón común
- French: bactériose du haricot; brûlure bactérienne du haricot
- Portuguese: crestamento commun
Local Common Names
- Germany: Bacterielle brandfleckigkeit; Fettfleckenkrankheit
- India: sem ki jeewanik angmaari
- Italy: batteriosa del fagiola; nebbia batterica
- Japan: hayake-byo
- Turkey: fasulye adi yaprak yanikligi
- XANTPH (Xanthomonas axonopodis pv. phaseoli)
Taxonomic TreeTop of page
- Domain: Bacteria
- Phylum: Proteobacteria
- Class: Gammaproteobacteria
- Order: Xanthomonadales
- Family: Xanthomonadaceae
- Genus: Xanthomonas
- Species: Xanthomonas axonopodis pv. phaseoli
Notes on Taxonomy and NomenclatureTop of page
X. axonopodis pv. phaseoli can be divided into three groups. X. axonopodis pv. phaseoli var. fuscans produces a diffusible brown pigment in culture that is enhanced by tyrosine (Goodwin and Sopher, 1994a), X. axonopodis pv. phaseoli var. indica produces a diffusible brown pigment in culture that is not enhanced by tyrosine (Bradbury, 1986) and X. axonopodis pv. phaseoli does not produce a brown pigment in culture. The brown pigment of X. axonopodis pv. phaseoli var. fuscans results from the secretion and oxidation of homogenistic acid, an intermediate in the tyrosine catabolic pathway (Goodwin and Sopher, 1994a). The fuscans variety is widespread and causes similar symptoms to X. axonopodis pv. phaseoli and was not considered to have any special taxonomic status (Bradbury, 1986). Common blight caused by the fuscans variety is known as fuscous blight of beans. There have been a number of reports indicating that the X. axonopodis pv. phaseoli var. fuscans isolates tend to be more pathogenic to beans, such as the demonstration that eight of the ten most aggressive isolates to Phaseolus vulgaris were the fuscous type (Opio et al., 1996). Several studies using RFLPs (Lazo et al., 1987), DNA-DNA homology (Hilderbrand et al., 1990), amplified DNA polymorphisms (Xue and Goodwin, 1993), RAPDs (Birch et al., 1997) and macrorestriction DNA polymorphisms (Chan and Goodwin, 1999) have consistently demonstrated that the fuscans strains are genetically distinct. It has been proposed that the fuscans variety should be considered a separate subspecies from those not producing a brown pigment in culture (Chan and Goodwin, 1999).
DescriptionTop of page
See also Hayward and Waterston (1965a, b).
DistributionTop of page
Records for Eritrea and Bermuda cited in previous editions of the Compendium (IMI, 1996) were based on old references (Petri, 1932; Waterston, 1947) and no recent records could be found for these countries.
Distribution TableTop of page
The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.Last updated: 12 May 2022
|Continent/Country/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
|Central African Republic||Present|
|Congo, Democratic Republic of the||Present|
|Morocco||Absent, Formerly present|
|South Africa||Present, Widespread|
|Hong Kong||Present, Few occurrences|
|Israel||Absent, Formerly present|
|Malaysia||Present, Few occurrences|
|Taiwan||Present, Few occurrences|
|United Arab Emirates||Present|
|Yemen||Present, Few occurrences|
|Austria||Present, Few occurrences|
|Bosnia and Herzegovina||Present|
|Czechia||Present, Few occurrences|
|Union of Soviet Socialist Republics||Present|
|Finland||Absent, Unconfirmed presence record(s)|
|Germany||Present, Localized||First reported: 196*|
|Netherlands||Present, Few occurrences||present, only in some areas where host plants are grown, at low prevalence.|
|Norway||Absent, Invalid presence record(s)|
|Poland||Present, Few occurrences|
|-Russian Far East||Present|
|-Southern Russia||Present, Localized|
|Serbia and Montenegro||Present, Localized|
|Spain||Present, Few occurrences|
|Sweden||Absent, Unconfirmed presence record(s)|
|Switzerland||Present, Localized||First reported: 198*|
|Bermuda||Absent, Unconfirmed presence record(s)|
|-Prince Edward Island||Present|
|Saint Vincent and the Grenadines||Present|
|Trinidad and Tobago||Present, Few occurrences|
|United States||Present, Localized|
|-Colorado||Present, Few occurrences|
|-Nebraska||Present, Few occurrences|
|-Texas||Present, Few occurrences|
|-Wyoming||Present, Few occurrences|
|-New South Wales||Present, Widespread|
|New Zealand||Present, Few occurrences|
|Papua New Guinea||Present|
|-Rio de Janeiro||Present|
|-Rio Grande do Sul||Present|
|Chile||Present, Few occurrences|
|Uruguay||Present, Few occurrences|
Risk of IntroductionTop of page
ECONOMIC IMPORTANCE High
SEEDBORNE INCIDENCE High
SEED TRANSMITTED Yes
SEED TREATMENT Yes
OVERALL RISK Low
Notes on phytosanitary risk
The distribution of infected seed has already spread common blight throughout the world, and X. axonopodis pv. phaseoli is probably present almost everywhere that susceptible beans are grown.
Hosts/Species AffectedTop of page
Helianthus annuus, Lupinus polyphyllus, Phaseolus acutifolius and P. coccineus have been reported as hosts by artificial inoculation (Bradbury, 1986). Senna (Cassia) hirsuta and Digitaria scalarum are reported as symptomless hosts (Opio et al., 1996).
Host Plants and Other Plants AffectedTop of page
|Lablab purpureus (hyacinth bean)||Fabaceae||Other|
|Lupinus polyphyllus (garden lupin)||Fabaceae||Main|
|Phaseolus coccineus (runner bean)||Fabaceae||Main|
|Phaseolus lathyroides (Phasey bean)||Fabaceae||Main|
|Phaseolus lunatus (lima bean)||Fabaceae||Other|
|Phaseolus vulgaris (common bean)||Fabaceae||Main|
|Pisum sativum (pea)||Fabaceae||Main|
|Vigna aconitifolia (moth bean)||Fabaceae||Main|
|Vigna mungo (black gram)||Fabaceae||Other|
|Vigna radiata (mung bean)||Fabaceae||Other|
|Vigna umbellata (rice bean)||Fabaceae||Main|
|Vigna unguiculata (cowpea)||Fabaceae||Other|
Growth StagesTop of page
SymptomsTop of page
Stem infection is less common. It begins as a water-soaked spot, which becomes a reddish-brown lesion, usually without chlorosis. Stem girdling may develop at the cotyledonary node. The bacteria can invade the xylem, and wilting may occur if sufficient bacterial numbers develop in the xylem.
Pod lesions begin as water-soaked spots which become sunken and dark to red brown. Under humid conditions, a yellowish bacterial ooze will develop from the lesion.
Severely infected seed may be shrivelled and show poor germination or produce weakened plants. On white-seeded varieties, yellow or brown spots may appear on the seed coat, particularly near the hilum area. On dark-seeded varieties, this discoloration is not visible. Infected seeds may also be symptomless (Yoshii, 1980).
List of Symptoms/SignsTop of page
|Fruit / lesions: black or brown|
|Fruit / lesions: on pods|
|Fruit / ooze|
|Leaves / abnormal colours|
|Leaves / necrotic areas|
|Seeds / discolorations|
|Seeds / lesions on seeds|
|Stems / discoloration of bark|
|Stems / internal red necrosis|
|Stems / wilt|
Biology and EcologyTop of page
X. axonopodis pv. phaseoli can survive in seed, infected plant debris and epiphytically on host and non-host plants (Saettler, 1991). Seed is particularly important in long-distance spread and for introducing X. axonopodis pv. phaseoli into new areas. Seed can also be important in long-term survival as X. axonopodis pv. phaseoli can remain viable in bean seeds for as long as 30 years (Trujillo and Saettler, 1980). Approximately 1000 to 10,000 bacteria per seed is the minimum needed to produce infected plants under field conditions (Weller and Saettler, 1980). Infection of flower buds and young pods can result in the transmission of X. axonopodis pv. phaseoli through the vascular system to the seed, leading to internal infection (Aggour et al., 1989). Plants grown from infected seed may have lesions of cotyledons, nodes or primary leaves which will then serve as sites for the spread of secondary infection. Seed may also become contaminated from affected plant residues during harvest.
Plant debris is probably more important as a source of inoculum in warmer climates where several susceptible crops can be planted in one year (Saettler, 1989). However, X. axonopodis pv. phaseoli populations will drop sharply in plant debris as other micro-organisms colonize and break down the material (Saettler, 1991). If infected bean debris is tilled into the soil it will greatly increase the rate of decay of the tissue and enhance the decline in X. axonopodis pv. phaseoli populations since it appears to be unable to survive for long in soil as free bacteria.
X. axonopodis pv. phaseoli can grow on the surface of both host and non-host plants (Cafati and Saettler, 1980). Growth on host-plant surfaces may provide a sizeable population with the potential to infect under appropriate conditions. The importance of epiphytic populations on non-host plants is not clear and Angeles-Ramos et al. (1991) found that weeds may not be not an important source of X. axonopodis pv. phaseoli inoculum because the pectolytic xanthomonads isolated from weeds were non-pathogenic to beans.
X. axonopodis pv. phaseoli can be spread by planting infected seed: plants grown from infected seed usually show lesions on the cotyledon or primary leaves (Zaumeyer and Thomas, 1957). Bacteria in lesions, infected plant debris and epiphytic populations can be spread by splashing or wind-blown rain. Overhead irrigation can be very effective in secondary spread of the bacteria. Surface irrigation water may also spread the bacteria (Steadman et al., 1975).
X. axonopodis pv. phaseoli can also be disseminated on the bodies of insects, such as Diaprepes abbreviatus and Cerotoma ruficornis, which may also create wounds by feeding, thus producing favourable sites for infection (Kaiser and Vakili, 1978). Whiteflies (Bemisia tabaci), grasshoppers (Melanoplus sp.), Mexican bean beetles (Epilachna varivestris) and leaf miners have also been reported to transmit X. axonopodis pv. phaseoli (Zaumeyer and Thomas, 1957; Sabet and Ishag, 1969).
Infection occurs through natural openings and wounds. Severe epidemics can occur following storms with wind-blown rain, which can force the bacteria through openings, such as stomata, into the intercellular spaces. Wounds due to hail or insect feeding can create favourable sites for infection. Once inside the plant, X. axonopodis pv. phaseoli multiplies rapidly in the intercellular spaces and it can take as little as 10-14 days from initial infection until secondary spread occurs (Saettler, 1991). The optimal temperatures for disease development are 28-32°C (Saettler, 1989). Relatively high populations, 100,000 to 1,000,000 c.f.u./g of host tissue, are required before symptoms become visible (Gilbertson and Maxwell, 1992). Plants appear to be more susceptible in the reproductive stage than in the vegetative stage. As the bacterial population increases, it can ooze onto the leaf surface and be spread further by water. The polysaccharide covering of the bacteria helps to prevent desiccation (Wilson et al., 1965) and may assist the bacteria to attach to plant surfaces. The bacteria can also enter the vascular system of many varieties of beans and then spread systemically in the plant (Zaumeyer and Thomas, 1957). Wilting can result from vascular infection. Bacteria in the vascular system can also enter the developing pods and pass into the seeds (Aggour et al., 1989). Infection of the seed coats can also occur from pod infections.
Natural enemiesTop of page
|Natural enemy||Type||Life stages||Specificity||References||Biological control in||Biological control on|
Seedborne AspectsTop of page
X. axonopodis pv. phaseoli (Xap) and X. axonopodis pv. phaseoli var fuscans (Xapf) is extensively seedborne on Phaseolus spp. A survey of navy bean (P. vulgaris) seeds showed that approximately 35% had internal contamination with X. axonopodis pv. phaseoli (Saettler and Perry, 1972). 34 bean seed lots were collected in Paraná, Brazil from 1998 to 1999 and, when studied, 50% of the seedlots were infected with Xap with incidences ranging from 0.1 to 1.7% (Torres et al. 2009). Assays conducted on seed collected from naturally contaminated commercial fields in Serbia indicated presence of Xap in 20 of the 23 cultivars from which collections were made from (Popovic et al., 2010). Five out of seven seed samples collected in Egypt tested positive for the presence of Xap (Abd-Alla et al., 2010). Incidences of seed infection by X. axonopodis pv. phaseoli as high as 16.1%, with populations of the bacterium averaging between 100,000 and 1,000,000,000 c.f.u./100 seeds, were recorded in farmers commercial and research seeds in Africa (Opio et al., 1993). Approximately 1 in 10,000 seeds is capable of causing an outbreak of blight (Sutton and Wallen, 1970), or 1000 to 10,000 bacteria per seed is the minimum needed to produce infected plants under field conditions (Weller and Saettler, 1980).
Infection of flower buds and young pods can result in the transmission of X. axonopodis pv. phaseoli through the vascular system to the seed, leading to internal infection (Aggour et al., 1989). X. axonopodis pv. phaseoli is a systemic pathogen that penetrates the ovule through the funiculus. It enters the sutures of the pod from the vascular system of the pedicel and passes into the raphe leading into the seed coat. Bacteria also may enter though the micropyle (Burkholder, 1921; Zaumeyer, 1930). X. axonopodis pv. phaseoli is capable of long-term survival having been recovered from 15 year-old bean seeds (Schuster and Sayre, 1967).
Surface and internal populations of X. axonopodis pv. phaseoli were investigated on and in flower buds, flowers and pods of five susceptible (Pinto UI114, Cranberry Taylor Hort, Charlevoix, Black Magic and C20) and two resistant (Valley and I84100) Phaseolus vulgaris genotypes (Mabagala, 1997). Plants were grown in the field in Tanzania, and artificially inoculated at the seedling stage (18 day old). The pathogen was recovered in high numbers from flower buds, flowers, pods and seed of resistant and susceptible genotypes. Significant differences (P=0.05) in population levels of X. axonopodis pv. phaseoli between stages of reproductive tissue development were observed. Infected seed from resistant genotypes had no visible symptoms. It is suggested that these seeds may play an important role in the epidemiology of common bacterial blight disease because they are difficult to detect and may occur at low frequency in seed lots.
Effect on Seed Quality
Severely infected seed may be shrivelled and show poor germination or produce weakened plants On white-seeded varieties, yellow or brown spots may appear on the seed coat, particularly near the hilum area. On dark-seeded varieties, this discoloration is not visible (Zaumeyer, 1957). Infected seeds may also be symptomless (Yoshii, 1980).
Seeds are the primary source of inoculum for X. axonopodis pv. phaseoli. Plants grown from infected seeds frequently bear lesions on the cotyledons or primary nodes. These lesions enlarge and under humid conditions, slimy masses of bacteria accumulate on the leaf surface. These are then spread to healthy plants (Zaumeyer, 1957). Approximately 1000 to 10,000 bacteria per seed is the minimum needed to produce infected plants under field conditions (Weller and Saettler, 1980). A range of tolerances, ranging from 0 seeds infected in samples of 4000-45,000, are used in seed quality programmes in US seed companies to ensure that the pathogen is not transmitted by commercial seed lots (Maddox, 1997).
The most effective survival mechanism for Xap is to inhabit infected bean seed (Cafati & Saettler 1980a, Gilbertson et al., 1990; Arnaud-Santana et al., 1991; Opio et al., 1993), within which bacteria may survive for up to thirty six years (Allen et al., 1998). Marques et al. (2005) indicated that the survival of seedborne X. axonopodis pv. phaseoli was reduced from 64 to 36-37% incidence during the first 6 months; however, seed stored at –18 and 5 ºC maintained the contamination rate at 30 and 60 months, and it was concluded that the optimum temperatures for storing seed is similar to conditions favourable for Xap longevity (Marques et al., 2005). Seed contamination may be internal or external (Saettler 1989; Allen et al., 1998) and even symptomless (Thomas & Graham, 1952; Weller and Saettler, 1980a), which has serious implications for seed certification schemes. It has, however, been indicated that the development of X. axonopodis pv. phaseoli epidemics are depended on the level of horizontal resistance and climatic conditions rather than the population size of Xap in bean seeds (He, 2010). He (2010) found that Xap had a significant higher percentage of seed to seedling transmission than Xap. Although both variants reduced seedling emergence , Xapf was more severe and the incidence of typical CBB lesions were higher in seedlings infested with Xapf than Xap.
Infected crop residues and other susceptible hosts are also sources of inoculum for X. axonopodis pv. phaseoli (Zaumeyer, 1957; Khlaif and Qadous, 1995). Conflicting reports exist on the ability of Xap to survive in infested soil and plant debris (Schuster & Coyne 1976; Saettler et al., 1986; Gilbertson et al., 1990). Gilbertson et al. (1990) found Xap populations to overwinter in bean debris on no-tillage plots. Non-pathogenic pectolytic strains of X. campestris were also consistently isolated. Experiments conducted in the Dominican Republic indicated that Xap survived up to 7 months on infected debris on the soil surface, but not in buried debris after 30 days (Arnaud-Santana et al. 1991). Xap survival studies conducted over ten years in Michigan indicated that infected crop debris is not the primary inoculum source for CBB (Saettler et al., 1986). Infected bean debris may be more important as an inoculum source in tropical and sub-tropical than in temperate areas (Gilbertson et al., 1990).
Survival of Xap is greater under dry conditions (Schuster & Coyne 1977a) as bacteria decline rapidly under moist conditions (Allen et al. 1998). Sabet & Ishag (1969) reported that Xap survived in press-dried bean leaves for more than 18 months in the laboratory, while Gilbertson et al. (1988) found Xap to remain viable in dry-leaf inoculum after 6 years. The longer survival under laboratory conditions as opposed to that in the field could be attributed the presence of antagonists, such as protozoa, in the soil (Habte & Alexander 1975).
Xap also survives on weeds and other host plants (Cafati & Saettler 1980b; Angeles-Ramos et al., 1991; Opio et al., 1995). Certain weed species may harbor the pathogen for up to 6 months (Opio et al. 1995). Angeles-Ramos et al. (1991) isolated epiphytic, pectolytic Xanthomonads from symptomless weeds where pathogenic strains were isolated from within infected fields. Epiphytic colonies survive on a wide range of plant species in the families Amaranthaceae, Commelinaceae, Compositae, Cruciferae, Gramineae, Oxalidaceae and Portulaceae in addition to various legumes (Allen et al., 1998). Epiphytic Xap populations are important in the epidemiology of CBB on dry beans (Ishimaru et al., 1991) and are differentially affected in hosts of different genotypes (Cafati & Saettler 1980a).
Seed infection by X. axonopodis pv. phaseoli can be external or internal. Both hot water and dry heat have been successful in treating bean seeds for X. axonopodis pv. phaseoli (Gondreau and Samson, 1994). This involves either incubating for 20 minutes in 52°C water or 23-32 hours in 60°C dry air at 45-55% RH. The latter treatment does not appear to affect seed viability. Treatment with an antibiotic such as streptomycin may be used to control external contamination with X. axonopodis pv. phaseoli, and streptomycin in polyethylene glycol may reduce, but not eliminate, internal populations of X. axonopodis pv. phaseoli (Liang et al., 1992).
The application of Rhizobium leguminosarum bv. phaseoli to common bean seeds reduced common bacterial blight severity and imnproved plant growth, both in field and greenhouse conditions (Osdaghi et al., 2011).
Under greenhouse conditions, bean seeds treated with a solution of tolylfluanid (1.20 g/L water) did not show any symptoms of infection until 30 days after sowing, and the overall prevelance of X. axonopodis pv. phaseoli was lower than in plants grown form untreated seeds (Lopes et al., 2008).
No methods are available to eradicate internal seed populations. However, external contamination may be controlled by streptomycin sulphate and sodium hypochlorite (Liang et al. 1992). Liang et al. (1992) investigated the potential of osmotic conditioning in reducing internal Xap populations from seeds, using polyethylene glycol (PEG) and glycerol as antibiotic carriers. They found that tetracycline and chlorotetracycline in PEG solutions effectively reduced Xap, but were phytotoxic. PEG solutions containing streptomycin reduced, but did not eradicate internal bacterial populations from naturally infected seeds with few phytotoxic effects. Tolylfluanid and Streptocycline were also reported as effective seed treatments (He, 2010).
Streptomycin is rapidly absorbed into bean stems and translocated to leaves but there is no indication that antibiotics are translocated downward through stems, trifoliate leaves or peduncle into the pod (Mitchell et al. 1954). Antibiotics should not be applied to leaves as resistant mutants may develop (Saettler 1989). Development of resistance to chemicals (Romeiro et al. 1998), costs involved and efficacy limit use of chemical control which may be feasible under certain circumstances, such as seed production or as a component of an integrated control strategy (Allen et al., 1998).
Seed Health Tests
Selective medium (Mohan and Schaad, 1987; Higley et al., 1993)
- Soak seeds for 20 h in a solution of 0.85% NaCl and 0.01% Tween 20 (SST) at 5°C.
- Aliquots of the soak solution are spread on a semi-selective medium (M-SSM) developed by (Mabagala and Saettler, (1992).
- Bacterial colonies that develop are isolated and tested for pathogenicity by inoculation onto greenhouse-grown bean plants (cv. Yelloweye).
As indicated by Maddox (1997) samples sizes can vary greatly (4000-45,000 seeds) in routine seed quality tests for this pathogen.
Dome test (Venette et al., 1987)
In this test, bean seeds are soaked in water. The soaked bean infusion is then vacuum-infiltrated into pregerminated seeds of the same lot. Symptoms are then observed after seedlings are grown out at high humidity.
Phage (Kahveci and Maden, 1994)
High degrees of specificity and reliability for detection of seedborne X. axonopodis pv. phaseoli was claimed for 2 of 16 bacteriophages obtained from X. axonopodis pv. phaseoli.
Serology (Wong, 1991)
Immunoflourescence and ELISA methods have been described.
DNA hybridization (Audy et al., 1996)
X. axonopodis pv. phaseoli can be detected in bean seeds after DNA extraction of intact or crushed seed followed by a PCR assay using pathovar-specific primers (Audy et al., 1996). Another PCR assay has been developed that is specific to X. axonopodis pv. phaseoli var. fuscans and thus will not detect X. axonopodis pv. phaseoli (Toth et al., 1998).
Nondestructive assay (Higley et al., 1993)
Various non-destructive methods detected X. axonopodis pv. phaseoli in bean seeds.
Real-time PCR (He, 2010)
Real-time PCR offers a sensitive, reliable and fast approach to diagnosis of Xap and Xapf in seeds (He, 2010). Assay specificity has been tested against DNA of several Xanthomonas species and pathovars of X. axonopodis. None of the closely related Xanthomonas strains were amplified using this PCR assay.
Plant TradeTop of page
|Plant parts liable to carry the pest in trade/transport||Pest stages||Borne internally||Borne externally||Visibility of pest or symptoms|
|Bark||Yes||Yes||Pest or symptoms usually visible to the naked eye|
|Fruits (inc. pods)||Yes||Yes||Pest or symptoms usually visible to the naked eye|
|Leaves||Yes||Pest or symptoms usually visible to the naked eye|
|Seedlings/Micropropagated plants||Yes||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||Yes||Pest or symptoms usually visible to the naked eye|
|Plant parts not known to carry the pest in trade/transport|
ImpactTop of page
Greater damage is more likely when early plant infection occurs. This is due to premature defoliation, which reduced the photosynthetic area available, interferes with translocation and reduces seed number and size. Lesions on seed and pods reduces quality. In the USA, crops with pods having 4% blemishes caused by blight are substandard and may not be harvested for seed, resulting in substantial economic losses. In 1983 in Uganda, there was a bacterial blight outbreak at the main seed multiplication site. This caused the operation to be abandoned and delayed the release of seed to farmers (Allen and Lenne, 1998).
This disease is most severe in tropical conditions, where it favours the high temperature and rainfall. In Colombia, a yield loss of 22% was reported for naturally infected fields and 45% for inoculated fields (Yoshi et al., 1976), while in north-western Argentina, it is one of the main diseases of Phaseolus vulgaris (Irigoyen and Garbagnoli, 1997).
In Mexico in 1988, X. axonopodis pv. phaseoli was found at a frequency of 37% in two locations. An inverse correlation was found between disease incidence and yield; disease incidence and severity were shown to increase after rain (Diaz-Plaza et al., 1991). In another trial in Mexico, the occurrence of bean common blight was associated with late flowering of P. vulgaris (Moran-Medina and Barrales-Dominguez, 1990). In field trials conducted during 1989-90 in Mexico, X. axonopodis pv. phaseoli was again found to be one of the most important diseases (Flores-Revilla et al., 1993). In the spring-summer and summer-autumn seasons of 1991, common blight had the highest incidence, severity and relative increase rate when compared with Macrophomina phaseolina and bean common mosaic virus in two experiments. It increased in incidence and severity with increases in temperature and rainfall (Mayek-Perez et al., 1995). In a separate set of trials in 1990 and 1991, disease severity was 53% and severity was 20%. Bacterial blight severity was negatively correlated with yield (Pedrosa et al., 1994). X. axonopodis pv. phaseoli severity has been shown to decrease under drought conditions in Mexico (Diaz-Plaza et al., 1991).
In Colombia, yield losses of 22 and 45% were estimated from natural and artificial infection, respectively. Studies in the 1980s showed yield losses of 20-47% (Allen and Lenne, 1998).
In the Caribbean, X. axonopodis pv. phaseoli is reported as frequently limiting bean yield (Beaver, 1999).
In Ethiopia, P. vulgaris is the most important legume crop and over 300,000 ha are grown annually by smallholders. Average yields vary between 500 to 1000 kg/ha, the reasons for the low productivity being abiotic and biotic factors; X. axonopodis pv. phaseoli is considered to be a major disease (Ohlander, 1980; Girna et al., 1994). In Kenya, X. axonopodis pv. phaseoli is again a constraint to bean production. Percentage crop losses of between 10 and 75% have been reported (Makini and Danial, 1994). Intercropping bean with maize was shown to reduce the severity of common bacterial blight during 1987-88 in Tanzania (Kiroka et al., 1989).
In Uganda, P. vulgaris is also the most important legume crop. Production is low, varying between 400 and 700 kg/ha, attributed to biotic and abiotic constraints. X. axonopodis pv. phaseoli is widespread and can cause considerable decreases in yields along with several other diseases (Opio et al., 1994). Recent losses are estimated at 40%. Work in Uganda has also shown that for each 1% increase in the incidence of common blight during reproductive growth there is a yield loss of 3.5-11.5 kg/ha, depending on the season (Allen and Lenne, 1998).
Major losses have also occurred in temperate climates. In southern Ontario, Canada, a yield loss of 38% was recorded in 1971-72. Yield losses of bean crops in 1968, 1970 and 1972 were 762,000, 1,256,000 and 217,000 kg, respectively (Wallen and Jackson, 1975). In white pea bean lines from a P. vulgaris/P. acutifolia cross, susceptible lines showed an average yield loss of 25% when disease free and inoculated plots in Ontario were compared. Resistant lines recorded little or no yield loss. The most severely infected lines tended to have the greatest loss in yield (Scott and Michaels, 1992). In Michigan, USA, a 1976 outbreak of common blight affected 75% of a 263 000 ha bean crop causing an estimated yield loss of 10-20% (Allen and Lenne, 1998).
Experimental studies have also reported on the losses due to this disease and the factors that can affect disease severity or incidence. Disease intensity and yield loss (green pods) varied from 4 to 71 and 1 to 84%, respectively, for two P. vulgaris cultivars infected with X. axonopodis pv. phaseoli at different growth stages. Early inoculation resulted in greater losses (Kishun et al., 1988).
Treatment of P. vulgaris plants with antibiotics at the beginning of anthesis was shown to reduce seed infection by X. axonopodis pv. phaseoli and increased yield by 14.4% (Tsvetkov and Donev, 1984). Increasing cumulative sulfur dioxide concentrations resulted in a significant decrease in the rate of lesion appearance. However, while the sulfur dioxide effectively inhibited disease development, it also led to a reduction in yield (Reynolds et al., 1987).
Results from laboratory, greenhouse and field experiments with seed from two cultivars of P. vulgaris inoculated with X. axonopodis pv. phaseoli showed that the inoculum could be localized on the seed surface or internally. A high degree of seed transmission of the pathogen was recorded. Seedling emergence was not affected up to a frequency of 10% infected seeds, but infection levels of 5% or more reduced crop yield (Valarini et al., 1996).
Inoculation of P. vulgaris with X. axonopodis pv. phaseoli by sandblast injury in the field resulted in 26-28% blight of leaves compared with 8-16% under natural infections. However, disease incidence and severity were low, and were not correlated with yield in this experiment (de Fario and de Melo, 1989).
The effect of hydrogen fluoride (HF) on field-grown beans spray-inoculated with an antibiotic-resistant strain of X. axonopodis pv. phaseoli was assessed. Final disease severity was not affected by exposure to HF, but the apparent infection rate increased with an increase in concentration of HF. In 1984, bean yield was not affected by HF, but in 1985 yield decreased with an increase in foliar tissue fluoride concentration (Reynolds and Laurence, 1988).
DiagnosisTop of page
Many different diagnostic methods have been developed for X. axonopodis pv. phaseoli and X. axonopodis pv. phaseoli var. fuscans (Xapf). Xap and Xapf can be easily isolated from CBB symptoms on leaves and pods using general isolation media (Schaad & Stall 1988). On media such as sucrose peptone agar (SPA), colonies are circular, smooth and mucoid with a yellow pigment referred to as xanthomonadin. Intensity of this yellow colour varies with medium used (Moffet & Croft 1983). Isolation media containing tyrosine differentiates between Xap and Xapf in that the latter produces a brown diffusible pigment (Basu & Wallen 1967). Goodwin & Sopher (1994) found this pigment to be produced due to secretion and subsequent oxidation of homogentisic acid rather than tyrosine activity.
Selective media are more effective for isolating specific bacteria from diseased material when selective at species level (Claflin, et al., 1987). Examples of semi-selective media for X. axonopodis pv. phaseoli are MXP (Claflin et al., 1987), milk-tween agar (Goszczynska and Serfontein, 1998) and M-SSM (Mabagala and Saettler, 1992). Remeeus and Sheppard (2006) recommended the use of semi-selective media MT and XCP1 for the detection of Xanthomonas axonopodis pv. phaseoli. They indicated that although MT was not as effective than XCP1 a larger number of suspected and confirmed colonies of Xanthomonas axonopodis pv. phaseoli were found on MT versus XCP1. A number of semi-selective media are, however, available to isolate Xap and Xapf (Kado & Heskett 1970, Schaad & White 1974, Trujillo & Saettler 1979, Claflin et al. 1987, Dhanvantari & Brown 1993, Jackson & Moser 1994).
Apart from using selective media, techniques such as bacteriophage typing (Katznelson et al. 1954, Sutton & Wallen 1967), serology testing (Trujillo & Saettler 1979), host inoculation (Saettler 1971), ELISA (Wong 1991) and immunofluorescent staining (Malin et al. 1983), can be used to detect and identify Xap and Xapf. These techniques are time consuming and labourious. More sensitive, rapid and specific detection of the pathogen is often needed. This is particularly important when identification is complicated by epiphytic Xap strains (Gilbertson et al., 1989, Audey et al., 1994) that may confuse seed certification (Wong 1991, Audey et al. 1994).
Starch is frequently included in the medium to help detect X. axonopodis pv. phaseoli because it hydrolyses starch, an uncommon trait among bacteria. Various diagnostic physiological tests are available for use with pure cultures but a pathogenicity test is required for detection at the pathovar level (Sheppard et al., 1989). Detection with phages specific to X. axonopodis pv. phaseoli have also been attempted, but not all strains are susceptible to a particular phage and phages are usually not truly species-specific (Sheppard et al., 1989).
Immunosassays are available for X. axonopodis pv. phaseoli but most have not been sufficiently specific for accurate diagnosis. However, monoclonal antibodies are now available for X. axonopodis pv. phaseoli and have been used in immunofluorescence microscopy and an ELISA to detect it in seed (Wong, 1991). This monoclonal antibody appears to be highly specific to X. axonopodis pv. phaseoli and only cross-reacts with X. axonopodis pv. malvacearum (Wong, 1990).
Specific diagnosis of X. axonopodis pv. phaseoli was achieved with a DNA hybridization probe (Gilbertson et al., 1989). Based on this probe, another highly specific PCR probe, for Xap detection, was developed to detect as few as 10 colony forming units (CFU), using ethidium bromide-stained agarose gel (Audey et al. 1994). Audey et al. (1996) developed a rapid, sensitive PCR assay for detection of seedborne Xap in large bean seed samples containing as few as one infected in 10 000 healthy seeds. Birch et al. (1997) used RAPD-PCR to differentiate between Xap and Xapf. Toth et al. (1998) used primers which amplified a DNA fragment from all Xapf-isolates used, while no amplification products were obtained from Xap-isolates. These primers, therefore, provide a rapid, improved method to differentiate between these two variants. More recent techniques used to differentiate Xap and Xapf include restriction fragment length polymorphism (RFLP) analyses (Chan et al., 1990; Gilbertson et al., 1991, Lazo et al., 1987), DNA-DNA hybridization (Hildebrand et al., 1990), pulse field gel electrophoresis (Chan et al., 1990) and rep-PCR (López et al., 2006; Mahuku, 2006; Mkandawire et al., 2004).
The pathogen can be readily isolated from symptomatic tissue (Dreo et al., 2003). A comparison of various extraction processes is given in He and Munkvold, 2012).
The method of identification issued by the International Seed Federation in 2006 is successfully used in Balaz et al. (2008). X. axonopodis pv. phaseoli was isolated from bean seeds and isolated on a semiselective media. ELISA and PCR can then be used to confirm the identity of resulting isolates. Milk Tween Agar (MTA) and Modifies Sucrose Peptone Agar (MSPA) are both suitable for isolation of P. savastanoi pv. phaseolicola.
The development and use of a PCR-based method for detection of X. axonopodis pv. phaseoli in pomegranate is described in Mondal et al. (2012).
Seed Health Tests
A sample, usually 1000 to 10,000 seeds, is washed in running water and then placed in a container to which a solution of phosphate-buffered saline or liquid semi-selective media is added. The samples are then incubated at room temperature for 2-3 hours. Longer incubation times can increase the number of saprophytic bacteria and reduce the number of aerobic xanthomonads. The solution, known as the seed-soak wash, is diluted and plated onto various culture media. The bacteria are then identified by physiological traits, phage tests, pathogenicity tests and/or immunoassays (Sheppard et al., 1989). The seed-soak wash can also be directly injected into susceptible plants or vacuum-infiltrated into pregerminated bean seeds, which are then observed for typical common blight symptoms (Venette et al., 1987; Sheppard et al., 1989).
A comparison of extraction methods showed that crushing seed in phosphate-buffered saline was more effective in releasing bacteria than soaking intact seeds (Maringoni et al., 1998). X. axonopodis pv. phaseoli can also be detected in seed after DNA extraction of intact or crushed seed followed by a PCR assay using specific primers (Audy et al., 1996). Results from studies conducted by He (2010) to assess the influence of different extraction steps on the sensitivity of Xap indicated that vacuum extraction and centrifugation of seed extracts increased sensitivity, and the highest sensitivity was obtained with the 3-hour vacuum extraction at room temperature followed by centrifugation.
For infected plant tissue other than seed, the material should be washed thoroughly in running water and dried. Surface sterilization may be necessary as a general hygiene precaution to avoid surface contaminants. The margin of a lesion is excised with a sterile blade and mixed with a small amount of sterile water or phosphate-buffered saline to allow the bacteria to diffuse out into the solution for 15-30 minutes. A sterile loop is then used to streak the solution onto agar media (Schaad and Stall, 1988).
Detection and InspectionTop of page
Similarities to Other Species/ConditionsTop of page
The symptoms of common blight can also be confused with those of chocolate spot of bean caused by Pseudomonas syringae pv. syringae.
Prevention and ControlTop of page
Due to the variable regulations around (de)registration of pesticides, your national list of registered pesticides or relevant authority should be consulted to determine which products are legally allowed for use in your country when considering chemical control. Pesticides should always be used in a lawful manner, consistent with the product's label.Use of Disease-Free Seed
Approximately 1 in 10,000 seeds is capable of causing an outbreak of blight (Sutton and Wallen, 1970). A survey of navy bean (Phaseolus vulgaris) seeds showed that approximately 35% had internal contamination with X. axonopodis pv. phaseoli (Saettler and Perry, 1972). The level of infected seed will vary depending on the level of common blight occurring during seed production.
Use of disease-free seed with a strict seed certification programme is important to reduce the amount of initial inoculum. Beans for seed production should be grown in an isolated area separated from production fields and should be inspected for common blight. Growing beans in irrigated desert areas greatly reduces the chances of spread by splashing rain, and in the USA and Canada, bean seed is grown in areas such as Idaho where bacterial blights are minimal and field inspections are strict (Sheppard, 1983). Seed should be tested for internal infection as the presence of symptoms is not a reliable indicator of infection (Sheppard et al., 1989). Seed may also be treated with bactericides, such as streptomycin or sodium hypochlorite, to eliminate surface contamination by X. axonopodis pv. phaseoli (Liang et al., 1992). Treating seed may reduce disease levels in a subsequent crop but should not form the basis of quarantine measures.
For further information on seed health tests, see Diagnostic Methods.
Seed infection by X. axonopodis pv. phaseoli can be external or internal. Thermotherapy is widely applied for the control of seedborne bacteria (Gondreau and Samson, 1994). Both hot water and dry heat have been successful in treating bean seeds for X. axonopodis pv. phaseoli (Gondreau and Samson, 1994). This involves either incubating for 20 minutes in 52°C water or 23-32 hours in 60°C dry air at 45-55% RH. The latter treatment does not appear to affect seed viability. Treatment with an antibiotic such as streptomycin may be used to control external contamination with X. axonopodis pv. phaseoli, and streptomycin in polyethylene glycol may reduce, but not eliminate, internal populations of X. axonopodis pv. phaseoli (Liang et al., 1992).
Cultural practices are important in controlling common blight. Eliminating weeds, volunteer beans and other potential hosts of X. axonopodis pv. phaseoli will reduce disease incidence. Good weed control will not only remove potential sources of epiphytic X. axonopodis pv. phaseoli populations, but will also improve aeration around the crop so that the plants dry faster, thus reducing the chances for bacterial spread and infection. X. axonopodis pv. phaseoli is readily spread by water, and walking or working in the field while plants are wet will splash the bacteria and create wounds. Plants should be allowed to dry before allowing workers or machinery to enter. Eliminating infected plant debris is very important, particularly in tropical regions (Saettler, 1991). A rotation of at least 2 years between bean crops will give time for the X. axonopodis pv. phaseoli population to decline in the debris. Deep ploughing will also encourage the breakdown of infected plant debris and reduce the population of X. axonopodis pv. phaseoli (Gilbertson et al., 1990). Another option is to burn the crop debris to eliminate infected plant material. The incidence of X. axonopodis pv. phaseoli can also be reduced if beans are grown with maize rather than in a monoculture (Van Rheenen et al., 1981). The maize appears to provide a physical barrier to the movement of X. axonopodis pv. phaseoli between bean plants.
There are no reports of high resistance to X. axonopodis pv. phaseoli in P. vulgaris. However, many lines of P. vulgaris show some resistance to X. axonopodis pv. phaseoli and these varieties may be planted if available. Increased resistance can be developed by selecting for horizontal rather than vertical resistance (Garcia-Espinosa, 1997). Partial resistance to X. axonopodis pv. phaseoli in P. vulgaris has been linked to delayed flowering under long photoperiods (Coyne et al., 1973). P. acutifolius is highly resistant to X. axonopodis pv. phaseoli and partial resistance has been transferred from this genotype to P. vulgaris (Goodwin et al., 1995). There are also several other reports of resistance transferred from P. acutifolius to P. vulgaris (Thomas and Waines, 1984; Park et al., 1998; Yu et al., 1998). Resistance in P. acutifolius is controlled by one or two dominant genes and is related to the hypersensitive response (Zapata, 1998; Urrea et al., 1999). In addition, crosses between P. coccineus and P. vulgaris also showed resistance to X. axonopodis pv. phaseoli (Zapata et al., 1985; Yu et al., 1998). A number of P. vulgaris lines with varying levels of resistance to X. axonopodis pv. phaseoli have been registered (for example, Beaver et al., 1999; Miklas et al., 1999).
A number of molecular markers have been developed for resistance genes to X. axonopodis pv. phaseoli. A growing number of DNA markers, particularly RAPDs, have been found to be linked to common blight resistance genes and are being used in marker assisted selection (Jung et al., 1996; Miklas et al., 1996; Park et al., 1998; Tar'-an et al., 1998; Yu et al., 1998; Ariyarathne et al., 1999).
Several factors should be taken into consideration in evaluating resistance to X. axonopodis pv. phaseoli. Not all plant parts react similarly and the level of resistance to X. axonopodis pv. phaseoli has been found to be different in foliage and pods, each of which are determined by different genes (Schuster et al., 1983). Variation in the virulence of X. axonopodis pv. phaseoli has been observed frequently and isolates from tropical regions appear to be more virulent than those from temperate areas (Schuster et al., 1973). Environmental conditions can also affect resistance. Some cultivars which are moderately resistant in temperate regions become susceptible in the tropics, probably because of their poor adaptation to tropical growing conditions (Webster et al., 1983). Common blight is more severe under the higher temperatures and shorter photoperiods, which are found in subtropical and tropical areas (Arnaud Santana et al., 1993).
Chemical control may reduce leaf infection but usually has little improvement on yield. Copper compounds may be used (Weller and Saettler, 1976). Foliar antibiotic treatment can provide some control but is undesirable because it can result in antibiotic-resistant mutants of X. axonopodis pv. phaseoli.
ReferencesTop of page
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