Bacterial leaf streak (BLS), caused by Xan-thomonas translucens (ex Jones, Johnson and Reddy 1917) Vauterin, Hoste, Kersters and Swings 1995, is the major bacterial disease of wheat. It occurs over a range of very different conditions, such as sprinkler-irrigated fields in temperate climates, high-rainfall subtropical highlands and warmer environments characterized by cool nights or frequent climatic changes and sudden temperature variations. The disease, called black chaff when on the glumes, is seed-borne and a constraint for international germplasm exchange. Bacterial leaf streak was first identified on barley (Hordeum vulgare L.) (Jones et al., 1917) and later on wheat (Triticum aestivum L.) (Smith et al., 1919), rye (Secale cereale L.) (Reddy et al., 1924), grasses (Wallin, 1946), and finally on triticale (X Triticosecale Wittmack) (Zillinsky and Borlaug, 1971). Different names have been proposed, depending on the host plant, for the closely related cereal streak pathogens also often grouped together under the name 'translucens group'. The taxonomy of this group of bacteria and the phytopathological relevance of the classification has been recently revisited in detail and clarified (Vauterin et al., 1995; Bragard et al., 1997). The name X. translucens pv. undulosa is used to refer to the pathogen that causes BLS on wheat.
DISTRIBUTION
Bacterial leaf streak has a wide geographical distribution (Duveiller et al., 1997). In North America, it has been reported in the United States, Canada and Mexico. In South America, it occurs in Argentina, Bolivia, parts of Brazil, Paraguay, Peru and Uruguay (Mehta, 1990; Mohan and Mehta, 1985; Duveiller et al., 1991; Frommel, 1986; Tessi, 1949). The disease is known on wheat in China (Sun and He, 1986), Pakistan (Akhtar and Aslam, 1986) and Iran (Alizadeh et al., 1995), and on triticale in India (Richardson and Waller, 1974). In the Near and Middle East, it affects durum (T. turgidum var. durum L.) and bread wheat in irrigated areas of Syria (Mamluk et al., 1990), Israel (CIMMYT, 1977), Turkey (Demir and Üstün, 1992) and Yemen (Bragard et al., 1995). The disease currently seems to be absent from Western Europe (Paul and Smith, 1989), probably due to unfavourable environmental conditions, particularly low temperatures. In Africa, BLS has been found in Kenya (Burton, 1931), Ethiopia (Korobko et al., 1985), South Africa (Smit and Van A. Bredenkamp, 1988), Tanzania (Bradbury, 1986), Libya and Madagascar (Bragard et al., 1995), Morocco (Sands and Fourest, 1989) and Zambia (Bragard et al., 1997). In Australia, BLS has been recorded on wheat and rye in New South Wales (Noble, 1935).
IMPORTANCE
Little quantitative information is available on losses caused by BLS. Yield losses as high as 40 percent have occurred in the most severely diseased fields in Idaho, United States, although losses are generally 10 percent or less (Forster et al., 1986). In severe cases, 5 to 10 percent of the wheat spikes may be sterile due to infection (Forster and Schaad, 1988), and the disease may attack a complete nursery so severely that nothing can be harvested (Burton, 1931).
Data from Mexico indicated that, on average, losses below 5 percent could be expected when the percent infected flag leaf area is less than 10 percent. Yield loss is a linear function of the percent infected flag leaf area, and even a small-infected leaf area has an effect on yield. The disease mainly affects grainfilling, but grain number was significantly correlated with BLS severity levels two out of three years under Mexican conditions (Duveiller and Maraite, 1993).
SYMPTOMS
Typical symptoms on the leaf consist of elongated, light brown lesions, several centimetres long, which are initially distinct but later coalesce to cover larger solid areas. Early symptoms are characterized by translucent stripes that are easily seen under incident light. Initially, lesions are water-soaked and produce honey-like exudates giving a milky slime under humid conditions (Smith, 1917). If undisturbed, the exudates harden into yellowish, resinous granules or scales studding the surface of the lesions and are easily detachable (Plate 48, Plate 49).
When on the glumes, BLS is characterized by black, longitudinal, more or less parallel stripes that are more numerous and conspicuous on the upper parts (Smith, 1917). Bacterial leaf streak can be recognized by a greasy appearance or alternating bands of diseased and healthy areas on the awns (Plate 50). Purple-black symptoms may extend to the peduncle between the inflorescence and the flag leaf, and may sometimes present a yellow centre (Forster et al., 1986).
Several authors have found that susceptibility to melanism on the spike, also referred to as black chaff, is often inherited from stem rust-resistant parents. Johnson and Hagborg (1944) showed that high temperature conditions, especially when combined with high humidity, favoured the development of melanic areas on the glumes, lemmas, peduncles and internodes of rust-resistant cultivars. As brown melanosis is known to be associated with the Sr2 gene for stem rust resistance, it is possible that what early reports of black chaff were really describing was pseudo-black chaff not caused by bacteria.
Sharp discoloured interveinal streaks on the glumes suggest the presence of Xanthomonas, particularly if also irregularly distributed on the spike and if there is abundant BLS on the leaves. In contrast, melanosis on the peduncle, which occurs on the same side of most culms in a field as a result of exposure to sunshine, is indicative of brown melanosis.
EPIDEMIOLOGY AND BIOLOGY
Survival
Seed is the most important source of primary inoculum, and large-scale transmission of BLS is due to its seed-borne nature (Smith et al., 1919). Depending on storage conditions, it is estimated that the bacterium will die in 63 to 81 months (Forster and Schaad, 1990). Black chaff of wheat has a very low transmission rate, i.e. low levels of seed contamination will not result in field disease (Schaad, 1988a). In Idaho, more than 60 percent of all spring wheat seed lots were found to be contaminated, and seed lots with less than 1 000 colony-forming units (cfu) per gram do not cause field epidemics. This suggests that methods for detecting the pathogen on the seed do not have to be very sensitive (Schaad, 1987a; Forster and Schaad, 1987). However, the situation may vary from one environment to another, and the pathogen's multiplication capacity should not be underestimated.
The bacterium survives poorly in soil, but does better when crop debris is present (Boosalis, 1952). Also, plant stubble usually decays very fast in warm, humid climates, and wheat pathogenic bacteria cannot survive in decomposing debris.
Xanthomonas translucens pv. undulosa can survive on weeds and grasses due to its broad host range; however, this is probably not significant on annual hosts. Epiphytic populations of the pathogen have been detected in Idaho on grasses near spring wheat fields (Thompson et al., 1989). Wallin (1946) gathered evidence that X. translucens can overwinter on perennial hosts, such as smooth brome (Bromus inermis Leyss.) and timothy (Phleum pratense L.), which gives the pathogen the opportunity to spread to nearby cereals. The bacterium also seems to overwinter on winter wheat and rye (Boosalis, 1952).
FIGURE 20.1
Disease cycle of Xanthomonas translucens pv. undulosa and possible
ways disease may spread
Conditions conducive to epidemics
Bacterial leaf streak outbreaks are characterized by sporadic epidemics and are usually observed by farmers relatively late in the growing season (Forster et al., 1986). Moisture facilitates the pathogen's release from the seed and contributes to leaf colonization and invasion of leaf tissue. Free water allows the pathogen to spread in the field and to disperse on the leaf, thus increasing the number of lesions. Bacteria enter through the stomata and multiply in large masses in the parenchyma. This causes elongated streaks limited by the veins, which act as barriers. Later milky or yellow exudates form on the surface of lesions. Rain and wind greatly influence the spread of these exudates and of the disease from leaf to leaf throughout the field (Figure 20.1). Micro-injuries to awns and leaves caused by hail or wind may contribute to bacterial penetration.
The BLS-inducing pathogen tolerates a wide range of temperatures (15° to 30°C) (Duveiller et al., 1991). Recent studies (Duveiller and Maraite, 1995) have shown that temperature has a major impact on epidemics. Pathogen multiplication in leaf tissue is directly dependent on temperature, and dry air conditions (relative humidity less than 30 percent) do not limit disease progress. Symptoms only occur when temperature allows the bacterial population to reach an estimated threshold of 108 cfu/leaf. Low temperatures retard the multiplication of the pathogen and disease progress.
Epiphytic populations may be important for understanding the aetiology of BLS and discovering why the disease is sporadic. In Mexico, it was possible to monitor a X. t. pv. undulosa population in plots of symptomless genotypes contrasting in their field resistance to the pathogen (Duveiller, 1994a). The population of pathogenic bacteria decreased after a heavy rainfall, which suggests that epiphytic X. t. pv. undulosa are present on wheat leaves before they actually penetrate the parenchyma.
Strains of X. translucens express ice nucleation activity at temperatures from -2°C to -8°C (Kim et al., 1987). Damage caused to plant tissue by the ice provides conditions suitable for pathogen invasion and multiplication. Frost conditions may thus explain the frequent incidence of BLS in high elevation environments or in regions, such as southern Brazil, where wheat is grown during the winter season. Waller (1976) reported that a slight frost precipitated a BLS outbreak in the Toluca Valley of Mexico (2 600 masl) in 1973. However, frost conditions are not common in this area during the summer season when plants present BLS symptoms, and ice nucleation is thus not necessary to induce an epidemic (Duveiller et al., 1991).
Spread of the pathogen in the field
Pathogen transmission by rain and dew and plant-to-plant contact explains local dissemination (Boosalis, 1952). In addition, visitors to demonstration plots, particularly in the morning when dew is at its maximum, increase the spread of bacteria. The disease may spread in the direction of the prevailing wind and driven rain; however, the movement of the disease in space under other conditions proved to be limited. In Brazil, Mehta (1990) indicated that the spread of BLS from one field to another is limited, and disease spread through splashing rain is restricted to distances as short as 4 to 5 m. The role of aphids in long distance transmission of the disease is probably limited.
THE PATHOGEN
Isolation and pathogenicity test
Xanthomonas translucens pv. undulosa grows fastest in vitro at 28° to 30°C. The bacterium can be cultivated on common media, such as nutrient agar and Wilbrink's medium (Sands et al., 1986). These culture media are not semi-selective and can be used for a wide range of bacteria. Semi-selective media include KM-1, XTS and WBC (Duveiller et al., 1997). When no selective medium is available, Wilbrink's medium is preferred, given that the pathogen's typical yellow mucoid colony is best distinguished from saprophytes on this non-selective medium.
Forcing a 105 cfu/ml bacterial suspension into the leaf whorl of young plants (four to five leaves) or into the boot of older plants with a hypodermic syringe is a very effective inoculation method (Bamberg, 1936) for testing pathogenicity. This was confirmed at the In-ternational Maize and Wheat Improvement Center (CIMMYT), where plants are usually incubated for five days in a humid chamber after inoculation (Duveiller, 1994b). In some cases, water soaking is observed as early as three to four days.
Diversity of X. translucens strains that attack wheat and other small grains
Host range and other host-pathogen relationships
Since BLS was first identified on wheat, several pathovar names have been used for X. translucens strains isolated from small grains; however, these strains have not always been subjected to differential host range pathogenicity tests. This has caused confusion: in the first place, strains having different names (based on the host plant from which they were isolated) may be similar; second, many authors use the name X. t. pv. translucens in a general sense for any cereal streak pathogen (Bradbury, 1986), although the names of four BLS-inducing pathovars of X. translucens are currently included in the most recent International Society for Plant Pathology (ISPP) list of plant pathogenic bacteria (Young et al., 1996). These names are:
· Xanthomonas translucens pv. translucens (Jones, Johnson and Reddy 1917) Vauterin, Hoste, Kersters and Swings 1995;
· Xanthomonas translucens pv. cerealis (Hagborg 1942) Vauterin, Hoste, Kersters and Swings 1995;
· Xanthomonas translucens pv. secalis (Reddy, Godkin and Johnson 1917) Vauterin, Hoste, Kersters and Swings 1995;
· Xanthomonas translucens pv. undulosa (Smith, Jones and Reddy 1919) Vauterin, Hoste, Kersters and Swings 1995.
If the ISPP rules are followed correctly, the pathovar name X. t. pv. translucens should be reserved for strains pathogenic on barley only (Jones et al., 1917). Xanthomonas translucens pv. undulosa designates strains pathogenic on wheat and triticale and can be isolated from several hosts including wheat, barley, triticale and rye. Hence, its host range is not only broader than that of X. t. pv. translucens, but also covers the host range of X. t. pv. cerealis as defined by inoculation tests conducted on oat, rye and Bromus (Boosalis, 1952; Bragard et al., 1995). Based on amplified fragment length polymorphism (AFLP) and fatty acid methyl ester (FAME) analysis, Bragard et al. (1997) showed that strains pathogenic on barley, but not on wheat, clustered in a genetically different group. Results indicated that the pathovars cerealis, translucens and undulosa correspond to true biological entities. Xan-thomonas translucens pv. secalis, which has been described as pathogenic on rye (Reddy et al., 1924), appears closely related to X. t. pv. undulosa (Bragard et al., 1997).
Clear differences in aggressiveness were noted among X. t. pv. undulosa strains from various geographical origins. Whereas typical strains induce extensive stripe symptoms on wheat and barley, strains from some areas induce limited symptoms (Bragard and Maraite, 1994). No evidence of strong race specialization has as yet been found on wheat, as indicated by highly non-significant cultivar x strain interaction (Milus and Chalkley, 1994).
Biochemical and physiological traits
Simple tests can be used to identify X. trans-lucens from other bacterial species (Schaad, 1988b). The BLS pathogen is non-sporing, rod-shaped, Gram-negative and motile by a single polar flagellum. It is further characterized by rods 0.4 to 0.8 µm x 1.0 to 2.5 µm, singly or in pairs, except in peptonized nutrient broth with 2 percent sodium chloride (NaCl) in which long non-motile chains are formed (Jones et al., 1917; Dowson, 1939). Xanthomonas translucens is oxidative, and no nitrate to nitrite reduction is observed. The reaction for Kovacs' oxidase and arginine dihydrolase is also negative. There is no 2-ketogluconate production, and esculin hydrolysis is positive. Hypersensitivity on tobacco is positive (Mohan and Mehta, 1985), but this reaction is not always clear, and a cool incubation temperature is suitable. Unlike X. campestris, X. translucens strains do not hydrolyse starch and do not use lactose (Schaad, 1987b). These tests, as well as immunological methods, do not help to differentiate among the various pathovars of X. translucens from cereals.
CONTROL STRATEGIES
Rotations
Since the major source of inoculum is infected seed, rotations may not play a key role in controlling the disease. Straw can harbour viable inoculum from season to season and cause initial infection in the field, but the number of viable bacteria in infested, overwintered straw is reduced when the straw is incorporated into the soil (Boosalis, 1952). Moreover, it should be stressed that survival of the pathogen on plant debris seems improbable due to its extreme susceptibility to antagonistic bacteria.
Seed health
Dilution plating
The best way to limit BLS is to avoid sowing infected seed. A seed wash test on a semi-selective medium after dilution plating is the normal non-destructive procedure used in pathogen-free seed certification. The method has the advantage of detecting living pathogens. The number of colony-forming units per gram of seed gives an estimate of the number of bacterial cells present in the sample. The number of single colonies growing on the agar medium has to be counted, and representative colonies have to be cloned and proven pathogenic on wheat.
Several semi-selective media have been developed:
Schaad and Forster (1985) developed the XTS medium: glucose, 5.0 g; nutrient agar (Difco), 23.0 g; cycloheximide, 200 mg; cephalexin, 10.0 mg; gentamycin, 8.0 mg; and distilled water, 1 litre. To perform the test, take 120 ml of sterile, cold 0.85 percent NaCl (saline) containing 0.02 percent v/v Tween 20 and add to 120 g seed (about 3 000 seeds). After shaking vigorously for three to five minutes, let settle for one minute and prepare tenfold dilutions to 10-3 using cold sterile saline. Transfer 0.1 ml of each dilution onto each of three plates of XTS agar and spread with an L-shaped rod. Examine the plates after five days of incubation at 30°C. Colonies of X. translucens are 1 to 2 mm in diameter, yellow, clear, round, convex and smooth (Schaad and Forster, 1993). Streak a known culture onto XTS for comparison. Positive colonies are tested for pathogenicity by injecting a bacterial suspension (approximately 105 cfu/ml) into leaf whorls of susceptible wheat seedlings. Disease symptoms appear after incubating for five to seven days in a dew chamber at 26°C. Zero tolerance is not necessary where BLS is endemic (Schaad, 1988a), but infested seed should not be used for germplasm exchange.
Another seed test medium that proved to be effective under Mexican and other conditions is WBC medium, a modification of Wilbrink's medium (Duveiller, 1990a). WBC medium is Wilbrink's medium amended with boric acid (0.75 g/L) and cephalexin (10 mg/L). It does not contain gentamycin but includes 75 mg/L cycloheximide to reduce fungal growth.
Recently, Maes et al. (1996) developed a method for recognizing BLS-causing Xan-thomonas pathogens using ribosomal DNA spacer sequences and polymerase chain reaction (PCR) (Maes and Garbeva, 1994). The tests proved to be quick (results can be obtained in five hours, compared to several days using the dilution and plating method) and relatively sensitive (2 x 103 cfu/g of seed), indicating the technique might be useful for detecting those pathogens in seed without isolation. However, this method also detects five other Xanthomonas with a host range restricted to forage and some ornamental grasses. No data are available on the survival of these grass pathogens on non-host plants, especially in seeds of cereals such as wheat.
Serodiagnostic assays
Serological methods have been developed for identifying strains grown as pure cultures and for detecting the black chaff-inducing pathogen in seed. Using rabbit polyclonal antibodies for detecting X. translucens in wheat seed, Claflin and Ramundo (1987) were only able to obtain positive readings with a dot-immunobinding assay (DIA) when cell concentration was 105 cfu/ml or higher.
Bragard and Verhoyen (1993) developed monoclonal antibodies reacting positively with X. t. pv. undulosa, X. t. pv. cerealis, X. t. pv. translucens and X. t. pv. secalis, which proved more specific than polyclonal rabbit antiserum. Monoclonal antibody AB3-B6 was used in both immunofluorescence (IF) and DIA to detect pathogens in aqueous seed extracts. These techniques were compared to dilution plating of a seed wash. Seed lots contaminated with a high (greater than 104 cfu/g) population of bacteria were consistently identified with all three methods. Immunofluorescence was more sensitive than DIA and gave more reproducible results. The DIA method is simple and requires inexpensive equipment, but the detection threshold is high (105 cfu/ml), making it more appropriate for the identification of pure strains. On the other hand, although IF requires more expensive equipment, it is relatively quick and sensitive. The detection threshold is 103 to 104 cfu/ml. Immunofluorescence-positive seed lots should not be used for sowing in areas that favour disease development since a pathogen concentration of 1 x 103 cfu/g of seed is likely to induce an epidemic (Duveiller and Bragard, 1992).
Techniques using seedlings
Determining the percent infected seedlings after growing naturally contaminated seed on sterile soil in a moist chamber (100 percent relative humidity; 22° ± 3°C) did not prove workable under Mexican conditions. No symptoms were found on more than 13 000 seedlings grown from heavily infected seed (greater than 106 cfu/g) after four weeks of incubation in trays where each seed was put in a 1 cm deep hole in sterile soil (Duveiller, 1994b).
The modified injection technique proposed by Mehta to detect the presence of X. t. pv. undulosa on wheat, triticale and rye seed and X. t. pv. translucens on barley seed may be used for quarantine purposes (Mehta, 1990). Shake the seed (20 g) thoroughly for 90 minutes in 20 ml of sterile saline; then remove seeds and inoculate the suspension into 20-day old seedlings with a hypodermic syringe. Xanthomonas streak symptoms are assessed 7 to 12 days after inoculation. This method is recommended when immunofluorescence microscopy is not available (Bragard et al., 1993).
Seed treatments
Since no pesticide effectively controls the disease in the field, research on chemical control focuses on seed disinfection. However, the disease cannot be controlled by seed treatments alone, although several studies report partial effectiveness of various compounds (Sands et al., 1986).
Results of chemical seed treatments are contradictory. It was believed that the high BLS incidence recorded during the 1980s was due to the ban on mercurial compounds, but as shown by Forster and Schaad (1988), these products proved to be ineffective for controlling the disease. Cupric hydroxide (Kocide SD), non-volatile mercury (Mist-O-Matic) and volatile organic mercury compound (Panogen 15) are also unsatisfactory (Jons et al., 1982; Forster, 1982). Mehta (1986) reduced transmission of X. t. pv. undulosa by 80 percent with 300 ml/100 kg seed of Guazatine Plus (syn. Panoctine Plus) in an experiment with naturally infected seed of wheat genotype IAPAR-Caete. The treatment is effective if applied at least five months before sowing, but usually ineffective if applied a month before sowing. Also, a few heavily contaminated seeds may escape the product during the procedure and remain contaminated (Mehta and Bassoi, 1993). Seed treatment with acidified cupric acetate (0.5 percent) at 45°C for 20 minutes significantly reduced the amount of black chaff in the field. Stand count can be reduced significantly compared to other treatments when seed treated with acidified cupric acetate is planted; however, this level of phytotoxicity is considered acceptable for a foundation seed health programme (Forster and Schaad, 1988).
Fourest et al. (1990) recommend treating the seed at 72°C for seven days. This method allows treating larger amounts of seed, but experiments conducted at CIMMYT indicate that the method is not completely effective, particularly on seed samples larger than 100 g (Duveiller et al., 1997).
Bactericide seed treatments were also evaluated at CIMMYT using naturally infected wheat genotype Alondra, which is very susceptible to BLS. Results indicated that hot cupric acetate, Panoctine Plus (a.i. guazatine 300 g/L and imazalil 20 g/L), formaline and dry heat consistently reduced the amount of bacteria in the seed as determined using the dilution plate method with WBC agar. However, although the effects of bactericide treatments were significant (P=0.05), control of BLS in the field was not possible (Duveiller et al., 1991). When seed with high levels of X. t. pv. undulosa is used, bacteria surviving the treatments can multiply to reach the level found in leaves when symptoms develop (greater than 108 cfu/leaf). The incomplete effect of Panoctine Plus and dry heat was confirmed using heavily contaminated seed (100 g samples), but Panoctine Plus was used only a few days before planting. Nevertheless, even if not completely satisfactory, seed treatment with dry heat or a product such as Panoctine Plus is recommended.
The amount of bacteria in the seed, as well as its heterogeneous distribution, may partly explain contradictory results, particularly if the samples used are small. If a seed lot contains 105 cfu/g, a 99 percent effective bactericide will still allow 1 000 cfu/g to survive, which is the threshold for an epidemic (Forster and Schaad, 1985).
Seed multiplication in a disease-free area
Clean seed should be used to produce foundation seed, and multiplication should be conducted in a disease-free area under dry conditions without overhead irrigation. Although a seed multiplication field may not show black chaff symptoms, the pathogen may increase on the leaf and head surfaces, resulting in contaminated seed. In contrast, a high percentage of disease in the field does not necessarily result in a higher amount of bacteria in harvested seed lots (Mehta, 1990).
BREEDING FOR RESISTANCE
Since controlling black chaff through seed treatment is not easy, breeding resistant genotypes appears to be the best way to reduce the risk of yield losses. Screening for resistance is essential for breeding. The material to be screened must be uniformly exposed to the pathogen, and this is only possible through artificial inoculation. Epidemics are sporadic, and natural homogeneous infection in the field is too unreliable to allow adequate evaluation. Lines identified as susceptible under natural conditions may be infected as a result of higher seed infection levels. Also, disease-free genotypes may not really be resistant but may have simply escaped infection.
Greenhouse tests can be conducted on seedlings and young plants by infiltrating low cell concentrations into the leaf. It is important to use low bacterial concentrations to be able to detect measurable differences in resistance. A concentration of 104 cfu/ml of a young culture (24 hour) on agar medium is usually appropriate. The concentration can be adjusted with the help of a Petroff-Hausser counting chamber (Duveiller et al., 1997).
In another inoculation technique, the seedling pseudostem is filled with sterile water, and then a needle dipped in a young bacterial culture is passed through it. After five to seven days of incubation in a humid chamber, ideally at 24° to 26°C, disease is scored on the emerging leaf.
The major problem with screening at the seedling stage in the greenhouse is the fairly high degree of data variation. To minimize this variation, inoculum concentration, infiltration into confined portions of the leaf blade and moisture distribution in the dew chamber have to be carefully standardized. Also, the correlation between disease scores on seedling and field data is not always clear (Duveiller et al., 1997). Therefore, evaluating resistance based on adult plant response in the field is recommended.
Field inoculation can be done by spraying a concentrated (109 cfu/ml) bacterial suspension on plants at the tillering stage. This should be done in the afternoon to take advantage of night-time dew formation, which increases the chances of successful infection through leaf stomata. Approximately 200 Petri dishes containing a pure culture of a single X. t. pv. undulosa strain cultured on Wilbrink's medium are needed to inoculate half a hectare. After two days of incubation at 30°C, wash the agar and suspend the bacteria in water to produce highly concentrated inoculum. The inoculum can be prepared in the laboratory or in the field. The dilution factor necessary to prepare a 109 cfu/ml inoculum suspension for spraying needs to be established. Inoculum calibration can be done in the laboratory by doing a cell count with the help of a Petroff-Hausser counting chamber, or by estimating the number of Petri dishes covered with a 48-hour culture that are necessary to prepare the final inoculum. After adjusting the concentration, add Tween 20 (0.02 percent) to the inoculum to facilitate the spread of the liquid over the leaf. The inoculum suspension (approximately 20 ml/m2) is applied using a backpack sprayer at 1.4 kg/cm2 pressure. Inoculation can be carried out at Zadoks' DC 30 to 35 stages (Zadoks et al., 1974; Duveiller, 1990b) and may be repeated if necessary.
The disease progresses up a vertical gradient as shown by a smaller damaged leaf area on the flag leaf than on the flag leaf minus one. The disease progresses upward, and disease severity is assessed at flowering (Zadoks' DC 64). The scale proposed by Saari and Prescott (1975) for evaluating the intensity of foliar diseases in wheat can be used for screening purposes. However, new scales have been proposed to score severity of leaf damage in wheat and several other small grain cereals, such as triticale, barley and rye (Duveiller, 1994c). Disease rating can be done at flowering and again at the early dough stage (Zadoks' DC 80).
Immunity does not occur with BLS. Disease may occur even in seemingly resistant parents, provided inoculum pressure is sufficiently strong and the disease has enough time to develop. Although BLS resistance has been identified globally in wheat (Akhtar and Aslam, 1985; Bamberg, 1936; Boosalis, 1952; Thompson and Souza, 1989; Duveiller, 1990b; El Attari et al., 1996; Hagborg, 1974; Milus and Mirlohi, 1994; Milus et al., 1996), very little information is available on its mode of inheritance. Recent research conducted in the field in Mexico showed that five genes condition BLS resistance in five wheat lines (Turaco, Alondra, Angostura, Mochis and Pavon). Cultivars Pavon and Mochis showed the highest level of resistance. None of the five genotypes contained the full set of identified resistance genes, which suggests there are cultivars with more resistance than Pavon and Mochis (Duveiller et al., 1993).
CONCLUDING REMARKS
Bacterial leaf streak is a sporadic but wide-spread disease of wheat that can cause significant losses. The major problem is that the disease is seed-borne. Although zero tolerance of bacteria in the seed is not required due to its low transmission rate, there is a very real possibility that primary inoculum may increase during seed multiplication. The risk of disease is variable in many wheat-growing areas of the world, but the possibility of it occurring in areas where it is not usually found should not be overlooked. Fortunately, a specific succession of events is necessary to induce an epidemic. If one of the events required for disease development does not occur, the epidemic may not materialize. Black chaff incidence, severity and distribution may thus vary from year to year, even in disease-prone areas.
Epidemics of bacterial leaf streak may occur in various scenarios. This explains why the disease has a global distribution and is sporadic in areas as different as sprinkler-irrigated wheat fields in the United States, Mexican highlands characterized by marked daytime temperature changes and the Southern Cone countries of South America, where warm and cloudy days may occur alternately. Because disease occurrence is sporadic, research on epidemiology and resistance is particularly difficult and, consequently, advances in controlling BLS are slow.
Discarding infected seed prior to planting should be the primary control measure, since sowing pathogen-free seed is the first logical step in avoiding an outbreak. Seed indexing procedures are not routinely practised in many places but should be encouraged. The apparent absence of races and the wide-spread distribution of the pathogen are not convincing reasons for not implementing seed health procedures to limit the initial inoculum. Foundation seed should be multiplied in disease-free areas where climatic conditions are unfavourable for the development of epidemics. Seed should be disinfected before sowing even if currently available seed treatments are not fully satisfactory.
The most economical and environmentally friendly way of controlling BLS is through genetic resistance, and sources of incomplete genetic resistance have been identified. Differences in the degree of susceptibility are more easily observed in the field in disease-prone areas where artificial epidemics allowing the consistent differentiation between susceptible and resistant genotypes can be induced. Screening for resistance should be encouraged in areas where pathogen populations present the most variation.
REFERENCES
Akhtar, M.A. & Aslam, M. 1985. Bacterial stripe of wheat in Pakistan. Rachis, 4: 49.
Akhtar, M.A. & Aslam, M. 1986. Xan-thomonas campestris pv. undulosa on wheat. Rachis, 5: 34-37.
Alizadeh, A., Barrault, G., Sarrafi, A., Rahimian, H. & Albertini, L. 1995. Identification of bacterial leaf streak of cereals by their phenotypic characteristics and host range in Iran. Europ. J. Plant Pathol., 101: 225-229.
Bamberg, R.H. 1936. Black chaff disease of wheat. J. Agric. Res., 52: 397-417.
Boosalis, M.G. 1952. The epidemiology of Xanthomonas translucens (J.J. and R.) Dowson on cereals and grasses. Phytopathology, 42: 387-395.
Bradbury, J.F. 1986. Guide to plant pathogenic bacteria. Farnham House, Slough, UK, Mycological Institute, CAB International. 332 pp.
Bragard, C. & Maraite, H. 1994. Pathogenic variation in Xanthomonas campestris pv. undulosa. In M. Lemattre, S. Freigoun, K. Rudolph & J.R. Swings, eds. Proc. 8th Int. Conf. Plant Pathogenic Bacteria, Versailles, France, Les Colloques 66, p. 807-812. Paris, INRA/ORSTOM.
Bragard, C. & Verhoyen, M. 1993. Monoclonal antibodies specific for Xan-thomonas campestris bacteria pathogenic on wheat and on other small grains, in comparison with polyclonal antisera. J. Phytopathol., 139: 217-228.
Bragard, C., Mehta, Y.R. & Maraite, H. 1993. Serodiagnostic assays vs. the routine techniques to detect Xanthomonas campestris pv. undulosa in wheat seeds. J. Phytopathol., 18: 42-50.
Bragard, C., Verdier, V. & Maraite, H. 1995. Genetic diversity among Xanthomonas campestris strains pathogenic for small grains. Appl. Environ. Microbio., 61: 1020-1026.
Bragard, C., Singer, E., Alizadeh, A., Vauterin, L., Maraite, H. & Swings, J. 1997. Xanthomonas translucens from small grains: diversity and phytopathological relevance. Phytopathology, 87: 1111-1117.
Burton, G.J.L. 1931. Annual report of the senior plant breeder 1931. Kenya Dept. Agric. Ann. Rep., 176-209.
CIMMYT. 1977. Israel. In CIMMYT Report on Wheat Improvement, p. 237. Mexico, DF.
Claflin, L.E. & Ramundo, B.A. 1987. Evaluation of the dot-immunobinding assay for detecting phytopathogenic bacteria in wheat seeds. J. Seed Tech., 11: 52-61.
Demir, G. & Üstün, N. 1992. Studies on bacterial streak disease (Xanthomonas campestris pv. translucens (Jones et al.) dye of wheat and other gramineae. J. Turk. Phytopathol., 21: 33-40.
Dowson, W.J. 1939. On the systematic position and generic names of the Gram negative bacterial plant pathogens. Zentralblatt für Bakt. etc. II. Abt. Bd. 100, 9/13: 177-193.
Duveiller, E. 1990a. A seed detection method of Xanthomonas campestris pv. undulosa, using a modification of Wilbrink's agar medium. Parasitica, 46: 3-17.
Duveiller, E. 1990b. Screening criteria for bacterial leaf streak in bread wheat, durum wheat and triticale in CIMMYT. In Z. Klement, ed. Proc. 7th Int. Conf. Plant Pathogenic Bacteria, Budapest, II, p. 1011-1016. Budapest, Akadémiai Kiadó.
Duveiller, E. 1994a. A study of Xanthomonas campestris pv. undulosa populations associated with symptomless wheat leaves. Parasitica, 50: 109-117.
Duveiller, E. 1994b. Bacterial leaf streak or black chaff of cereals. Bull. OEPP/EPPO Bull., 24: 135-157.
Duveiller, E. 1994c. A pictorial series of disease assessment keys for bacterial leaf streak of cereals. Plant Dis., 78: 137-141.
Duveiller, E. & Bragard, C. 1992. Comparison of immunofluorescence and two assays for detection of Xanthomonas campestris pv. undulosa in seeds of small grains. Plant Dis., 76: 999-1003.
Duveiller, E. & Maraite, H. 1993. Study of yield loss due to Xanthomonas campestris pv. undulosa in wheat under high rainfall temperate conditions. J. Plant Dis. Prot., 100(5): 453-459.
Duveiller, E. & Maraite, H. 1995. Effect of temperature and air humidity on multiplication of Xanthomonas campestris pv. undulosa and symptom expression in susceptible and field-tolerant wheat genotypes. J. Phytopathol., 143: 227-232.
Duveiller, E., Bragard, C. & Maraite, H. 1991. Bacterial diseases of wheat in the warmer areas - reality or myth? In D. Saunders, ed. Wheat for the Non-traditional Warm Areas. Proc. Int. Conf., Iguazu Falls, Brazil, p. 189-202. Mexico, DF, CIMMYT.
Duveiller, E., van Ginkel, M. & Tijssen, M. 1993. Genetic analysis of resistance to bacterial leaf streak caused by Xan-thomonas campestris pv. undulosa in bread wheat. Euphytica, 66: 35-43.
Duveiller, E., Fucikovsky, L. & Rudolph, K., eds. 1997. The bacterial diseases of wheat: concepts and methods of disease management. Mexico, DF, CIMMYT. 78 pp.
El Attari, H., Sarrafi, A., Garrigues, S., Dechamp-Guillaume, G. & Barrault, G. 1996. Diallel analysis of partial resistance to an Iranian strain of bacterial leaf streak (Xanthomonas campestris pv. cerealis) in wheat. Plant Pathol., 45: 1134-1138.
Forster, R.L. 1982. The status of black chaff disease in Idaho. In Idaho wheat (Dec. issue). Owyhee Plaza Hotel, Boise, ID, USA, Idaho State Wheat Growers Association. 20 pp.
Forster, R.L. & Schaad, N.W. 1985. Evaluation of seed treatments for eradication of Xanthomonas campestris pv. translucens from wheat seed. Phytopathology, 75: 1385.
Forster, R.L. & Schaad, N.W. 1987. Tolerance levels of seed borne Xanthomonas campestris pv. translucens, the causal agent of black chaff of wheat. In E.L. Civerelo, A. Collmer, R.E. Davis & A.G. Gillaspie, eds. Proc. 6th Int. Conf. Plant Pathogenic Bacteria, Maryland, USA, p. 974-975. Dordrecht, Netherlands, Martinus Nijhoff Publishers.
Forster, R.L. & Schaad, N.W. 1988. Control of black chaff of wheat with seed treatment and a foundation seed health program. Plant Dis., 72: 935-938.
Forster, R.L. & Schaad, N.W. 1990. Longevity of Xanthomonas campestris pv. translucens in wheat seed under two storage conditions. In Z. Klement, ed. Proc. 7th Int. Conf. Plant Pathogenic Bacteria, Budapest, Part A, p. 329-331. Budapest, Akadémiai Kiadó.
Forster, R.L., Mihuta-Grimm, L. & Schaad, N.W. 1986. Black chaff of wheat and barley. Current Information Series No. 784, p. 2. University of Idaho, College of Agriculture.
Fourest, E., Rehms, L.D., Sands, D.C., Bjarko, M. & Lund, R.E. 1990. Eradication of Xanthomonas translucens from barley seed with dry heat treatments. Plant Dis., 74: 816-818.
Frommel, M.I. 1986. Xanthomonas campestris pv. translucens, causal agent of bacterial streak of wheat (Triticum aestivum). Montevideo, Uruguay, Dirección de Sanidad Vegetal (in Spanish).
Hagborg, W.A.F. 1974. Notes on bacterial diseases of cereals and some other crop plants. Can. Plant Dis. Surv., 54: 129-151.
Johnson, T. & Hagborg, W.A.F. 1944. Melanism in wheat induced by high temperature and humidity. Can. J. Res., 22(C): 7-10.
Jones, L.R., Johnson, A.G. & Reddy, C.S. 1917. Bacterial blight of barley. J. Agric. Res., 11: 625-643.
Jons, V.L., Hosford, R.M., Jr. & Lamey, H.A. 1982. Brown streak of wheat leaves caused by Xanthomonas campestris pv. undulosa in North Dakota. In R.M. Hosford, ed. Proc. Tan Spot of Wheat and Related Diseases Workshop, p. 110-113. Fargo, ND, USA, North Dakota Agricultural Experiment Station, North Dakota State University.
Kim, H.K., Orser, C., Lindow, S.E. & Sands, D.C. 1987. Xanthomonas campestris pv. translucens strains active in ice nucleation. Plant Dis., 71: 994-997.
Korobko, A.P., Wondimagegne, E. & Anisimoff, B.V. 1985. Bacterial stripe and black chaff of wheat in Ethiopia. Ambo, Ethiopia, Scientific Phytopathological Laboratory. 5 pp.
Maes, M. & Garbeva, P. 1994. Detection of bacterial phytopathogens based on nucleic acid technology. Parasitica, 50 (1-2): 75-80.
Maes, M., Garbeva, P. & Kamoen, O. 1996. Recognition and detection in seed of the Xanthomonas pathogens that cause cereal leaf streak using rDNA spacer sequences and polymerase chain reaction. Phytopathology, 86: 63-69.
Mamluk, O.F., Al-Ahmed, M. & Makki, M.A. 1990. Current status of wheat diseases in Syria. Phytopath. Medit., 29: 143-150.
Mehta, Y.R. 1986. Effect of Guazatin Plus to control Xanthomonas campestris pv. undulosa in wheat. In Reunion Nacional de Pesquisa em Trigo, Brazil, p. 56. Londrina, Brazil, IAPAR (in Portuguese).
Mehta, Y.R. 1990. Management of Xan-thomonas campestris pv. undulosa and hordei through cereal seed testing. Seed Sci. Tech., 18: 467-476.
Mehta, Y.R. & Bassoi, M.C. 1993. Guazatine Plus as a seed treatment bactericide to eradicate Xanthomonas campestris pv. undulosa from wheat seeds. Seed Sci. Tech., 21: 9-24.
Milus, E.A. & Chalkley, D.B. 1994. Virulence of Xanthomonas campestris pv. translucens on selected wheat genotypes. Plant Dis., 78: 612-615.
Milus, E.A. & Mirlohi, A.F. 1994. Use of disease reactions to identify resistance in wheat to bacterial streak. Plant Dis., 78: 157-161.
Milus, E.A., Duveiller, E., Kirkpatrick, T.L. & Chalkey, D.B. 1996. Relationships between disease reactions under controlled conditions and severity of wheat bacterial streak in the field. Plant Dis., 80: 726-730.
Mohan, S.K. & Mehta, Y.R. 1985. Studies on Xanthomonas campestris pv. undulosa in wheat and triticale in Paraná State. Fitopatologia Brasileira, 10: 447-453 (in Portuguese).
Noble, R.J. 1935. Australia: notes on plant diseases recorded in New South Wales for the year ending 30th June 1935. Int. Bull. Plant Prot., 12: 270-273 (abstract: RAM 15, 280).
Paul, V.H. & Smith, I.M. 1989. Bacterial pathogens of gramineae: systematic review and assessment of quarantine status for the EPPO region. Bull. OEPP/EPPO Bull., 19: 33-42.
Reddy, C.S., Godkin, J. & Johnson, A.G. 1924. Bacterial blight of rye. J. Agric. Res., 28: 1039-1040.
Richardson, M.J. & Waller, J.M. 1974. Triticale diseases in CIMMYT trial locations. In Triticale: Proc. Int. Symp., El Batan, Mexico, Monograph 024e, p. 193-199. Ottawa, Canada, International Development Research Center.
Saari, E.E. & Prescott, J.M. 1975. A scale for appraising the foliar intensity of wheat diseases. Plant Dis. Rep., 59: 377-380.
Sands, D.C. & Fourest, E. 1989. Xan-thomonas campestris pv. translucens in North and South America and in the Middle East. Bull. OEPP/EPPO Bull., 19: 127-130.
Sands, D.C., Mizrak, G., Hall, V.N., Kim, H.K., Bockelman, H.E. & Golden, M.J. 1986. Seed transmitted bacterial diseases of cereals: epidemiology and control. Arab J. Plant Prot., 4: 127-125.
Schaad, N.W. 1987a. The use and limitations of methods to detect seed borne bacteria. In Seed pathology, vol. 2, International advanced course, p. 324-332. Passo Fundo, RS, Brazil, University of Passo Fundo.
Schaad, N.W. 1987b. Problems with the pathovar concept. In E.L. Civerelo, A. Collmer, R.E. Davis & A.G. Gillaspie, eds. Proc. 6th Int. Conf. Plant Pathogenic Bacteria, Maryland, USA, p. 783-785. Dordrecht, Netherlands, Martinus Nijhoff Publishers.
Schaad, N.W. 1988a. Bacteria. In Symp. Inoculum Thresholds of Seed-borne Pathogens. 76th Annual Meeting of the American Phytopathological Society. Phytopathology, 78: 872-875.
Schaad, N.W., ed. 1988b. Laboratory guide for identification of plant pathogenic bacteria. Moscow, ID, USA. 164 pp.
Schaad, N.W. & Forster, R.L. 1985. A semi-selective agar medium for isolating Xanthomonas campestris pv. translucens from wheat seeds. Phytopathology, 75: 260-263.
Schaad, N.W. & Forster, R.L. 1993. Black chaff. In S.B. Mathur & B.M. Cunfer, eds. Seed-borne diseases and seed health testing of wheat, p. 129-136. Frederiksberg, Denmark, Jordbrugsforlaget.
Smit, I.B.J. & Van A. Bredenkamp, T. 1988. Items from South Africa: international nurseries. Ann. Wh. Newsl., 34: 84.
Smith, E.F. 1917. A new disease of wheat. J. Agric. Res., 10: 51-54.
Smith, E.F., Jones, L.R. & Reddy, C.S. 1919. The black chaff of wheat. Science, 50: 48.
Sun, F. & He, L. 1986. Studies on determinative techniques for resistance of wheat to black chaff (Xanthomonas translucens f. sp. undulosa). Acta Phytophylacica Sinica, 13: 109-115 (in Chinese; English summary).
Tessi, J.L. 1949. Current status of work on Xanthomonas translucens var. cerealis. Presented at the 4th Wheat, Oats, Barley and Rye Meeting, Castellar, Argentina, p. 200.
Thompson, D.C. & Souza, E.J. 1989. Reaction of spring wheat cultivars to black chaff, 1988. American Phytopathological Society. Biol. Cult. Tests, 4: 51.
Thompson, D.C., Schaad, N.W. & Forster, R.L. 1989. New perennial hosts of epiphytic populations of Xanthomonas campestris pv. translucens. Phytopathology, 79: 1168 (abstr.).
Vauterin, L., Hoste, B., Kersters, K. & Swings, J. 1995. The relationship within genus Xanthomonas and a proposal for a new classification. Int. J. Syst. Bacteriol., 45: 472-489.
Waller, J.M. 1976. The influence of climate on the incidence and severity of some diseases of tropical crops. Rev. Plant Pathol., 55: 185-194.
Wallin, J.R. 1946. Parasitism of Xanthomonas translucens (J.J. and R.) Dowson on grasses and cereals. Iowa St. Coll. J. Sci., 20: 171-193.
Young, J.M., Saddler, G.S., Takikawa, Y., De Boer, S.H., Vauterin, L., Gardan, L., Gvozdyak, R.I. & Stead, D.E. 1996. Names of plant pathogenic bacteria 1864-1995. Rev. Plant Pathol., 75: 721-763.
Zadoks, J.C., Chang, T.T. & Konzak, C.F. 1974. A decimal code for the growth stages of cereals. Weed Res., 14: 415-421.
Zillinsky, F.J. & Borlaug, N.E. 1971. Progress in developing triticale as an economic crop. Int. Maize Wheat Improv. Cen. Res. Bull., 17: 18-21.
