Xanthomonas translucens pv. translucens (bacterial leaf streak of barley)
- Summary of Invasiveness
- Taxonomic Tree
- Notes on Taxonomy and Nomenclature
- Distribution Table
- History of Introduction and Spread
- Risk of Introduction
- Hosts/Species Affected
- Host Plants and Other Plants Affected
- Growth Stages
- List of Symptoms/Signs
- Biology and Ecology
- Means of Movement and Dispersal
- Seedborne Aspects
- Pathway Causes
- Pathway Vectors
- Plant Trade
- Wood Packaging
- Impact Summary
- Risk and Impact Factors
- Uses List
- 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 translucens pv. translucens (Jones et al. 1917) Vauterin et al. 1995
Preferred Common Name
- bacterial leaf streak of barley
Other Scientific Names
- Bacterium translucens Jones, Johnson & Reddy 1917
- Phytomonas translucens (Jones, Johnson & Reddy) Bergey et al.
- Pseudomonas translucens Jones, Johnson & Reddy 1917
- Xanthomonas campestris pv. hordei (Hagborg 1942) Dye 1978
- Xanthomonas campestris pv. translucens (Jones et al. 1917) Dye 1978
- Xanthomonas translucens (Jones, Johnson & Reddy) Dowson 1939
- Xanthomonas translucens f.sp. hordei-avenae Hagborg (1942)
- Xanthomonas translucens pv. hordei (Egli et al. 1975) Vauterin et al. 1995
International Common Names
- English: bacterial leaf streak of cereals; bacterial leaf streak of small grain cereals; bacterial leaf stripe; BLS; cereal black chaff; xanthomonas leaf streak
- Spanish: gluma negra; quemadura bacteriana de los cereals; quemadura bacteriana del trigo; quemaduras bacterianas de la cebada
- French: brûlure bactérienne de l'orge; brûlure bactérienne du blé; glume noire des céréales
- German: Bakterienbrand: Gerste; Schwarzspelzigkeit: Getreide; Spelzenbräune: Getreide
Summary of InvasivenessTop of page
Bacterial leaf streak caused by X. translucens pv. translucens is an economically important disease of barley around the world. The disease occurs in many countries with particular importance in regions characterized by high precipitation. As a seedborne pathogen, X. translucens pv. translucens is included in the A2 list of quarantine pathogens by the European and Mediterranean Plant Protection Organization (EPPO).
Taxonomic TreeTop of page
- Domain: Bacteria
- Phylum: Proteobacteria
- Class: Gammaproteobacteria
- Order: Xanthomonadales
- Family: Xanthomonadaceae
- Genus: Xanthomonas
- Species: Xanthomonas translucens pv. translucens
Notes on Taxonomy and NomenclatureTop of page
Bacterial leaf streak of barley was first reported in 1917 in the USA (Jones et al., 1917). The causal agent was named Bacterium translucens because of the translucent lesions on symptomatic leaves. However, according to Bamberg (1936), the occurrence of a leaf streak-like disease on wheat and barley had been observed and recorded as early as 1893. Subsequently, a similar disease was reported on wheat by Smith et al. (1919) and the pathogen was named Bacterium translucens var. undulosum because it morphologically resembled the barley pathogen and was able to infect barley through artificial inoculation. Once Dowson (1939) created the genus Xanthomonas, the cereal pathogens were reclassified as X. translucens.
Subsequently, Hagborg (1942) subdivided X. translucens into five formae speciales on the basis of natural host as well as the ability to infect hosts using artificial inoculation. Accordingly, the species X. translucens was divided into f.sp. hordei (infecting barley), f.sp. undulosa (infecting wheat, barley and rye), f.sp. secalis (infecting rye), f.sp. hordei-avenae (infecting barley and oat) and f.sp. cerealis (infecting wheat, barley, rye and oat). Fang et al. (1950) combined f.sp. hordei-avenae and hordei to be a single taxon. During the 1970s and 1980s, the entire X. translucens members were transferred into X. campestris as different pathovars, which included X. campestris pvs. cerealis, hordei, secalis, translucens and undulosa (Young et al.,1978; Dye et al., 1980).
On the basis of host specificity, the International Society for Plant Pathology (ISPP) has recognized five Xanthomonas campestris pathovars (cerealis, hordei, secalis, translucens and undulosa) (Young et al., 1978; Dye et al., 1980) and these pathovar names have been retained (Vauterin et al., 1995). However, these names have been used rather indiscriminately in the literature. Duveiller (1994a) and Bragard et al., (1995) demonstrated the need for a revision of pathovar names with amalgamation as synonyms of these pathovars which attack small grains. The taxonomy and nomenclature of Xanthomonas spp. were revised in 1995 on the basis of DNA-DNA hybridization and substrate utilization, along with some data on fatty acid analysis and SDS-PAGE of proteins. Thus, Vauterin et al. (1995) have re-established all X. campestris pathovars infecting cereals, and the strains causing diseases on members of the Poaceae were grouped as Xanthomonas translucens (Jones et al., 1917; Vauterin et al., 1995).
Subsequently, Bragard et al. (1995, 1997) suggested that X. translucens pv. translucens is a synonym of X. translucens pv. hordei, and the translucens group of X. translucens contains three true biological entities, cerealis, translucens and undulosa, with pv. translucens pathogenic on barley, pv. undulosa pathogenic to both barley and wheat, and pv. cerealis pathogenic to barley, wheat, oat and bromegrass. During the past few years, the taxonomy of X. translucens has been refined using multilocus sequence analysis and typing (MLSA/MLST) as well as complete genome sequence-based phylogenetic analyses (Wichmann et al., 2013; Gardiner et al., 2014; Peng et al., 2016; Curland et al., 2018, Khojasteh et al., 2019). These studies have confirmed the pathovar status of the xanthomonad pathogens infecting cereals. Other references on taxonomy and nomenclature include Jones et al. (1917), Smith (1917), Reddy et al. (1924), Hagborg (1942) and Vauterin et al. (1992).
DescriptionTop of page
X. translucens pv. translucens is an aerobic, Gram-negative rod with one polar flagellum. Cells measure 0.5 x 1.0 µm. On YDC agar medium, the colonies are pale yellow, mucoid, convex and smooth with entire margins (Wiese, 1987). On X. translucens agar isolation medium, colonies have a similar appearance except that they tend to be clearer (less mucoid), and some colonies appear flattened rather than convex (Schaad and Forster, 1989).
DistributionTop of page
Due to continuous taxonomic complexities within the species X. translucens since the first description of the disease in 1917, the exact distribution of each pathovar has not yet been determined. The information provided in the literature before the reclassification of the species in 1995 (Vauterin et al., 1995) might have referred to bacterial leaf streak as a disease complex instead of determining pathovar status of the pathogen. Even after the reclassification of the species in 1995, most descriptions of the pathogen in areas with no history of the disease did not accurately determine the status of the pathogen to pathovar level. Hence, the information provided in the following table does not necessarily refer to the geographic distribution of the barley pathogen X. translucens pv. translucens. Rather, it shows the global distribution of bacterial leaf streak disease caused by the complex species X. translucens.
See also CABI/EPPO (1998, No. 288).
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: 20 Nov 2020
|Continent/Country/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
|Libya||Absent, Unconfirmed presence record(s)||Original citation: Duveiller, 1994a|
|Belgium||Absent, Formerly present|
|Bulgaria||Absent, Formerly present||1981|
|France||Absent, Formerly present|
|Poland||Absent, Unconfirmed presence record(s)||Original citation: Duveiller, 1994a|
|-Central Russia||Present, Localized|
|-Southern Russia||Present, Localized|
|Spain||Absent, Intercepted only|
|Sweden||Absent, Formerly present|
|United States||Present, Widespread|
|-New South Wales||Present|
|-Mato Grosso do Sul||Present|
History of Introduction and SpreadTop of page
Due to continuous taxonomic complexities within the species X. translucens since the first description of the disease in 1917, the exact distribution of each pathovar has not yet been determined. The information provided in the literature before the reclassification of the species in 1995 (Vauterin et al., 1995) may refer to bacterial leaf streak as a disease complex instead of determining the pathovar of the pathogen. Even after the reclassification of the species in 1995, most descriptions of the pathogen in areas with no history of the disease do not accurately determine the status of the pathogen to pathovar level. Hence, the information provided in this datasheet does not necessarily refer to the geographic distribution of the barley pathogen X. translucens pv. translucens, rather, it shows the global distribution of bacterial leaf streak disease caused by the species complex X. translucens.
Risk of IntroductionTop of page
RISK CRITERIA CATEGORY
ECONOMIC IMPORTANCE Moderate
SEEDBORNE INCIDENCE Moderate
SEED TRANSMITTED Yes
SEED TREATMENT Yes
OVERALL RISK Moderate to high depending on the climate and environmental conditions
Notes on phytosanitary risk
X. translucens pv. translucens is listed as a prohibited organism on any plant material in the EPPO region and has been included in the EPPO A2 list since 1993 and the IAPSC (Inter-African Phytosanitary Council) A2 list since 1989. Several other countries in Asia have listed the pathogen as a regulated quarantine agent (https://gd.eppo.int/taxon/XANTTR/categorization).
Hosts/Species AffectedTop of page
Although different pathovars of the complex species X. translucens are capable of infecting a wide range of Poaceae and dicotyledonous plants, e.g. pistachio, the host range of X. translucens pv. translucens is limited to barley (Roman-Reyna et al., 2020). Mellano and Cooksey (1988) determined that independent positive factors determine host range in X. translucens rather than an avirulence gene system, such as that determining race specificity in other plant pathogens. Waney et al. (1991) identified host-specific virulence loci from X. translucens. Inactivation of these host-specific virulence genes narrowed the host range by one plant genus and did not affect the non-host hypersensitive response. Existence of these host-specific virulence genes may explain why the pathogens of small grains have over-lapping host ranges. Recently, Khojasteh et al. (2020) investigated the transcription activator-like effectors (TALE) diversity in Iranian strains of X. translucens which reflected the TALE-based pathovar differentiation and host speciation of X. translucens pv. translucens and X. translucens pv. undulosa.
Host Plants and Other Plants AffectedTop of page
|Hordeum vulgare (barley)||Poaceae||Main|
Growth StagesTop of page
SymptomsTop of page
Field symptoms of bacterial leaf streak caused by X. translucens pv. translucens are similar to those initiated by the wheat and barley pathogen X. translucens pv. undulosa, hence the two pathovars are indistinguishable using symptoms on host plants (Khojasteh et al., 2019). Under field conditions, symptoms may be found at all growth stages, but are most frequently observed after flowering. On leaves, bacterial streak symptoms begin as small, light-brown, water-soaked streaks that tend to be confined by the veins. Often the lesions will begin at the edges or midrib of leaves. On very susceptible hosts under moist conditions, the streaks will elongate, parallel to the veins, and develop a dark-brown water-soaking that is translucent when held up to light. Bacterial exudate in the form of small yellow droplets may be associated with the streaks when leaves are wet. Under dry conditions, the exudate will appear as small yellow granules, or thin shiny scales. On less susceptible hosts, or in dry conditions, individual streaks are likely to remain small and turn chlorotic or necrotic within a few days. However, numerous streaks may coalesce to kill large portions of a leaf, or even the entire leaf. Bacterial exudate is not conspicuous on less susceptible hosts, especially under dry conditions.
Symptoms on the heads are called 'black chaff' and appear as distinct thin black streaks between the veins beginning at the tips of the glumes and lemmas. When severe, the streaks may coalesce giving the heads a dark appearance, and the grain may be brown and shrivelled. The pathogen may also attack the stem and peduncle, causing black streaks or uniformly black lesions. On certain cultivars, black peduncle lesions may have light-coloured centres.
Seeds from diseased plants may be shrivelled and usually have reduced test weight. Shane et al. (1987) reported up to a 34% loss in kernel weight. No information is available on the effects of infection on germination or seedling emergence, but it is not likely that these variables would be affected greatly unless the seed was severely shrivelled.
List of Symptoms/SignsTop of page
|Inflorescence / lesions on glumes|
|Leaves / abnormal colours|
|Leaves / necrotic areas|
|Leaves / odour|
|Seeds / discolorations|
|Seeds / galls|
|Seeds / lesions on seeds|
|Stems / discoloration of bark|
Biology and EcologyTop of page
Unless otherwise stated, the biological and ecological features among the three pathovars X. translucens pv. translucens, X. translucens pv. undulosa and X. translucens pv. cerealis are similar (Duveiller and Bragard, 2017). Each X. translucens pathovar can cause disease on one or more members of the Poaceae, including barley, triticale, rye and wheat. X. translucens is adapted to grow as an epiphyte on wheat (Azad and Schaad, 1988a; Duveiller, 1994c), Bromus inermis and Phleum pratense (Wallin, 1946), Elymus repens (Boosalis, 1952), Brachiaria plantaginea, Cenchrus echinatus, Digitaria horizontalis and Eleusine indica, and tomato (Timmer et al., 1987). Epiphytic growth of plant pathogenic bacteria is probably a prerequisite to infection (Leben, 1965; Mew et al., 1986; Osdaghi et al., 2016; Osdaghi et al., 2018a; Zarei et al., 2018). There is probably little saprophytic growth and multiplication of X. translucens in the absence of green plants.
Transmission by seed is the most likely means of long-distance dissemination and survival from year to year (Sands et al., 1986; Mehta, 1990; Mehta et al., 1992; Milus and Mirlohi, 1995). The pathogen can remain viable in infested seeds for more than 5 years (Forster and Schaad, 1990). Little is known about the location of the pathogen in seed, but it is suspected that the most important inoculum is carried internally.
Schaad and Forster (1985) determined that seed lots with fewer than 1000 c.f.u./ml in seed washing (approximately 42 c.f.u./seed) were unlikely to develop disease under irrigated growing conditions in Idaho, USA. Mehta (1990) determined that seed production fields with less than 10% bacterial streak severity at the soft dough stage were likely to produce seeds with seedborne inoculum levels low enough to be suitable for planting under most conditions in Brazil. Milus and Mirlohi (1995) were able to demonstrate transmission rates of 4 to 25%, depending on the seed lot, for a rifampicin-resistant strain on a susceptible wheat cultivar under growth chamber conditions, but barely detectable transmission under field conditions with the same seed lots. More information is needed on thresholds for transmission of seedborne inoculum.
Over-wintering in infested crop debris and on alternative hosts may be possible in colder climates where spring small grains are grown (Boosalis, 1952), but the pathogen does not survive hot and dry over-summering periods where small grains are grown during cooler times of the year (Milus and Mirlohi, 1995). The pathogen does not survive long periods in soil unless it is associated with infested crop debris (Boosalis, 1952; Cunfer, 1988).
It has been suggested that the bacterial leaf streak pathogen enters plant tissues through stomata and mainly colonizes mesophyll. However, Sapkota et al. (2020) showed that using the spray inoculation method for X. translucens pv. translucens, little or no disease developed on barley leaves, suggesting that it may have a different tissue specificity. Leaf clipping and dip inoculation was successfully used by Pesce et al. (2017) to induce disease on barley. Within fields, X. translucens is most effectively disseminated by wind-driven rain, but the pathogen can also be disseminated by aphids, wind, infested crop debris, and plant-to-plant contact (Boosalis, 1952; Cunfer, 1987). X. translucens is the only known Xanthomonas species capable of ice nucleation, and may play a role in frost injury on host and non-host plants (Kim et al., 1987; Azad and Schaad, 1988a).
ClimateTop of page
|C - Temperate/Mesothermal climate||Preferred||Average temp. of coldest month > 0°C and < 18°C, mean warmest month > 10°C|
|Cs - Warm temperate climate with dry summer||Preferred||Warm average temp. > 10°C, Cold average temp. > 0°C, dry summers|
|Cw - Warm temperate climate with dry winter||Preferred||Warm temperate climate with dry winter (Warm average temp. > 10°C, Cold average temp. > 0°C, dry winters)|
|Cf - Warm temperate climate, wet all year||Preferred||Warm average temp. > 10°C, Cold average temp. > 0°C, wet all year|
Means of Movement and DispersalTop of page
Transmission by seed is the most likely means of long-distance dissemination and survival from year to year.
Milus and Mirlohi (1995) demonstrated transmission rates of 4 to 25%, depending on the seed lot, for a rifampicin-resistant strain on a susceptible wheat cultivar under growth chamber conditions, but transmission was barely detectable under field conditions with the same seed lots.
Seedborne AspectsTop of page
Different pathovars of X. translucens are seedborne in wheat, barley, rye and triticale as well as some grasses (Mehta, 1990; Richardson, 1990). However, X. translucens pv. translucens is mainly restricted to barley and less likely to be transmitted by the seeds of other small grain cereals. Producing seed in dry environments using furrow irrigation will reduce bacterial streak and the level of seedborne X. translucens (Forster and Schaad, 1988) but some contamination is likely unless the production field is planted with seeds free of the pathogen and is well isolated from other sources of inoculum. Seed produced under such conditions would probably have a low risk of initiating an epidemic.
Bacterial streak can be responsible for great damage in winter cereals. Infected seeds and crop debris are the main form of survival and dissemination of these pathogens. Malavolta et al. (2000) carried out a study to determine the period of survival of X. translucens in wheat crop debris and wheat and triticale seeds, from naturally infected crops at Palotina, Brazil. In seeds of 10 genotypes of wheat and three of triticale, the pathogenic bacteria were slightly reduced in the first 2 years after harvest. After this period a rapid reduction of viable bacteria occurred, with no recovering of the pathogen 42 months after harvest, except for in one genotype of wheat. Pathogen survival in crop debris was longer than 30 months under laboratory conditions and shorter than 8 months under field conditions. Although there is rapid decline under field conditions, an overlapping of the pathogen survival period and a new planting may occur.
Effect on Seed Quality
Seeds from diseased plants may be shrivelled and usually have reduced test weight. Shane et al. (1987) reported losses of up to 34% in kernel weight. No information is available on the effects of infection on germination or seedling emergence, but it is not likely that these variables would be affected greatly unless the seed was severely shrivelled.
Transmission by seed is the most likely means of long-distance dissemination and survival from year to year (Sands et al., 1986; Mehta, 1990; Mehta et al., 1992; Milus and Mirlohi, 1995). Milus and Mirlohi (1995) were able to demonstrate transmission rates of 4 to 25%, depending on the seed lot, for a rifampicin-resistant strain on a susceptible wheat cultivar under growth chamber conditions, but transmission was barely detectable under field conditions with the same seed lots. When single wheat seeds were assayed using a combination of semi-selective enrichment and ELISA, there was a good correlation between the percentage of infected seeds and the percentage of diseased seedlings in a greenhouse assay (Frommel and Pazos, 1994). By correlating seedborne inoculum with development of the disease in field plantings, Schaad and Forster (1985) determined that seed lots with fewer than 1000 c.f.u./ml in seed washings (approximately 42 c.f.u./seed) were unlikely to develop disease under irrigated growing conditions in Idaho, USA. Mehta (1990) determined that seed production fields with less than 10% bacterial streak severity at soft dough stage were likely to produce seeds with seedborne inoculum levels low enough to be suitable for planting under most conditions in Brazil. However, more information is needed on thresholds for transmission of seedborne inoculum.
Tubajika et al. (1998) examined the relationship between foliar disease symptoms on parent plants, seed contamination by the causal bacterium (X. translucens pv. translucens) and subsequent development of bacterial leaf streak in wheat in microplots and in the laboratory to determine the role of seed transmission in disease epidemiology. Microplot experiments were carried out during the 1994-95 and 1995-96 growing seasons using seed harvested in Baton Rouge, Louisiana, USA, in 1994 and 1995, respectively. Treatments were seed lots from plants with differing levels of bacterial leaf streak severity on the flag leaves of the parent tillers. X. translucens pv. translucens was detected in 1 to 20% of seed from susceptible cultivars Florida 304 and Savannah collected from plants with leaf streak symptoms. Correlations between seed contamination and disease on plants that developed from this seed were detected only when seed came from parent tillers that expressed flag leaf disease severity of 15 to 20% in 1994-95 and 30 to 35% in 1995-96. However, symptoms of bacterial leaf streak on plants that developed from these seed were evident on only 3% of plants. Results suggest a possible threshold level for bacterial leaf streak on flag leaves is necessary before X. translucens pv. translucens can be detected in seed. Seedling emergence in microplots correlated negatively with leaf streak severity on parent tiller flag leaves. Artificial infestation of seed with X. translucens pv. translucens also reduced seed germination, but this was more evident in Savannah than in Florida 304.
Over-wintering in infested crop debris and on alternative hosts may be possible in colder climates where spring small grains are grown (Boosalis, 1952), but the pathogen does not survive hot and dry over-summering periods where small grains are grown during cooler times of the year (Milus and Mirlohi, 1995). The pathogen does not survive long periods in soil unless it is associated with infested crop debris (Boosalis, 1952; Cunfer, 1988).
There are no seed treatments that eradicate X. translucens from seed without excessive damage to the seed. Hot, acidified cupric acetate (Forster and Schaad, 1988), dry heat (Fourest et al., 1990) and Guzatine Plus (Mehta and Bassoi, 1993) have been shown to greatly reduce seedborne populations of X. translucens and bacterial streak in the field. However, acidified cupric acetate and dry heat are best suited to small seed lots, and none of the treatments are 100% effective in eradicating the pathogen or preventing transmission to plants (Duveiller, 1994a). Sands et al. (1986) reported a hot-water treatment at 53°C for 10 minutes followed by immediate cooling and drying.
Sun et al. (1988) reported control of black chaff with TF-128 in China, and de Luz et al. (1993) reported control of X. translucens with probenazole in Brazil, but no other information is available about these compounds.
Seed Health Tests
Duveiller and Bragard (2017) have provided a detail-oriented description of the seed test protocols used for the detection of the bacterial leaf streak pathogen. To find a selective medium for the pathogen, see Forster and Schaad (1996).
Extraction of the bacteria
- Add 1 drop of Tween 20 with a Pasteur pipette to 120 ml of sterile cold aqueous saline (0.85% NaCl) in a 250 ml Erlenmeyer flask and swirl to disperse.
- Add 3000 seeds (120 g) and shake vigorously on a rotary shaker at approximately 200 r.p.m. for 3-5 min at room temperature.
- Allow to settle for 1 min and make 10-fold dilutions to 103 using cold saline.
- Store an undiluted sample at 2°C for possible future use.
Culturing the bacteria
- Transfer 0.1 ml of the undiluted and 101, 102 and 103 dilutions onto three plates each of modified XTS agar (Schaad and Forster, 1985).
- Spread samples with an alcohol flamed, L-shaped glass rod and incubate plates at 30°C for 5 days.
- Streak or spread a suspension of a known culture of X. translucens onto modified XTS for comparison.
- Colonies of X. translucens pv. translucens are 1-2 mm in diameter, yellow, clear, round, convex to somewhat flat, and smooth.
- Count the total number of colonies of X. translucens and record as c.f.u./ml from 3000 seeds.
- Transfer several suspected colonies to YDC agar plates and incubate at 30°C for 2 days.
Identification of suspected colonies by pathogenicity tests
- Prepare inoculum of approximately 106 c.f.u./ml of X. translucens pv. translucens by suspending a 2 mm diameter colony growing on YDC in 2-3 ml of saline and making a 100-fold dilution in saline.
- Using a 26 gauge needle and syringe, inject inoculum into a susceptible seedling of a susceptible cultivar of barley at the 1-2 leaf stage about 1 cm above the soil line (i.e., at or slightly below the point where the first leaf starts to separate from the base of the seedling).
- Include a positive and negative check isolate in the test.
- Place inoculated plants and uninoculated checks on a greenhouse bench at 26-28°C with intermittent misting sufficient to keep the leaves continuously wet for the first 24 hours; thereafter, the misting may be eliminated. Alternatively, plants may be kept in a dew chamber at 26-28°C for the first 24 hours before being placed on a greenhouse bench.
- Observe water soaked streaks emanating from the needle hole in leaves after 8-10 days
Injection technique (Mehta, 1990)
Seed (20g) are shaken in 20 ml sterile saline for 90 minutes. The suspension is then inoculated into 20-day-old seedlings of a susceptible host. Occurrence of bacterial streak symptoms 7-12 days after inoculation indicated the presence of the pathogen in the seed lot. Mehta believed the technique to be efficient, inexpensive and suitable for quarantine purposes.
Serological methods (Frommel and Pazos, 1994)
This technique uses a combination of semi-selective enrichment and ELISA.
Duveiller and Bragard (2017) have described the following immunofluorescence technique to be used for the detection of the pathogen:
Pipette 40 μl of seed wash water directly from the flask into a 6-mm well on a multi-window slide and then fix with hot air from a hairdryer. Expose wells for 60 min to monoclonal antibody AB3-B6 [UCL, Belgium] diluted 100 times in phosphate buffered saline (PBS; 8 g of NaCl, 2.7 g of Na2HPO4. 2 H2O and 1 L of distilled water, pH 7.2); use 20 μl of antibody per well. Rinse wells with PBS and expose them for 30-60 min in the dark to a mouse anti-rat Mab conjugated with fluorescent isothiocyanate MARM4-FITC (IMEX, UCL, Brussels) diluted 1:100 in PBS. The optimum dilution will vary with each batch of conjugated antibody and therefore must be determined empirically on a known positive sample. Use 20 μl of antibody per well. Then, rinse with PBS, add three drops of buffered glycerine (100 mg of diphenylamine in 10 ml of PBS, pH 9.6, and 90 ml of glycerol); slip on a cover glass. Observe under immersion oil using a microscope equipped with a high pressure mercury ultraviolet lamp HBO-50 and Carl Zeiss filter combination x1000. If the preparation cannot be observed on the same day, the multi-window slides can be stored in the dark for later observation.
Maes et al. (1996) developed a PCR protocol for the detection and discrimination of cereal-pathogenic xanthomonads in seed lots. The PCR primers designed to exploit the variability of DNA sequences situated in the spacer segment between the 16S and 23S rRNA genes and flanking an alanine-tRNA gene. The primers amplify a 139-bp fragment from strains of leaf streak pathogens. The assays proved to be fast and relatively sensitive (28 × 103 c.f.u./g of seed) indicating the technique might be useful for detecting pathogens in seed (Duveiller and Bragard, 2017).
Pathway CausesTop of page
Pathway VectorsTop of page
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|
|Flowers/Inflorescences/Cones/Calyx||Yes||Yes||Pest or symptoms usually visible to the naked eye|
|Leaves||Yes||Yes||Pest or symptoms usually visible to the naked eye|
|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|
Wood PackagingTop of page
|Wood Packaging not known to carry the pest in trade/transport|
|Loose wood packing material|
|Processed or treated wood|
|Solid wood packing material with bark|
|Solid wood packing material without bark|
Impact SummaryTop of page
ImpactTop of page
Yield losses due to bacterial streak and black chaff were estimated to be 30-40% in the most severely diseased wheat fields in Idaho, USA (Forster and Schaad, 1988). Studies have indicated that yield loss due to bacterial leaf streak is generally negatively correlated with disease severity on flag leaves (Shane et al., 1987; Duveiller and Maraite, 1993; Sapkota et al., 2020). For spring wheat in Minnesota, USA, Shane et al. (1987) determined that 50% disease severity on the flag leaf reduced kernel weight by 8-13%, and 100% disease severity on the flag leaf resulted in a 13-34% loss. Duveiller (1992) reported a 20% average yield loss for wheat with 50% diseased leaf area.
Risk and Impact FactorsTop of page
- Invasive in its native range
- Proved invasive outside its native range
- Has a broad native range
- Abundant in its native range
- Highly adaptable to different environments
- Reproduces asexually
- Has high genetic variability
- Host damage
- Negatively impacts agriculture
- Negatively impacts animal/plant collections
- Negatively impacts trade/international relations
- Highly likely to be transported internationally accidentally
- Highly likely to be transported internationally deliberately
- Highly likely to be transported internationally illegally
- Difficult to identify/detect as a commodity contaminant
- Difficult to identify/detect in the field
- Difficult/costly to control
Uses ListTop of page
- Laboratory use
- Research model
DiagnosisTop of page
Dilution plating on a semi-selective medium has been the traditional method for confirming X. translucens as the pathogen. Schaad and Forster (1985) developed X. translucens agar medium for quantifying X. translucens in seed lots, and the medium is also useful for isolating the pathogen from all plant parts. However, the concentration of gentamicin in the original medium was toxic to some strains of X. translucens (Duveiller, 1989) and Forster et al. (1995) later modified the recipe (23 g Difco nutrient agar, 2.5 g glucose in 1 litre of distilled water plus 200 mg cycloheximide, 10 mg cephalexin, and 1450 µg gentamicin after autoclaving).
X. translucens also can be isolated on modified WBC agar (Duveiller, 1990; Duveiller and Bragard, 2017). See Duveiller (1990) for further details.
WBC medium: Bacto peptone 5 g; Sucrose 10 g; K2HPO4 0.5 g; MgSO4 7H2O 0.25 g; Na2SO3 (anhydrous) 0.05 g; Agar 15 g; Distilled water 850 ml. Mix with the following solution (autoclaved separately): Boric acid 0.75 g; Distilled water 150 ml. After cooling to 45ºC, add cycloheximide (in 2 ml of 75% ethanol) 75 mg and cephalexin (1 ml of a 10 mg/ml stock 10 mg solution in 75% ethanol).
Plates should be incubated at 25-30°C. Colonies should appear in 2-3 days, but it may take a few more days to develop their characteristic appearance. X. translucens colonies are pale yellow, convex and smooth with entire margins. It is best to dilution plate a known pathogenic strain under the same conditions as the strain being tested, for comparison.
To confirm cultures from single colonies as X. translucens, a quick pathogenicity test can be performed on a large number of isolates using seedlings of a susceptible host as described by Milus and Mirlohi (1993). Infiltrate primary leaves of 10-day-old seedlings with water using a 1-ml disposable syringe modified with a short piece of rubber tubing over the tip. Puncture the water-soaked area of a leaf with a needle carrying cells of each strain to be tested. Positive (known strain) and negative (sterile needle) checks should be included. Incubate plants at 25°C and 100% humidity for 12 hours followed by 48 hours at 25°C and ambient humidity. Pathogenic strains will cause water-soaking by the end of the incubation period. Longer incubation at 100% humidity will cause false positives. Another pathogenicity test using detached leaves on agar containing benzimidazole was developed by Colin et al. (1990).
Fatty acid analysis can be used to differentiate cultures of X. translucens from other bacteria (Stead, 1989; Yang et al., 1993) but this technique requires specialized equipment and computer software. Bacteriophages can also be used to differentiate X. translucens from other bacteria (Katznelson and Sutton, 1953; Mohan and Mehta, 1985) but some bacteriophages are specific to certain X. translucens strains which may limit their usefulness. Serological methods can be used to differentiate X. translucens (Elrod and Braun, 1947; Fang et al., 1950; Azad and Schaad, 1988b). X. translucens can also be characterized by restriction fragment length polymorphisms (Berthier et al., 1993).
Mehta (1990) proposed an injection technique to detect X. translucens in seed lots. Twenty grams of seeds were shaken in 20 ml sterile saline for 90 minutes. The suspension was then inoculated into 20-day-old seedlings of a susceptible host. Occurrence of bacterial streak symptoms 7-12 days after inoculation indicated the presence of the pathogen in the seed lot. Mehta believed the technique to be efficient, inexpensive and suitable for quarantine purposes.
Frommel and Pazos (1994) developed an ELISA procedure utilizing polyclonal antiserum that was able to detect 5000 c.f.u./ml of X. translucens. When semi-selective enrichment was performed on the sample before ELISA testing, the limit of detection was lowered to 500 c.f.u./ml. Single wheat seeds were assayed using a combination of semi-selective enrichment and ELISA, and there was a good correlation between the percentage of infected seeds and the percentage of diseased seedlings in a greenhouse assay. Possible disadvantages of this particular procedure for seed health testing are the use of polyclonal antiserum which is not readily reproducible, and the testing of individual seeds which is time-consuming. However, the procedures could probably be modified to work with bulk samples of seed. On the other hand, Bragard and Verhoyen (1993) developed monoclonal antibodies to X. translucens. One monoclonal was used with indirect immunofluorescence to detect X. translucens in several seed lots known to be infected. While this technique is still experimental and requires expensive equipment, it has the advantage of using monoclonal antibodies that can be reproduced indefinitely.
Langlois et al. (2017) developed diagnostic loop-mediated isothermal amplification (LAMP) primers based on genomic differences of representative strains within X. translucens. The LAMP technique clearly distinguishes strains that cause disease on cereals, such as pvs. translucens, undulosa, hordei and secalis, from strains that cause disease on non-cereal hosts, such as pvs. arrhenatheri, cerealis, graminis, phlei and poae. These primers will be instrumental in diagnostics when implementing quarantine regulations to limit further geographic spread of X. translucens. Multilocus sequence analysis (MLSA) of housekeeping genes is a reliable method for identification and discrimination of taxonomically complex xanthomonads e.g. members of X. euvesicatoria that cause disease on tomato, pepper and lucerne (Osdaghi et al., 2017; Yaripour et al., 2018; Osdaghi et al., 2018b). Recently, it has been showed that the MLSA of four housekeeping genes (i.e. dnaK, fyuA, gyrB, and rpoD) is suitable for the identification of cereal-pathogenic xanthomonads to pathovar level (Curland et al., 2018; Khojasteh et al., 2019).
Detection and InspectionTop of page
Inspect heads, peduncles and uppermost leaves for characteristic symptoms and dried exudate between the flowering and soft dough stages. It may be necessary to use a hand lens and hold the specimen at the proper angle to observe thin scales of dried bacterial exudate.
Suspect lesions on leaves can be examined for bacterial streaming to confirm the bacterial nature of the symptoms. Cut across the lesion perpendicular to the veins, mount the cut tissue in water between a glass slide and cover slip, and observe under low magnification (approximately 40 X). Fresh water-soaked lesions caused by X. translucens will ooze masses of bacterial cells that are easily seen streaming from the cut ends of the lesion. A cell suspension can be dilution plated to isolate the pathogen (see Diagnosis).
Older necrotic leaf lesions, and lesions on glumes and peduncles, will stream a few bacterial cells or none at all, but it is still possible to isolate the pathogen from these tissues by dilution plating. It is not possible to inspect seeds visually for the pathogen.
Similarities to Other Species/ConditionsTop of page
X. translucens pv. translucens is not likely to be confused with other pathogens of small grains, but may be confused with other Xanthomonas spp. (Schaad and Stall, 1988), or with various saprophytic bacteria commonly associated with plants or seeds. A pathogenicity test (see Diagnosis) should be performed on suspected colonies to differentiate the pathogen from saprophytes. Adriko et al. (2014) developed a set of genus-specific PCR primers for the detection and identification of xanthomonads. Hence, these primers could be used for the preliminary identification of yellow-pigmented bacterial strains isolated from cereals.
Leaf symptoms on wheat have been mistaken for symptoms of Septoria leaf blotch caused by Mycosphaerella graminicola and Phaeosphaeria nodorum. Leaf blotch caused by M. graminicola has linear lesions similar to bacterial streak, but usually there are abundant black pycnidia in the lesions. Leaf blotch caused by P. nodorum lacks conspicuous pycnidia, but the lesions tend to be lens-shaped with a chlorotic margin.
Black chaff symptoms on glumes have been confused with glume blotch caused by S. nodorum, and basal glume rot caused by Pseudomonas syringae pv. atrofaciens. Glume blotch generally causes an irregular grey-to-black lesion on the side of the glumes, and pycnidia generally can be seen through a hand lens. Basal glume rot causes irregular grey-brown discoloured areas at the base of the glumes. Brown melanosis or pseudo black chaff can be found on wheats with the Hope (Sr 2) resistance to stem rust when grown under high temperature and humidity (Waldron, 1929; Johnson and Hagborg, 1944). Wheat American striate mosaic virus also can cause brown melanosis on glumes of certain wheat cultivars (Seifers et al., 1995). Brown melanosis can be very similar to black chaff, and X. translucens should be isolated to differentiate the physiological disorder from black chaff.
Lesions on the stems and peduncles can be confused with lesions caused by M. graminicola or Fusarium spp., and it may be necessary to isolate the pathogen to be sure of the causal agent.
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.
Where bacterial streak does not yet occur, it may be possible to exclude the pathogen through diligent enforcement of quarantine measures for the import of seed. Where bacterial streak is well established, no one method is likely to provide complete control, and several methods appropriate to specific situations should be integrated into a programme designed to limit economic losses.
There are no seed treatments that eradicate X. translucens without excessive damage to the seed. Hot, acidified cupric acetate (Forster and Schaad, 1988; Duveiller, 1989), dry heat (Fourest et al., 1990) and Guzatine Plus (Mehta and Bassoi, 1993) have been shown to greatly reduce seedborne populations of X. translucens and bacterial streak in the field. However, acidified cupric acetate and dry heat are best suited to small seed lots, and none of the treatments are 100% effective for eradicating the pathogen or preventing transmission to plants (Duveiller, 1994a). Sands et al. (1986) reported a hot-water treatment at 53°C for 10 minutes followed by immediate cooling and drying.
Sun et al. (1988) in China reported control of black chaff with TF-128, and de Luz et al. (1993) in Brazil reported control of X. translucens with probenazole, but no other information is known about these compounds.
Producing seed in dry environments using furrow irrigation will reduce bacterial streak and the level of seedborne X. translucens (Forster and Schaad, 1988), but some contamination is likely unless the production field is planted with seed free of the pathogen and is well isolated from other sources of inoculum. Seed produced under such conditions would probably have a low risk of initiating an epidemic.
Considering the higher economic impact of the wheat and barley pathogen X. translucens pv. undulosa compared to the barley pathogen X. translucens pv. translucens, most breeding programmes have focused on the former pathogen. On the other hand, in some of the germplasm screening and evaluation projects, the identity of the pathogen (either used for inoculation under controlled environments or naturally occurred in the field plots) itself was confusing. For instance, Tubajika et al (1998) investigated the relationship between flag leaf symptoms caused by X. translucens pv. translucens and subsequent seed transmission in wheat. They isolated the X. translucens pv. translucens strains Xtt 90-1, 41 and 42 from diseased wheat plants in East Baton Rouge, Acadia and Concordia parishes, Louisiana, respectively, during the spring of 1989 and 1990. However, no discriminative assay was performed to determine the pathovar status of the strains. X. translucens pv. translucens strains should not have been pathogenic on wheat.
Furthermore, Kandel et al. (2012) screened spring wheat germplasm for resistance to bacterial leaf streak caused by X. campestris pv. translucens in the USA. Considering the host range of X. translucens pv. translucens (formerly known as X. campestris pv. translucens) which is restricted to barley, it could be hypothesized that unappropriated strains of the pathogen were used in the mentioned screening programme. It is recommended that the taxonomic status of the bacterial strains used in screening programmes is accurately determined to avoid subsequent confusion.
There have been a number of research studies reporting sources of resistance against X. translucens pv. translucens in barley. For instance, Alizadeh et al. (1994) reported partial resistance of some barley cultivars against X. translucens pv. translucens (formerly X. campestris pv. hordei) in Iran.
Khojasteh et al. (2019) investigated the genetic diversity of the worldwide population of X. translucens pathogenic on small grain cereals using the MLSA and MLST technique. It was shown that strains isolated in the Iranian Plateau, considered the centre of origin of the cultivated small grain cereals, were genetically more diverse than strains isolated in the New World, i.e., the USA. These findings raise questions about whether there is any source of resistance among the wild population of cereal species in the centre of origin of this crop, and emphasize the need for a more detailed investigation in this regard.
Source of resistance can be detected using inoculations in greenhouse or growth chamber tests (Milus and Mirlohi, 1994) but it is best to select for resistance in the field under high disease pressure (Milus et al., 1996). Pictorial disease assessment keys (Duveiller, 1994b) may be useful for rating disease severity in the field. Resistance is probably due to reduced ability of the pathogen to multiply in the host (El-Banoby and Rudolph, 1989; Milus and Mirlohi, 1994).
X. translucens may survive in volunteer crop plants, on crop debris or on alternative hosts (see Biology and Ecology), but these possible sources of inoculum are likely to be much less important than seedborne inoculum (Mehta et al., 1992). Therefore, crop rotation, tillage and weed control are not likely to have a large impact on disease severity, but these practices may be useful when attempting to produce seed with low levels of the pathogen.
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17/10/20 Updated by:
Ebrahim Osdaghi, Department of Plant Protection, University of Tehran, Karaj 31587-77871, Iran.
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