Open access peer-reviewed chapter

Bacterial Pathogens of Wheat: Symptoms, Distribution, Identification, and Taxonomy

Written By

James T. Tambong

Submitted: 06 January 2022 Reviewed: 25 January 2022 Published: 16 March 2022

DOI: 10.5772/intechopen.102855

From the Edited Volume

Wheat - Recent Advances

Edited by Mahmood-ur-Rahman Ansari

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Bacterial pathogens are significant biotic factors of wheat, a globally important source of carbohydrates. The diseases caused by these pathogens are reported to reduce annual wheat production by about 10% and up to 40% in severe infections occurring early in the growth period. This chapter presents current information on the symptoms, distribution, identification, and taxonomy of key bacterial pathogens of wheat with a focus on the seed-borne bacterium, Xanthomonas translucens pv. undulosa, the causative agent of the leaf streak and black chaff disease. Other wheat-pathogenic bacterial pathogens addressed in the chapter are Pseudomonas syringae pv. syringae, the causal agent of bacterial leaf blight; P. syringae pv. atrofaciens that cause the basal glume rot; Pseudomonas fuscovaginae, the causal agent of the bacterial brown sheath; Erwinia rhapontici, the causal agent of the pink seed of wheat; Pseudomonas cichorii, the causative agent of wheat stem melanosis; Clavibacter tessellarius is responsible for the bacterial mosaic of wheat as well as other minor bacterial pathogens. Finally, the chapter proposed the use of genome-based tools for the accurate identification and classification of bacterial pathogens of wheat.


  • wheat
  • bacterial pathogens
  • Xanthomonas translucens
  • Pseudomonas syringae
  • Clavibacter tessellarius
  • bacterial leaf streak and black chaff disease
  • Erwinia
  • Pseudomonas fuscovaginae
  • Rathayibacter
  • Pseudomonas cichorii

1. Introduction

Wheat is the most important food grain source in the human diet and is considered a global primary commodity ([1]; The “big three” cereal crops, wheat, maize, and rice account for 75% and 50%, respectively, of the carbohydrate and protein intakes by humans [2, 3, 4]. Of the three cereal crops, wheat is the most nutritious with a protein content of about 11–14% even though it is low in some essential amino acids, e.g. lysine [1, 5, 6]. Wheat, as an essential staple food, provides about 20% of the calories intake of about 40% of the global population [2].

Notwithstanding the significance of wheat grain and products to humans globally, the worldwide increment in wheat production is third behind maize and rice. Between 1961 and 2013, the total cultivated land area allotted to bread and durum wheat increased from 204 Mha to 218 Mha, an increment of only 6.8%, but recorded a 321% increase in production from 222 MT to 713 MT [4]. In 2017, global wheat production was 757 MT harvested from 220 M ha [4]. An increase of 30% in wheat production to 1 billion tons, as suggested by Bockus et al. [2] is required by 2030 to feed the nine billion estimated population. It is unclear how this can be achieved in the agricultural world threatened by global warming. This is compounded by the fact that annually, about 25–30% of wheat crop yield loss is incurred due to abiotic and biotic stresses.

Biotic stresses are incited by diseases and insect pests of wheat. One of these groups is bacteria. Bacteria are prokaryotic microorganisms that have a nucleus or organelles not bound to a membrane. Plant pathogenic bacteria are unicellular and may be motile due to the presence of one or more flagella; and the mode of reproduction is by binary fission, in which the cell divides into two similar daughter cells with new generations occurring in less than 2 h in some species. Due to their small size, measuring up to about 2 μm, light (at least 400×) and electron microscopes are required for detailed studies and investigations of the cell morphology and structure. Bacteria that are pathogenic to wheat are categorized into two main groups: Gram-positive and Gram-negative based on the reactions to Gram’s stain, a reaction during which strains of the former group stains dark purple while the latter group appears red by taking up the counterstain.

Bacterial pathogens are significant biotic factors of wheat, a globally important source of carbohydrates. The diseases caused by these pathogens are reported to reduce annual wheat production by about 10% and up to 40% in severe infections occurring early in the growth period. This chapter presents current information on the typical symptoms, distribution, identification, and taxonomy of key bacterial pathogens of wheat. The pathogens to be profiled include Xanthomonas translucens, the causal agent of the bacterial leaf streak and black chaff disease; the causal agent of bacterial leaf blight, Pseudomonas syringae pv. syringae; Clavibacter michiganensis subsp. tessellarius (bacterial mosaic of wheat) as well as other minor bacterial pathogens.


2. Wheat bacterial pathogens

2.1 Xanthomonas translucens pv. undulosa (Xtu), the causal agent of bacterial streak and black chaff disease of wheat

2.1.1 Disease symptoms, importance, and distribution

The leaf streak and black chaff (BLS) is the most important bacterial disease of wheat. Symptoms appear on the leaves and/or spikes of wheat plants. The disease starts with water-soaked necrotic streaks that eventually change to translucent lesions (Figure 1). Under favorable temperature and humidity, yellow exudates of the bacteria can be seen oozing out on the surfaces of infected leaves. The black chaff phase of the disease is seed-borne and as such may pose restrictions for germplasm exchange internationally and the trade of wheat grain [7]. In addition, losses in yield resulted mainly from the poor filling of grains and can be as high as 40% [8] but if the heads are heavily diseased no marketable grain is yielded [2]. Black chaff is often confused with the pseudo-chaff or brown melanosis conditions caused by abiotic stress [9, 10] as well as fungal pathogens and genetic factors [7, 11]. In fact, the black chaff was reported to be a composite disease involving three major factors: the bacterial black chaff, alternaria blotch and favorable environmental conditions for melanism [10, 12].

Figure 1.

Symptoms associated with bacterial leaf streak disease of wheat caused by Xanthomonas translucens pv. undulosa (a) typical longitudinal necrotic lesions; (b) completely diseased or dead flag leaves (center of the photo); and (c) typical black chaff symptom of the spikes. Photographs (a) and (b) are courtesy of Dr. M. Harding, Alberta Agriculture and Forestry; and (c) was kindly provided by Dr. S. Wegulo, University of Nebraska-Lincoln.

BLS disease has been confirmed in 34 countries of wheat-growing regions worldwide. Based on a recent global database of the European Plant Protection Organization (EPPO;, accessed 22 November 2021), the disease is present in Africa (Ethiopia, Kenya, Madagascar, Morocco, South Africa, Tanzania, Tunisia, and Zambia), Australia, South America (Bolivia, Argentina, Brazil, Peru, Paraguay, and Uruguay), North America (Mexico, Canada, and United States,), several European countries (Russia, Romania, Ukraine, Turkey), and Asia (China, India, Iran, Israel, Japan, Kazakhstan, Malaysia, Azerbaijan Pakistan, Syria, Georgia, Yemen). The western part of Europe appears to be shielded from the BLS disease which might be due to unfavorable environmental factors coupled with aggressive quarantine efforts [13, 14]. In North America, the first reports of BLS were in barley and wheat farms in the Midwestern United States [15]. Figure 2 shows the global distribution of X. translucens pv. translucens (Xtt). It is worthwhile to note that some plant pathologists had referred to the BLS disease causal agent of wheat as X. t. pv. translucens (see the section on taxonomy below).

Figure 2.

Global distribution of Xanthomonas translucens pv. translucens. CABI, 2020. X. translucens pv. translucens (bacterial leaf streak of barley). In: Crop Protection Compendium. Wallingford, UK: CAB International. [accessed: 17/12/2021]. Note that wheat BLS pathogen has also been referred as X. t. pv. translucens [2].

The disease spread to many other counties with outbreaks and epidemics occurring in southeastern regions of United States [16]. Between 2018 and 2012, the BLS disease incidence increased significantly in the Upper Midwestern states of Minnesota [17], South Dakota [18], and North Dakota [8]. The earliest available record of BLS in Canada was in 1934 [10]. In recent years, no outbreaks have been reported in Canada but the frequency of annual occurrences is becoming high, especially in Alberta and Saskatchewan provinces of Canada. In Mexico, the earliest record of BLS was reported in 1931 in the northeastern part of the country [19]; and currently, high elevation temperate and humid areas of Toluca, 2650m above sea level, seem to have high disease pressure [20].

Although the BLS is one of the most important bacterial diseases of wheat and is considered a potential biothreat, recent data or reports on yield losses do not exist. This is compounded by the fact that the importance of the disease may significantly vary in the different wheat-growing areas which might partly be dependent on the tolerant/resistant levels of the cultivars grown as well as prevalent environmental conditions [14]. Wheat yield grain losses attributed to BLS disease are largely about 10% or lower, but yield losses on highly susceptible cultivars can be as high as 40% [21, 22]. A 20% yield loss as a function of leaf streak severity on bread wheat was reported in Mexico [23]. Duveiller and Maraite [23] and Shane et al. [24] reported that the BLS severity on the flag leaves correlates negatively with the BLS yield loss, and suggested that BLS infection of 50% leaf surface area of flag leaves may lead to a yield reduction of up to 20%. Shane et al. [24] reported that a BLS severity of 100% on flag leaves led to up to 34% yield wheat grain loss. Yield loss due to the severity of BLS disease can be attributed to the reduction in seed weight and the number of seeds per spike [25]. Also, there is a reduction in the quality as BLS infection seems to change the protein content of wheat grains [24].

2.1.2 Taxonomy

The taxonomy of the “translucens” group is not clear due to the pathovar naming system that is based on the first host from which the bacteria was isolated. This section provides some historic and current nomenclature of the group. The bacterial pathogen of the BLS disease on wheat was originally named Bacterium translucens pv. undulosa [26]. Since then, the nomenclature of the pathogen has evolved through a series of names including Xanthomonas campestris pv. translucens and X. campestris pv. undulosa [27]. The pathogen has also been referred in many publications as X. translucens pv. translucens [2], the valid name for the pathovar that infects barley. The most recent and valid nomenclature for this pathogen is X. translucens pv. undulosa (Smith et al.) Vauterin, Hoste, Kersters and Swings 1995. Xtu is reported to infect both wheat and barley while Xtt is pathogenic to only barley [8, 28]. The complete current taxonomy of Xtu is Kingdom Bacteria; Phylum Proteobacteria; Class Gammaproteobacteria; Order Xanthomonadales; Family Xanthomonadaceae; Genus Xanthomonas; and Species X. translucens. The translucens group is divided into 10 pathovars: undulosa, translucens, cerealis, hordei, secalis, arrhenatheri, graminis, poae, phlei and pistaciae. Pathovar cerealis (Xtc) has also been reported to cause BLS disease symptoms [29, 30], but Xtc has a broad host range than Xtu that includes wheat (Triticum spp.), barley (Hordeum vulgare), oats (Avena spp.), triticale (Triticosecale), and rye (Secale cereale) [8, 31, 32].

2.1.3 Isolation and identification

The type and quality of the collected BLS diseased leaf sample and culture media are crucial to isolating Xtu strains and could, also, depend on the sample type, leaf, or seeds. Generally, a leaf sample with young BLS lesions is selected, thoroughly washed in running water, surface disinfected, and blotted dry. About 0.25 cm2 sections of symptomatic leaf tissue are aseptically excised, bisecting, at least, one lesion and immersed in a 100 μl droplet of sterile water, and incubated at room temperature for 5 min [28]. Using a 10-μl loop, the suspensions are streaked onto appropriate non-selective culture media e.g. nutrient agar (NA), Wilbrink’s medium, King’s B, and Yeast peptone glucose agar plates, and incubated at 28–30°C for up to 72 h. Semi-selective culture media, KM-1 [33], XTS [34] and Wilbrink’s boric acid-cephalexin (WBC) [35] have also been developed. KM-1 and XTS have been proposed for isolating Xtt while WBC is a useful medium for isolation of X. t. pv. undulosa as well as related pathovars. The composition of these culture media is given in Table 1.

KM-1 [33]XTS agar [34]Wilbrink’s boric acid-cephalexin (WBC) [35]
Lactose10.0Glucose5.0Bacto peptone5.0
D (+) trehalose4.0Nutrient agar (Difco)23.0Sucrose10.0
Thiobarbituric acid0.2Distilled H2O (ml)978.0K2HPO40.50
K2HPO40.8MgSO4 .7H2O0.25
KH2PO4. 7H2O0.8Na2SO3 (anhydrous)0.05
Yeast extract1.0Agar15.0
NH4Cl1.0Distilled water (ml)850
Bacto agar (Difco)15.0
Distilled water1.0 L
Dissolve the ingredients completely and adjust pH to 6.6 using 1 N NaOH before adding agar. Autoclave at 121°C for 20 min., 15 psi; and cool to to 50°C prior to adding:
  • Cycloheximide (dissolved in 95% ethanol) 100.0 mg

  • Ampicillin (dissolved in 50% ethanol) 1.0 mg

  • Tobramycin (dissolved in 50% ethanol) 8.0 mg

  • Useful for isolation of X. t. pv. Translucens

Autoclave at 121°C for 20 min, 15 psi and cool to 45°C. Then add:
  • Cycloheximide (20 ml of a 100 mg/ml 200.0 mg stock solution in 75% ethanol)

  • Cephalexin (1 ml of a 10 mg/ml 10.0 mg stock solution in 75% ethanol)

  • Gentamycin (0.8 ml of a 10 mg/ml 8.0 mg stock solution in 75% ethanol)

  • developed for X. t. pv. translucens.

Autoclave at 121°C for 20 min. 15 psi and mix with the following solution (autoclaved separately):
  • Boric acid 0.75 g in 150.00 ml DH2O

Allow to cool to 45°C, and then add:
  • Cycloheximide (in 2 ml of 75% ethanol) 75.00 mg

  • Cephalexin (1 ml of a 10 mg/ml stock 10.00 mg solution in 75% ethanol)

  • Developed for isolation of X. t. pv. undulosa and similar pathovars.

Table 1.

Semi-selective media for isolation Xanthomonas translucens pv. undulosa and related pathovars.

Xtu can be identified and differentiated from Xtt and X. translucens pv. cerealis (Xtc) by pathogenicity tests. Application of the bacterial inoculum in the soil and/or seeds is not effective in testing pathogenicity [7, 10]. Artificial inoculation of wheat plants at the 4–5 leaf stage through the injection of bacterial suspension using a hypodermic syringe is the most reliable and effective method [19, 36]. As indicated above, Xtu is reported to infect wheat and barley, Xtt is pathogenic to barley only while Xtc has a broad host range that includes several other cereals. Well-conducted pathogenicity tests could be useful in the identification and differentiation of Xtu.

The pathogen can be distinguished from other pathogenic bacteria of wheat by biochemical and physiological trains. The bacterium causes hypersensitivity on tobacco [37]. Strains of Xtu are non-sporulating, rod-shaped Gram-negative bacteria and use a polar flagellum for motility. This bacterium is oxidative (producing acid from glucose under aerobic conditions) [7]. It does not reduce nitrate to nitrite [38] while esculin hydrolysis is positive but no 2-ketogluconate is produced [39, 40]. Strains of X. translucens, unlike X. campestris, do not use lactose nor hydrolyze starch [41]. Very few biochemical and physiological tests exist for pathovar differentiation. Metabolic fingerprinting based on BIOLOG MicroPlatesTM on carbohydrates and amino acid utilization may not be effective in the identification of pathovars [7].

Molecular techniques are useful for identification of Xtu but very few records are published. Repetitive element palindromic PCR (rep-PCR) and amplified fragment length polymorphism (AFLP) have been used to partly taxonomically differentiate some X. translucens pathovars [32, 42]. Maes et al. [43] reported the development of assay based on rDNA spacer sequences for detection X. translucens pathovars causing cereal leaf streak in the seed Iqbal et al. [44] developed a 2-step conventional PCR assay specific for detection and identification of Xtu in wheat in Pakistan. Recently, genome-based and multilocus molecular typing and identification of X. translucens pathovars have been reported [17, 28, 45, 46, 47, 48]. Langlois et al. [49] characterized the X. translucens complex by sequencing and analyzing 16 draft genomes and exploited the genetic variability for the development of diagnostic loop-mediated isothermal amplification (LAMP) assays that can distinguish pathovars (pvs. undulosa, translucens, hordei, and secalis) that cause disease on cereals from noncereal pathovars (arrhenatheri, cerealis, graminis, phlei, and poae). Genome sequencing and comparative genomics have become more reliable and effective tools in the identification of plant pathogenic bacteria including members of the X. translucens group (see Section 3).

2.1.4 Management strategies

Since the BLS pathogen is seed-borne, one of the best control strategies is to avoid planting infected seeds. It is, thus, important to do a simple seed wash test [7]. The test is conducted by adding 120 g of seeds in 120 ml aqueous saline solution containing 0.02% v/v Tween 20, shake vigorously for 1 minute, and perform serial tenfold dilutions up to 10−3 before plating on semi-selective media e.g. KM-1 and WBC, described above. Levels of 1000 colony forming units per gram or less in seed washes have been reported to result in little or no disease [34]. Avoid using infested seeds for germplasm exchange [7] to minimize the spread of the pathogen.

Crop rotations have been reported as another management strategy but little data exist on how this measure works in reducing the BLS and black chaff epidemics. Given that the main source of spread of the pathogen is through infested seeds, crop rotation may not be a key management strategy. However, the bacterium can survive in wheat straw and can induce initial field infections. Boosalis [50] noted that the viable number of the pathogen is low in overwintered straw and greatly reduced when the straw are buried in the soil. Also, pathogen survival seems to be almost impossible under non-host rotations as well as the high susceptibility of the bacteria to antagonistic bacteria [34].

2.2 Pseudomonas syringae pathovars

2.2.1 Pseudomonas syringae pv. syringae, the causal agent pathogen of the leaf blight Disease symptoms, distribution, and importance

P. syringae pv. syringae is the pathogen that causes the bacterial leaf blight [51] or leaf necrosis [52] disease of wheat. During the initial infection, the symptoms are small, water-soaked spots that expand into characteristic large lesions, blotches, or streaks often observed under rainy and high humidity conditions [2]. Under low humidity, the water-soaked lesions quickly turn into characteristic gray-green. Within 3 or 4 days, the lesions coalesce to form irregularly shaped blotches and become tanned or white. Whole leaves might be necrotic while the heads and lower leaves are without any symptoms [2].

The disease was first reported in the USA in the following states, South Dakota, North Dakota, and Nebraska [53], Minnesota [51] and Montana [54]; and 2 Canadian provinces of Saskatchewan and Alberta [52]; Italy [55], South Africa [56]; Pakistan [57] and Argentina [58]. Besides wheat, Pseudomonas syrinage pv. syringae is a widespread plant epiphytic bacterium with a very wide host range consisting of many herbaceous and woody plants including corn (Holcus spot), sorghum (bacterial leaf blight), and lilac (bacterial blight). Otta [52] found that strains of P. syringae pv. syringae isolated from corn, foxtail, sorghum, and peach are pathogenic on wheat seedlings. This opportunistic bacterium has been reported to cause disease in various host plants of agricultural importance as well as many weeds [59, 60] in Europe, Asia, Oceania, South America, Caribbean, and North and Central America. Its worldwide distribution is well documented as the causal agent of the bacterial canker or blast disease of stone and pome fruits (Figure 3).

Figure 3.

Distribution of Pseudomonas syringae pv. syringae Van Hall worldwide. CABI, 2020. P. syringae pv. syringae (bacterial canker or blast (stone and pome fruits)). In: Crop Protection Compendium. Wallingford, UK: CAB International. [accessed: 17/12/2021] [61]).

The importance of the disease on wheat is dependent on specific weather conditions. The disease is more often observed during several days of high humidity, cool temperatures (15–25°C), and heavy rainfall [62]; and plants are more susceptible at the boot stage. The disease is generally considered to be of minor importance with minimal impact on yield losses. However, Otta [52] reported high yield losses in South Dakota where an epidemic outbreak resulted in very high infection severity in fields with 75% or more necrotic leaves. Also, in North America, the cultivation of susceptible cultivars, e.g. Chris, Era, Scout 66, and Winoka, led to significant yield losses. These susceptible cultivars have been replaced with resistant wheat cultivars and as such the disease is only sporadic in North America [2].

The current and complete taxonomic standing of the bacterium is Kingdom Bacteria; Phylum Proteobacteria; Class Gammaproteobacteria; Order Pseudomonadales; Family Pseudomonadaceae; Genus, Pseudomonas; and Species, P. syringae; pathovar syringae.

There are no traditional management strategies for the bacterial leaf blight disease caused by P. syringae pv. syringae. It is, however, recommended to avoid planting very susceptible cultivars.

2.2.2 Pseudomonas syringae pv. aatrofaciens, the incitant of the basal glume rot disease Disease symptoms, distribution, and importance

P. syringae pv. atrofaciens (McCulloch) Young, Dye, & Wilkie is the incitant of basal glume rot of wheat. Typical and characteristic symptoms of the disease are dull brownish to blackish discoloration, usually, on the lower part of the wheat glume (Figure 4a). It is more visible in the inner than the outer side of the glume. At times, water-soaked sections can be seen around the lesions [63]. Seeds in florets showing typical symptoms of the disease are often brownish to blackish in color (Figure 4b). The grains may be shriveled when wet darkish green lesions appear on the peduncle and surrounding it fully [2]. Also, small (2–10 mm) wet, darkish green lesions may appear on the leaves which then could quickly become necrotic [62]. Symptoms on the glume and peduncles can be confused with black-chaff caused by X. translucensor pseudo-chaff, a disorder associated with abiotic stress factors. Seed symptoms are often confused with the black point disease. Wet climatic conditions after heading favor the development of the disease.

Figure 4.

(a) Typical dark brown to black discoloration and (b) diseased seed symptoms caused by Pseudomonas syringae pv. atrofaciens; and (c) global distribution of the causal agent of the wheat basal glume rot disease. Photos: Centro Internacional de Mejoramiento de Maíz y Trigo (CIMMYT) ( Distribution map from CABI, 2020. In: Crop Protection Compendium. Wallingford, UK: CAB International. [accessed: 17/12/2021].

The basal glume rot disease caused by P. s. pv. atrofaciens was first described in America [64]. It has since been reported in almost all wheat-growing regions of the world [65]. The basal glume rot disease of wheat has been reported in the USA and Canada [64]; Mexico [66]; Ukraine [67]; Bulgaria [68]; Australia [69]; New Zealand [70]; South Africa [56, 71]; Germany and Denmark [72]; and Belgium [62]. Figure 4c shows the global distribution of the disease.

The basal glume rot occurs only sporadically. It is generally thought to be a minor disease and as such, the impact on wheat grain yield has not been well studied. However, P. syringae pv. atrofaciens is reported to cause yield losses exceeding 50% in marshy soils of Germany [73]. Also, 7-year outbreaks of leaf necrosis led to leaf infections of over 75% [52] and can cause a severe reduction in grain quality [74, 75].

Since the bacterium is seed-borne, contaminated seeds remain the most important source of infections to new wheat plantings. No effective seed treatment exists for the management of P. s. pv. atrofaciens. Also, wheat genotypes may vary in resistance but little is known of the level of resistance of present-day cultivars [2]. Wheat growers are recommended to avoid planting seeds harvested from infected fields.

2.2.3 Isolation and differentiation of P. syringae pv. syringae and P. syringae pv. atrofaciens

Epiphytic populations of P. syringae pv. atrofaciens and P. syringae pv. syringae are extensive and easily isolated from wheat plants. As such, isolating these bacteria is not definite prove that the observed symptoms are induced by P. syringae pv. atrofaciens or P. syringae pv. syringae [62]. However, if the concentrations of the bacteria in diseased leaf tissues are high, e.g. 108 colony forming units per gram fresh weight, then they might play somewhat a likely causal role [62].

These two pathovars cannot be differentiated based on colony morphology or physiological traits [53, 62, 76], nor by using serological methods [52, 77] while Iacobellis et al. [76] reported that P. syringae pv. syringae and P. syringae pv. atrofaciens could not be differentiated by genetic features. However, advances in whole-genome sequencing and comparative genomics analysis have made it possible to accurately identify and differentiate these two pathovars (see Section 3). Genome-based DNA-DNA relatedness calculations between P. syringae pv. syringae and P. syringae pv. atrofaciens revealed 82.2% hybridization values, suggesting a genetic difference of about 18% (Tambong, unpublished). Accurate and reliable pathovar identification/differentiation is only possible based on the disease symptoms induced on cereals [78, 79].

The isolation of P. syringae pv. syringae and P. syringae pv. atrofaciens has been routinely achieved using King’s medium B. Colonies are whitish-gray in color and are circular and convex in shape after 24 h of incubation; and exhibit blue fluorescence under ultra-violet light, which turns green after 48 h [62]. No semi-selective media have been reported for use in the isolation of these P. syringae pathovars. However, KBC medium, King’s B medium amended with boric acid, cephalexin, and cycloheximide [80] showed high selectivity for P. syringae pv. atrofaciens even though the bacterium seemed to grow slightly slower compared to King’s medium B.

2.3 Pathogens of the genus “Clavibacter

2.3.1 Clavibacter tessellarius, the causative agent of bacterial mosaic of wheat

The wheat bacterial mosaic disease was first reported in Nebraska (USA) in 1976 and taxonomically described in 1982 [81]. The disease is caused by Clavibacter michiganensis subsp. tessellarius (Carlson & Vidaver) Davis et al.. The taxonomy has evolved from Corynebacterium michiganese subsp. tessellarius through C. michiganensis subsp. tessellarius to Clavibacter tessallarius based on whole-genome sequence analysis [82]. The geographic distribution is restricted to North America: USA [81] and Canada [13]. The pathogen is specific to wheat but seems to be related to C. michiganensis [81]. It can, however, be differentiated from other plants pathogenic corynebacterium by bacteriocin typing. The disease is sporadic, annually, and also occurs in triticales. The economic significance of the disease is yet to be documented.

The bacterial mosaic disease caused by C. tessellarius is a foliar disease characterized by a mosaic of small yellow lesions that resemble infections by viral pathogens. The small lesions on the leaf may coalesce to form streaks [7]. Under greenhouse temperature conditions of 20–22°C, artificially inoculated seedlings develop typical mosaic-like symptoms in three to 5 days [81]. The pathogen is seed-borne [83] and as such control strategies should include removing contaminated seeds and the development of tolerant/resistant genotypes. The available wheat genotypes seem to respond differentially to the pathogen. This could be an indication that genetic improvement is possible [7].

2.3.2 Clavibacter iranicus, the pathogen of gumming disease of wheat spikes

C. iranicus is the causative agent of the gumming disease of wheat spikes and has only been reported in Iran [84]. The taxonomic name of the pathogen was initially Corynebacterium iranicum [84] followed by C. iranicus (ex Scharif) Davis, Gillaspie, Vidaver & Harris 1984 while Zgurskaya et al. [85] proposed the renaming of this bacterial pathogen as Rathayibacter iranicus. The economic significance of the pathogen to wheat is unknown.

There are no control or management strategies proposed in the literature for the gumming disease of wheat spikes.

C. iranicus (Syn. R. iranicus) exhibits a close relatedness with Rathayibacter tritici but they are different [2].

2.3.3 R. tritici, causative agent of spike blight of wheat

R. tritici is the causal agent of spike blight of wheat and was first reported in India in 1917 as the causal agent of the tundu disease [2]. It is also known as the yellow ear rot or yellow slime rot disease; and also pathogenic to several grasses, e.g. barley. The spike blight is considered a disease complex involving the bacterium R. tritici and a nematode, Aguina tritici, the causal agent of the seed galls, also known as ear cockle, in some wheat cultivars [86].

The current taxonomic nomenclature of the pathogen is R. tritici (Carlson & Vidaver) Zgurskaya, Evtushenko, Akimov & Kalakoutskii. Previous scientific names, from latest, included Clavibacter tritici (Carlson & Vidaver) Davis, Gillaspie, Vidaver & Harris, Corynebacterium michiganense pv. tritici (Hutchinson) Dye & Kemp Corynebacterium tritici (Hutchinson) Burkholder, Phytomonas tritici (Hutchinson) Bergey and Pseudomonas tritici Hutchinson.

The geographic distribution includes 14 countries: Egypt, Ethiopia, Morocco, Zambia, Afghanistan, China, India, Iran, Iraq, Pakistan, Cyprus, Australia [60, 86, 87, 88, 89].

Initial field symptoms of the spike blight disease include parallel white or yellow streaks generally along the veins of leaves. Later, this is transformed into a sticky mass, yellow gummosis on wheat spikes. The spikes and peduncles (necks) are often distorted when they emerge from the whorl. Also, early leaves may also be twisted or wrinkled. When the sticky mass is dry, the gummois becomes pale yellow-colored flecks on the spikes and the adaxial leaf surface [2]. Since the bacterial sticky mass is watery under wet weather conditions and dry when the RH is low, the hardened gummy substance mechanically causes the leaves, spikes, and peduncles to be distorted. The symptoms that are caused by A. tritici (nematode) are part of the spike blight disease complex.

R. tritici is reported to persist in crop residues in moist soils. To facilitate the colonization of wheat, the pathogen has to be carried by the nematode A. tritici into the whorl enclosures. Generally, the juveniles of A. tritici are contaminated by the cells of R. tritici in the soil. This helps with the dissemination of the bacterial cells of R. tritici on seeds, in seed galls, and in soil. The A. tritici and R. tritici can survive in seed galls for over 5 years [90].

The management of spike blight disease is not well studied. However, growing wheat on well-drained soils significantly reduces spike blight disease. Also, management strategies used to control A. tritici would be helpful in reducing the spike blight incidence. A 2- or 3-year crop rotation of wheat with non-grass crops is another control method.

2.4 Other bacterial pathogens

This section focuses on other bacterial causal agents of wheat diseases that are rarely observed and are pathogenic to a variety of host plants. These are considered to be less specialized pathogens some of which are epiphytes or opportunistic organisms that cause disease to the wheat only under favorable unusual conditions [91]. Since the diseases caused by these bacteria have not been extensively studied very limited information exists. The pathogens described here include Pseudomonas fuscovaginae, Pseudomonas cichorii, and Erwinia rhapontici.

2.4.1 Pseudomonas fuscovaginae, causal agent of the bacterial brown sheath

P. fuscovaginae (ex Tanii et al.; Miyajima et al.) is the causal agent of the bacterial sheath rot, also referred to as brown sheath rot, of wheat. Tanii et al. [92] were the first to report this pathogen on rice but have since been isolated from several other cereal crops including wheat [93]. A recent genome-based taxonomic study re-classified P. fuscovaginae as a later heterotypic synonym of Pseudomonas asplenii [94].

The bacterial brown sheath rot disease caused by P. fuscovaginae (heterotypic synonym of P. asplenii) after the first report in Japan in 1976 on rice, it has been reported in 33 other countries, e.g. Burundi, Mexico, (Figure 5) on a variety of crops and grasses including wheat, barley, maize, oats, bentgrasses, bromegrass, perennial ryegrass, smooth meadow-grass, rye, and sorghum (CABI, accessed November 17, 2021). Little information exists on the effects of the pathogen on the yield of wheat. But severity seems to vary with the genetics of the host plants. Two wheat cultivars, Anahuac and Seri 82 suffered severe damage in Mexico in 1990 with 18–20% infections of the tillers [2, 93]. In Nepal, the disease was not observed on genotype RR21 while four other cultivars (WK685, Annapurna-1, Annapurna-2, and Annapurna-3) were heavily infested [95].

Figure 5.

Global distribution of Pseudomonas fuscovaginae. CABI, 2021. P. fuscovaginae (sheath brown rot). In: Crop Protection Compendium. Wallingford, UK: CAB International. [accessed: 17/12/2021].

The symptoms are characterized by black brown lesions of angular to irregular shapes on the leaves. The lesions have blackish-purple water-soaked discolored borders [96]. The adaxial surface of the leaf sheath is often where the initial infection starts [97]. Plants with severe incidences may show poor spike emergence and even sterility. P. fuscovaginae is disseminated by seed and plant susceptibility is dependent on the developmental stage. If the pathogen is present, infections are highly favorable during the flowering stage at a temperature range of 17–25°C with 100% relative humidity (RH) [2].

Management of the disease is very difficult since some of the factors involved cannot be controlled easily. Given that the pathogen is seed-borne [93], preventive measures are key to reducing the incidence of the disease. Avoid sowing contaminated seeds or susceptible cultivars, especially in growing areas under low temperatures and high RH microclimates.

2.4.2 Pseudomonas cichorii, the causative agent of wheat stem melanosis

The Gram-negative bacterium P. cichorii is a ubiquitous organism that is pathogenic to several host plants worldwide [60]. The only report of P. cichorii on wheat is that of Piening et al. [98] isolated from the Park spring wheat cultivar grown in copper-deficient soil in Canada (Alberta). No recent report of its occurrence on wheat exists and data on the economic importance of the disease is not available.

The symptoms were first observed in 1965 in central Alberta as irregularly shaped, sharply defined, dark patches in fields of cv. Park spring wheat at the milky stage of growth [98]. The initial symptoms start at the milky stage with the development of small lesions of light brown coloration under the lower two nodes which later darken and coalesce on the stem, rachis, and peduncle [91]. The spikes (also called the ear or head) are bleached and the grains are shriveled [2]. Even though the epidemiology is not understood, high temperatures (29°C) and relative humidity are conducive for the spread of the pathogen; and the disease has, also, been associated with soils that are cooper-deficient [99, 100]. There is no known management strategy for stem melanosis. However, the application of Cu chelate as amendments at the rate of 2.4 kg/ha to copper-deficient soils has been reported to reduce the severity of stem melanosis and improve wheat grain yield [99].

The current taxonomy of the pathogen is Kingdom, Bacteria; Phylum, Proteobacteria; Class, Gammaproteobacteria; Order, Pseudomonadales; Family, Pseudomonadaceae; Genus, Pseudomonas; and Species, P. cichorii.

2.4.3 Erwinia rhapontici, the causal agent of the pink seed of wheat

E. rhapontici (Millard) Burkholder emend. Hauben et al., a homotypic synonym to Pectobacterium rhapontici (Millard) Patel & Kulkarni, Erwinia carotovora var. rhapontici (Millard) or Pseudobacterium rhapontici (Millard) Krasil’nikov, is the causal agent of the pink seed of wheat. Heterotypic synonymic names include Bacterium rhaponticum, Phytomonas rhapontica, Aplanobacter rhaponticum, and Xanthomonas rhapontica.

It has been reported in Canada [101], France [102], England [103], USA [104], Belgium [105] and Russia and Ukraine [106]. It is also reported to be pathogenic to pea [107], onion [108], garlic [109], common bean [110], lentil [111], rhubarb [7] and several other plant species.

This pathogenic bacterium is opportunistic as it affects mainly injured kernels caused by the gall midge (family Cecidomyiidae). Seeds/kernels infected by E. rhapontici exhibit pinkish discoloration and slightly shriveled compared normal kernels. The germination vigor of infected seeds is poor. The market value of pink seeds is significantly low and often not used in pasta production [104].

The infrequent and somewhat random occurrence of pink seed on wheat makes it difficult to understand the disease and devise effective prevention strategies.


3. Genome-based identification of key bacterial pathogens of wheat

With advances in whole-genome sequencing (wgs) and bioinformatics tool developments, genome-based methods of classification and identification of prokaryotes including wheat pathogens are fast replacing traditional methods such as colony morphology, conventional polymerase chain reaction-based assays, multilocus sequence analysis, and wet-lab DNA-DNA hybridization similarity values. Some of the traditional approaches, e.g., wet-lab DDH have inherent drawbacks such as irreproducibility between laboratories [112]. Whole-genome sequencing provides complete and draft chromosome data that can be used to better understand the evolutionary and taxonomic relationships in bacteria, in general [113, 114]. Genome sequencing and comparative genomics are powerful technologies in the accurate identification and classification of bacterial pathogens of wheat.

Table 2 shows the number of publicly available whole-genome sequences (wgs) of each of the bacterial pathogens of wheat profiled in this chapter. Application of taxogenomics approach to identifying bacterial pathogens of wheat can be achieved by computing genome-to-genome distance (GGDC; [115]); average nucleotide identity (ANI; [116]), MuMmer-based average nucleotide identity (ANIm; [117]; tetranucleotide usage patterns (TETRA; [118]; and codon usage [119] as well as supertree analysis and other genomic signatures [116]. The three most commonly used genome-based parameters in bacterial identification are GGDC, ANI, and TETRA. GGDC outputs a pairwise genome-based digital DNA-DNA hybridization (dDDH) value between two bacterial strains with a species-level cut-off value of 70%; the ANI similarity value based on the BLAST or MuMmer approach has a species delineation threshold of 95–96%; and TETRA cut-off value of 0.998 for same species assignment of two bacterial strains. It is key that the whole genome sequence of the unknown strain is compared to the type strain or pathotype of the target bacterial species.

PathogenWheat diseaseNumber of sequenced genomesGenome size (Mb)G + C content (%)Number of Protein-coding sequences (CDS)
X. t. pv. undulosaLeaf streak and spike black chaff12 (8)4.667.933406
P. s. pv. syringaeLeaf blight76 (8)5.9959.084652
P. s. pv. atrofaciensBasal glume rot7 (1)5.9159.15109
P. fuscovaginaeBrown sheath9 (1)6.5461.885693
P. cichoriiStem melanosis51 (3)5.8958.284990
Erwinia rhaponticiPink seed8 (5)5.3154.044648
C. tessellariusBacterial mosaic2 (0)3.3173.453028
R. iranicusGumming6 (1)3.3367.183066
R. triticiSpike blight6 (1)3.2369.782982

Table 2.

Number of publicly available whole-genome sequences (wgs) of each of the bacterial pathogens of wheat. X. t., Xanthomonas translucens; P. s., Pseudomonas syringae; C. Clavibacter; and R., Rathayibacter. wgs presented are from wheat as well as other hosts, and the number in brackets indicate genomes having complete chromosomes instead of draft genomes.

The procedure can be summarized into 5 crucial steps: (1) bacteria isolation from plant tissues. Single colony purification, by repetitive streaking, is important to ensure that only one bacterial species is present. The presence of two or more species will result in poor genome assembly; (2) genomic DNA extraction and quantification. High purity genomic DNA is required as well as the quantity which is dependent on the genome sequencing method to be used; (3) library preparation. During this step, DNA is fragmented into short-reads, end-repaired, and adapter-ligated. Adapters allow the attachment of sequences to the flow cell, to identify samples, and to permit multiplexing; (4) Next-generation genome sequencing (NGS). Also referred to as high-throughput sequencing, enables sequence profiling of genomes. In a relatively short time, NGS generates large amounts of short or long sequence data; (5) Genome assembly and analysis. This step involves the use of a specialized Bioinformatics tool to assemble the raw reads into contigs, scaffolds, or chromosomes. These tools can be user-friendly or command-line accessed. The assembled whole-genome sequences (contigs, scaffolds, or complete chromosomes) can then be utilized as input in other bioinformatics tools to compute dDDH, ANI, or ANI similarity values between the type strain of the target bacterial species and the newly isolated or unknown bacterial strains. There are several user-friendly bioinformatics pipelines that can be routinely used by biologists, plant pathologists, or other scientists with basic or little bioinformatics knowledge. Examples of these pipelines are PATRIC ([120]; and galaxy@pasteur [121];


4. Conclusion

The purpose of this chapter was to summarize the disease symptoms, distribution, taxonomy, and identification methods of key bacterial pathogens of wheat. Occurrences of wheat diseases caused by bacteria are sporadic or confined to limited ecological niches due to favorable weather conditions. However, with the advancement of climatic change some of these diseases might become more prevalent and as such the information provided here should be helpful to agriculturists, biologists, plant pathologists as well as wheat breeders with limited experience to recognize and identify these pathogens. This is particularly important since some of these pathogens, e. g., X. translucens pv. undulosa, are seed-borne and efforts are being made to minimize spread through germplasm exchange. It is highly recommended to confirm the pathogenicity of bacterial isolates prior to using appropriate determinative tests to validate species-level identification. Accurate identification of the causal pathogen(s) of field observed symptoms is key to the development or selection of appropriate management strategies. With significant reductions in bacterial genome sequencing costs, as low as $220 per strain, and the availability of user-friendly bioinformatics tools, biologists and plant pathologists are encouraged to adopt the genome-based approach in the identification of pathogens of wheat. Whole-genome sequences parameters provide reproducible data leading to reliable and accurate identification of bacteria, in general, and wheat pathogens, in particular.



Funding for this work was provided by Agriculture and Agri-Food Canada through projects # J-002272 and # J-002749. The author is grateful to Dr. M. Harding and Dr. S. Wegulo for providing some of the photographs. Also, the author is thankful to CAB International (England) for authorizing the use of distribution maps generated in Plantwise.


  1. 1. Shewry PR. Wheat. Journal of Experimental Botany. 2009;60:1537-1553. DOI: 10.1093/jxb/erp058
  2. 2. Bockus WW, Bowden RL, Hunger M, Morrill WL, Murray TD, Smiley RW. Compendium of Wheat Disease. 3rd ed. St Paul, Minnesota, U.S.A.: The American Phytopathological Society; 2010. p. 117
  3. 3. Shewry PR, Hey SJ. The contribution of wheat to human diet and health. Food and Energy Security. 2015;4:178-202
  4. 4. Igrejas G, Branlard G. The importance of wheat. In: Igrejas G, Ikeda TM, Guzmán C, editors. Wheat Quality for Improving Processing and Human Health. Switzerland: Springer Nature; 2020. pp. 1-7. DOI: 10.1007/978-3-030-34163-3_1
  5. 5. Nigro D, Gadaleta A, Mangini G, Colasuonno P, Marcotuli I, Giancaspro A, et al. Candidate genes and genome-wide association study of grain protein content and protein deviation in durum wheat. Planta. 2019;249(4):1157-1175. DOI: 10.1007/s00425-018-03075-1
  6. 6. FAO/UN. FAO Food and Nutrition Paper 92. Dietary protein quality evaluation in human nutrition (PDF). Rome, Italy: Food and Agriculture Organization of the United Nations; 2013 [Accessed: November 22, 2021]
  7. 7. Duvieller E, Bragard C, Mariate H. Bacterial leaf streak and black chaff caused by Xanthomonas translucens. In: Duvieller E, Fucikovsky L, Rudolph K, editors. The Bacterial Diseases of Wheat: Concepts and Methods of Disease Management. Mexico: CIMMYT; 1997. pp. 25-47. Available from:
  8. 8. Adhikari TB, Gurung S, Hansen JM, Bonman JM. Pathogenic and genetic diversity of Xanthomonas translucens pv. undulosa in North Dakota. Phytopathology. 2012;102:390-402
  9. 9. Broadfoot WC, Robertson HT. Pseudo-black chaff of reward wheat. Science in Agriculture. 1933;13:512-514
  10. 10. Hagborg WAF. Black chaff, a composite disease. Canadian Journal of Research. 1936;14:347-359
  11. 11. McMullen M, Adhikar T. Bacterial leaf streak and black chaff of wheat. In: Plant Disease Management. Fargo, ND, USA: North Dakota State University Extension Publication; 2011. p. 1566
  12. 12. Johnson T, Hagborg WF. Melanism in wheat induced by high temperature and humidity. Canadian Journal of Research. 1944;22:7-10
  13. 13. Paul VH, Smith I. Bacterial pathogens of Gramineae: Systematic review and assessment of quarantine status for the EPPO region. Bulletin OEPP. 1989;19:33-42. DOI: 10.1111/j.1365-2338.1989.tb00126.x
  14. 14. Sapkota S, Mergoum M, Liu Z. The translucens group of Xanthomonas translucens: Complicated and important pathogens causing bacterial leaf streak on cereals. Molecular Plant Pathology. 2020;21:291-302. DOI: 10.1111/mpp.12909
  15. 15. Jones LR, Johnson AG, Reddy CS. Bacterial blight of barley. Journal of Agricultural Research. 1917;11:625-643
  16. 16. Tubajika KM, Tillman BL, Russin JS, Clark CA, Harrison SA. Relationship between flag leaf symptoms caused by Xanthomonas translucens pv. translucens and subsequent seed transmission in wheat. Plant Disease. 1998;82:1341-1344
  17. 17. Curland RD, Gao L, Bull CT, Vinatzer BA, Dill-Macky R, Van Eck L, et al. Genetic diversity and virulence of wheat and barley strains of Xanthomonas translucens from the upper midwestern United States. Phytopathology. 2018;108:443-453. DOI: 10.1094/PHYTO-08-17-0271-R
  18. 18. Kandel YR, Glover KD, Tande CA, Osborne LE. Evaluation of spring wheat germplasm for resistance to bacterial leaf streak caused by Xanthomonas campestris pv. translucens. Plant Disease. 2012;96:1743-1748
  19. 19. Bamberg RH. Black chaff disease of wheat. Journal of Agricultural Research. 1936;52:397-417
  20. 20. Duveiller E. Research on ‘Xanthomonas translucens’ of wheat and triticale at CIMMYT. Bulletin. 1989;19:97-103
  21. 21. Waldron R. The relationship of black chaff disease of wheat to certain physical and pathological characters. Science. 1929;70:268
  22. 22. Forster RL, Schaad NW. Control of black chaff of wheat with seed treatment and a foundation seed health program. Plant Disease. 1988;72:935-938
  23. 23. Duveiller E, Maraite H. Study of yield loss due to Xanthomonas campestris pv. undulosa in wheat under high rainfall temperate conditions. Journal of Plant Diseases Protection. 1993;100:453-459
  24. 24. Shane WW, Baumer JS, Teng PS. Crop losses caused by Xanthomonas streak on spring wheat and barley. Plant Disease. 1987;71:927-930
  25. 25. Tillman BL, Kursell WS, Harrison SA, Russin JS. Yield loss caused by bacterial streak in winter wheat. Plant Disease. 1999;83:609-614
  26. 26. Smith EF, Jones LR, Reddy CS. The black chaff of wheat. Science. 1919;50:48
  27. 27. Dye DW, Bradbury JF, Goto M, Hayward AC, Lelliott RA, Schroth MN. International standards for naming pathovars of phytopathogenic bacteria and a list of pathovar names and pathotype strains. Review Plant Pathology. 1980;59:153-168
  28. 28. Tambong JT, Xu R, Gerdis S, Daniels GC, Chabot D, Hubbard K, et al. Molecular analysis of bacterial isolates from necrotic wheat leaf lesions caused by Xanthomonas translucens, and description of three putative novel species, Sphingomonas albertensis sp. nov., Pseudomonas triticumensis sp. nov. and Pseudomonas foliumensis sp. nov. Frontiers in Microbiology. 2021;12:666689. DOI: 10.3389/fmicb.2021.666689
  29. 29. Pesce C, Bolot S, Cunnac S, Portier P, Fischer-Le Saux M, Jacques MA, et al. High quality draft genome sequence of the Xanthomonas translucens pv. cerealis pathotype strain CFBP 2541. Genome Announcements. 2015;3:e01574-ee1614
  30. 30. Shah SMA, Haq F, Ma W, Xu X, Wang S, Xu Z, et al. Tal1NXtc01 in Xanthomonas translucens pv. cerealis contributes to virulence in bacterial leaf streak of wheat. Frontiers in Microbiology. 2019;10:2040. DOI: 10.3389/fmicb.2019.02040
  31. 31. Cunfer BM, Scolari BL. Xanthomonas campestris pv. translucens on triticale and other small grains. Phytopathology. 1982;72:683-686
  32. 32. Bragard C, Singer E, Alizadeh A, Vauterin L, Maraite H, Swings J. Xanthomonas translucens from small grains: Diversity and phytopathological relevance. Phytopathology. 1997;87:1111-1117
  33. 33. Kim HK, Sasser M, Sands DC. Selective medium for Xanthomonas campestris pv. yranslucens. Phytopathology. 1982;72:936
  34. 34. Schaad NW, Forster RL. A semi-selective agar medium for isolating Xanthomonas campestris pv. Translucens from wheat seeds. Phytopathology. 1985;75:260-263
  35. 35. Duveiller E. A seed detection method of Xanthomonas campestris pv. undulosa, using a modification of Wilbrink’s agar medium. Parasitica. 1990a;46:3-17
  36. 36. Duveiller E. Bacterial leaf streak or black chaff of cereals. Bulletin OEPP/EPPO Bulletin. 1994;24:135-157
  37. 37. Mohan SK, Mehta YR. Studies on Xanthomonas campestris pv. undulosa in wheat and triticale in Paraná state. Fitopatologia Brasileira. 1985;10:447-453 (in Portuguese)
  38. 38. Dye DW. The inadequacy of the usual determinative tests for the identification of Xanthomonas spp. New Zealand Journal of Science. 1962;5:393-416
  39. 39. Miyajima K. Occurrence of bacterial blight Xanthomonas campestris pv. Hordei (Hagborg) Dye of orchard grass in Hokkaido prefecture. Annual Report Plant Protection North Japan. 1980;31:74-77 (in Japanese)
  40. 40. Bradbury JF. Family 1: Pseudomonaceae. Genus II: Xanthomonas Dowson 1939. In: Krieg NR, Holt JG, editors. Bergey’s Manual of Systematic Bacteriology. 1st ed. Baltimore/London: The Williams and Wilkins Co.; 1984. pp. 199-210
  41. 41. Schaad NW. Problems with the pathovar concept. In: Civerolo EL, Collmer A, Davis RE, Gillaspie AG, editors. Proceedings of the 6th International Conference on Plant Pathogenic Bacteria. Maryland. Dordrecht (The Netherlands): Martinus Nijhoff Publishers; 1987. pp. 783-785
  42. 42. Rademaker JLW, Norman DJ, Forster RL, Louws FJ, Schultz MH, De Bruijn FJ. Classification and identification of Xanthomonas translucens isolates, including those pathogenic to ornamental asparagus. Phytopathology. 2006;96:876-888
  43. 43. Maes M, Garbeva P, Kamoen O. Recognition and detection in seed of the Xanthomonas pathogens that cause cereal leaf streak using rDNA spacer sequences and polymerase chain reaction. Phytopathology. 1996;86:63-69. DOI: 10.1094/Phyto-86-63
  44. 44. Iqbal MA, Ullah I, Shahbaz MU, Kamran M, Saleem K. Biochemical and molecular identification of Xanthomonas translucens pv. Undulosa causing bacterial leaf streak of wheat in Punjab, Pakistan. Archives of Phytopathology and Plant Protection. 2013;47:417-424. DOI: 10.1080/03235408.2013.811030
  45. 45. Khojasteh M, Taghavi SM, Khodaygan P, Hamzehzarghani H, Chen G, Bragard C, et al. Molecular typing reveals high genetic diversity of Xanthomonas translucens strains infecting small-grain cereals in Iran. Applied and Environmental Microbiology. 2019;85:e01518-e01519. DOI: 10.1128/AEM.01518-19
  46. 46. Wichmann F, Vorholter FJ, Hersemann L, Widmer F, Blom J, Niehaus K, et al. The noncanonical type III secretion system of Xanthomonas translucens pv. Graminis is essential for forage grass infection. Molecular Plant Pathology. 2013;14:576-588
  47. 47. Gardiner DM, Upadhyaya NM, Stiller J, Ellis JG, Dodds PN, Kazan K, et al. Genomic analysis of Xanthomonas translucens pathogenic on wheat and barley reveals cross-kingdom gene transfer events and diverse protein delivery systems. PLoS One. 2014;9:e84995
  48. 48. Peng Z, Hu Y, Xie Z, Potnis N, Akhunova A, Jones J, et al. Long read and single molecule DNA sequencing simplifies genome assembly and TAL effector gene analysis of Xanthomonas translucens. BMC Genomics. 2016;17:21
  49. 49. Langlois PA, Snelling J, Hamilton JP, Bragard C, Koebnik R, Verdier V, et al. Characterization of the Xanthomonas translucens complex using draft genomes, comparative genomics, phylogenetic analysis, and diagnostic lamp assays. Phytopathology. 2017;107:519-527. DOI: 10.1094/PHYTO-08-16-0286-R
  50. 50. Boosalis MG. The epidemiology of Xanthomonas translucens (J.J. and R.) Dowson on cereals and grasses. Phytopathology. 1952;42:387-395
  51. 51. Sellam MA, aWilcoxson RD. Bacterial leaf blight of wheat in Minnesota. Plant Disease Report. 1976;60:242-245
  52. 52. Otta JD. Pseudomonas syringae incites a leaf necrosis on spring and winter wheats in South Dakota. Plant Disease Report. 1974;58:1061-1064
  53. 53. Otta JD. Occurrence and characteristics of isolates of Pseudomonas syringae on winter wheat. Phytopathology. 1977;67:22-26
  54. 54. Scharen AL, Bergman W, Burns EE. Leaf diseases of winter wheat in Montana and losses from them in 1975. Plant Disease Report. 1976;60:686-690
  55. 55. Varvaro L. Una batteriosi del Frumento duro (Triticum durum Desf.) causata da Pseudomonas syringae pv. atrofaciens (McCulloch) Young et al. in Italia. Informatore Fitopatologico. 1983;33:49-51
  56. 56. Smith J, Hattingh MJ. Fluorescent pseudomonads associated with diseases of wheat in South Africa. Journal of Phytopathology. 1991;133:36-48
  57. 57. Akhtar MA. Outbreaks and new records. Pakistan. Bacterial gumming disease of wheat. FAO Plant Protection Bulletin. 1987;35:102
  58. 58. Teyssandier E, Sands DC. Wheat leaf blight caused by Pseudomonas syringae in Argentina. Phytopathology. 1977;87:1766
  59. 59. Elliot C. Manual of Bacterial Plant Pathogens. 2nd ed. Waltham (Massachusetts): Chronica Botanica; 1951. p. 186
  60. 60. Bradbury JF. Guide to Plant Pathogenic Bacteria. CAB International Mycological Institute: Farnham House, Slough, UK; 1986. p. 332
  61. 61. CABI/EPPO. Pseudomonas syringae pv. syringae. Distribution Maps of Plant Diseases No. 336. Wallingford, UK: CABI Head Office; 2012
  62. 62. von Kietzell J, Rudolf K. Wheat diseases caused by Pseudomonas syringae pathovars. In: Duveiller E, Fucikovsky L, Rudolf K, editors. The Bacterial Diseases of Wheat: Concepts and Methods of Disease Management. Mexico City: CIMMYT; 1997. pp. 49-57
  63. 63. Toben H. Untersuchungen über die basale Spelzenfäule an Weizen und Gerste, hervorgerufen durch Pseudomonas syringae pv. atrofaciens ((McCull.) Young, Dye, Wilkie) [diploma thesis]. Göttingen, Germany: Universität Göttingen, Institut für Pflanzenpathologie und Pflanzenschutz; 1989
  64. 64. MCculloch L. Basal glume rot of wheat. Journal of Agricultural Research. 1920;18:543-551
  65. 65. Matveeva IEV, Pekhtereva ES, Polityko VA, Ignatov AN, Nikolaeva EV, Schaad NW. Distribution and virulence of Pseudomonas syringae pv. atrofaciens, causal agent of basal glume rot, in Russia. In: Rudolph K, Burr TJ, Mansfield JW, Stead D, Vivian A, and von Kietzell J, editors. Pseudomonas syringae and Related Pathogens. The Netherlands: Kluwer Academic Publishers. (2003). P. 97-105. Available from:
  66. 66. Duveiller E. Collaborative research network on bacterial diseases of wheat. In: 3rd Annual Report on CIMMYT Project Bacterial Diseases Network of Wheat. Mexico: CIMMYT; 1990. pp. 5-6
  67. 67. Galatchian MJ. Testing the pathogenicity of strains of Bacterium atrofaciens (McCull.) under laboratory conditions. C.R. pan. Sov. V.I. Lenin Academy of Agricultural Science, Moscow. 1941;6:40-43 (in Russian)
  68. 68. Karov S, Vassilev VI. Studies on wheat bacteriosis in Bulgaria. Scientific Works. 1981;26:85-90 (in Bulgarian)
  69. 69. Noble RJ. Basal glume rot. Agricultural Gazette New South Wales. 1933;44:107-109
  70. 70. Wilkie JP. Basal glume rot of wheat in New Zealand. New Zealand Journal of Agricultural Research. 1973;16:155-160
  71. 71. Dippenaar BJ. ‘n Bakteriese wat‘n verdorring van die kaffies van koringare veroorsaak. South African Journal of Science. 1931;28:280-283 (in Africaans)
  72. 72. Rudolph K, Mavridis A. Über die Isolation von Pseudomonas syringae pv. atrofaciens aus Winterweizenähren in Schafhausen (Sindelfingen). Phytomedizin, Mitt. Deutsch. Phytomed. Ges. 1987;17:20
  73. 73. Toben H, Mavridis A, Rudolph WE. On the occurrence of basal glume rot of wheat and barley caused by Pseudomonas syringae pv. Atrofaciens in West Germany. Journal of Plant Disease and Protect. 1991;98:225-235 (in German)
  74. 74. Vassilev VI, Karov S. Effect of cereal basal bacteriosis on the economic physico-technological and biochemical properties of wheat grain and flour. Plant Science. 1985;22:13-20 (in Bulgarian)
  75. 75. Mavridis A, Meyer D, Mielke H, Steinkampf G. Zum Auftreten und zur Schadwirkung der basalen Spelzenfäule beim Sommerweizen. Kali- Briefe (Büntehof). 1991;20:469-473
  76. 76. Iacobellis NS, Figliuolo G, Janse J, Scortichini M, Ciuffreda G. Characterization of Pseudomonas syringae pv. Atrofaciens. In: Rudolph K, Burr TJ, Mansfield JW, Stead D, Vivian A, von Kietzell J, editors. Pseudomonas syringae Pathovars and Related Pathogens. Dordrecht (The Netherlands): Kluwer Academic Publishers. 1997. pp. 500-504.
  77. 77. Claflin LE, Ramundo BA. Evaluation of the dot-immunobinding assay for detecting phytopathogenic bacteria in wheat seeds. Journal of Seed Technology. 1987;11:52-61
  78. 78. Toben H, Mavridis A, Rudolph KWE. Basal glume rot (Pseudomonas syringae pv. Atrofaciens) on wheat and barley in FRG and resistance screening of wheat. Bulletin OEPP/EPPO Bulletin. 1989;19:119-125
  79. 79. Maraite H, Weyns, J. Pseudomonas syringae pv. aptata and pv. atrofaciens, specific pathovars or members of pv. syringae?. In: Rudolph K, Burr TJ, Mansfield JW, Stead D, Vivian A, von Kietzell J, (eds Pseudomonas syringae Pathovars and Related Pathogens.). Dordrecht (The Netherlands):Kluwer Academic Publishers, 1997. pp. 515-520
  80. 80. Mohan SK, Schaad NW. An improved agar plating assay for detecting Pseudomonas syringae pv. syringae and P. s. pv. phaseolicola in contaminated bean seed. Phytopathology. 1987;77:1390-1395
  81. 81. Carlson RR, Vidaver AK. Bacterial mosaic, a new corynebacterial disease of wheat. Plant Disease. 1982;66:76-79
  82. 82. Li X, Tambong J, Yuan KX, Chen W, Xu H, Lévesque CA, et al. Re-classification of Clavibacter michiganensis subspecies on the basis of whole-genome and multi-locus sequence analyses. International Journal of Systematic and Evolutionary Microbiology. 2018;68:234-240. DOI: 10.1099/ijsem.0.002492
  83. 83. McBeath JH, Adelman M, Jackson L. Screening wheat germplasm for Corynebacterium michiganense subsp. tessellarius. Phytopathology. 1988;78:1566
  84. 84. Scharif G. Corynebacterium iranicum sp. nov. on wheat (Triticum vulgare L.) in Iran and a comparative study of it with C. tritici and C. rathayi. Entomologie et phytopathologie appliquées. 1961;19:1-24
  85. 85. Zgurskaya HI, Evtushenko LI, Akimov VN, Kalakovtskii LV. Rathayibacter gen. nov., including the species Rathayibacter rathayi comb. nov., Rathayibacter tritici comb. nov., Rathayibacter iranicus comb. nov., and six strains from annual grasses. International Journal of Systematic Bacteriology. 1993;43:143-149
  86. 86. Bamdadian A. The importance and situation of wheat diseases in Iran. In: CENTO Panel on Pests and Diseases of Wheat, Tehran University, College of Agriculture; 5-7 February 1973; Karaj, Iran. Tehran Iran: Central Treaty Organization (CENTO); 1973. pp. 57-63
  87. 87. Paruthi IS, Bhatti DS. Estimation of loss in yield and incidence of Anguina triticina on wheat in Haryana (India). International Nematology Network Newsletters. 1985;2:13-16
  88. 88. Benjama A. Bacterial diseases of cereals in Morocco. Cahiers Agricultural. 1997;6:605-610
  89. 89. Fattah FA. Effects of inoculation on the incidence of ear-cockle and 'tundu' on wheat under field conditions. Plant and Soil. 1988;109:195-198
  90. 90. Riley IT, Reardon TB. Isolation and characterization of Clavibacter tritici associated with Anguina tritici in wheat from Western Australia. Plant Pathology. 1995;44:805-810
  91. 91. Fucikovsky L, Duvieller E. Other plant pathogenic bacteria reported on wheat. In: Duvieller E, Fucikovsky L, Rudolph K, editors. The Bacterial Diseases of Wheat: Concepts and Methods of Disease Management. Mexico: CIMMYT; 1997. pp. 59-64
  92. 92. Tanii, Miyajima K, Akita T. The sheath brown rot disease of rice plant and its causal bacterium Pseudomonas fuscovaginae A. Tanii, K. Miyajima and T. Akita sp. nov. Annals Phytopathology Socical Japan. 1976;42:540-548
  93. 93. Duveiller E, Maraite H. Bacterial sheath rot of wheat caused by Pseudomonas fuscovaginae in the highlands of Mexico. Plant Disease. 1990;74:932-935
  94. 94. Tohya M, Watanabe S, Tada T, Tin HH, Kirikae T. Genome analysis-based reclassification of Pseudomonas fuscovaginae and Pseudomonas shirazica as later heterotypic synonyms of Pseudomonas asplenii and Pseudomonas asiatica, respectively. International Journal of Systematic and Evolutionary Microbiology. 2020;70:3547-3552. DOI: 10.1099/ijsem.0.004199
  95. 95. Anonymous. Occurrence of Sheath Rot, a New Disease in Wheat. Vol. 1. Pokhara, Nepal: Lumle Newsletter, Lumle Agricultural Research Centre; 1995. p. 3
  96. 96. Klement Z. Tobacco (HR) test for the quick demonstration of pathogenicity. In: Klement Z, Rudolph K, Sands DC, editors. Methods in Phytobacteriology. Akadémiai Kiadó: Budapest; 1990. pp. 101-102
  97. 97. Cottyn B, Ceres MT, Van outryve MF, Baroga J, Mew TW. Bacterial diseases of rice. I. Pathogenic bacteria associated with sheath rot complex and grain discoloration of rice in the Philippines. Plant Disease. 1996;80:429-437
  98. 98. Piening LJ, MacPherson DJ. Stem melanosis, a disease of spring wheat caused by pseudomonas cichorii. Canadian Journal of Plant Pathology. 1985;7:168-172
  99. 99. Piening LJ, MacPherson DJ, Mahli SS. The effect of copper in reducing stem melanosis of park wheat. Canadian Journal of Plant Science. 1987;67:1089-1091
  100. 100. Piening LJ, MacPherson DJ, Mahli SS. Stem melanosis of some wheat, barley, and oat cultivars on a copper deficient soil. Canadian Journal of Plant Science. 1989;11:65-67
  101. 101. Howe ET, Simmonds PM. Bacterial pink blotch of wheat. In: Conners IG, Graigie JH, Vanterpool TC, editors. Proceedings of the Canadian Phytopathological Society; June 28-30, 1937; Ottawa, Canada. 17th ed. 1937. p. 6
  102. 102. Luisetti J, Rapilly F. Sur une altération d’origine bactérienne des graines de blé. Ann. Epiphyt. Phytogenet. 1967;18:483-493
  103. 103. Roberts P. Erwinia (Millard)Burkholder associated with pink grain of wheat. The Journal of Applied Bacteriology. 1974;37:353-358
  104. 104. McMullen MP, Stack RW, Miller JD, Bromel MC, Youngs VL. Erwinia rhapontici, a bacterium causing pink wheat kernels. In: Proceedings of the North Dakota Academy of Sciences; Grand Forks, ND. Vol. 38. 1984. p. 78
  105. 105. Dutrecq A, Debras P, Stevaux J, Klaessens D. Estimation de la flore bactérienne sur les épis de froment échaudés. Parasitica. 1990;46:69-84
  106. 106. Diekmann M, Putter CAJ. FAO/IPGRI Technical Guidelines for the Safe Movement of Germplasm. No. 14. Small Grain Temperate Cereals. Rome: FAO/IPGRI; 1995. p. 67
  107. 107. Huang HC, Phillippe RC, Phillippe LM. Pink seed of pea: A new disease caused by Erwinia rhapontici. Canadian Journal of Plant Pathology;12:445-448
  108. 108. Ohuchi A, Ohsawa T, Nishimura J. Two pathogenic bacteria, Erwinia rhapontici (Millard 1924) Burkholder 1948 and Pseudomonas marginalis pv. Marginalis (Brown 1918) Stevens 1925, causing a soft rot of onion. Japanese Annals of the Phytopathological Society. 1983, 1990;49:619-626
  109. 109. Choi JE, Han KS. Studies on the bacterial soft rot disease of Liliaceaecrops in Korea: 2. Soft rot of garlic caused by Erwinia. Korean Journal of Plant Pathology. 1989;5:271-276
  110. 110. Huang HC, Erickson RS, Yanke LJ, Mündel H-H. First report of pink seed of common bean caused by Erwinia rhapontici. Plant Disease. 2002;86:921
  111. 111. Huang HC, Erickson RS, Yanke LJ, Hsieh TF, Morrall RAA. First report of pink seed of lentil and chickpea caused by Erwinia rhapontici in Canada. Plant Disease. 2003;87:1398
  112. 112. Stackebrandt E. The richness of prokaryotic diversity: There must be a species somewhere. Food Technology and Biotechnology. 2003;41:17-22
  113. 113. Tambong JT. Taxogenomics and systematics of the genus Pantoea. Frontiers in Microbiology. 2019;10:2463
  114. 114. Thompson CC, Chimetto L, Edwards RA, Swings J, Stackebrandt E, Thompson FL. Microbial genomic taxonomy. BMC Genomics. 2013;14:913. DOI: 10.1186/1471-2164-14-913
  115. 115. Meier-Kolthoff JP, Auch AF, Klenk HP, Goker M. Genome sequence-based species delimitation with confidence intervals and improved distance functions. BMC Bioinformatics. 2013;14:60. DOI: 10.1186/1471-2105-14-60
  116. 116. Jain C, Rodriguez RL, Phillippy AM, Konstantinidis KT, Aluru S. High throughput ANI analysis of 90K prokaryotic genomes reveals clear species boundaries. Nature Communications. 2018;9:5114
  117. 117. Goris J, Konstantinidis KT, Klappenbach JA, Coenye T, Vandamme P, Tiedje JM. DNA-DNA hybridization values and their relationship to whole-genome sequence similarities. International Journal of Systematic and Evolutionary Microbiology. 2007;57:81-91. DOI: 10.1099/ijs.0.64483-0
  118. 118. Teeling H, Meyerdierks A, Bauer M, Amann R, Glockner FO. Application of tetranucleotide frequencies for the assignment of genomic fragments. Environmental Microbiology. 2004;6:938-947
  119. 119. Duret L. Evolution of synonymous codon usage in metazoans. Current Opinion in Genetics & Development. 2002;12:640-649
  120. 120. Wattam AR, Abraham D, Dalay O, Disz TL, Driscoll T, Gabbard JL, et al. PATRIC, the bacterial bioinformatics database and analysis resource. Nucleic Acids Research. 2014;42:581-591
  121. 121. Afgan E, Baker D, Batut B, van den Beek M, Bouvier D, Čech M, et al. The galaxy platform for accessible, reproducible and collaborative biomedical analyses. Nucleic Acids Research. 2018;46:W537-W544. DOI: 10.1093/nar/gky379

Written By

James T. Tambong

Submitted: 06 January 2022 Reviewed: 25 January 2022 Published: 16 March 2022