In Great Britain and/or Ireland:
Foodplant / pathogen
Xanthomonas campestris pv. campestris infects and damages live, yellow-blotched leaf of Crambe maritima

Foodplant / pathogen
Xanthomonas campestris pv. campestris infects and damages live, yellow-blotched leaf of Brassicaceae

Foodplant / pathogen
Xanthomonas campestris pv. campestris infects and damages live, yellow-blotched leaf of Brassica
Other: major host/prey


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Xanthomonas campestris

Xanthomonas campestris is bacterial species that causes a variety of plant diseases. Available from the NCPPB and other international Culture collections such as ICMP, ATCC, and LMG in a purified form, it is used in the commercial production of a high-molecular-weight polysaccharide - xanthan gum - that is an efficient viscosifier of water and that has many important uses, especially in the food industry. It causes spots on the infected plant.

Types of Xanthomonas campestris[edit]

(pv. means pathovar, a type of classification based on the host plant that is attacked by Xanthomonas campestris)

  • Xanthomonas campestris pv. armoraciae
  • Xanthomonas campestris pv. begoniae A
  • Xanthomonas campestris pv. begoniae B
  • Xanthomonas campestris pv. campestris
  • Xanthomonas campestris pv. carota
  • Xanthomonas campestris pv. corylina
  • Xanthomonas campestris pv. dieffenbachiae
  • Xanthomonas campestris pv. graminis
  • Xanthomonas campestris pv. hederae
  • Xanthomonas campestris pv. hyacinthi
  • Xanthomonas campestris pv. juglandis - the walnut blight
  • Xanthomonas campestris pv. malvacearum or Xanthomonas citri subsp. malvacearum [1]
  • Xanthomonas campestris pv. musacearum
  • Xanthomonas campestris pv. nigromaculans
  • Xanthomonas campestris pv. pelargonii
  • Xanthomonas campestris pv. phaseoli
  • Xanthomonas campestris pv. poinsettiicola
  • Xanthomonas campestris pv. prunii
  • Xanthomonas campestris pv. raphani
  • Xanthomonas campestris pv. sesami
  • Xanthomonas campestris pv. tardicrescens
  • Xanthomonas campestris pv. translucens
  • Xanthomonas campestris pv. vesicatoria
  • Xanthomonas campestris pv. viticola[2]

The former pv. citri, which causes citrus canker, was reclassified as X. axonopodis in 1995 Xanthomonas campestris#cite note-0. In 2006, the species designations for pv. citri and malvacearum were revised to X. citri and these pathovars are now referred to as subspecies Xanthomonas campestris#cite note-1.


  • Gerhard Reuther, Martin Bahmann: Elimination of Xanthomonas campestris pv. pelargonii by Means of Micropropagation of Pelargonium Stock Plants; In: 3rd International Geranium Conference, 1992. Proceedings, Ball Publishing Batavia, IL. USA ; (1992),


  1. ^
  2. ^ Outbreak of grapevine bacterial canker disease in India. R. Chand and R. Kishun, Vitis, 1990, volume 29, pages 183-188 (article)
  • Schaad NW, Postnikova E, Lacy GH, Sechler A, Agarkova I, Stromberg PE, Stromberg VK, Vidaver AK (2006). "Emended classification of xanthomonad pathogens on citrus." Syst Appl Microbiol 29(8): 690-695.
  • Vauterin L, Hoste B, Kersters K, and Swings J (1995). "Reclassification of Xanthomonas." Int J Syst Bacteriol 45: 472-489.
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Banana Xanthomonas wilt

Banana Xanthomonas Wilt (BXW), or banana bacterial wilt (BBW) or enset wilt is a bacterial disease caused by Xanthomonas campestris pv. musacearum.[1] After being originally identified on a close relative of banana, Ensete ventricosum, in Ethiopia in the 1960s,[2] BXW emanated in Uganda in 2001 affecting all types of banana cultivars. Since then BXW has been diagnosed in Central and East Africa including banana growing regions of: Rwanda, Democratic Republic of the Congo, Tanzania, Kenya, Burundi, and Uganda.[3]

Of the numerous diseases infecting bananas, BXW alongside banana bunchy top virus has been the most devastating in recent years. Global concern arose over the livelihoods of African banana farmers and the millions relying on bananas as a staple food when the disease was at its worst between the years 2001 and 2005. It was estimated that in Central Uganda from 2001 and 2004, there was a 30-52 % decease in banana yield due to BXW infection.[4] Although extensive management of the disease outbreaks has helped reduce the impact of Banana Xanthomonas Wilt even today BXW continues to a pose a real problem to the banana farmer of Central and East Africa.


BXW symptoms can be sorted into two domains: symptoms on the inflorescence and symptoms on the fruit. Symptoms on the fruit are usually used to distinguish BXW from alternative banana diseases. A bacterial ooze is excreted from the plant organs and this is a mandatory sign that BXW may be present. Common symptoms on the fruit include internal discoloration and premature ripening of the fruit. A cross section of the BXW infected banana is characterized by the yellow- orange discoloration of the vascular bundles and dark brown tissue scaring.[5] Symptoms on the inflorescence include a gradual wilting and yellowing of the leaves plus wilting of the bracts and shriveling of the male buds.[6] Many factors may affect the combination of disease symptoms on show. These include the particular cultivar infected, how the disease has been transmitted and the current growing season.



Soil is one of the main sources for Xanthomonas campestris pv. musacearum inoculum.[6] Xanthomonas campestris pv. musacearum may contaminate the soil for four months and more. BXW awareness campaigns have helped reduce the numbers of farmers growing bananas on contaminated plantains aiding in the control of the disease overall. Transmission of contaminated disease itself is thought to be low.


It widely thought that Xanthomonas campestris pv. musacearum bacteria is transmitted to airborne vectors through exposed male flowers (see plant reproductive morphology). Xanthomonas campestris pv. musacearum bacteria has been isolated from the ooze and nectar excreted from openings of fallen male flowers.[7] Insects, namely stingless bees (Apidae), fruit flies (Drosophilidae) and grass flies (Chloropidae), transmit the disease from banana to banana after being drawn to the infected nectar.[8] If the disease has been transmitted by insects the symptoms tend to first appear on the male buds of the banana plant.


The knife (panga) is used almost universally in African agriculture. Use of contaminated knives was a common method for disease spread when the disease first originated but increased knowledge of BXW transmission has led to increased numbers knives being disinfected after use. Herbicides are now advised as a more economical and effective way of destroying infected banana crop.[9]

Infected plant material[edit]

BXW infects all parts of the plant. Disease spread has been primarily linked with the transport of plants shoots for replanting.[8] Other parts of the plant such as the male buds (used in banana beer production) and mulch (banana waste material) can also expose novel regions to the disease.[8]

Disease management[edit]

Control of BXW is based upon a variety of methods to help prevent the spread of the disease. Vigilance and the quick removal of infected plants remain critical to minimising spread of the disease.

Infected plants can be removed using herbicides or more commonly by cutting the plant into small fragments and decomposing. The risk of infection can be lowered by removal of the male bud ('debudding') but many farmers believe this is essential to the quality of the banana fruit. The risk of infection decreases if the plants are not covered with topsoil.[3] However the risk of disease should be balanced against the resulting decrease in yield of the banana plantain. A major part of disease control is the disinfecting of the tools used.

Much of the work in controlling BXW has been done through educational campaigns raising awareness of the disease to the banana farmers. For example: in Uganda and Tanzania where the government has actively worked alongside farmers to help limit spread of the disease, over 90% control of BXW has been reported.[6] Moreover much of the information taught to the farmers can be used in the control of other banana infecting diseases.

BXW resistant banana[edit]

No banana cultivars in Central and Eastern Africa have shown any resistance to BXW despite some varieties, such as those in the 'Pisang Awak' region, showing increased susceptibility. Scientists have recently transferred two genes from sweet green pepper to bananas in order to confer resistance to BXW.[10][11] This is a promising step forward in circumventing the time consuming and expensive practices of disease management such as 'debudding'.

Pflp and Hrap genes encoding the proteins plant ferredoxin-like amphipathic protein (pflp) and hypersensitive response-assisting protein (hrap) were isolated from sweet pepper and introduced to the genome of East African bananas using genetic engineering. The two proteins induced a hypersensitive response and systemic acquired resistance within the banana plant after being exposed to the bacterial pathogen. It was reported that over half of the transgenic bananas were resistant to BXW,[10] resistance that was also found in field trials.[12]


  1. ^ Tushemereirwe, W. Kangire, A. Ssekiwoko, F. Offord, L.C. Crozier, J. Boa, E. Rutherford, M. Smith J.J. (2004). "First report of Xanthomonas campestris pv. musacearum on banana in Uganda". Plant Pathology 53: 802. doi:10.1111/j.1365-3059.2004.01090.x. 
  2. ^ Bradbury, J.F. Yiguro, D. (1968). "Bacterial wilt of Enset ("Ensete ventricosa") incited by "Xanthomonas musacearum".". Phytopathology 58: 111–112. 
  3. ^ a b Mwangi, M. Bandyopadhyay, R. Ragama,P. Tushemereirwe, R.K. (2007). "Assessment of banana planting practices and cultivar tolerance in relation to management of soilborne Xanthomonas campestris pv. musacearum". Crop Protection 26: 1203–1208. doi:10.1016/j.cropro.2006.10.017. 
  4. ^ Karamura, E. et al. (2010). "Assessing the Impacts of Banana Bacterial Wilt Disease on Banana(Musa spp.) Productivity and Livelihoods of Ugandan Farm Households.". Acta Horticulture (ISHS) 879: 749–755. 
  5. ^ Biruma, M. et al. (2007). "Banana Xanthomonas wilt: a review of the disease, management strategies and future research directions". African Journal of Biotechnology 6: 953–962. 
  6. ^ a b c Abele, s. et al. (2009). "Xanthomonas Wilt A threat to banana production in East and Central Africa". Plant Disease: 439–451. 
  7. ^ Tinzaaara, W. et al. (2006). "The possible role of insects in the transmission of Banana Xanthomonas Wilt". p. 60. 
  8. ^ a b c Smith, J.J. et al. (2008). "An analysis of the risk from Xanthomonas campestris pv. musacearum to banana cultivation in Eastern, Central and Southern Africa". Biodiversity International. 
  9. ^ Blomme, G. Turyagyenda, L.F Mukasa, H. Eden-Green, S. (2008). "The effectiveness of different herbicides in the destruction of Banana Xanthomonas Wilt infected plants". African Crop Science Journal 16: 103–110. 
  10. ^ a b Namukwaya, B. et al. (2012). "Transgenic banana expressing Pflp gene confers enhanced resistance to Xanthomonas wilt disease". Transgenic Resistance 21: 855–865. doi:10.1007/s11248-011-9574-y. 
  11. ^ Tripathi, L. Mwaka, H. Tripathi, J.N. Tushemereirwe, W.K. (2010). "Expression of sweet pepper Hrap gene in banana enhances resistance to Xanthomonas campestris pv. musacearum". Molecular Plant Pathology 6: 721–731. 
  12. ^ Tripathi, L., Tripathi, J. N., Kiggundu, A., Korie, S., Shotkoski, F., & Tushemereirwe, W. K. (2014). "Field trial of xanthomonas wilt disease-resistant bananas in east africa.". Nat Biotech 32: 868–870. doi:10.1038/nbt.3007/>. 
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Xanthomonas campestris pv. campestris

Black rot, caused by the bacterium Xanthomonas campestris pv. campestris (Xcc), is considered the most important and most destructive disease of crucifers, infecting all cultivated varieties of brassicas worldwide.[1][2] This disease was first described by botanist and entomologist Harrison Garman in Lexington, Kentucky, USA in 1889.[3] Since then, it has been found in nearly every country in which vegetable brassicas are commercially cultivated.[4]

Host infection by Xcc can occur at any stage of the plant life cycle. Characteristic symptoms of black rot caused by Xcc are V-shaped chlorotic to necrotic lesions extending from the leaf margins and blackening of vascular tissues.

The pathogen thrives in warm and humid climates and is rapidly disseminated in the field. Use of clean seed, crop rotation, and other cultural practices are the primary means of control of black rot. However, in developing countries such as those in South and Eastern Africa, black rot remains the greatest impediment to cabbage cultivation due to unreliable "clean" seed, multiple croppings annually, and high susceptibility of popular local cultivars to the disease.[5]

Hosts and symptoms[edit]

Members of the plant family Brassicaceae (Cruciferae), which includes cabbage, broccoli, cauliflower, kale, turnip, oilseed rape, mustard, radish, and the model organism Arabidopsis thaliana are affected by black rot.[1][6][7][8][2]

Host infection by Xcc causes V-shaped chlorotic to necrotic foliar lesions, vascular blackening, wilting, stunted growth, and stem rot symptoms.[1] As the pathogen proceeds from the leaf margins towards the veins, water stress and chlorotic symptoms develop due to occlusion of water-conducting vessels by bacterial exopolysaccharides and components of degraded plant cell walls.[1][6] The darkening of vascular tissues following bacterial invasion gives the black rot disease its name.[2] Lesions produced by Xcc may serve as portals of entry for other soft-rot pathogens such as Pectobacterium carotovorum (formerly Erwinia carotovora) and Pseudomonas marginalis.[1][2][8]

These symptoms may be confused with fusarium wilt of cabbage (fusarium yellows), caused by the fungus Fusarium oxysporum f. sp. conglutinans. In contrast to black rot, in which the pathogen invades leaf margins and causes chlorotic to necrotic symptoms that progress downwards in the plant, fusarium wilt symptoms first develop in the lower portions of the plant and move upwards.[9] Furthermore, leaf veins invaded by Xcc turn black compared to the dark brown vein discoloration found in fusarium wilt.[10][11]

Symptoms of black rot may vary widely among different species of crucifers. On cauliflower, Xcc infection via stomates causes black or brown specks, scratched leaf margins, black veins, and discolored curds.[12] Additionally, the severity of symptoms and aggressiveness of the disease varies between different strains of the Xcc pathogen.[1] The isolates can be differentiated into races based on the reaction of several Brassica lines after inoculation. A race structure including 5 races (0 to 4) was first proposed in 1992;[13] a revised classification model with 6 races was proposed in 2001[14] and, more recently, the model was expanded to include nine races.[15][16]

V-shaped chlorotic to necrotic lesion on cabbage leaf caused by the black rot pathogen Xanthomonas campestris pv. campestris
V-shaped chlorotic to necrotic lesion on cabbage leaf, symptomatic of infection by the black rot pathogen Xanthomonas campestris pv. campestris. Photo by David B. Langston, University of Georgia.

Disease cycle[edit]

Life cycle of the black rot pathogen Xanthomonas campestris pv. campestris
Life cycle of the black rot pathogen Xanthomonas campestris pv. campestris by G. Kwan.

The primary source of inoculum is Xcc infected seed.[1] During germination, the seedling becomes infected through the epicotyl [1] and cotyledons may develop blackened margins, shrivel, and drop.[6] The bacteria progress through the vascular system to the young stems and leaves, where the disease manifests as V-shaped chlorotic to necrotic lesions extending from the leaf margins. Under humid conditions, bacteria present in guttation droplets can be spread by wind, rain, water splashes, and mechanical equipment to neighboring plants.[1][6]

The natural route of invasion by Xcc is through the hydathodes, though leaf wounds caused by insects and plant roots may also be portals of entry.[1] Occasionally, infections occur through stomata. Hydathodes provide the pathogen a direct path from the leaf margins to the plant vascular system and thus systemic host infection. Invasion of the suture vein leads to production of Xcc infected seed.

Xcc can survive in plant debris in soil for up to 2 years, but not more than 6 weeks in free soil.[1] Bacteria present in plant debris can serve as a source of secondary inoculum.


Warm and wet conditions favor plant infection by Xcc and the development of disease.[6][8] Free moisture is required for host invasion, considering that the natural route of infection is through the hydathodes.

The optimum temperature range for bacterial growth and host symptom development is between 25° to 30 °C . A slower rate of growth is observed at temperatures as low as 5 °C and up to 35 °C.[6] However, infected hosts are symptomless below 18 °C.[17]


Management of black rot relies heavily on cultural practices:[6][7]

  • Use of certified disease-free seeds and transplants
  • Hot water treatment of non-certified seeds; chemical treatments with sodium hypochlorite, hydrogen peroxide, and hot cupuric acetate or zinc sulfate may also be used
  • Control of insects
  • Crop rotation with non-cruciferous plants (3-4 years)
  • Removal of crop debris after harvest
  • Control of cruciferous weeds that may serve as reservoir for the pathogen
  • Sanitation (e.g., clean equipment, avoiding work in wet fields, etc.)

The development and use of black rot resistant cultivars has long been recognised as an important method of control, but in practice has had limited success. Resistance to the most important pathogenic races of Xcc is rare in B. oleracea (e.g., cabbage, broccoli, cauliflower); the most common and potentially useful sources of black rot resistance occur in other brassica genomes including B. rapa, B. nigra, B. napus, B. carinata and B. juncea.[18]

Resistant or tolerant cabbage cultivars are available and include:[6][8]

  • Atlantis
  • Blueboy
  • Bravo
  • Bronco
  • Cecile
  • Defender
  • Dynasty
  • Gladiator
  • Guardian
  • Hancock
  • Ramada


Economic impact[edit]

Cabbage cultivation is a multi-billion dollar industry worldwide, reflecting its value as a vegetable crop, source of vegetable oil, component of fodder crop for livestock feed, and ingredient in condiments and spices. In 2007, the cabbage crop in the US exceed $413M (1.4M+ tons).[19] Black rot is considered the most important disease of cabbage and other crucifers because Xcc infections may not become apparent until the warm summer months (well after planting), the pathogen spreads rapidly, and losses due to the disease may exceed 50% in warm, wet climates.[6] The importance of using disease-free seed and/or transplants is highlighted by the fact that “as few as three infected seeds in 10,000 (0.03%) can cause black rot epidemics in a field.[6]” In transplant beds, an initial infection level of 0.5% can rise to 65% in just three weeks.[2] In fact more recent work [20] indicates that spread can be much more rapid than this: with overhead gantry irrigation, spread of the pathogen greatly exceeded symptom spread to the extent that in one experiment almost 100% of the transplants were infested in a block of 15 module trays (approx. 4500 plants) six weeks after sowing from a single primary infector. Modelling of the rate of spread in transplants indicates that the widely used tolerance standard for seed health testing (0·01%) should be revised to 0·004% [21]



Xanthan is an exopolysaccharide produced by Xcc. Commercially produced xanthan is used as a thickening food additive and lubricant, amongst other industrial applications.[2]


The genomes of three Xcc strains — ATCC 33913, B100, and 8004 — have been fully sequenced and are publicly available.[22][23][24]


  1. ^ a b c d e f g h i j k Alvarez AM. "Black rot of crucifers." In: Slusarenko AJ, Fraser RSS, van Loon LC (Eds.) Mechanisms of Resistance to Plant Diseases. Dordrecht, The Netherlands: Kluwer Academic Publishers, 2000. pp 21-52.
  2. ^ a b c d e f Williams PH. "Black rot: a continuing threat to world crucifers." Plant Disease 64.8 (1980): 736-742.
  3. ^ Garman H. "A bacterial disease of cabbage." Kentucky Agric Exp Stat Rep 3 (1890):43-46.
  4. ^ Chupp C. “Black rot of cabbage.” Manual of Vegetable Plant Diseases. New Delhi, India : Discovery Publishing House, 2006. p. 132-133.
  5. ^ Massomo SMS, Mabagala RB, Swai IS, Hockenhull J, Mortensen CN . “Evaluation of varietal resistance in cabbage against the black rot pathogen, Xanthomonas campestris pv. campestris in Tanzania.” Crop Protection 23,4(2004): 315-325.
  6. ^ a b c d e f g h i j "Black rot of cabbage and other crucifers." Integrated Pest Management. University of Illinois Extension. Dec 1999.
  7. ^ a b Miller SA, Sahin F, and Rowe RC. "Black rot of crucifers." Extension fact sheet HYG-3125-96. Ohio State University Extension. 1996.
  8. ^ a b c d Seebold K, Bachi P, and Beale J. "Black rot of crucifers ." UK Cooperative Extension Service . University of Kentucky. Feb 2008.
  9. ^ Sherf, A. "Fusarium yellows of cabbage and related crops." New York State Cooperative Extension. Cornell University. Jan 1979.
  10. ^ Sherf, A. "Fusarium yellows of cabbage and related crops." New York State Cooperative Extension. Cornell University. Jan 1979.
  11. ^ "Black rot of cabbage and other crucifers." Integrated Pest Management. University of Illinois Extension. Dec 1999.
  12. ^ Miller SA, Sahin F, and Rowe RC. "Black rot of crucifers." Extension fact sheet HYG-3125-96. Ohio State University Extension. 1996.
  13. ^ Kamoun S, Kamdar HV, Tola E, Kado CI. “Incompatible interactions between crucifers and Xanthomonas campestris involve a vascular hypersensitive response: Role of the hrpX locus.” Molecular Plant-Microbe Interactions 5 (1992): 22-33.
  14. ^ Vicente JG, Conway J, Roberts SJ, Taylor JD. “Identification and origin of Xanthomonas campestris pv. campestris races and related pathovars.” Phytopathology 91 (2001): 492-499.
  15. ^ Jensen BD, Vicente JG, Manandhar HK, Roberts SJ. “Occurrence and diversity of Xanthomonas campestris pv. campestris in vegetable brassica fields in Nepal.” Plant Disease 94 (2010): 298-305.
  16. ^ Fargier E, Manceau C. “Pathogenicity assays restrict the species Xanthomonas campestris into three pathovars and reveal nine races within X. campestris pv. campestris.” Plant Pathology 56 (2007): 805-818.
  17. ^ Carisse O, Wellman-Desbiens E, Toussaint V, Otis T. "Preventing black rot." Government of Canada. Horticultural Research and Development Centre. 1999.
  18. ^ Taylor JD, Conway J, Roberts SJ, Vicente JG. “Sources and origin of resistance to Xanthomonas campestris pv. campestris in Brassica genomes.” Phytopathology 92 (2002): 105-111.
  19. ^ United States. Department of Agriculture. U.S. Cabbage Statistics - U.S. fresh cabbage: Area, yield, production, & value, 1960-2007. May 2008.
  20. ^ Roberts SJ; Brough J; Hunter PJ (2007) Modelling the spread of Xanthomonas campestris pv. campestris in module-raised brassica transplants. Plant Pathology, 56 (0): 391-401.Modelling the spread of Xanthomonas campestris pv. campestris in module-raised brassica transplants - Roberts - 2006 - Plant Pathology - Wiley Online Library
  21. ^ Roberts SJ (2009) Transmission and spread of Xanthomonas campestris pv. campestris in brassica transplants: implications for seed health standards. In: Biddle AJ; Cockerell V; Tomkins M; Cottey A; Cook R; Holmes W; Roberts SJ; Vickers R, Seed Treatment and Production in a Changing Environment. pp 82-85.[1]
  22. ^ da Silva AC, et al. "Comparison of the genomes of two Xanthomonas pathogens with differing host specificities." Nature 417(2002):459-63.
  23. ^ Vorhölter FJ, Schneiker S, Goesmann A, Krause L, Bekel T, Kaiser O, Linke B, Patschkowski T, Rückert C, Schmid J, Sidhu VK, Sieber V, Tauch A, Watt SA, Weisshaar B, Becker A, Niehaus K, Pühler A. "The genome of Xanthomonas campestris pv. campestris B100 and its use for the reconstruction of metabolic pathways involved in xanthan biosynthesis." Journal of Biotechnology 134(2008): 33-45.
  24. ^ Qian W, Jia Y, Ren SX, He YQ, Feng JX, Lu LF, Sun Q, Ying G, Tang DJ, Tang H, Wu W, Hao P, Wang L, Jiang BL, Zeng S, Gu WY, Lu G, Rong L, Tian Y, Yao Z, Fu G, Chen B, Fang R, Qiang B, Chen Z, Zhao GP, Tang JL, He C. "Comparative and functional genomic analyses of the pathogenicity of phytopathogen Xanthomonas campestris pv. campestris." Genome Research 15.6 (2005): 757-67.
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Bacterial wilt of turfgrass

Bacterial wilt of turfgrass is the only known bacterial disease of turf. The causal agent is the Gram negative bacterium Xanthomonas campestris pv. graminis. The first case of bacterial wilt of turf was reported in a cultivar of creeping bentgrass known as Toronto or C-15, which is found throughout the midwestern United States. Until the causal agent was identified in 1984, the disease was referred to simply as C-15 decline. This disease is almost exclusively found on putting greens at golf courses where extensive mowing creates wounds in the grass which the pathogen uses in order to enter the host and cause disease.[1]

Hosts and symptoms[edit]

Creeping bentgrass (Agrostis stolonifera) and annual bluegrasses (Poa annua) are the makeup of most putting greens, as well as the preferred hosts of this pathogen. Specifically, Toronto (C-15), Seaside, and Nemisilla are the cultivars of creeping bentgrass most commonly affected.[2] The bacteria enter the plant host and interfere with water and nutrient flow, causing the plant to look drought stressed and to take on a blueish-purple color. Additionally, symptoms of bacterial wilt of turf grass include yellow leaf spots, tan or brown spots, water soaked lesions, elongated yellow leaves and shriveling of aforementioned blue or dark green leaves.Since putting greens are not a pure stand of turf, some grass blades may be resistant to the bacterium and thus remain unharmed while the surrounding turf dies, rendering the putting surface inconsistent and unsightly, especially at high-end golf courses.[3]

Disease cycle[edit]

The bacterium overwinters in diseased plants and thatch and is disseminated by water through rain splash, or mechanically by mowers, hoses, other gardening equipment, and golf shoes. The pathogen can also be present in the host at planting in infected sprigs, sod, or plugs.[4] Unlike fungi, bacterial plant pathogens are unable to wound or mechanically probe plant hosts on their own. Instead, these pathogens enter through wounds inflicted on plants through verticutting, cultivation, sand, or through natural openings such as stomates and hydathodes.[5] Once they have successfully entered and colonized the plant host, bacterial plant pathogens commandeer the nutrient and water supplies of plant cells to aid their own reproduction. In a study conducted by Zhou et. al (2013), the time between inoculation and wilt of host plants was found to depend heavily on temperature, with higher temperatures resulting in more rapid host decline. The researchers report the time between inoculation and wilting of host plants to be between 9–42 days. Bacteria can also spread to the roots of nearby plants underground. Within the plant host itself, bacteria spread by multiplying through binary fission and taking over more and more of the host by simply increasing their numbers.[6]


Prolonged periods of wetness and/or poorly drained soils, followed by warm, sunny days and cool nights constitute the optimal conditions for bacterial wilt. Thus, transitions between spring and summer, and summer and autumn are usually accompanied by increases in bacterial wilt of turf grass as these seasonal changes, especially in the early fall, bring sustained rainfalls and longer, cooler nights. Bacterial wilt of turf grass has been reported in several regions of Illinois, as well as other Midwestern states such as Michigan, Ohio, and Wisconsin. Antiserum produced from an isolate of this pathogen in Illinois was found to have reacted to another such isolate from Europe, suggesting that the pathogen was brought over to North America from Europe.[4] In fact, a survey of wheat and rye fields in Western Scotland demonstrated Xanthomonas campestris pv. graminis infection in 71% of the fields, with rye grass bearing the most infection. Meadow fescue from the same fields also demonstrated susceptibility to the pathogen under laboratory conditions, further pointing to Europe as the potential origin of this pathogen in the United States.[7]


General biocides such as copper, Junction, or ZeroTol offer a potential solution to bacterial wilt of turf grass, however such chemical control ages must be applied after every mowing which may be economically impractical and ultimately phytotoxic.[8] If bacterial wilt is present of the golf course, the best option may be to designate a mower for use on infected greens only in order to prevent the spread of the pathogen to other greens. Other viable methods include simply limiting the number of wounds the plant incurs, thereby limiting entry sites for the pathogen. A simple example would be less frequent mowing. It has also been proven that the disease is most devastating in grass cut to a length of between 1/8 and 3/16 of an inch, but less so in grass over 1/4 of an inch in length or longer, which presents an additional argument for limiting mowing. Another example is limiting sand topdressing as this is also a very abrasive technique which can create small wounds which allow entry of bacteria into the plant.[4]

A major factor complicating the control of Xanthomonas campestris pv. graminis is weather. While it is not possible to control the weather per se, a study found great decreases in pathogen efficacy at temperatures below 20 °C, suggesting that cooling measures may be effective in combating this pathogen.[6]

Ideally, resistant strains of the host plant should be used to control such a plant pathogen, however no resistant cultivars of turf grass have been identified to date. While no completely resistant cultivars exist, golf course owners can find solace in the fact that certain cultivars such as Penncross and Penneagle are more resistant to bacterial wilt and may thus reduce the need for frequent chemical applications and other cultural controls.[4] Researchers are making gains towards the identification of resistant cultivars as evidenced by the finding that variation in genetic linkage groups 1, 4, and 6 accounted for over 43% of resistance among Italian rye grass.[9]

A 1987 study found evidence of a possible biocontrol strategy for bacterial wilt of turf grass. The researchers found that antiserum to Psuedomonas fluorescens or Erwinia herbicola from hosts which have survived infections by the corresponding pathogens is capable of reducing wilt symptoms in turf grass caused by Xanthomonas campestris pv. graminis. The researchers did note, however, that while it is important to ensure the presence of a higher number of competing bacterial cells in order to reduce symptoms, one should take care to avoid over-infecting the host with a new bacterial pathogen. [10]

Further gains towards host resistance were made in 2001 when researchers found that inoculation of meadow fescue during breeding with a single aggressive strain of the bacterial wilt pathogen greatly increased resistance in offspring, thereby demonstrating the potential of selective breeding to reduce bacterial wilt pathogenesis on turf and rye grasses.[11]


The impact of bacterial wilt of turf grass was perhaps most poignant when the disease destroyed the Toronto greens at the Butler National Golf Course in Illinois, just days before the PGA Western Open was set to take place there in 1980. While bacterial wilt of turf has plagued golf courses for many years, ongoing climate change may exacerbate its prominence, as the causal agent, Xanthomonas campestris pv. graminis, prefers the persistent rainfalls and cool nights which tend to accompany weather changes.[12]


  1. ^ Dernoeden, Peter H. et al. (January 2003) Bacterial Wilt: An enigmatic annual bluegrass disease of putting greens. Golf Course Magazine.
  2. ^ Latin, Rick and Martin, Bruce. "Bacterial Decline on Creeping Bentgrass-North and South Perspectives".
  3. ^ Bacterial Wilt — Xanthomonas translucens pv. poae. Michigan State University.
  4. ^ a b c d RPD No. 414 – Bacterial Wilt and Decline of Turfgrasses. University of Illinois-Extension, Oct. 1987.
  5. ^ Fech, John C., and Roch E. Gaussoin. "The Control Center: Bacterial Wilt." Golf Course Superintendent Association of America. N.p., July 2010. Web.
  6. ^ a b Bacterial Wilt. University of Connecticut. December 3, 2013.
  7. ^ Channon, A. G.; Hissett, R. (1984). "The incidence of bacterial wilt caused by Xanthomonas campestrispvgraminisin pasture grasses in the West of Scotland". Plant Pathology 33: 113. doi:10.1111/j.1365-3059.1984.tb00594.x.  edit
  8. ^ Bacterial Wilt of Annual Bluegrass. UMass Extension, Aug. 2011. Web. December 3, 2013.
  9. ^ Studer, B; Boller, B; Herrmann, D; Bauer, E; Posselt, U. K.; Widmer, F; Kölliker, R (2006). "Genetic mapping reveals a single major QTL for bacterial wilt resistance in Italian ryegrass (Lolium multiflorum Lam.)". Theoretical and Applied Genetics 113 (4): 661–71. doi:10.1007/s00122-006-0330-2. PMID 16799808.  edit
  10. ^ Schmidt, D. (1988). "Pseudomonas fluorescens and Erwinia herbicola Reduce Wilt of Grasses Caused by Xanthomonas campestris pv. graminis". Journal of Phytopathology 122 (3): 245. doi:10.1111/j.1439-0434.1988.tb01013.x.  edit
  11. ^ Michel, V. V. (2001). "Interactions Between Xanthomonas campestris pv. graminis Strains and Meadow Fescue and Italian Rye Grass Cultivars". Plant Disease 85 (5): 538. doi:10.1094/PDIS.2001.85.5.538.  edit
  12. ^ Occurrence of Bacterial Wilt On Poa Annua and Other Turfgrasses. Michigan State University, n.d. Web. December 3, 2013..
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