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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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. Programme and Abstract Book of the 4th International Bacterial Wilt Symposium. 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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Bacterial wilt of turfgrass

Bacterial wilt of turfgrass is a disease caused by the bacterium Xanthomonas campestris pv. graminis. The cause of the disease was initially unknown, and the malady was referred to as ‘C-15 (the cultivar of creeping bentgrass known as Toronto) decline’.” (Dernoeden, et. Al.) This disease is almost exclusively found on putting greens at golf courses with a budget large enough to allow the right management practices for the bacteria to get into the plant.

Hosts and symptoms[edit]

Creeping bentgrass (Agrostis stolonifera) and annual bluegrass (Poa annua) are the two grasses affected, as they are what makes up most putting greens. “Disease is reported more frequently on a few modern cultivars. However, reports exist where more traditional cultivars are involved.” (Latin) Bacterial wilt of turf takes hold when there is an extended period of wetness followed by warm, sunny days and cool, wet nights. The bacteria get into the plant and affect the water and nutrient flow, causing the plant to look drought stressed, it appears a blueish-purple color. “Small, yellow leaf spots, streaked tan to dark brown spots, dark green, water soaked lesions, shriveled blue to dark green leaves, and yellow elongated leaves are all symptoms that have been associated with bacterial wilt.” (Michigan State) Since putting greens aren’t a pure stand of turf, some grass may be resistant to the bacteria, remaining healthy, while the turf surrounding it dies, rendering the putting surface inconsistent.

Disease cycle[edit]

"The bacterium overwinters in diseased plants and thatch and is disseminated by rain splash or flowing water, by physical transmission on mowers, hoses, other turfgrass equipment, or shoes, and by planting infected sprigs, sod, or plugs.” (Illinois) Bacteria, as is true of all other pathogens, need a host in order to cause disease. “Verticutting, cultivation and sand topdressing have the capacity to wound turf plants, creating additional disease entry points to existing stomates and hydathodes.” (Fech, Superintedent Magazine) They enter the plants and take over cells in order to take water and nutrients and multiply. In a study conducted by Zhou and Neal, “the time periods from inoculation to wilt and death of plants varied from 9-42 days, depending on temperature with higher temperature producing a more rapid response.” (Zhou) Bacteria can also move to other plants via the roots where they are touching underground, which is one way that it can spread. In the plant itself, bacteria spread by multiplying their numbers Other ways the bacteria can spread is on mowing equipment, and also foot traffic, but not as much as the mowers.


Prolonged periods of wetness and/or poorly drained soils, followed by warm, sunny days and cool nights are the optimal condition for bacterial wilt. As mentioned in the disease cycle, the bacteria enter the plants from wounds caused by maintenance practices and via natural openings. Late spring/early summer and early fall are the times when bacterial wilt comes into play due to more rainfall for prolonged wetness with warm days and cool nights.


Bacterial wilt hasn’t really been much of a problem in turf until recently, but this is due to the fact that the last three summers have been some of the hottest on record. In research trials, antibiotics have been used as a control method but they are illegal to use to that doesn’t do much good for golf courses. “General biocides such as copper (Kocide), JunctionTM (mancozeb), or ZeroTolTM (hydrogen peroxide) can reduce bacterial populations on leaf surfaces, but must be applied very frequently (after each mowing) which would result in severe phytotoxicity.” (UMass). If bacterial wilt is present of the golf course, the best option is to designate a mower for those greens in order to prevent its spread to other greens. A walk mower is the best for doing this. Another way to help with the “wounding” of the turf is to not mow greens every day, but instead change to every other and just use a roller to keep greens speed up when not mowing. Sand topdressing is a very abrasive technique and can create small wounds perfect for bacteria to enter into the plant. Higher end courses, with larger budgets, incorporate this into the routine much more often than lower end, so this will mostly apply to them. While Xanthomonas has a bad reputation of wilting the poa on putting greens, tees and fairways, what if it could be used as a way to rid putting greens of poa and establish a more pure stand of turf that can resist poa invasion in the future? A trial was done with a Japanese isolate of Xanthomonas and the results provided an interesting argument for poa control. “At 25°C/20°C (day/night) annual bluegrass wilted severely in 7–10 days, but lower temperatures caused a loss of efficacy, showing that temperature is an important factor for effective control. Inoculation tests in zoysia (Zoysia tenuifolia) greens, in the fall, with a rate of 109CFU/ml at 400 ml/m2, resulted in over 90% disease severity the following spring.” (Imaizumi)


This disease is becoming more of an issue as the climate changes and the earth experiences more of the extremes. In the past, turf managers were not experienced with bacterial wilt of turf and the disease had effects all the way into the PGA Tour. David L. Roberts of Michigan State University stated, “…disease that destroyed the 'Toronto' greens two weeks prior to the 1980 PGA Western Open at the Butler National Golf Course in Oak Brook, IL.” (Roberts) As more and more research is done about bacterial wilt of turf, turf managers will learn more about ways to manage and prevent it. Bruno Studer, in a genetic mapping study says, “Moreover, there is a lack of knowledge of the relationship between seedling resistance observed in the glasshouse and adult plant resistance in the field.” (Studer)


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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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