This is an article containing relevant information for pepper growers about Pepper mild mottle virus (abbreviated PMMoV).

For a .pdf version of this article, please click here.

Plants Pepper mild mottle virus (PMMoV) can infect ^{3, 11}

• Sweet and chilli peppers, Capsicum annuum
• Cayenne pepper, C. baccatum
• Hot bonnet peppers, C. chinense
• Tabasco or cayenne peppers, C. frutescens
• Most other domesticated or wild capsicum species
• Petunia, Petunia hybrida (asymptomatic)
• Basil, Ocimum basilicum L. (local lesions)
• Plants of mainly academic/research interest including: tobacco varieties (Nicotiana clevelandii Gray, N. debneyi, N. glutinosa, N. sylvestris, N. tabacum); Datura metel, D. stramonium, Chenopodium amaranticolor, C. quinoa

Plants PMMoV cannot infect: ^{3, 4}

• Tomato, Solanum lycopersicum L.
• Eggplant/aubergine/brinjal, Solanum melongena L.
• Cucumber, Cucumis sativus
• Beans, Phaseolus vulagris
• Many others

Symptoms in peppers ^{1, 3, 4, 7, 11}

Symptoms will vary depending upon pepper variety, virus isolate, plant age when infected, and environmental conditions. Symptom pictures of PMMoV in Carolina Reaper peppers can be found below.

Leaf symptoms:

• Mild mottle
• Mosaic (light and dark yellow or green patches) can appear 3+ weeks after infection in new apical leaves
• Necrosis (browning)
• Minor or no symptoms
• Stunting

Fruit symptoms:

• Deformed and irregular shapes
• Sunken yellow or white spots and stripes
• Premature flower and fruit abscission leading to no fruit development

PMMoV symptoms in Capsicum chinense ‘Carolina Reaper’

Note: symptoms will vary depending on numerous factors and your plants may not show these exact symptoms.

Only 4 leaves on the entire plant are showing symptoms at 2 weeks after inoculation, pictured are the most severe symptoms of a brown (necrotic) leaf pattern. This leaf falls off by the following week but you can track the lower left leaf through 10 weeks in the photos that follow.

Inoculated plant has an increase in the number of leaves with brown patterns and some drop off the plant.

Healthy plant is growing rapidly and has numerous flowers and fruit set.
PMMoV inoculated plant has foliar symptoms of mottling and necrotic (brown) patterns in new growth and is stunted. Flower buds are just starting to appear.

Healthy plant has several ~1" diameter fruit and numerous smaller fruit.
PMMoV inoculated plant is stunted, leaves are mottled and have extensive necrotic patterns. Leaves and flowers fall off and there are no fruit of any size on the infected plant.

Note yellow tone due to HPS lighting.

Yield impact

       PMMoV causes high losses because plants infected by PMMoV are often stunted and if any fruit are produced they are not saleable due to their irregular shape and color. In 2002 jalapeno peppers in Georgia, USA were infected by PMMoV which resulted in a yield loss of 50-100%.⁷
       PMMoV is distributed world-wide and can be difficult to detect early due to minor leaf symptoms.⁴ Producers of transplants should be aware of PMMoV because it is easily transmitted by infected sap and can rapidly infect an entire bench of pepper transplants if preventative cultural control methods are not practiced.

Transmission and spread of PMMoV

       PMMoV can be transmitted through infected sap spread from an infected plant to healthy plants during: grafting, transplanting, pruning, trellising, or harvesting.
       PMMoV is efficiently transmitted by seed with up to 100% seed infection rates.⁵ Although some seed lots may be heavily infested with PMMoV, even a low seed transmission rate is enough to start an epidemic if preventative cultural control methods are not practiced.

Detection

       PMMoV can be tested for on the farm by using rapid immunochromatographic dipsticks obtained online from commercial sellers or plant leaves or seeds can be mailed or dropped off at a plant disease diagnostic clinic.

Control

       The most effective control method for PMMoV is to use preventative control measures to prevent PMMoV from infecting plants because there are no curative chemicals for viral infections in plants.

Seed control of PMMoV ^{5, 9}

       Seeds without a coating can be disinfected using 2% lye (sodium hydroxide or caustic soda; NaOH) for 2 minutes at room temperature (~68°F)¹⁰ or with 10% trisodium phosphate (Na₃PO₄) for 2.5 hours at room temperature.
       Both treatments should be mixed during soaking and then rinsed thoroughly with tap water and sown immediately. These treatments may reduce germination and increase the number of abnormal seedlings so it is best to test a portion of your seed lot with these treatments to determine if any effects occur with your specific variety.⁵
       These chemical seed treatments will greatly reduce the amount of PMMoV on infested seeds but may not completely eliminate it on heavily infested seeds. These treatments will also eliminate other seed borne viral pathogens.⁹

Tool sterilization to prevent the spread of PMMoV ^{6, 8}

      Tools and glove covered hands can be sterilized by dipping in a 10% solution of bleach (use bleach with a 5.25-6% NaOCl concentration and change the solution frequently, minimally every 2 hours) or 2% Virkon S solution between each plant or variety. Disinfection is immediate, no tool soaking is required. It is recommended to dry tools and hands before handling plants as the chemicals may damage plant tissue. Sap residue must be removed from tools as it can harbor infectious virus, even after a dip. These treatments also prevent transmission of other viruses and plant pathogens.
       String used for trellising should not be reused. If wire must be used again, sterilize it by completely submerging in a 5% solution of trisodium phosphate (Na3PO4) for 10 minutes, or 0.1% caustic soda (also called lye and sodium hydroxide with a chemical formula of NaOH) for 10 minutes, or use dry heat (oven) of a minimum of 266° F (130° C) for 15 minutes.

Plant resistance to control PMMoV ²

      Resistant pepper varieties could be selected for use in the following years if PMMoV has been diagnosed-however there have been resistance breaking PMMoV strains identified that can infect and cause symptoms in peppers containing resistance genes. These four genes are called the L1-4 resistance genes and were identified from wild capsicum species. There have been no large trials to test the resistance claims that are made for pepper varieties but Cornell University has one of the most complete disease resistance charts for varieties currently on the market that can be used to choose resistant varieties.

Link here: http://vegetablemdonline.ppath.cornell.edu/Tables/TableList.htm

Look-alikes

      Other viruses cause similar symptoms to PMMoV but have different hosts and management methods. Testing and identification of the agent causing symptoms will allow for effective management solutions to be taken to reduce damage in the current growing season and the following ones as well. Mixed infections with more than one virus species present are also possible and can be identified by testing symptomatic plants.

       Reference to commercial products or trade names is made with the understanding that no discrimination is intended of those not mentioned and no endorsement is implied for those mentioned.

This work was supported by the USDA NIFA AFRI ELI project award#: 2017-67012-26090.

      This article was written for that project to inform and educate pepper growers on how to recognize, detect, and manage Pepper mild mottle virus in peppers.

All photos taken by Sara Bratsch. For non commercial use only.

      Please contact regarding all other uses including but not limited to: data for computational algorithms, books, pamphlets, articles, apps, websites, etc.

Cite this article:

Bratsch, Sara. 2018. "Pepper mild mottle virus". Web article. (date accessed). http://plantpathlesstraveled.com/pepper-mild-mottle-virus

Citations

1. Adkins, S., Lamb, E. M., Roberts, P. D., Gooch, M. D., Breman, L., and Shuler, K. D. 2001. Identification of Pepper mild mottle virus in commercial bell pepper in Florida. Plant Disease. 85:679.
2. Antignus, Y., Lachman, O., Pearlsman, M., Maslenin, L., & Rosner, A. 2008. A new pathotype of Pepper mild mottle virus (PMMoV) overcomes the L 4 resistance genotype of pepper cultivars. Plant Disease, 92(7), 1033-1037.
3. Feldman, J. M., & Oremianer, S. (1972). An unusual strain of tobacco mosaic virus from pepper. Journal of Phytopathology, 75(3), 250-267.
4. Green, S. K. 2003. Pepper mild mottle virus. Pages 32-33 in: Compendium of Pepper Diseases. K. Pernezny, P. D. Roberts, J. F. Murphy, and N. P. Goldberg, eds. American Phytopathological Society, St. Paul, MN.
5. Jarret, R. L., Gillaspie, A. G., Barkley, N. A., & Pinnow, D. L. 2008. The occurrence and control of pepper mild mottle virus (PMMoV) in the USDA/ARS Capsicum germplasm collection. Seed technology, 26-36. https://naldc.nal.usda.gov/download/26970/PDF
6. Li, R., Baysal-Gurel, F., Abdo, Z., Miller, S. A., & Ling, K. S. (2015). Evaluation of disinfectants to prevent mechanical transmission of viruses and a viroid in greenhouse tomato production. Virology journal, 12(1), 5.
7. Martínez-Ochoa, N., Langston, D. B., Mullis, S. W., & Flanders, J. T. 2003. First report of Pepper mild mottle virus in Jalapeno pepper in Georgia. Plant health progress, 12, 1-2.
8. Nitzany, F. E. (1960). Tests for tobacco mosaic virus inactivation on tomato trellis wires. Ktavim, 10, 59-61.
9. Rast A.T.B., Stijger C.C.M.M. (1987). Disinfection of pepper seed infected with different strains of capsicum mosaic virus by trisodium phosphate and dry heat treatment. Plant Pathology 36: 583-588.
10. Salamon P. Kaszta M. (2000). Investigation on the transmission of some Tobamoviruses by pollen and seed in pepper (Capsicum annuum L.). In: 8th International Pollination Symposium, Mosonmagyarovar, Hungary, 10–14 July, 2000. International Journal of Horticultural Science, 6: 127 –131
11. Wetter, C., Conti, M., Altschuh, D., Tabillion, R., & Van Regenmortel, M. H. V. (1984). Pepper mild mottle virus, a tobamovirus infecting pepper cultivars in Sicily. Phytopathology, 74(4), 405-410.

Life cycle

       Some slime molds like the dog vomit slime mold are visible in their plasmodium stage because it is brightly colored. However, the turfgrass slime mold is clear or watery-white during this life stage and is hard to observe.⁶ For the larger dog vomit slime mold it may be possible to observe where it has been as the mold will leave a trail of slime behind it.
       The slime mold plasmodium life stage is different from most larger organisms as it is simply a ball of cellular fluids, nuclei, and other organelles (also called the protoplasm) contained by only a stretchable cell membrane.⁶ Most larger organisms consist of numerous cells with the protoplasm broken into separate compartments by cell walls and membranes.
       Under proper conditions the plasmodium will form fruiting bodies which release spores.⁶ Physarum cinereum uses blades of grass as a tall structure to release spores from and is not eating the grass.⁷ The microscopic spores are spread by the wind which makes a tall grass blade a perfect spot to release spores from. These spores allow the slime mold to survive unfavorable conditions and spread to new food sources.
       The sporulating life stage is commonly observed for the turf grass slime mold.⁷

Microscopic view of slime mold, Physarum cinereum, spores in structures on the surface of the grass.

Life cycle continued⁶

When appropriate growing conditions return, the spores will break open and release cells called protoplast which develop flagella. These cells will move towards other cells and will fuse to form myxamoebae which then grow and becomes a new plasmodium. Although slime molds produce spores, they are not fungi and are actually classified as Protists.

Habitat

      In addition to turf grass different species of slime mold can be observed on wood chips, timber retaining walls, fallen trees, leaves, and even fish tanks.^{2, 3, 6, 7} Slime molds can be found throughout the world, often appearing after rains, in damp areas with dead material to eat. In this case the slime mold was likely consuming the dead thatch during the plasmodium stage and then used the blades of grass as aerial structures to release spores from.⁷

Control ⁷

      There are no control methods for slime molds as they do not damage the grass, clover, or other plants they may cover. If desired, the slime mold could be removed by brushing the affected leaves or mowing to remove the most heavily infested leaves. Spraying with water is not recommended because that will favor the development of new plasmodium from the newly released spores.

Other interesting information

      If you are so inclined, you can actually purchase a kit containing a slime mold to keep as a pet or use for an educational tool.
      Slime molds have gained popularity with videographers so it is possible to find videos and documentaries on various species of slime molds.
      Computer scientists have made a simple biological computer with logical circuits using living slime mold tubes, instead of solid silicone, to process information.¹
       Another area of study with slime molds is their ability to solve mazes and optimize routes.^{4, 5}

All photos taken by Sara Bratsch. For non commercial use only.

Please contact regarding all other uses including but not limited to: data for computational algorithms, books, pamphlets, articles, apps, websites, etc.

Cite this article:

Bratsch, Sara. 2018. "Turf grass slime mold". Web article. (date accessed). http://plantpathlesstraveled.com/turf-grass-slime-mold

Citations

1. Adamatzky, A., & Schubert, T. (2014). Slime mold microfluidic logical gates. Materials Today, 17(2), 86-91.
2. Hidden Forest. (2010). Species: Physarum cinereum. http://www.hiddenforest.co.nz/slime/family/physaraceae/physa16.htm
3. Kasper. (2013). Slime mold in the Aquarium. https://fishtankdk.blogspot.com/2013/01/slime-mold-in-aquarium.html
4. Nakagaki, T., Yamada, H., & Tóth, Á. (2000). Intelligence: Maze-solving by an amoeboid organism. Nature, 407(6803), 470.
5. Nakagaki, T., Kobayashi, R., Nishiura, Y., & Ueda, T. (2004). Obtaining multiple separate food sources: behavioural intelligence in the Physarum plasmodium. Proceedings of the Royal Society of London B: Biological Sciences, 271(1554), 2305-2310.
6. Shen, Y. F. (1964). A study on the life cycle of Physarum cinereum grown in culture. Taiwania, 10(1), 63-71.
7. Vargas, J. M. (1993). Management of turfgrass diseases. CRC Press. 65-66.

Plants Tomato mosaic virus has been shown to infect ^{2, 5, 6, 8, 15 * +}

      Tomato mosaic virus (abbreviated ToMV) can infect a wide range of economically important plant species including: sweet, chilli, and hot peppers (Capsicum annuum, C. frutescens); quinoa (Chenopodium quinoa); tomato (Solanum lycopersicon L.); tobacco (Nicotiana benthamiana, N. clevelandii, N. glutinosa, N. megalosiphon, N. rustica, N. tabacum); tomatillo and cape gooseberry/ground cherry (Physalis ixocarpa, P. peruviana); eggplant (Solanum melongena); and potato (S. tuberosum).
       Weedy plant species that have been identified with ToMV infection include: redroot pigweed (Amaranthus retroflexus); common lamb’s quarters and nettle leaf goosefoot (Chenopodium album ssp. amaranticolor, C. murale); physalis (P. floridana); and black nightshade (Solanum nigrum).
Ornamental plants that can be infected with ToMV include: angel’s trumpet (Datura metel); petunia (Petunia x hybrida); and red bitter berry (Solanum giganteum).
       Fruits including cherry (Prunus avium and P. cerasus), pear (Pyrus communis), apple (Malus sylvestris), and grape (Vitis vinifera) can also be infected.

* ToMV has been inoculated into healthy plants of these species and infected them, field samples may have tested positive for ToMV. See citations for more details.
+ This does not exclude plant species not listed-these are the only species research has been completed and published on.

Plants Tomato mosaic virus has not been shown to infect ^{2, 5, 6, 8, 15 ** +}

      ToMV has not been shown to infect muskmelon (Cucumis melo); squashes, pumpkins and zucchini (Cucurbita pepo spp.); green bean (Phaseolus vulgaris); and cowpea (Vigna unguiculate).

** ToMV has not been isolated from field samples or infected laboratory inoculated plants of these species.
+ This does not exclude plant species not listed-these are the only species research has been completed and published on.

Symptoms in tomatoes ^{5, 6, 8, 15}

      This virus causes a wide variety of symptoms dependent upon: plant variety, temperature, light intensity, plant age, virus strain, and more. It can also show no symptoms in many plants including tomatoes.

Leaf symptoms:

• thin, thread-like leaves
• leaf malformation
• mosaic (mottled areas of light green, dark green, yellow, or white)
• brown or yellow spots, stripes, or line patterns on leaves or stems
• stunting
• wilting
• complete plant death

Fruit symptoms:

• yellow spots or stripes that do not ripen
• internal browning
• yellow or brown rings
• brown circles or spots
• line patterns

Yield impact

      Tomato mosaic virus can cause yield losses of 20-100% due to reduced fruit set or symptoms that make the fruit unsaleable.⁴ Infected plants should be pulled, attempting to get the majority of the roots, and burnt or otherwise removed from the growing area. ToMV can survive and be infectious in tomato root debris at 47 inches (120 cm) deep in fallow soil after 22 months and for over 2 years in soil under black plastic.⁴ It can also survive winter conditions in soil in Wisconsin.⁹ ToMV is a very stable virus, dried leaf tissue that was stored at room temperature was still infectious after 24 years.⁷ It is very important to test and identify ToMV infected plants so they can be removed promptly, reducing the amount of virus inoculum left in the growing area.

Spread

      ToMV is easily spread by any contact that wounds or creates microwounds in plant leaves, stems, or roots including transplanting, pruning, trellising, harvesting, and bumble bee pollination.^{5, 13, 15} Chewing insects such as grasshoppers and locusts can also spread the virus from infected plants to other plants including weeds.¹⁶ Other insects such as aphids have been shown to spread ToMV by breaking leaf hairs.⁵ Water used in hydroponics can also spread ToMV so it must be sterilized if recirculated.⁵
      ToMV can have up to a 94% seed transmission rate although less than a 1% seed transmission rate is enough to serve as inoculum for an epidemic due to the ease of mechanical transmission.^{5, 6}

Detection

      ToMV can be tested for on the farm by using rapid immunochromatographic dipsticks obtained online from commercial sellers or infected plant tissue can be mailed or dropped off at a plant disease diagnostic clinic. Search online to find your local plant disease diagnostic clinic or consult your extension agent for your closest clinic.
      Tomato mosaic virus is closely related to Tobacco mosaic virus, Tomato brown rugose fruit virus, Tomato mottle mosaic virus, and other Tobamoviruses which have been reported causing disease in Solanaceae crops.^{14, 1} Antibody based tests for any of these viruses will react with other viruses in this family (Tobamovirus).¹ Specific RT-PCR and or sequencing of a portion of the virus genome is required to obtain a species level identification.¹ These can be completed by a plant disease diagnostic clinic.

Control

Seed treatment ^{3, 5}

      The major control method for ToMV is to use certified virus free seed or to disinfest seeds yourself. The hot water bath method does not reduce the amount of ToMV on tomato seeds.³
      Seeds should be soaked with a 10% solution of trisodium phosphate=trisodium orthophosphate=sodium phosphate tribasic (Na₃PO₄) for 30 minutes and mixed several times, rinsed and dried. Note this solution should be made with warm water (this chemical does not dissolve easily so warm water makes it faster) and the trisodium phosphate should be stored in an air tight container to prevent it from solidifying (otherwise it may turn into a hard brick before it is used again). After rinsing and drying, follow with a 1-2 minute soak in a 1:4 dilution of bleach, using 1 gallon of diluted bleach per pound of seed. Use bleach with 5.25-6% sodium hypochlorite concentration (NaOCl) and do not reuse. Pour seed into a fine mesh and rinse thoroughly with cool water. After air drying the seeds can be coated with fungicides following manufacturer’s recommendations to control seed rot and damping-off.³ Trisodium phosphate and bleach will control anthracnose, bacterial spot, and will greatly reduce or eliminate seed coat carried ToMV.³ A trisodium phosphate soak alone will greatly reduce seed coat carried ToMV only.³
      ToMV can also be transmitted inside of the seed up to 25%, which is not destroyed by the soaking methods.⁵ ToMV is not always transmitted inside of the seed, so it is important to treat your seed to reduce the potential for seed to be a source of virus introduced to your growing area.⁵ Scouting of plants and testing to identify the biotic or abiotic issue causing symptoms is very important, even if using treated seeds.
      Experiments are on-going to determine a protocol for using sonication, a non-chemical seed disinfesting method. This article will be updated with a link to that protocol when it is completed.

Tool sterilization ^{10, 12}

      Always sterilize tools and glove covered hands by dipping in a 10% solution of bleach with a 5.25-6% NaOCl concentration (change frequently, minimally every 2 hours) or 2% Virkon S between each plant.¹⁰ Disinfection is immediate, no tool soaking is required. It is recommended to dry tools and hands before handling plants as the chemicals may damage plant tissue. Sap residue must be removed from tools as it can harbor infectious virus, even after a dip. These treatments also prevent transmission of Tobacco mosaic virus, Pepino mosaic virus, and Potato spindle tuber viroid.¹⁰
      Minimally tools and hands should be sterilized in between varieties to help reduce the chance of the entire crop becoming infected.
      String used for trellising tomatoes should not be reused. If wire must be used again, sterilize it by completely submerging in a 5% solution of trisodium phosphate (Na₃PO₄) for 10 minutes, or 0.1% caustic soda (also called lye and sodium hydroxide with a chemical formula of NaOH) for 10 minutes, or use dry heat of 266° F (130° C) for 15 minutes.¹²

Resistance ¹¹

      Resistant tomato varieties could be selected for use in the following years if ToMV has been diagnosed-however there have been resistant breaking ToMV strains identified that can cause symptoms in tomatoes containing resistance genes.¹¹ ToMV and Tobacco mosaic virus (TMV) have different resistance genes so it is best to obtain a diagnosis to the species level to make choosing resistant tomato varieties effective. There have been no large trials to test the resistance claims that are made for tomato varieties but Cornell University has one of the most complete disease resistance charts click here to follow the link to their page for varieties currently on the market that can be used to choose resistant varieties.
      Resistant tomato varieties in many cases can still be infected by and allow the reproduction of (and an increase in concentration) of both ToMV and TMV but do not show symptoms.

Look-alikes

      Herbicide damage can also resemble virus symptoms but some diagnostic laboratories can test for common herbicides-consult them for specific sampling and shipping details.
      Other viruses and viroids can also cause similar symptoms to ToMV but have different management methods. Testing and identification of the agent causing symptoms will allow for effective management solutions to be taken to reduce damage in the current growing season and the following ones as well.

Reference to commercial products or trade names is made with the understanding that no discrimination is intended of those not mentioned and no endorsement is implied for those mentioned.

This work was supported by the USDA National Institute of Food and Agriculture AFRI ELI Post-doctoral fellowship project award#: 2017-67012-26090.

This article was written for that project to inform and educate tomato growers on how to recognize, detect, and manage Tomato mosaic virus in tomatoes.

All photos taken by Sara Bratsch. For non commercial use only.

Please contact regarding all other uses including but not limited to: data for computational algorithms, books, pamphlets, articles, apps, websites, etc.

Cite this article:

Bratsch, Sara. 2017. "Tomato mosaic virus in tomatoes". Web article. (date accessed). http://plantpathlesstraveled.com/tomato-mosaic-virus

Citations

1. Adams, M.J., Adkins, S., Bragard, C., Gilmer, D., Li, D., MacFarlane, S.A., Wong, S-M., Melcher, U., Ratti, C., Ryu, K.H., and ICTV Report Consortium. 2017, ICTV Virus Taxonomy Profile: Virgaviridae. Journal of General Virology, 98: 1999-2000.
2. Aghamohammadi, V., Rakhshandehroo, F., Shams-Bakhsh, M., & Palukaitis, P. (2013). Distribution and genetic diversity of tomato mosaic virus isolates in Iran. Journal of plant pathology, 339-347.
3. Babadoost, M. (1992). University of Illinois Extension. RPD No. 915
4. Broadbent, L., Read, W., Last, F. (1965). The epidemiology of tomato mosaic: Persistence of TMV-infected debris in soil and the effects of soil partial sterilization. Ann. Appl. Biol. 55:471-83.
5. Broadbent, L. (1976). Epidemiology and control of tomato mosaic virus. Annual review of Phytopathology, 14(1), 75-96.
6. Brunt, A.A., Crabtree, K., Dallwitz, M.J., Gibbs, A.J., Watson, L. & Zurcher, E.J. (eds.). (1997). `Plant Viruses Online: Descriptions and Lists from the VIDE Database. http://sdb.im.ac.cn/vide/descr832.htm
7. Caldwell, J., 1959. Persistence of tomato acuba mosaic virus in dried leaf tissue, Nature (London) 183: 1142.
8. Hollings, M., & Huttinga, H., (1976). Tomato mosaic virus, CMII AAB Descriptions of Plant Viruses No. 156.
9. Hoggan, I., & Johnson, J. (1936). Behavior of the ordinary tobacco mosaic virus in the soil. J. Agr. Res, 52, 271-294.
10. Li, R., Baysal-Gurel, F., Abdo, Z., Miller, S. A., & Ling, K. S. (2015). Evaluation of disinfectants to prevent mechanical transmission of viruses and a viroid in greenhouse tomato production. Virology journal, 12(1), 5.
11. Luria, N., Smith, E., Reingold, V., Bekelman, I., Lapidot, M., Levin, I., ... & Ezra, N. (2017). A new Israeli Tobamovirus isolate infects tomato plants harboring Tm-22 resistance genes. PloS one, 12(1), e0170429.
12. Nitzany, F. E. (1960). Tests for tobacco mosaic virus inactivation on tomato trellis wires. Ktavim, 10, 59-61.
13. Okada, K., Kusakari, S. I., Kawaratani, M., Negoro, J. I., Satoshi, T. O., & Osaki, T. (2000). Tobacco mosaic virus is transmissible from tomato to tomato by pollinating bumblebees. Journal of General Plant Pathology, 66(1), 71-74.
14. Sui, Xuelian, et al. "Molecular and Biological Characterization of Tomato mottle mosaic virus and Development of RT-PCR Detection." Plant Disease 101.5 (2017): 704-711.
15. Van Regenmortel, M. H., & Fraenkel-Conrat, H. (Eds.). (1986). The Plant Viruses: The Rod-shaped Plant Viruses. Springer Science & Business Media.
16. Walters, H. J. (1951). Grasshopper transmission of three plant viruses. Science, 113(2924), 36-37.

Introduction

Poinsettias, also known by their scientific name of Euphorbia pulcherrima Wild, are native to tropical Mexico, Belize, Guatemala, El Salvador, Honduras, Nicaragua, and Costa Rica. They were grown for their flowers which bloomed during December-January, dyes, and latex. It was not until 1825 that poinsettias were introduced to the USA by Joel Poinsette, who the common name of poinsettia (used in the USA) is derived from. Poinsettias were originally sold mainly as cut flowers due to the fact that the plants grew quite large and had limited branching (10, 12).

A ~6 foot tall poinsettia in Costa Rica blooming on one year’s growth. Note the lack of branching and thin leaves at the top (the "flowers") compared to modern varieties.

Variety development

Horticulturalists rapidly developed new varieties of poinsettias and today they are grouped into two types. One type is used for flowers and resembles the wild variety that has limited branching and can grow into a shrub. The cut flower poinsettia was the most common variety grown before the 1960’s (6).

The most commonly sold type today is used as a potted plant and is dwarfed in size and nicely branched to provide a very floriferous and attractive plant. The first report of a shorter poinsettia was in 1923 from the USA. In 1945 another greatly improved poinsettia was introduced again from the USA that had even better branching and flowering. In 1967 a superior variety was introduced from Norway which had an even greater amount of branching than observed in previous cultivars and retained its leaves much longer than previous varieties (15).

Greenhouse grown wild type Euphorbia pulcherrima plant stretching to over 10 feet tall in the greenhouse.

Early observations

Plant breeders would routinely treat their new poinsettia varieties with heat or tissue culture techniques to eliminate any viruses that had been introduced during the breeding process (8). However, they noted something critical to their new varieties would be eliminated after this virus cleaning process: the increased branching and dwarfing observed in the plant before treatment was eliminated and did not return. In 1983 a paper describing the ability to restore the branching habit by grafting a clean poinsettia after virus treatment onto a branching poinsettia for a rootstock was published (14).

Other early studies into what was causing the dwarfing and increased branching of some poinsettias suggested that Poinsettia mosaic virus or poinsettia cryptic virus were causing the symptoms, however the viruses could also be found in the taller wild varieties (7, 8, 9, 4, 5).

Potted poinsettias used for holiday decoration. Note the small stature and branching making a very floriferous plant compared to the wild type.

Identification of symptom causing microbe

No other microbe was observed until 1995 when a group of scientists working with the commercial horticulture industry sequenced pieces of DNA from a new phytoplasma they called Poinsettia branch inducing phytoplasma (PoiBI) (11). This group also showed it was possible to transmit the phytoplasma to a healthy (non-branchy) poinsettia and it would induce the plant to begin branching (10, 11).

Phytoplasmas are a group of microbes that are similar to bacteria. Unlike bacteria, phytoplasmas do not have a cell wall and cannot be cultured in media. Phytoplasmas require a living plant host to survive and can be transmitted by grafting, a parasitic plant called dodder, or their specific insect vector (leafhoppers, planthoppers, and psyllids) (2). Similar to bacteria, phytoplasmas can be killed with antibiotics and when branching poinsettias were treated with antibiotics this caused the plants to lose their free-branching habit (3).

Further molecular based research has shown that all branching and dwarfed poinsettias have at least one type of phytoplasma present in them, with some species of phytoplasma that are not closely related to the PoiBI species (1, 10, 13). At this time it is unknown how the novel free branching poinsettia varieties were infected by these phytoplasmas. Many plant breeders also utilized mutation breeding (chemical or radiation) to produce new poinsettia varieties which would also cause mutations in the phytoplasma(s) present (12). This makes it difficult to identify the specific population of phytoplasmas that cause the attractive branching of potted poinsettias today, and it is likely that there are several different groups of phytoplasmas that can cause dwarfing and branching in poinsettias (12).

Although many plant diseases have a bad reputation, Poinsettia branch inducing phytoplasmas are a highly desired plant pathogen for poinsettia breeders and buyers.

Potted poinsettias used for holiday decoration. Note the small stature and branching making a very floriferous plant compared to the wild type.

All photos taken by Sara Bratsch. For non commercial use only.

Please contact regarding all other uses including but not limited to: data for computational algorithms, books, pamphlets, articles, apps, websites, etc.

Cite this article:
Bratsch, Sara. 2017. "Poinsettia Phytoplasmas". Web article. (date accessed). https://www.plantpathlesstraveled.com/poinsettia-phytoplasmas/

Citations

1. Abad, I., Randal, J.A. and Moyer, J.W. 1997. Genomic diversity and molecular characterization of poinsettia phytoplasmas. Phytopathology 87:S1.
2. Bertaccini, A., & Duduk, B. 2010. Phytoplasma and phytoplasma diseases: a review of recent research. Phytopathologia mediterranea, 48(3), 355-378.
3. Bradel, B.G., Preil, W., and Jeske, H. (2000). Remission of the free-branching pattern of Euphorbia pulcherrima by tetracycline treatment. Journal Of Phytopathology 148, 587-590.
4. Dole, J. M., Wilkins, H. F., and Desborough, S. L. 1993. Investigation on the nature of a graft-transmissible agent in poinsettia. Can. J. Bot. 71:1097-1101.
5. Dole, J. M., Wilkins, H. F. 1994. Graft-transmissible branching agent. The scientific basis of poinsettia production. Strømme E., ed. The Agricultural University of Norway. 121: 45-48.
6. Ecke Jr., P., Matkin, O. A., and Hartley, D. E. 1990. The Poinsettia Manual, 3rd ed. Paul Ecke Poinsettias, Encinitas, CA.
7. Fulton, R.W. and Fulton, J.L. 1980. Characterization of a tymo-like virus in poinsettia. Phytopathology 70:321-324.
8. Koenig, R. and Lesemann, D.E. 1980. Two isometric viruses in poinsettia. Plant Disease 64:782-784.
9. Koenig, R., Lesemann, D. E., and Fulton, R. W. 1986. Poinsettia mosaic virus. AAB Descriptions of Plant Viruses, No. 311.
10. Lee, I.-M., Tiffany, M., Gundersen, D. E., and Klopmeyer, M. 1995. Phytoplasma infection: A beneficial factor for production of commercial branching poinsettia cultivars? Phytopathology 85:1179.
11. Lee, I.-M., Klopmeyer, M., Bartoszyk, I. M., Gundersen-Rindal, D. E., Chou, T.-S., Thomson, K. L. and Eisenreich, R. 1997. Phytoplasma induced free-branching in commercial poinsettia cultivars. Nature Biotechnology 15:178-182.
12. Nicolaisen, M. 2001. phytoplasma with the focus on Euphorbia pulcherrima and other ornamental plants’.
13. Pondrelli, M., Caprara, L., Bellardi, M. G., & Bertaccini, A. (2000, May). Role of different phytoplasmas in inducing Poinsettia Branching. In X International Symposium on Virus Diseases of Ornamental Plants 568 (pp. 169-176).
14. Stimart, D. P. 1983. Promotion and inhibition of branching in poinsettia in grafts between self-branching and non-branching cultivars. J. Amer. Soc. Hort. Sci. 108:419-422.
15. Strømme, E. 1994. The Scientific basis of poinsettia production. The Agricultural University of Norway, Aas-NLH, Norway.

History

Rose rosette disease (RRD) was first recorded in 1940-1941 in Manitoba Canada, and California, Nebraska, and Wyoming USA causing symptoms of excessive shoots, thorns, and red pigmentation on roses (3). This disease was of particular interest for use as a potential biocontrol agent for the invasive multiflora rose. Multiflora roses were promoted for use from the 1930’s-1970’s as an attractive living fence to control soil erosion and provide food for animals (rose hips and leaves) (7). Multiflora roses were quite hardy and prolifically reproduced by both seed, crown growth, and from branches that bent over and rooted severely reducing pasture and park quality. Physical removal, herbicides, and overgrazing by goats are expensive control measures so a method that would kill the plants without requiring a lot of physical labor or chemicals was desired (7).

RRD was studied for its use as a biocontrol of multiflora roses and was found to kill multiflora roses in 2-3 years for single crowned plants and 4-5 years for multicrowned plants (7). RRD was found to infect only wood’s rose, multiflora rose, macartney rose, hybrid tea rose (and all other cultivated roses including shrub, floribunda, grandiflora, miniature), and sweetbriar and was recommended for use to control multiflora roses (7). This may be surprising because of the high popularity of cultivated roses today, however it is only with the release of the Knock Out® shrub rose line in 2000 that consumers embraced the floriforous, disease resistant, cold hardy, heat resistant new generation of roses buying 4 million Knock Out® roses a year for the next four years (10) (followed by many more shrub roses as new varieties were released). Prior to the Knock Out® shrub rose line the majority of cultivated roses required pesticide sprays to control various pathogens, special pruning, and protection to survive winter.

The organism causing RRD was not described until 2011 when Rose rosette virus had the genome sequenced and in 2015 the experiments were completed to show it caused RRD (8, 6). RRV can be found throughout the USA, Canada, and was recently identified in India (3, 9, 4).

Transmission

RRV has been shown to be transmitted by the eriophyid mite, Phyllocoptes fructiphilus, and through grafting (1, 2). This species of mite is 140-170 micrometers (0.0055-0.0067 inches) long by 43 micrometers (0.0017 inches) wide and is found in rose shoots and buds so is difficult to observe without magnification (2). These tiny mites are thought to be passively moved by air currents or humans (2).

Studies conducted in the field in West Virginia revealed that plants infected by eriophyid mites carrying RRV took from 30-279 days to express symptoms (1, 2).

Symptoms

Early RRV symptoms include mosaic leaf patterns. Later symptoms include excessive thorns on canes, high levels of red pigmentation in canes and leaves, excessive lateral shoot growth, and dense masses of shoots (witches’ brooming). RRV infections will kill infected plants in 2-5 years (7).

A Knock Out® rose with healthy growth on the left and RRV infected growth showing excess red pigmentation and witches’ brooming of a flower shoot.

A “White Smith Perish” rose infected with RRV showing excessive lateral shoot growth and witches’ brooming.

A Knock Out® rose cane with leaves removed to show excessive thorniness and red pigmentation.

Not symptoms: new growth

Many varieties of roses have been released that have very dark red or purple new growth that fades to green within a few weeks. If the rose variety is unknown check for the presence of excessive thorns among the reddened foliage. RRV infected roses with reddened tissue will not fade in color.

A Knock Out® rose with new red growth. This plant is healthy and RRV negative.

Not symptoms: herbicide exposure

Accidental herbicide exposure can cause distorted, RRD-like symptoms that will disappear with later growth. If there are any other plants that have odd growth it's worth investigating if a neighbor, lawn care company, or city/county sprayers were out spraying chemicals. Symptomatic tissue can be submitted to an herbicide testing lab to get confirmation that herbicide exposure caused the symptoms.

Diagnosis

Current testing methods to diagnose RRV are based on amplifying a portion of the virus genome by reverse transcription polymerase chain reaction (RT-PCR) (5). This molecular test can be conducted by a plant disease clinic or diagnostic lab. Work is on-going to produce a rapid, protein based method for use in the field or garden.

Management

Large and valuable rose collections should be observed frequently for symptoms of RRD. Plants showing symptoms can be submitted to a plant disease clinic or diagnostic lab for testing. There is no cure for a plant virus infection and the plant must be destroyed. Infected plants should be bagged (to prevent infected mites from escaping) and removed (including roots) from the garden. Pruning symptomatic portions out of a bush does not cure it as the plant is infected systemically (from the roots to the leaves). Any neighboring roses should be watched for the development of symptoms and removed.

Work is currently ongoing to determine if any cultivated rose varieties are resistant to RRV. When choosing new rose plants for your garden chose roses that have tested negative for RRV or have no symptoms.

All photos taken by Sara Bratsch. For non commercial use only.

Please contact regarding all other uses including but not limited to: data for computational algorithms, books, pamphlets, articles, apps, websites, etc.

Cite this article:

Bratsch, Sara. 2017. "RRV". Web article. (date accessed). http://plantpathlesstraveled.com/RRV/

Citations

1. Allington, W. B., R. Staples and G. Viehmeyer. 1968. Transmission of rose rosette virus by the eriophyid mite Phyllocoptes fructiphilus. J. Econ. Entomol. 61:1137-1140.
2. Amrine Jr, J. W., Hindal, D. F., Stasny, T. A., Williams, R. L., & Coffman, C. C. (1988). Transmission of the rose rosette disease agent to Rosa multiflora by Phyllocoptes fructiphilus (Acari: Eriophyidae). Entomological news (USA). 99:239-252.
3. Conners, L. 1941. Twentieth Annual Report of the Canadian Plant Report Survey, 1940. 98.
4. Chakraborty, P., Das, S., Saha, B., Karmakar, A., Saha, D., & Saha, A. (2017). Rose rosette virus: An emerging pathogen of garden roses in India. Australasian Plant Pathology, 46(3), 223-226.
5. Dobhal, S., Olson, J. D., Arif, M., Suarez, J. A. G., and Ochoa-Corona, F. M. 2016. A simplified strategy for sensitive detection of Rose rosette virus compatible with three RT-PCR chemistries. J.Virol. Methods.232:47-56.
6. Di Bello, P. L., Ho, T., & Tzanetakis, I. E. (2015). The evolution of emaraviruses is becoming more complex: seven segments identified in the causal agent of Rose rosette disease. Virus research, 210, 241-244.
7. Epstein, A. H., & Hill, J. H. (1998). Status of rose rosette disease as a biological control for multiflora rose. Plant disease, 83(2), 92-101.
8. Laney, A.G., Keller, K.E., Martin, R.R. and Tzanetakis, I.E. 2011. A discovery 70 years in the making: characterization of the Rose rosette virus. J. Gen. Virol. 92:1727-1732.
9. Martin, C.W. 2014. Rose Rosette Disease and the Impacts on Propagation©. Acta Hortic. 1055:319-321.
10. Pemberton, H. B., and Karlik, J. F.2013. A Recent history of changing trends in USA garden rose plant sales, types, and production methods.VI International Symposium on Rose Research and Cultivation. 1064:223-234.

Introduction

TSWV is not a recently described plant pathogen, the first formal publication of the disease was observed on tomatoes in 1915 in Australia (Brittlebank, 1919). TSWV belongs to the virus genus Tospovirus (INSV, impatiens necrotic spot virus, also belongs to this virus group) (Pappu et al., 2009). TSWV utilizes a small insect called thrips to spread from plant to plant. TSWV is a common viral pathogen found in greenhouses and field crops (Pappu, 2012).

TSWV is now present throughout the world along with its thrip vectors (Oliver & Whitfield, 2016). It has been shown to infect over 1,090 plant species from 15 families of monocots, 69 families of dicots, and one family of pteriodophytes (Parrella et al., 2003). These species include such plants as numerous weeds; tomatoes, peppers, potatoes, lettuce, peanuts; and flowering plants including chrysanthemum, petunia, impatiens, and zinnia (Pappu, 2012).

Symptoms

TSWV causes a wide range of symptoms depending upon environmental growing conditions (temperature), age a plant was infected, the genotype of plant, and virus isolate. Young tomatoes may wilt and die, while tomato plants infected later in their life can show brown leaf spots, purple/bronzing of leaves, or chlorotic patterns and ringspots on the fruit. Depending on the strain of TSWV infecting tomatoes they may also show brown streaking along stems and along leaf veins. Symptoms expressed by ornamental plants also vary greatly as the following photos show.

Chrysanthemum infected with TSWV with symptoms of red splotches on the leaves.

A flat of zinnia infected with TSWV showing symptoms of necrotic rings on lower leaves and distorted, yellowed top leaves.

Tomato fruit from a plant infected with TSWV with characteristic symptoms of yellow rings. Fruit can also have necrotic (brown) rings.

Transmission

Several species of thrips in the order Thysanoptera can transmit TSWV (Mound 1996). The western flower thrip (Frankliniella occidentalis) is a frequent vector in the USA and Canada (Rosello et al 1996; Allen & Broadbent, 1986).
TSWV is transmitted by thrips in what is called a persistent propagative manner. After a thrip has fed on an infected plant, the virus will pass through its digestive system into the circulatory system. The virus particles are then circulated into the salivary glands where TSWV replicates and can be transmitted to other plants the thrip feeds on. Since the virus is replicating in the insect vector, an infected insect can transmit the virus until it dies. Depending on thrip species it can take as little as 15 minutes of feeding on virus infected plants for them to acquire the virus (Sakuma, 1962). Thrips can only acquire the virus when they are larvae, after pupating into their adult form they cannot acquire and transmit TSWV (Wijkamp, 1993). Both adults and young thrips can transmit TSWV so the presence of both should be monitored.

TSWV is not easily mechanically transmissible and is also not transmitted by seed (Pappu, 2012).

Testing

Symptomatic plants can be tested for the presence TSWV in the field with an immunostrip/lateral flow test or can be sent into a plant diagnostic clinic.

Control

Control methods to limit losses by TSWV infections rely upon good management practices including plant resistance, vector control, and quarantining new arrivals. Resistant varieties of tomatoes, peppers, and peanut have been developed and can be used to limit TSWV losses. Plant resistance to TSWV varies for different strains of TSWV so varieties with multiple resistance genes should be used (Mandal et al., 2006). Greenhouses receiving live transplants should quarantine new arrivals to check for thrips (which may be carrying TSWV) or for the development of symptoms. Thrip levels should be monitored with sticky traps or physically examining plants. When needed insecticides should be used carefully to ensure the thrip population is reduced after application.

Production areas outside of enclosed greenhouses should also quarantine new transplants to ensure no symptoms develop. Silver reflective mulch has been shown to keep thrip populations low, decreasing the incidence of TSWV infected tomato plants (Riley & Pappu, 2004). Weeds and native plants should be managed as 113 plant species native to Ontario (in 35 plant families) can be infected by TSWV and act as a source of virus to infect crops (Stobbs et al., 1992).

Plants cannot be cured once infected by a virus and should be destroyed. It is useful to identify the virus causing symptoms so that an effective management plan can be created and followed to limit production losses the following year.

All photos taken by Sara Bratsch. For non commercial use only.

Please contact regarding all other uses including but not limited to: data for computational algorithms, books, pamphlets, articles, apps, websites, etc.

Cite this article:

Bratsch, Sara. "TSWV". Web article. (date accessed). http://plantpathlesstraveled.com/TSWV/

Citations

Allen, W. R., & Broadbent, A. B. (1986). Transmission of tomato spotted wilt virus in Ontario greenhouses by Frankliniella occidentalis. Canadian Journal of Plant Pathology, 8(1), 33-38.

Mandal, B., Pappu, H. R., Csinos, A. S., & Culbreath, A. K. (2006). Response of peanut, pepper, tobacco, and tomato cultivars to two biologically distinct isolates of Tomato spotted wilt virus. Plant Disease, 90(9), 1150-1155.

Oliver, J. E., & Whitfield, A. E. (2016). The Genus Tospovirus: Emerging Bunyaviruses that Threaten Food Security. Annual Review of Virology, 3, 101-124.

Pappu, H. R., Jones, R. A. C., & Jain, R. K. (2009). Global status of tospovirus epidemics in diverse cropping systems: successes achieved and challenges ahead. Virus research, 141(2), 219-236.

Pappu, H. R. (2012). "Tomato spotted wilt virus." http://plantpath.wsu.edu/wp-content/uploads/2012/10/Pappu-final-version.pdf

Parrella, G., Gognalons, P., Gebre-Selassie, K., Vovlas, C., & Marchoux, G. (2003). An update of the host range of Tomato spotted wilt virus. Journal of Plant Pathology, 227-264.

Riley, D. G., & Pappu, H. R. (2004). Tactics for management of thrips (Thysanoptera: Thripidae) and tomato spotted wilt virus in tomato. Journal of economic entomology, 97(5), 1648-1658.

Stobbs, L. W., Broadbent, A. B., Allen, W. R., & Stirling, A. L. (1992). Transmission of tomato spotted wilt virus by the western flower thrips to weeds and native plants found in southern Ontario. Plant Disease, 76(1), 23-29.

Wijkamp, I., van Lent, J., Kormelink, R., Goldbach, R., & Peters, D. (1993). Multiplication of tomato spotted wilt virus in its insect vector, Frankliniella occidentalis. Journal of General Virology, 74(3), 341-349.

Causes of variegation

There are several factors that can cause variegation in plants. Differential gene regulation occurs in some variegated plants where the plant cells contain the same genomic sequence, but different cells of the plant express the gene(s) at different rate(s). This type of variegated plant will grow true to seed and can be observed in striped watermelon rinds, petunias with bicolor flowers, or the face on a pansy (Marcotrigiano 1997).

Physical structures of the plant leaf can also cause variegation. In this case, leaf cells physically separate from the layers above or below them, forming in essence, a plant blister (Hara 1957). This type of physical variegation causes silver spots or streaks and can be observed with Pilea cadieri, the aluminum plant.

The presence of viruses also causes variegation and other leaf or flower patterns. Virus induced variegation is observed in two popular mint varieties with golden veins, ‘golden ginger mint’ and ‘emerald mint’. These varieties have been shown to contain several viruses (Tzanetakis et al 2010). Eliminating the viruses from these cultivars also eliminates the presence of the attractive golden veins (Tzanetakis et al 2010). The presence of novel variegation and leaf distortion in some hosta spp. has also been attributed to the presence of a virus called Hosta virus X or HVX (Currier and Lockhart 1996; Blanchette and Lockhart 2003).

Hosta with virus caused leaf pattern. White edges are normal in this variety.

Genetic mosaicism or chimeras are also a cause of plant variegation. Plants with chimeric variegation may be transient and present in only one plant section, or stable and able to be passed down to progeny. A chimeric plant has genetically different cells in different regions. This type of variegation can arise by transposable genetic elements (Doodeman et al 1984), spontaneous mutations, or induced mutations in the nuclear or chloroplast genome (Tilney-Bassett 1986). Chimeric variegation can be stable and easily propagated or can be unstable and present in only an isolated portion of a plant.

Pepper plant with a transient chimeric leaf variegation in young leaves.

Stable chimeric variegation observed in variegated spider plant, Chlorophytum comosum. Yellow tint due to artificial lighting.

Differentiating between virus induced symptoms and natural genetic mutations in some plant species is challenging, time consuming, and requires laboratory testing. New hosta varieties are now vigorously tested to verify new leaf color patterns are caused by chloroplast mutations and not viruses.

Plant variegation is an interesting pattern of pigmentation in plants that can be caused by viruses, but in many cases is simply differential gene regulation, physical deformity, or a genetic mutation.

All photos taken by Sara Bratsch. For non commercial use only.

Please contact regarding all other uses.

Cite this article:

Bratsch, Sara. "Plant variegation". Web article. 27 February 2017. http://plantpathlesstraveled.com/plant-variegation/

Citations

Blanchette, B., & Lockhart, B. (2003). Hosta virus X: A three-year study. Hosta J, 35, 19-23.

Currier, S., & Lockhart, B. E. L. (1996). Characterization of a potexvirus infecting Hosta spp. Plant disease, 80(9), 1040-1043.

Doodeman, M., Boersma, W., Koomen, W., Bianchi, F. (1984). Genetic analysis of instability in Petunia hybrida l. A highly unstable mutation induced by a transposable element inserted at the An1 locus for flower colour. Theor. Appl. Genet. 67:345-455.

Hara, N. (1957). Study of the variegated leaves with special reference to those caused by air spaces. Jap. J. Bot, 16, 86-101.

Marcotrigiano, M. (1997). Chimeras and variegation: patterns of deceit. HortScience, 32(5), 773-784.

Tilney Bassett, R. A. (1986). Plant chimeras. Cambridge, UK: CUP.

Tzanetakis, I. E., Postman, J. D., Samad, A., & Martin, R. R. (2010). Mint viruses: Beauty, stealth, and disease. Plant disease, 94(1), 4-12.

CLYMV was first reported to occur in the USA in 1942 and was further characterized in 1962 (Johnson 1942, Agrawal et al. 1962).

ClYMV has a positve sense single stranded RNA genome and is a potexvirus. It can be detected by using antibodies (for Immunosorbant Electron Microscopy, ISEM, or ELISA) or through molecular methods (by reverse transcription polymerase chain reaction, RT-PCR).

Symptoms

In clover and alfalfa ClYMV produces mosaics or leaf streaking. It is also commonly found co-infecting clovers with White Clover Mosaic Virus (Agrawal et al. 1962). In pea ClYMV produces necrotic lesions and mosaic symptoms. ClYMV produces slight symptoms in verbena that include stunting, yellowing, and leaf distortion of young plant leaves (Baker et al. 2004).

ClYMV symptoms in red clover. The faint streaking may be mistaken for nutrient deficiency.

Vector

The vector (method of spread) of ClYMV is mechanically through infected sap or through seed.

Control

Seed for planting new clover or alfalfa fields should be tested for ClYMV to minimize the virus from the fields. Due to being spread mechanically, what was originally a small percentage of ClYMV infected clover or alfalfa plants can rapidly increase when the field is mowed or harvested.

Ornamental plants with virus like symptoms growing near symptomatic fields of clover or alfalfa should be tested for viruses to determine if they are infected with ClYMV.

Where can I see this virus?

ClYMV along with White Clover Mosaic Virus can be easily found in red clover and alfalfa that are growing in ditches or other areas. Plants will have streaky leaves and may be stunted.

All photos taken by Sara Bratsch. For non commercial use only.

Please contact regarding all other uses.

Cite this article:

Bratsch, Sara. "Clover yellow mosaic virus". Web article. 24 June 2016. http://plantpathlesstraveled.com/clover-yellow-mosaic-virus/

Citations

Agrawal, H., M. Chessin, and L. Bos. "Purification of clover yellow mosaic virus." (1962): 408-409.

Baker, C. A., K. Beckham, and E. Hiebert. "A virus related to Clover yellow mosaic virus found east of the Mississippi River in Verbena Canadensis in Florida." Plant Disease 88.2 (2004): 223-223.

Ford, Richard E. "Concentration and purification of clover yellow mosaic virus from pea roots and leaves." Phytopathology 63 (1973): 926-930.

Johnson, Folke. The complex nature of white-clover mosaic. Ohio State University, 1942.

Hampton, R., et al. "Host reactions of mechanically transmissible legume viruses of the northern temperate zone." Phytopathology (1978).

Pratt, Michael J. "Studies on clover yellow mosaic and white clover mosaic viruses." Canadian Journal of Botany 39.3 (1961): 655-665.

Rao, D.V., Hiruki, C. and Matsumoto, T. (1980). Phytopath. Z. 98: 260.

Sit, Tim L., et al. "Complete nucleotide sequence of clover yellow mosaic virus RNA." Journal of General Virology 71.9 (1990): 1913-1920.

Introduction

Impatiens necrotic spot virus (abbreviated INSV) is a prevalent plant virus found in horticultural production facilities. INSV was originally described as a strain of Tomato spotted wilt virus (TSWV) and was called the Tomato spotted wilt virus-Impatiens strain (TSWV-I) (Law & Moyer 1990). This name is rarely used today but can be observed in literature prior to the early 1990's.

Differences in TSWV strains were first reported in 1955 (Best & Gallus 1955). INSV responded weakly or not at all with antiserum to TSWV (Law & Moyer 1990; Wang & Gonsalves 1990). In addition, distinct structures could be seen in thin sections of plants infected with INSV but not TSWV (Law & Moyer 1990, Urban et al. 1991). This data drove the research that characterized INSV and today allows it to be detected separately from TSWV.

INSV is classified in the genus Tospovirus and has a single stranded RNA genome consisting of three segments. These segments are packaged into quasi-spherical particles (Law & Moyer 1991).

Symptoms

The symptoms caused by INSV vary widely and range from purple spots, necrotic (dying) spots, stunting, yellow concentric rings, wilting, stem lesions, plant death, and others. Some plants can exhibit no symptoms of INSV infection (they are asymptomatic). Asymptomatic plants can act as a virus reservoir where thrip vectors can acquire it. INSV symptoms can vary based on plant age when infected, growth conditions, plant growth rate, plant species/cultivar, and more.

Asymptomatic african violet infected with INSV.Some stunting is observed but this may be due to poor growth conditions.

Symptoms can be easily attributed to nutrient, chemical, environmental issues or confused with fungal or bacterial pathogens.

INSV has been found in over 350 species of plants including ornamental bedding plants, vegetables, weeds, and field crops originating from greenhouse transplants that were contaminated (Daughtrey et al. 1997).

Uneven symptom distribution among plants could be attributed to thrip feeding patterns. There have been no extensive surveys published since 1997 on the prevalence of INSV in horticulture of vegetable crops in the US (Daughtrey et al. 1997).

INSV symptoms in Nicotiana benthamiana. Note the characteristic necrotic rings formed where a thrip vector carrying INSV fed.

INSV symptoms in Nicotiana benthamiana, 1 day later. Note the appearance of water marking around the necrotic rings. The plant will soon wilt and die.

INSV symptoms in dahlia.

The Vector

INSV is primarily spread by thrips. The most prevalent INSV thrip vector in the USA is the western flower thrip, Frankliniella occidentalis (Van De Wetering et al 1996). Adult thrips are 1 mm in size and can be difficult to detect. Western flower thrip larva acquire INSV and can transmit the virus through their adult life (Van De Wetering et al 1996).

INSV can also be transmitted mechanically but this is not the prevalent route of spread in horticultural production facilities (Daughtrey et al. 1997).

Control

The best defense against INSV is to invest in virus free propagating material. Any new plants brought into the production facility should be inspected closely for symptoms of viral pathogens or thrip vectors. New plants should be isolated until they are verified to be INSV and thrip free. Thrip populations can be monitored with blue sticky traps placed at plant height. High thrip populations can be managed through use of insecticides but care must be taken to properly apply them due to the fact that thrips prefer to hide inside flowers and leaf buds. Insecticides should also be rotated to decrease the risk of allowing insects to become resistant to them as some thrip populations are already resistant to various chemicals (Daughtrey et al. 1997).

It is important to remember that even if thrips are present, if the plants are free from virus no losses will be attributed to INSV.

Transgenic crops were developed in the 1990's to resist INSV (and other Tospoviruses) but they have not been utilized (Gonsalves et al. 1995).

Testing

INSV can be detected through the use of immuno-strips in the field. These can be purchased directly by growers or plants can be submitted to a local plant disease clinic for testing. Symptomatic tissue should be used for testing.

INSV can also be detected through the use of indicator plants such as Nicotiana benthamiana, Petunia hybrida, and dwarf fava bean (Daughtrey et al. 1997).

All photos were taken by Sara Bratsch, for non commercial use only.

Please contact regarding all other uses.

Cite this article:

Bratsch, Sara. “Impatiens Necrotic Spot Virus” Web Article. 31 May 2016. http://plantpathlesstraveled.com/impatiensnecroticspotvirus/

Citations

Avila, A. D., Haan, P. D., Kitajima, E. W., Kormelink, R., Resende, R. D. O., Goldbach, R. W., & Peters, D. (1992). Characterization of a Distinct Isolate of Tomato Spotted Wilt Virus (TSWV) from Impatiens sp. in The Netherlands. Journal of Phytopathology, 134(2), 133-151.

Best, R.J. & Gallus, H.P. (1955). Strains of tomato spotted wilt virus. Australian Journal of Science. 15, 212-214.

Daughtrey, M. L., Jones, R. K., Moyer, J. W., Daub, M. E., & Baker, J. R. (1997). Tospoviruses strike the greenhouse industry: INSV has become a major pathogen on flower crops. Plant Disease, 81(11), 1220-1230.

de Haan, P., de Avila, A. C., Kormelink, R., Westerbroek, A., Gielen, J. J., Peters, D., & Goldbach, R. (1992). The nucleotide sequence of the S RNA of Impatiens necrotic spot virus, a novel tospovirus. FEBS letters, 306(1), 27-32.

Gonsalves, D., Pang, S. Z., Gonsalves, C., Xue, B., Yepes, M., & Jan, F. J. (1995). Developing transgenic crops that are resistant to tospoviruses. Tospoviruses and Thrips of Floral and Vegetable Crops 431, 427-431.

Law, M. D., & Moyer, J. W. (1990). A tomato spotted wilt-like virus with a serologically distinct N protein. Journal of General Virology, 71(4), 933-938.

Law, M. D., Speck, J., & Moyer, J. W. (1991). Nucleotide sequence of the 3? non-coding region and N gene of the S RNA of a serologically distinct tospovirus. Journal of General Virology, 72(10), 2597-2601.

Law, M. D., Speck, J., & Moyer, J. W. (1992). The M RNA of impatiens necrotic spot Tospovirus (Bunyaviridae) has an ambisense genomic organization. Virology, 188(2), 732-741.

Urban, L. A., Huang, P. Y., & Moyer, J. W. (1991). Cytoplasmic inclusions in cells infected with isolates of L and I serogroups of tomato spotted wilt virus. Phytopathology, 81(5), 525-529.

Van De Wetering, F., Goldbach, R., & Peters, D. (1996). Tomato spotted wilt tospovirus ingestion by first instar larvae of Frankliniella occidentalis is a prerequisite for transmission. PHYTOPATHOLOGY-NEW YORK AND BALTIMORE THEN ST PAUL-, 86, 900-905.

Wang, M., & Gonsalves, D. (1990). ELISA detection of various tomato spotted wilt virus isolates using specific antisera to structural proteins of the virus. Plant Disease, 74(2), 154-158.

Introduction

Dwarf mistletoes belong to the genus Arceuthobium and are obligate parasitic plants on conifers and cypress trees and shrubs in North America. In the USA one can find dwarf mistletoe in the western states (Rocky mountain area) and northern states. Although the majority of dwarf mistletoes are found natively in North America they can also be found in Central America, Asia, Europe, and northern Africa. They can be identified by their camouflaged shoots of thin green, yellow, or brown stems. Depending upon the species the stems can be as short as 0.5 inch or as long as 28 inches. Another symptom produced by some species is the formation of witches’ brooms, or a proliferation of shoots (Hawksworth et al. 1998).

Tree with numerous dwarf mistletoes camouflaged. Photo taken along the South Rim of the Grand Canyon National Park in Arizona.

Close up showing the yellow shoots.

Dwarf mistletoes are characterized as plant pathogens because they derive most of their nutrition directly from the host plant. They are able to do this by using a specialized vascular system similar to plant roots. This system consists of tubes called haustoria which anchor the dwarf mistletoe into the host plant and allow it to take nutrients and water from the host. Dwarf mistletoe are also able to increase the translocation rate from the host plant to ensure a constant flow of water and nutrients (Wanner and Tinnin 1986). Dwarf mistletoe does have chlorophyll, but at only a level of 10-25% of what the host plant contains (Hull and Leonard 1964b). Reports suggest different dwarf mistletoe species are able to photosynthesize from 10-50% of their needed carbohydrates (Rey et al. 1991).

Life Cycle

Dwarf mistletoe is a dioecious plant having separate male and female plants. A life cycle can take from two to eight years to complete. Once a mature seed lands on a susceptible host it will germinate and attach to the host branch. After several years of growth inside the host the mistletoe plant will start growing shoots which can live for five or more years. Female plants will produce flowers either in early summer or late summer to fall. The seeds are dispersed by hydrostatic explosive discharge allowing a maximal spread of 52 feet, although distances of 30 feet are more likely (Robinson and Geils 2006). Squirrels and birds are also able to spread the seeds but it is not the major dispersal method (Nicholls et al. 1991). The seeds are coated with a sticky coating (viscin) so they do not bounce off of the needles they hit-but rather adhere to them. Despite these adaptations less than 10% of seeds land on a suitable infection site, but large numbers of seeds are produced per plant to ensure continued survival (Hawksworth 1965).

Female plants will produce seeds at the tips of their branches. Photo taken near the Cloquet Forestry Center in Minnesota.

Impact

Dwarf mistletoes are considerable plant pathogens because they reduce: the growth rate, quality of wood (broken branches/dying tops), resistance to other diseases and insects (Kenaley et al. 2006), drought stress tolerance; and can increase the rate of wildfire spread due to the increase in dead branches in infected trees. However dwarf mistletoe is beneficial to numerous forest animals including: birds which use seeds as food and witches’ brooms as nesting sites; mammals which eat seeds, young shoots, and also use witches’ brooms as nesting sites and for cover; and for a variety of insects, mites, and spiders which eat the shoots, fruits, seeds, and other organisms present (Hawksworth et al. 1998).

Some trees produce witches’ brooms when infected with dwarf mistletoe. Due to the high nutrient requirements dwarf mistletoe can weaken and kill the upper crown of the host tree as seen here. Photo taken near the Cloquet Forestry Center in Minnesota.

Control

Numerous control measures have been developed for this plant pathogen with the simplest method being the removal of the infected branch or tree. This is not a perfect method however as it is difficult to determine the extent of infection in the tree and the entire dwarf mistletoe plant may not be removed. For larger areas chemical treatment with ethephon formulations (causes release of ethylene) will cause the plants to fall off of the host. This does not kill the dwarf mistletoe plant but does prevent it from spreading via seed. Applications must be repeated every one to three years for continued control. The final control measure is through controlled fires which will kill trees weakened by dwarf mistletoe and the dwarf mistletoe. The most extreme measure, which may be required for heavily infested forests, is to clear cut large areas to remove the dwarf mistletoe plants and seeds.

On your holiday travels this year make sure to take a look out the windows-you may spot an interesting plant pathogen decorating the trees.

All photos were taken by Sara Bratsch. For non commercial use only.

Please contact regarding all other uses.

Cite this article:

Bratsch, Sara. “Dwarf Mistletoe”. 23 December 2015. http://plantpathlesstraveled.com/dwarfmistletoe/

Article Citations

Hawksworth, F. G. (1965). Life Tables for Two Species of Dwarfmistletoe I. Seed Dispersal, Interception, and Movement. Forest Science, 11(2), 142-151.

Hawksworth, F. G., Brian, F. G. H. D. W., & Wiens, D. (1998). Dwarf mistletoes: biology, pathology, and systematics. DIANE Publishing.

Hoffman, J. (2004). Management Guide for Dwarf Mistletoe. Forest Health Protection and State Forestry Organizations, 14.

Hull, R. J., & Leonard, O. A. (1964). Physiological aspects of parasitism in mistletoes (Arceuthobium and Phoradendron). II. The photosynthetic capacity of mistletoe. Plant Physiology, 39(6), 1008.

Kenaley, S. C., Mathiasen, R. L., & Daugherty, C. M. (2006). Selection of dwarf mistletoe-infected ponderosa pines by Ips species (Coleoptera: Scolytidae) in northern Arizona. Western North American Naturalist, 66(3), 279-284.

Nicholls, T. H., Egeland, L., & Hawksworth, F. G. (1989). Birds of the Fraser Experimental Forest, Colorado, and their role in dispersing lodgepole pine dwarf mistletoe. Colorado Field Ornithology Journal, 23, 3-12.

Rey, L., Sadik, A., Fer, A., & Renaudin, S. (1991). Trophic relations of the dwarf mistletoe Arceuthobium oxycedri with its host Juniperus oxycedrus. Journal of plant physiology, 138(4), 411-416.

Robinson, D. C., & Geils, B. W. (2006). Modelling dwarf mistletoe at three scales: life history, ballistics and contagion. Ecological modelling, 199(1), 23-38.

Wanner, J., & Tinnin, R. O. (1986). Respiration in lodgepole pine parasitized by American dwarf mistletoe. Canadian Journal of Forest Research, 16(6), 1375-1378.

Introduction

Squash mosaic virus (abbreviated SqMV) is a common virus found in cucurbit plants. SqMV was first described by a scientist named Kendrick in 1934 but it was not until 1941 that Freitag identified the causal virus. SqMV continues to be found in cucurbit crops today. SqMV is a spherical virus with particles ~30 nm in diameter (Campbell 1971).

Squash mosaic virus from a partially purified squash extract that is stained with PTA. This image was taken at 53,000 times magnification. The virus particles are the round structures.

Symptoms

Infected plant leaves may be symptomless or show ring patterns, chlorotic mottle, deformation, vein clearing, or streaking. Fruit that is infected at or before blossoming will show symptoms including color change, bumps, and other distortion. Plants infected after fruit have set will show symptoms in the leaves but the fruit will not develop symptoms.

Zucchini infected with SqMV showing severe foliar symptoms of deformation, bumps, and chlorotic mottling.

‘Jack be little’ pumpkin infected with SqMV. Note the variation in leaf symptoms between old leaves (right side) and newer leaves (left side).

Yellow squash fruit that tested positive for SqMV showing bumps and color change.

Host

The natural host range of SqMV is extremely limited and includes only the Cucurbitaceae family (squashes, pumpkins, melons, gourds) (Campbell 1971). Experimentally SqMV has been transmitted to the plant families: Amaranthaceae, Chenopodiaceae, Hydrophyllaceae, Leguminosae, and Umbelliferae (Frietag 1956). One report from Morocco has also transmitted SqMV by mechanical inoculation to Chenopodium quinoa Willd. where it systemically infected the plants and produced symptoms (Lockhart, Ferji, Hafidi 1982).

Control

SqMV is seed transmitted with a transmission rate ranging from 1%-94% transmission (Grogan, Hall & Kimble 1959; Rayder, Fitzpatrick & Hildebrand 1947). It can be transmitted within a field from infected plants by various beetle species including: western striped cucumber beetle (Acalymma trivittata Mann.), western twelve-spotted cucumber beetle (Diabrotica undecimpunctata), banded cucumber beetle (D. balteata), 28-spotted ladybird beetle (Henosepilachna vigintioctopunctata) and other beetles (Freitag 1941, Sittterly 1960). The beetles can obtain SqMV by feeding for only a brief period of time, and are able to transmit the virus immediately for up to 20 days. The length of time beetles feed on infected plants determines how long they are able to transmit the virus, longer feeding times mean the beetles will transmit the virus for a longer period of time. SqMV can also be transmitted mechanically (Campbell 1971).

Control measures for SqMV include using only virus-free certified seed, and maintaining low beetle populations before fruit sets. If plants show symptoms early in the growing season it is important to remove them and destroy the tissue so that they do not serve as a source of inoculum to infect the remaining plants in the field.

Virus Specifics

Squash mosaic virus is a spherical virus that belongs to the family Secoviridae, genus Comovirus. This group of viruses have linear single stranded (positive sense) RNA genomes enclosed in non enveloped capsid (King et al. 2011).

All photos were taken by Sara Bratsch. For non commercial use only.

Please contact regarding all other uses.

Cite this article:

Bratsch, Sara. “Squash Mosaic Virus Part 2 (SqMV)”. 31 October 2015. http://plantpathlesstraveled.com/squash-mosaic-virus-part-2-sqmv/

Citations

Campbell, R. N. (1971). Squash mosaic virus. CMI/AAB Descriptions of plant viruses, (43), 4.

Freitag. (1941). Insect transmission, host range and properties of squash-mosaic virus.
Phytopathology 31: 8.

Freitag, J. H. (1956). Beetle Transmission, Host Range, and Properties of Squash Mosaic Virus.
Phytopathology, 46(2), 73-81.

Grogan, R. G., Hall, D. H., & Kimble, K. A. (1959). Cucurbit mosaic viruses in California.
Phytopathology, 49(6), 366-376.

Kendrick, J. (1934). Cucurbit mosaic transmitted by melon seed. Phytopathology, 24.7: 820-823.

King, A. M., Adams, M. J., & Lefkowitz, E. J. (2011). Virus taxonomy: classification and nomenclature of viruses: Ninth Report of the International Committee on Taxonomy of Viruses (Vol. 9). Elsevier.

Lockhart, B. E. L., Ferji, Z., & Hafidi, B. (1982). Squash mosaic virus in Morocco. Plant Disease, 66(12), 1191-1193.

Rayder, Wm. E., Fitzpatrick, H. F., Hildebrand, E. M. (1947). A seed-borne virus of muskmelon.
Phytopathology 37: 809-816.

Introduction

Odontoglossum ringspot virus (abbreviated ORSV) is one of the most common and important viruses infecting orchids worldwide. The other major viral pathogen of orchids is Cymbidium mosaic virus. ORSV was first described in 1951 as a viral pathogen of Odontoglossum grande orchid plants (Jensen, Gold 1951). ORSV in the past was also called the orchid strain of tobacco mosaic virus due to the similarity in virus particle shape and size. ORSV has rod-shaped particles that are 300 nanometers long and 18 nanometers wide.

Negatively stained transmission electron microscopy of ORSV isolated from a Cymbidium hybrid orchid by Sara Bratsch.

Symptoms

ORSV infecting Odontoglossum grande plants cause symptoms of small necrotic spots that appear on older leaves while younger leaves have light green or pale yellow rings. The rings continue to grow in size and can cover the entire leaf. Flowers may exhibit color breaking or necrotic spotting (Jensen, Gold 1951). Mild flower symptoms on both flower petals and sepals include irregular areas that are lighter or darker in color than normal. Severe flower color break will cause white variegation in both petals and sepals or brown necrotic streaking (McMillan, Vendrame 2005). Previous studies have shown no obvious foliar symptoms in cymbidium orchids mechanically inoculated with ORSV and followed for 4 years. The plants did have a reduction in growth along with several plants dying (Pearson, Cole 1991). The absence of obvious leaf symptoms in ORSV infected plants increases the likelihood of the virus infection going undetected.

Symptoms of ORSV infection in several orchid species. A-Bollea violaceae showing very slight symptoms (which could also be due to lack of fertilizing). B-Epidendrum raniferum leaves showing extreme chlorosis and necrotic spotting. C-Cymbidium sp. with pin-prick chlorotic spots. D-Miltonia sp. showing necrotic spotting. E-Brassia verrucosa with chlorotic spots and leaf yellowing.

Control

ORSV can be transmitted mechanically with infected sap on tools. Seed or insect vector transmission of ORSV has not been extensively studied. At this time the major transmission method of ORSV is mechanical transmission or propagation by tissue culture of an infected mother plant.

Several studies have been done to determine the incidence of ORSV infection rate in several countries. A 1993 orchid survey by the University of Hawaii of 3,600 orchid plants from 3 collections, 22 commercial farms, and 6 nurseries identified a plant infection rate of 17% by ORSV and 15% infection by both ORSV and Cymbidium mosaic virus (CymMV) (CymMV was found in 45%)(Hu et al. 1993). A similar study by the Department of Botany in Singapore of 1146 orchid plants from 4 commercial cut flower farms identified a 4% ORSV infection rate and 14.2% infection rate for ORSV and CymMV (54.6% were infected with CymMV alone)(Wong et al. 1994). A more recent survey from Thailand of 280 Dendrobium orchids from cut flower farms in Thailand identified a 0% infection rate by ORSV (65.4% CymMV infection rate)(Khentry et al 2006). A study from Sikkim, India showed 42 of 100 orchids sampled were infected with ORSV with the majority of plants showing no symptoms (Sherpa et al. 2006). These results indicate that it is likely plants being sold are infected with ORSV but it is possible to keep the incidence to 0% through sterilization methods (as seen by the 0% infection rate from cut flower farms).

There has been some work done using tissue culture methods to eliminate ORSV from cultured plant material. These methods could be used to obtain virus free plantlets from an infected mother plant. ORSV is not easily eliminated through meristem tissue culture, however it is possible to obtain some virus free plantlets using tissue culture with chemotherapy (antiviral chemicals added to the media). ORSV has been shown to be present in the lateral and apical meristems of ORSV infected Cymbidium orchids making meristem culture alone an inefficient method of producing virus-free plantlets (Toussaint et al. 1984). Meristem tissue culture combined with antiviral chemicals (like ribavirin/virazole) have been successful in producing ORSV free Cymbidium plantlets, however the efficiency was dependent upon media and concentration of chemical used (Freitas-Astua et al. 1998). Further studies with Mokara orchids in tissue culture showed similar results with certain concentrations of ribavirin working more efficiently to eliminate ORSV from meristem or thin section cultures (Lim et al. 1993).

Testing methods for ORSV include indicator plants, reverse transcription polymerase chain reaction (RT-PCR), and antibody based methods including ELISA, immunosorbant transmission electron microscopy, and rapid antibody tests. The commercially available rapid antibody tests from Agdia for ORSV, which are available for orchid growers of all sizes, utilize a method similar to pregnancy tests with a band indicating a positive reaction from a sample of ground plant tissue. Plant Pathology diagnostic labs can also be consulted for orchid virus (and other pathogen) tests.

Control measures for ORSV include testing and isolating or destroying infected plants. Plants that will be used for mass propagation by tissue culture should be tested for viruses before they are used for propagation, even if they do not show symptoms, as not every plant will express symptoms. Strict sterilization methods are required for working with more than one orchid plant. Tools should be single use like individual razor blades which are disposed after using on one plant. Otherwise tools should be disinfected by dipping in a 1% NaOH (bleach) solution between plants to inactivate any virus present (Hu 1994). It is also important to sterilize pots, benches, and the working area to eliminate the risk of transmitting any virus present.

In the future it may be possible to buy genetically engineered orchids with resistance to ORSV and CymMV, however there are currently no resistant orchids being sold. The only way to protect your collection of orchids is to maintain sterilization techniques between plants and when purchasing new plants to buy only ones that have been virus tested. To determine if your plant has been virus tested you can look on the label or contact the seller directly to learn what viruses they are testing for. It is important to remember that although a plant may be expressing no symptoms, it may still contain a virus that can slowly weaken plants and be spread between plants.

For information on CymMV check out my posting about it here: https://plantpathlesstraveled.com/cymbidium-mosaic-virus/

All photos were taken by Sara Bratsch. For non commercial use only.

Please contact regarding all other uses.

Cite this article:

Bratsch, Sara. “Odontoglossum ringspot virus”. 12 June 2015. http://plantpathlesstraveled.com/odontoglossum-ringspot-virus/

Citations:

Freitas-Astua, J., & Rezende, J. A. M. (1998). Odontoglossum ringspot virus-free Cymbidium obtained through meristem tip chemotherapy. Fitopatologia Brasileira, 23(2), 158-160.

Hu, J. S., Ferreira, S., Xu, M. Q., Lu, M., Iha, M., Pflum, E., & Wang, M. (1994). Transmission, movement and inactivation of cymbidium mosaic and odontoglossum ringspot viruses. Plant disease, 78(6), 633-636.

Khentry, Y., Paradornuwat, A., Tantiwiwat, S., Phansiri, S., & Thaveechai, N. (2006). Incidence of Cymbidium mosaic virus and Odontoglossum ringspot virus in Dendrobium spp. in Thailand. Crop Protection, 25(9), 926-932.

Lim, S. T., Wong, S. M., & Goh, C. J. (1993). Elimination of cymbidium mosaic virus and odontoglossum ringspot virus from orchids by meristem culture and thin section culture with chemotherapy. Annals of applied biology, 122(2), 289-297.

McMillan Jr, R. T., & Vendrame, W. A. (2005). Color break in orchid flowers. In Proceedings of the Florida State Horticultural Society (Vol. 118, pp. 287-288).

Pearson, M. N., & Cole, J. S. (1991). Further observations on the effects of Cymbidium mosaic virus and Odontoglossum ringspot virus on the growth of Cymbidium orchids. Journal of Phytopathology, 131(3), 193-198.

Sherpa, A. R., Bag, T. K., Hallan, V., & Zaidi, A. A. (2006). Detection of Odontoglossum ringspot virus in orchids from Sikkim, India. Australasian Plant Pathology, 35(1), 69-71.

Toussaint, A., Dekegel, D., & Vanheule, G. (1984). Distribution of Odontoglossum ringspot virus in apical meristems of infected Cymbidium cultivars. Physiological plant pathology, 25(3), 297-305.

Wong, S. M., Chng, C. G., Lee, Y. H., Tan, K., & Zettler, F. W. (1994). Incidence of cymbidium mosaic and odontoglossum ringspot viruses and their significance in orchid cultivation in Singapore. Crop Protection, 13(3), 235-239.

Introduction

The orchid family is a diverse plant family valued for their exotic flowers, scents, and flavor. Orchids were first collected in the early 1800's by British explorers in South America and brought back to glass greenhouses in England. The ensuing "orchid fever" among hobbyists drove a lucrative business in procuring, rearing, and selling rare orchids collected from the wild. Small-scale production of orchids was common up until the 1950's when research studies revealed successful tissue culture protocols, planting media, environmental conditions, and fertilization methods to use for rapid Cattleya and Phalaenopsis production (Griesbach 2002).

Today the major countries producing orchids for potted plants or cut flowers include: China, Germany, Japan, Netherlands, Taiwan, and the USA. The USDA Floriculture Crops 2013 Summary shows that in the U.S. 170 businesses produced 30.3 million pots of orchids with a wholesale value of $246 million. In contrast to this in the cut flower orchid market 32 businesses produced 5.5 million stems of orchids with a wholesale value of $5.6 million.

As is often the case with plants that are globally traded and incredibly popular there are several pathogens that can damage or kill orchid plants. These include viruses (Cymbidium Mosaic Virus, Odontoglossum Ringspot Virus, Orchid Fleck Virus, others); bacteria causing soft rots and spots; and fungi causing wilts, root rots, and leaf spots. See Phelps Farms for more detailed pictures. (PDF file)

Virus details

Cymbidium mosaic virus (CymMV) was first described in 1951 and is also referred to as Cymbidium black streak virus, and Orchid Mosaic Virus (Jensen 1951). CymMV is a potexvirus with filamentous particles, 13 nm wide by 475 nm long, enclosing an RNA genome. CymMV and Odontoglossum Ringspot virus are the most common viral pathogens of orchids. CymMV and odontoglossum ringspot virus are the most common viral pathogens of orchids.

Transmission electron microscope image of CymMV particles at 66,000 times magnification.

Symptoms

CymMV causes brown necrotic spots on orchid leaves and flowers, but can also be symptomless. The major transmission method is mechanically through tools contaminated with virus infected plant sap or propogation by tissue culture of an infected mother plant. There are no known insect vectors and seed transmission is only likely at a very low rate or may not be possible. The possibility of seed transmission of CymMV has not been extensively studied.

Diagnostics

Testing methods for CymMV include indicator plants, reverse transcription polymerase chain reaction (RT-PCR), and antibody based methods including ELISA, immunosorbant transmission electron microscopy, and rapid antibody tests. The commercially available rapid antibody tests from Agdia for CymMV, which are available for orchid growers of all sizes, utilize a method similar to pregnancy tests with a band indicating a positive reaction from a sample of ground plant tissue. Plant Pathology diagnostic labs can also be consulted for orchid virus (and other pathogen) tests.

Orchids infected with and showing symptoms of CymMV. Plants were tested by dipstick and ISEM. A. Vanda tricolor orchid showing older leaves with more brown streaks (right) and younger leaves behind (left) just beginning to show symptoms. B. Cattleya bicolor showing streaking symptoms on the canes and leaves. C. Ludisia discolor, jewel orchid, showing mosaic symptoms in the new growth. Due to the great genetic variation in the orchid family it is not wise to base a disease diagnosis on symptoms alone.

Control

Control measures for CymMV include testing and isolating or destroying infected plants. Plants that will be used for mass propagation by tissue culture should be tested for viruses before they are used for propagation even if they do not show symptoms. Strict sterilization measures are necessary for all growth stages of orchids to prevent virus infections as CymMV can remain infectious in plant sap for up to a week at room temperature (Gibbs 1970). Tools should be single use like individual razor blades disposed after using on one plant, or thoroughly sterilized by dipping in ethanol and flaming for 15 seconds or dipping in 1% NaOH (bleach) between individual plants (Hu 1994). Skim milk does not inactivate CyMV and should not be used. NaOH concentrations higher than 1% inactivate CymMV but also cause phytotoxic damage on orchids (Hu 1994). It is important to sterilize repotting tools, pots, and benches as they can also serve as inoculation points for orchids.

It is important to choose virus-tested orchids when purchasing a new plant as that is the only way you can guarantee it does not contain a virus. Implementing sterilization techniques between your plants will limit the spread of any virus already in your collection.

All photos were taken by Sara Bratsch. For non commercial use only.

Please contact regarding all other uses.

Cite this article:

Bratsch, Sara. “Cymbidium mosaic virus”. 10 March 2015. http://plantpathlesstraveled.com/cymbidium-mosaic-virus/

Citations:

Gibbs, A.J.; Harrison, B.D.; Murant, A.F. (1970). CMI/AAB Descriptions of Plant Viruses. Commonwealth Mycological Institute, England. 27.

Griesbach, R.J. (2002). Development of Phalaenopsis Orchids for the Mass-Market. p. 458Ð465. In: J. Janick and A. Whipkey (eds.), Trends in new crops and new uses. ASHS Press, Alexandria, VA.

Hu, J. S., Ferreira, S., Xu, M. Q., Lu, M., Iha, M., Pflum, E., & Wang, M. (1994). Transmission, movement and inactivation of cymbidium mosaic and odontoglossum ringspot viruses. Plant disease, 78(6), 633-636.

Jensen DD (1951). Mosaic or black streak disease of Cymbidium orchids. Phytopathology 41: 401-414.

http://www.phelpsfarm.com/OrchidPestsandDiseases.pdf

Cucumber Mosaic Virus

Cucumber mosaic virus (CMV) can infect over 1,200 plant species and results in a wide range of symptoms depending on host type and at what age the plant was infected. Plants that are infected from a young age will have more severe symptoms. CMV can be spread though infected seed, aphid vectors, and mechanical damage. As CMV has a wide host range an important source of virus inoculum can come from weedy plant species growing outside of a field of pumpkins.

CMV infected cucurbits will experience stunting, malformed leaves, bumps on fruits, and a mosaic yellow and green pattern on the leaves. CMV symptoms in other plant families vary.

Close up view of a symptomatic malformed bush zucchini leaf showing rugosity (roughening). The plant was inoculated 38 days previously.

CMV can be managed by using resistant varieties of cucumber, squash, and melons. If resistant varieties are not available floating row covers can be applied to susceptible crops, which will prevent aphids from infecting young plants.^[1]

Watermelon Mosaic Virus

Watermelon mosaic virus (WMV) can infect all cucurbits and several weedy plant species. Typical early symptoms include vein clearing. Vein clearing is when the veins of plant leaves become translucent or clear. Mosaic patterns, mottling, and stunting of leaves will also occur later in an infection.

WMV can be spread by aphids so early detection and roguing (pulling out) infected plants is necessary. Protective systemic insecticides can also be used to reduce aphid populations limiting virus spread, however this will not completely eliminate virus spread as the aphids only need to probe a leaf once to infect a new plant.^[2]

Close up view of a stunted zucchini leaf infected with WMV showing a mottled light/dark green pattern.

Squash Mosaic Virus

Squash mosaic virus (SqMV) infects various members of the cucurbit family, several beans (Leguminosae), several members of the carrot family (Umbelliferae), and several weedy plant species.

Symptoms of SqMV infection include a dark green mosaic, vein clearing, and malformed leaves and fruit.

SqMV can be spread by seeds and though beetle spit transferred while feeding. Infected beetles can transmit the virus for up to 20 days. SqMV however is not spread by aphids. To avoid a SqMV infection plant only certified virus-free seed and limit beetle populations.^[3]

Close up view of zucchini leaves infected with SqMV showing mosaic and rugosity. Yellowish tint is from artificial lighting.

All photos were taken by Sara Bratsch. For non commercial use only.

Please contact regarding all other uses.

Cite this article:

Bratsch, Sara. "Pumpkin Viruses! Happy Halloween!" 31 October 2015. http://plantpathlesstraveled.com/pumpkin-viruses-happy-halloween/

Citations:

Zitter, Thomas A., and J. F. Murphy. "Cucumber mosaic virus." The Plant Health Instructor 10 (2009): 1094. http://www.apsnet.org/edcenter/intropp/lessons/viruses/Pages/Cucumbermosaic.aspx^[1]

http://www.ipm.ucdavis.edu/PMG/r116101811.html^[2]

http://www.pestid.msu.edu/PlantDiseases/SquashMosaicVirus/tabid/291/Default.aspx^,[3]`

Mutinus elegans can first be spotted as an egg-shaped mass on the ground. If one of the eggs is cut open it will reveal a liquid filled mini-mushroom. The egg will sprout the main bright orange stalk which sports a grey, slimy mass of spores on the tip. The scent emitted attacks flies and other insects which carry the spores to other locations.

Here you can see several eggs, which will emerge into mushrooms in the next several days. There are also several withered mushrooms from the previous day.

From egg, to sprout, to mushroom.

Mature fruiting stinkhorn.

Stinkhorn mushrooms are aptly named as they have a strong odor and intense coloration. Check out your lawn and mulch beds this fall to see if you have any to observe up close. This mushroom is not poisonous, and the immature eggs can be eaten-however they are not as delicious as a morel! If the mushrooms disturb you, it is possible to remove them by raking them out of your lawn or mulch.

All photos were taken by Sara Bratsch. For non commercial use only.

Please contact regarding all other uses.

Cite this article:

Sara Bratsch. "Stinkhorn mushroom: Mutinus elegans". 1 October 2014. http://plantpathlesstraveled.com/stinkhorn-mushroom-mutinus-elegans/

Citations:

Info from:
http://www.mushroomexpert.com/mutinus_elegans.html