Posts Tagged: Bt
Why This Corn Pest May Lose Its Nickname of 'Billion-Dollar Beetle'
The Western corn rootworm, known as “the billion-dollar beetle” because its...
A cornfield in Franklin, Pa. (Photo by Fishhawk of Flickr, Creative Commons)
Corn rootworm adult feeding on silk. (Photo by Sarah Zukoff)
Corn rootworm damage. (Photo by Keith Waldron)
Insect resistance to biopesticides
Mechanisms of insecticide resistance in insects.
Use of biopesticides or non-chemical pesticides is encouraged as a part of integrated pest management (IPM) for environmental and human safety and to reduce the risk of insecticide resistance. With the increase in biopesticide use in both organic and conventional cropping systems, it is a good time to review the potential of insect resistance to botanical and microbial pesticides.
Insects and mites develop resistance to chemical pesticides through genetic, metabolic, or behavioral changes resulting in reduced penetration of toxin, increased sequestration or excretion, reduced binding to the target site, altered target site that prevents binding of the toxin, or reduced exposure to the toxin through modified behavior. When the active ingredient is a toxic molecule and has the mode of action similar to that of a chemical compound, regardless of the plant or microbial origin, arthropods are more likely to develop resistance through one or more of the abovementioned mechanisms. When the mode of action is infection by a microorganism, rather than a toxin, arthropods are less likely to develop resistance. Under natural circumstances, plants, insects, natural enemies, and beneficial or harmful microbes continuously co-evolve and adapt to changing environment. When there is a higher selection pressure, such as indiscriminate use of chemical pesticides, increased mutagenesis can lead to resistance issues. A good understanding of insect resistance to biopesticides will help minimize potential risks and improve their efficient use in IPM.
Resistance to botanical pesticides
Nicotine, an alkaloid from Nicotiana spp., is one of the earlier botanical pesticides known. Although nicotine is not currently used as an insecticide, its synthetic alternatives – neonecotinoids – are commonly used against several pests. Botanical insecticide pyrethrum, extracted from the flowers of Chrysanthemum cinerariaefolium, contains insecticidal pyrethrins (synthetic pyrethrins are referred to as pyrethroids). Although insect resistance to pyrethrum or pyrethroid compounds has been known (Whitehead, 1959; Immaraju et al., 1992; Glenn et al., 1994), they have been effectively used against a number of pests through careful placement in IPM, organic, or conventional management strategies. Additionally, pyrethrin products have been effectively used along with piperonyl butoxide, which acts as a synergist and resistance breaker (Gunning et al. 2015).
Another botanical insecticidal compound, azadirachtin, is a tetranortriterpenoid limonoid from neem (Azadirachta indica) seeds, which acts as an insecticide, antifeedant, repellent and insect growth regulator. While neem oil, which has a lower concentration of azadirachtin, has been used in the United States as a fungicide, acaricide, and insecticide for a long time, several azadirachtin formulations in powder and liquid forms have become popular in recent years and were found effective in managing important pests (Dara 2015a and 2016). Feng and Isman (1995) reported that the green peach aphid, Myzus persicae developed resistance to pure azadirachtin under artificially induced selection pressure after 40 generations, but did not develop resistance to a refined neem seed extract. They suggested that natural blend of azadirachtin compounds in a biopesticide would not exert selection pressure that could lead to resistance. Additionally, Mordue and Nisbet (2000) discussed that azadirachtin can play a role in insecticide resistance management because it reduces the detoxification enzyme production as a protein synthesis inhibitor. Azadirachtin also improved the efficacy of other biopesticides in multiple studies (Trisyono and Whalon, 2000; Dara, 2013 and 2015b).
Insects feeding on plant allelochemicals can develop cross-resistance to insecticides (Després et al., 2007). For example, overproduction of detoxification enzymes such as glutathione S-transferases and monooxygenases in the fall armyworm, Spodoptera frugiperda,when it fed on corn and cowpea, respectively, imparted cross-resistance to various chemical pesticides. It is important to keep this in mind when botanical pesticides are used to detect potential resistance issues.
Resistance to bacterial biopesticides
Bacillus thuringiensis (Bt)is a gram-positive soil bacterium, which contains crystalline toxic protein that is activated upon ingestion by an insect host, binds to the receptor sites in the midgut, and eventually causes insect death. Since the mode of action involves a toxin rather than bacterial infection, several insects developed resistance to Bt pesticides or transgenic crops that contain Bt toxins (Tabashnik et al., 1990; McGaughey and Whalon, 1992; Tabashnik, 1994; Iqbal et al., 1996). However, Bt pesticides are still very popular and used against a variety of lepidopteran (Bt subsp. aizawai and Bt subsp. kurstaki), dipteran (Bt subsp. israelensis and Bt subsp. sphaericus), and coleopteran (Bt subsp. tenebrionis) pests.
Spinosad is a mixture of macrocyclic lactones, spinosyns A and spinosyns D, derived from Saccharopolyspora spinosa, an actinomycete gram-positive bacterium, and is used against dipteran, hymenopteran, lepidopteran, thysanopteran, and other pests. Spinosad products, while naturally derived are registered as chemical pesticides, not as biopesticides. Insect resistance to spinosad later led to the development of spinetoram, which is a mixture of chemically modified spinosyns J and L. Both spinosad and spinetoram are contact and stomach poisons and act on insect nervous system by continuous activation of nicotinic acetylcholine receptors. However, insect resistance to both spinosad (Sayyed et al., 2004; Bielza et al., 2007) and spinetoram (Ahmad and Gull, 2017) has been reported due to extensive use of these pesticides. Cross-resistance between spinosad and some chemical insecticides has also occurred in some insects (Mota-Sanchez et al., 2006; Afzal and Shad, 2017).
Resistance to viral biopesticides
Baculovirus infections in lepidoptera have been known for centuries, especially in silkworms. Currently, there are several commercial formulations of nucleopolyhedroviruses (NPV) and granuloviruses (GV). When virus particles are ingested by the insect host, usually lepidoptera, they invade the nucleii of midgut, fatbody, or other tissue cells and kill the host. Baculoviruses are generally very specific to their host insect species and can be very effective in bringing down the pest populations. However, variations in the susceptibility of certain insect populations and development of resistant to viruses has occurred in several host species (Siegwart et al., 2015). Resistance to different isolates of Cydia pomonella granulovirus (CpGV-M, CpGV-S) in codling moth (Cydia pomonella) populations is well known in Germany and other parts of Europe (Sauer et al., 2017a & b).
Resistance to fungal biopesticides
There are several fungi that infect insects and mites. Fungal infection starts when fungal spores come in contact with an arthropod host. First, they germinate and gain entry into the body by breaching through the cuticle. Fungus later multiplies, invades the host tissues, kills the host, and emerges from the cadaver to produce more spores. Entomophthoralean fungi such as Entomophthora spp., Pandora spp., and Neozygites spp. can be very effective in pest management through natural epizootics, but cannot be cultured in vitro for commercial scale production. Hypocrealean fungi such as Beauveria bassiana, Isarea fumosorosea, Metarhizium brunneum, and Verticillium lecanii,on the other hand, can be mass-produced in vitro and are commercially available. These fungi are comparable to broad-spectrum insecticides and are pathogenic to a variety of soil, foliar, and fruit pests of several major orders. Since botanical, bacterial, and viral biopesticides have insecticidal metabolites, proteins, or viral particles that have specific target sites and mode of action, insects have a higher chance of developing resistance through one or more mechanisms. Although fungi also have insecticidal proteins such as beauvericin in B. bassiana and I. fumosorosea and dextruxin in M. anisopliae and M. brunneum, their mode of action is more through fungal infection and multiplication and arthropods are less prone to developing resistance to entomopathogenic fungi. However, insects can develop resistance to entomopathogenic fungi through increased melanism, phenoloxidase activity, protease inhibitor production, and antimicrobial and antifungal peptide production (Wilson et al., 2001; Zhao et al., 2012; Dubovskiy et al., 2013). It appears that production of detoxification enzymes in insects against fungal infections can also impart resistance to chemical pesticides. Infection of M. anisopliae in the larvae of greater wax moth, Galleria mellonella, increased dexotification enzyme activity and thus resistance to malathion (Serebrov et al., 2006).
These examples show that insects can develop resistance to biopesticides in a manner somewhat similar to chemical pesticides, but due to the typically more complex and multiple modes of action, at a significantly lesser rate depending on the kind of botanical compound or microorganism involved. Resistance to entomopathogenic fungi is less common than with other entomopathogens. Since biopesticide use is not as widespread as chemical pesticides, the risk of resistance development is less for the former. However, excessive use of any single tool has the potential for resistance or other issues and IPM, which uses a variety of management options, is always a good strategy.
Acknowledgements: Thanks to Pam Marrone for reviewing the manuscript.
References
Afzal, M.B.S. and S. A. Shad. 2017. Spinosad resistance in an invasive cotton mealybug, Phenacoccus solenopsis: cross-resistance, stability and relative fitness. J. Asia-Pacific Entomol. 20: 457-462.
Ahmad M. and S. Gull. 2017. Susceptibility of armyworm Spodoptera litura (Lepidoptera: Noctuidae) to novel insecticides in Pakistan. The Can. Entomol. 149: 649-661.
Bielza, P., V. Quinto, E. Fernández, C. Grávalos, and J. Contreras. 2007. Genetics of spinosad resistance in Frankliniella occidentalis (Thysanoptera: Thripidae). J. Econ. Entomol. 100: 916-920.
Dara, S. K. 2013. Strawberry IPM study 2012: managing insect pests with chemical, botanical, and microbial pesticides. UCANR eJournal Strawberries and Vegetables March 13, 2013 (//ucanr.edu/blogs/blogcore/postdetail.cfm?postnum=9595).
Dara, S. K. 2015a. Root aphids and their management in organic celery. CAPCA Adviser 18(5): 65-70.
Dara, S. K. 2015b. Strawberry IPM study 2015: managing insect pests with chemical, botanical, microbial, and mechanical control options. UCANR eJournal Strawberries and Vegetables November 30, 2015 (//ucanr.edu/blogs/blogcore/postdetail.cfm?postnum=19641).
Dara, S. K. 2016. Managing strawberry pests with chemical pesticides and non-chemical alternatives. Intl. J. Fruit Sci. https://doi.org/10.1080/15538362.2016.1195311
Després, L., J.-L. David, and C. Gallet. 2007. The evolutionary ecology of insect resistance to plant chemicals. Trends in Ecol. Evol. 22: 298-307.
Dubovskiy, I. M., M.M.A. Whitten, O. N. Yaroslavtseva, C. Greig, V. Y. Kryukov, E. V. Grizanova, K. Mukherjee, A. Vilcinskas, V. V. Glupov, and T. M. Butt. 2013. Can insects develop resistance to insect pathogenic fungi? PloS one 8: e60248.
Feng, R. and M. B. Isman. 1995. Selection for resistance to azadirachtin in the green peach aphid, Myzus persicae. Experientia 51: 831-833.
Glenn, D. C., A. A. Hoffmann, and G. McDonald. 1994. Resistance to pyrethroids in Helicoverpa armigera (Lepidoptera: Noctuidae) from corn: adult resistance, larval resistance, and fitness effects. J. Econ. Entomol. 87: 1165-1171.
Gunning, R., G. Moores, and M. Balfe. 2015. Novel use of pyrethrum to control resistant insect pests on cotton. Acta Hort. 1073: 113-118. https://doi.org/10.17660/ActaHortic.2015.1073.16
Immaraju, J. A., T. D. Paine, J. A. Bethke, K. L. Robb, and J. P Newman. 1992. Western flower thrips (Thysanoptera: Thripidae) resistance to insecticides in coastal California greenhouses. J. Econ. Entomol. 85: 9-14.
Iqbal, K., R.H.J. Vekerk, M. J. Furlong, P. C. Ong, S. A. Rahman, and D. J. Wright. 1996. Evidence for resistance to Bacillus thuringiensis (Bt) subsp. kurstaki HD-1, Bt subsp. aizawai and Abamectin in field populations of Plutella xylostella from Malaysia. Pest Manag. Sci. 48: 89-97.
McGaughey, W. H. and M. E. Whalon. 1992. Managing insect resistance to Bacillus thuringiensis toxins. Science 258: 1451-1455.
Mordue, A. J. and A. J. Nisbet. 2000. Azadirachtin from the neem tree Azadirachta indica: its action against insects. An. Soc. Entomol. Bras. 29: 615-632.
Mota-Sanchez, D., R. M. Hollingworth, E. J. Grafius, and D. D. Moyer. 2006. Resistance and cross-resistance to neonicotinoid insecticides and spinosad in the Colorado potato beetle, Leptinotarsa decemlineata (Say) (Coleoptera: Chrysomelidae). Pest Manag. Sci. 62: 30-37.
Sauer, A. J., E. Fritsch, K. Undorf-Spahn, P. Nguyen, F. Marec, D. G. Heckel, and J. A. Jehle. 2017a. Novel resistance to Cydia pomonella granulovirus (CpGV) in codling moth shows autosomal and dominant inheritance and confers cross-resistance to different CpGV genome groups. PLoS ONE 12(6): e0179157.
Sauer, A. J., S. Schulze-Bopp, E. Fritsch, K. Undorf-Spahn, and J. A. Jehle. 2017b. A third type of resistance of codling moth against Cydia pomonella granulovirus (CpGV) shows a mixture of a Z-linked and autosomal inheritance pattern. Appl. Environ. Microbiol. AEM-01036.
Sayyed, A. H., D. Omar, and D. J. Wright. 2004. Genetics of spinosad resistance in a multi-resistant field-selected population of Plutella xylostella. Pest Manag. Sci. 60: 827-832.
Serebrov, V. V., O. N. Gerber, A. A. Malyarchuk, V. V. Martemyanov, A. A. Alekseev, and V. V. Glupov. Effect of entomopathogenic fungi on detoxification enzyme activity in greater wax moth Galleria mellonella L. (Lepidoptera, Pyralidae) and role of detoxification enzymes in development of insect resistance to entomopathogenic fungi. Anim. Human Physiol. 33: 581-586.
Siegwart, M., B. Graillot, C. B. Lopez, S. Besse, M. Bardin, P. C. Nicot, and M. Lopez-Ferber. 2015. Resistance to bio-insecticides or how to enhance their sustainability: a review. Front. Plant Sci. 6:381. https://doi.org/10.3389/fpls.2015.00381
Tabashnik, B. 1994. Evolution of resistance to Bacillus thuringiensis. Annu. Rev. Entomol. 39: 47-79.
Tabashnik, B. E., N. L. Cushing, N. Finson, and M. W. Johnson. 1990. Field development of resistance to Bacillus thuringiensis in diamondback moth (Lepidoptera: Plutellidae). J. Econ. Entomol. 83: 1671-1676.
Trisyono, A. and M. E. Whalon. 2000. Toxicity of neem applied alone and in combination with Bacillus thuringiensis to Colorado potato beetle (Coleoptera: Chrysomelidae). J. Econ. Entomol. 92: 1281-1288.
Whitehead, G. B. 1959. Pyrethrum resistance conferred by resistance to DDT in the blue tick. Nature 184: 378-379.
Wilson K., S. C. Cotter, A. F. Reeson, and J. K. Pell. 2003. Melanism and disease resistance in insects. Ecology Letters 4: 637-649.
Zhao, P., Z. Dong, J. Duan, G. Wang, L. Wang, Y. Li, Z. Xiang, and Q. Xia. 2012. Genome-wide identification and immune response analysis of serine protease inhibitor genes in the silkworm, Bombyx mori. PloS one 7: e31168.
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Amaryllis Help
Advice for the Home Gardener from the Help Desk of the
UC Master Gardener Program of Contra Costa County
MGCC's Help Desk Response: Thank you for contacting the UC Master Gardener Program Help Desk about the issues with your precious amaryllis bulb.
Two things that we discussed when reviewing the status of your prized amaryllis:
1. The container may be too large for the bulb. Amaryllis bulbs like to be potted in narrow and shallow containers.
2. The soil in your container appears to be moist. As mentioned in our previous emails on this subject, amaryllis do not like wet feet. They like to be kept in fast draining soil and only watered when the top two inches of soil feels dry.
We suggest that you correct these issues first in order to successfully grow your bulb.
For potted amaryllis, you don't need a great quantity of potting mix, because the bulbs grow and flower best when not over-potted. A 4-inch bulb only needs a pot 6 inches wide by 6 inches deep. The top one-third of the bulb should not be covered with soil, but left bare and protruding above the soil. As the bulb grows, move it gradually to larger pots, but still with a pot not much larger than the bulb itself. This prevents over-watering and waterlogged soil, which would rot the bulb.
We identified your flying insect as a fungus gnat. They are common in poorly drained soil or soil that is constantly wet (over-watered). Fungus gnats are a type of fruit fly that unfortunately either already exist in other potted plants in your home or patio, or may have come in with a new bag of potting soil.
Most of the fungus gnat's life is spent as a larva and pupa in organic matter or soil, so the most effective control methods target these immature stages rather than attempting to directly control the mobile, short-lived adults.
Physical and cultural management tactics—primarily the reductions of excess moisture and organic debris (such as dead leaves)—are key to reducing fungus gnat problems. We also suggest considering the use of the biological insecticide Bacillus thuringiensis (BT). This product is readily available in retail nurseries and garden centers, so it may be the most convenient for you to use. Just ask your favorite nursery for BT and then follow the package directions.
Here is additional information about fungus gnat control and prevention:
http://ipm.ucanr.edu/PMG/PESTNOTES/pn7448.html
Also, please see the attached information on proper use of pesticides:
http://ipm.ucanr.edu/PMG/PESTNOTES/pn74126.html
The beauty of amaryllis bulbs is that they are forgiving and easy to grow, so we want to help get you to the point of little worry and care for your grandmother's bulb!
If you have more questions, please do not hesitate to contact us again.
Help Desk of the UC Master Gardener Program of Contra Costa County (SLH)
Note: The UC Master Gardeners Program of Contra Costa's Help Desk is available year-round to answer your gardening questions. Except for a few holidays, we're open every week, Monday through Thursday for walk-ins from 9:00 am to Noon at 75 Santa Barbara Road, 2d Floor, Pleasant Hill, CA 94523. We can also be reached via telephone: (925)646-6586, email: ccmg@ucanr.edu, or on the web at http://ccmg.ucanr.edu/Ask_Us/ MGCC Blogs can be found at http://ccmg.ucanr.edu/HortCoCo/ You can also subscribe to the Blog (//ucanr.edu/blogs/CCMGBlog/).
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Corn Rootworms Outwit Genetically Modified Seed
My genealogical roots deep run through many a corn field since I was born from a Nebraska-farm girl mother and an Indiana-raised father. Field corn. Popcorn. Sweet corn. You name the corn, my relatives planted it. Several decades back I even grew ornamental corn one summer to decorate Christmas wreaths. So needless to say I was riveted to a recent Wall Street Journal article about how Mother Nature is outsmarting genetically modified corn seed. (Ian Berry, “Pesticides Make a Comeback”, Wall Street Journal, May 22, 2013, p. B-1.)
Seems that entomologists at the University of Illinois and Iowa State University found corn rootworms immune to Monsanto’s Bt (Bacillus thuringiensis) gene. That gene was originally designed to shield corn crops from this pest that feeds on leaves, tassels and silks; injures roots and can stunt or kill young shoots and plants.
The article also points out that last year America’s farmers planted 97 million acres in corn based on increasing prices and EPA approval touting reduced insecticide use that would give growers and farm workers “greater safety, protect water bodies from runoff and mitigate” harm to wildlife.
“Some of those gains are quickly being reversed,” said Michael Gray, a UI entomologist quoted in the story, who went on to say that next year over a quarter of corn farms plan to use insecticides as “cheap insurance.”
Makes senses now why sales are up for pesticide producers. To read the entire article, log on to WSJ online or review a similar story “Pesticides make a comeback against Monsanto seed” at
http://www.bignewsnetwork.com/index.php/sid/214688991/scat/3de2685784ae51b
Frankly, I wanted to know exactly what this crawly critter chomping on corn crops looked like. During my research of “corn rootworms,” I discovered crop damage is not limited to larvae but includes the adult — two familiar beetles often found in our own backyard vegetable patch that also feeds on cucurbits, legumes and grasses — the Western striped cucumber beetle (Acalymma trivittatum) and the Western spotted cucumber beetle (Diabrotica undecimpunctata undecimpunctata). (See UC IPM Pest Management Guidelines: Corn, plus UC ANR Publication 3443. Also UC IPM Pest Management Guidelines: Curcurbits, plus UC ANR Publication 3445.)
Photos below are of the Western striped cucumber beetle and the Western spotted cucumber beetle).
And there’s more. In fact, there’s also a banded cucumber beetle (Diabrotica balteata) and a spotted cucumber beetle (Diabrotica undecimpunctata howardi), also known as the Southern corn rootworm. In addition, there’s the Northern corn rootworm (Diabrotica barberi Smith & Lawrence), and the Western corn rootworm (Diabrotica virgifera virgifera LeConte).
When you compare the above two photos with photographs on Purdue University’s IPM website, you’ll notice that our Western spotted cucumber beetle looks identical to the Southern corn rootworm and that our Western stripped cucumber beetle appears the same or similar to the female Western corn rootworm. Plus, there’s a photo of the larvae -- the actual rootworm. Here’s the Purdue IPM link:
http://extension.entm.purdue.edu/fieldcropsipm/insects/corn-rootworms.php