- Author: Surendra K. Dara
- Author: Suchitra S. Dara
- Author: Stefan Jaronski, USDA-ARS
The western Grapeleaf skeletonizer (WGLS), Harrisina metallica Stretch (Lepidoptera: Zygaenidae), previously known to cause severe defoliation to vineyards and backyard grapevines appears to be re-emerging in California. Since its first detection in San Diego in 1941, WGLS spread through commercial vineyards and backyard grapes becoming a serious problem. Although two biological control agents from Arizona and Mexico were introduced in California for WGLS control, a naturally occurring granulovirus (Harrisina brillians granulovirus) nearly eradicated WGLS populations and kept them under control. WGLS has not been a problem especially in conventional vineyards. However, based on some unpublished observations, WGLS populations are emerging in organic vineyards and backyard grapevines.
WGLS lives up to its name by skeletonizing and defoliating grape leaves. Organic vineyards are especially at risk and uncontrolled populations can destroy vineyards resulting in significant losses. Metallic bluish or greenish black moths lay barrel shaped yellowish eggs on the lower side of the leaves. There are five larval instars. Early instars are cream colored and develop black and purple bands in later stages. Pupation occurs in a whitish cocoon. Upon hatching, larvae start feeding side by side in a row on the lower side of leaf. Damage by younger larvae appears as whitish leaf area containing veins and the upper cuticle, which eventually turn brown. Older larvae skeletonize leaves leaving larger veins. Larvae may also feed on fruit leading to bunch rot. Severe damage can cause defoliation and sunburn of the exposed fruit.
A study was conducted to evaluate the efficacy of six non-chemical control options that included formulations of spinosad, two subspecies of Bacillus thuringiensis, and a botanical insecticide/growth regulator along with two unformulated entomopathogenic fungal isolates native to California. Larvae were collected from an infested, untreated backyard grapevine and maintained in one gallon plastic tubs with screened lids on infested leaves. Fresh, untreated grape leaves from uninfested vines were provided daily for 3 days before starting the assay. For each treatment, five 4-5 instar larvae were placed on a grape leaf disc (rinsed in water and dried) in a Petri plate (100 mm dia) with a moist filter paper. Larvae were treated by spraying 1 ml of the treatment solution (containing Dyne-Amic as a surfactant at 0.125% vol/vol). Application rates for commercial formulations were determined based on label recommendations for 100 gallons of spray volume. Entomopathogenic fungal concentrations were also determined based on the label rates for similar commercial products. Treatments were replicated four times and the assay was conducted twice. Larval mortality was observed daily and dead larvae were removed and incubated separately. Fresh leaf discs were provided as needed to the remaining larvae. Actual and corrected (for control mortality) total mortality were calculated.Data were arcsine-transformed for statistical analysis and significant means were separated using Tukey's HSD test.
Both cumulative daily mortality and total mortality significantly (P < 0.0001) differed among treatments. Entrust and M. anisopliae resulted in the highest mortality followed by B. bassiana, Neemix, and Agree. In general, feeding reduced or ceased in all larvae following treatment and could have contributed to a lower mortality in B. thuringiensis treatments. Entomopathogenic fungi emerged from all the cadavers from respective treatments. Microbial and botanical options provided good control of WGLS. These non-chemical alternatives can be effectively used in both organic and conventional vineyards. California isolates of B. bassiana and M. anisopliae demonstrated good control efficacy and the potential to be developed as microbial pesticides.
Acknowledgements: Thanks to the technical assistance of Alor Sahoo in carrying out these assays, and Certis and Corteva for providing the pesticide formulations.
Federici, B. A. and V. M. Stern. 1990. Replication and occlusion of a granulosis virus in larval and adult midgut epithlium of the western grapeleaf skeletonizer, Harrisina brillians. J. Invertebr. Pathol. 56: 401-414.
- Author: Melissa O'Neal, Marrone Bio Innovations
- Author: Surendra K. Dara
Biopesticides are based on naturally occurring microorganisms, plant extracts or other materials and are regulated by the United States Environmental Protection Agency (EPA)'s Biopesticide Division. Biopesticides have been safely used for over 63 years and are generally subjected to reduced regulation compared to conventional chemical pesticides.
Biopesticides can be developed from plant extracts or entomopathogenic microorganisms. Graphic: Surendra Dara
The active ingredient in microbial pesticides consists of a microorganism, such as a bacterium, fungus, nematode, protozoan or virus. While microbials are capable of assisting in the management of many different types of pests, each type of microorganism tends to be relatively specific for a target pest or group of pests. Biochemical pesticides are based on naturally occurring substances, which function by providing pest management through non-toxic mechanisms. Biochemical pesticides may function by disrupting or interfering with mating, such as in the case of insect sex pheromones or various plant extracts which serve as insect attractants used with traps. Conventional pesticides, by contrast, are generally synthetic materials that directly kill or inactivate the pest (Leahy et al., 2014).
Typically, samples of microorganisms or infected arthropods are collected from natural environments. Samples are taken to the laboratory and plated on media; thereafter, various colonies form from the collected samples. Individual colonies of interest may be selected, suspended, and examined for pesticidal activity during laboratory bioassays (Taylor, 1988). As part of the laboratory bioassay process, researchers screen candidates against a number of potential targets, which may vary widely, depending upon institutional goals and availability.
A key initial task is identification and characterization of the pesticidal compounds sourced from the plants or microbes collected in natural settings (Strobel and Daisy, 2003). Part of this process involves isolating and eliminating any compounds which have potential human health implications or may negatively impact non-targets organisms (USDA, 2017b). Additionally, analytical assays based on bioactive chemistry are developed to ensure quality control during the manufacturing process (Strobel and Daisy, 2003).
Several steps are involved with product and process development. First, user-friendly formulations are developed in both lab and pilot facilities. Next, manufacturing processes are developed and scaled in arenas including lab, pilot, and manufacturing facilities (Strobel and Daisy, 2003). Thereafter, field studies are conducted and data are gathered for the regulatory submissions which support product registration (USDA, 2017a).
Biopesticide registration process
A special committee has been established within the EPA due to the fact that it is often challenging to determine whether a substance meets the criteria for classification as a biochemical pesticide (Leahy et al., 2014). The Biopesticide Pollution Prevention Division (BPPD) of the EPA is charged with data review required for registration. Requirements for registration include acute studies consisting of oral, inhalation, intravenous, and dermal tests, in addition to eye and skin studies in rodents. A product chemistry review involving a five-batch analysis is also required by BPPD. Microbiology and quality control investigations assure that material is free of human pathogens. Ecological effects, including impact on non-target birds, fish, Daphnia, honeybees, lacewings, ladybeetles, and parasitic wasps is additionally determined. The review process is taken one step further during the endangered species review. Finally, the matter of the Exemption from Tolerance Petition for Food Use is addressed (EPA, 2017). It should be noted that efficacy data are required in addition to the aforementioned topics when attempting to register a new biopesticide in California (CDPR, 2017). There are several examples of successful pesticides which are sourced from natural products and registered as chemical pesticides (Fig. 1).
Fig. 1. Chemical pesticides developed from natural sources. Graphic: Melissa O'Neal
Abamectin is an insecticide/miticide derived from Streptomyces avermitilis, a microorganism found in soil. Its mode of action involves interference with neurotransmission (CDPR, 1993). Tebufenozide is an insect growth disruptor which interferes with insect molting hormones (Smagghe et al., 2012). The spinosyns are a family of chemicals produced by fermentation of Saccharopolyspora bacteria which are toxic due to disruption of neurotransmitters in both target and non-target organisms (Kirst, 2010). Azoxystrobin is a synthetic material derived from phytotoxic compounds which naturally occur in the mushrooms Oudemansiella mucida and Strobilurus tenacellus. Its mode of action is disruption of energetic reactions involving ATP synthesis (AgChemAccess, 2015). Finally, pyrethrins are naturally occurring materials derived from the chrysanthemum (Chrysanthemum cinerariaefolium) flowers and acts as a contact nerve poisons (Extoxnet, 1994).
The following tables 1-5 provide an overview of some of the commercial biopesticides currently registered in the US and other countries for controlling insects, mites, plant pathogenic fungi, and plant parasitic nematodes.
Table 1. Microbial insecticides andacaracides.
Table 2. Plant extract and oil insecticides and acaricides.
Table 3. Microbial fungicides.
Table 4. Non-microbial fungicides.
Table 5. Bionematicides.
AgChemAccess. 2015. Azoxystrobin. http://www.agchemaccess.com/Azoxystrobin.
(CDPR). California Department of Pesticide Regulation. 1993. Abamectin Avert Prescription Treatment 310 (Section 3 Registration) Risk Characterization Document. http://www.cdpr.ca.gov/docs/risk/rcd/abamectin.pdf
(CDPR). California Department of Pesticide Regulation. 2017. How to apply for pesticide product registration. http://www.cdpr.ca.gov/docs/registration/instructions.htm
(EPA). U.S. Environmental Protection Agency. 2017. Biopesticides. https://www.epa.gov/pesticides/biopesticides#what
Extoxnet. 1994. Pesticide information profile: Pyrethrins. http://pmep.cce.cornell.edu/profiles/extoxnet/pyrethrins-ziram/pyrethrins-ext.html
Kirst, H.A. 2010. The spinosyn family of insecticides: realizing the potential of natural products research. J Antibiot 63(3): 101-11. doi: 10.1038/ja.2010.5.
Leahy, J., M. Mendelsohn, J. Kough, R. Jones, and N. Berckes. 2014. Biopesticide oversight and registration at the U.S. Environmental Protection Agency. In Biopesticides: State of the Art and Future Opportunities; Coats, et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2014.
Smagghe, G., L.E. Gomez, and T.S. Dhadialla. 2012. Insect growth disruptors. Adv Ins Phys 43: 1-552.
Strobel, G. and B. Daisy. 2003. Bioprospecting for microbial endophytes and their natural products. Microbiol Mol Bio Rev 67(4): 491-502.
Taylor, J.K. 1988. Quality assurance of chemical measurements. Chelsea, MI: Lewis.
(USDA). United States Department of Agriculture. 2017a. About AMS.
(USDA). United States Department of Agriculture. 2017b. USDA FY 2016 avoiding harm from invasive species (USDA Do No Harm 2016 Report, Part 2). https://www.invasivespeciesinfo.gov/docs/resources/usdanoharm20170119.docx
- Author: Surendra K. Dara
- Author: Sumanth S. R. Dara
- Author: Suchitra S. Dara
- Author: Ed Lewis
Eggs, nymphs, and adult silverleaf whitefly on zucchini. Photo by Surendra Dara
A study was conducted in the summer of 2017 to evaluate the efficacy of various chemical, botanical, and microbial pesticides against arthropod pests on zucchini. Zucchini plants initially had a high aphid infestation, but populations gradually declined due to natural control by lady beetle activity. However, heavy silverleaf whitefly (Bemisia tabaci) infestations developed by the time the study was initiated. Other pests that were present during the study period were aphids (possibly melon aphids), the western flower thrips (Frankliniella occidentalis), and the pacific spider mite (Tetranychus pacificus).
Pacific spider mite (egg, male, and females), western flower thrips larva, and unknown aphids on zucchini. Photo by Surendra Dara
Experiment was conducted using a randomized complete block design with 10 treatments. Each treatment had two 38” wide and 300' long rows of zucchini replicated four times. Treatments included i) untreated control, ii) Sivanto 200 SL (flupyradifurone) 14 fl oz/ac, iii) Sequoia (sulfoxaflor) 2.5 fl oz/ac, iv) Venerate XC (heat-killed bacterium, Burkholderia rinojensis strain A396) 4 qrt/ac, v) PFR-97 20% WDG (entomopathogenic fungus, Isaria fumosorosea Apopka strain 97) 2 lb/ac, vi) I1800AA (undisclosed botanical extract) 10.3 fl oz/ac, vii) I1800A 12.7 fl oz, viii) I1800A 17.1 fl oz, ix) I1800A 20.5 fl oz, and x) VST-00634LC (based on a peptide in spider venom) 25%. A spray volume of 50 gpa for all treatments except for VST-00634LC, which had 25 gpa. Treatments were applied on 28 August and 4 September, 2017 using a tractor-mounted sprayer with three Teeject 8003vs flat spry nozzles that covered the top and both sides of each bed.
Pest populations were counted before the first spray application and 4 days after each application. On each sampling date, one mid-tier leaf was collected from each of the five randomly selected plants within each plot. A 2-square inch disc was cut out from the middle of each leaf and the number of aphids, eggs and nymphs of silverleaf whitefly, larvae of western flower thrips, and eggs and mobile stages of pacific spider mite were counted under a dissecting microscope. Data were analyzed using Statistix software and Tukey's HSD test was used to separate significant means.
Spraying, sampling, and counting
Efficacy varied among different treatments and for different pests.
Aphid: There was a general decline in aphid populations during the study period and there was no difference (P > 0.05) among the treatments (Fig. 1).
Fig. 1. Aphid numbers and percent change from pre-treatment counts
Western flower thrips: Nymphal numbers declined in most of the treatments during the observation period (Fig. 2). However, significant differences (P = 0.0220) only after the second spray application where Sivanto treatment had significantly fewer thrips than Venerate treatment (Fig. 2). There was a 92.5% decline by the end of the study, compared to the pre-treatment counts, from PFR-97 application, followed by 88.1% decline in Sivanto, 85.4% in VST-00634, and 82.9% in I1800AA at 10.3 fl oz.
Fig. 2. Western flower thrips larvae and percent change from pre-treatment counts
Pacific spider mite: There was an increase in mite eggs in all treatments after the first spray application followed by a decline after the second one without significant differences (P > 0.05) (Fig. 3) Similar trend was also seen in mobile stages in some treatments. Number of mobile stages was significantly different (P = 0.0025) only after the first spray where untreated control, PFR-97, Venerate, and I1800AA at 20.5 fl oz had the lowest. When percent change in egg numbers from the pre-treatment counts, only I1800AA treatments reduced egg numbers after the second spray with a 33.8% decline at 10.3 fl oz rate, 35.7% at 20.5 fl oz, and 60% at 17.1 fl oz. There was also a decline in the mobile stages after the second spray with 54.1% reduction in untreated control to 67.7% in PFR-97 treatment.
Fig. 3. Pacific spider mite egg and mobile stages and percent change from pre-treatment counts
Silverleaf whitefly: There was a general increase in the egg and nymphal stages of whitefly during the study (Fig. 4). Significant differences were seen pre-treatment counts of egg (P = 0.0330) and nymphal stages (P = 0.0011), and after the second spray in nymphal stages (P = 0.0220). Compared to the untreated control, both Sivanto and Sequoia resulted in a significant reduction in egg numbers after the first spray, whereas Sequoia, Venerate, and I1800AA at 20.5 fl oz reduced nymphal stages after the second spray. When the percent change from the pre-treatment counts was compared, only Sivanto and Sequoia reduced whitefly egg numbers after both sprays. There was also a reduction in eggs after the first spray from I1800AA at 17.1 fl oz. However, there was a reduction in nymphal stages after the first spray in Sivanto, I1800AA at 17.1 fl oz, and VST-00634, and after the second spray in Sivanto, Sequoia, Venerate, and I1800AA at 17.1 and 20.5 fl oz.
Fig. 4. Silverleaf whitefly egg and nymphal stages and percent change from pre-treatment counts
All arthropod pests: When all data were combined for different pests and their life stages, Sivato, Sequia, and PF-97 resulted in a significant (P = 0.0001) decline in pest numbers compared to untreated control after the first spray. Only Sivanto and Sequoia caused such a reduction (P = 0.0048) after the second spray.
Fig. 5. All arthropod pest numbers and percent change from pre-treatment counts
In general, both the chemical pesticides (Sivanto and Sequoia) provided a very good pest control. The efficacy of the botanical extract was moderate to good depending on the pest, life stage, or the application date. Spider venom-based product also provided a good control while microbial products had a moderate impact. Although chemical pesticides appeared to be very efficacious, non-chemical alternatives were also effective. It is important to consider all these options to apply in combinations or rotations to obtain desired pest suppression without posing the risk of insecticide resistance.
Acknowledgements: Thanks for the financial support of Arysta LifeScience, CertisUSA, Dow AgroSciences, and Vestaron, and the technical assistance of Neal Hudson.
- Author: Melissa O'Neal, Marrone Bio Innovations
- Author: Surendra K. Dara
Biopesticide refers to a pesticide which originates from animals, microorganisms, or plants. In addition to preventing yield losses through pest and disease control, biopesticides improve environmental and human health by contributing to the reduction of chemical pesticides as well as by improving the quality of produce (Popp et al., 2012). Additionally, these products have the potential to improve harvest and shipping flexibility, assist with environmental stewardship, and assist growers to achieve sustainability goals. Biopesticides are also important tools in integrated pest management (IPM) programs and reducing the risk of resistance to chemical pesticides (Pretty and Bharucha, 2015), improving worker safety through short restricted entry intervals (Valland, n.d.), conserving natural enemies, and maintaining environmental health (EPA, 2017a).
Biopesticides are inherently less toxic than conventional pesticides. Most affect only the target pest and closely related organisms, in contrast to broad spectrum conventional pesticides that may affect nontarget organisms such as beneficial insects, birds, wildlife, aquatic animals, and mammals. The majority of biopesticides often rapidly decompose, resulting in decreased exposure as well as preventing many pollution problems commonly associated with conventional pesticides. Although relatively safer than chemical pesticides, users or applicators should follow safety guidelines and wear personal protective equipment according to the label directions (EPA, 2017a). It is also important to follow guidelines for spray volume, application rates, droplet size, water pH, compatibility with tank-mix partners, time and frequency of application, and other details to ensure efficacy of the biopesticides (van Zyl et al., 2010; Wang & Liu, 2007; Whitford et al., 2009).
Biopesticides use has been increasing in the recent years. They can be used as standalone treatments or combined or rotated with other pesticides in both organic and conventional production systems. The fact that there are no residues is a huge benefit for exported commodities, as maximum residue limit issues continue to be a challenge in this arena (Berger, 2013).
In expanding upon the role of biopesticides in biocontrol, the topic of resistance management is a key consideration. Pest resistance to conventional chemical pesticides is a significant concern. Scientific research has repeatedly demonstrated that continuous use of the same class of pesticides, especially those reliant on a single mode of action, will result in the emergence of a pest population resistant to those products (Osteen et al., 2012). Populations of insect pests, plant pathogens, nematodes, and weeds all have the ability to develop resistance quickly, even to different types of functionally similar chemistries. This phenomenon is called cross-resistance and is caused by multi-chemistry detoxification mechanisms present in many pest populations (Horowitz and Ishaaya, 2009).
Because of the increasing number of novel, low-impact chemistries available, educators and growers have additional tools to manage resistance within IPM programs (EPA, 2017a). Biopesticides have long been used in combination with synthetic chemistries to provide the basis for excellent control programs that effectively manage resistance. Additionally, they typically have modes of action that are different from synthetic pesticides and do not rely on a single target site for efficacy. Properly used, these products have the potential to extend the effective field life of all products by curtailing the development of resistant pest populations (Horowitz and Ishaaya, 2009).
According to the United States Environmental Protection Agency (EPA), “IPM is an effective and environmentally sensitive approach to pest management that relies on a combination of common-sense practices” (2017b, p. 1). The University of California Statewide Integrated Pest Management Program (UCIPM) (2017) defines the IPM approach as combining prevention, cultural, physical, biological and chemical means to control pests, all the while minimizing economic, public health, beneficial as well as non-target organism, and environmental risks. Biopesticides are noted among the low-risk and most highly effective tools for achieving crop protection in IPM systems. The challenges of farming require that IPM systems actively integrate multiple management approaches to balance optima productivity with sustainability (BPIA, 2017).
Biopesticides should be considered as a component of a holistic total program and used at an appropriate time and pest density. Today, many forward looking IPM professionals are incorporating biopesticides into traditionally conventional pest management strategies (EPA, 2017b). However, education and training are needed to address biopesticide best use practices, the methods of integrating them into IPM programs; as well as instruction to promote an understanding of their unique modes of action (EPA, 2017b). Part of the educational process involves research through fair and realistic field trials that evaluate biopesticides both as standalone treatments as well as in combination and rotation with other options with an objective of improving IPM practices (Abler et al., 1992; Kumar and Singh, 2015). All of these learning experiences are useful in demonstrating the science of biopesticide use and establishing best use practices. A better understanding of biopesticide potential and the mode of action of different active ingredients, increased grant support to promote biopesticide research, and productive grower-industry-researcher collaborations to generate applied research data and design IPM strategies are necessary to make the best use of biopesticides and for environmental sustainability.
Abler, D.G., G.P. Rauniyar, and F.M. Goode. 1992. Field trials as an extension technique: The case of Swaziland. NJARE 21(1): 30-35.
Berger, L. 2013. MRL issues and international trade commodity perspectives, pp 3-48. In Proceedings: Idaho Pesticide MRL Workshop, 2 December 2013, Boise, ID. AgBusiness Resources, Visalia, CA.
(BPIA). Biological Products Industry Alliance. 2014. Biopesticides in a program with traditional chemicals offer growers sustainable solutions. http://www.bpia.org/wp-content/uploads/2014/01/grower-final.pdf
(BPIA). Biological Products Industry Alliance. 2017. Benefits of biological products. http://www.bpia.org/benefits-of-biological-products/
(EPA). U.S. Environmental Protection Agency. 2017a. Biopesticides. https://www.epa.gov/pesticides/biopesticides#what
(EPA). U.S. Environmental Protection Agency. 2017b. Integrated pest management principles. https://www.epa.gov/safepestcontrol/integrated-pest-management-ipm-principles
Horowitz, A. and I. Ishaaya. (2009). Biorational control of arthropod pests: Application and resistance management. Springer, New York, NY.
Kumar, S., and A. Singh. 2015. Biopesticides: Present status and the future prospects. J Fertil Pestic 6: e129. doi:10.4172/2471-2728.1000e129
Osteen, C., J. Gottlieb, and U. Vasavada (eds.). 2012. Agricultural Resources and Environmental Indicators. EIB-98, U.S. Department of Agriculture, Economic Research Service, August 2012.
Popp, J., K. Peto, and J. Nagy. 2012. Pesticide productivity and food security: A review. Agron Sustain Dev 33: 243–255. DOI 10.1007/s13593-012-0105-x.
Pretty, J. and Z.P. Bharucha. 2015. Integrated pest management for sustainable intensification of agriculture in Asia and Africa. Insects 6(1): 152–182.
(UCIPM). University of California Statewide Integrated Pest Management Program. 2017. What is integrated pest management (IPM)? http://www2.ipm.ucanr.edu/WhatIsIPM/
Vallad, G.E. n.d. Use of biopesticides for the management of vegetable diseases. University of Florida Gulf Coast Research and Extension Center. http://ipm.ifas.ufl.edu/pdfs/Bio-Pesticides_Slides_IPM_site.pdf
van Zyl, S.A., J. Brink, F.J. Calitz, S. Coertze, and P.J. Fourie. 2009. The use of adjuvants to improve spray deposition and Botrytis cinerea control on Chardonnay grapevine leaves. Crop Prot 29(1): 58-67. https://doi.org/10.1016/j.cropro.2009.08.012
Wang, C.J. and Z.Q. Liu. 2007. Foliar uptake of pesticides: Present status and future challenge. Pest Biochem Phys 87(1): 1-8. https://doi.org/10.1016/j.pestbp.2006.04.004
Whitford, F., D. Penner, B. Johnson, L. Bledsoe, N. Wagoner, et al. 2009. The impact of water quality on pesticide performance. Purdue Extension Publication PPP-86. https://www.extension.purdue.edu/extmedia/ppp/ppp-86.pdf
- Author: Surendra K. Dara
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.
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 (http://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 (http://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.