Grower Notes and Pest News
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
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.
The western tarnished plant bug or lygus bug (Lygus hesperus) continues to be a major pest of strawberry on the Central Coast. Most growers typically rely on chemical or biological pesticides to manage pest. Some growers also use tractor-mounted vacuums to remove the pest, but the western tarnished plant bug is a major concern as it causes significant losses to marketable yields by deforming developing berries. Considering the status of the pest, having additional control options is critical both to reduce yield losses and also to strengthen the current integrated pest management (IPM) strategies.
A solar-powered UV light trap was reported to be a potential tool for controlling a variety of coleopteran, lepidoptera, hemipteran and other pests including the western tarnished plant bug. To evaluate its role as a potential IPM tool for strawberry pests a study was in conducted in fall-planted organic and conventional strawberry fields in Santa Maria.
Specifications of the trap: UV light trap known as Solar Powered Pest Control Machine (Model GFS-8) is manufactured by GreenFuture Equipment based in Sacramento, CA. It has a 30W solar power panel and a 12V battery to power a dual color UV light bulb and a rotating grill/grid for two nights on one day of charging. The grill surrounds the light produces 3600 volts of electricity with a surface area of 2.37 sqft and electrocutes insects as they are attracted to the light. Each light trap is supposed to cover 3-4 acres of area. A rubber flap brushes off insects into the container in the bottom of the trap as the grill rotates periodically.
Solar-powered UV light traps in conventional (left) and organic (middle) fields and insects collected in the container (right)
Experimental set up: One light trap was set up in a conventional strawberry field on West Main St (Manzanita Berry Farms) and another one in an organic field on Solomon Rd (Eraud Farms) in late March, 2017. Contents of the container were collected each week in a bag, taken back to the laboratory, and pest and beneficial insects were categorized and enumerated. Observations were made on 13 sampling dates between 2 May and 26 July, 2017.
Results: There were several groups of beneficial and pest insects were found attracted to the light trap. However, the western tarnished plant bugs were not seen throughout the observation period although field scouting indicated their presence. In general, the western tarnished plant bug infestations were lower this year and grower was able to manage pest populations by regular vacuuming. Pesticides were not used to control this pest during this period.
Among the pest insects trapped, corn earworm (Helicoverpa zea) adults were the only ones known to be a pest of strawberry in California. Insects that are generally recognized as pests included tiger moths, owlet moths, corn earworm adults, eucalyptus moths, sphinx moths, and mosquitoes while the beneficial insects included crane flies, lady beetles, parasitic wasps, neuropterons (such as lace wings), and soldier beetles. Some crane flies are important in the ecosystem as a prey for some animals and birds or through the activity of the larvae on decaying organic matter in the soil. However, their impact in strawberry is unknown.
Number of various pes (above) and benefifcial (below) insects collected on each sampling date
All pest and beneficial insects collected on different sampling dates (blue line: organic, orange line: conventional)
The number of both pest and beneficial insects was higher in the organic field than in the conventional field. Seasonal average of all insects per sampling date was 177 for the organic field and 98 for the conventional field. The proportion of the beneficial insects was about 27 in the organic field and 5 in the conventional field.
Seasonal average of various insects collected (Click on the image for a bigger version)
Although the role of UV light traps as a control option for the western tarnished plant bug could not be determined, it appeared to be a good tool for trapping corn earworm adults and other moths. This light trap could probably be useful for managing lepidopteran pests in strawberry or other corps.
Acknowledgements: Thanks to Dave Peck for his collaboration, GreenFuture Equipment for donating the light traps, and Maria Murrietta, Tamas Zold, and Chris Martinez for their technical assistance.
California offers ideal weather conditions for both nursery plant and strawberry fruit production. Variations in weather conditions in three strawberry production regions in California complement fruit production from each other and help avoid market glut. The warmer Oxnard area, the milder Santa Maria area, and the colder Watsonville area with minimal overlapping of their peak fruit production seasons allow yearlong strawberry production.
Weather influences on strawberry have been well documented in various strawberry producing regions across the world (Palencia et al., 2013; Li et al., 2010; Waister et al., 1972). Examples of key weather parameters correlated with strawberry yield include temperature, precipitation, solar radiation, relative humidity, and wind speed. Crop growth is weather dependent, thus, it is a common practice to estimate fruit yield based on weather variables. Since strawberry production spreads across 4–5 months, evaluating relationships between meteorological parameters and strawberry yield can provide valuable information and early indications of yield estimations that growers can utilize to their advantage. Objective of this research was to evaluate correlations of meteorological parameters on strawberry yield for Santa Maria region and to develop weather based statistical yield forecasting models for strawberries.
Strawberry yield data
Daily strawberry yield data for the Santa Maria region was obtained from published sources (California Strawberry Commission). This information is publically available and is originally compiled from the United States Department of Agriculture Market News/Fruits & Vegetables website. Daily strawberry yield data for the month of April through July were aggregated to weekly values. For this analysis we used weekly strawberry yield data for 2009 through 2015.
Weather data were obtained from the California Irrigation Management Information System (http://www.cimis.water.ca.gov/), a network of over 145 automated weather stations in California. Specific meteorological parameters used in this study were net radiation, air temperature (minimum and maximum), relative humidity (minimum and maximum), dew point temperature, soil temperature (minimum and maximum), vapor pressure (minimum and maximum), reference evapotranspiration, and average wind speed.
Weekly values of meteorological parameters from October of the year prior to harvest to February of current year of strawberry harvest were correlated with weekly strawberry yield from April through July and tested for significance. Each meteorological variable was correlated with strawberry yields from April to July. This thorough correlation analysis was done in order to understand influence of meteorological parameters on strawberry yield on a more detailed basis.
Fall and winter weather conditions during the vegetative growth period of strawberry have a significant influence on the fruit yields in the following spring and summer for the Santa Maria region. Results show that net radiation, relative humidity, vapor pressure, wind speed, and temperature showed significant correlations with strawberry yields at various temporal scales. In general, it was evident that many meteorological parameters during the early stages of strawberry growth and development phase exhibit statistically significant correlation with strawberry yields during the peak fruit production period. This finding is consistent with the findings of Lobell et al. (2006) for strawberry and other crops in California.
Table 1. Correlation matrix of monthly meteorological parameters and strawberry yields that were statistically significant.
Statistical yield model
Weather parameters that showed significant correlations were used to develop strawberry yield forecasting model. Instead of using weather parameters as explanatory variables, they were transformed into principal components to develop yield–forecasting models.
Figure 1 shows observed versus predicted yields for April (top) and June (bottom)
It is important to note that there are limitations on how much variability in yield data that can be explained by meteorological parameters as many other factors such as management practices, pests, diseases, varieties, other stress factors can also influence yield variability. Additionally, historic strawberry yield data provided an average estimate for the region and might not represent accurate observations.
These results demonstrate the potential to predict strawberry yield using weather variables relevant to the Santa Maria strawberry growing region. In order to make these results usable for decision–making, it could be refined to be utilized at the field scale. Additionally, skills of these models can be further improved by combining weather parameters and relevant physiological parameters of strawberry at the field scale.
A full version of this article (Pathak et al., 2016) can be viewed at: https://www.hindawi.com/journals/amete/2016/9525204/
Palencia, P., F. Martínez, J.J. Medina, and J. López– Medina. 2013. Strawberry yield efficiency and its correlation with temperature and solar radiation. Hortic. Bras.31(1): 93–99
Li, H., T. Li, R.J. Gordon, S.K. Asiedu, and K. Hu. 2010. Strawberry plant fruiting efficiency and its correlation with solar irradiance, temperature and reflectance water index variation. Environ. Exp. Bot. 68, 165–174.
Waister, P. D. 1972. Wind as a limitation on the growth and yield of strawberries. hort. Sci. 47: 411 – 418.
Lobell, D.B., K. Cahill, and C. Field. 2006. Weather–based yield forecasts developed for 12 California crops. California Agriculture, 60(4): 211–215.
Pathak, T.B., Dara, S.K., Biscaro, A. 2016. Evaluating correlations and development of meteorology based yield forecasting model for strawberry. Advances in Meteorology, vol. 2016, article ID 9525204. doi:10.1155/2016/9525204
Species of the genus Entomophthora cause epizootics in various insects around the world, but such infections are less common in California. Burger and Swain (1918) reported Entomophthora chromaphidis infections in the walnut aphid, Chromaphis juglandicola, in Southern California. With infections as high as 95%, the fungus was a significant mortality factor in aphid populations. In 2011, nearly a century after the epizootics in the walnut aphid, a single strawberry aphid, Chaetosiphon fragaefolii, was found infected with a species of Entomophthora in an organic strawberry field (Eraud Farms) in Santa Maria. The strawberry aphid is an occasional and minor pest in strawberry in California.
Entomophthora planchoniana infection in strawberry aphid
Attempts to in vitro culture the fungus were unsuccessful, but microscopic measurements of conidial size and shape indicate that the causal agent could be Entomophthora planchoniana. Bell-shaped conidia measured 17.3 μm or micrometers (14.8-20.1) long and 14.6 μm (12.7-17.7) wide (based on the measurements of 100 conidia) and had a broad base (papilla) and a pointed apex. Conidia also appeared to have 4-6 nuclei.
E. planchoniana and E. chromaphidis are closely related and were previously considered as synonymous species (MacLeod et al., 1976; Waterhouse and Brady, 1982). Humber and Feng (1991) later described these two as separate species due to the variation in conidial size, geographic distribution, host range, in vitro culturing techniques, and other characteristics. While E. planchoniana is commonly found in Europe, E. chromaphidis is reported elsewhere.
The following are the characters of E. chormaphidis and E. planchoniana from Keller's (2002) description of 22 species of Entomophthora.
First found in walnut aphids in California. Primary conidia are 11-14 μm long and 10-11 μm wide with 4-6 nuclei and contain a single large oil globule. Resting spores are 30 μm. Insect host is attached to the plant surface with fungal rhizoids (bundles of modified hyphal bodies).
First found on unidentified aphids on elder in western Europe. Primary conidia are 15-20 μm long and 12-16 μm wide with 4-11 nuclei [species description has 4-11 and key has 5-9 nuclei in Keller (2002)]. Resting spores are 31-38 μm. The fungus produces rhizoids to attach the host insect to the plant surface.
Keller (2002) noted an overlap in the number of nuclei between these two species and suggested the size of primary conidia as the distinguishing character. Hence the fungus found in the strawberry study is considered E. planchoniana, which was first reported in 1948 in the strawberry aphid, then known as Pentatrichopus fragariae (Petch, 1948; Leatherdale, 1970). Cédola and Greco (2010) reported E. planchoniana as a major mortality factor of the strawberry aphid in Argentina.
Life cycle of E. planchoniana
The infection process starts when a conidium (single spore) comes in contact with the insect cuticle, produces a germ tube and gains entry into the insect body with the help of a penetration peg and cuticle degrading enzymes. The fungus multiplies inside the insect body as protoplasts (cells without a cell wall) and invade the tissues. Vegetative growth stops when nutrients are depleted and the insect is dead. The fungus then produces conidiophores that emerge from the cuticle, produce bell-shaped primary conidia that are forcibly discharged. A halo of protoplasm (cellular contents) is often seen around the primary conidium, which may either produce the germ tube to cause infection or a secondary conidium. Secondary conidia are smaller than primary conidia, have rounded or less pointed apices, and more rounded basal papillae.
This is the first report of the occurrence of E. planchoniana in the strawberry aphid in California. Although strawberry aphid is not an important pest in California and only one infected aphid was found, this finding is important to record the distribution of E. planchoniana.
Burger, O. F. and A. F. Swain. 1918. Observations on a fungus enemy of the walnut aphis in Southern California. J. Econ. Entomol. 11: 278-289.
Cédola C. and N. Greco. 2010. Presence of the aphid, Chaetosiphon fragefolii, on strawberry in Argentina. J. Ins. Sci. 10: 1-9.
Humber, R. A. and M.-G. Feng. 1991. Entomophthora chromaphidis (Entomophthorales): the correct identification of an aphid pathogen in the pacific northwest and elsewhere. Mycotaxon 41: 497-504.
Leatherdale, D. 1970. The arthropod hosts of entomogenous fungi in Britain. Entomophaga 15: 419-435.
MacLeod, D. M., E. Müller-Kögler, and N. Wilding. 1976. Entomophthora species with E. muscae-like conidia. Mycologia 68: 1-29.
Petch, T. 1948. A revised list of British entomogenous fungi. Trans. Brit. Mycol. Soc. 31: 286-304.
Waterhouse, G. M. and B. L. Brady. 1982. Key to the species of Entomophthora sensu lato. Bull. Brit. Mycol. Soc. 16: 113-143.