Spotted lanternfly (Lycorma delicatula). Photo by Surendra Dara

The spotted lanternfly (SLF) [Lycorma delicatula (Hemiptera: Fulgoridae)] is an invasive planthopper, which causes a significant damage to apples, grapes, stone fruit, trees used for timber, and other hosts (Dara et al. 2015).  Native to China, SLF was first reported in 2014 in Pennsylvania and has been rapidly spreading in the eastern United States and moving westward.  California has 22 cultivated and about 70 wild hosts of SLF and include several high value crops such as apples, cherries, grapes, and plums.  The tree-of-heaven, an invasive species, is a favorite host of SLF and is widely distributed in California.  SLF is also a nuisance pest with 100s or 1000s of individuals infesting landscape trees and hosts in residential areas.  This pest deposits eggs on inanimate objects such as vehicles, furniture, stones, and packages and thus spread to other areas through the movement of these objects.  Awareness of the pest and its damage potential, ability of Californians to recognize and report the pest if found, and the knowledge of control practices will help prevent accidental transportation of eggs or other life stages from the infested areas to California and prepare the citizens to take appropriate actions.  Outreach efforts have been made in California since 2014 through extension articles, presentations at extension meetings, videos, social media posts, and personal communication (Dara, 2014).

Wakie et al. (2020) modeled the establishment risk of SLF in the United States and around the world and indicated that many coastal regions and the Central Valley of California are among the high-risk areas.  Considering the risk to several high-value commodities and the presence of several wild hosts that are distributed all over California, mapping of the risk-prone areas based on the cultivated hosts, their acreage and value in different counties, and the distribution of wild hosts was done to help both growers and other Californians to prepare for potential invasion of SLF. 

Methodology

The list of SLF hosts is continuously evolving with host specificity studies in various places.  Based on two published resources (Dara et al. 2015; Barringer and Ciafré 2020), 22 cultivated and 70 wild hosts appear to be present in California.  Plant species that support some of the feeding life stages or all life stages were included in preparing these lists.  The cultivated hosts include apples, apricots, basil, blueberries, butternut, cherries, cotton, grapes, hibiscus, hops, mock orange, nectarine, peaches, pears, persimmon, plums, pomegranates, roses, soybean, sponge gourd, tea, and walnuts; and the wild hosts include Acacia sp., American hazelnut, Amur corktree, American linden, American sycamore, arborvitae, Argentine cedar, Asian white birch, bee balm, big-toothed aspen, black gum, black hawk, black locust, black walnut, Bladder senna, boxelder, chestnut oak, chinaberry tree, Chinese boxwood, Chinese juniper, Chinese parasol tree, Chinese wingnut, devilwoods, dogwood, Eastern white pine, edible fig, false spiraea, fireweed, five-stamen tamarisk, flowering dogwood, Forsythia, Glossy privet, greater burdock, grey alder, hemp, hollyhocks, honeysuckle, hornbeam, Japanese angelica, Japanese boxwood, Japanese maple, Japanese snowball, Japanese zelkova, jujubes, Kobus magnolia, Northern spicebush, Norway maple, lacquer tree, perennial salvia, Persian silk tree, plane tree, Poinsettia, poplars, princess tree, red maple, sapphire dragon tree, sassafras , sawtooth, serviceberry, silver maple, slippery elm, snowbell, staghorn sumac, sugar maple, tree-of-heaven, tulip tree, Virginia creeper, white ash, wild grape, and willows.

The summary of county crop reports from the California Department of Food and Agriculture (CDFA 2018) was used to determine the value and acreages of the cultivated hosts.  To determine the distribution of wild hosts various online resources were used.  SLF risk levels were determined as very low, low, moderate, high, and very high for the number of hosts, acreage and value of each cultivated host, and other such parameters within each county.  The highest risk value within each parameter was used to determine ‘very high' category and 4/5, 3/5, 2/5, and 1/5 were used for high, moderate, low, and very low categories, respectively.  In other words, 0-20% risk was considered very low, 21-40% as low, 41-60% as moderate, 61-80% as high, and 81-100% as very high for each measured parameter.  Data were entered into a spreadsheet and maps were generated using QGIS open-source cross-platform geographic information system application.

Risk-prone areas in California

The following maps show areas in California that are prone to SLF risk based on the distribution of cultivated and wild hosts, and the acreage and value of important cultivated crops.

Areas where grapes, the most important cultivated host of SLF, are grown in California

Distribution of tree-of-heaven, an invasive tree and the favorite host of SLF

 

Based on these maps, the entire state of California is at some level of risk.  In addition to the commercially produced crops, several backyard or landscape plant species such as roses, grapes, peaches, plums, and others are present throughout the state and can harbor SLF.  Such host plants in residential and urban landscapes can serve as SLF sources for commercial crops.  The tree-of-heaven is present throughout California and several such uncultivated hosts can serve as sources of undetected infestations.  While researchers are working on appropriate biocontrol solutions such as releasing natural enemies, other control options such as synthetic and microbial pesticide applications, sticky traps, removal of egg masses and wild hosts, and other strategies can help manage SLF.  In the meantime, Californians will benefit by knowing about this pest and its potential risk to the state.  The ability to identify, destroy or capture, and report the pest to county and state departments or University of California Cooperative Extension offices will help prevent or delay SLF invasion and spread in California.

Conclusion

California is at the risk of SLF invasion and spread.  Depending on the number of cultivated crops, their acreage, value, and the distribution of wild hosts, the risk level varies in various counties throughout the state. Outreach efforts are helping to alert Californians about SLF and its damage to cultivated crops and nuisance in urban and residential areas.

Additional resources

Refer to https://ucanr.edu/spottedlanternfly for additional information about the pest.  If you happen to see this pest in California, please contact your local Agricultural Commissioner, California Department of Food and Agriculture, or UC Cooperative Extension office to report.

Acknowledgments

Thanks to the California Department of Food and Agriculture for funding this study.

References

Barringer, L. and Ciafré, C. M.  2020.  Worldwide feeding host plants of spotted lanternfly, with significant additions from North America.  Environ. Entomol. 49: 999-1011.

CDFA (California Department of Food and Agriculture).  2018.  California County Agricultural Commissioners' Report Crop Year 2016-2017 (https://www.cdfa.ca.gov/statistics/pdfs/2017cropyearcactb00.pdf).

Dara, S. K.  2014.  Spotted lanternfly (Lycorma delicatula) is a new invasive pest in the United States.  UCANR eJournal Pest News (https://ucanr.edu/blogs/blogcore/postdetail.cfm?postnum=15861).

Dara, S. K.  2018.  An update on the invasive spotted lanternfly, Lycorma delicatula: current distribution, pest detection efforts, and management strategies.  UCANR eJournal Pest News (https://ucanr.edu/blogs/blogcore/postdetail.cfm?postnum=26349).

Dara, S. K., Barringer, L. and Arthurs, S. P.  2015.  Lycorma delicatula (Hemiptera: Fulgoridae): a new invasive pest in the United States.  J. Integr. Pest Manag. 6: 20.

Wakie, T. T., Nevin, L. G., Yee, W. L. and Lu, Z.  2020.  The establishment risk of Lycorma delicatula (Hemiptera: Fulgoridae) in the United States and globally.  J. Econ. Entomol. 113: 306-314.

Larvae of the western grapeleaf skeletonizer feeding on a grape leaf. (Photo by Surendra Dara)

The western grapeleaf skeletonizer (WGLS), Harrisina metallica, is a pest of vineyards in some parts of California.  Larval feeding skeletonizes grape leaves and uncontrolled populations can lead to a complete loss of foliage, fruit damage, and yield reduction.  WGLS populations are usually suppressed with standard pest management practices used against it or other pests.  However, considering regular WGLS infestations in the past few years especially in organic vineyards in warmer parts of the state warrant development of a good monitoring and integrated pest management strategy to improve the pest control efficacy and to minimize the risk of resistance development from potential overuse of limited organic pesticides.  An earlier bioassay with biologicals showed azadirachtin, spinosad, Bacillus thuringiensis subsp. aizawai, and entomopathogenic fungi Beauveria bassiana and Metarhizium sp. as potential control options (Dara et al., 2019).  The potential of Harrisina brillians granulovirus, a naturally occurring virus that previously suppressed WGLS populations a few decades ago is also explored as a natural solution (Federici and Stern, 1990).

WGLS has 2-3 generations per year with late spring-early summer and mid-late summer infestations in the coastal regions (Fig. 1).  Based on the detection of shiny black moths and growing degree-day calculations, pesticide applications can be timed to target hatching larvae.  Growing degree-day calculations were made using a model provided by Pest Prophet and temperature data from GreenCast. Good monitoring tools such as traps equipped with lures can be useful to improve the monitoring accuracy especially when the adult activity spreads over multiple weeks for each generation.  A study was conducted to assist with the development of new lures for WGLS.

Fig. 1. Growing degree-days indicating WGLS sping and summer generations and thresholds for larvae and adults

Methodology

An organic Cabernet Sauvignon vineyard (San Juan North) in Shandon was used for the study conducted between May and July 2021.  Treatments included a blank lure, Pherocon WGLS, TRE 2500, and TRE 2501.  The last two are developmental formulations.  The pheromone components of the lures are different combinations of 2S-butyl Z7-tetradecenoate, 2-butyl decanoate, 2-butyl dodecanoate, and isopropyl Z7-tetradecenoate to attract male moths and the latter two are new combinations of active ingredients.  Each treatment was replicated six times in a randomized complete block design.  Within each treatment, a lure was placed in Pherocon VI Delta trap with an adhesive, replaceable liner and tied in the top part of the canopy.  A 30 m distance was maintained between the traps with and between replications.  Traps were first set up with new lures and liners on 8 May 2021.  Adhesive liners were observed every week between 15 May and 3 July 2021 on eight observation dates to count the number of moths.  Lures were replaced on 5 June 2021 and adhesive liners were replaced every week or every other week as needed.  Data were analyzed using Statistix software and Tukey's HSD test was used to separate significant means.

Pheromone infused lure surrounded by the western grapeleaf sekeltonizer male moths (Above photo by Surendra Dara and below photo by Carson DiCicco)

Results

Moth counts significantly (P < 0.0005) varied among the lures on all observation dates (Table 1 and Fig. 2).  In general, TRE 2501 lure attracted significantly higher number of moths for most of the observation period.  Due to a logistics issue, adhesive liners were not replaced after the moth counts were made on 12 June and those numbers were detected from the next count to derive 19 June moth counts.  A lack of space on the liner was probably the reason for having lower moth numbers on 19 June 2021 in TRE 2501. Pherocon WGLS, which is commercially available in the market, was generally less attractive than the developmental formulations.  Pheromone combination in the TRE 2501 can be considered for the new formulation for improved monitoring efficacy.  Compared to visual monitoring of moth activity, using lures appeared to be an effective strategy for monitoring WGLS, which helped the grower to make effective treatment decisions.  On average, 287 moths were captured per each TRE 2501 lure during the 8-week observation period.  Considering that each moth can deposit 300 eggs in its lifetime, trapping adults during monitoring can also contribute to reduction in their offspring.  In addition to serving as a monitoring tool, lures can also be a control option, if economical.

Acknowledgments: Thanks to Trece for providing lures and traps for the study.

References

Dara, S. K., S. S. Dara, and S. Jaronski.  2019.  Biorational control options for the western grapeleaf skeletonizer, a re-emerging pest in California.  eJournal of Entomology and Biologicals. https://ucanr.edu/blogs/blogcore/postdetail.cfm?postnum=29081

Federici, B. A. and V. M. Stern.  1990.  Replication and occlusion of a granulosis virus in larval and adult midgut epithelium of the western grapeleaf skeletonizer, Harrisina brillians.  J. Invertebr. Pathol. 56: 401-414.

https://ucanr.edu/articlefeedback

Biopesticides contain active ingredients of natural or biological origin that include plant extracts, microorganisms, microbial metabolites, organic molecules, minerals, or other such natural materials that have pesticidal properties.  Pests such as herbivorous arthropods, pathogens, parasitic nematodes, mollusks, rodents, and weeds cause significant crop damage when they are not managed.  Pest suppression is a critical part of crop production to maintain plant health, prevent yield losses, and optimize returns.  As agriculture advanced from subsistence farming to a global enterprise, crop protection also evolved over millennia.  When farming was less organized, nature maintained a balance and provided solutions initially.  Then natural solutions were actively implemented until industrialization led to the use of synthetic inputs in the 20^th century.  While synthetic fertilizers and pesticides contributed to a tremendous improvement in the yield potential, the indiscriminate use of some of them and the resulting damage to the environment and human health steered food production in the recent past towards organic farming with the use of nature-based solutions. 

Although biopesticides have been around for a few decades, the growth of organic farming gave an impetus to the biopesticide industry during the past few years resulting in the development of new active ingredients and improved formulations.  Now, biopesticides are considered an important part of integrated pest management (IPM) strategies in both organic and conventional systems.  With a considerable industry investment in research and development, the quality and efficacy of biopesticides have also significantly improved.  This has also contributed to optimizing the cost of some formulations.  However, there is still a need to fill the knowledge gaps in biopesticides and their use.  Depending on the active ingredient, the mode of action for biopesticides, their target pests, their storage and handling, and the use strategies are quite diverse, and a thorough understanding of these aspects is critical for their successful use.  As emphasized in the new IPM model (Dara, 2019), while biopesticide use is an integral part of crop protection, understanding the pest biology, using biopesticides appropriate for the target life stage of the pest, applying them at the right time and rate using the right technology, avoiding incompatibility issues, building and sharing effective use strategies, and continuously investing in research and outreach are essential elements of biopesticide use.  Biopesticides also play an important role in insecticide resistance management (IRM) to address resistance issues associated with synthetic pesticides.  This article provides an overview of various biopesticide categories and general strategies for their successful use for IPM and IRM.

Biopesticides can be used for managing arthropod pests, bacterial or fungal pathogens, plant-parasitic nematodes, weeds, and snails and slugs.  Some formulations or active ingredients have multiple roles and can be effective against more than one category of pests.  While some active ingredients are very specific to a particular pest or related species, others have a broad-spectrum activity.  Based on the source, biopesticides can be placed in four broad categories: i) botanicals, ii) microbials, iii) toxins, and iv) minerals and other natural materials.

Botanical extracts: Plants are a rich source of numerous phytochemicals or secondary metabolites that have a wide range of properties including pesticidal activity.  Acids, alkaloids, flavonoids, glycosides, saponins, and terpenoids in plant extracts or oils obtained from seeds and other plant parts are some of the compounds present in various biopesticides (Pino et al., 2013).  Azadirachtin, BLAD (polypeptide from sweet lupine seeds), citric acid, essential oils, pyrethrins, soybean oil, and extract of the giant knotweed are used for their acaricidal, insecticidal, fungicidal, nematicidal, or herbicidal properties.  

Microbials: Some of the microbial pesticides have live microorganisms (such as entomopathogens, Bacillus spp., Streptomyces spp., and Trichoderma spp.) while others (such as Burkholderia rinojensis and Chromobacterium subtsugae)have heat-killed microorganisms and fermentation solids as the active ingredients.  Entomopathogenic microorganisms [Bacillus thuringiensis (bacterium), Beauveria bassiana and Cordyceps fumosorosea (fungi), Heterorhabditis spp. and Steinernema spp. (nematodes), and granuloviruses and nucleopolyhedroviruses] primarily kill their hosts through infection; microbe-based fungicides antagonize plant pathogens through competitive displacement and production of toxic metabolites; nematophagous fungi parasitize plant-parasitic nematodes; and plant pathogenic bacteria, fungi, and viruses infect and suppress weeds.  Bacteriophages, which are viruses that parasitize bacteria, are used against the plant pathogenic species of Clavibacter, Erwinia, Pseudomonas, Xanthomonas, Xylella, and other genera.    

Toxins and other organic molecules: There are multiple examples of toxic organic molecules derived from various organisms.  Avermectins from the bacterium Streptomyces avermitilis and spinosad from the bacterium Saccharopolyspora spinosa, strobilurin from the mushroom Strobuluris tenacellus, and cerevisane from the yeast Saccharomyces cerevisae are some of the microbial toxins that are effective against insects, plant-parasitic nematodes, or snails and slugs.  A venom peptide from the Blue Mountains funnel-web spider, Hadronyche versuta, from Australia is a recently developed insecticide active ingredient with its unique mode of action class.  Chitosan, a polysaccharide from the exoskeleton of shellfish, is a fungicide.

Minerals and other natural materials: Diatomaceous earth, mineral oil, and minerals such as sulfur are used for controlling multiple categories of pests.  Potassium salts of fatty acids of plant or animal origin, known as insecticidal soap, have insecticidal and fungicidal properties.  Organic acids such as acetic acid and citric acid are derived from plants and have fungicidal and herbicidal properties.  Since these are different from other botanical extracts, they are placed in this category.

Except for the microbial pesticides that have live microorganisms, most biopesticides have chemical molecules of microbial, fungal, botanical, or mineral origin and work through various modes of action similar to synthetic pesticides.  Several synthetic pesticides are developed from natural molecules.  Abamectin, pyrethroids, neonicotinoids, spinetoram, and storbulurins are synthetic analogs based on avermectins, pyrethrins, nicotine, spinosad, and strobulurin, respectively, and were developed for improved stability, safety, or ease of commercial-scale production.

Integrated pest management and resistance management: Biopesticides are very diverse in their origin and mode of action and have been successfully used in several cropping systems for managing a variety of pests.  They have complex interactions with plants, soil microbiota, pests, and environmental conditions.  It is critical to have a good understanding of the source of biopesticides and how they act on their target pests.  Certain biopesticides may have special storage and handling requirements or tank-mixing restrictions.  It is essential to refer to the manufacturer's guidelines or label instructions to avoid incompatible tank-mix combinations, understand proper application sequences, and to store, transport, and apply under unfavorable conditions.  While it is very important to use biopesticides as a part of the IPM program and tools for IRM, caution is warranted to avoid repeated use of the same or a similar type of biopesticide.  Pests can develop resistance to biopesticides just as they do to synthetic pesticides (Dara, 2020).  

Mechanisms of resistance to pesticides

Strategies for using biopesticides: From the seed or transplant treatment to soil or foliar application, biopesticides can be used throughout crop production.  Certain combinations can have an additive or a synergistic effect on pest suppression.  At the same time, certain inputs or practices can negatively impact biopesticide efficacy.  For example, alkaline tank-mix components breakdown the protein coat of entomopathogenic viruses and Bacillus thuringiensis.  Botanical oils can be incompatible with cold water.  Some fungicides such as captan and thiram are incompatible with entomopathogenic fungi like Beauveria bassiana while several others are compatible (Dara et al., 2014).

Investing in biopesticides: Environmental safety and resistance development are two major concerns for excessive use of synthetic pesticides and incorporating biopesticides into IPM will help address both issues.  Substituting biopesticides for synthetic pesticides will reduce the total amount of the latter during a production season and their potential negative impact on the environment and human health.  Several biopesticides are not harmful to pollinators and in some production systems, pollinators are used to deliver biopesticides to the crops they pollinate.  Adding biopesticides to the standard crop protection program will also increase pest control efficacy.  Additionally, by not continuously using synthetic pesticides, the risk of resistance will be reduced and thus their efficacy will continue to be maintained.  Although some biopesticides can be more expensive than synthetic pesticides, investing in them will be a good strategy for both the short-term benefit of effective pest suppression and the long-term benefit of a healthy and resilient ecosystem.  Since pests do not have boundaries, area-wide implementation of good agricultural practices with a balanced use of synthetic and natural inputs is necessary for maintaining the productivity of the cropping systems. 

Productive collaborations among the pesticide industry, researchers, extension educators, and the grower community are critical for successfully using biopesticides for sustainable food production.  While research helps to develop effective formulations and their use strategies, outreach helps with the implementation of those strategies.

References
Dara, S.S.R., S. S. Dara, A. Sahoo, H. Bellam, and S. K. Dara.  2014.  Can entomopathogenic fungus Beauveria bassiana be used for pest management when fungicides are used for disease management?  UCANR eJournal of Entomology and Biologicals. https://ucanr.edu/blogs/blogcore/postdetail.cfm?postnum=15671

Dara, S. K. 2019.  The new integrated pest management paradigm for the modern age.  J. Integr. Pest Manag. 10 (1): 12. https://doi.org/10.1093/jipm/pmz010

Dara, S. K.  2020.  Arthropod resistance to biopesticides.  Organic Farmer 3 (4): 16-19. https://organicfarmermag.com/2020/08/arthropod-resistance-to-biopesticides/

Pino, O. Y. Sánchez, and M. M. Rojas.  2013.  Plant secondary metabolites as an alternative in pest management.  I: Background, research approaches and trends.  Rev. ProtecciónVeg. 28 (2): 81-94.

https://ucanr.edu/articlefeedback

Brussels sprouts field in Santa Maria

The diamondback moth, Plutella xylostella (Lepidoptera: Plutellidae) is an important pest of broccoli, Brussels sprouts, cabbage, cauliflower, collards, kale, and other cruciferous crops.  It exclusively feeds on cultivated and weedy crucifers and has a worldwide distribution.  Larvae feed on foliage and growing parts of young plants or bore into the heads or flower buds resulting in skeletonization of leaves, stunting of the plants, or failure of head formation in some hosts.  In warmer areas, the diamondback moth has up to 12 generations per year.  While multiple species of parasitoids and predatory arthropods provide some level of natural control, insecticidal applications are a primary means of diamondback moth management.  Although several synthetic and biological pesticides are effective against the diamondback moth, resistance to Bacillus thuringiensis (Ferré et al. 1991), abamectin (Pu et al. 2009), emamectin benzoate, indoxacarb, and spinosad (Zhao et al. 2006), pyrethroids and other pesticides (Leibee and Savage 1992; Endersby et al. 2011) has been well-known from around the world.  Excessive use of any kind of pesticide leads to resistance problems (Dara 2020; also see a video presentation) to an individual pesticide or multiple pesticides.  Integrated pest management (IPM) strategy encourages the use of various control options both for maintaining pest control efficacy and reducing the risk of resistance development (Dara 2019).  Regularly monitoring the pest populations to make treatment decisions, rotating pesticides with different modes of action, exploring the potential of biocontrol agents, and other non-chemical control approaches such as mating disruption with pheromones are some of the IPM strategies for controlling the diamondback moth.  While sex pheromones effectively used to manage several lepidopteran pests and are proven to be a critical IPM tool, mating disruption is not fully explored for controlling the diamondback moth.  A study was conducted in Brussels sprouts to evaluate the efficacy of a sprayable pheromone against the diamondback moth and to enhance the current IPM strategies.

Methodology

The study was conducted on a 10-acre Brussels sprouts field in Santa Maria.  Cultivar Marte was planted in early July 2020 with expected harvesting in mid to late December.  A typical diamondback control program includes monitoring diamondback moth populations with the help of sticky traps and lures and applying various combinations of biological and synthetic pesticides at regular intervals.  This study evaluated the efficacy of adding CheckMate DBM-F to the grower standard practice of monitoring the diamondback moth populations with traps and lures and applying pesticides.  Treatments included i) grower standard pesticide program (Table 1) and ii) grower standard pesticide program with two applications of 3.1 fl oz of CheckMate DBM-F on 9 August and 11 September.  Treatment materials were applied by a tractor-mounted sprayer using a 100 gpa spray volume and necessary buffering agents and surfactants.  Each treatment was 5 ac and divided into four quadrants representing four replications.  In the middle of each quadrant, one Suterra Wing Trap was set up with a Trécé Pherocon Diamondback Moth Lure.  Lures were replaced once a month in early September and early October.  Sticky liners of the traps were replaced every week to count the number of moths trapped.  Traps were placed on 1, 12, and 24 August, 1, 11, 18, and 27 September, and 6 October and the moth counts were taken from respective traps on 8 and 20 August, 1, 11, 18, and 27 September, 6 and 15 October.  CheckMate DBM-F was applied at 3.1 fl oz/ac on 9 August and 11 September.  The number of larvae and their feeding damage on a scale of 0 to 4 (where 0=no damage, 1=light damage, 2=moderate damage, 3=high damage, 4=extensive/irrecoverable) were recorded from 25 random plants within each replication on 30 August and 6 and 18 October.  Data were subjected to analysis of variance using Statistix software and significant means were separated using Tukey's HSD test.  The retail value of various pesticides was also obtained to compare the cost of treatments. 

Table 1.  Pesticides, buffering agents, and surfactants, their active ingredients, rates/ac (along with the IRAC mode of action groups), and retail pricing for those applied in the grower standard diamondback moth control program.

Wing trap with septa lures

When CheckMate DBM-F [(Z)-11-Hexadecenal (3) , (Z) - 11 - Hexadecen-1-yl Acetate (1)] was applied the first time on 9 August, Dibrom 8 Emulsive was replaced with Warrior II, the buffering agent Quest was not used, and the surfactant Dyne-Amic was replaced with Induce (dimethylpolysiloxane) to avoid potential compatibility issues.  The impact of this substitution is expected to be negligible within the scope of this study.  The retail cost of 3.1 fl oz CheckMate DBM-F is $45.60.  The cost of lures and traps would be about $4-8 per acre for a six-month crop like Brussels sprouts.

Results and Discussion

Moth populations: Traps in replication 4 in both treatments on 8 August and replication 1 in the grower standard were missing, probably knocked down by a tractor.  The day before CheckMate DBM-F was first applied, the mean number of adult diamondback moths caught were 227 in the grower standard and 271 in the plots that would receive the pheromone application.  There was a gradual decline in moth counts during the rest of the observation period in both treatments.  However, the decline was higher in the plots that received CheckMate DBM-F.  The number of moths per trap were about 19% higher in the pheromone-treated plots compared to the grower standard before the study but were nearly 98% lower by the end of the study.  The reduction in moth populations from mating disruption was significant on 18 September (P =0.039) and 15 October (P = 0.006).

Adult diamondback moths on the last observation date in treatments with and without the pheromone

Larval populations: The mean number of larvae per 25 plants in a replication was zero on all observation dates except for 0.01 on 30 August in the plots that received CheckMate. Four insecticide applications by the time the study was initiated, and the remaining six applications effectively suppressed larval populations.

Damage ratings: Larval feeding damage ratings were consistently low (P < 0.0001) in the plants that did not receive CheckMate DBM-F.  The damage was limited to the older leaves at the bottom of the plants and must have been from early feeding before the initiation of the study.  The lack of larvae and the evidence of new feeding damage also confirm that the crop remained healthy and pest-free.

Since frequent pesticide applications effectively suppressed larval populations and prevented their feeding damage, the effectiveness of mating disruption in reducing yield losses could not be determined in this study.  Since larval counts were not made weekly or between pesticide applications, those that were probably present between the pesticide applications could not be determined.  Moths found in the traps probably developed from the larvae in the field or could have been those that flew in from other areas.  However, lower moth populations in CheckMate DBM-F treatment demonstrated the overall influence of mating disruption and pest suppression.

It is common to make about 10-12 pesticide sprays during the 6-month crop cycle of Brussels sprouts.  The cost of each application varied from about $73 to $192 depending on the materials used with an average cost of about $128 per application in this study.  The cost of two CheckMate DBM-F applications is $91.  If diamondback moth populations could be reduced with mating disruption, it is estimated that 2-3 pesticide applications could be eliminated.  That results in $164 to $292 of saving for the pesticide costs and additional savings in the application costs per acre by investing $91 in the mating disruption.  Since the diamondback moth can develop resistance to several chemical and natural pesticides, eliminating some applications as a result of mating disruption also contributes to resistance management along with potential negative impact of pesticides on the environment.  Compared to other mating disruption strategies, a sprayable formulation compatible with other agricultural inputs is easier and cost-effective to use.

This study demonstrated that mating disruption with CheckMate DBM-F will significantly enhance the current IPM practices by reducing pest populations, contributing to insecticide resistance management, and reducing pest management costs.  Additional studies, with fewer pesticide applications that allow larvae to survive and cause some damage, might further help understand the role of mating disruption where pest populations are not managed as effectively as in this field.

Watch a presentation of this study

Acknowledgments: Thanks to the PCA and the grower for their research collaboration, Tamas Zold for his technical assistance in data collection, Ingrid Schumann for market research of pesticide pricing, and Suterra for the financial support.

References

Dara, S. K.  2019.  The new integrated pest management paradigm for the modern age.  J. Int. Pest Manag. 10: 12.

Dara, S. K. 2020.  Arthropod resistance to biopesticides.  Organic Farmer 3 (4): 16-19.

Endersby, N. M., K. Viduka, S. W. Baxter, J. Saw, D. G. Heckel, and S. W. McKechnie.  2011.  Widespread pyrethroid resistance in Australian diamondback moth, Plutella xylostella (L.), is related to multiple mutations in the para sodium channel gene.  Bull. Entomol. Res. 101: 393.

Ferré, J., M. D., Real, J. Van Rie, S. Jansens, and M. Peferoen.  1991.  Resistance to the Bacillus thuringiensis bioinsecticide in a field population of Plutella xylostella is due to a change in a midgut membrane receptor.  Proc. Nat. Acad. Sci. 88: 5119-5123.

Leibee, G. L. and K. E. Savage.  1992.  Evaluation of selected insecticides for control of diamondback moth and cabbage looper in cabbage in Central Florida with observations on insecticide resistance in the diamondback moth.  Fla. Entomol. 75: 585-591.

Pu, X., Y. Yang, S. Wu, and Y. Wu.  2009.  Characterisation of abamectin resistance in a field-evolved multiresistant population of Plutella xylostella.  Pest Manag. Sci. 66: 371-378.

Zhao, J-Z., H. L. Collins, Y-X. Li, R.F.L. Mau, G. D. Thompson, M. Hertlein, J. T. Andaloro, R. Boykin, and A. M. Shelton.  2006.  Monitoring of diamondback moth (Lepidoptera: Plutellidae) resistance to spinosad, indoxacarb, and emamectin benzoate. J. Econ. Entomol. 99: 176-181.

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