The western flower thrips, Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) is one of the major pests of lettuce in California. It has a wide host range including several vegetable, ornamental, and other cultivated or wild plants. Native to North America, the western flower thrips is also known as alfalfa thrips, California thrips, and maize thrips among others. This article provides a general overview of the pest, its biology, damage, and management.
Eggs are small, oval, and inserted into plant tissue. Nymphs are slender and have four instars. The first two - larva I and II – feed on plant tissues while the latter two - prepupa and pupa – are non-feeding stages that are often found in the soil. Larvae are wingless and white initially and turn yellow or orange once they start feeding. Adults are small (< 2 mm), slender, and have two pairs of long, narrow wings with a fringe of hairs. The western flower thrips can occur in different color morphs such as yellow or orange, brown, and black.
The western flower thrips prefers flowers, but also feeds on developing buds, fruits, and foliage. Larvae and adults rupture the leaf surface with their rasping mouthparts and feed on plant juices. Feeding damage results in silvery appearance of the leaf surface, which later turns brown. The presence of dark fecal specs indicates thrips occurrence. In lettuce, the western flower thrips transmits Tomato spotted wilt virus and is the sole vector of Impatiens necrotic spot virus. Only the larval stages acquire these tospoviruses and the adults transmit the viruses to other plants as they spread in the field.
Integrated pest management approach is critical for successful pest management. It involves regular monitoring, exploring the potential of multiple options including cultural and biological solutions, and proper timing and application of various strategies among others. The western flower thrips is one of the pests where insecticide resistance is a common problem. To reduce the risk of resistance development, it is necessary to explore the potential of multiple control options and rotate insecticides with different modes of action. This is essential to suppress pest populations to desired levels and also to maintain control efficacy of existing pesticides.
Cultural control – Remove weed and other hosts that harbor thrips or viruses. Sprinkler irrigation can help reduce thrips populations. Plow down lettuce crop residue to destroy surviving stages. In general, maintaining good plant health with optimal nutrition and irrigation practices helps plants withstand pest damage. Silicate products can improve the structural strength of plant tissues and reduce pest damage and/or populations. Several biostimulants or biological soil amendments can also help activate plant's natural defenses against pest infestations. Consider using them to improve overall plant health and yields, and to protect plants from biotic and abiotic stresses.
Biological control – Predators such lacewings (Chrysopa spp. and Chrysoperla spp.), minute pirate bugs (Orius spp. and Anthocoris spp.), predatory mites (Amblyseius swirski, Ablyseius andersoni, Neoseiulus cucumeris and Stratiolaelaps scimitus), and rove beetles (Dalotia coriaria) attack thrips. Conserve natural enemies with insectary plants and applying safer pesticides, and augment natural populations by releasing commercially reared species.
Microbial control – Entomopathogenic fungi such as Beauveria bassiana and Cordyceps (Isaria) fumosorosea, products based on bacteria such as Burkholderia rinojensis and Chromobacterium subtsugae, and entomopathogenic nematodes such as Heterorhabditis spp. and Steinernema feltiae can be used against one or more life stages. Entomopathogenic nematodes are more effective against pupae in soil because they actively search for and infect their hosts. Entomopathogenic fungi can be used against all life stages.
Botanical control – Azadirachtin alone or in combination with entomopathogenic fungi or insecticides can also be used against multiple life stages. Azadirachtin is an insecticide, antifeedant, and a growth regulator. Similarly, pyrethrins derived from chrysanthemum flowers can be used alone or with other biological or synthetic insecticides. Pyrethrins are nerve poisons. Other botanical insecticides that contain soybean oil, rosemary oil, thymol, and neem oil (which also has a low concentration of azadirachtin) also provide control against thrips through insecticidal, repellency, and antifeedant activities.
Other control options – Insecticidal soaps and mineral oils can be used against different life stages of thrips. Spinosad, a popular insecticide of microbial origin and a mixture of two chemicals spinosyn A and spinosyn D, is very effective against thrips. However, overuse of spinosad can lead to resistance issues in thrips and other insects.
Chemical control – There are several synthetic insecticides that are effective against thrips. It is important to rotate chemicals among different mode of action groups to reduce the risk of insecticide resistance. The following are some synthetic active ingredients and their mode of actions groups in parenthesis that can be used for thrips control: methomyl (1A), bifenthrin (3A), lambda-cyhalothrin (3A), zeta-cypermethrin (3A), clothianidin (4A), spinetoram (5), and cyantraniliprole (28).
Depending on the level of control needed, combinations of products from different categories can improve control efficacy. For example, a combination of entomopathogenic fungi and nematodes can be applied to the soil for controlling prepupae and pupae. While the soil-dwelling predatory mite S. scimitus and the rove beetle, D. coriaria, can be used against pupal stages, other natural enemies can be used against nymphs and adults. A combination of entomopathogenic fungi and azadirachtin can be applied both to the soil or foliage for controlling different life stages. Similarly, various biological and synthetic insecticides can be applied in combination or rotation to obtain desired control.
The categories presented above are based on the source or nature of the active ingredients and do not indicate their organic or conventional label status. Please check the product labels for their appropriateness for managing thrips in lettuce, for use in organic farms, and guidelines for storage, handling, and field use. Entomopathogenic nematodes, fungi, and other biologicals are compatible with several synthetic agricultural inputs, but verify the label guidelines for specific instructions.
Dara, S. K. 2019. The new integrated pest management paradigm for the modern age. JIPM 10: 1-9. https://doi.org/10.1093/jipm/pmz010
Dara, S. K. 2021. Biopesticides: categories and use strategies for IPM and IRM. UC ANR eJournal of Entomology and Biologicals. https://ucanr.edu/blogs/blogcore/postdetail.cfm?postnum=46134
Natwick, E. T., S. V. Joseph, and S. K. Dara. 2017. UC IPM pest management guidelines: lettuce. UC ANR Publication 3450. https://www2.ipm.ucanr.edu/agriculture/lettuce/Western-flower-thrips/
Riley, D. G., S. V. Joseph, R. Srinivasan, and S. Diffie. 2011. Thrips vectors of tospoviruses. JIPM 2: I1-I10. https://doi.org/10.1603/IPM10020
- Author: Surendra K. Dara
The diamondback moth (DBM), Plutella xylostella, is a small plutellid moth of European origin that has been in North America for nearly two centuries. It is currently present in many parts of the world feeding exclusively on cruciferous hosts such as broccoli, cabbage, and cauliflower. DBM has multiple generations per year and can cause significant yield losses when populations are not controlled. Increasing temperatures that shorten pest life cycle, changing climatic patterns and milder winters in many areas, the ability of adult DBM to disperse, and the presence of cultivated and wild cruciferous crops year-round are worsening the pest problem and require continuous application of pesticides and other control options. Insecticide resistance is also a common problem in DBM where very high levels of resistance to some commonly used pesticides in field populations were reported. Although DBM infestations are common in cruciferous vegetable production, many parts of California and Arizona have seen a significant increase in DBM populations in the past few months. Year-round production of cruciferous vegetables supports DBM populations with as many as 12 generations per year and requires regular application of pesticides. Frequent pesticide applications can lead to insecticide resistance, ineffective pest suppression, and higher yield losses. A good integrated pest management (IPM) strategy is critical to address a pest like DBM.
Biology: A female moth deposits an average of 150 eggs over about 10 days (Capinera, 2018). Eggs are deposited in small batches in depressions on leaf surfaces. Small, green larvae actively feed on the foliage, first instars in mines and the remaining three on the surface. Pupation usually occurs on the lower side of the leaf surface in a loosely spun cocoon. Adult moths are slender, greyish brown with conspicuous antennae. The light-colored diamond pattern on the wings when the moth is resting gives the name diamondback moth.
Damage: Larval feeding on foliage and growing parts of young plants causes skeletonization of leaves. Larvae can also bore into the heads and flower buds resulting in the failure of head formation and stunting of plant growth. Uncontrolled populations cause significant yield losses.
A sound IPM strategy involves regular monitoring of pest infestations, a good understanding of the pest life cycle, and using multiple tactics that target one or more life stages (Dara, 2019). The following recommendations are developed based on the new IPM model and its different components.
A. Pest Management: Some pests can be effectively controlled by one or two tactics, but a difficult pest like DBM with its increasing threat needs a variety of tactics to achieve maximum control.
i) Host plant resistance: Planting cultivars that tolerate or resist DBM damage is the first line of defense. For example, cabbage cultivars with glassy leaves (Dickson et al., 1990) and a specific glucosinolate profile (Robin et al., 2017) are resistant to larval damage. On glassy leaf surfaces, larvae spend less time feeding and more time searching for a suitable spot to feed. The presence or higher levels of glucobrassicin, glucoiberin, and glucoiberverin and the absence or lower levels of 4-hydroxyglucobrassicin, glucoerucin, glucoraphanin, and progoitrin showed resistance to larval feeding in cabbage (Robin et al., 2017).
ii) Cultural control: Maintaining a brassica-free period or rotating with non-brassica crops will help break the pest cycle. Removal of weedy hosts can also reduce the source of infestation, but DBM adults can disperse in search of their hosts. Good agronomic practices can ensure optimal plant health and compensate for potential yield losses when infestations are low. Certain biostimulants can induce systemic resistance or strengthen plant tissues and further contribute to the plant health under pest attack.
iii) Biological control: Various species of natural enemies contribute to the control of DBM (Sarfraz et al., 2007). The egg parasitoid Trichogramma pretiosum and the larval parasitoids Cotesia plutellae, Diadegma insulare, Diadromus subtilicornis, and Microplitis plutellae, predatory ground beetles, hemipterans, syrphid fly larvae, and spiders are some of the natural enemies of DBM. Depending on the availability, parasitoids of other Cotesia spp. and Oomyzus spp. can also be used. Conserving these natural enemies by providing strips of insectary plants in the field along with releasing commercially available natural enemies will provide the necessary biological control of DBM.
iv) Behavioral control: Mating disruption with sex pheromone is the most effective behavioral control tactic for DBM. Using pheromones confuses the male moth in finding its female mate, reduces mating, and thus the next generation individuals. A recent study in a commercial Brussels sprouts field demonstrated the potential of mating disruption with a sprayable pheromone (Dara, 2020). Studies conducted in different countries explored the potential of various antifeedants against DBM larvae and when commercially available, such materials can contribute to DBM IPM. A triterpenoid saponin from the crucifer Barbarea vulgaris in Japan (Shinoda et al., 2002), momordicine I and II from the cucurbit Momordica charantia in China (Ling et al., 2008), and the extracts of Acalypha fruticosa (family Euphorbiaceae) in India (Lingathurai et al., 2011) are some examples of the antifeedant materials investigated against DBM.
v) Physical control: Depending on the field size, crop stage, and affordability, row covers can be used to exclude DBM.
vi) Microbial control: DBM is susceptible to naturally occurring bacterial, fungal, and viral pathogens, but biopesticides based on the bacterium Bacillus thuringiensis and the bacterial toxin spinosad are the most common microbial control options for DBM in the United States. Baculovirus-based products are available for DBM control in other countries.
vii) Chemical control: Application of chemical pesticides of natural and synthetic origin is the most commonly used tactic for DBM control. Azadirachtin, pyrethrins, and synthetic pesticides from different mode of action groups can be used against DBM. Studies conducted in Ethiopia (Begna and Damtew, 2015), India (Devi and Tayde, 2017), and Thailand (Kumrungsee et al., 2014) explored the potential of various botanical extracts against DBM with varying levels of efficacy. Vegetable oils, mineral oils, neem oil, and others can also be used as both ovicides and larvicides.
B. Knowledge and Resources: The most important aspect of IPM is to develop a good understanding of the pest life cycle, seasonal trends, host preference, feeding behavior, response to environmental conditions, and biotic and abiotic stressors. This knowledge helps to identify vulnerable stages of the pest and develop appropriate control strategies. For example, mating disruption to target adults, biocontrol agents against multiple life stages, especially eggs and larvae, oils as ovicides, and other pesticides against larvae and other life stages can tackle each stage effectively. Modern tools such as smart traps to monitor pest populations and drones for releasing natural enemies can also help improve the IPM program.
C. Planning and Organization: Since insecticide resistance is a common problem with DBM, rotating pesticides (both biological and synthetic) among different mode of action groups and avoiding repetitive application of the same or a similar pesticide are critical for resistance management. Making appropriate treatment decisions based on infestation levels and the life stage of the pest, regularly monitoring for potential resistance issues, and keeping track of combination and rotation programs that worked well are all a part of effective planning and information management that improve pest control efficacy. If necessary, aggressive area-wide management plans should be developed using one or more control options.
D. Communication: When dealing with an important pest such as DBM, effective communication will help address the knowledge gaps and contribute to effective pest management. Pest control professionals and growers can explore new DBM control options by contacting researchers, attending extension meetings, or reading various articles that can be accessed through internet, university resources, or local governmental agencies. Growers can also exchange information and develop IPM strategies that best suit their situation through a collaborative effort.
Since the field conditions, infestation levels, resistance in DBM populations, and availability and affordability of control options vary, growers should customize their IPM program to suit their local needs.
Begna, F. and T. Damtew. 2015. Evaluation of four botanical insecticides against diamondback moth, Plutella xylostella L. (Lepidoptera: Plutellidae) on head cabbage in the central rift valley of Ethiopia. Sky J. Agrl. Res. 4: 97-105. http://www.skyjournals.org/sjar/pdf/2015pdf/Aug/Begna%20and%20Damtaw%20pdf.pdf
Capinera, J. L. 2018. Diamondback moth. Featured Creatures, University of Florida Publication EENY-119. https://entnemdept.ufl.edu/creatures/veg/leaf/diamondback_moth.htm
Dara, S. K. 2019. The new integrated pest management paradigm for the modern age. JIPM 10: 12, 1-9. https://doi.org/10.1093/jipm/pmz010
Dara, S. K. 2020. Mating disruption as an IPM tool in diamondback moth management. UCANR eJournal of Entomology and Biologicals. https://ucanr.edu/blogs/blogcore/postdetail.cfm?postnum=44160
Devi, H. D. and A. R. Tayde. 2017. Comparative efficacy of bio-agents and botanicals on the management of diamondback moth (Plutella xylostella Linn.) on cabbage under Allahabad agroclimatic conditions. Int. J. Curr. Microbiol. App. Sci. 6: 711-716. https://doi.org/10.20546/ijcmas.2017.607.088
Dickson, M. H., A. M. Shelton, S. D. Eigenbrode and M. L. Vamosy. 1990. Selection for resistance to diamondback moth (Plutella xylostella) in cabbage. HortSci. 25: 1643-1646. https://doi.org/10.21273/HORTSCI.25.12.1643
Kumrungsee, N., W. Pluempanupat, O. Koul, and V. Bullangpoti. 2014. Toxicity of essential oil compounds against diamondback moth, Plutella xylostella, and their impact on detoxification enzyme activities. J. Pest Sci. 87: 721-729. https://doi.org/10.1007/s10340-014-0602-6
Ling,B., G.-c. Wang, J. Ya, M.-x. Zhang, and G.-w. Liang. 2008. Antifeedant activity and active ingredients against Plutella xylostella from Momordica charantia leaves. Agrl. Sci. China 7: 1466-1473. https://doi.org/10.1016/S1671-2927(08)60404-6
Lingathurai, S., S. E. Vendan, M. G. Paulraj, and S. Ignacimuthu. 2011. Antifeedant and alrvicidal activities of Acalypha fruticosa Forssk. (Euphorbiaceae) against Plutella xylostella L. (Lepidoptera: Yponomeutidae) larvae. J. King Saud Univ. Sci. 23: 11-16. https://doi.org/10.1016/j.jksus.2010.05.012
Robin, A.H.K., M. R. Hossain, J.-I. Park, H. R. Kim and I.-S. Nou. 2017. Glucosinolate profiles in cabbage genotypes influence the preferential feeding of diamondback moth (Plutella xylostella). Fron. Plant Sci. 8: 1244. https://doi.org/10.3389/fpls.2017.01244
Sarfraz, M., A. B. Keddie, and L. M. Dosdall. 2007. Biological control of the diamondback moth, Plutella xylostella: a review. Biocon. Sci. Tech. 15: 763-789. https://doi.org/10.1080/09583150500136956
Shinoda, T., T. Nagao, M. Nakayama, H. Serizawa, M. Koshioka, H. Okabe, and A. Kawai. 2002. Identification of a triterpenoid saponin from a crucifer, Barbarea vulgaris, as a feeding deterrent to the diamondback moth, Plutella xylostella. J. Chem. Ecol. 28: 587-599. https://doi.org/10.1023/A:1014500330510
- Author: Surendra K. Dara
- Author: Roland C. Bocco
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.
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.
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.
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.
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.
Thanks to the California Department of Food and Agriculture for funding this study.
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.
- Author: Surendra K. Dara
- Author: Carson DiCicco, Vina Quest
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
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)
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.
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.
- Author: Surendra K. Dara
- Author: Dave Peck, Manzanita Berry Farms
Botrytis cinerea infection appears as wilted flowers and a layer of spores on ripe fruit. Photo by Surendra Dara
Botrytis fruit rot or gray mold caused by Botrytis cinerea is an important disease of strawberry and other crops damaging flowers and fruits. Pathogen survives in the plant debris and soil and can be present in the plant tissues before flowers form. Infection is common on developing or ripe fruits as brown lesions. Lesions typically appear under the calyxes but can be seen on other areas of the fruit. As the disease progresses, a layer of gray spores forms on the infected surface. Severe infection in flowers results in the failure of fruit development. Cool and moist conditions favor botrytis fruit rot development. Sprinkler irrigation, rains, or certain agricultural practices can contribute to the dispersal of fungal spores.
Although removal of infected plant material and debris can reduce the source of inoculum in the field, regular fungicide applications are typically necessary for managing botrytis fruit rot. Since fruiting occurs continuously for several months and fungicides are regularly applied, botrytis resistance to fungicides is not uncommon. Applying fungicides only when necessary, avoiding continuous use of fungicides from the same mode of action group (check FRAC mode of action groups), exploring the potential of biological fungicides to reduce the risk of resistance development are some of the strategies for effective botrytis fruit rot management. In addition to several synthetic fungicides, several biological fungicides continue to be introduced into the market offering various options for the growers. Earlier field studies evaluated the potential of various biological fungicides and strategies for using them with synthetic fungicides against botrytis and other fruit rots in strawberry (Dara, 2019; Dara, 2020). This study was conducted to evaluate some new and soon to be released fungicides in fall-planted strawberry to support the growers, ag input industry, and to promote sustainable disease management through biological and synthetic pesticides.
This study was conducted at the Manzanita Berry Farms, Santa Maria in strawberry variety 3024 planted in October, 2020. While Captan and Switch were used as synthetic standards, a variety of biological fungicides of microbial, botanical, and animal sources were included at various rates and different combinations and rotations. Products and active ingredients evaluated in this study included Captan Gold 4L (captan) from Adama, Switch 62.5 WG (cyprodinil 37.5% + fludioxinil 25%) from Syngenta, NSTKI-14 (potassium carbonate 58.04% + thyme oil 1.75%) from NovoSource, A22613 [A] (botanical extract) from Syngenta, Regalia (giant knotweed extract 5%) from Marrone Bio Innovations, EXP14 (protein 15-20%) from Biotalys, Gargoil (cinnamon oil 15% + garlic oil 20%) and Dart (caprylic acid 41.7% + capric acid 28.3%) from Westbridge, Howler (Pseudomonas chlororaphis strain AFS009), Theia (Bacillus subtilis strain AFS032321), and Esendo (P. chlororaphis strain AFS009 44.5% + azoxystrobin 5.75%) from AgBiome, ProBlad Verde (Banda de Lupinus albus doce – BLAD, a polypeptide from sweet lupine) from Sym-Agro with Kiplant VS-04 (chitosan 2.3%) or Nu-Film-P spreader/sticker, AS-EXP Thyme (thyme oil) from AgroSpheres, and AgriCell FunThyme (thyme oil) provided by AgroSpheres.
Table 1. List of treatments color coded according to the kind of fungicide (light blue=synthetic fungicide; dark blue=synthetic+biological fungicide active ingredient; peach=synthetic and biological fungicides alternated; green=biological fungicides)
Excluding the untreated control, rest of the 24 treatments can be divided into synthetic fungicides, a fungicide with synthetic + biological active ingredients (a formulation with two application rates), synthetic fungicides alternated with biological fungicides, and various kinds of biological fungicides (Table 1). Treatments were applied at a 7-10 day interval between 22 April and 17 May, 2021. Berries for pre-treatment disease evaluation were harvested on 19 April, 2021. Each treatment had a 5.67'X15' plot replicated four times in a randomized complete block design. Strawberries were harvested 3 days before the first treatment and 3-4 days after each treatment for disease evaluation. On each sampling date, marketable-quality berries were harvested from random plants within each plot during a 30-sec period and incubated in paper bags at outdoor temperatures under shade. Number of berries with botrytis infection were counted on 3 and 5 days after harvest (DAH) and percent infection was calculated. This is a different protocol than previous years' studies where disease rating was made on a 0 to 4 scale. Treatments were applied with a backpack sprayer equipped with Teejet Conejet TXVK-6 nozzle using 90 gpa spray volume at 45 PSI. Water was sprayed in the untreated control plots. Dyne-Amic surfactant at 0.125% was used for treatments that contained Howler, Theia, Esendo, AgriCell FunThyme, AS-EXP Thyme, and EXP 14. Research authorization was obtained for some products and crop destruction was implemented for products that did not have California registration.
Percent infection data were arcsine-transformed before subjecting to the analysis of variance using Statistix software. Significant means were separated using the least significant difference test.
Pre-treatment infection was very low and occurred only in some treatments with no statistical difference (P > 0.05). Infection levels increased for the rest of the study period. There was no statistically significant difference (P > 0.05) among treatments for disease levels 3 or 5 days after the first spray application. Differences were significant (P = 0.0131) in disease 5 DAH after the second spray application where 13 treatments from all categories had significantly lower infection than the untreated control. After the third spray application, infection levels were significantly lower in eight treatments in 3 DAH observations (P = 0.0395) and 10 treatments in 5 DAH observations (P = 0.0005) compared to untreated control. There were no statistical differences (P > 0.05) among treatments for observations after fourth spray application or for the average of four applications. However, there were numerical differences where infection levels were lower in several treatments than the untreated control plots.
In general, the efficacy of both synthetic and biological fungicides varied throughout the study period among the treatments. When the average for post-treatment observations was considered, infection was numerically lower in all treatments regardless of the fungicide category. Multiple biological fungicide treatments either alone or in rotation with synthetic fungicides appeared to be as effective as synthetic fungicides.
Botanical and microbial fungicides can be effective against either for using alone or in rotation with synthetic fungicides for suppressing botrytis fruit rot in strawberry. Additional studies can help optimize the application rates and use strategies for those fungicides that were not as effective as others. Sanitation practices and use of synthetic and biological fungicides help manage botrytis fruit rot.
Acknowledgements: Thanks to AgBiome, AgroSpheres, Biotalys, NovaSource, Sym-Agro, Syngenta, and Westbridge for funding and Chris Martinez for his technical assistance.
Dara, S. K. 2019. Five shades of gray mold control in strawberry: evaluating chemical, organic oil, botanical, bacterial, and fungal active ingredients. UCANR eJournal of Entomology and Biologicals. https://ucanr.edu/blogs/blogcore/postdetail.cfm?postnum=30729
Dara, S. K. 2020. Evaluating biological fungicides against botrytis and other fruit rots in strawberry. UCANR eJournal of Entomology and Biologicals. https://ucanr.edu/blogs/blogcore/postdetail.cfm?postnum=43633