Insects – Page 6 – Maryland Agronomy News

Special Alert: Growers Need to Scout for Soybean Podworms and Sorghum Headworms

Kelly Hamby¹, Galen Dively¹, David Owens², Ben Beale³, Peter Coffey³, Andrew Kness³, Alan Leslie³, Erika Taylor³, Kelly Nichols³, Matthew Morris³, and Emily Zobel³

¹University of Maryland Department of Entomology
²University of Delaware Extension Entomology
³University of Maryland Extension

Moth flight activity for the corn earworm, also known as the soybean podworm and one species of sorghum headworm, has increased during the past week. Pest pressure varies across the state and Delmarva region. The early surge in activity is attributed to the record temperatures during June and July, which have accelerated larval development and shortened the generation time of this insect. Corn earworm has already caused significant damage to ears of sweet corn and early planted field corn. Note that Bt hybrids expressing single or multiple Cry proteins no longer control earworms due to the development of resistance; only hybrids expressing the Vip3a Bt protein provide good ear protection. These hybrids represent a relatively small portion of the planted acreage. Thus, significantly more adult moths are now recruited in corn compared to levels a decade ago.

Corn earworms are strongly attracted to and prefer fresh corn silks for egg laying. Outbreaks in other crops often follow a midsummer drought, which causes the corn to ripen earlier and become less attractive to the moths. As early planted corn fields dry down, moths will move into other vegetable and grain crops.

Podworms in Soybeans:

Corn earworm feeding on soybean pod. Image: Clemson University — Corn earworm feeding on soybean pod.
Image: Clemson University

In soybeans, female corn earworm moths prefer to lay eggs in open-canopied, late-blooming fields, and are most attracted to soybeans for egg laying from flowering to early pod-set. Drought conditions also delay soybean maturity and prevent normal canopy growth, so peak moth activity is more likely to coincide with blooming of open-canopied fields. In irrigated fields, activity may be greater along pivot tracks and dry corners. Corn earworm larvae can damage flowers; however, because soybeans produce more flowers than needed, flowering sprays are rarely necessary. Podworms cause the most damage when large larvae are feeding on full seed pods with large seeds (see information from NCSU).

What to do? Scout bean fields, paying special attention to those fields with a more open canopy in areas where the nearby maturing corn is no longer attractive to earworm moths.

Sampling should start during mid-August and be repeated at least weekly in each field until a spray decision is made or the pods reach full maturity. Most fields are planted as narrow-row beans, so a 15-inch sweep net is the most practical way to sample for earworms. Walk along the rows, swinging the sweep net so that the opening passes through the foliage. The net is turned 180 degrees after each sweep as you advance with each step to swing the net through the foliage in the opposite direction. Each stroke is counted as one sweep. A series of 25 sweeps should be taken at each of 5 sites in every 40 acres.

Treatment is recommended when counts exceed 3 medium to large podworms per 25 sweeps in narrow row fields, or 5 podworms per 25 sweeps in wide row fields (20 inches or greater). The timing strategy is to wait until most of the larvae are 3/8 inch or more in length, and then treat when pod damage is first evident. This allows for most egg laying and hatching to occur before treatment and reduces the chances of a second spray being needed later. These static thresholds are based on long-term averages for control costs and soybean prices. North Carolina State extension has developed a dynamic online threshold calculator for corn earworm in soybeans that takes into account the sampling method (uses a 15 sweep rather than a 25 sweep sample), row spacing, cost for control, and the value of soybeans, which can be found at:

https://www.ces.ncsu.edu/wp-content/uploads/2017/08/CEW-calculator-v0.006.html

Since the 2008 season, numerous reports of control failures with pyrethroids (Group 3A) used for earworm control have been reported from the Mid-Atlantic region and states to our south. This insect has developed moderate to high levels of resistance to this class of insecticides, so growers need to consider other modes of action. If a pyrethroid (e.g., Brigade, Warrior, Mustang Maxx, Hero, Baythroid, Tombstone) is used, the highest labeled rate timed for small to medium, rather than large worms, is recommended. Alternative classes such as diamides (Group 28; e.g., Coragen, Prevathon), oxadiazines (Group 22A; e.g., Steward) and spinosyns (Group 5; e.g., Blackhawk, Radiant) will be most effective. These materials are also generally softer on beneficial insects which prey upon other late season soybean pests, such as soybean looper and stink bug. ALWAYS read and follow instruction on the pesticide label; the information presented here does not substitute for label instructions.

Headworms in Sorghum:

Corn earworm female in sorghum. Image: John C. French Sr

Headworms (corn earworm, fall armyworm, and sorghum webworm) are caterpillar pests that infest grain heads. Flowering or heading sorghum is attractive to corn earworm females for egg laying, and headworm issues have been reported in Southern Maryland this year. Headworms feed on the flowers and developing kernels and large larvae can cause significant yield loss.

What to do? Scout sorghum fields from the end of flowering until hard dough.

Sample heads by bending them into a clean white 5 gallon bucket and beating them to dislodge the headworms. Sample 10 heads per location and sample multiple locations per field. If most larvae are small (up to ¼ inch) sample the field again in 3 to 4 days.

Thresholds vary by the size and species of larvae and sorghum value. In general, 2 corn earworm larvae per head would warrant treatment, and Texas A&M has developed a dynamic online threshold calculator that incorporates cost of control, grain value, anticipated yield (heads/acre), and larval size, which can be found at:

https://agrilife.org/extensionento/sorghum-headworm-calculator/

As mentioned above, pyrethroids (Group 3A) offer poor to moderate control of corn earworm in the Mid-Atlantic, and will not control heavy infestations or large worms. If a pyrethroid (e.g., Brigade, Warrior, Mustang Maxx) is used, the highest labeled rate is recommended. Alternative classes such as diamides (Group 28; e.g., Prevathon), spinosyns (Group 5; e.g., Blackhawk, Tracer), or carbamates (Group 1A; e.g., Sevin, Lannate) will be most effective. Selective insecticides that are less damaging to beneficials are recommended, such as Prevathon (most recommended) or Blackhawk. ALWAYS read and follow instruction on the pesticide label; the information presented here does not substitute for label instructions.

While scouting for headworms, growers are encouraged to look for sugarcane aphid in sorghum as well. Virginia Tech reported the first confirmed identification of white sugarcane aphid in Amelia County on August 1^st. For more information on sugarcane aphid see Agronomy News Volume 8 Issue 1. If sugarcane aphids are also present, we strongly advise using selective insecticides to preserve the natural enemies that slow sugar cane aphid population growth.

Further Resources:

North Carolina State Podworm Factsheet:

https://soybeans.ces.ncsu.edu/corn-earworm/

University of Delaware Weekly Crop Update:

https://sites.udel.edu/weeklycropupdate/

University of Delaware Insect Management:

http://extension.udel.edu/ag/insect-management/field-vegetables-fruit/

Virginia Tech Pest Management Field Crops Guide:

https://www.pubs.ext.vt.edu/456/456-016/456-016.html (Sorghum Headworm Section)

Sorghum Checkoff Headworm Guide:

https://www.sorghumcheckoff.com/newsroom/2016/03/28/headworms/

Sugarcane aphid found in VA sorghum – 2019:

https://blogs.ext.vt.edu/ag-pest-advisory/sugarcane-aphid-found-in-va-sorghum-2019/

Agronomy News Sugarcane Aphid Article:

https://extension.umd.edu/sites/extension.umd.edu/files/_docs/AgronomyNewsApril2017.pdf

Sulfoxaflor Registered for New Uses

The U.S. Environmental Protection Agency (EPA) has just issued a long-term approval for the insecticide sulfoxaflor, which the Agency has characterized as “an effective tool to control challenging pests with fewer environmental impacts.” The following information is from today’s EPA OPP Update.

“After conducting an extensive risk analysis, including the review of one of the agency’s largest datasets on the effects of a pesticide on bees, EPA is approving the use of sulfoxaflor on alfalfa, corn, cacao, grains (millet, oats), pineapple, sorghum, teff, teosinte, tree plantations, citrus, cotton, cucurbits (squash, cucumbers, watermelons, some gourds), soybeans, and strawberries.

EPA is providing long-term certainty for U.S. growers to use an important tool to protect crops and avoid potentially significant economic losses, while maintaining strong protection for pollinators,” said Alexandra Dapolito Dunn, assistant administrator for EPA’s Office of Chemical Safety and Pollution Prevention. “Today’s decision shows the agency’s commitment to making decisions that are based on sound science.”

Sulfoxaflor is an important and highly effective tool for growers that targets difficult pests such as sugarcane aphids and tarnished plant bugs, also known as lygus. These pests can damage crops and cause significant economic loss. Additionally, there are few viable alternatives for sulfoxaflor for these pests. In many cases, alternative insecticides may be effective only if applied repeatedly or in a tank mix, whereas sulfoxaflor often requires fewer applications, resulting in less risk to aquatic and terrestrial wildlife.

EPA’s registration also includes updated requirements for product labels, which will include crop-specific restrictions and pollinator protection language.

*Background*

In 2016, following a 2015 decision of the Ninth Circuit Court of Appeals vacating the registration of sulfoxaflor citing inadequate data on the effects on bees, EPA reevaluated the data and approved registrations that did not include crops that attract bees. The 2016 registration allowed fewer uses than the initial registration and included additional interim restrictions on application while new data on bees were being obtained. Today’s action, adding new uses, restoring previous uses, and removing certain application restrictions is backed by substantial data supporting the use of sulfoxaflor.

For additional information, please visit the EPA website.

Arrest These Early Season Soil Critters: Wireworm and White Grub Management

Edwin Afful, Nurani Illahi and Kelly Hamby
University of Maryland, Department of Entomology

white grub next to corn plant in field — *Figure 1. Grub (circled) uncovered in field corn.*

Spending most of their lives in the soil, they feed on our cherished seeds, and cause stand reductions that affect yield. Who are these critters and what can we do to save our seeds? White grubs and wireworms are a part of soil insect pest complex known to be culprits in the “covert stand reduction operation” in small grain and corn fields. Seed corn maggots, slugs, and cutworms can also cause early season stand reduction, and their damage can be distinguished from grubs and wireworms. The early developmental stages of grubs and wireworms occur on our blindside—in the soil. Larva of both species are the perpetrators of the illegal acts against our seeds, and there are no effective rescue treatments once damage is visible. Economic populations of these pests vary by season, depend upon a variety of factors, and are sporadic. In our recent studies we have not seen economic damage; however, control decisions should be based on pest history at your site and sampling information.

Identification

Close-up of white grub — Figure 2. Raster (arrow) of white grub.

White grubs are the immature stages of scarab beetles, and multiple species (1-3 year life cycle depending on species) occur and feed upon plant roots as they develop. They have a characteristic “C” shape (Fig. 1, circled), have 3 pairs of legs immediately behind their head, and the entire grub body measures 0.25-1 inch in length. They can be identified to species based on the pattern of the hairs on the underside of their end (“raster”, Fig. 2 arrow). For further information on identification and life cycles, see resources from Ohio State⁶and Purdue⁴.

Wireworms are the immature stages of click beetles, and multiple species with different life histories occur in most grain growing regions. Wireworms chew into seeds and stems, leaving holes, dead spots, or hollowed out seeds on seedlings. They are 0.25-0.75 inches long and have slender semi-cylindrical or cylindrical bodies that can be a white, yellowish, or coppery color with three small pairs of legs behind the head. On their end, they have a flattened segment with a “keyhole-shaped” notch that can look similar to the chewing mouthparts on the head (Fig. 3).

Damage

Grubs and wireworms both cause stand reduction; however, there are some distinguishable differences. Damage by wireworms and grubs are usually confined to certain areas of a field where populations are high or where soil conditions were optimum for egg laying and larval development.

White grubs feed on roots, chewing off the fine hairs on the roots. This reduces root uptake of water and phosphorus, resulting in aboveground symptoms of wilting and purpling of the stem. Severely infested fields often suffer stand loss when injured plants die.

Wireworms typically feed on the germ of corn kernels or hollow out the kernel, killing the plant. Wireworms may also cut off small roots or tunnel into the underground portions of the root or stem of young corn plants. If feeding is above the growing point, holes will appear in the leaves above ground.

Sampling

Sampling must occur before tilling and planting and should be done once it warms up enough for grubs and wireworms to be active at the soil surface (soil temperature at 6 inch depth >45 °F)^{3, 7}. One to two samples should be taken per 10 acres with no less than 5 locations per field. To sample, dig out a 1-foot square 6 inches deep and dump the soil in a tray or sift it with ¼” hardware cloth to look for grubs and wireworms. While economic thresholds have not been established, an average of 1 white grub and/or wireworm per square foot would warrant an at-planting or seed treatment insecticide^{3, 7}.

Management

Cultural management: Rotation is the most effective, and often the lowest cost, cultural tactic for reducing many grub and wireworm problems. Both grubs and wireworms prefer grass hosts, and rotation of corn and small grains with a non-grass crop reduces populations; however, weeds must be controlled to ensure there are no host plants. Planting under warm conditions allows seeds to germinate rapidly and plants may outgrow wireworms. Certain species of wireworms are abundant only in poorly drained soils; therefore, proper drainage of soils can reduce the wireworm threat. The use of starter fertilizer, timely planting, and effective weed management will also help reduce white grub and wireworm damage².

Biological control: Organisms such as birds, parasitic nematodes, and fungal pathogens prey on white grubs and wireworms. The effectiveness of two beneficial nematodes were studied on turfgrass against grubs in California with some success¹. Further research is required to effectively use biological control for these pests.

Figure 4. Mean control rating against white grubs across a varying number (N) of trials. Unpublished data from Dr. Dominic Reisig at North Carolina State University.

Chemical Control: There are no effective rescue treatments once white grub and wireworm damage is visible. However, commercial seed treatments and at-planting insecticides have varying effectiveness for grub and wireworm protection prior to damage. Dr. Dominic Reisig at North Carolina State University has been performing efficacy trials in field corn with grub and wireworm pressure, and has rated a variety of products against each based on his results. The N values indicate the number of trials each product was used in, and the values are a rating from 0 (no control) to 9 (excellent control), with the best performance products in dark red. Most products and rates provide fair control of grubs⁵, with heavy grub pressure requiring a higher seed treatment rate or in-furrow applications of Capture® (pyrethroid insecticide; Fig. 4). Wireworms require at least a 500 rate of seed treatment for fair control, with slightly better control at the 1250 rate (Fig. 5).

Figure 5. Mean control rating against wireworms across a varying number (N) of trials. Unpublished data from Dr. Dominic Reisig at North Carolina State University.

Further Reading/ References

Koppenhöfer,A.M., Wilson,M., Brown, I., Kaya, H.K., and R. Gaugler, Biological Control Agents for White Grubs (Coleoptera: Scarabaeidae) in Anticipation of the Establishment of the Japanese Beetle in California, Journal of Economic Entomology, Volume 93, Issue 1, 1 February 2000, Pages 71–80, https://doi.org/10.1603/0022-0493-93.1.71

NC State extension: Managing Insect Pests in Organically Certified Corn: https://entomology.ces.ncsu.edu/managing-pests-in-organically-certified-corn/

Owens, D., and B. Cissel. Insect Management in Field Corn – 2018: https://cdn.extension.udel.edu/wp-content/uploads/2018/05/02095804/Insect-Management-In-Field-Corn-20181.pdf

Purdue University Field Crops IPM. White Grubs: https://extension.entm.purdue.edu/fieldcropsipm/insects/corn-whitegrubs.php

Reisig, D., and E. Goldsworthy. Efficacy of Insecticidal Seed Treatments and Bifenthrin In-Furrow for Annual White Grub, Arthropod Management Tests, Volume 43, Issue 1, 2018, tsx135-136

https://academic.oup.com/amt/article/43/1/tsx135/4781660

https://academic.oup.com/amt/article/43/1/tsx136/4781661

https://academic.oup.com/amt/article/43/1/tsx137/4781662

Shetlar, D.J., and J. Andon. Identification of White Grubs in Turfgrass: https://ohioline.osu.edu/factsheet/hyg-2510

Whalen, J., and B. Cissel. Soil Insect Management in Field Corn: http://extension.udel.edu/factsheets/soil-insect-management-in-field-corn-2/

Guess the Pest! Week #24 Answer: European Corn Borer

Bill Cissel, Extension Agent – Integrated Pest Management, University of Delaware; bcissel@udel.edu

Congratulations to Grier Stayton for correctly identifying the insect as a European corn borer and for being selected to be entered into the end of season raffle for $100 not once but five times. Everyone else who guessed correctly will also have their name entered into the raffle. Click on the Guess the Pest logo to participate in this week’s Guess the Pest challenge!

Guess the Pest Week #24 Answer: European Corn Borer

It’s hard to believe that a pest that once caused an estimated annual economic loss of $1 billion dollars in the United States is now a rare occurrence. The European corn borer (ECB), as the name implies, is actually native to Europe and was introduced into North American in the early 1900s. In addition to being a pest of corn (field corn and sweet corn), it is also considered a pest of many vegetable and field crops. Since the adoption of transgenic corn hybrids in the mid-1990s, losses due to ECB have been virtually eliminated in Bt crops and significantly reduced in other vegetable and non-Bt field crops. This is one of the pests that the UD Insect Trapping Program monitors with black light traps. The reason we continue to monitor ECB populations throughout the state is because even though generally speaking, populations have been low, there are still local pockets where ECB is causing damage. The photo above of the ECB larva was taken on the Eastern Shore of VA by Helene Doughty from a non-BT sweet corn plot that was 100% infested with ECB.

For information on the benefits of Bt adoption, read this article: Regional pest suppression associated with widespread Bt maize adoption benefits vegetable growershttp://www.pnas.org/content/early/2018/03/06/1720692115

Guess the Pest! Week #22 Answer: Helicoverpa zea, Corn Earworm

Bill Cissel, Extension Agent – Integrated Pest Management, University of Delaware; bcissel@udel.edu

Congratulations to Amanda Heilman for correctly identifying the insect as an adult corn earworm and for being selected to be entered into the end of season raffle for $100 not once but five times. Everyone else who guessed correctly will also have their name entered into the raffle. Click on the Guess the Pest logo to participate in this week’s Guess the Pest challenge!

Guess the Pest Week #22 Answer: Helicoverpa zea, commonly known as corn earworm

The moth in the photograph is an adult Helicoverpa zea, commonly referred to as a corn earworm. The adult moth is a nectar feeder and not considered a pest. However, corn earworm larvae are considered by some to be the most economically important crop pest in North America. They are highly polyphagous meaning they feed on many different species of plants. Corn, especially sweet corn, is a preferred host plant. However, they also attack soybean, sorghum, snap bean, tomato, and cotton to name a few. Larvae prefer to feed on reproductive plant structures including blossoms, buds, and fruits. It is because of this large host range, and the fact that Helicoverpa zealarvae are so destructive that they are known by several other common names including tomato fruitworm, cotton bollworm, and podworm.

For the latest trap counts for corn earworm in your region, visit mdmothmap.com.

Guess the Pest! Week #20 & 21 Answer: Trissolcus japonicus

Bill Cissel, Extension Agent – Integrated Pest Management, University of Delaware; bcissel@udel.edu

Congratulations to Greg Hawn (week 20 winner) and to Joseph Streett (week 21 winner) for correctly identifying the insect as Trissolcus japonicus and for being selected to be entered into the end of season raffle for $100 not once but five times. Everyone else who guessed correctly will also have their name entered into the raffle. Click on the Guess the Pest logo to participate in this week’s Guess the Pest challenge!

Guess the Pest Week #20 – 21 Answer: Trissolcus japonicus
Bill Cissel, Extension Agent – Integrated Pest Management, Joe Kaser, Research Associate, USDA-ARS Beneficial Insects Introduction Research, and David Owens, Extension Entomologist

Trissolcus japonicus, a tiny wasp commonly referred to as the Samurai wasp, is an egg parasitoid of the invasive brown marmorated stink bug (BMSB). This particular species is native to Asia and has been in quarantine since 2007 and under evaluation for potential release as a classical biological control agent. In 2014, wild populations of Trissolcus japonicus, slightly different from the ones that were in quarantine, were detected in Beltsville, MD and since, additional discoveries have been made throughout the region, including Washington, D. C., Virginia, West Virginia, Pennsylvania, New Jersey, Ohio, and Delaware. It is believed that Trissolcus japonicus may have hitchhiked a ride in a BMSB egg mass that was on plant cargo shipped from Asia, but it is difficult to say exactly how it got here. In fact, it appears that the samurai wasp has hitchhiked here more than once!

A single Trissolcus japonicus female is capable of parasitizing an entire BMSB egg mass which typically contains ~28 eggs. When the male parasitoids emerge, they wait on the egg mass for the females to emerge so they can mate. They are capable of having up to ten generations per year.

To help with reducing BMSB populations in Delaware, we partnered with some of the folks at the USDA Beneficial Insects Introduction Research Laboratory in Newark, DE to redistribute Trissolcus japonicas throughout the state. When I share that we are releasing a parasitic wasp to help with BMSB control, the first reaction that I typically get is, “Will it sting me?” If you look at the photo with some wasps on the dime, you will understand why this is not a concern. Hopefully, this tiny wasp will live up to its name as the Samurai wasp and do its part in controlling BMSB.

Fun Entomology Fact: A female Trissolcus japonicus will chemically mark the BMSB eggs that she laid eggs in and defend them against other parasitoids.

Here is a link to a very informative fact sheet from UF on Trissolcus japonicus: http://entnemdept.ufl.edu/creatures/beneficial/wasps/Trissolcus_japonicus.htm

Guess the Pest! Week #18 Answer: Western Bean Cutworm

Bill Cissel, Extension Agent – Integrated Pest Management, University of Delaware; bcissel@udel.edu

Congratulations Bob Leiby for correctly identifying the moth as a western bean cutworm and for being selected to be entered into the end of season raffle for $100 not once but five times. Everyone else who guessed correctly will also have their name entered into the raffle. Click on the Guess the Pest logo to participate in this week’s Guess the Pest challenge!

Guess the Pest Week #18 Answer: Western Bean Cutworm
by Bill Cissel, Extension Agent, Integrated Pest Management and David Owens, Extension Entomologist

The western bean cutworm (WBC) is native to the western United States where it is considered a pest of corn and dry beans. Despite the name, they actually do not “cut” plants. Western bean cutworm are univoltine, meaning they have a single generation per year and overwinter as pre-pupa. In the spring, they pupate and adult moths emerge in early June. Female moths will lay eggs throughout July and August on both wild and cultivated plants. Field corn in the whorl stage prior to pollination is a preferred oviposition site. Eggs are typically laid on the upper leaf surface near the whorl in masses of 20-200 eggs which take approximatley 7 days to hatch. Larvae undergoe six instars before burrowing into the soil to pupate. Since the early 2000s, WBC has spread, causing economic damage as far east as NY, MI, OH, WI, and Ontario. Studies conducted in Nebraska and Iowa suggest an infestation averaging one larva per ear can cause yield losses reaching as high as 4 bu/A. Larvae bore through the side of the ear and open the ear up to mycotoxin-causing fungal colonization. Most Bt traits do not adequatley control this pest.

We first detected WBC in Delware in 2011 after capturing a few moths in a pheromone trap in New Castle Coutny. We captured 14 moths in 2012 and have not trapped for this pest since 2012 until 2018. This year, we have been monitoring 10 pheromone traps located throughout the state and have captured four moths to date. We will continue to monitor for this pest throughout the growing season but at this point, it appears that WBC populations remain low for us in Delaware. By comparison, states where western bean cutworm causes signficant injury to corn catch dozens of moths per week in a single trap.

2012 Western Bean Cutworm Trap Summary: http://s3.amazonaws.com/udextension/ag/files/2012/06/2011WesternBeanCutwormTrapSummary.pdf

2013 Western Bean Cutworm Trap Summary: http://s3.amazonaws.com/udextension/ag/files/2012/06/2012-Western-Bean-Cutworm-Trap-Summary2.pdf

Here is a link to a Fact Sheet from Purdue University with more detailed information on the identification, biology, and damage of the Western Bean Cutworm: https://extension.entm.purdue.edu/fieldcropsipm/insects/western-bean-cutworm.php

Fun Entomology Fact: It is not unusual to find an ear infested with multiple western bean cutworm larvae because they are not canabalistic like corn earworms.

Guess the Pest! Week #17 Answer: Soybean Leafminer

Bill Cissel, Extension Agent – Integrated Pest Management, University of Delaware; bcissel@udel.edu

Congratulations Julie Knudson for correctly identifying the damage in the photo as soybean leafminer damage and for being selected to be entered into the end of season raffle for $100 not once but five times. Everyone else who guessed correctly will also have their name entered into the raffle. Click on the Guess the Pest logo to participate in this week’s Guess the Pest challenge!

Guess the Pest Week #17 Answer: Soybean Leafminer
By David Owens, Extension Entomologist, owensd@udel.edu

This week’s Guess the Pest is an interesting but rather unimportant member of the defoliating insect complex. The soybean leafminer (Odontota horni) adult is a beautiful red, flattened, rectangular beetle with red wings and prothorax and black head, antennae, legs, and a black stripe down the middle of the back. The black stripe doesn’t reach all the way to the end of the wings. Adults are active beginning around early to mid-June. They lay eggs on the underside of leaves, and the larvae immediately mine into the leaf. Larvae spend their entire lives between the upper and lower leaf surface, leaving a quarter sized brown blotch. When larvae complete development, they pupate in the mine. Immatures require 30 – 40 days to fully develop into adults. There is only one generation per year. Beetles will continue to lightly skeletonize leaves and over the course of their adult life might feed on the equivalent of one leaflet until they migrate out of fields to find overwintering shelter in late summer. Beetles and larvae are never present in any significant populations.

There are a couple of other beetle leafminers that you may see this summer. The most obvious and abundant is the locust leafminer, Odontota dorsalis. It pretty much stays confined to locust and can cause a large amount of locust defoliation by August. So, as you drive along the highway and notice trees with a brown cast to them, you may be seeing locust leafminer. Locust trees can handle the defoliation and leaf back out.

Guess the Pest! Week #16 Answer: Spider Mite

Bill Cissel, Extension Agent – Integrated Pest Management, University of Delaware; bcissel@udel.edu

Congratulations Jeffrey O’Hara for correctly identifying the damage in the photo as spider mite damage and for being selected to be entered into the end of season raffle for $100 not once but five times. Everyone else who guessed correctly will also have their name entered into the raffle. Click on the Guess the Pest logo to participate in this week’s Guess the Pest challenge!

Guess the Pest Week #16 Answer: Spider Mite

The damage in the photo is from two-spotted spider mite (TSM) feeding on soybean. Hot, dry weather favors TSM and drought can trigger outbreaks. TSM populations are held in balance by natural enemies and the weather. Under high temperatures, the amount of time required for TSM to complete its lifecycle is shortened, allowing more generations to be completed in a shorter period of time. A female TSM can produce 300 offspring in her lifetime (~30 days) and most of the individuals in the population are female. Dry conditions also diminish the activity of fungal diseases that often play a key role in keeping outbreaks from occurring.

So, if it rains, does it mean we don’t need to worry about TSM? Precipitation not only favors spore formation and mite infection but also reduces plant stress. This however isn’t always a silver bullet and TSM populations can continue to increase even after rain events, especially if the weather returns to being hot and dry. Cool nights and humid conditions promote the fungal disease that infects TSM.

Below is a graph showing rain events and observed TSM populations in the untreated check from a TSM field trial conducted in Georgetown, DE in 2017:

Observed Influence of Precipitation on Two-Spotted Spider Mite Populations, 2017

Weather data obtained from the Delaware Environmental Observing System (DEOS): http://www.deos.udel.edu/data/agirrigation_retrieval.php

As you can see in the graph, TSM populations continued to increase and remained high despite rain events occurring on 7/22, 7/23, 7/25, and 7/28.

To scout for TSM, examine the underside of 5 leaflets in 10 locations for mites, noting the presence of mite eggs and the amount of leaf damage. The threshold for TSM during bloom to podfill is 20-30 mites per leaflet and 10% of plants with 1/3 or more leaf area damaged.

Concentrate scouting efforts on field edges for initial detection, especially edges bordered by grass and road ditches (it’s not unusual to also find hot spots in the interior portions of the field). TSM typically develop on grasses and other plants on field borders before ballooning into fields. Once TSM are detected, scout the interior portions of the field to determine if they have spread throughout the entire field. If only concentrated on field edges, spot treating may be an option. If spot treating on field edges, extend the treated area about 100 feet further into the field from the damaged area.

Here is a link to our Soybean Insecticide Recommendations for chemical control options if your field is at threshold for TSM: https://cdn.extension.udel.edu/wp-content/uploads/2018/05/02102500/Insect-Control-in-Soybeans-2018.pdf

Can Timing and Method of Barley Cover Crop Termination Impact Pests and Beneficials within a Subsequent Soybean Planting?

Alan W. Leslie, Armando Rosario-Lebron, Guihua Chen and Cerruti R. R. Hooks
Department of Entomology, College of Computer, Mathematical, and Natural Sciences

Summary

This extension article is meant to serve as a condensed write-up of a completed field study. Full-text of the published work can be viewed via open access at http://www.mdpi.com/2073-4395/8/6/87. Cover cropping has long been used as a method of reducing soil erosion, increasing soil quality, and suppressing weeds. However, impacts of cover crops in cropping systems differ and can be affected by timing and method of their termination. Field trials were conducted over two field seasons and at two sites in Maryland to examine how varying the date and method of terminating a barley (Hordeum vulgare) winter cover crop affects arthropods (insects and spiders) in succeeding no-till soybean (Glycine max) plantings. Experimental treatments included early-kill with pre- and post-emergent herbicides (EK), late-kill with pre- and post-emergent herbicides (LK), late-kill with a flail mower and pre-emergent herbicide (FM), and a fallow/bare-ground check with pre- and post-emergent herbicides (BG). Terminating barley late (i.e., just prior to soybean planting) resulted in significantly greater biomass accumulation in LK and FM than EK. However, method and timing of termination had no effect on communities of pest and beneficial arthropods in the soybean canopy. Results from this experiment suggest that terminating the cover crop early or late or using a mower or burn-down herbicide to kill the cover crop will result in similar species and number of arthropods within the soybean canopy.

Introduction

Cover cropping can be a viable weed management tool in conservation agriculture systems. When cover crops are terminated in reduced- and no-till cropping systems, resulting residues that remain on the soil surface can help prevent weed establishment. Thus, it is well known that cover crop residue impacts weed populations. More specifically, some of these studies were designed to examine how method and timing of cover crop termination practices impact weed populations in grain crops. However, impacts of these practices on arthropod populations are rarely considered. Despite this, studies have shown that cover crops can impact arthropod numbers in succeeding agronomic crops. Some insect pests shown to be impacted by cover crop residue include the potato leafhopper (Empoasca fabae), bean leaf beetle (Cerotoma trifurcata), and Japanese beetle (Popillia japonica) in soybean as well as thrips in cotton (Gossypium hirsutum). In addition to insect pests, their natural enemies may be influenced by cover crop residue.

The overall goal of this study was to investigate how different cover crop termination practices impact populations of insect pests and their natural enemies within no-till soybean plantings. Specific objectives were to compare the influence of termination method (chemical versus mechanical) and timing (early versus late) on arthropod populations. Barley was chosen as the test cover crop partially because of its accessibility and popularity among producers.

Materials and Methods

Field experiments were conducted at the University of Maryland’s Central Maryland Research and Education Center at the Upper Marlboro and Beltsville farm sites in 2013 and 2014. Each field experiment consisted of four treatments, including three cover crop termination methods and a fallow/bare-ground control. The three cover crop treatments included: (1) early-kill (EK), in which the cover crop was sprayed with post- and pre-emergent herbicides in mid-April; (2) late-kill (LK), in which the cover crop was sprayed with post- and pre-emergent herbicides in late May; and (3) flail-mowed (FM), in which the cover crop was sprayed with a pre-emergent herbicide and mowed in late May. An early-kill, flail-mowed treatment was not included in the experiment because mowing typically does not kill cover crops at early stages of development and farmers do not use this method. The bare-ground treatment (BG) remained fallow after the previous crop was harvested and received the same post- and pre-emergent herbicide applications as LK.

The EK treatment was sprayed with a post- and pre-emergent herbicide mixture on 15 April at Beltsville and 16 April at Upper Marlboro in 2013 and on 18 April at both sites in 2014. The LK treatment was sprayed with a post- and pre-emergent herbicide mixture on the day soybeans were planted. The BG treatment received the same spray protocol as LK. On the day soybeans were planted, the FM treatment was sprayed with a pre-emergent herbicide and the cover crop was mowed. The soybean was planted on 21 May at Beltsville and 20 May at Upper Marlboro in 2013 and 27 May 2014 at both sites. Soybeans were planted in wide rows [76 cm (30 inch) inter-row spacing] at Beltsville and narrow rows [18 cm (7 inch) inter-row spacing] at Upper Marlboro. A late-season herbicide application was applied to all plots at the Beltsville location as a “rescue” herbicide treatment primarily for large crabgrass (Digitaria sanguinalis).

Data collection. Data on vegetative (cover crop and weed) biomass, abundance of weeds and arthropods, soil moisture as well as yield were collected during this investigation. Data on weed population, soil moisture and yield will be presented in a future edition of Agronomy News. To quantify cumulative barley and weed biomass production, cover crop and weed biomass were measured in each plot just prior to their termination. Arthropod abundances were monitored with the use of a sweep net from the R1 through R5 soybean growth stages. Arthropods collected were divided into i) natural enemies (predators – arthropods that prey on herbivores & parasitoids – insects, especially wasps, that complete their development within the body of another insect eventually killing it) and ii) herbivores (insects that feed on plants). Arthropods were separated further according to seven feeding habits (guilds). The seven feeding habits that we used included 1) chewing predators, 2) sucking predators, 3) parasitoids, 4) plant-sucking herbivores, 5) pod feeders, 6) foliage feeders and 7) spiders. Though they are predators, spiders were placed into a separate predatory feeding guild.

Results

Vegetative biomass. At each farm site, flail-mowed (FM) and late-killed (LK) treatments had higher plant biomass than early-killed (EK) or bare-ground (BG) treatments (Table 1). Total barley biomass in LK and FM treatments were more than two times greater at Beltsville than Upper Marlboro. No differences were detected in plant biomass between BG and EK treatments within each site, but there was greater weed biomass in the BG treatment at Beltsville than Upper Marlboro (Table 1).

Table 1. Cover crop and weed dry biomass just prior to their termination.

Site	Treatment	Mass ± SEM (kg ha−1)
Beltsville	Early Kill	160.1	±	60.5	cd1
	Late Kill	2211.9	±	83.2	a
	Flail Mow	2123.4	±	112.9	a
	Bare Ground	896.0	±	254.3	bc
Upper Marlboro	Early Kill	85.8	±	21.2	d
	Late Kill	753.4	±	100.9	b
	Flail Mow	851.8	±	62.7	b
	Bare Ground	120.4	±	27.2	d

1Different letters indicate that means are significantly different.

Arthropod Counts. In total, 54 families of arthropods were collected from sweep samples which included a total of 11,344 specimens (Table 2). Approximately 98% of arthropods collected could be assigned to one of the seven feeding guilds used. Three feeding guilds, which included plant-sucking herbivores (25%), foliage-feeding herbivores (24%), and sucking predators (21%), accounted for 70% of the entire arthropod community sampled.

The abundance of arthropods from each feeding guild was similar among treatments. However, there was a significant effect of soybean development stage on all feeding guilds. In general, parasitoid, chewing predator, and sucking predator guilds reached greatest abundance later in the season (R4 or R5 stage). In contrast, numbers of foliage feeding and plant sucking herbivores peaked earlier in the growing season at the R2 or R3 stage (Table 3). Sucking predators and spiders were found in greater numbers in Beltsville than Upper Marlboro across all soybean growth stages.

Table 2. Arthropod feeding guilds, families and their abundances. Numbers represent total abundance across all sample dates.

		Beltsville		Upper Marlboro
Feeding Guild	Family	2013	2014	2013	2014
Spider	Salticidae	25	47	24	48
	Araneidae	5	69	0	37
	Oxyopidae	149	101	42	94
	Thomisidae	18	16	35	11
	Lycosidae	0	12	0	26
	Clubionidae	0	3	0	0
	Ctenidae	0	1	0	0
	Tetragnathidae	0	8	0	9
	Linyphiidae	0	4	0	2
	Pholcidae	0	1	0	0
Parasitoid	Platygastridae	159	11	57	0
	unspecified1	0	407	0	104
	Sceleonidae	10	23	1	15
	Chalcididae	0	2	0	3
	Proctotrupidae	0	1	0	2
	Braconidae	0	76	0	29
	Eulophidae	0	18	0	5
	Ichneumonidae	0	8	0	3
	Tiphiidae	0	178	0	25
	Aphelinidae	0	1	0	1
	Encyrtidae	0	1	0	0
	Mymaridae	0	0	0	1
	Eurytomidae	0	0	0	1
	Trichogrammatidae	0	0	0	2
Chewing predator	Asilidae	5	6	0	0
	Mantidae	1	1	1	0
	Coccinellidae	21	262	36	193
	Carabidae	0	5	0	3
	Syrphidae	0	101	0	4
	Cantharidae	0	0	0	1
Sucking predator	Geocoridae	543	346	223	326
	Pentatomidae	3	8	1	1
	Chrysopidae	5	1	3	13
	Anthocoridae	48	166	225	161
	Nabidae	100	340	37	93
	Hemerobiidae	0	10	0	3
	Reduviidae	0	0	0	4
Foliage feeder	Coccinellidae	287	92	43	1
	Erebidae	346	756	274	455
	Meloidae	0	1	0	0
	Scarabaeidae	428	284	90	96
	Chrysomelidae	2	345	0	254
	Noctuidae	0	1	0	0
	Hesperiidae	0	3	0	0
Plant sucking	Cicadellidae	32	732	109	689
	Membracidae	22	0	40	18
	unspecified	404	0	896	0
	Aphididae	0	0	0	60
Pod feeder	Pentatomidae	33	164	69	115
	Miridae	112	102	108	229
Unassigned	unspecified	0	2	67	30
	Chrysomelidae	0	0	0	2
	Curculionidae	2	5	0	30
	Lampyridae	4	21	17	5
	Lygaeidae	0	0	0	0
	Elateridae	18	5	0	12
	Noctuidae	0	0	0	1
	Apidae	0	0	1	0
	Cynipidae	0	0	18	3
	Vespidae	0	0	5	8
	Chrysididae	0	0	3	3
	Pompilidae	0	0	1	0
	Scoliidae	0	0	1	0
	Thyreocoridae	0	0	0	14
	Berytidae	0	0	0	41
	Alydidae	0	0	0	2

1Unspecified taxa were not identified to the family level.

Table 3. Means (± standard errors) of feeding guilds within farm site and soybean development stage.

		Abundance per 10 Sweeps
Feeding Guild	Site1	R1				R2				R3				R4				R5
Spider	BV	1.22	±	0.26	a2	0.91	±	0.20	a	1.27	±	0.13	a	2.02	±	0.25	a	1.28	±	0.22	a
Spider	UM	1.19	±	0.33	a	0.58	±	0.11	a	0.96	±	0.10	a	1.19	±	0.18	a	1.25	±	0.19	a
Parasitoid	BV	0.97	±	0.20	b	1.28	±	0.28	ab	1.23	±	0.15	ab	3.33	±	0.52	a	4.81	±	0.92	a
Parasitoid	UM	0.22	±	0.11	b	0.56	±	0.12	ab	0.72	±	0.14	ab	0.97	±	0.16	ab	1.28	±	0.38	a
Chewing predator	BV	0.63	±	0.33	b	0.13	±	0.05	ab	0.82	±	0.15	ab	1.25	±	0.24	ab	1.56	±	0.30	a
Chewing predator	UM	0.28	±	0.10	b	0.02	±	0.02	ab	0.48	±	0.10	ab	1.59	±	0.31	ab	1.50	±	0.31	a
Sucking predator	BV	3.75	±	0.69	b	3.41	±	0.51	b	4.03	±	0.37	ab	8.27	±	0.65	a	6.41	±	0.73	ab
Sucking predator	UM	2.13	±	0.34	b	2.59	±	0.35	b	2.03	±	0.18	ab	7.84	±	1.05	a	5.91	±	0.64	ab
Foliage feeder	BV	8.78	±	0.92	b	7.50	±	0.86	ab	9.62	±	0.70	a	6.25	±	0.85	b	4.06	±	0.86	b
Foliage feeder	UM	2.81	±	0.95	b	1.84	±	0.26	b	5.17	±	0.41	a	2.89	±	0.36	ab	2.41	±	0.52	b
Plant sucking	BV	6.81	±	1.16	ab	5.66	±	0.61	a	2.73	±	0.21	ab	3.20	±	0.43	b	2.22	±	0.32	b
Plant sucking	UM	3.81	±	0.72	a	8.61	±	1.01	a	2.97	±	0.19	a	15.0	±	3.66	a	8.38	±	1.77	a
Pod feeder	BV	0.81	±	0.24	a	0.64	±	0.15	a	1.23	±	0.16	a	0.50	±	0.14	a	1.34	±	0.36	a
Pod feeder	UM	1.00	±	0.21	a	1.73	±	0.28	a	1.09	±	0.15	a	1.69	±	0.29	a	3.50	±	0.66	a

1BV = Beltsville, UM = Upper Marlboro

2Different letters within individual rows represent significant differences between growth stages.

Discussion

The objective of this study was to quantify the impact of cover crop termination method and timing on arthropods within soybean foliage. Cover crop termination practices are known to impact arthropods via resulting residues that remain on the soil surface. Thus, it was believed that different cover crop termination methods examined during this study would influence arthropod abundances differently. As expected, delaying the cover crop termination date resulted in significantly greater biomass of residue in late-kill (LK) and flail-mowed (FM) than in early-kill (EK) treatments. Averaged across years, delaying cover crop termination in FM and LK increased barley biomass relative to EK by 2007.5 kg ha−1 (1791 lbs/acre) at Beltsville and 716.8 kg ha−1 (639.5 lbs/acre) at Upper Marlboro. However, arthropod populations within the soybean foliage responded similarly to treatments regardless of plant biomass differences. Instead, arthropod abundances changed according to soybean growth stage. Chemical (LK) and mechanical (FM) termination tactics also had similar effects on arthropod abundances. This suggests that whether cover crops are killed early or late, or chemically or mechanically by mowing, the resulting arthropod community will be similarly impacted. The EK “early” (early April) and LK “late” (at soybean planting from mid to late May) treatments represent some of the most widely used practices for cover crop termination by Mid-Atlantic soybean producers. The results of our study suggest that these two practices are likely to result in similar species and number of foliar arthropods throughout the different soybean growth stages.

Conclusions

Overall, our results indicate that cover crop termination methods that result in greater cover crop biomass will have no effect on insects and spiders within the soybean foliage. However, if delaying cover crop termination results in greater weed suppression without impacting soybean productivity, this practice should nevertheless make soybean systems more resilient to pest pressure and acceptable by soybean producers.

Acknowledgments

We thank crews at the Upper Marlboro and Beltsville Research and Education Centers for logistics in establishing and completing field trials. This work or publication was supported by Hatch Project No. MD-ENTO-9107/project accession no. 227029 and the Crop Protection and Pest Management (CPPM), Extension Implementation Program (EIP) award number 2017-70006-27171 from the USDA National Institute of Food and Agriculture, and funding from the Maryland Soybean Board.

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