Read and follow all label requirements for insecticides.
Soybean: Continue checking for defoliators, such as bean leaf beetle, Japanese beetle, grasshoppers, and caterpillars. Control may be needed if there is 15% defoliation on plants from bloom through pod fill. As we move into the heat of the summer, keep an eye out on your aphid population, which may increase quickly. The summer generation of soybean podworms are emerging. Fields next to maturing corn, have open canopies, are drought-stressed, or have recently had insecticide applied are at high risk for podworms. While worm feeding on flowers will not impact yields, feeding during pod development can. North Carolina State University has a great economic threshold calculator (https://soybeans.ces.ncsu.edu/wp-content/uploads/2017/08/CEW-calculator-v0.006.html) .
Alfalfa: Continue scouting for leafhopper and blister beetles.
Sorghum: Sugarcane aphids showed up in fields last year in August. Check the underside of leaves for insects. The threshold is 50 aphids per leaf on 25 – 30% of plants. They have shown some resistance to pyrethroids.
Check head for head worms and fall armyworms once heads have started to flower. Check 10 spots per field, 5 plants per spot. An easy scouting method is to use a 5-gallon bucket and shake the head into it and then count the number of medium (1/4 – 1/2 inches) and large (> 1/2 inch) dislodged caterpillars. Texas A&M has a great threshold calculator that takes the grain price and treatment into consideration (https://extensionentomology.tamu.edu/sorghum-headworm-calculator/).
Adapted from Maryland State Horticulture Society Newsletter
After several years of debate in Annapolis to ban the use of this product in Maryland, the final decisions was made to allow the phase out of this product. The manufacturer will discontinue production of this product. Instead of an outright ban, MDA has developed the phase out process which is listed below. This can be found on page 442 of the Maryland Register, Volume 47, Issue 8 dated April 10, 2020.
.02 General Requirements for Applying or Recommending Pesticides.
A.—D. (text unchanged)
Restrictions on Use of Insecticides that Contain Chlorpyrifos.
(1) Aerial Applications Prohibited. A person may not conduct an aerial application of any insecticide containing Chlorpyrifos in the State.
(2) Other Applications Generally Prohibited After December 31, 2020.
(a) Except as provided in §E(2)(b) and (c) of this regulation, after December 31, 2020, a person may not apply an insecticide containing Chlorpyrifos or seeds that have been treated with Chlorpyrifos in the State for any use.
(b) Fruit Trees and Snap Bean Seeds. Until June 30, 2021, a person may use an insecticide containing Chlorpyrifos or seeds that have been treated with Chlorpyrifos in the State to treat snap bean seeds and the trunks and lower limbs of fruit trees. After June 30, 2021, such applications are prohibited unless authorized by the Secretary under §E(2)(c) of this regulation.
(c) Limited Particular Use Authorization. After December 31, 2020, a person may file a written application with the Department requesting authorization to use an insecticide that contains Chlorpyrifos or seeds that have been treated with Chlorpyrifos for a particular use. If the Secretary has determined that there are no effective alternatives for the particular use noted in the application, the Secretary may authorize such use for a specified period of time, which may not extend beyond December 31, 2021.
(3) Establishment of Committee. The Secretary shall establish a committee, with members appointed by the Secretary, to determine alternatives to using Chlorpyrifos or seeds that have been treated with Chlorpyrifos, which shall dissolve on December 31, 2021.
This allows the use of this product as listed above. Please note the important dates. Until June 30, 2021, a person may use an insecticide containing Chlorpyrifos or seeds that have been treated with Chlorpyrifos in the State to treat snap bean seeds and the trunks and lower limbs of fruit trees. After June 30,2021, such applications are prohibited unless authorized by the Secretary under §E(2)(c) of this regulation. Use these products carefully .
Kelly Hamby1, Maria Cramer1, Galen Dively1, Sarah Hirsh2, Andrew Kness2 Alan Leslie2, Kelly Nichols2, Emily Zobel2, and David Owens3 1University of Maryland, Department of Entomology | 2University of Maryland Extension 3University of Delaware Extension
A few hot spots where corn earworm (also known as tomato fruitworm, soybean podworm, and sorghum headworm) activity is starting to rise have been identified in central Maryland and Delaware. The warm 2019-2020 winter allowed for overwintering in our area, and some parts of the state experienced a higher than normal first flight in early June. The warm weather through June and July made for speedy development and earlier activity for the second summer generation. Because corn earworm has developed resistance to most Bt hybrids, significantly more adult moths are emerging compared to levels a decade ago. Some areas continue to capture few moths and are experiencing low pressure, while others have been experiencing moderate pressure that may continue to increase towards heavy pressure (>65 moths captured per 5 days). Captures for select sites in Maryland and Delaware are pictured below, and values within the gray box indicate low pressure (<7 for weekly captures, and <5 for four to five day captures).
Corn earworm larva feeding damage to corn
Although corn earworm prefer fresh corn silks for egg laying, they will lay eggs on wilted and brown silks if the plants remain green and unstressed. As corn matures further over the next several weeks, corn earworm activity will shift to other host plants including soybeans and vegetables. See last summer’s articles for scouting and management recommendations in vegetables as well as sorghum and soybeans.
Podworm outbreaks have historically occurred in growing seasons where the corn crop was drought and heat stressed, with corn senescing earlier than normal. However, narrow row spacing in soybeans makes the plants less attractive to female moths and increases the likelihood that fungal pathogens will infect the larvae. Therefore, it is important to scout bean fields, especially paying attention to those fields with a more open canopy in areas where the nearby maturing corn is no longer attractive to earworm moths. North Carolina State University has produced a helpful economic threshold calculator for podworm in soybean: https://www.ces.ncsu.edu/wp-content/uploads/2017/08/CEW-calculator-v0.006.html
Acknowledgements: Corn earworm trapping efforts in were supported by the Crop Protection and Pest Management Program [grant numbers 2017-70006-27171 and 2017-70006-27286] from the USDA National Institute of Food and Agriculture. Any opinions, findings, conclusions, or recommendations expressed in this publication are those of the author(s) and do not necessarily reflect the view of the U.S. Department of Agriculture.
Emily Zobel, Agriculture Agent Associate University of Maryland Extension, Dorchester County
Soybean: The usual defoliators are starting to arrive, including bean leaf beetle, Japanese beetle, grasshoppers, and caterpillars. Control may be needed if there is 30% defoliation during the seedling and vegetative stages and 15% defoliation once plants start to bloom through pod fill.
Adult Dectes Stem borer will be emerging over the next several weeks. Chemical control is not recommended since it would require multiple applications to reduce larval infestations, which is not economical. If a high number of adults are found, harvesting that field as soon as it matures will reduce losses associated with lodged plants.
Fields that have an open canopy, drought-stressed, or have recently had an insecticide applied are at higher risk for corn earworm (CEW). CEW larva can feed on flowers without impacting yields because soybeans overproduce flowers. However, feeding during pod development can affect yield. An economic threshold calculator is available to assist with management decisions: https://soybeans.ces.ncsu.edu/wp-content/uploads/2017/08/CEW-calculator-v0.006.html.
Field Corn: As corn ears begin to form, check for stink bugs. Stink bugs will gather around the edges of fields, so scouting should be done at least 15 rows in. Thresholds are 1 stink bug per 4 plants when the ear is forming, and 1 stink bug per 2 plants from pollen shed to blister stage. Treatment is not recommended past the blister stage. Japanese beetles are minor defoliators and will clip corn silks, but control is not needed unless silks are cut back to less than ½ inch, and less than half the field has been pollinated.
Alfalfa: Once plants have hopper burn, there is no way to undo it, so continue scouting for leafhopper. Since infestations are highly variable, individual fields should be scouted. If you are planning on selling your hay for horse feed, check for blister beetle as well since they produce cantharidin, which causes skin blisters on humans and can make horses sick.
Sorghum: Sugarcane aphids were found on the Eastern Shore last year and typically show up in fields late July and August. Check underside of leaves for insects. Honeydew will turn leaves shiny and is an easy to see indicator that aphids are present. Sugarcane aphids are light yellow with black cornicles, antennae, and feet. Thresholds depend on plant growth stage; at boot to milk, thresholds are 50 aphids per leaf on 25 – 30% of plants. There is documented resistance to resistance to pyrethroids.
Laura C. Moore^,*, Alan W. Leslie#, Cerruti RR Hooks$,* and Galen P. Dively+,* Former graduate student^, Associate Professor and Extension Specialist$, Professor Emeritus+, CMNS, Department of Entomology*, Agriculture Extension Agent, Charles County#
Introduction
Increasing floral diversity within agricultural fields has been proposed as a method to bolster natural enemies and subsequently reduce pest populations. A key factor that enhances predator and parasitoid populations is the availability of nectar and/or pollen food subsidies from flowering plants. Many natural enemies, particularly hymenopteran (wasps) parasitoids, require carbohydrates for successful reproduction and overall fitness. However, monoculture cropping systems are relatively weed-free and generally lack floral resources required by many natural enemies. A literature review showed that the successful establishment of certain parasitoids in cropping systems depended on the presence of nectar-bearing weeds. In addition to providing natural enemies nectar and pollen to eat, flowering plants can supply alternative hosts or prey, shelter, overwintering sites and a more suitable microclimate.
Many conservation projects have been implemented by farmers to increase beneficial services on arable lands. In Maryland, the opportunity to practice conservation biological control exists within the Conservation Reserve Enhancement Program (CREP). Conservation biological control is a pest management approach that manipulates agricultural systems so as to promote pest suppression by naturally occurring predators, parasitoids and pathogens. The CREP seeks to establish riparian buffers in Maryland to improve water quality, filter sediments and nutrients from runoff and provide wildlife habitat. However, these buffers can be engineered to support communities of natural enemies and serve as corridors for their movement into neighboring crops.
The aim of this study was to determine whether buffer strips could be used as insectary plants to enhance beneficial arthropods (insects and spiders) within neighboring soybeans. Insectary plants are plants grown with cash crops to attract, feed and shelter insect parasitoids and predators so as to enhance their natural control of insect pests. We monitored pest and beneficial arthropods in the buffer insectary plants and neighboring soybean plantings and tried to link arthropods found in buffer plants with pest management in neighboring soybeans.
Abbreviated Experimental Procedures
Insectary buffer test plants. Partridge pea and purple tansy are commonly used to enhance floral resources along field margins for pollinator plantings and to enhance communities of natural enemies in adjacent crops. Partridge pea is a native annual legume and is widely used in seed mixes of CREP riparian buffers because it readily reseeds itself, is competitive when grown with grass mixes, and provides nutritional seed for game birds. As an insectary plant, partridge pea has a long bloom period and each leaf petiole has an extrafloral nectary at its base, which produces nectar throughout the growing season. A diverse assemblage of pollinators and natural enemies are attracted to partridge pea. Purple tansy has a long flowering period, high-quality nectar and pollen production, and is reported as being a valuable insectary plant. Proso millet is a warm season annual grass that lacks floral resources. As such, it served as a grass control to determine how added vegetation diversity in the absence of floral resources would impact natural enemies.
Experimental design. Field experiments were conducted over two years at the Central Maryland Research and Education Center in Beltsville, MD. In year 1, 16 plots of soybean were seeded on May 11. Each plot consisted of 20 soybean rows spaced 35 cm (15 in) apart and bordered on each side by an insectary buffer strip (Fig. 1). The test buffer strips consisted of 1) purple tansy, 2) partridge pea, 3) 50:50 seed mixture of purple tansy + partridge pea, or 4) proso millet. Each soybean plot-buffer combination was replicated four times. Seeds of partridge pea, purple tansy, and proso millet were planted with a no-till drill in rows 23 cm (9 in) apart at a rate of roughly 12,000 seeds per ha (4856 per ac) on the day soybeans were planted.
Fig. 1. Illustration of a soybean-buffer treatment plot in year 1. Soybean plots were bordered on each side with buffer insectary plants. Buffers included purple tansy, partridge pea, 50:50 seed mixture of partridge pea/purple tansy or proso millet.
The year 1 study showed that purple tansy was unsuitable for the hot summer conditions in Maryland Thus, it was not used in the year 2 experiment, which focused solely on partridge pea as the insectary buffer plant. The year 2 experiment included 14 strip plantings of full season soybean at five different locations (Fig. 2). Soybeans were planted no-till in 75 cm (30 in) wide rows during May. Each strip was bordered at one end with a partridge pea buffer and at the other end with a mixed grass border of fescue (Festuca spp.) and orchardgrass (Dactylis spp.).
Fig. 2. Aerial view of experimental layout in year 2. Study consisted of 14 contour strips of full-season soybeans and adjoining partridge pea buffers (indicated by black polygons) at one end of each strip. Grassy areas were on opposite ends of soybean strips without a buffer.
Arthropod population assessments. Abundances of arthropods active in the plant canopy were measured with yellow sticky cards secured to bamboo poles. Further, sweep-net samples were taken in July and August to estimate green cloverworm (Hypena scabra) numbers. The green cloverworm served as a bioindicator of changes in pest populations potentially caused by enhanced natural enemy activity. The larger field size in year 2 allowed sticky cards to be placed throughout the soybean strip. One card was placed in the center of each partridge pea buffer, and additional cards were placed at distances of 3, 6, 12, 18 and 24 m (10 ft to 79 ft) from the border on both sides of each soybean strip (total of 10 sticky cards per strip). Sampling was conducted weekly or biweekly. In year 2, pitfall traps were also installed in the ground adjacent to each sticky card to estimate the abundance of surface-dwelling arthropods over 7-day intervals.
Summary of Results
Year 1 Study – comparison of four insectary buffers parasitoid abundance. Three families of parasitoids Mymaridae, Scelionidae and Trichogrammatidae comprised 83.9% of the total of parasitic wasps captured on sticky cards. Families Ceraphronidae, Braconidae and Eulophidae comprised an additional 12.5% of the wasp parasitoid group. Of these parasitoids, mymarids were the most abundant and there were 73-78% higher sticky card captures of this wasp in partridge pea compared to purple tansy and millet buffers. However, significantly fewer mymarids were captured in soybeans adjacent to partridge pea than adjacent to purple tansy or millet. Scelionid parasitoids were more abundant in millet and purple tansy buffers but their numbers were similar in soybeans regardless of the neighboring buffer type. Trichogrammatid abundance was greatest in millet early in the season and in buffers with partridge pea by season end. Two families of fly parasitoids (Tachinidae and Sarcophagidae) averaged 9.4 and 4.4 flies per sticky card in insectary buffers and soybean plots, respectively. The abundance of sarcophagid flies was significantly higher in buffers with partridge pea than millet or purple tansy alone. Similarly, soybeans adjacent to partridge pea were inhabited by more tachinids and sarcophagids than soybeans adjacent to millet or purple tansy.
Predator abundance. Overall predator abundance was significantly higher in purple tansy and millet compared to partridge pea or mixed (partridge pea + purple tansy) buffers. Mean captures per card were 5.0 in partridge pea, 6.8 in mixed, 8.5 in purple tansy, and 10.3 in millet. However. similar predator numbers were captured in soybean plots adjacent to all four buffer types.
Insect herbivores (plant feeders). Sweep net counts of green cloverworm were statistically similar in soybean plots adjacent to the four different buffer types. Overall numbers per 10 sweeps averaged 24.6, 27.0, 18.0, and 23.0 in soybeans adjacent to millet, purple tansy, mixed and partridge pea buffers, respectively. The bulk of other insect herbivores captured on sticky cards were mainly aphids, leafhoppers, planthoppers and plant bugs. Mean numbers captured per card were 86.1 (millet), 113.2 (purple tansy), 57.6 (mixed) and 53.7 (partridge pea).
3.2. Year 2 Study partridge pea vs. natural grass vegetation
Parasitoid abundance. The most abundant parasitoids belonged to families Mymaridae, Trichogrammatidae and Scelionidae in order of abundance, and together comprised 84.3% of the total hymenopteran parasitoids captured. Each family responded differently to the partridge pea treatment. Mymarid abundance was higher overall in partridge pea buffers but did not enhance their abundance in neighboring soybeans (Fig. 3). Significantly fewer trichogrammatids were captured in partridge pea compared to numbers captured in soybean with and without the partridge pea buffer. Mean captures of dipteran parasitoids per sticky card abundance were significantly higher in soybean neighboring partridge pea, with the exception of the first and last sampling dates.
Fig. 3. A) Mean number (±SE) of mymarid parasitoids captured per sticky card in partridge pea buffer, soybean neighboring buffer, and soybean without buffer in year 2. Data for soybean were averaged over all sampling distances (3, 6, 12, 18 and 24 m) from the field edges. B) Mean number in soybean at different distances from field edges with and without a partridge pea buffer.
Predator abundance. Long-legged flies, minute pirate bugs, and big-eyed bugs comprised 81.4% of the total predatory arthropods captured. Soldier beetles, fireflies and lady beetles represented an additional 11.6%. Mean abundance of predators per sticky card was 11.5 ± 1.1 in buffer, 4.1 ± 0.16 in soybean neighboring buffer and 4.9 ± 0.18 in soybean without buffer. Abundance of predators was significantly lower in soybean neighboring the partridge pea buffer (Fig. 4). However, this was largely due to the activity of long-legged flies, which were more attracted to the partridge pea buffer. Still, their numbers were significantly lower in soybean strips neighboring partridge pea compared to soybeans without partridge pea buffers.
Fig. 4. A) Mean number (±SE) of arthropod predators captured per sticky card in partridge pea buffer, soybean neighboring buffer and soybean without buffer in year 2. Data for soybean were averaged over all sampling site distances (3, 6, 12, 18 and 24 m) from the field edges. B) Mean number in soybean at different distances from field edges with and without a partridge pea buffer. Arthropod predator guild consisted of long-legged flies, minute pirate bugs, big-eyed bug, soldier beetles, fireflies and lady beetles.
Insect herbivores/pests. Thrips, leafhoppers, treehoppers, froghoppers and planthoppers comprised over 95% of herbivores captured on sticky cards. The total number of herbivores per sticky card averaged 108.2 in the partridge pea buffer, 96.3 in soybean neighboring buffer, and 96.4 in soybean without buffer. Thus, herbivore numbers did not differ significantly in the buffer and soybeans.
Pitfall trap predators. A total of 56,296 arthropods were identified from pitfall trap samples. Of predators captured in pitfall traps, ants, spiders, soldier beetle larvae, rove beetle adults and larvae, and ground beetle adults and larvae were the most abundant. Ant numbers were significantly lower in soybeans neighboring partridge pea on all sampling dates.
Discussion
Year 1 study was conducted to determine if pure and mixed buffer strips of partridge pea and purple tansy could attract greater numbers of beneficial arthropods than non-floral strips of millet, and whether these buffers enhance beneficial arthropod abundance in neighboring soybeans. Purple tansy was not a suitable insectary plant as it was not well adapted to the seasonal period of the study in Maryland. Furthermore, purple tansy would probably be less desirable to establish and maintain as a buffer strip due to its relatively high seed price, slow growth characteristic and greater susceptibility to weed competition. Moreover, purple tansy was quickly out-competed by partridge pea in the mixed planting to the extent that the pure and mixed buffers containing partridge pea attracted similar arthropod communities.
Overall, results consistently showed that partridge pea attracted and supported high populations of natural enemies and potential hosts and prey, with abundances significantly greater than levels found in adjacent soybeans. Sticky card captures of wasp and fly parasitoids in year 1 were more than 70% higher overall in buffers containing partridge pea compared to other buffer types. Similarly, populations of all beneficial arthropods captured by sticky card and pitfall sampling in year 2 were approximately 80 to 72% higher, respectively, in partridge pea buffers compared to the soybean crop.
Parasitoids. Mymarid wasps were notably the most common parasitoids captured on sticky cards and consistently more abundant in partridge pea compared to soybean. These tiny wasps parasitize insect eggs in concealed sites within plant tissues or the soil and are important natural control agents of economically important leafhopper pests. In year 1, mymarids reached levels in partridge pea buffers that were four-fold higher than those in soybean plots, yet significantly lower levels of mymarids were captured in soybean adjoining these buffers. This suggests that the partridge pea lured mymarids from neighboring soybeans. High numbers of mymarids were also captured in partridge pea in year 2 but their abundance in soybeans was not enhanced. This suggests that partridge pea may provide some parasitoids and their associated hosts with all resources required for survival and reproduction. This would in effect provide no incentive for these parasitoids to forage within neighboring crops.
Most fly parasitoids found on sticky cards were tachinids or sarcophagids. The vast majority of hosts of tachinid flies are plant-feeding insects. Their level of parasitism can vary greatly, from less than 1% to approaching 100%, depending on such factors as the size of a host and parasitoid population, and environmental conditions. During both study years, their overall abundance in partridge pea was 62.3% higher than levels in soybean. In year 2, this effect was heightened at the field edge next to buffers, suggesting that higher numbers of parasitic flies encroached into the neighboring soybeans but enter only a short distance within the crop.
Predators. In year 1, predators captured on sticky cards were 65% more abundant in the millet and purple tansy buffers. This response was mainly attributed to the abundance of long-legged flies. These predatory flies hover while searching for small, soft-bodied arthropods, particularly other flies, aphids, spider mites, larvae of small insects and thrips. However, abundances of long-legged flies in soybean plots were not affected by buffer type in year 1. Long-legged flies were also the predominant predators active in the plant canopy in year 2, with overall numbers 2-3 times higher in partridge pea buffers compared to levels found in soybeans. However, their abundance was significantly lower in soybean neighboring partridge pea, particularly at sampling sites closest to the field edge. This is further evidence that the partridge pea acted as a natural enemy sink.
Of the ground-dwelling predators captured by pitfall traps, ants were the predominant group and their abundance was significantly higher in partridge pea than adjoining soybeans. Their numbers were significantly lower in soybean plantings adjacent to partridge pea than grassy check treatment on all sampling dates, implying again that partridge pea acted as a natural enemy sink by luring ants away from soybean. Populations of other ground-dwelling predators, which consisted mainly of spiders, rove beetles, soldier beetles and ground beetles, showed a definite preference for partridge pea compared to soybeans. However, their abundances in the crop were not affected by partridge pea presence.
Herbivores. Sticky card captures each study year indicated that partridge pea harbored significantly more insect herbivores compared to soybean. The majority of herbivores were aphids, leafhoppers, planthoppers and plant bugs. In year 1, number of green cloverworm, as well as other herbivores in soybean were similar regardless of the buffer treatment.
Conclusion
This study demonstrated that partridge pea provides floral resources and alternative food for a diverse community of natural enemies and herbivores. However, its presence as a monoculture buffer did not result in increased number of major natural enemies in neighboring soybeans. Taken together, partridge pea planted as a monoculture acted more as a natural enemy sink by attracting beneficial arthropods away from soybean, potentially decreasing natural control efforts. For this reason, a monoculture of partridge pea may not be an ideal insectary planting if the ultimate goal is to maximize natural enemy efficacy in neighboring soybean fields.
In conservation reserve practices, monocultures of partridge pea are more commonly planted as a wildlife habitat to provide food for bobwhite quail and other wildlife and as flowering habitat for different pollinator taxa. Because the foliage is potentially poisonous to cattle and re-seeding plants can aggressively fill in voids when used as part of a seed mix, conservationists recommend for herbaceous riparian buffers that the total seed mix consist of no less than 1% and no more than 4% partridge pea. However, decisions about the deployment of insectary plants as a monoculture or part of a riparian buffer mix planting should take into consideration the attractiveness and resources provided to natural enemies and their hosts/prey by the insectary habitat in comparison to those provided by the neighboring cash crop. Simple addition of a highly attractive flowering buffer adjacent to a crop could be counterintuitive to natural biological control efforts.
Acknowledgements
Financial support for field studies and publishing results was provided by the Northeast Sustainable Agriculture Research and Education Grants Program, Maryland Soybean Board and USDA NIFA EIPM grant number 2017-70006-27171.
Kelly Nichols, Agriculture Agent Associate University of Maryland Extension, Frederick County
Warmer temperatures and higher humidity are slowly making their appearance. Unfortunately, this means mosquitoes will soon make their appearance as well. Mosquitoes not only are pesky as they fly around you, but are a health risk for humans and livestock, as they carry diseases such as West Nile Virus and Zika Virus. Female mosquitoes need stagnant, nutrient-rich water in order to lay their eggs. After the mosquito eggs hatch, the larvae and pupae will live in the water before becoming an adult. The easiest solution to get rid of mosquitoes is to remove places where this water can accumulate.
Items such as water troughs, buckets, wheelbarrows, and bird baths should be stored inside or upside down when not in use. If stagnant water does accumulate while these items are in use, dump the water and replace it regularly. Loader buckets should be tilted downwards when stored to prevent water accumulation. Get rid of unused tires, and do not pile tires outside. If tires are outside, including those on top of bunk silos, cut them in half to reduce water accumulation. Tarps that cover bales or equipment should be placed so that water can drain. Gutters should be cleaned regularly.
For water sources that can’t be removed, there are insecticides and products that create a film on top of the water. Be sure to read the label before use. Note that the pupae do not actually eat during this stage, and therefore cannot ingest an insecticide, so it is important to use the right product for the correct stage. Creating a film on top of the water’s surface prevents the pupae from being able to access air to survive. Crusts on manure pits can act as a film; however, keep in mind that if using an insecticide, the crust will prevent it from reaching the larvae. Reduce weeds around the pit to deter mosquito habitat. If ponding is occurring in fields and around the barnyard, determine the cause and take the necessary actions. These actions may include alleviating compaction, fixing gutters, or fixing drainage pipes.
Emily Zobel, Agriculture Agent Associate University of Maryland Extension, Dorchester County
Overwintering bean leaf beetles are emerging and starting to feed. Soybean seedlings can recover, with no yield loss, from 40% defoliation. Many small caterpillars, such as the green clover worms will also defoliate plants. However, once soybean plants start to bloom, you want to control defoliating insects when you have greater than 15% defoliation.
Check for cutworm and armyworms leaf feeding on young corn plants. The threshold for cutworms is when 10% of the field has feeding damage at 1-2 leaf, and 5% damage at 3-4 leaf or 4 larvae found per 100 ft. For armyworms, the treatment threshold is when 25% of the plants are infested and larvae are under a 0.75 inch long. Armyworms that are 1.25 inches are late instars and have likely completed their feeding.
No-till fields of both corn and soybean are at an increased risk of slug damage. Slugs feed at night, so you will likely not find them during daytime scouting. Their feeding damage will be found on the lower leaves of plants. The leaf will have narrow, irregular, linear tracks or scars of various lengths that may be eaten partly or entirely through the leaf. Peter Coffey wrote a great article about slug management, which can be found here (https://blog.umd.edu/agronomynews/2018/05/03/what-should-i-do-about-slugs/) .
As wheat gets harvested during the month, stink bugs may move into nearby cornfields. Their feeding could affect the developing ear and kernels. Populations will be highest around the edge of the field, and full-field control may not be needed. Field corn treatment threshold are when 25% of plants are infested with stink bugs before pollination, and 50% of plants are infested with stink bugs are after pollination up to early dough stage. Counts should be done on 10 plants in 10 different locations in the field. Do not count beneficial stinkbugs, such as the spined soldier bug.
Alan Leslie1, Agriculture Agent; Kelly Hamby2, Extension Specialist; and Galen Dively, Professor Emeritus2 1University of Maryland Extension, Charles County 2University of Maryland, Department of Entomology
This time of year, anyone growing small grains will be planning to apply fungicides to manage Fusarium head blight, and many will consider tank-mixing an insecticide to control any insect pest problems at the same time. These tank mixes are an appealing option to reduce the time, fuel, and damage to the crop from having to make a second pass over the field later on in the season. In addition, with many synthetic pyrethroids now available as cheaper generic versions, the costs associated with adding an insecticide to the tank may seem like cheap insurance against possible pest outbreaks. However, to ensure that this added investment gives you a return with increased yields, you should still follow an integrated pest management approach and base the decision to add an insecticide on scouting and documentation of an existing pest problem. Below, we outline several possible insect pests that could be controlled with an insecticide applied with fungicides over small grains, and summarize situations where that application may be warranted, and when it may not.
Aphids. Aphid populations need to be controlled in the fall to reduce Barley Yellow Dwarf Virus incidence in small grains. Spring insecticide applications will not reduce incidence of the disease. Only a few aphid species tend to feed on grain heads, and can reduce yield from head emergence through milk stage (Fig. 1). After the soft dough stage, no economic losses occur. Aphid populations are generally kept in check by insect predators and parasitoids, and thresholds for chemical control of aphids in the spring require at least 25 aphids per grain head (with 90% of heads infested) or 50 per head (50% heads infested) and low numbers of natural enemies. Applying a broad spectrum insecticide when aphid pressure is not above threshold tends to kill off beneficial predatory and parasitic species, which can allow aphid populations to flare up, as they are no longer being suppressed by their natural enemies.
Figure 1. Aphids feeding on wheat head.
True armyworm and grass sawfly. Both true armyworm (Fig. 2) and grass sawfly (Fig. 3) are sporadic pests of small grains and their pest pressure and feeding damage can vary widely from year to year. Automatically applying an insecticide to target these pests is not likely to be a cost-effective strategy since they are not pests that reliably cause economic injury. When these pests are present in high numbers, they are capable of causing significant yield loss through their behavior of clipping grain heads. Scouting should be done to check for the presence of these two pests and insecticide treatment is only needed if they exceed threshold values of one larva per linear foot for armyworm and 0.4 larvae per linear foot for grass sawflies.
Hessian fly. Cultural methods are the best way to control Hessian fly in small grain, such as planting after the fly-free date, selecting resistant varieties, and using crop rotation to disrupt their population growth. Spring feeding by the fly larvae can cause stems to break, reducing yields. There are no effective rescue treatments for Hessian fly; insecticides targeting fly larvae are ineffective since they are well protected from sprays by feeding inside of the leaf sheath (Fig. 4). If this year’s crop is damaged, it is imperative that fly-resistant varieties are planted after the fly-free date next year.
Cereal leaf beetle. This species is widespread in Maryland and is typically present in small grains, though it only occasionally reaches levels that injure crops. Cereal leaf beetle larvae chew the upper surfaces of leaves, leaving them skeletonized (Fig. 5). Larvae can cause yield loss if the flag leaf is severely skeletonized before grain-fill is completed. Insecticides with good residual activity tank mixed and applied with fungicides can potentially control populations of cereal leaf beetles, protect the flag leaf, and improve the yield of the crop if beetle pressure is high. However, predicting whether populations will reach damaging levels is not straightforward, and scouting should be used to guide spray decisions. If a field has 25 or more larvae plus eggs per 100 tillers, and there are more larvae than eggs, then chemical control is needed. In Maryland, a parasitoid wasp species (Anaphes flavipes) may parasitize 70-98% of cereal leaf beetle eggs, so if a field is dominated by eggs with few larvae, insecticide may not be needed. Additionally, feeding by cereal leaf beetle will not cause economic damage after the hard dough stage. So far, we have received no reports of economic levels of cereal leaf beetle in the region.
Figure 5. Cereal leaf beetle larva and feeding damage.
In conclusion, tank mixing an insecticide with your fungicide application can pay off if you have economically damaging levels of an insect pest, but applying any insecticide without a pest problem will not pay off. If populations are present, seem to be increasing, and you will not be harvesting soon, you could gamble. The risks of that gamble include losing money on an unnecessary input cost, secondary pest outbreaks if natural enemy populations are wiped out, or the target pest outbreaks anyway because the application was poorly timed. Scouting fields regularly to document pest pressure and using IPM thresholds as a guide for using chemical controls is the best way to hedge your bets when deciding whether to add an insecticide to the tank this spring.
For more information on tank-mixing insecticides with small grain fungicide applications, check out current research updates from Dr. Dominic Reisig at North Carolina State University: https://smallgrains.ces.ncsu.edu/2019/03/aphids-in-wheat/
Alan Leslie Agriculture Extension Agent, Charles County
After a relatively mild winter, we are getting reports of high numbers of alfalfa weevils causing damage to alfalfa fields in Southern MD. Eggs that were laid by alfalfa weevil adults last fall are all hatching now, and early spring is when the larvae can cause the most damage to alfalfa stands, reducing the yield and quality of the first cutting and potentially setting the entire stand back for the rest of the year. Now is the time to be scouting for larvae and to consider chemical treatment to prevent economic yield loss. Alfalfa weevil larvae look very similar to caterpillars, with a dark head and a green body with a white stripe running down the middle. Early signs of injury from alfalfa weevil larvae appear as pinholes in leaves, but extensive feeding will skeletonize leaves, leaving plants with a distinct gray color. Sweep nets do not do a very good job of catching small larvae, so the best way to scout your fields is to cut stems and beat them on the inside of a bucket to knock the larvae loose. To effectively sample a field, you should collect at least 30 stems from across the field, and beat them vigorously inside the bucket. Make sure to keep track of the exact number of stems you sample, so you can calculate the average number of larvae per stem. Thresholds for control depend on the average height of the stand at the time you sample, with the threshold increasing in taller plants (Table 1).
Table 1.Threshold numbers of larvae per plant for different average stand heights that would warrant chemical control
Stand height (inches)
Larvae per plant
0-11
0.7
12
1.0
13-15
1.5
16
2.0*
17-18
2.5*
*At these stand heights, prompt or early harvest can also control the larval infestation
We have also gotten reports of less than adequate control of alfalfa weevil populations using synthetic pyrethroids (group 3A; beta-cyfluthrin, zeta-cypermethrin, permethrin, lambda-cyhalothrin, cyfluthrin). Because synthetic pyrethroids have historically been such a cheap and reliable chemical class, they have been widely used on many pests, and there are many examples of insect species becoming resistant to this class of insecticide. Currently we do not have good data on the extent to which local populations of alfalfa weevil are becoming resistant to pyrethroids, but it is always a good idea to rotate modes of action of insecticides sprayed over your fields to help prevent resistant populations from developing. Other chemical classes that give effective control of alfalfa weevil include carbamates (group 1A; methomyl, carbaryl) organophosphates (group 1B; phosmet, malathion, chlorpyrifos), and oxadiazines (group 22A; indoxacarb). If you have any remaining chlorpyrifos in your inventory, this may be a good opportunity to use it up, since the state of Maryland will likely introduce legislation to ban its use this year or in the near future. Remember to always read and follow the label whenever making any insecticide applications; the label is the law!
Darsy Smith, Graduate Student & Dr. William Lamp, Professor University of Maryland, Department of Entomology
Figure 1. Yellow appearance of the alfalfa field that lead to the discovery of the cowpea aphid outbreak at BARC. Photo courtesy of Russell Griffith.
An unexpected outbreak of cowpea aphid, Aphid craccivora Koch, in Maryland was discovered last month by Terry Patton, who was contacted by Russell Griffith, tractor operator leader at Beltsville Agricultural Research Center (BARC), because of the yellow appearance of an alfalfa field (Figure 1) and the infestation of dark aphids (Figure 4). Since the 1990s, infestations of the cowpea aphid have been observed in Maryland alfalfa, but this is an unusually large outbreak. Stay alert to this emerging pest and learn how to identify it since it has a wide range of hosts and may damage crops.
Figure 2. Adult cowpea aphid. Note the cornicle (yellow arrow) is dark and long and the abdomen (red arrow) is distinctive dark and shiny. Photo courtesy of influentialpoints.comFigure 3. Adult and nymph cowpea aphids. Nymph color is opaque and varies (yellow arrow) from brown to gray while the adult (red arrow) has the distinctive dark, shiny abdomen. Photo courtesy of Andrew Jensen, https://aphidtrek.org/
Cowpea aphid identification and injury. Cowpea aphid is not generally an economic pest in alfalfa but learning how to identify the aphid and its injury can help you prevent losses. Cowpea aphid is easily differentiated from other aphids in alfalfa because its dark coloration, with the abdomen of the adults much darker and shinier than the rest of its body (Figure 2). In addition, the cornicle (or siphuncule) is dark and long (Figure 2). The nymph is less shiny (opaque) and varies from brown to gray (Figure 3). The legs and antennae of both adults and nymphs are pale with dark tips (this characteristic is more distinctive in adults).
The cowpea aphid is a sap-sucking feeder and damage caused in alfalfa by this pest results from the injection of a toxin into the phloem of the plant. With high population densities on plants, the aphid can cause stunting or plant death. In addition, it can cause yellowing in alfalfa leaves (Figure 1). Like other aphids, this insect produces honeydew that will benefit fungus growth and eventually cause sooty mold.
How to find them? Cowpea aphids are usually found in clusters on the alfalfa leaf and stems (Figure 4). It can also be found in vegetative growth and flower parts of a wide range of hosts. They are readily sampled with sweep nets.
Figure 4. Cowpea aphids on growing tip of alfalfa at BARC. Photo courtesy of Russell Griffith.
Host plants. Cowpea aphid is most commonly found in alfalfa, but may be found on other legumes, such as clovers. More uncommonly, the aphids occur on a variety of weeds and other plants in other plant families.
Management options. Unfortunately, there is not an economic threshold specified for this pest in Maryland alfalfa at this point. However, here are general guidelines for responding to the pest:
Conserve natural enemies. Natural enemies such as lady beetles, damsel bugs, and parasitoid wasps often locate, feed, and reproduce in conjunction with high densities of aphid in alfalfa. If you conserve natural enemies you might find aphids parasitized by parasitoid wasps (Figure 5). When scouting for aphids, watch for natural enemies to help control aphid populations. To help natural enemies stay in your alfalfa field you can use border-strip cutting while harvesting to provide refuge habitats. For more information of this practice, see “Harvest Scheduling and Harvest Impacts on IPM” at the end of this report.
Monitoring for decision-making. Early infestations in alfalfa can result from migration from southern areas. Pay attention to alfalfa fields in March and continue to monitor until fall. Since there are no thresholds developed for cowpea aphid, the thresholds for insecticide applications developed for the blue alfalfa aphid can be used: if alfalfa is 10 inches, then treat if there are 20 or more aphids per stem; if alfalfa is 20 inches tall, then treat if 50 or more aphids per stem.
Figure 5. Aphids parasitized or aphid mummies by a parasitoid wasp. Note the emergence hole (yellow arrows) of the parasitoid wasp in the mummy. Photo courtesy of Darsy Smith.
Potential reasons for the outbreak.
Researchers in the Lamp Lab, University of Maryland, have noted this pest in greenhouse settings but rarely observed them in alfalfa fields. Dr. Lamp suggests that the cowpea aphid may have migrated into Maryland because this species is more common in southern and western areas. Also, the mild winter may have allowed individuals that migrated last year to overwinter in Maryland. Additionally, lack of natural enemies in early spring can potentially lead to an outbreak. Conserving the natural enemies in alfalfa fields and neighboring areas can help decrease aphid abundance.
How can you help?
Records from your alfalfa fields and surrounding crops are valuable sources of information. The information is helpful to not just explain an outbreak but also to provide useful guidelines for farmers to manage the crop and avoid a future outbreak. Practical information that you can provide include any of the following:
Date of first time you have observed the pest
Date of outbreak
Plant height and phenology stage: when you observed the pest for the first time and during outbreak
Presence of natural enemies
Pesticide use and efficacy of application
Alfalfa cultivars/varieties planted during outbreak and previous year
Pictures of damage and estimate of loss
If you find cowpea aphid in your alfalfa field, please contact the nearest county extension office.
