Tag: management

Preparing for Tar Spot of Corn in 2023

Andrew Kness, Senior Agriculture Agent | akness@umd.edu
University of Maryland Extension, Harford County

Figure 1. Signs and symptoms of tar spot on corn. Black raised areas are tar spot and long rectangular grey lesions are from grey leaf spot.

Tar spot is a foliar disease of corn caused by the fungus Phyllachora maydis and we confirmed it for the first time in Maryland from a grower field in Harford County in 2022; however, it is likely that it has been present in fields at low levels earlier than the 2022 growing season. Weather conditions across northern Maryland and Southern PA in August and September were favorable for tar spot development and pockets of disease outbreaks were reported, leading to much discussion about the disease amongst farmers and ag service providers over the 2022-2023 winter months about what to do to manage this disease in the future.

The pathogen that causes tar spot is favored by cool, wet weather. Tar spot spores overwinter in old corn crop residue and it seems to survive our winters just fine, as demonstrated by winter survival in Pennsylvania, as well as many northern corn belt states.

Temperatures between 60-70°F, coupled with 7+ hours of leaf wetness from dew, humidity, rain, or irrigation, trigger sporulation and subsequent spore germination on susceptible corn plants. Roughly 14-21 days later, signs and symptoms of tar spot will develop on corn plants in the form of small, raised black spots that have the appearance of tar or splattered black paint (Figure 1). These spots are the reproductive structures which provide secondary inoculum that repeatedly infect more tissue for as long as temperature and moisture conditions remain conducive.

In the Midwest where tar spot has been present since 2015, yield losses have been reported upwards of 60 bushels in bad years. It is also important to note that tar spot can make corn plants senesce and dry down much faster than normal, going from green to brown in 10-14 days under optimum conditions. This can make silage harvest tricky, which is why scouting is so important.

We do not know how prevalent and severe this disease will become in Maryland, so I encourage farmers to diligently scout corn fields to get ahead of it and also to determine where the disease is distributed. Scouting will also help you determine if a fungicide application is warranted. Fortunately, most fungicides that are labeled for corn do a fairly good job of protecting against tar spot, but there is data that suggests that the two and three way mode of action (MOA) products work better than single MOA products.

Fungicides should be applied as close to disease onset as possible; for tar spot this can be tricky because it can infect corn at any growth stage and it can still have significant yield impact as late into the season as R4. University research in the corn belt has found that the best chance for an economic return on investment is a single application around VT-R1; however, there are some instances where a second application was necessary but these were only when weather conditions for tar spot remained favorable during these later reproductive growth stages.

A few things to consider for tar spot management as we go into the 2023 growing season are as follows:

  1. Avoid highly susceptible hybrids, especially in corn-on-corn fields or if you have a field with history of tar spot. There is no complete resistance to tar spot in commercial corn hybrids, but we do know there is some variability in susceptibility. Work with your seed dealers to try to identify your best hybrids and plant them in these fields where you think tar spot may be a problem.
  2. Tillage and residue management appears to play a minor role in the management of this disease. Tillage may slightly reduce primary inoculum, but we need to keep in mind that tar spot spores can blow in from neighboring fields; so, I would not roll out the heavy tillage and blow up your no-till system just to try managing tar spot because it will only have a marginal effect.
  3. Corn-on-corn has a higher risk for developing tar spot, especially if the previous corn crop was infected. Rotate with other crops to break up this cycle. P. maydis only infects corn (including dent, sweet, and popcorn); all other crops are not hosts.
  4. Hybrid maturity also plays a role in disease severity. Research from the Midwest has shown that longer maturing hybrids suffer greater yield loss than shorter maturing hybrids. This is because the longer you push the grain fill period into the cooler late summer/fall months, the more likely tar spot will infect during earlier grain fill growth stages.
  5. Scout fields this year starting a little before tasseling through to maturity. As mentioned above, this will help you determine if a fungicide is likely to pay off or not.

We will be conducting some field trials this year looking at tar spot management in Maryland with funds from the Maryland Grain Producer’s Utilization Board. Part of this project will also include a survey of corn across the state to determine the distribution of tar spot. If you think you find tar spot in a field this year, I would be interested in knowing about it. You can call or email me (410-638-3255, akness@umd.edu), or report a sighting at corn.ipmpipe.org.

Plant parasitic nematode genera in Maryland and Delaware Soybean fields and strategies for their management

Ramesh Pokharel1, Alyssa Koehler2, Sarah Hirsh3, Jim Lewis3, & Nidhi Rawat4
1USDA APHIS, Riverdale, MD
2Department of Plant and Soil Sciences, University of Delaware
3University of Maryland Extension
4Department of Plant Science and Landscape Architecture, University of Maryland, College Park

Small-scale soil surveys were conducted in the major soybean growing counties of Maryland and Delaware in summer 2019 with funding support from Maryland Soybean Board. The following eight genera of parasitic nematodes were observed in varying densities: root-knot (Meloidogyne), Cysts (Heterodera), root lesion (Pratylenchus), dagger (Xiphinema), stunt (Tylenchorhynchus), stubby root (Trichodorus), spiral (Helicotylenchus), lance (Hoplolaimus), and needle nematode (Longidorus). Out of these eight, soybean cyst, root-knot, and root lesion nematodes are the most economically significant to soybean production. It is important to note that nematode management in fields requires long-term application of management tactics, and should be adopted is the densities of problematic nematodes exceeds their Economic threshold levels. Economic Threshold Level (ETL) is defined as the density of nematodes at which the damage caused by a particular nematode exceeds the cost of managing it. Biology of major parasitic nematodes of soybean found in the mini-survey, as well as the management approaches recommended for them are described in this article.

Soybean Cyst Nematode (SCN)

Soybean cyst nematode (SCN) was first detected in Worcester County in Maryland in 1980 (Sardanelli et al., 1982) and subsequently spread to most eastern shore counties of MD and to DE.  In the present survey 41 and 57, samples from Maryland and Delaware, respectively were found positive for SCN, where 50 and 41 % of fields had SNC densities higher than ETL (60 individuals per 500 cc soil). This survey indicated that SCN remains an important problem in the eastern shore counties of MD and in DE.

Biology: Juvenile larvae (J2), the most infective stage, emerge from eggs and enter plant roots establishing feeding sites. Males become vermiform (worm-shaped) and exit the roots, whereas lemon-shaped young females slightly less than 1 mm in diameter, whitish-yellow in color appear on the roots. The adult females break through the root surface but remain attached to the root. Mating takes place on the root surface and females lay 50-100 eggs in an egg mass on the back of their body. The female then fills will more than 200 eggs and dies. The body walls harden to form a tough cyst around the eggs, which can remain viable in the soil for long periods, even in adverse conditions. The vermiform (worm-shaped) juvenile and adult males as well as the brown cysts (dead female body containing eggs), and freed eggs can be extracted from the soil. Often, cysts may get confused with nitrogen fixing bacteria Bradyrhizobium root nodules, which are pinkish in color and smaller than cysts and can easily be squeezed in between fingers producing pinkish fluid. Under favorable environmental conditions, SCN can complete a life cycle in four weeks, making it possible to have multiple life cycles over the course of a season.

Symptoms: SCN infected fields often lack obvious above ground symptoms, making SCN infestation difficult to predict unless soil samples are taken for nematode assessment. If symptoms are present, they may be confused with other diseases or abiotic stress. High SCN populations may cause yellowing or stunting of above ground plant parts and/or root distortion, dwarfing, stunting, reduction in nitrogen-fixing nodules, and increased susceptibility to other soil-borne plant pathogens.

Management strategies: SCN is often called the silent yield robber because it can be difficult to know that  populations are building. Since SCN can survive in the soil for many years even in adverse conditions, it will take consistent long-term efforts to reduce SCN problem in a field. Currently, ‘PI 88788’, ‘Peking’, and ‘PI 437654’ ‘Hartwig’ are the available sources of resistance. ‘PI 88788’ has been the primary source of resistance for decades and is found in over 95% of soybeans currently available. Long-term exposure to this resistance gene has selected for SCN populations that are able to reproduce at higher rates and overcome the ‘PI 88788’ source of resistance. Rotating resistance from ‘PI88788’ to ‘Peking’ has been reported to minimize losses, while plots with the continuous use of ‘PI 88788’ or ‘Peking’ had 5% and 8% lower yields, respectively. Race determination is important to decide the efficacy of resistant sources or cultivars. Soybean maturity group also affects SCN reproduction.

Late maturing groups remain in the field until late October or November, allowing an extra generation of SCN to develop. Selecting soybeans with an earlier maturity group can help reduce SCN population numbers the following year. Soybeans rotated with non-host crops such as corn, small grains, and alfalfa help to reduce the nematode build up in a field. All types of beans, lespedeza and hairy vetch, some ornamental plants and weeds such as henbit, purple deadnettle and common mullein can help maintain SCN populations and should be avoided. One year in a weed-free non-host crop can reduce SCN population up to 55%.

Since some of the eggs may remain unhatched in the cyst for years, it is impossible to completely destroy SCN populations by starvation. Seed treatments generally help to protect young roots from nematode infection, thereby reducing crop yield losses, but this will not reduce the SCN population in a field. Aveo and ILEVO are some of the products that are known to reduce nematode damage to the crop.

Root-knot nematode (RKN)

Root-knot nematode (RKN) may cause significant (more than 70%) yield loss, when a susceptible variety is planted in field with a high population density of root-knot nematodes. In less fertile soil, soybean and corn can be severely damaged, especially when they are mono-cultured for several years. Damage to these crops is more serious and wide-spread in years when soils where temperatures are unusually warm during the early part of the growing season. Several species of Meloidogyne are reported to infect soybean; M. incognita (southern root knot), M. enterlobii (guava root knot), M. javanica (Javanese root knot), M. hapla (northern root knot), and M. arenaria (peanut root knot). Of which, Meloidogyne incognita, is the most common root-knot nematode species found in Maryland and Delaware (Everts et al., 2006). In the present mini-survey of Maryland and Delaware, 58% and 17% of samples, respectively had RKN. Three percent of the fields tested in Maryland had RKN densities higher than ETL of 170 nematodes per 500 cc soil.

Biology: The life cycle of root knot nematodes (eggs, larvae and adults) is completed within 14-25 days depending upon several factors, especially soil temperature. Under favorable conditions, juveniles hatch immediately from eggs, otherwise eggs remain dormant and can survive in soil for several years until favorable conditions for hatching are available.

Symptoms: Plants infected with RKN exhibit symptoms varying from asymptomatic to non-uniform stunting, wilting, and chlorotic patches, mostly caused by high populations in the soil. When dug up, plants infected by RKN will exhibit root galling symptoms. Root galls caused by RKN may get confused with root nodules produced by nitrogen fixing bacteria, Bradyrhizobium. Root nodules are generally round and can easily be squeezed producing pink to reddish milky substances whereas nematode induced root galls vary in shape and size,are hard to squeeze, and do not produce milky substance when crushing.

Management strategies: Knowing where RKN is distributed in a field is an important first step to management. Nematodes can easily be carried field to field by equipment and vehicles, so sanitizing equipment between affected fields can minimize the change of moving RKN to new fields. Like many nematode genera, RKN is often worse in sandy-to-sandy loam soils.  Due to a wide host range, crop rotation is not always an effective management strategy for root-knot nematode. Rotation with poor host crops can help to decrease populations. Broccoli, cauliflower, grain sorghum or millet (for bird feed) can lower root-knot numbers, particularly if they are grown for two consecutive years. Rye, grown as a winter cover crop, may also help lower nematode populations. Delaying soybean planting until mid-June, as done in double-crop wheat/soybean systems will reduce the number of nematode reproductive generations.

With the phase out of many chemical nematicides, chemical control is not cost-effective for RKN currently. The most cost-effective management strategy is to select a RKN resistant variety.

Root Lesion Nematode (RLN)

Root-lesion nematode (RLN) is one of the most common nematodes in Maryland and Delaware in soybean crops. Jenkins et al. (1956) reported higher numbers of this genus in soybean samples than other crops. Later, Pratylenchus spp. were found in 78% of 362 samples in eight counties across MD and DE with P. penetrans being the most common species (Sindermann et al., 1993). In the present mini-survey, genus Pratylenchus was observed in 58% and 60% of the field samples in MD and DE, respectively. Densities higher than the ETL (>500 nematodes per 500 cc) soil were found in 5% of the fields.

Biology: Sexual reproduction is common in P. penetrans, but not in all species. Males are rare or absent in many species such as P. hexincisus, P. neglectus, and P. scribneri. The females lay eggs (after/ without mating) singly or in small groups in the host root or in the soil near the root surface. The first larval stage and molt occur within the egg. The second-stage larva emerges from the egg and undergoes three more molts before becoming an adult. The egg hatches within 1 to 3 weeks, depending on the soil temperature.

Symptoms: RLN produces characteristic necrotic lesions (darkened areas of dead tissue) on infected roots, which turn reddish-brown to black later. Nematode migration and feeding within the roots cause coalescence  of several small spots  into large necrotic areas that may eventually girdle the root. High nematode populations can cause stunting and necrosis of the root system. The extent of lesion formation can be accelerated by root infection by other soilborne plant pathogens, which may produce synergistic disease complexes.

Management strategies: Due to the wide host range, crop rotation is generally not effective for the management of RLN especially in high density populations. RLN can cause extensive yield loss in more than 160 host plants that includes both grasses and broadleaf plants including crops and weeds. Grain crops such as corn, wheat, and alfalfa are favorite hosts. If the nematode population is low, cover crops such as ryegrass or canola used as green manures may be useful to decrease population build up if successively incorporated into rotational cropping sequences (Everts et al., 2006). Limited information is available on soybean cultivars with  RLN resistance.

Dagger nematode (Xiphinema)

Dagger nematode is not an important problem in soybean in Maryland and Delaware at present because of lower densities and incidence as compared to other nematodes. The damage due to dagger The ETL of dagger nematode is population densities above 250 per 100 cc soil. Dagger nematode has a very wide host range and many grass species appear to be good hosts of dagger nematode. Species of Xiphinema are sensitive to changes in soil temperature and moisture and will migrate away from desiccating conditions in topsoil; most dagger nematodes can live and survive deep in the soil. Dagger nematodes feed on the outside of roots and root cells eventually collapse due to feeding. These nematodes are important in crop production because they vector some important plant viruses. Soybean severe stunt virus (SSSV) transmitted by this nematode has been reported in some fields in MD and DE.

Lance nematodes (Hoplolaimus spp.)

Lance nematodes feed on plant roots as ectoparasites (body remaining outside) or semi-endoparasites (body within roots). Previously this genus was observed in 23% of samples in 1956, and 38% in 1978 (Golden and Rebois,1978). Additionally, lance nematode were found in 43% of 199 soybean fields in eight counties of Maryland in 1993 (Sindermann et al., 1993). In the present mini-survey, Lance nematodes were found in 8% samples with only 2% fields having levels higher than ETL. Higher percentages of tested fields (about 25%) were positive for lance nematodes in Delaware, but only 2% of fields were above ETL. Lance nematodes have been associated with alfalfa, barley, carnation, clover, corn, grass, lespedeza, oat, pea, pepper, rye, soybean, sweet potato, timothy, tobacco, tomato, and wheat. No difference in the preference of soil type of this nematode was found and Hoplolaimus spp. were considered less common than other plant parasitic nematodes of the region.

Stubby root nematodes (Trichodorus and Paratrichodorus spp.)

Stubby root nematodes are difficult to manage in corn/soybean rotations, as both crops are susceptible.  Optimal growing condition, especially moisture and fertilizer, will help to support plant growth. Nematicide seed treatments are not effective if high populations of the nematodes are present. Stubby root nematodes may live below the depth that fumigants would be placed, so fumigant treatments are not recommended.

Concluding remarks: Mini-surveys of soybean fields conducted in summer 2019 in Maryland and Delaware soybean growing counties identified Soybean Cyst, Root knot, and Root lesion nematodes as predominant parasitic nematodes in the two states. Dagger and lance nematodes were found to be other potentially damaging nematode genera in variable densities, but not in all samples. Other minor nematode genera observed included Stunt nematode, Needle nematode, and Spiral nematode, with low densities and incidences.  By the time aerial symptoms become apparent, high densities of nematodes have already developed in the field. It is advisable to keep an annual track of problematic nematodes by soil sampling and scouting for root symptoms. Consistent long-term management strategies need to be applied to keep the nematode problem successfully and sustainably in control. Table 1 below summarizes the economic threshold levels and management strategies recommended for managing major problematic soybean nematode genera found in the 2019 mini-survey. It is important to note that the need to consider application of major management strategies should be based on population densities of the parasitic nematodes in your field. If the nematode population density in a field exceeds the ETL, suggested management strategies should be used. For making this decision, it is important to measure the density of particular nematodes in your fields.  The following article, “Sampling for plant parasitic nematodes” provides details about how to correctly collect representative soil samples for nematode determination in crop fields.

Table 1. Economic Thresholds and Management Strategies for Parasitic nematodes found in soybean fields in Maryland and Delaware during a mini-survey conducted in summer 2019.

Nematode ETL* per 500 cc soil Available management strategies
Soybean Cyst Nematode 60 1. Rotate soybeans with non-host crops like corn, small grains, and alfalfa.

2. Avoid lespedeza, hairy vetch, and beans which maintain SCN population.

3. Avoid weeds such as henbit, purple deadnettle, and common mullein.

4. Grow early maturing cultivars if your field has high population level of SCN.

5. Manage and rotate resistant cultivars as majority of ‘PI 88788’ resistant cultivars are becoming susceptible to SCN.

6. If severe nematode populations persist, apply seed treatments such as Aveo and ILEVO.
Root Lesion Nematode >500 1. Grow non-host crop such as ryegrass or canola.

2. Grow relatively tolerant soybean cultivar such as ‘Essex’ and avoid sensitive cultivars such as ‘Forrest’.

3. Delay soybean planting until mid-June, as done with double-crop wheat/soybean systems.

4. Avoid planting other suitable host crops such as tobacco and peanuts.
Root Knot Nematode 170 1. Plant resistant soybean cultivars.

2. Use cover crops ryegrass or canola and incorporate as green manures.

3. Rotate soybean with non-host crops such as Broccoli, cauliflower, grain sorghum or millet (for bird feed) and winter crops such as Rye.

4. If severe RKN persist, apply seed treatment.
Lance Nematode 250 1. Grow moderately tolerant cultivars such as ‘Northrup King’, ‘S83-30’, ‘Coker 368, ‘Centennial, ‘Hagood’ and ‘Maxcy’.

2. Grow small grains if nematode numbers are moderate-to-high.

3. Grow castor bean, rape seeds and sorghum sundangrass as green manure crops and incorporate after three months.

4. If high nematode population exists for several years apply chemical seed treatment.
Dagger Nematode 250 1. Practice crop rotation with corn or grain sorghum, wheat followed by ‘HT-5203’ soybean, or 2-year fallow
Stubby root Nematode >100 1. Grow moderately tolerant cultivars.

2. Avoid host crops such as corn, cotton, and soybean.

References:

Everts, K. L., Sardanelli, S., Kratochvil, R. J., Armentrout, D. K., & Gallagher, L. E. (2006). Root knot and root lesion nematode suppression by cover crops, poultry litter, and poultry litter compost. Plant Disease, 90, 487–492.

Golden, A. M., & Rebois, R. V. (1978). Nematodes on soybean in Maryland. Plant Disease Report, 62, 430–432.

Jenkins, W. R., Taylor, D. P., & Rohde, R. A. (1956). A preliminary report of nematodes found on corn, tobacco, and soybean in Maryland. Plant Disease Report, 40, 37–38.

Sardanelli, S. L., Krusberg, L. R., Kantzes, J. G., & Hutzell, P. A. (1982). Soybean cyst nematode, fact sheet 340. College Park: Maryland Cooperative Extension Service.

Sindermann, A., Williams, G., Sardanelli, S., & Krusberg, L. R. (1993). Survey for Heterodera glycines in Maryland. Journal of Nematology, 25, 887–889.

 

Scouting for Soybean Cyst Nematode

Alyssa Koehler, Extension Field Crops Pathologist
University of Delaware

soybeans showing no visible symptoms of SCN
Figure 1. Soybeans with healthy looking foliage, but high levels of SCN in the soil.

Soybean Cyst Nematode (SCN) consistently ranks as the most yield limiting pathogen of soybeans across the US, with average annual yield losses estimated over $1 billion dollars. SCN and other nematodes are often silent yield robbers, being present in the field without noticeable aboveground symptoms. If symptoms from SCN do occur, they can look similar to other production challenges like nutrient deficiency, soil compaction, drought stress, or other diseases. SCN can inhibit Rhizobium nodule formation, causing chlorosis or yellowing of soybeans in affected areas of the field. Due to the lack of consistent or obvious aboveground symptoms, it is very common for SCN to go unknown until severe infestation develops (Figure 1). Scouting soybean roots for SCN females in season and conducting fall soil samples are two ways to check your field for SCN. Yellow to white females can be found on roots from about six weeks after planting through the end of the season. While females on the roots confirm the presence of SCN, they do not provide information on the level of infestation. Soil samples are the best method to assess overall populations across the field. Soil sampling can be conducted at any time, but fall samples provide a good snapshot of end of season populations and can be collected when already out for routine fertility sampling. We will discuss the steps to collect soil samples for SCN in an August article. Today I will introduce the steps to scout for SCN females on roots:

When to sample: Scouting for SCN females on roots can occur 6 weeks after planting up until 3-4 weeks before harvest. Digging plants earlier in the season is generally more effective because new roots surrounding the base of the plant are easier to dig and not as far down into the soil profile.

Where to sample: When scouting a field that has never been checked for SCN, you can target any areas with yellowing or stunting, but it is also a good idea to include healthy looking plants since SCN can be present without any aboveground symptoms. Areas of the field that tend to be higher risk for SCN include: near a field entrance, areas that have been flooded, areas with pH greater than 7, areas where yield has historically been lower, areas where weed control is not as good.

SCN females on a soybean root
Figure 2. Soybean root system with SCN females indicated at arrows.

How to sample: Using a shovel, dig 6 to 8 inches from the base of the plant to try to remove as much of the root system as possible. (Avoid tugging or pulling on the plant since you will leave much of the root system behind in the soil.) Gently shake off the soil and check the root system for white to light-yellow lemon-shaped adult SCN females (Figure 2). SCN females are much smaller than the nitrogen-fixing nodules (Figure 3). A hand lens or magnifying glass can make looking for SCN females easier, especially when scouting in sandy soils where sand particles can resemble SCN females. Gently swirling roots in a bucket of water can help to remove soil particles without dislodging the females.

Comparison of larger nitrogen fixation nodules next to smaller SCN females on a soybean root
Figure 3. Soybean root system with nodulation (left arrow) and SCN females (right arrow).

What to do next: If you find SCN females or suspect nematodes are present in the field, a soil test is the next step to estimate population density in the field. For many years, nematode populations were managed through a single source of resistance, PI88788. Over the past few decades, we have seen a break down in this resistance and nematodes are reproducing at far higher rates. When a resistant variety is providing effective control, there should only be 10 to 20 SCN females on the roots. When digging some of our SCN trial plots this week we had plants with 150+ SCN females. If high levels of SCN are present, rotation of crop and variety are the best steps to reduce populations. Corn and wheat are both non-host options. While the PI88788 resistance gene still accounts for over 95% of soybean acreage, there are new resistance genes coming out on the market. Seed treatments are another control option. We are currently screening multiple seed treatment products for efficacy in our region and will post those results as they become available later this year.

 

Managing Fusarium Head Blight

Dr. Alyssa Koehler, Extension Field Crops Pathologist
University of Delaware

With the mild winter, wheat and barley are moving right along. Planting behind corn is common in our region, but this maintains inoculum for Fusarium Head Blight (FHB). Fusarium species that cause FHB can infect both corn and small grains. Walking through fields with corn stubble, you may see orange growth on old debris (Figure 1). Wet spring conditions favor fungal sporulation that can lead to infected wheat heads. As the pathogen grows on debris, spores are released that can be rain dispersed or moved through air currents. As the grain is flowering, spores land on the head or anthers, colonize these tissues, and move into the grain head. Once inside the grain, water and nutrient movement is disrupted, which results in the bleached florets we associate with FHB (Figure 2). Shriveled and wilted “tombstone” kernels can reduce yield and result in grain contaminated with mycotoxins. Deoxynivalenol (DON), also referred to as vomitoxin, is a health hazard to humans and animals. Wheat heads colonized later in development may not show dramatic symptoms, but can still have elevated DON.

Figure 1 (left). Corn stubble with Fusarium sporulation that can contribute to FHB in wheat. Figure 2 (right). Wheat head showing bleached florets from Fusarium Head Blight.

As we approach heading and begin to think about in-season disease management strategies, a well-timed fungicide application can help to reduce disease severity and DON levels. It is important to remember that fungicides can help to reduce disease levels and DON (traditionally around 50% reduction on a susceptible variety), but they do not eliminate FHB or DON. To try to maximize the efficacy of fungicides, it is important to apply at the correct timing. Fungicides for FHB are most effective when applied during flowering in wheat and at head emergence in barley. The Fusarium Risk Assessment Tool (www.wheatscab.psu.edu) is a forecasting model that uses current and predicted weather forecasts to predict FHB risk. The model is currently being configured for this season and should be accessible at the link above by the end of the first week of April. Historically about 70% accurate, this tool aids in assessing FHB risk as wheat approaches flowering and fungicide application decisions are made. The pathogen that causes FHB infects through the flower and rainfall 7 to 10 days prior to flower favors spore production and increases risk of infection. Optimal wheat fungicide application is at early flowering (10.5.1) to about 5 days after. Although new products like Miravis Ace can be applied earlier, it is still best to wait for main tillers to be at 10.5.1 or a few days beyond so that secondary tillers have a greater chance of being at 10.3-10.5.1. If you spray too early, heads that have not emerged will not be protected by the fungicide application. When wheat heads begin to flower, look for yellow anthers in the middle of the wheat head. When at least 50% of main stems are flowering, you will want to initiate fungicide applications. As the flowering period continues, anthers will emerge from the top and then the bottom of the wheat heads. Anthers can stay attached after flowering but usually become a pale white (Figure 3, next page). Triazole (FRAC group 3) fungicides that are effective on FHB include Caramba (metconazole), Proline (prothioconazole), and Prosaro (prothioconazole + tebuconazole). Miravis Ace (propiconazole + pydiflumetofen) offers a triazole + SDHI, FRAC group 7. As a reminder, fungicides containing strobilurins (QoI’s, FRAC 11) should not be used past heading because these fungicides can result in elevated levels of DON. Flat fan nozzles pointed 90° down are great at covering foliage but they do not provide good coverage on heads, which is the target for FHB management. Nozzles that are angled forward 30-45° down from horizontal (30 degrees is better than 45) or dual nozzles angled both forward and backward give better contact with the head and increase fungicide efficacy. For ground sprays, fungicides should be applied in at least 10 gallons of water per acre.

Figure 3. From left to right: Feekes 10.3, Anthesis; Feekes 10.5.1 (yellow anthers beginning flowering); 4 days after anthesis (white anthers post flowering). Image: A. Koehler, Univ. of Delaware.

Thinking beyond this season, an integrated approach can improve management of FHB and help to keep DON levels low. In your field rotation plan, avoiding planting small grains into corn residue will help to reduce the amount of initial inoculum in your field. If you have soybean fields that can be harvested early enough for a timely wheat planting, this rotation helps to break up Fusarium inoculum. In addition to rotation considerations, seed selection is another important piece of FHB management in wheat. There is no complete host resistance against FHB, but you can select wheat varieties with partial resistance. The University of Maryland sets up a misted nursery to compare FHB index and DON levels across local wheat varieties to aid in variety selection decisions. Results from 2019 can be found at https://scabusa.org/pdfs/UMD_Misted-Nursery_Factsheet-2019.pdf. Remember that these trials are conducted under extreme disease pressure and you want to look at relative DON performance. Unfortunately, barley does not have any resistance to FHB. In UMD’s 2019 trial, Calypso had the lowest DON content in local barley varieties tested.

 

Wheat Variety Selections—An Important Factor For Managing Head Blight

Andrew Kness, Agriculture Agent
University of Maryland Extension, Harford County

Compared to the 2018 wheat crop, 2019 was a much better year for Fusarium head blight (FHB, also known as head scab). Growing quality wheat in Maryland starts with proper variety selection. As you look ahead to the 2020 wheat crop, select wheat varieties that have good FHB ratings. There are no varieties with complete resistance to head scab; only varying degrees of susceptibility. Nevertheless, planting a somewhat resistant variety will go a long way in managing FHB and keeping vomitoxin levels (DON) lower in your grain compared to a susceptible variety.

To aid in your selection of wheat varieties, the University of Maryland screens several wheat varieties for their resistance to Fusarium graminearim, the causal agent of FHB. The results from the 2019 trials can be found here.

Additional considerations for FHB management include:

  • Planting behind soybeans rather than corn or other small grains. The FHB pathogen survives on residue of corn, wheat, barley, oats, and other grasses; however, it does not persist on soybean residue.
  • If planting into corn residue, consider tillage if it is an option for your farm. Sizing and burying corn residue will accelerate its decomposition and reduce the FHB pathogen survival.
  • Fungicides in spring 2020. Please note that fall fungicide applications do not have any effect on managing FHB. More information will be covered concerning fungicide recommendations in the spring, or read this article from earlier this year.

 

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 State6 and 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).

wireworm
Figure 3. Close up image of wireworm showing the “keyhole-shaped notch” that occurs at the rear.

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 insecticide3, 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/

 

Managing Fusarium Head Blight

Alyssa Koehler, Extension Plant Pathologist
University of Delaware

Andrew Kness, Agriculture Agent
University of Maryland Extension, Harford County

When it comes to controlling Fusarium Head Blight (FHB) and keeping deoxynivalenol (DON) levels low, it is important to have an integrated approach. Considering the disease cycle of FHB (Figure 1), the FHB pathogen (Fusarium graminearum and other Fusarium sp.) is able to grow on crop residues from corn and small grains. In your field rotation plan, try to avoid planting wheat or barley into corn residue; this will help to reduce the amount of initial inoculum in your field. As the pathogen grows on debris, it eventually releases spores that can be rain dispersed or moved through air currents. While the grain is flowering, spores land on the head or anthers, colonize these tissues, and move into the grain head. Once inside the grain, water and nutrient movement is disrupted which results in the bleached florets we associate with FHB (Figure 2). Shriveled and wilted “tombstone” kernels can reduce yield and result in grain contaminated with mycotoxins. DON, also referred to as vomitoxin, is a health hazard to humans and animals. Wheat heads colonized later in development may not show dramatic symptoms, but can have elevated DON.

Figure 1. Fusarium Head Blight Disease Cycle. For more information on the FHB disease cycle visit https://www.apsnet.org/edcenter/disandpath/fungalasco/pdlessons/Pages/Fusarium.aspx Image: apsnet.org.

In addition to rotation considerations, seed selection is another important piece of FHB management in wheat. There is no complete host resistance against FHB, but you can select wheat varieties with partial resistance. The University of Maryland sets up a misted nursery to compare FHB index and DON levels across local wheat varieties to aid in variety selection decisions https://scabusa.org/pdfs/UMD-UDE_Misted-Nursery_Factsheet-2018.pdf. Unfortunately, barley does not have any resistance to FHB. At this point in the season, rotation order and variety are established, but you can consider these factors as you plan for next season.

Figure 2. Wheat head with Fusarium head blight. Image: Andrew Kness, University of Maryland.

As we think about 2019 in-season disease management strategies, a well-timed fungicide application can help to reduce disease severity and DON levels. It is important to remember that fungicides can help to reduce disease levels and DON (traditionally around 50% reduction on a susceptible variety) but they do not eliminate FHB or DON. To try to maximize the efficacy of fungicides, it is important to apply at the correct timing. Fungicides for FHB are most effective when applied during flowering in wheat and at head emergence in barley. As wheat approaches heading, the Fusarium Risk Assessment Tool (www.wheatscab.psu.edu) is a forecasting model that uses current and predicted weather forecasts to predict FHB risk. This tool is historically about 70% accurate, and can help you assess your risk for developing FHB as your wheat approaches flowering. The pathogen that causes FHB infects through the flower, and rainfall 7 to 10 days prior to flowering increases spore production and risk of infection. Optimal wheat fungicide application is at early flowering (10.5.1) to about 5 days after. For initial signs of wheat heads beginning to flower, look for yellow anthers in the middle of the wheat head. When at least 50% of main stems are flowering, you will want to initiate fungicide applications. As the flowering period continues, anthers will emerge from the top and then the bottom of the wheat heads (Figure 3). Anthers can stay attached after flowering but usually become a pale white.

Method of fungicide application is also important. Flat fan nozzles pointed 90° down are great at covering foliage; however do not do a good job of covering the heads, which is where the product needs to be located. Use nozzles that are angled forward 30-45° down from horizontal (30 degrees is better than 45) or dual nozzles angled both forward and backward. Research has shown that a single forward-angled nozzle or nozzles angled forward and backward allow for significantly more product to contact the head and increase fungicide efficacy. Optimal spray volume is 10 gallons per acre.

Triazole (FRAC group 3) fungicides that are effective on FHB include Caramba (metconazole), Proline (prothioconazole), and Prosaro (prothioconazole + tebuconazole). This year, a new mixed mode of action product is on the market, Miravis Ace. This product contains propiconazole (DMI, FRAC 3) and pydiflumetofen (SDHI, Group 7). On the label, application can begin at Feekes 10.3 through 10.5.2. Although this product can be applied at the earlier timing, preliminary data has shown that optimal FHB control and lower DON levels are achieved at the 10.5.1 timing or a few days beyond this timing. If you spray too early, heads that have not emerged will not be protected by the fungicide application. Rainfall during flowering can increase levels of FHB and delay the ability to get into fields to apply fungicides. The expanded application window of Miravis Ace may offer options if periods of extended rainfall are in the forecast. However, if the weather allows, 10.5.1. to about 5 days after appears to provide the best control to reduced DON. We will be collecting local data on optimal application timing in Georgetown this spring. As a reminder, fungicides containing strobilurins (QoI’s, FRAC 11) should not be used past heading because these fungicides can result in elevated levels of DON.

Figure 3. From left to right Feekes 10.5, Feekes 10.5.1 (beginning flowering), Feekes 10.5.2 (flowering growth stage), Feekes 10.5.3 (full flower). Image: C. Knott, Univ. of Kentucky https://mccracken.ca.uky.edu/files/identifying_wheat_growth_stages_agr224.pdf.