Cucumber downy mildew can be transmitted in seed

October 20, 2014 in Vegetables

Cucumber downy mildew (CDM) caused by Pseudoperonospora cubensis  is an extremely damaging and important disease of cucurbits.  Our previous thinking was that P. cubensis could not be transmitted in seed, and instead epidemics were dependent on the pathogen overwintering on living host tissues and blowing into cooler production regions during the summer.  However, a recent study published in PLOS one has shown that this pathogen can be transmitted to seed of infected plants.

In the study, researchers found that 6-7% of fruits collected from infected plants, when ground and used to inoculate healthy cucurbits, could incite CDM.  A low amount of seed (roughly 1%) collected from fruit of infected plants developed CDM when planted.  Molecular assays determined that the pathogen can be found in various parts of the fruit and seed.  This finding could explain recent changes in population structure that plant pathologists have noticed in recent years.  To read more, clink this link: Seed transmission 17 10 2014 PolsOne

DON: Keeping a Mycotoxin in Check Through Ongoing Research, Sound Sampling & Producer Awareness

October 10, 2014 in Barley, Barley diseases, Wheat, Wheat Diseases

The following is a link to an article written by Don Lilleboe, an agricultural writer from North Dakota.  The article highlights the efforts of the USWBSI in combating Fusarium Head Blight in small grains.

 

Click here to access the article.

Anthracnose: foliar symptoms vs stalk infections

October 7, 2014 in Corn, Corn Disease Management

This season, growers in the Mid-Atlantic noticed an abundance of foliar anthracnose on their corn.  Growers also are likely aware that the same fungus that causes foliar anthracnose also be associated with stalk rots.  One question you may have is, “Do foliar symptoms of anthracnose indicate a stalk rot issue?

Internal and external symptoms of Anthracnose stalk rot on corn.  http://www.udel.edu/PR/UDaily/2007/aug/corn080906.html

Internal and external symptoms of anthracnose stalk rot on corn. http://www.udel.edu/PR/UDaily/2007/aug/corn080906.html

To answer that question we need to dig into the primary literature, meaning the peer-reviewed, replicated, basic research conducted by scientists published in reputable scientific journals.  The following information is from a few good papers on the subject.

Anthracnose is caused by the fungus Colletotrichum graminicola.  The fungus can be found on foliage as well as in stalks.  Foliar symptoms of anthracnose are often observed early in the season on young seedlings (prior to V6).  After the V6 stage the foliar phase of the disease can be greatly reduced in some hybrids because of the accumulation of defensive chemicals in leaves.  A second flush of foliar symptoms may be noticed on upper foliage after tasseling in some hybrids. C. graminicola can infect corn stalks through: 1) roots, 2) direct penetration of stalks and 3) wounds.

Research has shown that there is a relationship between insect injury and C. graminicola in stalks.  For example,  damage caused by the European corn borer provides points of entry that facilitates colonization of stalks by stalk rotting pathogens, including C. graminicola (e.g. Gatch and Munkvold 2002).  Studies indicate that infection is greatest if it occurs immediately after wounding, and decreases with time after wounding (Muimba-Kankolongo and Bergstrom,  2011).  This response is observed in susceptible and resistant hybrids.

C. graminicola can also enter stalks rots in the absence of insect feeding. Studies indicate that direct penetration of the stalk by the fungus ipossible, but is not efficient, occurring in only 20% of the time in inoculation studies (White and Humy 1976). In order to colonize the stalk directly the fungus enters and passes through the cells through very narrow threads of fungal tissue, eventually causing tissue discoloration and producing shiny black stromata, which give infected stalks the black appearance characteristic of the disease (Venard and Vaillancourt 2007).  Later in the season during grain fill the internal tissues of the stalk are digested and lodging may occur.  However, it must be noted that these studies were conducted on highly susceptible sweet corn hybrids.  The use of resistant varieties are highly effective in managing anthracnose stalk rot.  Muimba-Kankolongo and Bergstrom (2011) showed striking reductions in colonization of wounded, inoculated stalks (a worst case scenario) when comparing resistant to susceptible hybrids.  Even when the pathogen was directly introduced into stalks, C. graminicola failed to develop in resistant genotypes.  Anthracnose stalk rot is also, and perhaps more commonly, caused by infection of roots by the fungus present in and on soil residues.  For example, Sukno et al (2008) showed that 28% of corn plants challenged with C. graminicola  developed aanthracnose stalk rot without obvious symptoms.

In sum:  Foliar symptoms are not directly associated with stalk rot, although the presence of foliar symptoms may indicate that the field is at an increased risk for stalk-related issues.  Management for anthracnose is best achieved by resistant hybrids, residue management, crop rotation, and stress mitigation.  In addition, use of Bt hybrids and using good insect management practices will further reduce the potential for stalk-related anthracnose issues.

For more information on anthracnose in corn and its management see my factsheet.

One final point: stalk rots, including anthracnose stalk rot, are most severe and associated with high levels of plant stress, particularly around grain fill.  Any factor that limits carbohydrate production by the ear leaf can result in remobilization of carbohydrate reserves from the stalk into the grain, thereby weakening the stalk and predisposing the plant to the activities of stalk rotting pathogens.  For more information on stalk rots, see this factsheet.

 

 

References:

Bergstrom GC, and Nicholson RL. 1983. The biology ofr corn anthracnose-knowledge to exploit for improved management.  Plant Disease 83:pp 596-608.

Gatch EW, and Munkvold GP.  2002. A comparison of maize stalk rot occurance in Bt and non-Bt hybrids.  Plant Disease 86:pp 1149-1155.

Muimba-Kankolongo A, Bergstrom GC.  2011. Reduced anthracnose stalk rot in resistant maize is associated with restricted development of Colletotrichum graminicola in pith tissues.  Journal of Phytopathology 159:pp 329-341.

Sukno SA, Garcia VM, Shaw BD, Thon MR. 2008. Root infection and systemic colonization of maize by Colletotrichum graminicola.  Applied and Environmental Biology 74(3):pp 823-832.

Venard C, and Vaillancourt L. 2007. Penetration and colonization of unwounded maize tissues by the maize anthracnose pathogen Colletotrichum graminicola and the related pathogen C. sublineolum.  Mycologia, 99(3): pp 368-377.

White DG, and Humy C. 1976. Methods for inoculation of corn stalks with Colletotrichum graminicola.  Plant Disease Reporter 60:pp 898-899.

New work on chemical management of Fusarium wilt in watermelon

October 2, 2014 in Vegetables

Fusarium wilt of watermelon, caused by Fusarium oxysporum f.sp. niveum (FON), is the most important and severe soilborne disease of watermelons.  As growers know, this plant pathogen, once established in a field, is battled yearly.  This is because the fungus produces special overwintering structures that are resistant to environmental stress and persist in the soil for upwards of 10 years.  FON infects seedling roots, eventually moving into the vascular system.  The activity of the fungus eventually disrupts the function of the vascular system, resulting in wilt of vines.  Although single runners can be affected, it is not uncommon to see entire plants wilted and dead as a result of FON.  Plants that are able to survive infection often are stunted and produce fewer or smaller fruit.  For more information on Fusarium wilt of watermelons, see my article in Plant Health Progress.

Growers have limited options for managing Fusarium wilt.  The most common practice is to utilize diploid varieties with resistance/tolerance to FON race 1, the most commonly found pathogenic race; however, race 2 and 3, which are highly aggressive, are found at high levels in watermelon fields in Delaware and Maryland, as well as other areas where watermelons are produced.  These races can overcome resistance to FON race 1.  In addition, an increase in seedless watermelon production has resulted in more outbreaks of Fusarium wilt in watermelon.  This is because most of our seedless watermelon varieties lack resistance to FON race 1.  Other practices such as increasing the organic matter in soils through cover crops, avoiding excessive use of nitrate, and scouting transplants and fields can also help to reduce the impacts of Fusarium wilt on watermelon.

Research presented in a new paper published in the journal Crop Protection examined the efficacy and utility of Actigard and several fungicides in the greenhouse and field in several locations throughout the United States over a 3 year period.  A total of 13 fungicides were evaluated  on the FON susceptible cultivar Sugar Baby.  Fungicides with the greatest performance in greenhouse trials were evaluated in the field using different techniques, rates, and number of applications.  Overall, the study showed that prothioconazole and thiophanate-methyl were the most effective fungicides and that severity of Fusarium wilt can be significantly reduced when fungicides are applied through the drip.  The study indicates that three applications through the drip are likely to result in the greatest amount of disease suppression.  A supplemental label was recently recieved for Proline (prothioconazole) for use through the drip in some states, including Maryland and Delaware.  This gives growers an additional tool for managing Fusarium wilt in watermelon.

 

 

 

Nematode Control Suggestions for Vegetables

September 26, 2014 in Vegetables

The following is from Bob Mulrooney, Extension Plant Pathologist (ret.) and nematode enthusiast, UD.

 

Nematode surveys, grower sampling, and troubleshooting samples have demonstrated that varied populations of several plant parasitic nematodes occur on farms in Delaware. When nematode counts are at or above threshold levels, consider implementing control measures for susceptible crops. Many cultural practices (rotation, fallow, resistant and tolerant varieties, etc.) will reduce nematode populations. However, at times, such practices are not possible or feasible. When this is the case, consider using chemicals (nematicides) for nematode control.

Threshold Levels*

 

Damaging Nematode

   Fall Sampling

     Spring Sampling

Root-Knot 500+ 50+
Lesion 400+ 200+
Root-Knot 200+ 25+
Lesion 300+ 150+

*Above threshold levels per 250cc (1 cup) soil sample.

NOTE: Stunt, spiral, stubby-root, lance, dagger, and pin nematodes have also been detected in samples assayed from vegetable fields. At present it is not known if control measures are needed for these nematodes on vegetables in Delaware. Most nematologists would agree that populations above 1,000 would be troublesome. If control is considered advisable, recommendations for control of these nematodes will be indicated on assay report forms.

Nonchemical Management of Nematodes

Prevention of spread. Plant-feeding nematodes move only short distances – a few inches to a few feet – under their own power. Typically, nematodes are spread by the movement of infested soil and/or infected plants. Sanitation and good cultural practices are the best preventive measures against nematodes. Obtain nematode-free transplants from reputable sources. Wash soil from machinery and tools before using them at another location. Nematodes may also be spread by wind, water, soil erosion, and animals.

Crop rotation. Rotation of crops is an effective and widely used cultural practice to reduce nematode populations in the soil. To be most effective, crops that are poor hosts or nonhosts of the target nematodes should be included in the rotation sequence.

Root-knot and lesion nematodes have such a wide host range that a practical rotation plan to reduce these nematode populations for vegetable crops cannot be recommended at the present time. The exception to this is the use of root-knot resistant soybeans in a rotation. A few root-knot resistant cultivars are available.

Cover crops. Some plants commonly used as cover crops are naturally suppressive to certain nematode species, but no single crop is effective against all nematodes. The cover crop plant may be a nonhost and, therefore the nematodes starve, thus reducing their populations as with fallow. Nematodes invade the roots of certain other cover crop plants, but they fail to reproduce. Yet, other “antagonistic” plant species exude chemicals from their roots which are toxic to nematodes, including marigold and asparagus.

Soil amendments and green manures. In general, the incorporation of large amounts of organic matter into the soil reduces populations of plant-feeding nematodes. The decomposition products of some plants kill nematodes. These include butyric acid, which is released during the decomposition of rye and timothy, and isothiocyanates, released during the decomposition of rapeseed and other mustards in the genus Brassica. Maximum benefit of these”natural” nematicides is obtained when the plant material is incorporated into the soil as green manure. Green manure treatments are not equally effective against all plant- parasitic nematodes and therefore it is important to consult with a diagnostic lab or extension agent to make sure the treatment is appropriate for the nematode being controlled. For example, rapeseed is effective against dagger nematodes but not lesion nematodes. Also keep in mind that varieties of the same green manure crop can differ in the amount of toxic chemical components in their cell walls and therefore differ in the amount toxic by-products released during decomposition.

For dagger nematode control, two years of rapeseed green manure is desirable, but it may be possible to get the same benefit by growing two crops of rapeseed within one year. The following timetable is suggested for producing two rotations of rapeseed within one year:

  • Prepare seedbed and plant rapeseed by late April or early May. (Plant only recommended winter rapeseed varieties.)
  • Turn under green rapeseed by early September. Prepare seedbed and plant second crop by mid-September.
  • The second crop should be turned under in late spring after soil temperatures reach 45°F or higher.
  • Ideal conditions for incorporating the cover crop are similar to those required for obtaining the maximum benefit from fumigation (i.e., the soil should be above 45°F and moist).
  • Alternatively, planting dates may be reversed so that the first planting is in the fall followed by a second crop planted in the spring. This would end the rotation cycle in fall of the following year. Some rapeseed varieties are more effective at suppressing nematode populations than others, and some varieties will not over-winter or they bloom too early in summer to be useful. The winter varieties ‘Dwarf Essex’ and ‘Humus’ work well for both spring and fall planting dates. If planted in the spring, these varieties grow vigorously to crowd out weeds and do not go to seed.

 Tips:

  • Rapeseed requires a firm, smooth seedbed that is free of weeds, heavy residue, and large clods.
  • Seed may be drilled or broadcast. Seed at a depth of 3/8 inch and avoid planting too deep! If seed is broadcast, a cultipacker may be used to cover seed.
  • A seeding rate of 7–8 pounds per acre works well.
  • Rapeseed is sensitive to broadleaf herbicide carryover.
  • Fall-planted rapeseed should have 8–10 true leaves and a 5- to 6-inch tap root with a 3/8-inch diameter root neck before the ground freezes.
  • Sulfur is necessary for rapeseed to produce nematicidal compounds. Some soils may be deficient in sulfur. A soil test for sulfur may be beneficial. Keep in mind that some biofumigant crops like rapeseed and sorghum-sudangrass are hosts for nematodes and it is not until incorporated into the soil as green manure that they will suppress nematode populations.

Plant nutrition and general care of the plant. The harmful effects of nematodes on plants can be reduced by providing plants with adequate nutrition, moisture, and protection from stress. These tactics sometimes may be of limited usefulness, because, if susceptible crops are grown continuously, the nematode population may increase to levels that cause serious damage.

Resistance. Some vegetable varieties are available with resistance to root-knot nematode; e.g., tomato, pepper, and sweet potato. Some of these resistant varieties have limited horticultural use in our area. Check with seed company and Extension horticulturists for current variety suggestions.

Integrated management practices. Each of the practices mentioned above reduces the soil population of plant-feeding nematodes to varying degrees. Each practice has limitations. The degree of nematode control achieved depends on environmental factors, as well as the particular nematode and crop being considered. Maximum benefit is realized when several of these practices are employed in an integrated crop management program. Because the host range of different nematodes varies, the selection of cover crops, rotation crops, and green manures will be determined by the kinds of nematodes present. No single practice is a cure-all for all nematode problems.

Chemical Control Measures – Nematicides

Fumigants. Most pre-plant fumigants – dichloropropene (Telone II), chloropicrin, or metam-sodium (Busan, Nemasol, Vapam) – can be used for nematode control. Dosage, restrictions, and crop specificity are listed on the label. Follow the label instructions carefully to ensure satisfactory results. The ideal time to fumigate is when soil temperatures at a 6-inch depth range between 50-80 degrees F and the soil has moderate moisture. Fall months are generally best for fumigation. Pre-plant fumigation in the spring should be done with CAUTION. Some fumigants may linger in cool, wet soils and increase the possibility of injury to young plants.

Fields to be treated with soil fumigation must be prepared sufficiently to seed a vegetable crop. The soil should contain little or no crop debris, be free of clods, and soil moisture must be adequate to support seed germination. If soil moisture levels are low, fields should be irrigated to bring the moisture to a satisfactory level. If fields are not properly prepared, soil fumigation will be not effective due to lack of penetration of all soil particles by the gaseous fumigant.

Soil temperature at the 6-inch depth should be in the range of 50o to 80oF ( 10o to 26.7oC). Fall months are ideal for fumigation. Fumigation in the spring is less desirable because some fumigants may linger in cool, wet soils and increase the likelihood of reducing seed germination or injuring young plants.

The following multipurpose soil fumigants should be used to provide disease and nematode control:

chloropicrin – 50 gal/A, or

metam-sodium (Busan, Nemasol, Vapam HL) –37.5-75 gal/A

For nematodes only use one of the following:

 dichloropropene (Telone II)—9-12 gal/A, or

dichloropropene + chloropicrin (Telone C-17)–      11-17 gal/A, or

dichloropropene + chloropicrin (Telone C-35)–13-20.5 gal/A

Soil fumigants are injected to a depth of 6 to 8 inches. Immediately after application, soil should be dragged, rolled, or cultipacked to delay loss of fumigant. Metam-sodium is water soluble and can be injected and applied via irrigation systems (solid set sprinkler or drip/trickle). Metam-sodium must be injected for the entire time that the field is irrigated (apply an acre inch of water). Rinse the irrigation system with clean water only long enough to clear the system. Too much rinsing or a heavy rainfall within 24 hours of application will reduce the efficacy of the treatment.

At least 2 to 3 weeks should intervene between the application of most soil fumigants and the time a crop is planted. See manufacturer’s label recommendations for specific crops and fumigants.

One week after application, work soil to a depth of several inches so that gases may escape. Severe injury or killing of sensitive plants may occur if the fumigant has not sufficiently dissipated.

To determine if it is safe to plant into fumigated soil, collect a soil sample from the treated field (do not go below the treated depth). Place the sample in a glass jar with a screw top lid. Firmly press numerous seeds of a small seeded vegetable crop (lettuce, radish, etc.) on top of the soil and tighten the lid securely. Repeat the process in another jar with nonfumigated soil to serve as a check. Observe the jars within 1 to 2 days. If seeds have germinated, it is safe to plant in the field. If seeds have not germinated in the fumigated sample and have germinated in the nontreated sample, then the field is not safe to plant. Rework the field and repeat the process in a few days.

Note: Since nitrifying bacteria are reduced by the fumigants, at least 50 percent of the nitrogen in the initial fertilizer application should be in the nitrate form.

Nonfumigants. Several nonfumigant nematicides can be used on selected crops. These nematicides do not volatilize in the soil as the fumigants do. Therefore, soil temperatures and moisture requirements are not as critical for these chemicals. Be sure to check the label before using any of these materials.

 Nematicide                                             Vegetable Cleared for Use

Counter 20CR                                        Sweet corn

Mocap 10G, 15G and 6EC                    Cabbage, Snap bean, Lima bean, Sweet corn, Cucumber, Irish potato, Sweet Potato.

Do not use Mocap as a seed furrow treatment because crop injury may occur.

Vydate L                                                  Carrots, Cucumber, Cantaloupe, Eggplant, Honeydew Melon, Pepper (Bell & non-Bell), Sweet Potatoes, Squash, Tomatoes, Watermelon. Can be injected through trickle irrigation systems.

 

 

What you do in the fall can impact small grain diseases in the spring

September 26, 2014 in Barley, Barley diseases, Wheat, Wheat Diseases

Small grains will be planted in the next few weeks. The decisions made at planting can and do significantly impact many diseases of small grains in the ensuing season. The following is a brief review of some of the more important planting decisions growers will make in the next 3-4 weeks that can impact diseases in 2015.

Variety: Variety selection is the most important part of an integrated disease management program. No two varieties are created equal in terms of disease resistance or tolerance. Although it is not often possible to find a high-yielding variety with excellent resistance to all pathogens, it is possible to find some productive varieties with solid disease resistance common diseases with the potential to reduce yield such as powdery mildew and leaf blotch or contaminate grain such as Fusarium head blight. When selecting varieties, take into account what diseases tend to be issues in your fields and choose those that will help reduce the risk of severe disease epidemics through resistance. Disease resistance ratings for Mid-Atlantic small grain varieties can be found at the University of Delaware, The University of Maryland, and Virginia Tech Cooperative Extension websites. Seed companies also provide disease resistance ratings.

Planting date: Wheat planted before the Hessian fly-free date has a greater chance to be damaged by viruses such as Barley Yellow Dwarf. If you do plant before the Hessian fly-free date make sure you are planting a variety with tolerance to Barley Yellow Dwarf and follow IPM practices for aphid management. Early planting can also lead to higher levels of infection and overwintering of several foliar pathogens. This can result in more foliar and head diseases in the spring.

Stands and fertilization: Disease issues may occur in fields with excessive plant populations. Planting at excessive rates reduces airflow and increases canopy humidity, which favors the development of many diseases. Excessive fertilization promotes rapid, lush growth that can enhance disease issues. Not only does excessive fertilization result in dense canopies, but it also can cause internal metabolic shifts that influence the overall ability of the plant to defend itself against pests and plant pathogens.

 

Fusarium ear and kernel rot in corn

September 11, 2014 in Corn, Corn Disease Management

There have been a few reports of ear rots in some fields in Delaware and Maryland. The most common ear rot this season is Fusarium ear rot, which we have observed in trials at the Carvel Research Center in Georgetown, Delaware, and in variety trials in Middletown.

Fusarium ear rot tends to be more of a problem when conditions are dry and hot around flowering. Therefore, you may notice it more this year in dryland fields when compared to irrigated fields. Three species Fusarium are common throughout our corn: F. verticilloides, F. proliferatum, and F. graminearum . These fungi can produce mycotoxins under some conditions. Potential mycotoxins produced by Fusarium-ear-rotting-fungi include fumonisin, deoxynivelonol, and zereleone. Of these, fumonisins are the most important as they are known to cause equine leukoencephalomalacia, “blind staggers” in horses, pulmonary edema in swine, and have been linked to human esophageal cancers in other parts of the world.

Symptoms of Fusarium ear rot vary, but typically infected kernels are scattered throughout the ear. A white to pink fungal growth is sometimes observable on kernels and silks. Often, infected kernels have a starburst symptom, where fungal growth has damaged channels within the pericarp (Figure 1). The fungus overwinters in debris and produces spores under favorable conditions. Spores land on silks and grows into the ear as the silks senesce. Ears may also be infected through the shank or stalk. Insect and bird injury often enhances colonization of kernels.

Fusarium ear rot with characteristic starburst symptom.  Photo from: http://www.apsnet.org/publications/imageresources/PublishingImages/1999/Corn100.jpg

Fusarium Ear rot with characteristic starburst symptom. http://www.apsnet.org/publications/imageresources/PublishingImages/1999/Corn100.jpg

Harvesting early at high moisture can  help minimize potential mycotoxin production. If certain areas of the field are affected more than others, harvest these areas first and segregate this grain from cleaner grain.  Affected corn harvested for silage or grain should be dried to below 15% moisture within 1-2 days of harvest to halt the production of mycotoxins.   If you plan on storing the grain long term then it should be dried to 13% to decrease spoilage. Fusarium and other ear rotting fungi will continue to grow in higher moisture corn in bins.  Keeping bin moisture low is the easiest way to minimize mycotoxin development.

Glyphosate and Sudden Death Syndrome

September 2, 2014 in Soybean Disease Management

SDS close

A leaf showing symptoms of SDS. Photo by N. Kleczewski

Sudden Death Syndrome (SDS) is an important disease that impacts soybean growers in the Mid-Atlantic.  The disease is caused by a fungus that infects seedling roots early in the season.  After flowering, and when conditions are warm and wet, interveinal necrosis, defoliation, and plant death can occur.  For more information on SDS see my article from August 30th.

It has been suggested by some that glyposate use can exacerbate diseases of field crops.  Some suggest manganese may play a role in these putative effects.  A recent publication in the journal Plant Disease examined the effects of glyphosate on SDS, yield, and plant nutrition.  A total of 14 field experiments were conducted in the Midwest and parts of Canada from 2011 through 2013.  What did they find?

1) There were no effects of glyposate or herbicide use on SDS

2) Glyphosate use tended to be associated with increased yields

3) Glyposate did not impact plant manganese levels

4) SDS was worse in irrigated fields

 

In sum, these data indicate that glyposate use is not likely to increase SDS or alter manganese levels in plant tissues.  Glyphosate use does suppress weeds and increase yields.  If you have a field with a history of SDS, avoid over-irrigation, which favors infection and disease development.

 

Reference:  Kandal et al. 2014.  Effect of glyphosate application on sudden death syndrome of glyphosphate-resistant soybean under field conditions.  Plant Disease.http://dx.doi.org/10.1094/PDIS-06-14-0577-RE

Sudden Death Syndrome Popping Up in Mid-Atlantic Soybeans

August 30, 2014 in Soybean Disease Management

SDS dead mid

Figure 1. A soybean starting to show symptoms of SDS, including interveinal chlorosis. Photo by N. Kleczewski

Over the last two weeks we have been seeing an increase in the number of fields with Sudden Death Syndrome (SDS).  SDS is caused by a Fungus (Fusarium vurguliforme- yes another Fusarium causing problems in our crops).  The fungus is fairly unique as it is blue in color, making it  easy to diagnose if it is present on symptomatic plants. This disease starts in the soil, where the fungus overwinters as mycelium in residue, as thick walled resting spores, or even in cysts of the Soybean Cyst Nematode.  Infection occurs early in the growing season, often within the first 1-2 weeks of emergence. Cool, wet weather favors infection by the fungus. After the fungus infects the plant roots it remains fairly inactive until after flowering. Wet and warm conditions during the reproductive phases of soybean growth cause the fungus to produce toxins which move up the infected plants and eventually enter the leaves. The toxin builds up in foliage, causing foliar necrosis and defoliation (Figure 1).  The leaf veins tend to remain green for a longer period of time, giving the leaf a unique appearance (Figure 2).

SDS close

Figure 2. Foliar symptoms characteristic of SDS. Note that the veins remain gree. Photo by N. Kleczewski

sdsbluecircle

Figure 3. Blue fungal growth at the soil line or on roots is diagnostic for SDS. Photo by N. Kleczewski

 

The internal tissue of the lower stem and roots will be brown when compared to healthy plants and if you are lucky, blue fungal growth may be observed at the soil line or on the roots. The blue fungus is diagnostic for the disease (Figure 3).  Infected plants may also have compromised, rotten root systems.

As far as management- avoiding early planting dates, selecting a variety with good tolerance to SDS, and avoiding compaction are key factors to consider when planting soybeans into fields with a history of SDS.  SDS is not likely to be as big of an issue in double crop soybeans due to later planting dates.

Sclerotinia stem blight in Soybeans

August 26, 2014 in Soybean, Soybean Disease Management, Soybean diseases

We have had a few reports of sclerotinia stem blight (white mold) on soybeans grown in the mid-Atlantic. This is a cool season disease that we see almost every year, but only to a very small degree. Often you will see it in high yield environments and in shaded areas of the field, such as along wood edges. The fungus overwinters as small, pebble-like structures in the soil which germinate to directly infect plants or produce a mushroom-like structure that can produce millions of spores over several days.  Sclerotinia spores are wimpy and require dead/dying tissues to germinate and take hold.  This is why we see the fungus cause issues during flowering- spores will land on decaying flower petals and grow into the flower and eventually the stem.  Over time lesions with distinct margins develop and more of the black pebble-like structures may be observed in or on the stem.  Affected plants may wilt or mature prematurely.

Sclerotinia stem blight.  Note the black structures on the stem and sharp lesion edges.  Photo: C. Whaley.

Sclerotinia stem blight. Note the black structures on the stem and sharp lesion edges. Photo: C. Whaley.

 

If you notice Sclerotinia stem blight (white mold) in your field it is likely there to stay. The best management practice at this point in full season beans is to schedule infested fields to be harvested last to minimize spread of the pathogen from field to field on farm equipment.  Hot weather is not favorable for this disease and only a few days of hot dry weather are needed to burn out Sclerotinia.

 
In soybeans planted into fields with a known history of Sclerotinia stem blight, and when weather is cool and wet, applications targeting the R1-R2 stage are the most efficacious.   Example fungicides for SSB suppression include labeled group 1’s (Topsin), labeled group 3’s (Proline, Domark, Topguard), Fluazinam (Omega); labeled group 7’s (Endura) and labeled group 11’s (Aproach). Often fungicide applications are not justified in Delaware or Maryland but they can be beneficial under some circumstances. As usual, preventative applications are most efficacious. For more information on fungicide ratings, please see the NCERA 212/218 fungicide efficacy tables click this link: Soybean Fungicide efficacy table_FINALx

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