Sclerotinia sclerotiorum is a plant pathogenic fungus that causes important diseases known as white mold, Sclerotinia wilt or stalk rot, or Sclerotinia head rot on a wide variety of broadleaf crops grown in the North Dakota - Minnesota region. The pathogen is wide spread in the eastern half of North Dakota, especially in the Red River Valley. It is commonly found damaging dry beans, sunflowers, soybeans and canola. There are many other crops in this region that are susceptible such as field pea, potato, mustard, safflower, lentils, flax, borage, crambe, buckwheat, chickpea, lupine, faba bean and numerous vegetables. Some of these crops are rarely damaged by S. sclerotiorum, while other are quite susceptible. With the expansion of canola acreage and other susceptible crops, Sclerotinia will continue to increase in importance. This pathogen is known to infect about 408 species of plants. In the south eastern United States, the fungus is a problem on peanut and alfalfa. Numerous weeds such as marsh elder, lambsquarters, pigweed, Canada thistle, sow thistle, and wild mustard are also hosts in this area, and can play a role in disease cycles.
There are two other species of Sclerotinia, S. trifoliorum and S. minor. Neither of these are reported in North Dakota. S. trifoliorum is known only on alfalfa and forage legumes in the south east and eastern USA, while S. minor is primarily a lettuce and peanut pathogen found in the eastern and western coastal areas. S. trifoliorum is similar in biology and morphology to S. sclerotiorum, but S. minor produces much smaller sclerotia and generally does not produce apothecia in nature.
S. sclerotiorum has become an important pathogen in this area for numerous reasons. First, there is a large acreage of susceptible crops for the fungus to reproduce on, second, there are cold soil temperatures about half the year so microbial activity that could reduce survival of the pathogen is curtailed, third, our generally drier soil conditions favor survival, and fourth, our environmental conditions during the growing season favor infection and production of inoculum.
The primary survival (overwintering) structure of S. sclerotiorum is the sclerotium. A sclerotium is a hard resting structure consisting of a light colored interior portion called a medulla and an exterior black protective covering called the rind. The rind contains melanin pigments which are highly resistant to degradation, while the medulla consists of fungal cells rich in -glucans and proteins. The shape and size of sclerotia depend the host on where they are produced in or on infected plants.
What is the origin of sclerotia in a field? There are four primary methods that fields are infested with sclerotia. The most common is by susceptible crops or weeds being infected by ascospores coming from adjacent infested fields. The fungus then produces sclerotia on those plants and some are returned to the soil when the field is harvested. Wind transported soil or crop debris infested with sclerotia are also known to contaminate adjacent fields. Contaminated machinery can introduce sclerotia into a field. Surface irrigation water or rain water moving naturally between fields can also move sclerotia to previously clean fields. Seed contaminated with sclerotia is another method of introducing the fungus into clean fields.
The basic disease cycle of Sclerotinia diseases begins with the overwintering of sclerotia in the soil. In the growing season, overwintered sclerotia can germinate in one of two methods. Probably the most common is carpogenic germination which results in the production of a small mushroom called an apothecium. The apothecium forms spores called ascospores which are wind transported to susceptible plants. The other method of germination is myceliogenic, where the sclerotium produces mycelium. The only crop in this region where myceliogenic germination plays a major role in the disease cycle is in Sclerotinia wilt of sunflower. Sclerotinia wilt is caused by sclerotia germinating and infecting the sunflower roots. All the other Sclerotinia or white mold diseases, such as on dry beans, soybean, canola and sunflower head rot are initiated by carpogenic germination and infection of above group plants parts by ascospores (see the section by J. Venette).
How long do sclerotia survive in the soil? Few studies have quantified sclerotia survival in the field. There are many factors affecting survival such as soil type, previous crops, initial population of sclerotia and environmental conditions, but how and to what degree they affect survival is not well understood. High temperature and high soil moisture combined are probably the two most deleterious environmental factors. Microbial degradation, however, is the principal reason for a decline in the populations of sclerotia. There are many fungi, bacteria and other soil organisms that parasitize or utilize sclerotia as carbon sources. One reason that crop rotation is recommended for Sclerotinia is to allow the natural microbial population to degrade sclerotia. Two important fungal parasites in this area are Coniothyrium minitans and Sporidesmium sclerotivorum. Both these fungi have been touted as possible biocontrol agents for sclerotia, but their use to control Sclerotinia in this area has not been adequately studied and there are no commercial products yet available.
The effect of tillage on survival of sclerotia is poorly studied and no generalizations can be made to aid in management of the pathogen. There is evidence that leaving the sclerotia on the soil surface enhances degradation whereas burying the sclerotia enhances survival. It is thought that the more dramatic changes in temperature and moisture on the soil surface are deleterious to sclerotia.
Survival in North Dakota was studied in 32 naturally infested fields over the years 1980 to 1886. Inoculum densities ( number of sclerotia in a unit of soil) in fields ranged from <1 to 12 sclerotia/ 800 cm3 of soil. In fields in rotation to non-susceptible crops for 3 to 5 years, the average annual reduction in populations of sclerotia in the soil was about 22% per year (range 15-39%). Fields with populations as low as 1 sclerotium/800 cm3 of soil needed 5 to 6 years in rotation before detection of sclerotia was difficult by soil sampling. Unfortunately, even such a long rotation does not guarantee the elimination of Sclerotinia. Soil sampling can be used to estimate inoculum densities in fields, but Sclerotinia wilt of sunflower is the only disease where such information can be used to predict disease incidence. For Sclerotinia diseases initiated by ascospores, there is no model which can predict the relationship between inoculum density (sclerotia in soil) and disease. Part of the reason for this is that environment, specifically moisture, plays a critical role in ascospore production, infection and disease development.
Inoculum density is the most important factor affecting the development of Sclerotinia wilt of sunflower. An inoculum density of less then 0.5 sclerotia/800 cm3 of soil can result in significant incidence of disease. The effect of soil factors and environmental factors on Sclerotinia wilt are not understood. Sclerotinia wilt, however, develops on a variety of soil types and during both wet and dry years. Most sunflower hybrids are susceptible to Sclerotinia wilt, but hybrids are under development which are less susceptible than those currently on the market.
The first plants showing Sclerotinia wilt in a field generally do not appear until several weeks prior to flowering and most plants wilt after flowering. The higher the inoculum density the earlier wilt symptoms will appear in the field. The appearance of disease later in the season is related to root growth of sunflower. Maximum root growth occurs at flowering, thus maximum root-sclerotia contact does not occur until then. Root senescence begins after flowering, which may lower the innate ability of the roots to resist decay by the fungus. Also, contact between plants within a row does not begin until around 30 days after planting. These factors favor root infection and plant to plant spread of the fungus later in the season. When monitoring for Sclerotinia wilt, it is important to do the final monitoring as late in the season as possible, because an early monitoring can dramatically underestimate the total number of infected plants. Those infected plants will be returning sclerotia to the soil, thus affecting the inoculum for future susceptible crops.
Selected References:
Abawi, G. S., and Grogan, R. G. 1979. Epidemiology of diseases caused by Sclerotinia species. Phytopathology 69:899- 904.
Adams, P. B., and Ayers, W. A. 1979. Ecology of Sclerotinia species. Phytopathology 69:896- 899.
Boland, G. J. and Hall, R. 1994. Index of plant hosts of Sclerotinia sclerotiorum. Can J Plant Pathol. 16: 93-100.
Gulya, T. J. 1984. Incidence and severity of sunflower disease in the Dakotas and Minnesota during the 1984 growing season, p6. In: Proceedings Sunflower Research Workshop, Fargo, ND, Dec. 10-11, 1984.
Gulya, T., Fick, B. and Nelson, B. 1989. Sclerotinia head rot of sunflower in North Dakota: 1986 incidence, effect on yield and oil components, and sources of resistance. Plant Disease 73:504-507.
Holley, R. C., and B. Nelson. 1986. Effect of plant population and inoculum density on incidence of Sclerotinia wilt of sunflower. Phytopathology 76:71-74.
Nelson, B. 1983. Inoculum density of Sclerotinia sclerotiorum in naturally infested fields in North Dakota. Pages 22-23. In: Proceedings Sunflower Research Workshop, January 26, 1983, Minot, ND.
Nelson, B. 1984. The effect of plant stage and root growth on incidence of Sclerotinia wilt of sunflower. Phytopathology 74:813.
Nelson, B. D., Hertsgaard, D. M. and Holley, R. C. 1989. Disease progress of Sclerotinia wilt of sunflower at varying plant populations, inoculum densities and environments. Phytopathology 79:1358-1363.
Nelson, B., and A. H. Lamey. 1984. Sclerotinia Diseases of Sunflower. Cooperative Extension Service Circular PP-840. North Dakota State University. 8pp.
Purdy, L. H. 1979. Sclerotinia sclerotiorum: History, disease and symptomatology, host range, geographic distribution and impact. Phytopathology 69:875-880.
Introduction. Ascospores are the primary inoculum units for dispersal of Sclerotinia sclerotiorum, the fungal pathogen that causes white mold of dry bean, sunflower, canola, soybean, pea, lentil, flax, potato, and other broad-leaved crops. Changing agricultural practices, especially increased plantings of crops susceptible to Sclerotinia, has increased the risk of substantial economic losses. The pathogen can attack more than 360 species of plants including common weeds so it is difficult to control by rotation. Fungicides can control white mold on some crops, but the value of fungicides is dependent on protection of susceptible plant parts contaminated by airborne ascospores. The following discussion focuses on the processes in which Sclerotinia spores are formed, transported, and initiate infection.
Production of fruiting bodies. Sclerotia are the principal structures for overwintering and for long-term survival in field conditions. Large numbers of sclerotia can be produced in heavily infected crops. We have calculated that as many as 1 million sclerotia per acre were returned to the soil from dry beans severely infected by Sclerotinia. If nutrients are available, sclerotia can germinate directly with new mycelial growth (=mycelogenic germination). If nutrients are limited, sclerotia can germinate with the production of a specialized sexual fruiting body that eventually liberates numerous spores (=carpogenic germination). Fruiting bodies develop from sclerotia that are wet, conditioned, and are near the soil surface. Minimum or shallow cultivation places many of the sclerotia at depths of 0.25 in to 1.25 in which is optimum for emergence of sexual fruiting bodies called apothecia. Formation of apothecia generally occurs after a period of sclerotia dormancy during which the sclerotia are chilled or frozen. Cold temperature seems to be the predominate factor in "conditioning" sclerotia so that when soil conditions are suitable (moisture >50% field capacity, temperature 59-64 F) for 10-14 days, apothecia can form. Temperatures of 59-64 F are optimum, but apothecia can form in soils at temperatures of 40 to 86 F which means they can be produced throughout the growing season if moisture is sufficient. Carpogenic germination begins with active fungal growth in regions of the sclerotial cortex or medulla. Growing fungal cells form dense primordia which break through the rind of the sclerotium, and continue growth as tube-shaped stalks called stipes. Large sclerotia such as those from sunflower can produce as many as 40 stipes, although all the stipes may not be produced at the same time. Smaller sclerotia such as those from dry bean produce fewer stipes. In the field, we generally find 1 to 4 stipes associated with single sclerotia from bean. Stipes are negatively geotropic, and after the stipes emerge from the soil, they continue to grow upward to a height of about 1 cm. If the stipes are exposed to light, especially ultraviolet light (<390 nm), they differentiate into apothecia.
In the formation of an apothecium, the tip of the stipe expands to form a top surface made up of the hymenium and hypothecium. These layers are supported by the tapering tissues of the former stipe, now called the excipulum, giving the overall structure the appearance of a tan- to buff-colored trumpet bell or a golf tee. In the hymenium are born numerous asci and sterile supportive hairlike structures, the paraphyses. In the asci, sexual recombination occurs, and the products are eight ascospores, neatly lined-up near the tip of each ascus. A vacuole, which is responsible for increasing the hydrostatic pressure within the ascus, forms below the string of ascospores. As the hydrostatic pressure builds, the ascus expands, but its lateral expansion is restricted by the paraphyses and neighboring asci. Therefore, most expansion is near the tip, and expansion continues until each ascus protrudes beyond the paraphyses. At some point, the pressure exceeds that supportable by the stretched wall and the ascus explodes.
Puffing of spores into turbulent wind. Ascospores of Sclerotinia are physically shot out from the surface of the hymenium in an upward direction . The display is spectacular primarily because tens of thousands of asci fire their ascospores almost simultaneously producing a large puff of spores. In bright light, the hundreds of thousands of spores can be seen with the naked eye as a smoke-like cloud. The kick from the firing is often strong enough to shake the entire apothecium, and in quiet air, the spores are propelled several centimeters above the hymenium. Puffing is an adaptive feature because the simultaneous firing of asci creates air currents that help elevate the spores above the hymenium. In the field, spore discharge occurs for several hours beginning about noon or when the sun is near the zenith. Timing of spore discharge is an important adaptation for increased spore dispersal. Stipes are positively phototropic and align the hymenium with the strongest light source. The tips of the asci are also positively phototropic which helps fine-tune the trajectory of spores for maximum dispersal potential. At solar noon, radiation into upright canopies such as grain fields is greatest. The sun warms the soil creating thermal air turbulence that helps carry spores out of the canopy. The change in temperature also brings about a change in relative humidity which can trigger puffing.
Number of spores produced. Each apothecium can produce from 2 to 30 million spores over a period of several days. In some grain fields infested with sclerotia from diseased dry beans, we have assayed over 40,000 apothecia per acre. This means the potential spore load would equal a billion to a trillion spores from each acre of infested land. We have collected ascospores emanating from these fields with an Andersen air sampler, a vacuum device that draws air at 1 cubic ft/min through a series of stacked sieves beneath each of which is a petri dish filled with a Sclerotinia-selective medium. The largest spores are trapped beneath the largest sieve, and progressively smaller spores are trapped in the subsequent stages. By sampling for known periods of time, we can determine number of spores per unit volume of air as well as determine the spore size distribution within the air mass. Over a three year period, we have detected as many as 7,400 Sclerotinia ascospores per cubic meter of air at the margin of a grain field. This large number of spores represents a serious danger to nearby susceptible crops such as dry bean, sunflower heads, and canola. By collecting simultaneous samples at the margin of an infested grain field and distances of 10 m and 30 m downwind, we were generally unable to show reductions in spore numbers with distance, although spores collected at both of the distances were fewer than those at the source. Most spores were caught in stages 1 and 2 of the sampler, with progressively fewer in stages 3 and 4. None were caught in stages 5 and 6. Particles in stage 1 have an aerodynamic diameter greater than 8 m; in stage 2, 5 m; in stage 3, 3 m; and stage 4, 2 m. Spores in stages 1 and 2 probably represented clumps of Sclerotinia spores, and by their large mass would be less apt to be windblown long distances. Spores in stages 3 and 4 represent single spores, and spores with these small diameters could be blown great distances. Even particles in stage 2 would have a settling time of 1 mm per sec, meaning that a spore blown to a height of 20 ft would require over 100 minutes to settle to ground level. In a 15 mph wind, that time would allow the spore to travel 25 miles. When we sampled air 20 ft above the downwind edge of a grain field, we found spore numbers 70% to 85% those at the canopy level, indicating substantial populations available for mid-range transport.
Sticky spores. Ascospores of Sclerotinia sclerotiorum are covered with a sticky mucilage, the origin of which is unknown. It may be residue of the liquid from asci, or it may be part of the cell wall. Regardless of its origin, the mucilage is very sticky. The mucilage not only cements the spore to any object it contacts, it also glues spores together in clumps. We were able to capture ascospores on fine polymer threads and examine them microscopically for proportion of clumping. Eighty five percent of the spores were captured as single spores, 10% were in clusters of two, 3% were in clusters of three, and 2% were in clusters of four. If Sclerotinia is like another ascomycete, Sordaria, the viscosity of the mucilage will be greater at low temperatures which would favor spore clumping. The larger clumps of spores are probably deposited or impact near the apothecium. In cool, wet weather, clumps of spores may be more effective in establishing infections. When we placed graded spore doses on detached bean blossoms, we found single spores on each blossom produced 25% infection. Two spores per blossom produced 75% infection, and four spores per blossom produced 100% infection. Blossoms with the larger number of spores decayed more rapidly.
Spore survival. In laboratory conditions, dried frozen Sclerotinia ascospores have remained viable for years, but freshly produced spores generally survive 5 to 21 days depending on relative humidity. On foliage in the field, spores survive about 12 days. Little is known about spore survival while spores are airborne.
Infection of dry bean, soybean and canola. For dry bean, soybean, canola, and some other crops, ascospores first colonize dead or nearly dead plant material as a nutrient source. Senescing blossoms are the most important nutrient source although almost any damaged tissues can be colonized. After initial saprophytic growth, the fungus proceeds into healthy plant tissue by direct penetration. Wetness is important for both blossom colonization and infection of healthy tissue. In dry bean, for example, senescing blossoms must be wet for about 48 hours during which spores germinate and the fungal mycelium colonizes the blossom tissue. Ascospores germinate best at temperatures of 68 to 77 F. In wet conditions, the fungus can proceed from the old blossom into healthy plant tissues in about 16 to 24 hours. The fungus can colonize healthy tissue and produce new sclerotia in 10 to 14 days. In dry weather, disease progression in a plant can be slowed or stopped, but will resume when extended periods of plant wetness favor fungal growth. Closed canopies, narrow rows, restricted wind patterns, irrigation, prolonged dew, and frequent rains all contribute to extended periods of plant wetness. Sclerotinia can continue to cause damage to plants in windrows should the windrows remain wet.
The fungus can spread from plant to plant by contact, or if infected plant material is dispersed but this form of spread is limited. The fungus does not produce any important asexual spores, so epidemic outbreaks from secondary spores are not a concern.
Plant invasion. Sclerotinia is characterized as a necrotrophic fungus meaning that it lives on dead and dying plant tissue. Some of the earliest studies on the disease showed that the pathogen kills cells ahead of the advancing mycelium. Sclerotinia produces toxic oxalic acid which is partly responsible for plant cell death. In addition, the oxalic acid creates an acidic environment in which the many degradative enzymes produced by the fungus are most efficient.
Conclusion. Sclerotinia sclerotiorum is well adapted for pathogenesis on many broad-leaved crops. Sclerotia can survive for years in soil. Apothecia can produce millions of spores, and the fungus has adaptations for insuring spores are inserted into airstreams. Spores can be transported in the wind for miles and a sticky mucilage insures spores stick to any surface they contact. If spores contact weakened plant material, the fungus can grow saprophytically and then proceed into healthy plant material as a parasite. This mode of pathogenesis makes Sclerotinia very difficult to control, especially during prolonged periods of cool, wet weather. Agricultural practices such as increased irrigation, conservation tillage, and increased acreages of susceptible crops indicate Sclerotinia diseases will continue as an important threat to crop productivity.
Canola. Observations from eight trials by Thompson, Thomas and Evans in Alberta in 1981 demonstrated that there was a close relationship between Sclerotinia incidence (percent infected stems) and disease loss. In six trials with Benlate, yield differences between the sprayed and the unsprayed portions of fields ranged from 131 lb/A to 1,009 lb/A. Differences in incidence between sprayed and unsprayed ranged from 3% for the sprayed and 26% for the unsprayed to 6% for the sprayed and 66% for the unsprayed. By comparing the differences in incidence between sprayed and unsprayed with the yield differences for these same fields I developed a "loss factor". I calculated this as follows: loss factor = % yield difference (yield difference/treated yield)/difference in incidence (treated incidence - untreated incidence) where yield difference is the difference in yield between the treated and untreated portion of the field. For the six fields treated with Benlate, the loss factor was 0.75. Two other fields sprayed with Rovral had a loss factor of 0.65. The average loss factor for all eight fields was 0.72.
Observations of Lamey from irrigated trials at Oakes, ND in 1991 and 1993 showed a yield difference of 477 to 578 lb/A between the unsprayed and plots sprayed with Benlate or Rovral. The loss factor was 0.63. The average loss factor for all 10 trials in Alberta and North Dakota is 0.70. This factor is used in calculating 1997 losses in Minnesota and North Dakota.
Canola Losses in 1997. The average incidence in 63 fields surveyed in Bottineau, Rolette, Towner, Ramsey and Cavalier Counties of North Dakota was 14.2%, for a calculated loss of 10.1%. The average incidence for Cavalier County was 21.1% for a calculated loss of 14.8%. The average incidence for the 10 worst fields in North Dakota was 42%, for a calculated loss of 29.4%. Incidence in the worst field was 70% for a calculated loss of 49%.
The average incidence in 19 fields surveyed in Roseau and Lake of the Woods Counties of Minnesota was 18.8%, for a calculated loss of 13.2%. The average incidence for Roseau County was 24.2% for a calculated loss of 16.9%. The average incidence for the five worst fields in Minnesota was 42%, for a calculated loss of 29.4%. Incidence in the worst field was 75% for a calculated loss of 52.5%.
Dry Bean. In 1992 irrigated trials at Staples, MN, Meronuck recorded a yield of 1,174 lb/A on dry beans sprayed with fluazinam and 456 lb/A in the unsprayed, for a yield difference of 718 lb, or a 61% yield reduction. Total yield loss was greater than 61%, however, since there was 35% white mold in the treated plot at first white mold reading versus 89% in the untreated.
In 1986 and 1987 nonirrigated grower strip trials at Buxton and Northwood, ND, yield differences ranged from 523 to 720 lb/A when unsprayed strips were compared with strips sprayed with either Benlate or Topsin M. White mold was as high as 61% and 72% in two of the trials compared to 48% and 34%, respectively, for the treated strips. These are yield differences of 30% and 21%.
Soybean. Grau reported five epidemics of Sclerotinia stem rot on soybean in Wisconsin. His experimental data indicated a yield loss of 230 lb/A for each 10% of incidence in Wells II, a maturity II cultivar. Grau indicated that a more precise relationship was defined by using a 1-3 severity scale. Using the severity scale, he defined a linear relationship with a yield of 62 bu/A at 0% severity, 42 bu/a at 40% severity and 21 bu/A at 80% severity. Thus, losses at 40% severity would be 33% and losses at 80% severity would be 67%.
Sunflower. Separate assessments of losses can be made for Sclerotinia wilt and for head rot. Wilt can occur at any stage of growth, although the majority of wilt seems to occur following the initiation of bloom (anthesis). Wilt is especially important to the sunflower producer since it can occur whenever sunflower is planted into an infested field. Wilt occurrence is essentially independent of weather. Head rot, which develops from ascospore infections, is highly dependent on the occurrence of wet weather at flowering.
Wilt. In studies at Morden, MB, on oilseed sunflower, Dorrell and Huang sowed sclerotia along with the seed (1:3 weight ratio). They observed only 7% wilt at anthesis, but 60% wilt eight weeks after the initiation of flowering. They determined the seed yield, 1000 seed weight and seed oil content of plants infected at 1 through 8 weeks after anthesis and compared these data to data for apparently healthy plants. If wilt occurred within 4 weeks of anthesis, seed yield losses were 70% or greater. Seed yield losses were over 98% on plants infected one week after anthesis, about 70% if infected three or four weeks after anthesis, and about 12% if infected 8 weeks after anthesis. The 1000 seed weight was reduced from 76 g for the healthy to 27 g for plants infected 1 week after anthesis, for a 64% reduction in seed weight. The 1000 seed weight was 53.3 g when infection occurred four weeks after anthesis, an acceptable seed weight. There was a reduction in oil content from 46% for the healthy to 32% for plants infected two weeks after anthesis. This is a reduction of 14 percentage points, or a 30% reduction in oil content. There was no change in linoleic or stearic acids, a slight increase in palmitic acid when infection occurred early, and a slight reduction in oleic acid content. There was only a slight effect on meal protein. No similar data is available for confection sunflower. However, similar reductions in seed yield and seed weight might be expected.
Head Rot. Gulya, Vick and Nelson determined the effect of head rot on seed yield, seed weight and oil content on heads collected from 25 North Dakota fields, with 10 healthy and 10 infected heads collected from each field. On oilseed sunflower, they found the yield per head (after sclerotia were removed) reduced 31% by head rot, the 200 seed weight reduced 10% and the oil content reduced 2%. On confection sunflower, they found the yield per head reduced 36% and the 200 seed weight reduced 11%.
White mold [Sclerotinia sclerotiorum (Lib.) deBary] is a major concern to dry bean (Phaseolus vulgaris L.) growers in most of the production areas in the United States. A survey of dry bean growers in North Dakota and Minnesota found that respondents ranked white mold as the most serious disease problem, with fungicides used on 33% of the acreage in an attempt to control the disease. Crop losses averaging 30% in Nebraska with individual field losses as high as 92% have been observed. In North Dakota, estimates of economic loss caused by white mold have not been thoroughly investigated, but it has been estimated that yield loss and cost of fungicide applications can exceed $30 million annually.
Control of white mold is difficult. Application of Benzimidazole fungicides is costly, but has been the primary method of control. Timing of fungicide applications and the mechanism of application during the blossom period are critical to effect good control; applications may be hampered by wet weather conditions that favor disease development. Other cultural practices, such as crop rotation, tillage practices, and reduced seeding rates, recommended to control the pathogen, have met with little success.
Physiological resistance and plant architecture have both been identified as possible mechanisms to reduce white mold damage in dry bean. Because resistance in P. vulgaris is low and probably controlled by several genes, use of avoidance mechanisms, including upright and open plant structure, less dense canopies and branching patterns, elevated pod set, and reduced lodging have been suggested to reduce damage to the white mold pathogen. These mechanisms enhance penetration of the canopy by sun and aids air circulation, thereby creating a microclimate that is less conducive for infection and disease progression.
Evaluating material in the field is hampered by the vagaries of environmental conditions that exist at the test site, including temperature, moisture, and uniform distribution of sclerotia throughout the experimental site; these environmental factors also affect development of the plant canopy, which causes further variability in evaluation. Germplasm evaluation also is hampered by the avoidance mechanisms that may reduce injury of a fully susceptible plant. These traits may be useful in production fields that have not had a history of severe white mold damage; however, avoidance alone does not prevent disease development of susceptible materials in areas that have high white mold potential. Evaluating dry bean lines using various greenhouse and laboratory techniques has been only slightly successful; in general, these tests are very labor intensive and time consuming. With these tests, several replications and runs are necessary to achieve accurate results, thereby limiting their use to screen early generation breeding material for white mold resistance.
Previous work by the breeding program focused on developing improved screening procedures to evaluate for white mold resistance and to determine the mode of inheritance from three sources of resistance. A former graduate student at NDSU, Phil Miklas, worked on this project. An in vitro screening method was evaluated as a way to determine genetic resistance to white mold. Hypocotyl (stem) tissue was grown on a solid tissue culture medium amended with white mold culture or control filtrate. Putative resistance was identified as callus fresh weight, expressed as a percentage of control callus fresh weight.
A second laboratory procedure that Phil evaluated used excised stems from 28-day old plants inoculated with white mold mycelium below the sixth node was tested in an attempt to identify genetic resistance in dry bean. Within the navy bean market class, Bunsi had the lowest lesion length (LL) value, 68 mm, while D76125 and 'Upland' had the highest values (82.1 mm and 92.6 mm, respectively) [LSD 0.05 = 11.3]. The lesion length assay correlated well (r = 0.68, P < 0.02) with field disease incidence.
Lastly, we reported on inheritance of white mold resistance in bean. Means for lesion length and disease incidence were normally distributed, with transgressive segregation occurring for both traits. Heritability estimates for LL (0.27,0.38, and 0.66) were lower than DII (0.77, 0.58. and 0.70) for each population tested. Low genetic correlations between LL and DII suggested that selection for both traits is warranted.
Recently, a new inoculation test was described that is quick, easy, and highly repeatable. This test promises to be of major importance in evaluating dry bean germplasm for physiological resistance in the greenhouse. This "straw test" has been used to verify field response as well as evaluate germplasm for putative resistance.
Selection for genetic resistance to destructive pests and pathogens is important for yield stability and high quality. Genetic resistance is the most economically feasible method to control most pathogens; plant resistance also is useful in a sustainable agricultural system and reduces harm to the environment because of reduction or elimination of pesticide usage. Selection for improved white mold resistance in bean has been hampered by: 1) Lack of easy, reliable screening procedures in field and laboratory environments; and 2) minimal genetic variability for resistance. Recently, efforts to identify high levels of white mold resistance have intensified, making available several lines, notably in snap beans, with useable levels of resistance. Also, resistances have been identified in dry bean lines. Use of snap bean germplasm by dry bean breeding programs has been very limited because undesirable traits often are found in progenies from their crosses. Also, while my lab has focused on improving evaluation techniques to improve selection efficiency for white mold resistance, these techniques are labor intensive and require considerable greenhouse space. Combining phenotypic response and selection for QTL markers (from another study) across environments may improve the efficiency in breeding for resistance to this important pathogen.
NDSU navy bean breeding line 88-106-04 has exhibited considerable levels of resistance to white mold in field trials in North America from 1994 through 1997. This resistance was derived from the navy bean cultivar Bunsi, which exhibits high levels of resistance in both field and some greenhouse trials. We are transferring this resistance into adapted genotypes of pinto, pink, and other market classes by a series of hybridizations, followed by rigorous selection. In addition, other putative sources of white mold resistance continue to be evaluated and. If different sources of resistance are available, they will be hybridized with genotypes representing major market classes. To date, we have identified several lines with considerable resistance to white mold, including I9365-5, I9365-31, 92BG-7, ND91-047-04-01, ND91-076-01, and G122. (Interestingly, G122 also possesses high levels of resistance to root rots in Minnesota). While the breeding program has repeatedly used resistance sources, this project would focus exclusively on the transfer and selection of resistance into other seed classes, notably pinto bean.
Dr. Berlin Nelson supervised the evaluation of white mold on soybean cultivars at Page and Northwood, ND in 1996 and 1997. Forty commercially available public and private cultivars were evaluated in 1996 and 47 cultivars were evaluated in 1997. The percentage of plants with a lexion on the main stem was recorded for each of four replicates at each location. The correlation between locations for percent plants infected with white mold was r = 0.55 (P>0.001) in 1996 and r = 0.59 (P>0.001) in 1997. These inter-location correlations indicate that differences among cultivars was due to both genetic and environmental causes. The correlations between locations is a measure of how much of the variation among cultivars is due to the effects of genes.
In 1996 there were no significant correlations between lodging, date of physiological maturity, plant height, and grain yield with severity of white mold infection. In 1997 there was a correlation of r = 0.68 (P>0.001) between white mold infection and date of physiological maturity. This correlation indicates that early maturing cultivars tended to have less white mold infection compared to late maturing cultivars. However, the correlation between white mold infection and physiological maturity was significant only in 1997.
In 1997 there was a correlation of r = 0.67 (P>0.001) between lodging severity and white mold infection. Cultivars that were more susceptible to lodging had greater whitemold infection in 1997, but not in 1996. In 1997 there was a correlation of r = -0.75 (P>0.01) between grain yield and white mold infection. This correlation indicates that those cultivars that were more susceptible to whitemold infection tended to yield less in 1997, but not in 1996. The results indictae that in some years, cultivars that are early maturing, and lodging resistant tend to have less white mold infection and higher yield.
Some cultivars were susceptible to white mold in both 1996 and 1997. These cultivars should be considered most susceptible. Other cultivars were only susceptible in one of the two years and these cultivars may be susceptible only under specific environmental conditions. Some cultivars showed little white mold infection in both years. Based on the available data, growers that are likely to have white mold problems should plant cultivars that had the least white mold infection in both 1996 and 1997 seasons.
The NDSU soybean breeding program has a goal of developing high yielding cultivars that are stress tolerant. Iron chlorosis, drought tolerance, tolerance to waterlogged soils, and disease tolerance are all factors associated with stress. Experimental lines that are high yielding are evaluated for tolerance to these different stress factors. Soybean growers with a specific stress factor can then determine whether a cultivar is adapted to their conditions.
|
Table 1. White mold disease severity in 1996, 1997, two-year average and date of physiological maturity, averaged across Page and Northwood, ND for soybean cultivars. | |||||
|
|
|
|
|
|
|
|
Pioneer |
9004 |
3 |
1 |
2 |
Sept. 8 |
|
Novartis |
S03-C3 |
--- |
1 |
--- |
Sept. 14 |
|
Mycogen |
5006 |
--- |
1 |
--- |
Sept. 9 |
|
Dairyland |
006 |
--- |
2 |
--- |
Sept. 11 |
|
U of MN |
McCall |
4 |
2 |
3 |
Sept. 10 |
|
Pioneer |
9007 |
12 |
6 |
9 |
Sept. 12 |
|
Novartis |
S00-66 |
6 |
6 |
6 |
Sept. 17 |
|
DeKalb |
CX046 |
10 |
7 |
9 |
Sept. 24 |
|
Novartis |
Solano |
5 |
8 |
7 |
Sept. 16 |
|
Hyland |
Enterprise |
--- |
8 |
--- |
Sept. 24 |
|
Payco |
9802 |
--- |
11 |
--- |
Sept. 17 |
|
NDSU |
Traill |
--- |
14 |
--- |
Sept. 17 |
|
U of MN |
Glacier |
10 |
15 |
13 |
Sept. 13 |
|
Mycogen |
5072 |
--- |
15 |
--- |
Sept. 26 |
|
Wensman |
3067 |
--- |
16 |
--- |
Sept. 23 |
|
Pioneer |
9042 |
8 |
17 |
13 |
Sept. 23 |
|
Wensman |
3036 |
19 |
18 |
19 |
Sept. 24 |
|
NDSU |
Council |
10 |
18 |
14 |
Sept. 27 |
|
Terra |
E036 |
--- |
19 |
--- |
Sept. 27 |
|
U of MN |
Ozzie |
12 |
19 |
16 |
Sept. 25 |
|
Payco |
9803 |
--- |
20 |
--- |
Sept. 23 |
|
U of MN |
Proto |
8 |
21 |
15 |
Sept. 25 |
|
Hyland |
Maverick |
--- |
21 |
--- |
Sept. 25 |
|
Stine |
0470 |
25 |
21 |
23 |
Sept. 26 |
|
Payco |
9606 |
13 |
21 |
17 |
Sept. 25 |
|
Gold Country |
Bygland |
25 |
22 |
24 |
Sept. 23 |
|
AgriPro |
0110 |
--- |
22 |
--- |
Sept. 15 |
|
Mycogen |
5100 |
--- |
23 |
--- |
Sept. 30 |
|
DeKalb |
CX025 |
27 |
23 |
25 |
Sept. 23 |
|
Terra |
E067 |
--- |
24 |
--- |
Sept. 29 |
|
Sem. Prograin |
Korada |
9 |
24 |
17 |
Sept. 20 |
|
U of MN |
Lambert |
14 |
24 |
19 |
Sept. 28 |
|
Payco |
9804 |
--- |
24 |
--- |
Sept. 25 |
|
Stine |
0480 |
--- |
24 |
--- |
Sept. 26 |
|
Mycogen |
013 |
27 |
24 |
26 |
Sept. 24 |
|
Terra |
E047 |
--- |
25 |
--- |
Sept. 25 |
|
Stine |
0280 |
--- |
16 |
--- |
Sept. 25 |
|
U of MN |
Agassiz |
17 |
27 |
22 |
Sept. 17 |
|
NDSU |
Danatto |
9 |
27 |
18 |
Sept. 23 |
|
Ostland |
DG-3046 |
--- |
29 |
--- |
Sept. 24 |
|
Gold Country |
Tracker |
23 |
31 |
27 |
Sept. 24 |
|
Stine |
0653 |
18 |
33 |
26 |
Sept. 27 |
|
U of MN |
Minnatto |
6 |
35 |
21 |
Sept. 29 |
|
Wensman |
3075 |
--- |
39 |
--- |
Sept. 28 |
|
U of MN |
M301 |
30 |
42 |
36 |
Sept. 24 |
|
Payco |
9508 |
14 |
42 |
28 |
Sept. 25 |
|
Gold Country |
Windsor |
--- |
44 |
--- |
Sept. 26 |
The USDA-ARS genetics and pathology projects began a program several years ago to identify and develop germplasm with increased resistance to Sclerotinia stalk rot. The project was designed in three phases: 1) identify and select germplasm with increased resistance, 2) develop recurrent phenotypic selection procedures and populations by intermating resistant germplasm, and 3) cross highly resistant germplasms to develop new, improved germplasm with increased resistance. Natural infection, found in producers' fields with previous problems, was the primary tool to identify resistant germplasm. Artificial infection was an effective screening method to verify natural infection resistance and to assist in the recurrent selection procedure.
Three accessions originating from Russia, Ukraine, and Romania were found to have increased tolerance to Sclerotinia and were crossed with USDA derived materials to create populations for recurrent selection. Several lines have been derived from the populations and have been released to industry and public breeders and researchers. These lines were two restorer or male lines: FHA 408 and RHA 409; and three maintainer or female lines: HA 410, HA 411, and HA 412. Hybrids between these lines and testers produced hybrids which were approximately 15% infected with Sclerotinia stalk rot, whereas the check Hybrid 894 was approximately 35% infected. Total immunity to wilt has not been found in any line or population. The resistance observed in the lines appears to be polygenic, with population selection procedures effective in combining many genes to create genotypes with higher levels of resistance.
In the future, additional lines will be developed and released to industry programs derived from new introductions crossed with the released lines. These new lines will provide sunflower hybrids even more protection from Sclerotinia stalk rot. The challenge will be for industry companies to incorporate the resistance found in the USDA lines into their programs and offer new hybrids to producers.
White mold continues to cause serious loss to dry bean yield and quality. Bean varieties resistant to white mold are not yet available so disease management is based on cultural practices and on fungicides. Fungicides can reduce disease and improve yield, but fungicides are expensive and disease reduction is not consistent.
Previous experiments showed foliar applications of calcium solutions could reduce white mold. Reduction of white mold may be due to calcium's role in strengthening plant tissue. Calcium forms strengthening bridges especially in the pectate materials that form the middle lamellae of plant cells. Calcium is also important in maintaining selectivity of cell plasmalemmas and in binding the plasmalemma to the cell wall. Calcium also binds strongly to oxalic acid, an important toxin produced by Sclerotinia sclerotiorum
In 1997, calcium solutions were tested for white mold reduction in strip trials in an irrigated field near Crete, ND. Navy beans in strips 60 ft x 0.5 mile were band sprayed on 16 July and 23 July with Benlate 50W at 1.5 lb/A, calcium chelate at 2 gal/A, or with calcium sulfate at 5.5 lb/A in 50 gal water/A at 100 psi. One stip was left untreated. White mold was evaluated on 1 August at 12 sites, 200 ft apart in each strip and again on 8 August at 10 sites, 250 ft apart in each strip. At each site, 22 evaluations for white mold were made and percentage of infection was calculated. On 1 August, 9% of the plants treated with calcium chelate and 5% of the plants sprayed with calcium sulfate were infected. These infection rates were higher than that for Benlate-sprayed plants (2%) but were substantially lower than that of plants in the untreated strip (23%). By 8 August, disease progressed, and 3.0% of the plants treated with Benlate were infected. Plants treated with calcium sulfate (12.0% infection) or with calcium chelate (9.0% infection) had much less white mold than plants in the unsprayed strip (41% infection). Yields at Crete were not determined.
At the Carrington ND Research/Extension Center, a similar trial was made. Pinto beans in strips 20 ft x 150 ft were treated with Topsin M at 1.5 lb/A, calcium sulfate or calcium chelate at rates as before but in 37 gal water/A applied through drop nozzles at 35 psi. Sprays were applied on 11 July at 5-10% bloom and again 18 July. White mold was evaluated on August 4 and 11. In addition to percent plants infected, disease severity was rated and yields were determined. By mid-season, plants in the trial were severely affected by white mold. For example, on 4 August, 94% of the plants in the untreated strip were infected with an average severity of 50%. Topsin M reduced disease to 43% and severity to 30%. Plants treated with calcium sulfate had 83% infections and severity of 38%. The calcium chelate reduced infection by 9% and severity by 2%. By 11 August, only plants with Topsin M had reduced infection, but severity was reduced by all of the spray treatments. Untreated plants had the lowest yield (24.5 cwt/A) and Topsin M-treated plants had the greatest yield (36 cwt/A). Calcium compounds improved yield by 2-3 cwt/A.
In a study that combined calcium compounds with Topsin M for white mold control, heavy white mold disease pressure developed in a field trial at Carrington, ND. By the end of the season, 97 % of the unsprayed control plants were infected. Topsin M at 1.5 lb/A provided excellent white mold control resulting in a 10 cwt/A yield advantage. Among the calcium compounds tested as a supplement to Topsin M at 0.5 lb/A, calcium sulfate at 5.5 lb/A provided the best control. Control was significantly better than that provided by Topsin M alone at 0.5 lb/A and the calcium sulfate- Topsin M combination produced a yield advantage of 6 cwt/A over the untreated control. Combinations of calcium compounds with fungicides can provide suitable control at reduced cost. These tests indicate that foliar-applied calcium may be a nutritional supplement that increases plant resistance to white mold. Strip trial results were similar to those of earlier small-plot trials.
Application of fungicides to dry beans has been observed in dry beans in North Dakota producer strip trials since 1986. Several trials have included different methods of application.
A 1994 trial at Northwood, ND compared application of Benlate on Agri 1 navy beans planted on a 30 inch row spacing. Treatments included application with an Ag Cat airplane using 5 gallons/A (gpa), broadcast application using 100 psi and 24 gpa, application with drop nozzles (directed spray) using 175 psi and 16 gpa and air assist application using 32 psi and 15 gpa. White mold incidence in the untreated was over 90%; it was 35-40% in the treatments involving drop nozzles or the air assist. White mold in the broadcast treatment was intermediate between the untreated and the drop nozzle treatment. White mold in the aerial application treatment was nearly as high as the untreated. Yields ranged from slightly less than 2,000 lb/A in the untreated and the aerial application treatment to nearly 3,000 lb/A in the treatments with drop nozzles or the air assist. Yield in the broadcast treatment was intermediate. Dockage was highest in the untreated and lowest in the treatment with the air assist, followed by the drop nozzle treatment.
A producer in Grand Forks County, who was unable to use ground application due to excessive rainfall, had his field sprayed with Topsin M using aerial application at 7½ gpa. He had planted Othello pinto beans with a 36 inch row spacing. Topsin M applied 8 days after initiation of bloom resulted in yield increases of 600-700 lb/A in two different treatments. Two applications were not better than one. Application of Topsin M 16 days after early bloom resulted in a yield increase of 200-300 lb/A in two different treatments.
Another trial comparing equipment and application methods was conducted in 1995 at Hatton. The trial was on Upland navy beans with a 36 inch row spacing. Treatments included broadcast application at 40 psi and 24 gpa (Broad Lo), broadcast application at 100 psi and 24 gpa (Broad Hi), Broadcast application twice seven days apart at 100 psi and 24 gpa (Broad Hi2), directed (drop nozzles) spray at 40 psi and 15 gpa (Drop Lo), directed spray at 175 psi and 24 gpa (Drop Hi), directed spray at 175 psi and 15 gpa followed seven days later by broadcast application at 100 psi and 24 gpa (Drop Hi2), and an air assist application at 15 gpa (Air Asst). White mold incidence was 45% in the untreated, 21% in the Broad Lo, 12% in the Broad Hi, 17% in the Broad Hi2, 10% in the Drop Lo, 1% in the Drop Hi, and negligible in the Drop Hi2 and Air Asst. Most treatments had about a 500 lb/A increase in yield over the untreated check. The treatments with the lowest white mold did not out yield the other treatments due a heavy infestation of weeds in that part of the field.
Two trials were conducted at Cummings in 1996 comparing application timing of Topsin M. In a trial with Schooner navy bean planted on a 30 inch row spacing, Topsin M was applied by air using an Ag Cat and 7 gpa. The untreated had 32% white mold and a yield of 1,580 lb/A. Application at first bloom (0 days) resulted in 18% white mold and a yield of 1,894 lb/A; application at 4 days resulted in 14% white mold and a yield of 2,094 lb/A; application at 10 days resulted in 7% white mold and 2,231 lb/A. In this trial, delaying 4-10 days after early bloom was superior to application at early bloom.
The other trial at Cummings was with Envoy navy beans planted on a 22 inch row spacing. Topsin M was applied by air with an Ag Cat using 9 gpa. The untreated had 78% white mold and a yield of 1,295. Application at first bloom (0 days) resulted in 42% white mold and a yield of 1,849 lb/A; application at 3 days resulted in 40% white mold and a yield of 1,769 lb/A; application at 10 days resulted in 60% white mold and a yield of 1,269. In this trial application at 0 or 3 days was superior to application at 10 days. Several explanations may be involved: 1) Venette has shown that when infection is initiated by a larger population of spores, infection proceeds more rapidly, which implies that a fungicide must be applied more quickly; or 2) a delay in application may have made canopy penetration more difficult with 22 inch row spacing. Both trials at Cummings showed good white mold control and good yield response when Topsin M was applied 3-4 days after early bloom, regardless of row spacing or disease pressure.
An aerial application trial in 1996 at Mayville compared three different gallonages for application. The trial was conducted on RS 101 pinto beans planted on a 30 inch row spacing. Aerial application was with a turbine Air Tractor (approximate air speed = 130 mph, compared to 95 for a piston Ag Cat). White mold was 85% in the untreated strip with a yield of 3,010 lb/A. White mold in the 5 gpa treatment was 71% with a yield of 3,670 lb/A; white mold in the 7½ gpa treatment was 51% with a yield of 3,980 lb/A; white mold in the 10 gpa treatment was 58% with a yield of 3,960 lb/A. Use of 5 gpa provided some white mold control and a yield increase of over 600 lb/A, but use of 7½ or 10 gpa provided better white mold control and a yield increase of over 900 lb/A.
Summary Results. High pressure ground application with drop nozzles or an air assist unit has provided good white mold control and good yield increases. Low pressure ground application with drop nozzles or high pressure broadcast spraying has provided moderately good white mold control and yield. Aerial application has provided moderately good to good white mold control and yield. Use of 7½-10 gpa appears superior to use of 5 gpa.
Most existing data has been obtained on 30 or 36 inch row spacings. As sugarbeet acreage increases, 22 inch row spacing is becoming more common on dry beans. Since we have almost no data on application to this row spacing and canopy penetration will be more difficult, extrapolation of current data to use on 22 inch row spacing may not be reliable. A directed spray using drop nozzles is almost impossible. We cannot be certain that a broadcast spray or an air assist spray will penetrate the canopy as well with 22 inch row spacing as with wider row spacing, particularly if application is delayed past early bloom. We have one trial indicating that aerial application provided good white mold control and a good yield increase on a field with 22 inch rows, but do not know if canopy penetration is as good as with wider row spacings.
Spray Decision Guidelines. A white mold spray is likely to provide an economic return if: 1) There has been wet weather for 10-14 days before flowering, 2) it is wet and humid at flowering, 3) if white mold has been a problem in the planted field or nearby fields within the past 2-4 years, and 4) if the yield potential is 2,000 lb/A (which is likely if there has been cool wet weather and there is a good stand).
Meronuck's prediction data provides a producer with an experimentally based means of determining the potential of an economic return. A producer only needs to keep track of rainfall. If the rainfall (or rainfall and irrigation combined, for irrigated fields) from June 1 until 10 days after initiation of bloom has been 3-5 inches, a spray will be economic 20% of the time; if the rainfall during this period has been 5-7 inches, a spray will be economic 65% of the time; if the rainfall during this period has been over 7 inches, a spray will be economic 85% of the time.
Two applications. Data presented above do not show any advantage of two applications over a single application. Data from some 1997 trials (Venette) and some producer observations in northwestern Minnesota indicate that two applications (or even split applications = same total product as a single application, but delivered twice at half rates) may provide better performance than a single application at early bloom. Providing protection over a longer period of time may explain the better performance of two applications or of a split application in some cases.
Value of fungicides. The preponderance of scientific evidence indicates that fungicides have value in increasing yield of dry beans attacked by white mold. Unfortunately, growers often observe substantial numbers of diseased bean plants in fields following fungicide treatment and are frustrated that the apparent lack of control did not justify costs of application(s). When I examined 42 independent studies conducted in a number of states over the last 15 years, I found an average yield increase of 28.9% from the best treatment in each test.
The value of fungicide control is difficult to determine for a number of reasons. First, the wet conditions that favor white mold development also favor increased bean yields. Studies in Nebraska showed that bean yields responded to moisture as long as the crop had less than 40% molded plants. With more than 40% molded plants, the disease reduced yield.
Second, the yield potential of the crop needs to be sufficient to justify the costs of treatment. Dr. Lamey has used 2000 lbs/A yield potential as a lower limit in a white mold spray decision guide, and this yield seems to be practical. In none of the 42 fungicide trials I examined were potential yields determined.
Third, in some field trials reduction of white mold incidence (% of plants infected) and/or severity (the degree of disease progression) does not correlate with improved yield. We also have observed cases in which fungicides had no apparent effect on white mold, yet yields were significantly improved. Benzimidazole fungicides have the phytohormonal activity of a cytokinin, an anti-senesence material. In field trials, other diseases complicate test results, and white mold can develop in spots or patterns that make differences in yield due to fungicide application difficult to detect.
Fourth, disease pressure must be sufficient to justify application, but disease pressure is difficult estimate directly. In canola, petal assays are direct measures of disease pressure, and blossom assays are being developed for dry bean and soybean. An inherent problem with these assays is obtaining results in time for growers to make a spray decision. In his spray decision guide, Dr. Lamey has employed pathogen presence ("White mold in the area.") as well as environmental factors (described in another section) as indirect ways to estimate disease pressure. After-the-fact studies have shown little value in applying fungicides when fewer than 25% of the plants became infected.
Fifth, the economic value of fungicide treatment is clearly related to the market price of harvested beans. If beans are grown under contract, the selling price may be known. Otherwise, there is no good way to predict that yield increases will be economically justified. The fungicides most commonly applied by growers are expensive ($30+/A/application; less if band applied), and growers' reluctance to spray is understandable. If two applications of fungicide are band applied at a total cost of $40/A, a yield increase of 153 lbs/A sold at $26/CWT is sufficient to recover costs. In contrast, a yield increase of 267 lbs/A would be needed if the selling price was only $15/CWT.
Sixth, growers often do not have a good way to determine any increased yield as a result of fungicide treatment. Understandably, growers may be reluctant to leave an untreated check strip in a field for yield comparisons. Sometimes check strips are left, but they may not represent the rest of the field. For example, a check strip in a portion of a field exposed to the wind may have less disease than in a portion of the field protected by a shelterbelt. While comparisons with results on other fields are tempting, the huge array of factors that affect disease development make such judgements tenuous.
Registered dry bean fungicides. Fungicides registered for white mold control in dry beans are thiophanate methyl (Topsin M), benomyl (Benlate), and iprodione (Rovral). I know of no new products that will be available for the 1998 growing season. Grower surveys have shown that thiophanate methyl and benomyl are the fungicides most frequently applied for white mold control. Both of these materials are classified as benzimidazole fungicides. Both benomyl and thiophanate methyl break down to a fungitoxic compound (MBC, carbendazim) which is mostly responsible for the activity of the fungicides. Benomyl breaks down very rapidly to MBC in water, but thiophanate methyl is converted to a MBC through the actions of sunlight and metabolic processes of the plant and fungus. Both of these materials affect spore germination and cellular multiplication by interfering with DNA synthesis. Because the benzimidazoles have a single-site mode of action, resistance to the fungicides is well known in other disease systems, but Sclerotinia sclerotiorum resistance to benzimidazoles is not currently a concern. Systemic movement of the benzimidazoles is through the cell walls and through the water transport system; therefore the general movement is upward and outward in treated plants.
Iprodione (Rovral) has successfully controlled white mold in other areas, but we have not been able to demonstrate superior levels of control in any of our trials in North Dakota. Iprodione can break down in spray water with a high pH, and the breakdown may have affected our results. Iprodione belongs to the imide group of fungicides, but it is not generally considered systemic in plants. Iprodione has crop rotation restrictions.
Registered soybean fungicides. Thiophanate methyl (Topsin M) is now labeled for white mold control in soybean. It is to be applied at 3/4 to 1 lb/A at early bloom (R1 to R2) followed by a second application 7 to 14 days later if conditions are favorable for continued disease pressure.
Benomyl (Benlate) is cleared for application to soybean for control of certain stem and leaf diseases, but it is NOT cleared for white mold (Sclerotinia) control. For control of the stem and leaf diseases, benomyl is applied at early pod set (pods 1/8 to 1/4 inch long at one of four main stem upper nodes) and at a rate of 1 lb/A. A second application may be made 14 to 21 days later as needed. Apparently, benomyl has provided economic control of white mold on soybean even the rate seems low and the timing seems late.
Experimental fungicides that control white mold. We have shown in North Dakota tests that vinclozolin (Ronilan) is one of the most effective fungicides for control of white mold in bean. Vinclozolin is an imide fungicide, structurally similar to iprodione, that is not systemic but apparently has preventive and curative activity. Ronalin is cleared for application to canola in Canada, but its apparent toxicity to humans has slowed its registration for control of white mold on crops in the United States.
Fluazinam (Fluazinam) is a new generation fungicide with activity against Sclerotinia. We have obtained good to excellent control of white mold with fluazinam in North Dakota trials. Fluazinam was being developed by ISK Biosciences, but the company was sold and the parent company ISK Japan retained fluazinam. The potential for further testing and eventual registration remains unknown.
Azoxystrobin (Quadris) is a beta-methoxyacrylate fungicide which traces its hertiage back to a naturally produced fungicide from another fungus. This fungicide functions by attaching to a specific site on cytochrome B, inhibits electron transport from cytochrome B to cytochrome C1, and therefore inhibits production of ATP. We have not tested any of the fungicides in this class on dry beans; but in another state, Quadris showed excellent control of white mold on dry beans.
Canola. No fungicide is registered for Sclerotinia stem rot on canola. A request for a section 18 for the use of Benlate on canola in Minnesota and North Dakota in 1996 was denied by the Environmental Protection Agency (EPA). The Agency agreed that there was an emergency, but denied the request because the risk cup for Benlate was full. The risk cup is a new concept under the new Food Quality Protection Act (FQPA), which became effective in August 1996. FQPA requires EPA to consider all sources of risk from a product and other products with similar toxicity. Sources of risk considered include all foods which may be treated with the product, worker exposure, environmental exposure, home and garden or turf (includes golf courses) exposure, and exposure to products with related toxicity. EPA noted that both Benlate and Topsin M break down to methyl benzimidazole carbamate (MBC), and that this should be taken into consideration in risk assessment.
Benlate is in the IR-4 minor use registration program. Two years of residue trials have been conducted in North Dakota and several other states and a processing study has been done. The manufacturer of Benlate, du Pont, has the residue samples and expects to complete residue analysis and submit a complete registration package to IR-4 by April, so that IR-4 can request registration from EPA. du Pont has also done a risk analysis which they will provide to EPA. du Pont believes that the risk is less than that assumed by EPA.
Zeneca has a new product, Quadris, which has been tested extensively in Canada, but very little in the U.S. Trials in Manitoba, Saskatchewan and Alberta show good activity against blackleg and Alternaria black spot, and activity against Sclerotinia comparable or nearly comparable to Benlate. Quadris is a new class of chemistry that is considered environmentally friendly by EPA. It qualifies for "fast track" registration, which could result in registration taking 14-18 months instead of 3-4 years. Residue trials are complete and Zeneca expects to apply for Canadian registration soon. A request for U.S. registration may be submitted within the next two years.
Sunflower. No fungicide is registered for control of Sclerotinia head rot or wilt on sunflower. There are no immediate prospects of a fungicide being registered.
Registration of Topsin M as a seed treatment for seed borne Sclerotinia is being pursued through the IR-4 minor use program. It is highly effective against seed infections from Sclerotinia. This occurs when head rot is serious in seed production fields. The seed coats and outer portions of many seeds from these fields are invaded by the Sclerotinia fungus, resulting in poor germination. Seed treatment with Topsin M effectively restores good germination, if there are no other problems with the seed. Seed treatment with Topsin M will also provide early season protection against Sclerotinia wilt, but is not effective against most wilt infections, since most occur after the initiation of flowering.
Dorrell, D.G. and H.C. Huang, 1978. Influence of Sclerotinia Wilt on Seed Yield and Quality of Sunflower Wilted at Different Stages of Development. Crop Science 18:974-976.
Grau, C.R., 1988. Sclerotinia Stem Rot of Soybean. In T.D. Wyllie and D.H. Scott, eds., Soybean Diseases of the North Central Region. APS Press, The American Phytopathological Society, p. 56-66.
Gulya, T.J., B.A. Vick and B.D. Nelson, 1989. Sclerotinia Head Rot of Sunflower in North Dakota: 1986 Incidence, Effect on Yield and Oil Components, and Sources of Resistance. Plant Disease 73:504-507.
Hoes, J.A., and H.C. Huang. 1985. Effect of Between-row and Within-row Spacings on Development of Sclerotinia Wilt and Yield of Sunflower. Canadian Journal of Plant Pathology 7:98-102.
Thompson, J.R., P.M. Thomas and I.R. Evans, 1984. Efficacy of Aerial Application of Benomyl and Iprodione for the Control of Sclerotinia Stem Rot of Canola (Rapeseed) in Central Alberta. Canadian Journal of Plant Pathology 6:75-77.
There are numerous Web sites with information on Sclerotinia, but some are too far away to provide useful information for North Dakota and Minnesota. Following are two Canadian Web sites that provide good information on canola and one site from the University of Wisconsin Soybean Plant Health Department.
Canola Council - http://www.canola-council.org
Alberta Agriculture - http://www.agric.gov.ab.ca/index.html
UW Soybean Plant Health - http://www.plantpath.wisc.edu/soyhealth
Email: hbrown@ndsuext.nodak.edu
Published by the Department of Plant Pathology