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MAY BE RECALLED with earlier due date if requested. DATE DUE DATE DUE DATE DUE «WE: '“2 l! ”2007 2/05 p:/ClRC/DateDue.indd-p.1 STRATEGIES FOR MANAGING F USARIUM CROWN AND ROOT ROT ON ASPARAGUS By James W. Counts Jr. A THESIS Submitted to Michigan State University in partial fulfillment of the requirements for the degree of MASTER OF SCIENCE Department of Plant Pathology 2006 ABSTRACT STRATEGIES FOR MANAGING F USARIUM CROWN AND ROOT ROT ON ASPARAGUS By James W. Counts Jr. F usarium oxysporum f.sp. asparagi and F usarium proliferatum are soil—borne pathogens responsible for significant productivity declines in the asparagus industry. The objectives of this research were to evaluate chemical and biological controls applied as a crown dip or injected through drip irrigation along with different irrigation schedules and to evaluate commercial cultivars for resistance to F usarium spp. Two crown dip and two irrigation trials were established in fields with a known crop and disease history. Biological treatments include Trichoderma, nonpathogenic F usarium spp, and arbuscular mycorrhizae and the chemical treatments include thiophanate-methyl, benomyl, and fludioxonil. Treatments were applied at planting or to established plantings every 30- days via drip irrigation. Results between trials were not consistent. Phytophthora spp. were detected in one of the trials and would not be controlled by the treatments used When the oomycete fungicide mefenoxam was applied the number of fem with Phytophthora symptoms decreased. Twenty commercially available asparagus cultivars were screened for resistance to F usarium spp. Germinated seedlings were transferred to test tubes containing Hoagland’s media, allowed to grow for 14 days, and inoculated with F usarium spp. Disease assesments were made using a scale of 1 (healthy) to 6 (dead). Significant differences occurred between cultivars but no cultivar received a rating that would be considered F usarium resistant. ACKNOWLEDGEMENTS I would like to thank my major professor, Dr. Mary K. Hausbeck, for her guidance and support throughout this project. I would also like to thank the rest of my guidance committee, Dr. Mathieu Ngouajio and Dr. Gene Safir, for their assistance and suggestions. The Hausbeck lab, particularly Brian Cortright and Dr. Catarina Saude, provided many suggestions and hours in the field assisting in this research. Robert Podolsky was invaluable help in working through the many challenges of the statistical analysis. I would also like to extend my thanks to the growers (Oomen Brothers, Schafer’s, and Jerry Mahlburg) who allowed the research to be conducted on their farms. iii TABLE OF CONTENTS LIST OF TABLES ........................................................................................................ LIST OF FIGURES ....................................................................................................... LITERATURE REVIEW .............................................................................................. Introduction Cultivars and Cultural Practices Diseases Control Literature cited SECTION I CROWN DIP AND DRIP APPLIED FUNGICIDES AND BIOCONTROL AGENTS FOR MANAGING FUSARIUM CROWN AND ROOT ROT ON ASPRAGUS ............................................................................ Abstract Introduction Materials and Methods Results Discussion Literature Cited SECTION II SCREENING COMERCIALLY AVAILABLE ASPRAGUS CULTIVARS FOR SUSCEPTIBILITY TO F USARI UM SPP. IN A CONTROLLED ENVIRONMENT ..................................................................... Abstract Introduction Materials and Methods Results Discussion Literature Cited CONCLUSION ............................................................................................................. APPENDIX I ................................................................................................................. APPENDIX 11 ............................................................................................................... iv V vii \IUJOOANH Hg 24 25 27 29 38 47 51 55 56 57 6O 64 67 69 72 73 87 LIST OF TABLES Table Page 1. Crown Dip and Drip Applied Fungicides and Biocontrol Agents for Managing Fusarium Crown and Root Rot on Asparagus 1. Average monthly maximum and minimum temperatures and total monthly rainfall during the 2001 through 2005 growing season. ................ 3O 2. Average stand count and fern height for asparagus crowns treated at planting with fungicides or biocontrol agents in Crown Trial 1. ......... 39 3. Average stand count and fern height for asparagus treated at planting with fungicides or biocontrol agents in Crown Trial 2. ..................... 4O 4. Average stand count and fern height for asparagus plants treated with fungicides or biocontrol agents at planting and during the season in Irrigation Trial 1 .................................................................. 43 5. Average stand count of plants with Phytophthora symptoms when asparagus plants were irrigated and/or treated with fungicides during the growing season in Irrigation Trial 2 .................................................... 44 6. Average stand count of all plants when asparagus plants were treated with fungicides or biocontrol agents at planting and during the season in Irrigation Trial 2. ............................................................................ 45 7. Average stand count of total fern when asparagus plants were treated with the fungicide mefenoxam versus not treated Irrigation Trial 2 ................................................................................................... 46 II. Screening Commercially Available Asparagus Cultivars for Susceptibility to Fusarium spp. in a Controlled Environment. 8. Asparagus cultivars tested for susceptibility to F. oxysporum f. sp. asparagi and F. proliferatum. .............................................. 61 9. Scale of disease ratings on inoculated asparagus seedlings. ................................. 62 10. Average disease ratings for asparagus cultivars inoculated with F. oxysporum f.sp asparagi 2-1. .................................................................. 65 LIST OF TABLES (Continued) Table Page 11. Average disease ratings for asparagus cultivars inoculated with F proliferatum P-67. ................................................................................... 66 III. Appendix I 12. F umigation materials tested by product name and active ingredient for efficacy on Fusarium crown and root rot on asparagus seedlings. ................. 75 13. Weight evaluation on l-year-old asparagus seedlings produced in fumigated soil for managing Fusarium crown and root rot of asparagus ............. 79 14. Root length evaluation of asparagus seedlings produced in fumigated soil for managing Fusarium crown and root rot of asparagus. ............................. 80 15. Root lesion count evaluation of asparagus seedlings produced in fumigated soil for managing Fusarium crown and root rot of asparagus ............. 81 16. Fern lesion count evaluation of asparagus seedlings produced in fumigated soil for managing F usarium crown and root rot of asparagus. ........... 82 17. Root health evaluation of asparagus seedlings produced in fumigated soil for managing Fusarium crown and root rot of asparagus. ........... 83 18. F em health evaluation of asparagus seedlings produced in fumigated soil for managing Fusarium crown and root rot of asparagus. ........... 84 19. Plant vigor evaluation of asparagus seedlings produced in fumigated soil for managing Fusarium crown and root rot of asparagus. ........... 85 20. Weed evaluation of asparagus seedbeds treated with various fumigants. .......... 86 IV. Appendix II 21. Percent difference in stand count and fern height for asparagus treated at planting with fungicides or biocontrol agents in Crown Trial 2 ..................... 88 vi LIST OF FIGURES F i ure Page 1. Crown Dip and Drip Applied Fungicides and Biocontrol Agents for Managing Fusarium Crown and Root Rot on Asparagus. 1. Layout of a single treatment plot in Crown Trial 2 (A) and Irrigation Trial 2 (B) depicting which rows received treatment and the application interval .................................................................................. 33 2. Common Phytophthora symptoms found on asparagus: A) shepherd’s crooking of the fern, B) water soaking and necrosis of the crown and roots, C) water-soaked lesion on the spear found at the soil line, and D) crooking and water soaking observed on the spears. ................................................................. 41 11. Screening Commercially Available Asparagus Cultivars for Susceptibility to F usarium spp. in a Controlled Environment. 3. The F usarium symptoms observed on inoculated asparagus seedlings receiving a rating of a 2 (lefi) to a 6 (right). ......................................... 63 Images in this thesis are presented in color. vii LITERATURE REVIEW INTRODUCTION Asparagus oflicinalis is an herbaceous perennial and is a member of the Liliaceae family (Drost, 1997). Asparagus is considered a major vegetable crop around the world, although certain species are also used in landscaping. Asparagus is cultured around the world and is adapted to soils that are often not suitable for other crops (Hartung and Stephens, 1983). Native to the Mediterranean region of Europe, asparagus was initially cultivated by the Greeks and Romans. The Romans published information regarding asparagus care around 161 BC. (Elmer et. al., 1996). The Romans used asparagus for both culinary and medicinal purposes such as diuretics, sedatives, and painkillers (Drost, 1997). During the first century the culture of asparagus spread throughout Europe and in the 16th century the French Huguenots introduced asparagus into North America (Elmer et. al., 1996). Production Regions For many years, the United States was the largest producer of asparagus, mainly producing fresh green asparagus. Beginning around 1990, China and South American countries entered the production arena and became major exporters of white and green asparagus to the United States, Europe, and Japan (Calvin and Cooke, 1997; Elmer, 20013). Currently, China produces the most asparagus in the world with 5.5 million tons (Kosco and Jimenez, 2004). Peru produces approximately 185,000 tons and ranks second (Kosco and Jimenez, 2004). The United States now stands third and produced approximately 122,020 tons in 2003. Mexican production has recently increased to an estimated 53,286 tons and ranks fourth (Kosco and Jimenez, 2004). Spain is the fifth leading producer of asparagus and is also a major importer of fresh asparagus. Along with Spain, Germany is one of the largest growers of asparagus in Europe and is also a major consumer of fresh asparagus, importing much of their asparagus from Greece. In the United States, there are three main production areas: California, Washington, and Michigan. Michigan is the second largest producer in the United States with approximately 350 farms and 14,500 acres in asparagus production. Approximately 85% of the Michigan crop goes to processing (26.4 million pounds in 2004) with the remaining 15% sold on the fresh market (Michigan Asparagus Advisory Board, 2000; Wallingsford, 2004). The amount going to fresh market has increased over the last few years, reaching 5.4 million pounds in the 2005 production year (Kleweno, 2005). West central Michigan is the major producer for the processing market, while the fresh market sector is more evenly distributed throughout the state (Kleweno and Matthews, 2002). Physiology The asparagus crown includes the rhizome as well as the adventitious and lateral roots. The rhizome consists of buds that form in clusters (Drost, 1997). Lateral buds form new growing points along the sides of the rhizome, which produce spears. Spears that are not harvested develop into fern and replenish the crown’s carbohydrate supply for the following year (Drost, 1997). Asparagus leaves are reduced scales located on the central stem. At the axils, lateral branches may arise, which also contain scale leaves. Cladophylls are small, cylindrical, leaf-like structures that are actually modified branches. They form in the whorl of scale leaves at nodes and are the main photosynthetic portion of the plant (Drost, 1997; Gilman, 1999). However, all chlorophyll-containing parts of the plant are capable of photosynthesis. CULTIVARS AND CULTURAL PRACTICES Asparagus is a dieocious species with separate male and female plants. Historically, the asparagus cultivars Mary Washington or Martha Washington dominated the asparagus industry. However, these cultivars produce a large number of female plants which can decrease the productivity of the field since energy is transferred from replenishing food reserves to reproduction (Drost, 1997). As all-male hybrids became available, growers experienced higher yields. Many of these hybrids can be traced back to the Washington strains. In the United States, asparagus breeding programs have been established in California and New Jersey. The California lines tend to be more tolerant of hot, dry weather than the New Jersey lines. In California, the asparagus production regions occur in the desert valleys and the Delta and Central Costal region; both tend to favor the California cultivars (Mullen et. al., 1998). In the northern growing regions of the United States where the weather is cooler and wetter, the New Jersey breeding lines are favored. Production life for healthy asparagus fields can be as long as 15 or more years. Peak production begins in approximately the fourth growing season and continues for eight to ten years (Mullen et. al., 1998). Harvesting starts in the second year following planting but is limited to two to three pickings to allow field establishment. The number of harvests increases from eight to twenty pickings in the third and fourth years (Dartt et. aL,2002) Asparagus harvest occurs at various times during the year depending on geographical location. In Michigan, harvesting typically begins around the first week of May in southern Michigan and one to two weeks later in the northern regions. Harvesting concludes in late June or early July (Zandstra et. al., 1992). Depending on the weather, spears are harvested daily or every two to three days (Mullen et. al., 1998) by hand (cutting or snapping) or with a mechanical harvester (still in development). Once harvested, asparagus must be kept cool due to its high perishablity. Depending on the intended market, spears are either hydra-cooled for five minutes at 0° C and then stored (0-2.2° C) for up to three weeks, or stored in a cool area until processed (Zandstra et. al., 1992) Crown Production Asparagus crown production is a specialized industry that produces one-year-old crowns by a certified grower or nurseryman (Zandstra et. al., 1992; Motes et. al., 1996). Some asparagus producers also grow crowns (Motes et. al., 1996). Site selection is key for quality crown production and should include a virgin site with low weed and pathogen pressure. Bare-root crowns are produced in seedbeds that are established between mid- April and mid-May using seeds that are uniform in size and have a high germination rate (greater than 90%). Seed lots weighing 0.45 kg will plant a 0.40 ha seedbed, providing crowns for a two hectare production field (Zandstra et. al., 1992). In Michigan, seeds are planted at a spacing of 5.0 cm and at a depth of 2.5 to 5.0 cm (Zandstra et. al., 1992; Motes et. al., 1996). Rows are spaced 61.0 to 91.0 cm apart with seedlings emerging two to three weeks after planting (Motes et. al., 1996). Irrigation can improve crown size and quality, especially under drought conditions (Wilcox-Lee, 1987). Soil comprised of mostly loam, sandy loam or muck and is well-drained is desirable for a crown nursery (Zandstra et. al., 1992). Asparagus crowns require moderately high levels of nitrogen, phosphorus, and potassium (Zandstra et. al., 1992). Based on soil tests, phosphorus and potassium should be worked into the soil prior to planting. Nitrogen should be applied pre-plant (33.5 kg/Ha) and again when the plants are six to eight inches tall (56 kg/Ha) (Zandstra et. al., 1992; Motes et. al., 1996). Field Selection and Establishment When selecting an asparagus production site, well-drained sandy soils with a pH between 6.2-6.8 and a high organic matter content are preferred (Motes, 1996 et. al.; Sanders, 2001). Asparagus fields can be established with eight- to twelve-week-old greenhouse grown seedlings, one- to three-year-old field grown bare root crowns, or by direct seeding (Mullen et. al., 1998). Asparagus crowns and transplants are planted using similar methods. Furrows are spaced 100 to 180 cm apart using a middle buster plow (Zandstra et. al., 1992). Crowns can be planted in the ftu'row with either a single or double row configuration. In the single row configuration crowns are spaced 20 to 30 cm apart, whereas in the double row configuration crowns are normally planted every fifteen cm in a staggered pattern (Mullen et. al., 1998). Afier the crowns are placed in the furrow, they are covered with 5 to 7.5 cm of soil. Over the course of the summer, the furrow is gradually filled in completely with soil (Zandstra et. al., 1992; Sanders, 2001). Depending on the geographic location, crown planting can occur throughout the year. In Michigan it is important to plant the crowns as early in the spring as possible to ensure the new crowns are well established before winter (Zandstra et. al., 1992). Field Maintenance Once an asparagus field is established, fertilization is needed to optimize asparagus production. When comparing the annual removal of the three most common nutrients, nitrogen is the greatest at 7%, followed by potassium at 5%, and phosphorus at 2% (Drost, 1997). While all of the nutrients are important to asparagus, nitrogen needs to be reapplied yearly. Nitrogen applications afier harvest provide the greatest asparagus yield benefits (Drost, 1997). Although asparagus does not have high requirements for phosphorus and potassium, potassium has been shown to increase asparagus yields (Drost, 1997). Asparagus is a moderately drought tolerant crop (Wilcox-Lee, 1987; Drost, 1997). Irrigation can lessen potential spear damage due to blowing sand and can also slow fern development, thereby increasing spear quality during hot conditions (Drost, 1997). However, too much water during the harvest season is not always beneficial as cool soils can result in slow spear growth. After the harvest period, rain and irrigation during fern growth in July and August increases the carbohydrate reserves in the roots and crown (Drost, 1997; Marr and Tisserat, 1997). Water received during September or later can prompt a new flush of grth which reduces the carbohydrate reserves and can lower yields during the next harvest season (Drost, 1997). In some areas of the world water stress is used to initiate dormancy (Drost, 1997). The health of an asparagus crown is important to productivity. Monitoring the overall health of an asparagus field provides information on the optimal harvest duration and fertilization program (Drost, 1997). A typical way for a grower to monitor the field is a visual assessment of the fern and spears along with a nutrient analysis of the soil and plant tissue. A method to assess plant heath using the carbohydrate levels in the crowns has been developed (Wilson et. al., 2001). Early studies demonstrated that a reduced level of carbohydrate in the crown leads to a decline in productivity (Shelton, 1978). Researchers from New Zealand designed a web-based decision support system for growers (Wilson et. al., 2001) that uses Brix percentages to calculate the levels of available carbohydrate. Growers may use this method to determine harvest duration and assist in nutrient and water management decisions. DISEASES There are several foliar and soil-bome diseases of economic importance that affect asparagus in the United States. In Michigan, the foliar diseases including rust incited by Puccinia asparagi and purple spot incited by Stemphylium vesicarium occur annually (Elmer et. al., 1996; Anonymous, 2000). The soil-bome diseases have historically included F usarium crown and root rot (F usarium spp.). Phytophthora spear rot (Phytophthora megasperma) has been reported in asparagus producing regions (Falloon et. al., 1983; Elmer et. al., 1996; Anonymous, 2000; Elmer, 2001a), including Michigan (Saude et. al., 2005). Viral diseases are also a concern and include Asparagus virus 1 and 2 (Falloon et. al., 1986). Foliar diseases Puccinia asparagi infects asparagus crops in all parts of the world. Rust was first recorded in Europe in 1805 and was first identified in North America in 1896 (Louws et. al., 1994). Initial disease symptoms include light-green oval lesions resulting from infection by basidiospores. One to two weeks later, cream-colored aecia are produced which give rise to the uredinia stage (Elmer, 2001a). Aecia and uredinia spores spread and form new infection sites when weather conditions are wet and windy. Under favorable conditions the uredinia stage can result in new spores produced every 10-14 days (Elmer, 2001a). Teliospores are formed in uredinia lesions when temperatures are low or following extended dry periods (Louws et. al., 1994). Asparagus rust can be controlled using fungicides, resistant cultivars, and good sanitation practices (Louws et. al., 1994; Elmer et. al., 1996; Meyer et. al., 2000; Elmer, 2001a). Purple Spot (Stemphylium vesicarium (Wallr.) E.Simmons teleomorph Pleospora herbarum) was first reported in the United States and New Zealand in the early 19805 (Elmer et.al., 1996). Initial disease symptoms include small, elliptical, slightly sunken, purplish spots on spears. Under favorable environmental conditions (cool, wet weather with wind driven soil), 60 to 90 % of the harvested spears can exhibit purple spot lesions (Hausbeck et. al., 1999; Meyer et. al., 2000; Elmer, 2001a). S. vesicarium also develops on the fern causing tan to brown lesions with dark purple margins (Elmer et. al., 1996; Meyer et. al., 2000). Infection of the fern can cause premature defoliation of the Cladophylls, reducing photosynthetically active tissue and limiting the plant’s ability to replenish carbohydrate reserves for the next growing season (Elmer et. al., 1996; Meyer et. al., 2000). Fungicide applications prompted by a disease forecasting system such as Tom-Cast can effectively control purple spot (Meyer et. al., 2000). Cultural practices, including field maintenance and planting cover crops to reduce tissue damage on the stem from blowing sand, also reduce purple spot disease (Elmer et. al., 1996; Meyer et. al., 2000) Asparagus can also be infected by two viruses, the aphid-borne potyvirus Asparagus virus I and the seed-bome ilarvirus Asparagus virus 11 (Falloon et. al., 1986; Evans and Stephens, 1989). These viruses can be found worldwide. Asparagus virus 11 is considered more detrimental than Asparagus virus I, however, when both viruses are present, susceptibility to F usarium infection tends to be enhanced (Evans and Stephens, 1989). Virus symptoms are not visible on the fern, but overall decreased vigor may be observed (Elmer, 2001a). Soil-borne diseases Phytophthora spp., were first reported as a pathogen to asparagus in California in 1938; however, major impacts on production were not reported until the 19805 (F alloon et. al., 1983; Falloon and Grogan, 1988; Elmer, 2001a). The occurrence of Phytophthora spp. has been reported around the world. Multiple species including P. richardiae, P. megasperma, P. cactorum, and P. megasperma var. soj ae have been identified as causal agents of the disease (F alloon and Grogan, 1988). P. megasperma has been taxonomically reclassified into at least five new species (Hansen and Maxwell, 1991; Cooke et. al., 2000). The newly reclassified Phytophthora spp. demonstrate host specificity to crops such as soybean, clover, alfalfa, apple, cherry, and asparagus (Hansen and Maxwell, 1991; Cooke et. al., 2000). Phytophthora crown and spear rot occurs in warm climates, but can also occur in more northern areas (Elmer, 2001a). Between 2002 and 2005, Canada and Michigan first reported P. megasperma in asparagus fields (V ujanovic et. al., 2003; Saude et. al., 2005). Phytophthora spp. cause rot on the asparagus spears at the soil level. Initially, symptoms include gray-beige to brown water-soaked lesions slightly above or below soil level (Falloon et. al., 1983). Once the lesions begin to expand, the infected area turns light brown and the tissue collapses and shrivels (Elmer, 2001a). The tissue collapse causes the spear to become curved and may kill the spear or cause deformities to the developing fern. Infected crowns may have internal tissue that exhibit a yellow-brown color (Elmer, 2001 a). Phytophthora is found in both newly established fields and in older established 10 fields. Fungicides such as mefenoxam or metalaxyl (trade name Ridomil, manufactured by Syngenta Crop Protection, INC) currently assist in the control of Phytophthora (Falloon et. al., 1983; F alloon and Frasr-Kevern, 1996). Cultivar resistance trials have been conducted in California and New Zealand; no cultivars have shown resistance or tolerance to Phytophthora spp. (Falloon, 1985; Falloon, 1990; Elmer, 2001a). F usarium The genus F usarium includes major crop pathogens that result in yield losses around the world. The F usarium genus was first described 192 years ago by Link and is now one of the most studied fungal genera (Snyder, 1981). F usarium has been recorded on a wide variety of plant species and may have infected plants as long as 200 million years ago (Snyder, 1981). The classification of F usarium has been much debated over the years with at least three different methods used to delineate the F usarium genus. However, the most current classification is based on the research by Wollenweber in 1935 who divided the genus into 16 sections and 65 species, with further divisions into 55 varieties and 22 forms (Messiaen and Cassini, 1981). F. oxysporum is divided into formae speciales to delineate morphologically similar or indistinguishable isolates that cause disease on different plants (Messiaen and Cassini, 1981; Kistler, 1997). The F usarium genus causes two main types of plant disease: cortical rot and vascular wilt (Toussoun, 1981). Originally, F. oxysporum was thought to be strictly a vascular wilt causing organism, however, it has been shown to infect the cortex rather than the vascular tissue (Graham, 1955; Elmer, 2001b). F prolifleratum also causes a 11 wilt, although the mechanism is not fully understood (Guerrero et. al., 1999; Elmer, 2001b) The first reported case of F usarium on asparagus in the United States came from Massachusetts in 1908 (Sherf and MacNab, 1986). Cohen and Heald identified the first F usarium Sp. infecting asparagus in 1941. The formae specialis designation of “asparagz” was applied in 1959 by Grogan and Kimble (Elmer, 2001b). F usarium oxysporum f. sp. asparagi shows some specificity toward asparagus but also causes infection on other plants such as peas, celery, gladiolus, lupine, and onions (Elmer, 2001b) Disease symptoms vary depending on environmental conditions. Disease is usually readily visible in older fields afier harvest when the stalks turn yellow and die (Cohen, 1941; Elmer, 2001b). Symptoms consist of firm, reddish-brown elliptical lesions of varying sizes that form on feeder roots and also on storage or primary roots (Graham, 1955; Grogan and Kimble, 1959; Elmer, 2001b). Crown infection is initiated from root or stem infections. Vigorous crowns that become infected may also exhibit dry, discolored, corky areas that have been compartmentalized by zones of lignified tissue (Elmer, 2001b). These infected areas may become invaded by secondary pathogens that cause the crown to rot. Another significant F usarium pathogen of asparagus is F. proliferatum, which was first described by Snyder and Hansen as F moniliforme (Elmer, 2001b). F. proliferatum has a broad host range that includes crops such as figs, corn, rice, and other grain crops (Damicone et. al., 1988; Anonymous, 2002). F. proliferatum is often associated with wounded tissue and may be more parasitic in nature than F. oxysporum 12 (Graham, 1955; Messiaen and Cassini, 1981). F. proliferatum has been associated with multiple symptoms such as root tip necrosis and a brown, dry crown rot (Johnston et. al., 1979) CONTROL Management of F usarium is complicated by the soil-bome nature of the fimgus. Fungicide applications have proven ineffective or are impractical since the root zone where the fungicide is needed is hard to reach (Elmer, 2001b). Currently, preventive measures are recommended and include site selection, good pest management practices, and cultivar selection. Of all cultivars available, the all-male hybrids have shown some tolerance to F usarium (Ellison and Kinelski, 1985; Sonoda et. al., 2002). Prior to the use of commercial pesticides, rock salt (NaCl) was applied to asparagus fields for weed and disease suppression. This, however, lost favor with the advent of synthetic herbicides in the 1940’s (Elmer et. al., 1996). Recent studies in the laboratory and greenhouse environments demonstrated that NaCl could reduce the effect of F usarium on asparagus (Reid et. al., 2001). In a greenhouse study conducted by Elmer (1992), applications of NaCl increased the fresh weight of seedlings. The same study also demonstrated that NaCl suppressed root disease and the number of F usarium spp. colony forming units recovered per centimeter of root (Elmer, 1992). Elmer (1992) also demonstrated that spring applications of NaCl were capable of suppressing the disease and increasing marketable yields. When NaCl was applied in field situations significant positive results occurred in a severely declined research plot while no effects were noticed in a commercial field situation that was still productive (Reid et. al., 2001). 13 Other potential control methods include fungicide applications, biological agents, resistant varieties and good cultural practices. Fungicides have provided limited management of the pathogen, however, none have shown significant control in the field (Lacy, 1979; Reid, 2000; Elmer, 2001b). The fungicides benomyl, fludioxonil, and thiophanate-methyl have all exhibited activity in greenhouse trials (Reid, 2000). These products have not been tested in a field situation with sufficient disease pressure to determine whether they would be effective in a commercial planting (Reid et. al., 2002). F umigation is widely used on many fresh market crops to reduce pest pressure and enhance overall yield. Currently, fumigation is not used frequently in either asparagus production fields or seedbeds. When fumigation was tested in seedbeds the resulting crowns exhibited increased quality and vigor compared with crowns produced in non-fumigated soil (Lacy, 1979). While fumigation may not be cost effective for use in a production field, it has been shown to be beneficial in the establishment of new production fields (Manning and Vardaro, 1977; Lacy, 1979). Biological fumigants produced from organic matter have reduced diseases associated with apples, asparagus, potatoes, strawberries, and tomatoes (Blok et. al., 2000; Bello et. al., 2001; Mazzola and Mullinix, 2005). These biofumigants are primarily formed from members of the brassica family (Sarwar et. al., 1998; Kirkegaard et. al., 1999; Morra and Kirkegaard, 2002). When incorporated into the soil, these biofumigants release the chemical known as isothiocyanate (ITC), which is similar to metam sodium/potassium (Morra and Kirkegaard, 2002). Trials have shown that proper 14 incorporation followed by tarping can give results equivalent to that of methyl bromide (Bello et. al., 2001). Commercially available biocontrol products including arbuscular mycorrhizae (AM) and T richoderma spp. have been developed for use in other crops and omamentals to improve soil quality and inhibit pathogens (Menge, 1983; Sivan, 1987). Disease management is accomplished through several mechanisms that include competition and induction of a disease defense response in the host (Schenck, 1981; Howell, 2003). AM is best known for enhancing the ability of the plant to increase nutrient and water uptake and increase disease resistance and salt tolerance (Menge, 1983). It has been suggested that AM produces antibiotic compounds, volatile and nonvolatile fungistatic compounds that provide protection (Schenck, 1981). In greenhouse and field trials, AM enhances the asparagus growth and reduces disease (Dehne, 1982; Menge, 1983; Hussey et. al., 1984; Burrows et. al., 1990; Arriola et. al., 2000; Anonymous, 2002; Reid et. al., 2002). The mechanisms used by T richoderma includes mycoparasitism, antibiotic (toxin) production, competition, enzymes, induction of plant defense responses, metabolism of germination stimulants, and additional minor mechanisms (Howell, 2003). Using genetic mutations, known suppression mechanisms were turned off, however, disease suppression still occurred suggesting that there are more mechanisms used in disease suppression that are not defined. In greenhouse tests, Trichoderma was able to improve asparagus quality and also decrease disease severity (Reid et. al., 2002). Nonpathogenic F usarium spp. may suppress disease through mechanical exclusion, localized resistance, competition, or systemic acquired resistance (Damicone 15 and Manning, 1982). He et a1. (2002) used a split-root method to demonstrate that nonpathogenic F usarium strains induced a system disease resistance response in the laboratory or greenhouse. Field results from studies with nonpathogenic F usarium spp. have not been consistent and range from successful (Damicone and Manning, 1982) to ineffective (Damicone and Manning, 1982; Tu et. al., 1990; Reid et. al., 2002) suppression. Cultural practices can help limit F usarium crown and root rot. Selecting a cultivar that is well suited for the particular growing region is important. Today, all-male hybrids are typically used due to enhanced vigor and increased yields compared to the older cultivars. Despite attempts to breed cultivars with resistance to F usarium spp., resistance has not been identified. However, there are some lines that show a greater tolerance to F usarium than others (Stephens et. al., 1989; Dan and Stephens, 1995). This could be due to more vigorous growth making it possible to compensate for loss of root tissue. While A. oflicinalis has not shown any resistance, A. densiflorus (Kunth) Jessop ‘Sprengeri’ and ‘Myersii’ have been found to be more resistant than the other asparagus cultivars (Stephens et. al., 1989) Along with selecting a variety that suits the growing region it is also important to maintain a healthy field and keep stress to a minimum. It is important to maintain proper soil pH and nutrient levels in the soil (Louws et. al., 1994; Elmer, 2001b). By reducing stress on the plant a grower enhances the plant’s ability to ward off F usarium crown and root rot (N igh, 1990; Elmer et. al., 1996). 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Uragarni. 2002. Comparative evaluation of resistance of Asparagus oflicinalis L. cultivars and breeding lines to F usarium stem and crown rot. 10th International Symposium on Asparagus, ActaHorticulturae. 387-390. Stephens, C.T., R.M. DeVries, and KC. Sink. 1989. Evaluation of asparagus species for resistance to F usarium oxysporum f.sp. asparagz' and F. moniliforme. HortScience 24: 365-368. - Toussoun, T.A. 1981. Prologue. F usarium: Diseases, Biology, and Taxonomy. P.E. Nelson, Toussoun, T.A. and Cook, R.J. University Park, Pennsylvania State University Press.11-12. Tu, C.C., Y.H. Cheng, and AS. Cheng. 1990. Recent advance in biological control of F usarium wilt of asparagus in Taiwan. Acta Horticultureae 271: 353. Vujanovic, V., C.Hamel, S. Jabaji-Hare, and M. St-Amaud. 2003. First report of root rot on asparagus caused by Phytophthora megasperma in Canada. Plant Disease 87: 447. Wallingsford, B. 2004. Vegetables. USDA Agriculture Statistics Board. USDA. l4. http://usdamannlib.cornell.edu/reports/rlassr/fruit/pvg- bb/2004/vegeO904.pdf#search='asparagus%20production%203tatistics%202005'. Wilcox-Lee, D. 1987. Soil matric potential, plant water relations, and growth in asparagus. HortScience 22: 22-24. Wilson, D.R., C.G. Cloughley, and SM. Sinton. 2001. AspireNZ: a decision support system for managing root carbohydrate in asparagus. 10th International Conference on Asparagus, Japan, Acta Hort. 51-58. Zandstra, B.H., J .F. Kelly, M.K. Hausbeck, E.J. Grafius, and HG Price. 1992. Commercial vegetable recommendations: Asparagus. Michigan State University Extension. Michigan State University. 23 SECTION I Crown Dip and Drip Applied Fungicides and Biocontrol Agents for Managing Fusarium Crown and Root Rot on Asparagus 24 ABSTRACT F usarium oxysporum f. sp. asparagi is a common pathogen on asparagus (Asparagus oflicinalis) and has been associated with asparagus decline. In many areas, the amount of virgin asparagus land is limited, leading to fields being replanted to asparagus. The replanted fields often have a shortened production life. The objective of this study was to determine whether fungicides applied pre-plant or via drip irrigation during the growing season limit crown and root rot. In 2001, fungicides were applied as a crown dip or as in-furrow applications at planting and included the following: 1) untreated, 2) thiophanate-methyl, 3) benomyl, 4) Trichoderma harzianum, 5) arbuscular mycorrhizae (AM), 6) nonpathogenic F. oxysporum, and 7) fludioxonil (low and high rate). Significant differences (P<0.01) among treatments were observed for plant stand but not fern height (P<0.25). Thiophanate-methyl resulted in the densest plant stand followed by T. harzianum. In 2003, five treatments were investigated: 1) thiophanate- methyl, 2) fludioxonil, 3) nonpathogenic F. oxysporum, 4) T. harzianum, and 5) AM. Each treated plot was paired with an untreated control. Significant differences (P<0.73) in fern height or plant stand were not observed, but trends were noted. While application of fungicides via drip irrigation is promising as a means to transport treatments to the crowns, no significant differences were observed among treatments. A trend was observed whereby applications of nonpathogenic F. oxysporum and T. harzianum resulted in taller plants and higher stand counts when compared to the untreated control. In 2003 a trial was established to investigate 1) thiophanate-methyl, 2) fludioxonil, and 3) nonpathogenic F. oxysporum in combination with different irrigation schedules. Symptoms of Phytophthora megasperma were observed in 2004 following excessive 25 rainfall and mefenoxam was added to a single plant row in each of the irrigation treatments. The addition of mefenoxam significantly reduced the number of plants with Phtyophthara symptoms in 2004 when rainfall was excessive but not in 2005 when rainfall was limited. 26 INTRODUCTION Asparagus growers in Michigan expect fields to be productive for 10 to 15 years. However, Fusarium crown and root rot reduces the longevity of affected fields by 5 to 8 years (Elmer, 2001) and prevents growers from recouping field establishment costs [approximately $3952 US/ha (Dartt et. al., 2002)]. In the United States, F usarium oxysporum f. sp. asparagi and F. proliferatum have been implicated in the disease (Johnston et. al., 1979; Hartung et. al., 1990; Elmer, 2001; Reid et. al., 2002). Since Stone and Chapman’s initial finding (1908) in Massachusetts, Fusarium crown and root rot has been identified in the three major asparagus producing states; California, Washington, and Michigan (Cohen, 1941). Several asparagus cultivars have been screened for resistance to Fusarium crown and root rot but none have been identified (Louws et. al., 1994; He et. al., 2002). All-male hybrids have shown a greater tolerance to the disease than female lines (Stephens et. al., 1989; Dan and Stephens, 1995; Sonoda et. al., 2002) and are used by Michigan growers. Applications of biological or chemical fungicides could be beneficial in limiting Fusarium crown and root rot. The use of nonpathogenic F usarium spp. has been studied as a tool to manage Fusarium crown and root rot in both the laboratory and the field (Damicone and Manning, 1982; Tu et. al., 1990; Elmer, 2001; Reid et. al., 2002). Two commercially available biocontrol fungi, Trichoderma harzianum and arbuscular mycorrhizae (AM), were effective in managing F usarium disease in asparagus seedlings (Burrows et. al., 1990; Arriola et. al., 2000) and tomatoes (Dehne, 1982; Sivan, 1987; Larkin and F ravel, 1998). Trichoderma harzianum may protect plants against pathogens via mycoparasitism, competition with pathogenic fungi, and induction of plant defense 27 responses (Howell, 2003). In contrast, AM forms a mutualistic association with plants, helping the plant absorb nutrients and establish disease resistance (Dehne, 1982; Menge, 1983; Hussey et. al., 1984; Arriola et. al., 2000). There are few studies that have explored the use of fungicides as a means to manage F usarium spp. (Manning and Vardaro, 1977; Lacy, 1979). Benomyl is a member of the benzimidazole group of fungicides and has been helpful in managing F usarium spp. affecting asparagus, narcissus, and cyclamen (Manning and Vardaro, 1977; Hanks, 1996; Elmer and McGovern, 2004). The benzimidazoles also include tlriabendazole and thiophanate methyl. Benomyl and the relatively new fungicide, fludioxonil, increased root weight and decreased root disease when applied to asparagus seedlings inoculated with F usarium spp. in a greenhouse study (Reid et. al., 2002). Studies in field settings using benomyl showed that this fungicide increased fern fresh weight and decreased the incidence of fern yellowing (Manning and Vardaro, 1977; Lacy, 1979). While there was a benefit from using fungicides on crowns the level of protection may have varied based on the amount of F usarium infestation in the soil (Manning and Vardaro, 1977; Lacy, 1979) The objective of this study was to determine if biological or chemical treatments limit Fusarium crown and root rot of asparagus when applied in a commercial field setting to the crowns at planting or through a drip irrigation system. 28 MATERIALS AND METHODS All studies were conducted on commercial farms in Oceana County, Michigan, on one of three soil types: Perrinton loarn/Bono silt loam, Benona sand, or Spinks-Benona complex all at a zero to six percent slope. All locations had a history of asparagus production and plots were maintained according to commercial production standards. Fertilizer was applied at planting (DAP N 18-P205 46-K20 0; 224 kg/ha) and was side- dressed post harvest each subsequent year (N 6- P205 24- K20 24 and N 46- P205 0- K20 0; 224.3 kg/ha). Fern maintenance consisted of insecticide applications including diazinon (Diazinon AG500, Micro Flo Company LLC, Memphis, TN; 1.7 L/ha) and carbaryl (Sevin XLR Plus, Bayer CropScience, Research Triangle Park, NC; 3.4 L/ha) every fourteen days to control asparagus beetle (Crioceris asparagi) and asparagus miner (Ophiomyia simplex). The fungicides chlorothalonil (Bravo WeatherStik, Syngenta Crop Protection, Greensboro, NC; 3.4 L/ha) and tebuconazole (Folicur 3.6F, Bayer CropScience; 437.2 ml/ha) were applied every fourteen days for the management of asparagus rust (Puccinia asparagi) and purple spot (Stemphylium vesicarium). Weed control consisted of a pre-emergent application of metribuzin (Sencor DF 75%, Bayer CropScience; year 1, 560.2 g/ha; year 2, 1.5 kg/ha), diuron (Direx 80DF, Griffin LLC,Valdosta, GA; 1.1 kg/ha), and glyphosate (Roundup, Monsanto Company, St. Louis, M0; 3.4 L/ha) on 30 May 2003 and 20 April 2004, and sprays of halosulfuron-methyl (Sandea, Gowan Company, Yuma, AZ; 69.9 g/ha) on 17 June and 15 July 2003 and 22 June and 14 July 2004, clethodim (Select 2EC, Valent, Walnut Creek, CA; 584.4 ml/ha) on 22 June 2004 and fluazifop-P-butyl (Fusilade DX, Syngenta Crop Protection; 22.2 ml/3.8 L) on 15 July 2003 and 14 July 2004. Average monthly minimum and maximum 29 air temperatures and total rainfall during the growing season were obtained from the Michigan Automated Weather Network (www.agweather.geo.msu.edu/mawn/, Table 1). Table 1. Average monthly maximum and minimum temperatures and total monthly rainfall during the 2001 through 2005 growing seasons. 2001 2002 2003 2004 2005 Month Maxy Miny Rainz Max Min Rain Max Min Rain Max Min Rain Max Min Rain April 15.3 3.4 79 12.41.8 84 12.5 0.3 36 13.71.2 58 15.9 2.3 18 May 19.8 8.4 183 15.9 4.8 81 18.1 6.0 89 18.8 7.6 246 17.5 6.1 51 June 23.7 12.0 41 24213.5 74 23.7105 53 22.712.1 79 27.915.8 25 July 27.6 14.5 41 29.1 16.9 61 26.313.9 48 24.912.9 36 27.814.7 109 August 27.0 15.5 127 26.614.6 76 27.315.1 66 23511.2 53 26.3145 79 September 20.4 9.7 81 24.611.4 25 22.0 9.9 51 25210.4 15 24511.6 66 October 14.5 4.6 140 12.2 3.0 74 14.6 3.8 84 15.5 4.6 102 16.5 5.2 11 Average 21.2 9.7 98.9 20.7 9.4 67.9 20.6 8.5 61 20.6 8.6 84.] 22.3100 51.3 y Reported in degrees Celsius. 2 Reported in millimeters. Crown Treatments Crown Trial 1 was planted on 2 May 2001 using one-year-old ‘Jersey Supreme’ crowns obtained from a commercial grower. Products were applied directly to the crowns by dipping them prior to planting, or applied at planting as an in-furrow drench or as an in-furrow broadcast treatment. The following materials were applied to the crowns by dipping them in the solutions for 2 minutes: T. harzianum Rifai strain KRL-AGZ PlantShield HC; Bioworks, Inc., Geneva, NY) at 907.2 g/37.8 L; benomyl (Benlate SP; 30 Dupont, Wilmington, DE) at 22.6 g/37.8L; and thiophanate methyl (Topsin M 70WP; Cerexagri Inc., King of Prussia, PA) at 31.8 g/37.8 L. Fludioxonil (Medallion, Syngenta Crop Protection, Inc.) at 1.42 g/37.8 L (low rate) and 76.2 kg/ha (high rate) was applied as an in-furrow drench. Products applied as an in-furrow broadcast treatment included nonpathogenic F oxysporum at 427 kg/ha and AM (Micronized endo; Mycorrhizal Applications Inc., www.mycorrhizae.com) at 1.2 ml/plant. Nonpathogenic F. oxysporum was grown on sterile millet or in sterile asparagus broth. F. oxysporum strains F-21 and DI were used and provided protection against virulent strains of F. oxysporum f.sp. asparagi in a previous study (Reid, 2000). Strains are maintained in Dr. M. Hausbeck’s laboratory at Michigan State University. The nonpathogenic F. oxysporum strains were grown on carnation leaf agar (CLA) (Fisher et.al, 1982) at 20 to 25°C for 14 days prior to inoculation of the millet. To produce millet inoculum, the grain was mixed with water at a 10:1 ratio in a 4-L plastic jug and autoclaved for one hour. Following cooling, the mixture was inoculated with four, 7-mm mycelial plugs of each strain. Inoculated millet was incubated for 14 days at 20 to 25°C on a laboratory bench, mixed, and then transferred to brown paper bags and incubated at 20 to 25°C for 7 days. To produce asparagus broth, asparagus spears were cut into 3-cm cubes and blended in distilled water (1:1 ratio) for 45 seconds using a Waring blender (Waring Laboratory, CT). Fibrous asparagus material was strained out using a single layer of cheesecloth and the broth was diluted to a 2:1 ratio and autoclaved for 30 minutes. Broth was amended with 200 uL of 100-ppm ampicillin and inoculated with four, 7-mm mycelial plugs of F. oxysporum (either strain D-1 or F-21) per 500 ml of broth. Inoculated broth was incubated for one week at 20 to 25°C on a laboratory bench. 31 After incubation the mycelia were removed via straining using a single layer of cheesecloth. The broth of the F usarium strains was mixed in a 1 :1 ratio in a 4-L flask and then stored at 4-6° C until application. Treatments in Crown Trial 1 were replicated six times in a randomized complete block design with five, 12-m rows per treatment. Crown spacing within the row was approximately 19 cm and spacing between rows was 1.5 m. A 3-m fallow buffer was left between treatment blocks. On 10 September 2002, 16 September 2003, and 3 August 2004 fern height and plant stand counts were recorded from the center 6.1 m of the middle row of each block. Crown Trial 2 was planted on 22 and 23 April 2003 using one-year-old commercially produced ‘Jersey Giant’ crowns of a medium size that had been graded for uniformity. Crowns were planted every 19 cm in 12-m rows with 1.2 m between rows. Blocks consisted of 3 treated rows separated from 3 untreated rows with a 2-m fallow buffer (Figure 1). Treatment blocks were replicated six times in a randomized complete block design and the pairing of the treated and untreated plots was also randomized. Four of the products were applied by dipping the crowns for 2 minutes and included the following: AM (Micronized endo) at 59.5 g/37.8 L; T. harzianum Rifai strain KRL-AG2 (PlantShield HC) at 910 g/3 7.8 L; fludioxonil (Scholar; Syngenta Crop Protection, Inc.) at 22.7 g/37.8 L; and thiophanate-methyl (Topsin M 70WP) at 44.8 g/37.8 L. Millet inoculated with nonpathogenic F oxysporum was produced as previously described and was applied to the crowns at planting by broadcasting it in the furrow (397 g/12 row-m). A hand spreader (EarthWay ev-n-spread model 3500; Bristol, 32 IN) was used to broadcast the inoculum. F em height and stand counts were recorded on 11 June, 9 July, and 12 August 2003; 18 June, 15 July, and 20 August 2004; and 6 July and 10 August 2005 from the center 6.1 m of each row. Treated Untreated 7-day 30-day Figure 1. Layout of a single treatment plot in Crown Trial 2 (A) and Irrigation Trial 2 (B) depicting which rows received treatment and the application interval. 33 Drip Irrigation Treatments Irrigation Trial 1 was planted on 2 May 2001 using one-year-old commercially produced ‘Jersey Supreme’ crowns using a within-row spacing of approximately 19 cm. The plot was set up in a randomized complete block design with three replicates, with a single 30.5-m row per treatment in each replicate. There was a 3-m fallow buffer between rows and a 6-m fallow buffer between blocks. Fungicides and biocontrol products were applied at planting and reapplied every 30 days for a total of four applications per year. Products included fludioxonil (Scholar) at 9.9 g/30.5 m, benomyl (Benlate SP; Dupont, Wilmington, DE) at 5.67 g/30.5 m, nonpathogenic F. oxysporum at 20 kg/30.5 m (planting) and 993.3 ml/30.5 m (every 30 days), and T. harzianum Rifai strain KRL-AG2 (PlantShield HC) at 55.6 g/30.5 m. The nonpathogenic F. oxysporum was applied using inoculated millet (planting) or asparagus broth (post-planting applications) and was prepared as previously described. In 2001, treatments were initiated on 27 June and then applied every 30 days (3x in 2001, 4x in 2002, and 4x in 2003) through a drip irrigation system. The irrigation system consisted of a single drip line directly above the crowns with spring-loaded check valves in line to prevent treatment material from back flowing. Treatments were premixed with 7.6 L of water in a stainless steel container and then injected into the irrigation lines with a container operating pressure of 172 to 207 kPa and a line pressure of 83 to 103 kPa. The application occurred over a period of 60 minutes with the suspensions or solutions pulsed into the lines every 15 minutes. No applications were made in 2004. Fern height and plant stand count were taken from the middle 6.1 m of each treatment row on 12 September 2002, 23 September 2003, and 3 August 2004. 34 Irrigation Trial 2 was planted on 13 May 2003 using one-year-old commercially produced ‘Jersey Giant’ crowns that were selected as previously described. Crowns were planted in treatment plots consisting of five, 12.2-m long rows with a crown spacing of 19 cm and a between row spacing of 1.5 m. Treatment plots included the following: non- irrigated, irrigated (7- or 30-day intervals), nonpathogenic F. oxysporum at 9.8 L/305 m, fludioxonil (Scholar) at 22.7 g/37.8L, and thiophanate-methyl (Topsin M 70WP) at 560.2 g/ha. Irrigation was applied to two rows of each plot every 7 days or 30 days with treatments applied to all rows every 30 days (Figure 1). The center row of each treatment block did not receive any treatment and a 2.7-m fallow buffer was established between treatment plots. Treatments were replicated four times in a split plot design. Initial applications of the products were made at planting as an in-furrow drench using 8-L watering cans and as an irrigation injection every 30 days afier fern establishment. A total of three applications via irrigation were made during each of the 2003, 2004, and 2005 growing seasons. The irrigation system consisted of two drip lines 25.5 cm apart, 5 cm above the crowns, with spring-loaded check valves in line to prevent treatment material from back flowing. Treatments were premixed with 7.6 L of water in a stainless steel container and then injected into the irrigation lines with a container operating pressure of 172 to 207 kPa and a line pressure of 83 to 103 kPa. The application occurred over a period of 60 minutes with the suspensions or solutions pulsed into the lines every 15 minutes. In 2004 and 2005, mefenoxam (UltraFlourish; Nufarm Americas Inc., Burr Ridge, IL; 4.7 L/ha) was applied every 30 days (a total of 4 applications) to one row in the 7- and 30-day irrigation treatments. Evaluations were taken on 2 September 2003, 3 August 2004, and 24 August 2005 from the middle 6.] m 35 of row and included counting the plants exhibiting disease symptoms, number of living fern, number of dead fern, number of emerging spears, and the total number of fern. Statistical Analysis Data from all trials were analyzed using Statistical Analysis Software (SAS v.8.2). Analysis for Crown Trial 1 and Irrigation Trial 1 were conducted using the ‘Proc Mixed’ procedure and means were compared with Tukey’s HSD (P3005) with treatment being a fixed effect and replicate being a random effect. For Crown Trial 2, the analyses for counts were based on the analysis of a split plot design with treatments being the whole-plot factor and application (treated vs. untreated) the sub-plot factor. Rows were considered to represent sub-sampling so a random factor of treatment*replicate*application was included, leaving the row effect to be residual variance. The analysis was conducted using the ‘Proc Mixed’ procedure with treatment, application, and treatment*application being fixed effects, and replicate, treatment*replicate, and treatment"replicate*application being random effects. The analysis of the rating parameters for Irrigation Trial 2 was based on the analysis of a split plot design in which treatment was the whole-plot factor and row was the sub-plot factor. In doing the analysis, row effects (sub-plot factor) were coded as follows: for treated plots row 1 was 7r, row 2 was 7, row 3 was un, row 4 was 30, and row 5 was 30r; for the untreated plots rows were left as they occurred in the plot. If row differences were significant the rows were compared using the following contrasts: 1) treated compared to untreated; 2) irrigation every 7 days compared to irrigation every 30 days; 3) rows near the untreated control row compared to rows that were far from the untreated control row; and 4) the interaction of irrigation frequency and distance from the center untreated 36 control. All analyses were conducted using ‘Proc Mixed’ procedure where the treatment and row were included as fixed factorial factors and replicate and replicate*treatment interactions were included as random effects. 37 RESULTS In Crown Trial 1, disease pressure from F usarium spp. was severe. From 2003 to 2004, stand count and fern height in the untreated control was reduced by 50% and 24%, respectively. The plant stand resulting from asparagus crowns treated with fungicides or a biocontrol agent did not differ significantly from the untreated control in 2003 (Table 2). However, the stand count from the T. harzianum-treated asparagus crowns was significantly greater than that of the fludioxonil-treated crowns. In 2004, asparagus crowns treated with thiophanate-methyl produced a significantly better stand than the untreated and all other treatments with the exception of T. harzianum. F em height provided a means of assessing plant vigor and varied significantly among treatments in both years (Table 2). In 2003, asparagus treated with thiophanate-methyl had fern that was significantly taller than all other treatments, but was not significantly different from the untreated control. The asparagus fern in 2004 was tallest when crowns were treated with thiophanate-methyl, but did not differ significantly from the untreated or the T. harzianum and AM treatments. In Crown Trial 2, asparagus crowns that were treated did not differ significantly from the untreated in either plant stand or fern height (Table 3). A trend was noted whereby the asparagus crowns treated with AM produced a plant stand that was usually increased in number and height compared to the untreated. In the spring of 2004, disease symptoms other than those caused by F usarium spp. were noted on emerging spears and included distortion and necrosis. Other symptoms noted included a shepherd’s crook of the fern, shriveling of the spears, and water soaking of the spears or stem at the soil line (Figure 2). 38 Table 2. Average stand count and fern height for asparagus crowns treated at planting with fungicides or biocontrol agents in Crown Trial 1. Stand countI Fern height (cm)I Treatment and rate 2003 2004 2003 2004 Untreated ..................................................... 84.8 ab2 42.2 bc 105.4 ab2 80.8 ab Thiophanate-methyl 31.8 g/37.8 L .............. 93.0 ab 53.0 a 131.8a 109.7a T. harzianam 907.2 g/37.8 L ....................... 87.2 a 46.2 ab 100.1 be 83.8 ab Arbuscular mycorrhizae 1.2 mllplant .......... 83.3 ab 39.2 be 99.3 be 70.1 ab Benomyl 22.6 g/37.8 L ................................ 81.3 ab 39.2 be 88.1 be 66.8 b Fludioxonil 38.2 g/ha .................................. 77.0 ab 39.2 be 87.9 be 64.3 b Nonpathogenic F. oxysporum 427 kg/ha ....76.3 ab 36.2 c 80.0 be 61.0 b Fludioxonill.42g/37.8L ........................... 71.5 b 35.8 c 73.1 c 59.2 b TRatings were taken from the center 6.1 m of row on 16 September 2003 and 3 August 2004. 2Treatments with the same letter are not significantly different (Tukey’s HSD P<0.05). 39 Table 3. Average stand count and fern height for asparagus treated at planting with fungicides or biocontrol agents in Crown Trial 2. Stand countT Treatment and rate2 2003 2004 2005 Nonpathogenic F. oxysporum 397 g/12 m ........ 131.1 (-2.7)3 75.3(-7.9) 68.2(-3.8) Arbuscular mycorrhizae 59.5 g/37.8 L .............. 132.4(12.4) 77.6(4.6) 68.7(1.7) T. harzianam 910 g/37.8 L ................................ 124.3(1.2) 71.3(0.7) 65.0(2.0) Fludioxoni122.7 g/37.8 L ................................. 122.7(-5.6) 75.9(-8.7) 71.1 (-1.85) Thiophanate-methyl 44.8 g/37.8 L .................... 126.5(-4.3) 69.8(-3.8) 67.8(0.3) F em height (cm)I Nonpathogenic F. oxysporum 397 g/ 12 m ........ 53.3(2.3)3 122.9(-2.3) 90.5(-0.8) Arbuscular mycorrhizae 59.5 g/37.8 L ............. 50.8(1.8) 106.2(1.3) 80.6(6.9) T. harzianam 910 g/37.8 L ................................ 49.8(-0.8) 106.4(-3.6) 84.6(5.0) Fludioxonil 22.7 g/37.8 L ................................. 48.5(0.2) 108.7(0.9) 84.5(-7.1) Thiophanate-methyl 44.8 g/37.8 L .................... 55.6(4.6) 114.8(-2.6) 85.1(0.4) IRatings taken from the center 6.1 m of row on 12 August 2003, 20 August 2004, and 10 August 2005. 2Treatments were not significantly different at P>0.05 (Tukey’s HSD). 3Numbers in parentheses represent the difference between the treatment and the untreated control. 40 Figure 2. Common Phytophthora symptoms found on asparagus: A) shepherd’s crooking of the fern, B) water soaking and necrosis of the crown and roots, C) water-soaked lesion on the spear found at the soil line, and D) crooking and water soaking observed on the spears. 41 In Irrigation Trial 1, treatments did not differ significantly from the untreated control or from each other (Table 4). However, treatment with nonpathogenic F. oxysporum showed a trend of increased plant stand and fern height compared to all other treatments and the untreated control. In Irrigation Trial 2, disease symptoms other than those caused by F usarium spp. and similar to those observed in Crown Trial 2 were observed following heavy rains. Symptoms included curving of emerging spears or fern tops, shriveling of the stalk, and water-soaked lesions occurring slightly above or below the soil line (Figure 2). These symptoms were significantly reduced (> 28%) on asparagus treated with mefenoxam (Table 5). Based on fungal isolations from spears harvested in the spring, it was concluded that the observed disease symptoms on both spears and fern (Figure 2) were caused by Phytophthora megasperma (Saude et. al., 2005). In 2003, plant rows receiving only irrigation had significantly more fern than rows treated with thiophanate-methyl or fludioxonil (Table 6). However, in 2004 and 2005 there were no significant differences among any of the treatments. In 2005, mefenoxam treated rows had significantly more fern than plant rows that were not irrigated or irrigated but not treated with mefenoxam (Table 7). Maximum and minimum temperatures were approximately normal for all years based on the 30-year averages reported by the Michigan State Climatologist. Precipitation amounts varied among years with 2001 and 2004 receiving above normal precipitation in May, followed by below normal precipitation. Compared to the 30-year average, the precipitation in May was approximately 106 mm and 170 mm above normal in 2001 and 2004, respectively. 42 Table 4. Average stand count and fern height for asparagus plants treated with fungicides or biocontrol agents at planting and during the season in Irrigation Trial 1. Stand countl F ern height (cm)l Treatment and rate 2003 2004 2003 2004 Untreated2 ..................................................................... 83.03 54.0 90.2 89.7 Nonpathogenic F. oxysporum 993.3 ml/30.5 m4 .......... 85.3 68.0 121.6 121.4 Fludioxonil 9.9 g/30.5 m .............................................. 80.0 49.3 86.6 86.6 T. harzianam 55.6 g/30.5 m ......................................... 73.3 59.0 103.1 102.6 Benomyl 5.67 g/30.5 m ................................................ 72.7 51.3 91.2 89.7 lRatings were taken from the center 6.1 m of row on 23 September 2003 and 3 August 2004. 2Irrigation was applied every 30 days for 1 hour. 3 Treatments were not significantly different at P>0.05 (Tukey’s HSD). 4Treatments were applied every 30 days during irrigation. 43 Table 5. Average stand count of plants with Phytophthora symptoms when asparagus plants were irrigated and/or treated with fungicides during the growing season in Irrigation Trial 2. No. plants with Phytophthora symptoms/6.1 m Treatment and rate 2004 2005 Untreated .................................................. 35.4 b1 9.6 a 30-day treatments2 .................................... 36.2 b 12.9 a 7-day treatments3 ...................................... 29.7 b 16.3 a 30-day treatments + mefenoxam4 ............ 25.6 a 14.9 a 7-day treatments + mefenoxam ................ 24.1 a 14.3 a lTreatments with the same letter are not significantly different (Tukey’s HSD P<0.05). 23 O-day treatments were irrigated every 30-days and received a F usarium treatment every 30 days. 37-day treatments were irrigated every 7 days and received a F usarium treatment every 30 days. 4Mefenoxam was applied every 30 days to select rows of both 7 and 30 day F usarium treatments. 44 Table 6. Average stand count of all plants when asparagus plants were treated with fungicides or biocontrol agents at planting and during the season in Irrigation Trial 2. Total fern Treatment and rate 2003 2004 2005 Untreated ................................................................. 217.5 abl 164.4 a 224.8 a Irrigated2 .................................................................. 225.9 a 184.7 a 242.3 a Non-Pathogenic F usarium 9.8 L/305 m 3 ............... 207.9 ab 171.1 a 234.2 a Topsin 70WSB 560.2 g/Ha ..................................... 200.4 b 167.8 a 233.1 a Scholar 50WP 11.3 g/37.8 L ................................... 198.7 b 165.2 a 233.6 a lTreatments with the same letter are not significantly different (Tukey’s HSD P<0.05). 2Irrigation was applied every 7 or 30 days. 3 Treatments were applied every 30 days. 45 Table 7. Average stand count of total fern when asparagus plants were treated with the fungicide mefenoxam or not treated in Irrigation Trial 2. Total fern Treatments 2004 200 5 Untreated ...................................... 163.8 a‘ 199.6 a No Mefenoxam ............................ 172.5 a 226.9 a Mefenoxam2 ................................. 1 71 .8 a ' 249.0 b lTreatments with the same letter are not significantly different (Tukey’s HSD P<0.05). 2Mefenoxam was applied every 30 days. 46 DISCUSSION Michigan growers prefer virgin planting sites for the establishment of new asparagus production fields but are often forced to utilize fields that have previously hosted asparagus. Such replanted fields typically exhibit yield decline and Fusarium crown and root rot. The perennial nature of asparagus and its relatively deep root and crown system increases the complexity of implementing potential control measures for F usarium spp. Utilizing a crown dip at planting offers the easiest and least costly means of managing F usarium crown and root rot. However, the crown dip treatments included in our trials did not consistently produce results that were significantly different from the untreated control. In Crown Trial 1, treating asparagus crowns with thiophanate-methyl appeared to be helpful, whereas using the related fungicide benomyl was not. In Crown Trial 2, the inclusion of thiophanate-methyl produced results that were not comparable to those observed in Crown Trial 1. Possible reasons for the differences observed between these trials may include pathogen pressure at the two sites and the overall duration of the trials. For instance, treatments included in Crown Trial 1 were tested under extremely high Fuarium crown and root rot disease pressure that had resulted from three crops of asparagus previously grown at that site. Crown Trial 2 was conducted in a field that had previously hosted asparagus once. Also, disease symptoms in Crown Trial 2 were consistent with those caused by P. megasperma and F usarium spp. in 2004, indicating that this trial was confounded by a non-target pathogen that would not be limited by the fungicide treatments included. Because of the non-target pathogen, Crown Trial 2 was terminated afier only three years. Disease symptoms from Crown Trial 1 were consistent with F usarium crown and root rot and became especially severe in year 4 of the study. 47 This is consistent with grower reports and indicates that a minimum of 4 years is needed to assess treatment efficacy. Lacy (1979) had earlier explored the practice of dipping asparagus crowns in fungicide solutions. He determined that treating asymptomatic asparagus crowns grown in infested soil with fungicides, including benomyl and thiophanate-methyl, prior to planting in non-infested soil did not improve crown survival or yield compared to the untreated control. He attributed this lack of treatment benefit to naturally low disease pressure in his trial site and other cultural factors, including the lack of stressful growing conditions for the crowns that could exacerbate Fusarium crown and root rot. Manning and Vardaro (1977) determined that combining soil fumigation with pre-plant crown soaks with benomyl resulted in vigorous growth of a new asparagus planting in a F usarium-infested site. When potential F usarium-controlling products were applied via drip irrigation, the benefits were not conclusive. In Irrigation Trial 1, differences among the treatments and the untreated were not observed and may be attributed to ineffective product delivery (only a single drip line was used) and overall low disease pressure. However, Irrigation Trial 2 offered an interesting insight into the replant problem that some Michigan asparagus growers are experiencing. In particular, as noted in Crown Trial 2 and Irrigation Trial 2, disease symptoms observed included those caused by P. megasperma. Prior to 2004, this disease had not been noted to occur in Michigan. The addition of mefenoxam in the second treatment year significantly reduced symptoms of P. megasperma and in the third year increased the total number of fern stalks, verifying that applying products via drip irrigation can be effective. In this trial, two drip irrigation 48 tapes were used to deliver the products and applications were made throughout each fem- growing season. Asparagus yields can be reduced by at least 50% if Phytophthora is not managed (F alloon, 1990). Studies using metalaxyl, the precursor to mefenoxam, as a pre-plant crown dip or in-furrow spray significantly increased yields (F alloon et. al., 1985; Falloon et. al., 1991; Falloon and Fraser, 1991; Falloon and Fraser-Kevern, 1996). Of the two application methods, Falloon et. a1. (1991) reported that the crown dips provided better protection against Phytophthora infection than did the in-furrow sprays. In established fields Falloon et.al. (1985) found that banded sprays over the asparagus row significantly reduced the incidence of spear rot caused by Phytophthora. While Phytophthora can greatly reduce yields, the fungicide mefenoxam can limit the pathogen. Growers in Michigan do not use drip tape to produce asparagus, however, it offers a potentially effective and efficient means to deliver disease-controlling products and/or fertilizer directly to the crowns, as well as a way to provide water during periods of drought. Drip irrigation has proven to be effective in Peru, where the majority of the asparagus is grown in sandy soil that receives very little moisture (Casas, 2005). Unlike the northern production regions, were temperatures become low enough to initiate dormancy in asparagus, the southern production regions rely on drought stress, which makes irrigation an important component in Peruvian asparagus production. In European countries drip irrigation has been shown to increase overall asparagus yields (Panka and Rolbiecki, 2005). The use of sub-surface irrigation has been shown to increase asparagus yield and enhance field establishment (Wilcox-Lee, 1987; Sterrett et. al., 1989). Since low water potentials favor F usarium spp. (Cook, 1981), applying additional water to 49 established fields may reduce crown stress by keeping soil moisture at a level that is less favorable for F usarium development. 50 LITERATURE CITED Arriola, L.L., M.K. Hausbeck, J. Rogers, and GR. Safir. 2000. The effect of Trichoderma Hazianum and arbuscular mycorrizae on F usarium crown and root rot in asparagus. HortTechnology 60: 141-144. Burrows, R., F.L. Pfleger, and L. 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Grogan. 1991. Individual and combined effects of flooding, Phytophthora rot, and metalaxyl on asparagus establishment. Plant Disease 75: 514-518. Falloon, P.G., and HA. Fraser. 1991. Control of establishment failures in asparagus (Asparagus aflicinalis L.) caused by Phytophthora rot. New Zealand Journal of Crop and Horticultural Science 19: 47-52. Falloon, P.G., and HA. Fraser-Kevem. 1996. Effect of thiabendazole and metalaxyl on asparagus establishment in replant soil. VIII International Symposium on Asparagus, Acta Hort. 289-295. Fisher, N.L., L.W. Burgess, T.A. Toussoun, and RE. Nelson. 1982. Carnation leaves as a substrate and for preserving cultures of F usarium species. Phytopathology 72: 151- 153. Hanks, G.R. 1996. Control of F usarium oxysporum f.sp. narcissi, the cause of narcissus basal rot, with thiabendazole and other fungicides. Crop Protection 15: 549-558. Hartung, A.C., C.T. Stephens, and W.H. Elmer. 1990. Survey of F usarium populations in Michigan's asparagus fields. Proceedings of the 7th International Asparagus Symposium, Ferrara, Italy, Acta Horticulturae. 395-401. He, C.Y., T. Hsiang, and DJ. Wolyn. 2002. Induction of systemic disease resistance and pathogen defence responses in Asparagus oflicinalis inoculated with nonpathogenic strains of F usarium oxysporum. Plant Pathology 51: 225-230. 52 Howell, CR. 2003. Mechanisms employed by Trichoderma species in the biological control of plant diseases: the history and evolution of current concepts. Plant Disease 87 : 4-10. Hussey, R.B., R.L. Peterson, and H. Tiessen. 1984. Interactions between vesicular- arbuscular mycorrhizal fungi and asparagus. Plant and Soil 79: 409-416. Johnston, S.A., J.K. Springer, and GD. Lewis. 1979. F usarium moniliforme as a cause of stem and crown rot of asparagus and its association with asparagus decline. Phytopathology 69:778-780. Lacy, M.L. 1979. Effects of chemicals on stand establishment and yields of asparagus. Plant Disease Reporter 63: 612-616. Larkin, R.P., and DR. Fravel. 1998. Efficacy of various fungal and bacterical biocontrol organisms for control of F usarium wilt of tomato. Plant Disease 82: 1022-1028. Louws, F.J., P.R. Ragan, R.S. Vernon, A.R. Forbes, and J .A. Garland. 1994. Asparagus. Diseases and Pests of Vegetable Crops in Canada. R.J. Howard, Garland, J.A. and Seaman, W.L., Canadian Phytopathological Society and Entomology Society of Canada.43-49. Manning, W.J., and PM. Vardaro. 1977. Soil fumigation and preplant fungicide crown soaks: effects on plant growth and F usarium incidence in newly planted asparagus. Plant Disease Reporter 61: 355-357. Menge, J .A. 1983. Utilization of vesicular-arbuscular mycorrhizal fungi in agriculture. Canadian Journal of Botany 61: 1015-1024. Panka, D., and R. Rolbiecki. 2005. Health status of surface drip irrigated asparagus cultivars in central Poland conditions. 11th International Asparagus Symposium, Horst, Netherlands, Acta Hort. In Press. Reid, TC. 2000. Evaluation of strategies for the control of F usarium crown and root rot of asparagus. Plant Pathology. East Lansing, Michigan State University. Masters:131. Reid, T.C., M.K. Hausbeck, and K. Kizilkaya. 2002. Use of fungicides and biological controls in the suppression of F usarium crown and root rot of asparagus under greenhouse and growth chamber conditions. Plant Disease 86: 493-498. 53 Saude, C., M.K. Hausbeck, O. Hurtado-Gonzales, and K.H. Larnour. 2005. Detection of a Phytophthora sp. causing asparagus spear and root rot in Michigan. Plant Disease 89: 1011. Sivan, A. 1987. Biological control of F usarium crown rot of tomato by Trichoderma harzianum under field conditions. Plant Disease 71: 587-592. Sonoda, T., K. Tairako, and A. Uragami. 2002. Comparative evaluation of resistance of Asparagus oflicinalis L. cultivars and breeding lines to F usarium stem and crown rot. 10th International Symposium on Asparagus, ActaHorticulturae. 387-390. Stephens, C.T., R.M. DeVries, and KC. Sink. 1989. Evaluation of asparagus species for resistance to F usarium oxysporum f.sp. asparagi and F. moniliforme. HortScience 24: 365-3 68. Sterrett, S.B., B.B. Ross, and CR Savage Jr. 1989. Influence of irrigation method on establishment and yield of aspragus crowns and transplants. The Vegetable Growers News 44: 1-2. Tu, C.C., Y.H. Cheng, and AS. Cheng. 1990. Recent advance in biological control of F usarium wilt of asparagus in Taiwan. Acta Horticultureae 271: 353. Wilcox-Lee, D. 1987. Soil matric potential, plant water realtions, and grth in asparagus. HortScience 22: 22-24. 54 SECTION 11 Screening Commercially Available Asparagus Cultivars for Susceptibility to F usarium spp. in a Controlled Environment. 55 ABSTRACT Asparagus decline is caused by F usarr'um oxysporum f. sp. asparagi and F. proliferatum and reduces field productivity. These pathogens are long lived in the soil and limit the effectiveness of standard cultural and chemical strategies. The identification and/or development of resistant or tolerant asparagus could reduce losses resulting from F usarium infection. The objective of this research is to determine whether any commercially available cultivars exhibit reduced susceptibility to F usarium. Seeds of twenty cultivars were obtained and cleaned in a benomyl and acetone solution. Seeds were germinated and transferred to growth tubes containing Hoagland’s media, which were then inoculated with F oxysporum f.sp. asparagi and F. proliferatum after 14 days. Twenty-eight days after inoculation seedlings were rated on a scale of one (healthy, no disease) to six (dead). Ratings varied between trials and isolates had average ratings ranging from 2.5 to 5.6. Significant differences occurred among cultivars (p=0.05). However no cultivar consistently received a rating that would be considered F usarium resistant. 56 INTRODUCTION The asparagus industry has experienced a reduction in the production life of asparagus fields from a maximum of 15 years down to 5 to 8 years (Elmer, 2001). Asparagus decline has been attributed to Fusarium crown and root rot incited by F usarium oxysporum f.sp. asparagi and F usarium proliferatum. Since Stone and Chapman (1908) first reported F usarium spp. on asparagus in Massachusetts, F. oxysporum f.sp. asparagi and F. proliferatum have been documented in the three main asparagus producing states; California, Michigan, and Washington (Cohen, 1941). Chemical controls, including fumigants, have been tested for efficacy against F usarium spp. Fungicides have been applied to seed before planting, to the crowns at planting, and as soil drenches after the crowns have been planted (Manning and Vardaro, 1977; Lacy, 1979; Damicone et.al., 1981). Since asparagus is a deep-rooted perennial crop, the root zone is extensive, making it difficult to reach the target area with fungicide applications (Elmer, 2001). Lacy (1979) used seed treatments and crown dips but significant benefits were not observed. When crowns were grown in fumigated soil and then planted in a fumigated production field, significantly higher yields were observed compared to planting crowns in non-fumigated soil. When Manning and Vardaro (1977) combined firmigation with a crown dip in the fungicide benomyl the resulting planting had large vigorous crowns in a field known to be infested with F usarium spp. Damicone et.al. (1981) used benomyl in combination with a dilutent to eradicate F moniliforme and F. oxysporum from asparagus seed. They found that when diluting benomyl in acetone the number of F usarium colonies were significantly reduced. Studies have demonstrated that biological controls such as T richoa'erma and AM can increase initial growth 57 (Burrows et.al., 1990) and reduce incidence of Fusarium crown and root rot on seedlings (Arriola, 1997). While applications of biological and chemical fungicides to asparagus crowns and seeds may help limit the effects of F usarium (Manning and Vardaro, 1977; Lacy, 1979; Damicone et.al., 1981; Damicone and Manning, 1982; Hussey et.al, 1984; Tu et.al., 1990; Hanks, 1996; Elmer, 2002; Reid eta], 2002), a more effective and reliable management program is needed. The development of asparagus cultivars that are resistant or tolerant to Fusarium crown and root rot is a key component in disease management. Some of the commercial asparagus cultivars tend to show a higher tolerance to F usarium crown and rot root than do other cultivars (Stephens et.al, 1989; Sonoda et.al., 2002). In particular, the all-male hybrids demonstrated a greater tolerance than the mixed male/female cultivars such as Mary Washington. Asparagus densiflorus, a relative of A. oflicinalis, was identified as resistant to F usarium crown and root rot (Stephens et.al., 1989). However, resistance has not been observed in A. oflicinalis. Breeders have not been able to cross the related asparagus species due to genetic differences restricting the genetic pool to A. oflicinalis. While breeders have not been able to incorporate outside resistance sources, they have used different selection criteria within A. ofi‘icinalis to identify F usarium resistance. Pontaroli and Carmadro (2001) successfully used gametophyte selection to develop resistance that could be used on a large scale. Commercial seed companies continue to release new asparagus cultivars, however, few are screened for resistance to F usarium spp. The objective of the study was to screen commercially available cultivars for susceptibility to F usarium spp. in a 58 controlled environment. 59 MATERIALS AND METHODS Seeds of 20 asparagus cultivars (nineteen commercial and one experimental; Table 8) were weighed into 15 g lots. Individual seed lots were added to separate 100 ml solutions of 25,000-ppm benomyl in acetone and agitated on a rotary shaker (2000 rpm) for 24 hours. Following disinfection, the benomyl solution was decanted and the seeds were rinsed three times with acetone and then rinsed three times with sterile distilled water. One hundred milliliters of a 10% bleach solution was added to the seeds and then agitated at 2000 rpm for one hour for further disinfecting. Flasks were removed from the shaker and the bleach solution was decanted. Seeds were then rinsed three times with sterile distilled water. Rinsed seeds were allowed to dry on sterile papers towels in a flow hood for three to four hours before storing at 45° C. Based on previous studies, a pathogenic isolate of F. oxysporum and F. proliferatum were selected due to the high virulence that was exhibited (Reid, 2000). The isolate 2-1 was selected to represent F. oxysporum and the P-67 isolate was used to represent F. proliferatum. Both isolates are maintained in the culture collection of Dr. M. Hausbeck at Michigan State University. Isolates were grown on carnation leaf agar (CLA) (Fisher et.al., 1982) at 20 to 25° C for 14 days prior to inoculation of the seedlings. Seeds were germinated on sterile water agar plates that had been wrapped in aluminum foil and kept in the dark at 20 to 25° C for five to seven days. Germinated seedlings were then transferred to test tubes containing 20 ml of Hoagland’s media (Manning et.al., 1985; Stephens, 1988). Test tubes were placed in growth chambers set on a 12-hour day/night cycle with temperatures of 25° C (day) and 20° C (night). After 21 days, seedlings of each cultivar (5) were either inoculated with a 7-mm agar plug of isolate 2-1, 60 isolate P-67, or were uninoculated and served as a control. Agar plugs were placed touching the seedling with each test tube receiving 1 ml of sterile distilled water. Test tubes were then randomly placed in the growth chamber. Table 8. Asparagus cultivars tested for susceptibility to F. oxysporum f. sp. asparagi and F proliferatum. Cultivar Source 1 Jersey Giant Jersey Asparagus Farms, Inc. 2 Jersey Knight Jersey Asparagus Farms, Inc. 3 Jersey General Jersey Asparagus Farms, Inc. 4 Jersey King Jersey Asparagus Farms, Inc. 5 Jersey Supreme Jersey Asparagus Farms, Inc. 6 Jersey Gem Jersey Asparagus Farms, Inc. 7 Greenwich Jersey Asparagus Farms, Inc. 8 Gignlim Asparagus BV 9 K-5 78 Asparagus BV 10 Backlim Asparagus BV 11 Grolim Asparagus BV 12 Thelim Asparagus BV l3 UC157 California Asparagus Seed and Transplant 14 Apollo California Asparagus Seed and Transplant 15 Atlas California Asparagus Seed and Transplant 16 Grande California Asparagus Seed and Transplant 17 Dulce Verde California Asparagus Seed and Transplant 18 Millenium University of Guelph 19 Tiessen University of Guelph 20 MarLWashington Rupp Seeds 61 Following inoculation, seedlings were allowed to grow and infection to develop for four weeks in a grth chamber. At the end of this period, test tubes were removed from the chamber and seedlings were rated for disease on a scale of 1 (healthy, no disease) to 6 (plant death, severe disease) (Table 9, Figure 3). The trial was conducted two times. Statistical analysis was conducted using SAS (Statistical Analysis Software ver. 8.2). The PROC GLM method of SAS was used to perform a simple two-way factorial ANOVA to compare cultivars. Table 9. Scale of disease ratings on inoculated asparagus seedlings. Rating Symptoms 1 N0 disease 2 < 25 %‘ 3 26-50 %, fern yellowing 4 51-75 %, wilting and browning of fern 5 76-99 % 6 Plant death ' Indicates area of root system infected 62 Figure 3. The F usarium symptoms observed on inoculated asparagus seedlings receiving the rating of a 2 (left) to a 6 (right). 63 RESULTS Rusty brown elliptical lesions were first observed on the seedling roots 14 days after inoculation with mycelial growth on the stems also noted at this time. The cultivars screened had a minimum severity rating of 2.2. Higher severity ratings were noted for the F oxysporum isolate than the F proliferatum isolate in Trial 1 and Trial 2. In Trial 1 the cultivars Dulce Verde, Tiessen, Jersey Knight, and Thelim were significantly healthier than the cultivars Mary Washington or Jersey Giant when inoculated with F oxysporum (Table 10). In Trial 2 only the cultivar Jersey Giant was significantly better than some of the cultivars tested, including Thelim, Jersey Supreme, Millenium, K-578, Backlim, Jersey King, Mary Washington, Gijnlim, Atlas, Grande, Jersey Knight, Jersey General, and UC-157, when inoculated with F oxysporum (Table 10). In Trial 1, Greenwich was significantly less susceptible than Grande, Mary Washington, Jersey Knight, and Gijnlim when inoculated with F proliferatum (Table 11). In Trial 2, the cultivars Jersey General, Thelim, Dulce Verde, Jersey Giant, and Jersey Knight were significantly better than cultivars Apollo, Grande, Jersey King, Jersey Supreme, and K- 578 when inoculated with F proliferatum (Table 11). 64 Table 10. Average disease ratings for asparagus cultivars inoculated with F oxysporum f.sp. asparagi 2-1. Cultivar Trial 1 Trial 2 Jersey Giant ............. 4.0y be2 2.6 a Dulce Verde ............ 2.8 a 3.0 ab Tiessen .................... 2.8 a 3.2 abc Greenwich ............... 3.6 abc 3.8 abcd Grolim ..................... - - 3.8 abcd Jersey General ......... 3.0 ab 3.9 abcd Appolo ..................... 3.2 ab 4.0 abcde UC-157 .................... 3 .6 abc 4.2 cdef Jersey Knight ........... 2.6 a 4.4 cdef Atlas ........................ 3.2 ab 4.6 cdef Grande ..................... 3.6 abc 4.6 cdef Ginglim ................... 3.0 ab 4.8 def Backlim ................... 3.0 ab 5.0 def Jersey King .............. 3.6 abc 5.0 def Mary Washington... 4.6 c 5.0 def Jeresy Supreme ....... 3.2 ab 5.4 ef Millenium ................ 3.0 ab 5.4 ef K-578 ...................... - - 5.4 ef Thelim ..................... 2.6 a 5.6 f y Seedlings were rated based on the following scale, l=no disease, 2=<25 % of root system infested, 3=26-50 % of root system infected with fern yellowing, 4:51-75 % of root system infected with wilting and browning of fern, 5:76-99 % of root system infected with wilting and browning of fern, and 6=dead. z Cultivars with the same letter are not significantly different at the 0.05 level of Tukey’s HSD. 65 Table 11. Average disease ratings for asparagus cultivars inoculated with F proliferatum P-67. Cultivar Ratings Trial 1 Trial 2 Dulce Verde .................................. 3.2y abz 2.2 a Thelim ........................................... 3.2 ab 2.2 a Jeresy Giant ................................... 2.8 ab 2.4 a Jersey Knight ................................. 3.4 b 2.4 a J eresy General ............................... 2.9 ab 2.5 a Ginglim ......................................... 3.4 b 2.6 ab Tiessen .......................................... 2.8 ab 2.8 abc Atlas .............................................. 3.0 ab 3.0 abc Millennium .................................... 2.8 ab 3.0 abcd Backlim ......................................... 3.2 ab 3.2 abcd Greenwich ..................................... 2.4 a 3.2 abcd UC-157 .......................................... 3.2 ab 3.2 abcd Grolim ........................................... - 3.2 abcd Mary Washington .......................... 3.6 b 3.4 abcd Jersey Supreme ............................. 2.8 ab 3.8 bcd K-578 ............................................ - 3.8 bcd Grande ........................................... 3.6 b 4.0 cd Jersey King .................................... 3.0 ab 4.0 cd Apollo ........................................... 3.0 ab 4.2 d y Seedlings were rated based on the following scale, 1=no disease, 2=<25 % of root system infested, 3=26-50 % of root system infected with fern yellowing, 4:51-75 % of root system infected with wilting and browning of fern, 5=76-99 % of root system infected with wilting and browning of fern, and 6=dead. z Cultivars with the same letter are not significantly different at the 0.05 level of Tukey’s HSD. 66 DISCUSSION Identifying and developing an asparagus cultivar that is resistant or tolerant to F usarium has challenged asparagus breeders worldwide. While A. oflicinalis is not disease resistant, a relative species, A. densiflorus, has exhibited resistance to F usarium. The problem breeders face is the inability to crossbreed A. densiflorus and A. oflicinalis (Stephens et.al., 1989). Some of the cultivars screened in this trial had lower severity values than ‘Mary Washington’; however, these cultivars would not be considered to be F usarium resistant given their disease rating. Further, results were inconsistent between the two trials conducted. This variability in disease in our trial is likely due to the genetic variability that has been demonstrated for asparagus (Lassaga et.al., 1998). In a greenhouse study by Sonoda et. a1. (2002) also demonstrated that certain commercial and experimental cultivars had disease indices that were lower than the cultivar Mary Washington. All-male hybrids tend to have a greater tolerance to F usarium than the female cultivars (Ellison and Kinelski, 1985; Elmer, 2001). In trials, between 70 and 90 % of the cultivars had lower severity ratings than Mary Washington, suggesting a greater tolerance in the all-male hybrids (Ellison and Kinelski, 1985; Elmer, 2001). Even though disease severity values tended to be lower than Mary Washington, few of the male hybrids received a rating that was significantly lower. While F usarium resistance has not been identified in A. ofi‘icinalis, other crops such as tomatoes, collard, cucumber, and melons have identified lines that are resistant to F usarium spp. (Jones and Woltz, 1981; Pavlou et.al., 2002; Perchepied and Pitrat, 2004). Asparagus has been screened for resistance to Phytophthara spp. in greenhouse and field 67 trials (F alloon, 1985; Falloon, 1990; Falloon et.al., 2002). Several hybrids have been identified that contain durable resistance to Phytophthtora and have yield and quality characteristics equal to or better than the standard cultivars (Falloon et.al., 2002). There are many commercial cultivars that contain a range of traits that are suitable for different growing regions of the world. Although cultivars are often specific for a particular growing region, a resistant cultivar could be used to incorporate disease resistance into lines that are more suitable for the desired growing region. Future efforts should include screening asparagus gerrnplasm for resistance to both F usarium spp. and Phytophthora spp. 68 LITERATURE CITED Arriola, LL. 1997. Arbuscular Mycorrhizal fungi and Trichoderma harzianum in relation to border cell production and F usarium crown and root rot of Asparagus. Botany and Plant Pathology. East Lansing, Michigan State University. Masters. Burrows, R., F .L. Pfleger, and L. Waters. 1990. Growth of seedling asparagus inoculated with Glomusfasciculatum and phosphorus supplementation. HortScience 25: 519—521. Cohen, SJ. 1941. A wilt and root rot of asparagus caused by F usarium oxysporum Schlecht. Plant Disease Reporter 25: 503-509. Damicone, J .P., and W.J. Mamring. 1982. Avirulent strains of F usarium oxysporum protect asparagus seedlings from crown rot. Canadian Journal of Plant Pathology 4: 143-146. Damicone, J .P., D.R. Cooley, and W.J. Manning. 1981. Benomyl in acetone eradicates F usarium mom'liforme and F oxysporum from asparagus seed. Plant Disease 65: 892- 893. Ellison, J .H., and J .J . Kinelski. 1985 . 'Jersey Giant', An all-male asparagus hybrid. HortScience 20: 1141. Elmer, W.H. 2001. Fusarium diseases of asparagus. Fusarium: Paul E. Nelson Memorial Symposium. St. Paul, MN, APS Press.248-262. Elmer, W.H. 2002. Influence of forrnononetin and NaCl on mycorrhizal colonization and Fusarium crown and root rot of asparagus. Plant Disease 86: 1318-1324. Falloon, P.G. 1985. A method for screening asparagus (Asparagus officinalis L.) seedlings for resistance to Phytophthora rot. 6th International Asparagus Symposium, Guelph, Ontario, Canada, Acta Horticulture. 220-227. F alloon, P.G. 1990. Field screening of asparagus for tolerance to Phytophthora rot. Seventh International Asparagus Symposium, Ferrara, Italy, Acta Horticulturae. 69-73. 69 Falloon, P.G., L.M. Falloon, and A.M. Andersen. 2002. Breeding asparagus cultivars resistant to Phytophthora. 10th International Asparagus Symposium, Niigata, Japan, ACTA Hort. 185-191. Fisher, N.L., L.W. Burgess, T.A. Toussoun, and PE. Nelson. 1982. Carnation leaves as a substrate and for preserving cultures of F usarium species. Phytopathology 72: 151- 153. Hanks, GR. 1996. Control of F usarium oxysporum f.sp. narcissi, the cause of narcissus basal rot, with thiabendazole and other firngicides. Crop Protection 15: 549-558. Hussey, R.B., R.L. Peterson, and H. Tiessen. 1984. Interactions between vesicular- arbuscular mycorrhizal fungi and asparagus. Plant and Soil 79: 409-416. Jones, J .P., and SS. Woltz. 1981. F usarium-incited diseases of tomato and potato and their control. F usarium: Diseases, Biology, and Taxonomy. P.E. Nelson, Toussoun, T.A. and Cook, R.J. University Park, Pennsylvania State University Press.157-168. Lacy, M.L. 1979. Effects of chemicals on stand establishment and yields of asparagus. Plant Disease Reporter 63: 612-616. Lassaga, S.L., E.L. Camadro, and F .J . Babinec. 1998. Assessing genetic variablity for F usarium resistance in three asparagus populations with an in vitro assay. Euphytica 103:131-136. Manning, W.J, J .P. Damicone and R.L. Gilbertson. 1985. Asparagus root and crown rot: sources of inoculum for F usarium oxysporum and F usarium moniliforme. 6th International Asparagus Symposium, University of Guelph, Ontario, Canada, Acta Hort. 195-204. Manning, W.J., and PM. Vardaro. 1977. Soil fumigation and preplant fungicide crown soaks: effects on plant growth and F usarium incidence in newly planted asparagus. Plant Disease Reporter 61: 355-357. Pavlou, G.C., D.J. Vakalounakis, and BK. Ligoxigakis. 2002. Control of root and stem rot of cucumber, caused by F usarium oxysporum f.sp. radicis-cucumerinum, by grafting onto resistant rootstocks. Plant Disease 86: 379-382. 70 Perchepied, L., and M. Pitrat. 2004. Polygenic inheritance of partial resistatrrce to Fusarium oxysporum f.sp. melonis race 1.2 in melon. Phytopathology 94: 1331-1336. Pontaroli, AC, and EL. Camadro. 2001. Increasing resistance to F usarium crown and root rot in asparagus by gametophyte selection. Euphytica 122: 343-350. Reid, TC. 2000. Evaluation of strategies for the control of F usarium crown and root rot of asparagus. Plant Pathology. East Lansing, Michigan State University. Masters:l31. Reid, T.C., M.K. Hausbeck, and K. Kizilkaya. 2002. Use of Fungicides and Biological Controls in the Suppression of Fusarium Crown and Root Rot of Asparagus Under Greenhouse and Growth Chamber Conditions. Plant Disease 86: 493-498. Sonoda, T., K. Tairako, and A. Uragami. 2002. Comparative evaluation of resistance of Asparagus oflicinalis L. cultivars and breeding lines to F usarium stem and crown rot. 10th International Symposium on Asparagus, ActaHorticulturae. 387-390. Stephens, CT. 1988. An in vitro assay to evaluate sources of resistance in Asparagus spp. to F usarium crown and root rot. Plant Disease 72: 334-337. Stephens, C.T., R.M. DeVries, and KC. Sink. 1989. Evaluation of asparagus species for resistance to F usarium oxysporum f.sp. asparagi and F mom'liforme. HortScience 24: 365-368. Tu, C.C., Y.H. Cheng, and AS. Cheng. 1990. Recent advance in biological control of F usarium wilt of asparagus in Taiwan. Acta Horticultureae 271: 353. 71 CONCLUSION Asparagus decline has been a problem in the Michigan asparagus industry for many years and is caused by F oxysporum f.sp. asparagi and F proliferatum. The research presented gives some insight as to what can be done to manage asparagus decline. A first step in establishing a successful production field is to select a cultivar that is suited for the climate and is vigorous plants, with a low susceptibility to F usarium. Of the cultivars screened in this study using F oxysporum f.sp. asparagi, ‘Tiessen’ and ‘Dulce Verde’ could be considered depending on the growing region. After selecting a suitable cultivar, seedlings should be grown in a fumigated seedbed for one year. Resulting crowns should be graded for uniformity and diseased crowns should be discarded before being sent to the production grower. Crowns should be dipped in a fungicide and/or a biocontrol agent prior to planting. When a field has a history of F usarium infection, dipping the crowns in thiophanate-methyl, may increase plant stand and fern height. A sub-surface drip irrigation system installed in the production field would allow applications of a fungicide and/or biocontrol agent during the growing season. If a field is also infested with the soilbome pathogen, Phytophthora, F usarium treatments tend not to affect plant stand or fern height. A fungicide appropriate for Phytophthora will need to be implemented into the disease management program. F usarium treatments do not positively affect plant stand or fern height. Successful management of asparagus decline requires a multi-faceted approach that includes genetic resistance, cultural strategies, and fungicides. 72 APPENDIX I 73 ASPARAGUS (Asparagus oflicinalis ‘Mary J .W. Counts and M.K. Hausbeck Washington‘) Department of Plant Pathology Fusarium crown and root rot; F usarium sp. Michigan State University East Lansing, MI 48824 Evaluation of fumigants for managing Fusarium crown and root rot of asparagus, 2004. This study was conducted on a cooperator’s farm in Oceana County, MI on loamy fine sand (Spinks-Benona) with a known history of F usarium. The field was previously planted to asparagus, which was removed the fall of 2003 and then seeded with a rye cover crop. Prior to destruction of the asparagus field, areas of the field with missing asparagus plants were plotted in order to position the experiment in a uniform F usarium location. Rye was worked under early spring to a depth of 25 to 30.5 cm. with a field cultivator. Old asparagus crowns were removed from the surface of plot area, and the plot was reworked prior to bed formation on 26 May. Raised beds were formed (Rainflo 2550 bed former) and shank injected treatments (Table 12) were applied on 26 May. Treatments were applied using sweptback gas knives on 20-cm. spacing at a depth of 25 to 30.5 cm. from bed top. All treatment beds were covered with black-plastic mulch (1.25 mil embossed). Treatment beds consisted of paired, treated with an untreated control, 12-m rows that were 63.5 cm. wide at the top and 68.5 cm. wide at the bottom. Beds were 15 cm. high with a slight crown with a 162.5-cm. spacing of bed centers. Plastic mulch was held down on the bed with a 8-cm. overlap covered with soil. Two drip lines were installed 20-cm. apart and 2.5-cm. below the plastic during bed formation. The drip tape was 8-mil thick with 20-cm. emitter spacing with a flow rate of 5.6 LPM for 100 m of tape. Treatments were setup in a randomized complete block design with a total of four replicates. 74 Table 12. Fumigation materials tested by product name and active ingredient for efficacy on Fusarium crown and root rot on asparagus seedlings. Treatment Active ingredient Product/Ha Application Plant method back SEP 100 .................................................. sodium azide 20% 392.7 L Drip injection 21 day chloropicrin 100% 233.7 L Shank injected Chloropicrin 100% + K-pam TM ............. metam potassium 54% 561.0 L Drip injection 14 day K-pam TM ................................................ metam potassium 54% 280.5 L Drip injection 14 day K-parnTM ................................................. metam potassium 54% 561.0 L Drip injection 14 day PropozoneTM ........................................... propylene oxide 99.9% 748.0 L Shank injected 7 day Methyl Bromide/ methyl bromide 67% Chloropicrin (67/33) .............................. chloropicrin 33% 392.1 kg Shank injected 7 day 1,3-dichloropropene 63.4% Telone C35TM ......................................... chloropicrin 34.7% 327.2 L Shank injected 14-21 day Chloropicrin 100% ................................. chloropicrin 100% 28.0 kg Shank injected 7 day iodomethane 33% MidasTM/Chloropicrin 33/67 .................. chloropicrin 67% 336.1 kg Shank injected 7 day Drip treatments (Table 12) were applied on 28 May and 29 May using a stainless steel nitrogen apparatus. Applications were made using two drip tapes per bed at an operating pressure of 82-103.5 kPa. Beds were prewetted for 30 min. Treatments were premixed with water in a 1:1 ratio to form an emulsion before application into the drip tape system. Application of treatments occurred over a 90-min time span with containers being agitated every 15-20 min. Each drip treatment application was regulated with flow regulators (TeeJet CP49l6-16) operating at 172-207 kPa and material was not allowed to back-flow using spring load check valves. Upon completion of application, the irrigation was allowed to run for another 30 min to insure even dispersal of treatments with a total 75 of 1 in. of water being applied over 150 min. An additional 12.7 mm. of water was applied on 3 Jun to purge the treatments from the beds. Each treatment row had five sections (1 5-cm. wide by 2.4—m long) of plastic removed in the center of the beds, leaving 30-cm. strips to keep sides tight and reduce blowing of the plastic. The plastic was removed to facilitate planting. To aid in reducing washing and blowing of soil, a silt fence was placed around plot. Beds were planted on 15 Jun using a V-belt planter with an approximate seed spacing of 2.5-cm. and a depth of 2.5- cm. Weeds were controlled between rows with an application of Sandea 75WG (47 g/Ha) and Roundup (4.7 L/Ha) on 14 Jul and in rows by hand weeding. Diseases were controlled by applying Bravo Weatherstik 6SC (2.3 L/Ha) on 14, 29 Jul and 12, 24 Aug and Folicur 3.6 F (438.3 ml/Ha) on 14 Jul and 12 Aug. Insects were controlled with applications of Asana XL (701 ml/Ha) on 14 Jul, Sevin XLR Plus (3.5 L/Ha) on 29 Jul and 12, 24 Aug, and Diazinon AG500 (1.8 L/Ha) on 29 Jul and 12, 24 Aug. Irrigation was applied once during the growing season on 29 Jul at 12.7-mm per block. The percentage of bed covered with weeds was visually estimated on 13 Jul. All seedlings in the 12-m row were harvested on 28 Sep and transported in coolers to cold storage. Seedling weights, root lengths, number of root lesions, number of fern lesions, root health (1=healthy no lesions, 10=dead), fern health (1= health no lesions, 10= dead), and plant vigor (1= clean robust seedlings, 10= dead) were taken during the period from 30 Sep to 11 Oct. Seedling weights were gathered from 50 randomly selected seedlings from the row or as many seedlings were available in a row. Data from the trial were analde using Statistical Analysis Software (SAS v8.2.) with the ‘Proc Mixed’ 76 procedure. The analysis was based on a split plot design with treatments being the whole plot factor and application (treated vs. untreated) being the sub-plot factor. Treatment and application were considered fixed effects with rep and rep*treatment being random effects. It was first determined if significant differences occurred between the treated and untreated sections. If significant differences were present contrasts were used to determine which treatment showed the best differences between the treated and untreated section. Comparisons were done using the Sidak correction for multiple testing at P=0.05. RESULTS This plot received heavy rains prior to and after bed formation. Prior to planting the plot received 25.4-mm. of rain in approximately 15 min that washed over and destroyed several beds. SEP 100 showed early phytotoxicity and a reduced stand; however at the end of the study the seedlings recovered and were some of the healthiest. Weights for the seedling crowns showed significant differences between treatments (Table 12). Sep 100 had the greatest positive increase in seedling weight. K- pamTM 280.5 L, Telone C-35TM, Chloropicrin + K-pamTM and K-pamTM 561.0 L also had positive increases in seedling weight. Propozone showed the greatest negative weight difference on the seedlings compared to the untreated control. Methyl Bromide, MidasTM, Chloropicrin 100% also showed negative effects on seedling weight possible due to movement of gas down the row between the treated and untreated sections that compromised the untreated soil. There were clear differences between root lengths of treated and untreated plots (Table 13). Roots treated with K-pamTM 561.0 L had the greatest difference between 77 untreated and treated sections and was significantly better than PropozoneTM, Methyl Bromide, and MidasTM. Although not significantly different from the other treatments, SEP 100 had the second highest difference between results from treated and untreated plots. SEP 100 and K-pamTM 561.0 L had significantly fewer lesions on the roots and fern than did PropozoneTM (Table 14 & 15). Roots of plants treated with Sep 100 were significantly healthier than Propozone and Methyl Bromide (Table 16). K-pamTM 561.0 L had the greatest significant effect on the fern health followed by Chloropicrin + K- parnTM and both were significantly better than PropozoneTM (Table 17). For overall plant vigor SEP 100 was significantly better than PropozoneTM and Methyl Bromide (Table 18). K-pam TM 561.0 L and Chloropicrin + K-parnTM also had better vigor ratings but they were only significantly different from Propozone. There were no significant differences among treatments (P=0.50) for weed rating (Table 19). However, SEP-100, Kapam 280.5 L, Chloropicrin 100% + K-pamm, and PropozoneTM applications all were significantly better than the paired untreated control. 78 Table 13. Weight evaluation of 1-year-old asparagus seedlings produced in fumigated soil for managing F usarium crown and root rot of asparagus. Weight (g) Standard Sidak Treatment Rate/Ha untreated treated difference error 8TOUP Propozonem ................ 748 L 84.6 40.8 .43.7* 27.8 a" Methyl Bromide/ Chloropicrin (67/33) 392.1 kg 130.4 105.4 -25.0 19.7 a MidasTM /Chloropicrin (33/67) ......................... 336.1 kg 152.3 140.4 -11.9 19.7 ab Chloropicrin 100% ...... 28.0 L 122.2 110.6 -11.7 19.7 ab K-pamm ..................... 280.5 L 85.2 106.7 21.5 22.7 ab Telone c-35TM ............. 372.2 L 102.6 129.3 26.8 19.7 ab Chloropicrin 100% + 233.7 L k-pannTM ...................... 561.0 L 127.7 172.7 45.0 19.7 ab K-pamTM ..................... 561.0 L 82.0 133.7 51.7 19.7 ab SEP 100 ....................... 392.7 L 74.1 153.3 79.2 19.7 b * Negative numbers indicate untreated mean weight was greater. " Treatments with same letter are not significantly different at the 0.05 level for Sidak correction. 79 Table 14. Root length evaluation of asparagus seedlings produced in firmigated soil for managing Fusarium crown and root rot of asparagus. Length (cm) Standard Sidak Treatment Rate/Ha untreated treated difference error group Propozonem ................ 748.0 L 15.7 13.2 -2.5 r 27.8 a“ Methyl Bromide/ Chloropicrin (67/33) 392.1 kg 18.2 17.2 -0.9 19.7 a MidasTM /Chloropicrin (33/67) ......................... 336.1 kg 18.3 17.6 -0.7 19.7 a Telone c-35TM ............. 327.2 L 16.1 16.7 0.6 19.7 ab Chloropicrin 100% ...... 28.0 kg 15.9 17.4 1.5 22.7 ab K-parnTM ...................... 280.5 L 15.8 17.9 2.1 19.7 ab Chloropicrin 100% + 233.7 L K-pamm ...................... 561.0 L 16.9 19.3 2.3 19.7 ab SEP 100 ....................... 392.7 L 14.4 17.1 2.7 19.7 ab K—pamm ...................... 561.0 L 14.2 19.2 5.0 19.7 b * Negative numbers indicate untreated mean length was greater. " Treatments with same letter are not significantly different at the 0.05 level for Sidak correction. 80 Table 15. Root lesion count evaluation of asparagus seedlings produced in fumigated soil for managing F usarium crown and root rot of asparggus. Root lesions Standard Sidak Treatment Rate/Ha untreated treated difference error 8Y0“? Propozonem ................ 748.0 L 2.8 4.1 1.3 0.7 a* MidasTM /Chloropicrin (33/67) ......................... 336.1 kg 0.5 0.8 0.2 0.5 ab Telone c-35TM ............. 327.2 L 1.0 0.9 -0.1 u 0.5 ab Chloropicrin 100% ...... 28.0 kg 1.1 0.7 -0.5 0.5 ab Methyl Bromide/ Chloropicrin (67/33) .. 392.1 kg 1.4 0.7 -0.6 0.5 ab Chloropicrin 100% + 233.7 L K-pamm ...................... 561.0 L 1.6 0.1 -1.5 0.5 ab K-pannTM ...................... 280.5 L 1.8 0.1 -l.6 0.5 ab K-pamTM ..................... 561.0 L 2.1 0.3 -1.8 0.5 b SEP 100 ....................... 392.7 L 2.1 0.2 -1.9 0.5 b "' Treatments with same letter are not significantly different at the 0.05 level for Sidak correction. “ Negative numbers indicate untreated mean weight was greater. 81 Table 16. F em lesion count evaluation of asparagus seedlings produced in fumigated soil for managing Fusarium crown and root rot of asparagus. Fern lesions Standard Sidak Treatment Rate/acre untreated treated difference error group PropozoneTM ................ 748.0 L 1.5 1.9 0.4 0.2 a" MidasTM /Chloropicrin (33/67) ......................... 336.1 kg 0.7 0.5 -02 M 0.2 ab Methyl Bromide/ Chloropicrin (67/33) 392.1 kg 0.6 0.5 -0.1 0.2 ab SEP 100 ....................... 392.7 L 1.0 0.8 -0.2 0.2 ab Chloropicrin 100% ...... 28.0 kg 0.7 0.4 -0.3 0.2 ab Telone c-35TM ............. 327.2 L 1.0 0.6 -0.4 0.2 ab K-pamm ...................... 280.5 L 0.6 0.2 04 0.2 ab Chloropicrin 100% + 233.7 L K-parnTM ...................... 561.0 L 1.4 0.6 -0.8 0.2 b K-pamTM ...................... 561.0 L 1.4 0.3 -1.1 0.2 b "‘ Treatments with same letter are not significantly different at the 0.05 level for Sidak correction. " Negative numbers indicate untreated mean weight was greater. 82 Table 17. Root health evaluation of asparagus seedlings produced in fumigated soil for managing Fusarium crown and root rot of asparagus. Root healthz Standard Sidak Treatment Rate/acre untreated treated difference error ETOUP PropozoneTM ................ 748.0 L 2.2 2.5 0.3 0.3 ay MidasTM /Chloropicrin (33/67) ......................... 336.1 kg 1.3 1.4 0.1 0.2 a Telone C-35TM ............. 327.2 L 1.6 1.6 0.0 0.2 ab Methyl Bromide/ Chloropicrin (67/33) 392.1 kg 1.6 1.4 -0.2 " 0.2 ab Chloropicrin 100% ...... 28.0 kg 1.6 1.3 -0.3 0.2 ab Chloropicrin 100% + 233.7 L K-pamTM ...................... 561.0 L 1.7 1.1 -0.6 0.2 ab k-parnTM ...................... 280.5 L 1.9 1.1 -0.8 0.2 ab K-pamm ..................... 561.0 L 1.9 1.2 -0.7 0.2 ab SEP 100 ....................... 392.7 L 2.0 1.1 -0.9 0.2 b z l=healthy no lesions, 10=dead. y Treatments with same letter are not significantly different at the 0.05 level for Sidak correction. "Negative numbers indicate untreated mean was greater. 83 Table 18. Fern health evaluation of asparagus seedlings produced in fumigated soil for managing Fusarium crown and root rot of asparagus. F em healthz Standard Sidak Treatment Rate/acre untreated treated difference 9170" group PropozoneTM ................ 748.0 L 1.7 2.0 0.3 0.1 ay Methyl Bromide/ Chloropicrin (67/33) 392.] kg 1.4 1.3 -0.1 " 0.9 ab MidasTM /Chloropicrin (33/67) ......................... 336.1 kg 1.4 1.3 -0.1 0.1 abc Chloropicrin 100% ...... 28 kg 1.4 1.3 -0.1 0.1 abc SEP 100 ....................... 392.7 L 1.6 1.4 -0.1 0.1 abc Telone c-35TM ............. 327.2 L 1.6 1.5 -0.1 0.1 abc K-pamTM ...................... 280.5 L 1.4 1.1 -0.3 0.1 abc Chloropicrin 100% + 233.7 L k-pamTM ...................... 561.0 L 1.7 1.3 -0.4 0.1 bc K-pamm ...................... 561.0 L 1.7 1.2 -0.5 0.1 c ‘ l=healthy no lesions, 10=dead. ’ Treatments with same letter are not significantly different at the 0.05 level for Sidak correction. " Negative numbers indicate untreated mean was greater. 84 Table 19. Plant vigor evaluation of asparagus seedlings produced in fumigated soil for managing Fusarium crown and root rot of asparagus. Plant vigorz Standard Sidak Treatment Rate/acre untreated treated difference error group PropozoneTM ................ 748.0 L 5.2 7.3 2.1 0.5 ay Methyl Bromide/ Chloropicrin (67/33) 392.1 kg 5.0 5.7 0.7 0.4 ab Chloropicrin 100% ...... 28 kg 4.9 5.1 0.2 0.4 abc MidasTM /Chloropicrin (33/67) ......................... 336.1 kg 3.7 4.0 0.3 0.4 abc Telone C-35 ................ 327.2 L 4.9 5.1 0.2 0.4 abc K-pamm ...................... 280.5 L 5.2 5.2 0.0 0.4 abc Chloropicrin 100% 233.7 L + K-pamTM .................. 561.0 L 4.3 3.5 -0.8 " 0.4 bc K-parnTM ..................... 561.0 L 5.0 3.9 -l .1 0.4 bc SEP 100 ....................... 392.7 L 6.0 4.4 -l.6 0.4 c ‘ l= clean robust seedlings, 10= dead ’ Treatments with same letter are not significantly different at the 0.05 level for Sidak correction. "Negative numbers indicate untreated mean was greater. 85 Table 20. Weed evaluation of asparagus seedbeds treated with various fumigants. Weed ratingz Standard Sidak Treatment Rate/acre untreated treated difference error group Chloropicrin 100% ...... 28.0 kg 38.7 35.0 -3.7y 10.1 a" MidasTM /Chloropicrin (33/67) ......................... 336.1 kg 33.7 25.0 -8.7 10.1 a Telone c-35TM ............ 327.2 L 32.5 23.7 -8.8 10.1 a K-pantTM ..................... 561.0 L 20.0 3.7 -l6.3 Methyl Bromide/ Chloropicrin (67/33) 392.1 kg 33.7 16.2 -17.5 10.1 a PropozoneTM ................ 748.0 L 40.0 15.0 -25.0 r 10.1 a K-pamTM ...................... 280.5 L 31.2 3.7 -27.5 * 10.1 a SEP 100 ....................... 392.7 L 30.0 2.5 -275 r 10.1 a Chloropicrin 100% 233.7 L + K-pamm .................. 561.0 L 37.5 5.0 -325 * 10.1 a ’ Visual estimation of percentage of bed covered. ’ Negative numbers indicate untreated mean was greater. " Treatments with same letter are not significantly different at the 0.05 level for Sidak correction. * applications were significantly different (Sidak) This project was made possible by the support of lR-4. 86 APPENDIX 11 87 Table 21. Percent difference in stand count and fern height for asparagus treated at planting with fungicides or biocontrol agents in Crown Trial 2. Stand countl Treatment and rate 2003 2004 2005 Nonpathogenic F oxysporum 397 g/ 12 m 2 ...... -1.03 -5.0 -2.8 Arbuscular mycorrhizae 59.5 g/37.8 L .............. 4.9 3.0 0.1 T. harzianam 910 g/37.8 L ................................ 0.5 0.6 1.2 Fludioxonil 22.7 g/37.8 L ................................. -2.2 -5.6 -0.7 Thiophanate-methyl 44.8 g/37.8 L .................... -1.7 -2.6 1.4 Fern heightl Nonpathogenic F oxysporum 397 g/12 m ........ 1.4 4.3 -0.7 Arbuscular mycorrhizae 59.5 g/37 .8 L ............. 2.3 3.7 4.5 T. harzianam 910 g/37.8 L ................................ -1.0 -3.0 3.3 Fludioxonil 22.7 g/37.8 L ................................. -2.3 -1.6 -4.1 Thiophanate-methyl 44.8 g/37.8 L .................... 5.5 1.3 -0.3 1Ratings taken from the center 6.1 m of row on 12 August 2003, 20 August 2004, and 10 August 2005. 2Treatments were not significantly different at P>0.05 (Tukey’s HSD). 3Negative numbers indicate a percent reduction in plant stand and fern height when compared to the untreated. 88 1|lll‘lllllllIllllll 3 1293 0273