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Other___________________________________________________________________________ . Text follows. University Microfilms International ASPARAGUS VIRUSES AND THEIR CONTRIBUTIONS TO MICHIGAN ASPARAGUS DECLINE By Thomas Allen Evans A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of DOCTOR OF PHILOSOPHY Department of Botany and Plant Pathology 1985 ABSTRACT ASPARAGUS VIRUSES AND THEIR CONTRIBUTIONS TO MICHIGAN ASPARAGUS DECLINE BY Thomas Allen Evans Two viruses known to affect the vigor and productivity of commercial asparagus, Asparagus officinal is L., were investigated to determine their incidence and mode of transmission and spread within Michigan asparagus plantings. In addition, the potential of these viruses as biological stress factors leading to an increase in the severity of Fusarium crown and root rot was evaluated in the greenhouse and laboratory. Asparagus virus II was determined to be widespread within most asparagus plantings in Michigan with as many as 50 to 60% of the plants in most fields being infected. Asparagus virus I was isolated from only 5 of 60 asparagus plantings in Michigan. All five infected fields were in Oceana County and were within one mile of each other. These studies indicate that AV I can be transmitted within asparagus by the green peach aphid (Mvzus periscae Sulz.) but not by the European asparagus aphid (Brachycolus asparagi Mord.). The distribution of AV I-infected asparagus plants in the field is non-random as determined by ordinary run analysis. THOMAS ALLEN EVANS This research indicates that AV II can be mechanically spread within asparagus by sap transmission. Also, the exine of pollen from some AV II-infected asparagus plants was determined to be contaminated with infectious virions of AV II. The distribution of AV II-infected plants within a field was determined by ordinary run analysis to be random. In greenhouse studies, asparagus seedlings or clones infected with either AV I or AV II became more severely diseased when inoculated by either Fusarium oxysporum f.sp. asparagi (FOA) or F. moni1 iforme (FM) than did virus-free plants. Asparagus plants doubly infected both with AV I and AV II became the most severely diseased when inoculated with FOA. Infection with AV II was determined to affect the asparagus plant in two ways. First, there is an alteration in the composition of the root exudates of virus-infected asparagus plants. Electrolytes, glucose, total carbohydrates and amino acids were detected in much higher levels in the root exudates of virus-infected asparagus plants. These exudates were determined to positively affect the germination and subsequent germ tube growth of conidia from both FM and FOA. Secondly, AV II-infected asparagus plants were determined to be less able to wall-off and lignify the area surrounding the infection courts of either FM or FOA. ACKNOWLEDGEMENTS I wish to thank Dr. Christine Stephens, for serving as my major professor and for her support and guidance throughout my program. I am grateful to Dr. Donald Ramsdell, Dr. M e l v y n Lacy and Dr. Edward Grafius, members of my graduate committee, for their patience and guidance. I am a l s o d e e p l y Indebted to to Tom Stebblns and Jerri Gillett, for their technical assistance, to my wife, Sturges, for her constant encouragement and friendship, and to my many friends for their constructive criticisms. thank you all. I TABLE OF CONTENTS Page LIST OF TABLES............................................. vi LIST OF FIGURES............................................ vii CHAPTER I GENERAL INTRODUCTION AND LITERATURE REVIEW.. 4 ASPARAGUS VIRUSES.................................... 4 Occurrence and Distribution.................... 4 FUSARIUM CROWN AND ROOT R O T ......................... 7 History and Geographic Distribution........... The Causal Agents and Symptomatology.......... 7 11 VIRUS-FUNGUS INTERRELATIONSHIPS..................... 13 OBJECTIVES............................................ 17 LITERATURE 18 CHAPTER II C I T E D ................................. IDENTIFICATION, INCIDENCE AND DISTRIBUTION OF VIRUSES IN MICHIGAN ASPARAGUS............ 24 INTRODUCTION......................................... 24 MATERIALS AND METHODS................................ 27 Field Survey.................................... Isolates and Serology.......................... Physical Properties of Isolates............... Determination of Host Ranges.................. Virus Purification............................. Serologically Specific Electron Microscopy.... Electron Microscopy............................ Antisera Production............................ Gamma Globulin Purification.................... Elisa Conditions................................ 27 28 29 30 30 33 34 34 35 35 RESULTS............................................... Field Survey.................................... Serology........................................ Physical Properties and Host Ranges........... Serologically Specific Electron Microscopy.... Electron Microscopy............................ iii 37 37 41 41 45 49 Page DISCUSSION............................................. 54 LITERATURE CITED...................................... 56 CHAPTER III TRANSMISSION AND SPREAD OF ASPARAGUS VIRUSES WITHIN MICHIGAN ASPARAGUS.................. 58 INTRODUCTION........................................... 58 MATERIALS AND METHODS................................. 61 Aphid Transmission of AV I in Asparagus........ Mechanical Transmission of AV I in Asparagus... Mechanical Transmission of AV II in Asparagus.. Pollen Transmission of AV I I .................... Collection of Asparagus Pollen.............. Tests for Presence of Infective Viruses Identification of AV I I ...................... D e t e c t i o n of A V II A n t i g e n on the Surface of Asparagus Pollen....................... Field Indexing to Determine the Distribution and Extent of AV I and AV II Infection in an Asparagus Plantings.................... 61 61 62 63 63 63 64 RESULTS................................................ Aphid Transmission of AV I in Asparagus........ Mechanical Transmission of AV I in Asparagus... Mechanical Transmission of AV II in Asparagus.. Pollen transmission of AV II in Asparagus..... Association of AV II with Asparagus Pollen.. Localization of AV II Antigen in the Exine of Asparagus Pollen ........................ Extent and Distribution of AV I and AV II within one asparagus planting............... 65 67 70 70 70 70 72 72 75 75 DISCUSSION............................................. 80 LITERATURE CITED...................................... 83 CHAPTER IV INTERACTION BETWEEN ASPARAGUS VIRUSES AND PATHOGENIC FUSARIUM SPECIES IN ASPARAGUS 86 INTRODUCTION........................................... 86 MATERIALS AND METHODS................................. 89 Plant Materials.................................. Preparation of s e e d .......................... Preparation of seedlings..................... Preparation of tissue culture clones........ iv 89 89 89 90 Page Source and Maintenance of Fungi................ Inoculation Procedures.......................... Colonized millet inoculation technique....... Inoculation of plants........................ Preparation of Conidial Suspensions............ Evaluation ofInoculated Plants................. Influence of Virus Infection on Root Rot Severity...................................... Influence of Root Exudates from Healthy and Virus-Infected Asparagus on Root Rot Severity...................................... Collection of Root Exudates from Healthy and Virus-Infected Asparagus Clones............. Chemical Analysis of Root Exudates from Healthy and Virus-Infected Asparagus Clones........................................ The Effect of Root Exudates from Healthy and Virus-Infected Asparagus Clones on the Germination of Conidia and Growth of Germ Tubes......................................... The Effect of Virus Infection on Lignin Formation Within Asparagus Roots in Response to Infection by FOA and F M ...... 92 92 92 92 93 93 98 RESULTS................................................ 99 94 94 95 95 97 Influence of Virus Infection on Root Rot Severity.................................... 99 Influence of Leachates from Healthy and Virus-Infected Asparagus on Root Rot Severity.................................... 99 Changes in Root Exudates of Virus-Infected Asparagus Clones........................... 104 The Effect of Root Exudates from Healthy and Virus-Infected Asparagus on the Germination of Conidia and Germ Tube G r o w t h ........ 104 The Effect of Virus Infection on Lignin Formation Within Asparagus Roots in Response to Infection by FOA and F M .......... 109 DISCUSSION............................................. 114 LITERATURE CITED....................................... 117 V LIST OF TABLES Table 2.1. 2.2. 2.3. 2.4. 2.5. 3.1. 3.2. 3.3. 3.4. 4.1. Page Percent sap-transmissible virus infection as determined with Chenopodium quinoa indicator plants for 12 asparagus fields in Oceana County and 11 fields in V a n Buren C o u n t y ............ 38 Percent sap-transmissible virus infection as determined with Chenopodium quinoa indicator plants for 5 asparagus seedling nurseries in Oceana County, M I ............................ 39 Incidence of asparagus virus I (AV I) as determined with Chenopodium quinoa indicator plants and serologically specific electron microscopy in commercial asparagus plantings in M i c h i g a n a s p a r a g u s p l a n t i n g s .................. 40 Reactions of 12 plant species inoculated with one Michigan isolate of asparagus virus II (A V I I ) ......................................... 44 Reactions of 13 plant species inoculated with one Michigan isolate of asparagus virus I (AV I ) ........................................... 46 Mechanical transmission of asparagus virus II (AV II) in Asparagus officinalis L. using sap or concentrated purified virus preparations.. 71 Detection of antigens and infectious virions of asparagus virus II (AV II) on asparagus pollen and in pollen washes by enzyme-linked immuno­ sorbent assay (ELISA) and r u b - i n o c u l a t i o n ....... 73 Detection of asparagus virus II (AV II) antigens on asparagus pollen, in p ollen washes and within pollen by enzyme-1 inked immunosorbent assay (ELISA)............................................. 74 Localization of asparagus virus II (AV II) antigen in the exine wall of hand-collected asparagus pollen................................... 78 Disease severity of clones of Asparagus officinalis L. cultivar Viking KB3 with and without asparagus virus II (AV II) when inoculated with Fusarium oxysporum f.sp. asparagi. (FOA) and F. monl 1 i forme (FM)........ vi 100 Page 4.2. Effect of infection with asparagus virus I (AV I) or asparagus v ir u s II (AV II) a l o n e or in combinations on disease severity in seedlings of Asparagus officinal is when inoculated with Fusarium oxysporum f.sp. asparagi (FOA) or F. 5*°Si.iA£2HSl® ( F M ) .............................. 101 4.3. Effect of root exudates from healthy and AV IIinfected asparagus clones on the growth of Fusarium oxysporum f.sp. asparagi (FOA) and F. moniliforme (FM)................................... 4.4. Effect of infection with asparagus virus II (AV II) on 1 ignification within asparagus seedling roots in response to infection by Fusarium oxysporum f.sp. asparagi (FOA) or F. o r i e ( F M ) ............................... vii 112 113 LIST OP FIGURES Figure Page 2.1. O u c h t e r l o n y gel d o u b l e d i f f u s i o n test.......... 42 2.2. Ultraviolet absorption profile of a Michigan isolate of asparagus virus II (AV II) sedimented in a 0-30 percent linear-log sucrose density gradient............................................ 47 2.3. Results of immunosorbent electron microscopy with a Michigan isolate of asparagus virus I (AV I ) ............................................ 2.4. 3.1. 3.2. 3.3. 4.1. 50 Electron micrograph of Michigan isolate of asparagus virus II (AV II)........................ 52 Procedure for the localization of asparagus virus II (AV II) antigen on the surface of hand-collected pollen from greenhouse grown Asparagus officinalis L. using protein A-linked l a t e x a n t i s e r u m ( P A L L A S ) ...................... 68 Scanning electron microscopy of hand-collected asparagus pollen after exposure to latex spheres conjugated with asparagus virus II (AV II) antibodies................................. 76 Distribution of asparagus virus I (AV I) and a sparagus v i r u s II (AV II) in a 4 year o l d asparagus planting of v a r i e t y V i k i n g K B 3 ...... 79 Effect of leachates from AV II-infected and virus-free asparagus on the severity of Fusarium c r o w n a n d r o o t r o t ........................... 102 4.2. Increased exudation of electrolytes by AV IIinfected asparagus clones in liquid culture relative to the exudation of virus-free clones... 105 4.3. Increased exudation of glucose, total carbo­ hydrates, amino acids and proteins by AV IIinfected asparagus clones in liquid culture relative to the exudation of virus-free clones... 107 4.4. The effect of root exudates from virus-free and AV II-infected asparagus on the germination of conidia of Fusarium oxysporum f.sp. asparagi (FOA) and F. moniliforme (FM)........... viii 110 CHAPTER I GENERAL INTRODUCTION AND LITERATURE REVIEW Introduction Asparagus (Asparagus officinalis L.) Is an important commercial vegetable crop in a number of areas in Michigan. Michigan is a major producer of asparagus, ranking third in production of asparagus behind California and Washington. In 1983, 20,000 acres were in production in Michigan and seventeen and one-half million pounds of asparagus were harvested with a value of 10.9 million dollars (H. Poster, personal communication). Asparagus is a perennial vegetable crop grown on sandy soils in Michigan, and traditionally it has been thought that, if properly managed, a planting should persist for 16 or more years with yields of 2,000 lbs/acre or more. Today, fields are being removed after 12 to 15 years because plantings have become so sparse and yields so low that they are no longer economical to harvest. "Asparagus decline" occurs nationwide and results in a loss of productivity and longevity of established fields. Most fields are planted with approximately 10,000 crowns per acre but a 1978 survey of Michigan asparagus fields indicated that the average crown population was 3,153 crowns per acre (A. Putnam, personal communication). It Is the author's hypothesis that no single factor is responsible for this decline in vigor and longevity, but rather, that several cultural, environmental or biological stresses on the asparagus plant may predispose it to infection by the Fusarium crown or root rot organisms. Fusarium oxysporum (Schelcht.) emend. Snyder and Hansen f.sp. asparagi Cohen and F. m o n l llforme (Sheldon) emend. Snyder and Hansen are known to be present in all asparagus fields in Michigan and are believed to be the ultimate cause of “asparagus decline". Several viruses have been shown to be of importance in asparagus plantings in both Europe and North America (34, 35, 52, 64, 68). Asparagus vir u s I, a memb e r of the potyvlrus group, is aphid transmitted and widespread in Washington (60) and California (23) and has recently been determined to be present in New Jersey (20) and Michigan (Chapter 2). Asparagus virus II, an ilarvirus, Is widespread in Michigan (33), California (23) and Washington (60) . These viruses may constitute biological stresses to the asparagus plant. In Europe, Weissenfels and Schmelzer (64) reported a 20* reduction in yield in asparagus plantings infected with these viruses. Researchers in Washington (68) have found that plants infected with either asparagus virus I or asparagus virus II exhibited a mild reduction of vigor and productivity. Plants infected with both viruses showed severe decline and mortality in their second year in the field. It Is the purpose of these 3 investigations to evaluate the potential of these viruses as biological stress factors and to elucidate their role and relative importance in "asparagus decline" in Michigan. 4 Literature Review Asparagus Viruses Occurrence and distribution In 1960, Hein (34) first reported a virus isolated from asparagus which produced necrotic local lesions on several species of Chenopodium. This virus, which he designated as asparagus virus I (AV I) (35), was determined to be widespread in asparagus plantings in Germany. Hein (35) reported that asparagus virus I produced no symptoms on asparagus and that the virus was transmitted by several different aphids. Mink and Uyeda (47) in 1977 isolated a long flexous-rod shaped particle, from Washington asparagus. 700 to 880 nm in length Their isolate was mechanically transmissible only to species of Chenopodium and symptoms on C. quinoa and particle morphology were similar to those reported for asparagus virus I by Hein (34, 35). They reported that nearly every field grown asparagus plant tested contained asparagus virus I. Uyeda (60) reported that asparagus virus I was present in all fields surveyed, except one and that the incidence of infection was related to the age of the planting. In limited attempts Uyeda (60) was unable to transmit asparagus virus I with three aphid species; Myzus perslcae Sulz. (green peach aphid), Rhopa1osiphum pad! (L.) (oat-bird cherry aphid) and Cava r i e l la aeqopodll (Scopoll). Fujisawa et al (25) reported the occurrence of a long flexous rod-shaped virus particle in asparagus in Japan 5 similar to that reported by Hein (34). On the basis of particle morphology and its limited host range and symptomatology in those hosts, it was determined to be asparagus virus I. The Japanese isolate also produced laminate pinwheel inclusions within infected hosts and was transmissible by the green peach aphid. For these reasons, it was considered to be a member of the potyvirus group. Asparagus virus I was consistently isolated from asparagus in New Jersey by Davis and Garrision (20). They reported that asparagus plants grown in the field or greenhouse were Infected without obvious symptoms and there was no apparent correlation between the vigor of asparagus clones in tissue and infection by the virus. California asparagus for viruses, In a s u r v e y of Fa 1 loon et al (23) found asparagus virus I throughout the state and in all cultivars tested. The occurrence of asparagus virus II was first reported in Germany by Hein (35) and later by Weissenfels et al (64) and appears to be identical with asparagus latent virus described subsequently in Denmark (49). Uyeda and Mink (61) first reported the presence of asparagus virus II (AV II) in Washington asparagus and demonstrated that it was an isometric particle with a diameter of 23 to 33 nm and serologically related to citrus variagation virus, citrus leaf-rugose virus and elm mottle virus. Uyeda (60) demonstrated that AV II was widespread in Washington, Isolating it from all asparagus plantings surveyed and from more than half of the plants tested in 6 some fields. Uyeda and Mink (61) were unable to Infect asparagus seedlings in the greenhouse by manual inoculation. Further, they demonstrated that the virus was readily seed transmitted in asparagus and that it could be separated into three major nucleoprotein components in a rate zonal sucrose density gradient. Each component alone exhibited little or no infectivity and a maximum infectivity was achieved when all three components were present in the inoculation mixture. Also, they identified two serotypes of AV II and d e s i g n a t e d them A V II-S and A V II-P. diffusion tests, In agar gel d o u b l e the European isolate of A V II produced a precipitin line that coalesced with that of AV II-P but formed a spur with A V II-S w h e n tested against a n t i s e r u m to AV II-P and AV II-S. Because of these properties they placed A V II in the llarvlrus (isometric labile ringspot virus) group. Fujisawa et al (26) isolated AV II from symptomless asparagus plants in Japan and their isolate was determined to be more closely related serologically to AV II-S than AV II-P. Asparagus virus II was found to occur in almost all asparagus plantings in Hokkaido. They were unable to infect asparagus seedlings by manual Inoculation with sap from AV II-infected C. quinoa and tobacco but could infect asparagus with concentrated purified AV II. Asparagus shoots infected in this manner sometimes developed a light-green mosaic after about 20 days. Recent reports from California (23) and Michigan (33) 7 Indicate that AV II is widespread in the major asparagus growing regions of the United States and has also been detected at low levels in New Jersey asparagus plantings (R. Davis, personal communication). Fusarium Crown and Root Rot History and geographic distribution The first report of a Fusarium-incited disease of asparagus was made by Stone and Chapman in Massachusetts in 1908 (55). They observed a wilt of young shoots during the harvest season followed by a premature yellowing and rotting of the mature stalks or "ferns" later in the growing season. They consistently isolated an unidentified species of Fusarium from the affected plants. Cook, in 1923, described a similar disorder of asparagus in New Jersey, which he called "dwarf asparagus" (17). He reported a slow dying out of asparagus plants over circular areas and a concomitant stunting and premature yellowing of the affected plants. An unidentified Fusarium was always associated with the diseased plants. Over the next two decades numerous reports of Fusarium root and crown rots and wilts were made from New Jersey (62), Oregon (13), Massachusetts (12), North Carolina (2), South Carolina (4), Pennsylvania (41), Missouri (55), Illinois (1), and Idaho (10). In 1941, Cohen and Heald (16) observed a wilt and root rot of asparagus in many of the irrigated, sandy soil regions of Washington State. Their investigation demonstrated that the common soil-borne fungus, Fusarium oxysporum Schlecht., was the causal agent of the disease. Pathogenicity tests with their isolate of F. oxysporum from asparagus indicated that it was different from other isolates already described. Their asparagus Isolate was unable to produce disease in potato, tomato, carnation or onion and o n l y a s l i g h t wil t in A l a s k a n peas. It was a b l e to "vigorously" infect only asparagus and all asparagus varieties tested were equally susceptible. Cohen (15) distinguished the Isolate as F. oxysporum variety asparagi. Some years later Armstrong and Armstrong (5) concluded that this organism was the very specific pathogen of asparagus and called it F. oxysporum f.sp. asparagi. A Fusarium-Incited disease of asparagus seedlings was reported in Canada by Graham in 1955 (30). Seedling nurseries were so severely affected that they were destroyed and the land was used for other purposes. pathogen as F. oxysporum f.sp. redolens He identified the (Wr.) Gordon and first described the infection process in asparagus seedlings. The course of infection was determined by examining sections of several hundred rootlets. Penetration took place between the walls of epidermal cells primarily in the meristematic region of the root tip and through the stomata of the hypocotyl and coleoptile. This was subsequently confirmed by Shoemaker (57). Graham also suggested that the pathogen may take advantage of natural openings on more mature parts of seedlings. In mature crowns he demonstrated that vascular discoloration was associated primarily with a wound, leading him to suggest that this organism was a "wound parasite". Endo (personal communication) later demonstrated that asparagus storage roots were capable of rapidly walling off Fusarium infections and wounds which could serve as potential infection sites. Further, he suggested that stress from overpicking might reduce the ability of these roots to wall off invading pathogens. Graham was also able to Isolate F. monil lforme from diseased seedlings and demonstrated that this fungus caused root tip necrosis but also concluded that it was not Important in seedling blight. No comparable disease could be found in surrounding production fields, although growers had reported unexplained reductions in yield for years. Grogan and K i m b l e in 1959 (31) a l s o reported a s l o w decline in the productivity of asparagus plantings in California and an inability to re-establish productive plantings in these areas. The asparagus decline and replant problem in California was attributed to the most prevalent pathogenic organism associated with the disorder, F. oxysporum f.sp. asparagi which was determined to be present on the seed. They concluded that the Fusarium wilt of asparagus in California was probably identical with the disease described by Cohen and Heald (16), who, in fact, identified the causal organism from infected asparagus plants received from California. Lewis and Shoemaker (43) screened a number of plant introductions of the genus Asparagus for resistance to a New 10 Jersey Isolate of F. oxvsporum f.sp. asparagl and found all except A. sprengerl Regel to be highly susceptible. A. sprengerl was determined to be immune to their Isolate. The first association of F. mon i liforme with asparagus decline in California was made by Endo and Burkholder (22). F. mo n i Ilforme was consistently isolated from brown, dry rotting asparagus crowns and Johnston et al in 1979 (39) asserted that F. mo n i liforme should be considered the pathogen of this completely separate disease of asparagus. This disease is characterized by a total collapse of older root tissue and proposed calling this disease Fusarium stem and crown rot. Asparagus decline was first reported in Michigan by Lacy in 1979 (42) and he also attributed the decline ultimately to the presence of F. oxvsporum f.sp. asparagl and F. m o n i liforme. He suggested that the decline was due to environmental stresses caused primarily by management practices such as overpicking, unsatisfactory weed control, mechanical damage from disking prior to harvesting and lack of Irrigation which might predispose crowns to Infection by F. oxvsporum f.sp. asparagl and F. moniliforme. Asparagus seed has a very rugged surface and the epidermal cells of its seed coat are tubular in shape and are separated by very deep crevices. demonstrated, Inglis (38) in 1980 using scanning electron microscopy, that F. oxvsporum f.sp. asparagl and F. mo n i liforme were external seed contaminants and that the inablity to surface-disinfest asparagus seed might be due to the rough seed surface and 11 the lodging of fungal spores in these natural crevices and in tunnels produced by Insects. She further concluded that asparagus seeds became Infested by these fungi during the commercial seed extraction process. Damicone and Cooley (19) developed a reliable method for the production of large numbers of disease-free asparagus seedlings for use in the laboratory and greenhouse. They routinely used 25,000 mg/ml benomyl in acetone for 24 hours to surface-dlsinfest seed and found that both Fusarium species were eradicated from asparagus seeds with a concomitant increase in seed germination. Gilbertson and Manning in 1983 (28) confirmed what Booth had suggested in 1971 (4); that wind was an important vehicle for dissemination of F. mo n i 11forme spores. They concluded that airborne spores of this fungus were the source of contamination of asparagus flowers, fruit, and ultimately, the seed. They also determined that the sources of airborne inoculum Included asparagus stems and corn ears and stalks Infected with F. m o n i liforme and Gilbertson (27) reported that this potential inoculum on Infected asparagus stems increased with plant age. Manning et al (45) suggested that latent, undetected Infections of both Fusarium species may be prevalent in asparagus seedlings and that several environmental stresses may be responsible for expression of latent infections. The causal agents and symptomatol ogy According to Messiaen et al (46), Fusarium oxvsporum 12 (Schelcht.) emend. Snyder and Hansen and F. moniliforme (Sheldon) emend. Snyder and Hansen are ubiquitous pathogens and play a major role in reducing yields and quality of major food crops of the world. Fusarium spp. are facultative parasites which colonize living and non-living tissue and may invade non-host tissue (3, 36). characteristic, This along with their ability to form chlamydospores or other resting structures which can survive in soils for many years make these fungi especially persistent once established (32, 48). Typically, F. monil iforme forms sporodochla and macroconidia in small numbers and produce an abundance of microconidia in chains (46). Members of this species do not form chlamydospores, but instead form individual thickened hyphae as survival structures (48). F. oxvsporum is the most frequently isolated species of Fusarium in soils and is characterized by its ability to form chlamydospores, macroconida and microconidia (46). Both causal agents associated with asparagus decline cause a yellowing, stunting or wilting of mature stalks that may die at various stages of elongation but, they produce this effect by slightly different means (15, 22). Fusarium oxvsporum f.sp. asparagl is known to produce vascular discoloration within stems, roots and crowns as w ell as reddish eliptical lesions on roots and the basal part of stems (31). This results in a vascular wilt that inhibits the movement of water and carbohydrates within the affected plant. Fusarium moniliforme is primarily a root and crown 13 rotting organism that causes extensive dry crown rotting and a brown stem pith discoloration and a total root collapse in some asparagus seedlings (39). Infection of asparagus roots by F. moniliforme is not associated with vascular discoloration (15, 39). Virus-Fungus Interrelationships The effects of virus Infection on fungus disease has been characterized by numerous researchers. Fungal infection may be increased, decreased or not affected by virus infection of a plant dependent on the system studied. In general, Infection by facultative parasites of virus infected plants is increased and infection by obligate parasites, as the rusts, smuts and powdery mildews, is decreased. such The literature concerning virus-fungus interrelationships has been reviewed in detail by Beute (7). Virus infection was demonstrated to enhance post­ emergence damping-off caused by Rhizoctonla sp. in cucumber. Bateman (6) found that the incidence of post-emergence damping-off was 10* to 15* for virus-free cucumber seedlings and 60* to 87* in cucumber seedlings previously Inoculated with cucumber mosaic virus. The greatest synergism between pathogens occurred when small quantities of Rhlzoctonia inoculum were used. Cotyledons showed an increased rate of respiration 48 hours after inoculation with the virus and Bateman suggested that the movement of materials from the root to the virus-infected cotyledons may Increase the susceptibllty of the roots to attack by the fungus. 14 An increase in the severity of several different root rot diseases in virus-infected plants has been reported for wheat (58), corn (65), red clover (21, 63), white clover (44), white lupine (50) and sugarbeets (18) grown under field conditions. Smith (58) reported that root rots of wheat incited by Fusarium and Rhizoctonla occurred predominately in plants infected with barley yellow dwarf virus. Corn seedlings Infected with maize dwarf mosaic virus were determined to be more susceptible than virus-free seedlings to root rots, stalk rots and seedling blights incited by a number of pathogens (65). Also, Pythium qramlnlcola Subram. caused more severe root rot in corn and wheat seedlings infected with a virus resembling wheat streak virus. Similarly, Watson and Guthrie (63) observed a severe root rot, general decline of vigor, and degeneration of the root system of red clover plants under field conditions. They noted that severe root rot developed when plants were infected with clover yellow mosaic virus alone or with white clover mosaic virus and any one of several fungal parasites that were usually only mildly pathogenic to red clover. In 1964, Farley and Lockwood (24) suggested that viruses may p l a y a role in the d e v e l o p m e n t of fungus root rots in peas. They demonstrated that three pea varieties were more susceptible to two common fungus root rot diseases when plants were Inoculated with any of four different viruses. Similar results were obtained over a wide range of greenhouse environmental conditions with different strains of two viruses, plants of different ages and different time Intervals between virus and fungus inoculations. Lockwood (9), working with the same system, Beute and determined that the increased Fusarium root rot in peas infected with bean yellow mosaic virus was not due to an Increased susceptibllty of root tissue but rather that virus infection increased the exudation of nutrients from roots thereby increasing the inoculum potential of the pathogen in the rhizosphere. The roots of virus-infected pea plants released more electrolytes, carbohydrates, amino acids and nucleotides than did the roots of healthy plants. They attributed this Increased exudation to an Increase in the permeability of cell membranes of the root. Beute (8) demonstrated that decreased yield of flowers, spikes, corms and cormels and an increase in plant mortality in Gladiolus was a result of the increased susceptibility of virus-infected plants. Chronic infection with cucumber mosaic virus or tobacco ringspot virus increased the prevalence and sometimes the severity of Fusarium and Stromatinla root rot diseases, Curv ularla leaf spot disease, and storage rot of corms in Gl adiolus. He concluded that even though the extent of virus damage was not immediately apparent, it was without doubt a major factor in the decline of G l adiolus stock. The first interaction between systemic fungal Infection and viral pathogens in crucifers was reported by Reyes and Chadha (54). Their data indicated that the severity of cabbage yellows disease in Brasslea campestrls var. chinensls was increased when plants were inoculated with turnip 16 mosaic virus regardless of the interval between fungus and virus inoculations. Virus-infected plants became more severely stunted when inoculated with the fungus than plants infected with the virus alone. Pieczarka (51) noted a synergistic relationship between Rhizoctonia damping-off in peppers and the concomitant infection with one or more of the prevalent pepper viruses. However, the nature of the relationship varied depending on which pepper variety was used. Infection of pepper variety Early Calwonder with strain P of tobacco mosaic virus (TMV-P) resulted in an Increase in susceptibility to Rhizoctonia damping-off while mixed viral infections with TMV-P and pepper mottle virus (PeMV) were no more effective than a single infection in increasing pepper susceptibility to this fungal pathogen. In contrast to positive interactions discussed above, Wilson (66) noted a cross-protection of bean plants infected with T M V to bean rust disease. Goheen and Schnathorst (29) reported that virus leafroll in affected grapevines had an increased resistance to powdery mildew. Antagonistic relationships between viral infection and fungal disease have been reported by investigators In cucumber (37), tobacco (67), red clover (40), and pigeon pea (14). In all cases described above the cross-protection was afforded by an obligate parasite. It is clear that infection of some plant species with one or more plant viruses can increase the incidence and severity of certain root rot diseases. Because asparagus Is a perennial crop and F. oxysporum f.sp. asparagl and F. 17 moni1 iforme are ubiquitous pathogens, infection with one or more of the asparagus viruses could have severe consequences on the longevity and productivity of asparagus plantings. Objectives The objectives of this research were: to determine which asparagus viruses were present in Michigan asparagus plantings (Chapter 2), to determine their distribution and incidence in the asparagus growing regions of the state (Chapter 2), to evaluate possible methods of transmission and spread of these asparagus viruses within asparagus (Chapter 3), to evaluate the effect of virus infection on the severity of Fusarium crown and root rot (Chapter) and to investigate the mechanisms by which asparagus viruses may contribute to "asparagus decline" (Chapter 4). 18 Literature Cited 1. Anonymous. 1922. Diseases of field and vegetable crops In the U.S. In 1921: Asparagus root rot caused b Y F u s a r ium sp.. page 402 in: P l a n t Dls. Rep. Suppl. 22. G. H. Coons ed. 2. Anonymous. 1927. Diseases of vegetable and field crops other than cereals in the U.S. In 1926. page 325 in: P l a n t Dls. Rep. Suppl. 54. R. J. H a s k e l l and J. I. Wood, eds. 3. Alexander, biology. 4. Armstrong, G. M. 1930. Fusarium sp. on asparagus in South Carolina. Plant Dis. Rep. 14:197. 5. Armstrong, G. M., and Armstrong, J. K. 1969. Relationship of F. oxvsporum formae speciales apl l , asparagl, casslae, m e 1ongenae, and v aslnfectum race 3 as revealed by pathogenicity of common hosts. Phyto­ pathology 59:1256-1260. 6. Bateman, D. F. 1961. Synergism between cucumber mosaic virus and Rhizoctonia in relation to rhizoctonia damping-off of cucumber. (Abstr.) Phytopathology 5 1 :574 7. Beute, M. K. 1967. Mechanism of increased root rot in virus-infected peas. Ph.D. Thesis. Michigan State University. 89 pp. 8. Beute, M. K. 1970. Effect of virus infection on susceptiblity to certain fungus diseases and yield of Gladiolus. Phytopathology 60:1809-1813. 9. Beute, M. K., and Lockwood, J. L. 1968. M e c h a n i s m of Increased root rot in virus-infected peas. Phyto­ pathology 58:1643-1651. M. 1961. Introduction to soil micro­ John Wiley and Sons, New York. 345 pp. 10. Blodgett, E.C. Idaho in 1943. 29:465. 1945. Diseases of various crops in Miscellaneous crops. Plant Dis. Rep. 11. Booth, C. 1971. The Genus Fusarium. Commonwealth Mycological Institute, Kew, Surrey, England. 237 pp. 12. Boyd, 0. C. 1942. Recent obserevatlons on plant disease in Massachusetts. Plant Dis. Rep. 26:334. 13. Boyle, L. W. 1943. Plant disease survey of Oregon in 1943. Plant Dis. Rep. Suppl. 149:386-395. 19 14. Chadha, K. C., and Raychaudhuri, S. P. 1964. I n t e r ­ action between sterility virus and Fusarium udum Butl in Pigeon-pea. Indian J. of Agric. Sci. 36:133-139. 15. Cohen, S. I. 1946. A w i l t and root rot of Asparagus officinalis L. var. a t i lis L. (Abstr.) Phytopathology 36:347. 16. Cohen, S. I., and Heald, F. D. 1941. A w i l t and root rot of asparagus caused by Fusarium oxvsporum Schlecht. Plant Dis. Rep. 25:503-509. 17. Cook, M. T. 13:284. 18. Crane, G. L., and Calpouzos, L. 1969. S y n e r g i s m of Cercospora beticola and beet yellows virus in killing sugar beet leaves. Phytopathology 57:808-809. 19. Damicone, J. P., Cooley, D. R., and Manning, W. J. 1981. Benomyl in acetone eradicates Fusarium moniliforme and E\. oxvsporum from asparagus seed. Plant Dis. 65:892-893. 20. Davis, R. F., and Garrison, S. 1985. First report of a virus present in New Jersey. Plant Dis. 66:628. 21. Denis, S. J., and Elliott, E. S. 1967. D e c l i n e of red clover plants infected with red clover vein mosaic virus and Fusarium species. (Abstr.) Phyopathology 57:808-809. 22. Endo, R. M., and Burkholder, E. C. 1971. The a s s o c ­ iation of Fusarium m o n i 11forme with the crown rot com­ plex of asparagus. Phytopathology 99:122-125. 23. Falloon, P. G., Falloon, L. M., and Grogan, R. G. 1985. A survey of California asparagus for asparagus virus I (AVI), asparagus virus II (AVII) and tobacco streak virus (TSV). Plant Dis. (in press). 24. Farley, J. D., and Lockwood, J. L. 1964. Increased susceptibility of root rots in virus infected peas. Phytopathology 54:1279-1280. 25. Fujisawa, I., Goto, T., Tsuchizaki, T., and Iizuka, N. 1983. Host range and some properties of asparagus virus I isolated from Asparagus officinalis in Japan. Ann. Phytopath. Soc. Japan 49:299-307. 26. Fujisawa, I., Goto, T., Tsuchizaki, T., and Iizuka, N. 1983. Some properties of asparagus virus II iso­ lated from Asparagus officinal is in Japan. Ann. Phyto­ path. Soc. Japan 49:683-688. 1923. Dwarf asparagus. Phytopathology 20 27. Gilbertson, R. L. 1981. Sources of inoculum and disease increase of stem, crown and root rot of asparagus caused by Fusarium oxvsporum and Fusarium moniliforme. M.S. Thesis, University of Massachesuetts. 169 pp. 28. Gilbertson, R. L., and Manning, W. J. 1983. C o n t a m ­ ination of asparagus flowers and fruit by airborne spores of Fusarium monil iforme. Plant Dis. 67:10031004. 29. Goheen, A. C., and Schnathorst, W. C. 1961. R e s i s t a n c e to powdery mildew in leaf r o l 1 -affected grapevines. Plant Dis. Rep. 45:641-643. 30. Graham, K. M. 1955. Seedling blight, a fusarium disease of asparagus. Can. J. Bot. 33:374-400. 31. Grogan, R. G., and Kimble, K. A. 1959. The a s s o c ­ iation of Fusarium wilt with the asparagus decline and replant problem in California. Phytopathology 99:122125. 32. Hart, L. P., and Endo, R. M. 1976. The reappe a r a n c e of Fusarium yellows of celery in California. Plant Dls. Rep. 62:138-142. 33. Hartung, A. C., Evans, T. A., and Stephens, C. T. 1985. Occurrence of asparagus virus II in commercial asparagus fields in Michigan. Plant Dis. 69:501-504. 34. Hein, A. 1960. Uber das vorkommen einer vlrose an spargel. Z. Planzenkr. Pflanzenpathol. Pflanzenschutz 67:217-219. 35. Hein, A. 1963. Viroses an spargel. Mitt. Biol. Bundesanst. Land-Forstwirtsch, Berlin-Dahlem 108:70-74 (Rev. Appl. Mycol, 1964. 43:455). 36. Hendrix, F. F., and Nielson, L. E. 1958. I n v a s i o n and infection of crops other than formae suscept by Fusarium oxvsporum f. butas and other formae. Phytopathology 48:224-228. 37. Hopen, H. J., and deZeeuw, D. J. 1962. R e d u c t i o n of susceptibility to cucumber scab by cucumber mosaic virus. Plant Dis. Rep. 46:93-97. 38. Inglls, D. A. 1980. Contamination of asparagus seed by Fusarium oxvsporum f.sp. asparagl and Fusarium monil iforme. Plant Dis. 64:74-76. 21 39. Johnston, S. A., Springer, J. K., and Lewis, G. D. 1979. Fusarium moniliforme as a cause of stem and crown rot of asparagus and Its association with asparagus decline. Phytopathology 69:778-780. 40. King, L. N., Hamptom, R. E., and Diachun, S. 1964. Resistance to Erysiphe polygonl of red clover Infected with bean yell o w mosaic virus. Science 146:1054-1055. 41. Kirby, R. S. 1943. Reports on diseases of miscellan­ eous vegetable crops. Plant Dls. Rep. 27:447. 42. Lacy, M. L. 1979. Effects of chemicals on stand estab­ lishment and yields of asparagus. Plant Dls. Rep. 63:612-616. 43. Lewis, G. D., and Shoemaker, P. B. 1964. Resistance of asparagus species to Fusarium oxvsporum f.sp. asparagl. Plant Dls. Rep. 48:364-365. 44. McCarter, S. M., and Halpin, J. E. 1961. Studies on the pathogenicity of four species of soil fungi on white clover as affected by the presence of bean yellow mosaic virus under conditions of controlled temperature and light. (Abstr.) Phytopathology 51:644. 45. Manning, W. J., Damlcone, J. P., and Gilbertson, R. L. 1979. Asparagus root and crown rot: Sources of inoculum for Fusarium oxvsporum and Fusarium moniliforme. Pages 195-205 in: G. Reuther, ed. Proc. 5th Int. Asparagus Symp., Geisenhelm, West Germany. 46. Messiaen. C.M., and Cassini, R. 1981. Taxo n o m y of Fusarium. pages 427-445 in: Fusarium: Diseases, Biology and Taxonomy. P. E. Nelson, T. A. Touss o u n and R. J. Cook, eds. Pennsylvania State University Press, Univer­ sity Park 457 p. 47. Mink, G. I., and Uyeda, I. 1977. Three mechanically transmissible viruses isolated from a s p a r a g u s in Washington. Plant Dis. Rep. 61:398-401. 48. Nyvall, R. F., and Kommedahl, T. 1968. Individual thickened hyphae as survival structures of Fusarium m o n i liforme in corn. Phytopathology 58:704-707. 49. Paluden, N. 1964. Vlrussygdomme hos Asparagus officinalis Mannedsovers. Platesygd. 407:11-16. 50. Patil, P. L. 1973 Increased susceptibility to root and stem rot in a virus-infected white lupine (Lupinus a l bus L.) Maharashtra Vidnyas Mandlr Patrika 8:24-31. (Rev. Plant Pathol. 54:629). 22 51. Pieczarka, D. J., and Zitter, T. A. 1981. Effect of Interaction between two viruses and Rhizoctonia in pepper. Plant Dis. Rep. 65:404-406. 52. Posnette, A. F. 1969. Nematode transmitted viruses in asparagus. J. Hortica. Sci. 44:403-408. 53. Raju, D. G., Sill, W. H., an d Browder, L. E. 1969. The combined effects of two viral diseases and leaf rust on wheat. Phytopathology 59:1488-1492. 54. Reyes, A. A., and Chadha, K. C. 1972. I n t e r a c t i o n between Fusarium oxvsporum f.sp. conglutlnans and turnip mosaic virus in Brassica campestris var chinensis seedlings. Phytopathology 62:1424-1428. 55. Scott, I. T. 1927. Diseases of vegetable and crops other than cereals in the U.S. in 1927. Dis. Rep. Suppl. 61:294. field Plant 56. Schade, C. 1969. Viruskrankheiten des spargels. Nachrlchtenbl. Dtsch. Pflanzenschutzdienst. (Berlin), 23:38-40. 57. Shoemaker, P. B. 1965. A comparative histological study of the penetration and infection process in seedlings of Asparagus L. by two isolates of Fusarium oxvsporum f. asparagl. Cohen and Heald. M.S. Thesis, Rutger s - T h e State Univ., N. J. 65 pp. 58. Smith, H. C. 1962. Is barley yellow dwarf a pre­ disposing factor in common root rot of wheat in Canada? Can. J. P l a n t Dis. Surv. 42:143-148. 59. Stone, G. botanist. 60. Uyeda, I. 1978. Identification, characterization, and incidence of viruses isolated from asparagus. Ph.D. thesis, Washington State University. 115 pp. 61. Uyeda, I., and Mink, G. I. 1981. P roperties of asparagus virus II, a new member of the Ilarvlrus group. Phytopathology 71:1264-1269. 62. Walker, E. A. 1943. Reports of diseases of miscellaneous vegetable crops. Plant Dis. Rep. 27:447. 63. Watson, R. D., and Guthrie, J. W. 1964. V i r u s - f u n g u s interrelationships in a root rot complex in red clover. Plant Dis. Rep. 48:723-727. E., and Chapman, G. H. 1908. Report of the Massachusetts Agr. Exp. Stat. Rep. 20:127. 23 64. Weissenfels, M., and Schmelzer, K. 1976. Untersuchungen ueber das schadausmass durch viren am spargel (Asparagus officinalis L.). Arch. Phytopathol. Pflanzenschutz, Berlin. 12:67-73. 65. W illiams, L. E., and Alexander, L. J. 1965. M a i z e dwarf, a new corn disease. Phytopathology 48:127-128. 66. Wilson, E. M. 1958. Rust-TMV cross protection and necrotic-ring spot reaction in bean. Phytopathology 48:127-128. 67. Wood, H. N., and Braun, A. C. 1961. Studies on the regulation of certain essential biosynthetic systems in normal and crown-gall tumor tissue. Proc. Nat. Acad. Sci. 47:1907-1913. 68. Yang, H. 1979. Early effects of viruses on the growth and productivity of asparagus plants. HortScience 14:734-735. CHAPTER II IDENTIFICATION, INCIDENCE AND DISTRIBUTION OF VIRUSES IN MICHIGAN ASPARAGUS PLANTINGS Introduction Numerous viruses have been reported to Infect asparagus (Asparagus officinalis L.) In both Europe and North America (11, 12, 16, 20, 21). In 1960, H e i n (12) first reported a virus isolated from asparagus which produced necrotic local lesions on several species of Chenopodium. This virus, which he designated as asparagus virus I (AV I) (13), was determined to be widespread in asparagus plantings in Germany. Hein reported that A V I produced no symptoms in asparagus and suggested that it was transmitted by several different species of aphid (14). Mink and Uyeda (17) isolated a long flexous rod-shaped particle, length, from Washington asparagus in 1977. 700 to 880 nm in Based on particle morphology and symptomatology on C. qulnoa they determined it to be s i m i l a r to A V I r e p o r t e d in G e r m a n y and reported that nearly every field grown asparagus plant in Washington was Infected with AV I. In limited attempts, Uyeda (17) was u n a b l e to transmit A V I in a s p a r a g u s by gre e n pea c h aphid (Myzus perslcae Sulz.), padl oat-bird cherry aphid (Rhopa1osiphum (L.)) or Cav a r l e l la aegopodli (Scopoli). 24 25 Fujisawa et al (8) In 1983, reported the occurrence of a long flexous rod-shaped virus particle in asparagus in Japan similar to that reported by Hein (11) in Europe. basis of particle morphology, On the its limited host range and its ability to produce laminate pinwheel inclusions within infected hosts it was d e t e r m i n e d to be A V I and a member of the potyvirus group (8). Fujisawa et al (8) were able to transmit their isolate of AV I with green peach aphids (Mvzus persicae) but not with melon aphids (Aphis qossypii). Asparagus virus I was consistently isolated from numerous asparagus plantings in New Jersey (4), and detected in asparagus plantings in Michigan (this manuscript) and California (7). Asparagus virus I has also been isolated from asparagus plantings in New Zealand where its incidence has been reported to increase to 90 or 100% of a p l a n t i n g in 3 years (6). The occurrence of asparagus virus II (AV II) was first reported in 1963 by Hein (12) in Germany and later by Weissenfels and Schmelzer (20) and appears to be identical asparagus latent virus described subsequently in Denmark (15). In 1981, Uyeda and Mink (18) reported that AV II was widespread in asparagus in Washington and demonstrated that it was an isometric particle, 23 to 30 nm in diameter, seed transmitted and belonged to the ilarvirus (isometric labile ringspot virus) group. All attempts by Uyeda (17) to mechanically transmit AV II to asparagus were unsuccessful. Recent reports from California (7), Michigan (10), New Zealand (6), and Japan (9) indicate that AV II is widespread 26 in many of the major asparagus growing regions of the world. Tobacco streak virus (TSV), another ilarvirus, has been reported to be of importance in the profitable production of asparagus (15). Its occurrence within asparagus has been reported in Denmark (15), England (1) and Washington State (14) . It was the o b j e c t i v e of this study to assess the incidence and d i s t r i b u t i o n of A V I, A V II and TS V within Michigan asparagus plantings. 27 Materials and Methods Field Survey During the summers of 1982, 1983, 1984 and 1985, commercial asparagus plantings in Michigan were indexed for the presence of A V I, A V II and TSV. In the 1982 h a r v e s t season, 23 production plantings in four different age groupings were surveyed, Including 12 plantings in Oceana County and 11 plantings in Van Buren County. In addition, six seedling nurseries in Oceana County were indexed for virus. An experimental plot (100 x 100 m) was established 100 m d i a g o n a l l y in from a corner of each p l a n t i n g and 20 to 30 spears or, in the case of seedlings, at random from each plot. ferns were collected S a m p l e s wer e stored at 4 C until being triturated in 0.02 M sodium phosphate buffer, pH 7.0, and rub-inoculated to Chenopodlum qulnoa Willd. Indicator p l a n t s were m a i n t a i n e d in a g r e e n h o u s e at 21 to 24 C and assessed for symptom development 5 to 14 days after Inoculation (17). During the summers of 1983 and 1984, 20 randomly selected asparagus plants from each of 20 fields in Oceana County were Indexed for the p r e s e n c e of A V I and T S V as described above. During the harvest season of 1985, ten asparagus plantings in Oceana County and seven plantings in Van Buren were s u r v e y e d for the presence of A V I. One of the plantings in Oceana County suspected of being Infected with AV I was intensively surveyed to accurately assess the percent Infection of the virus within that field. One 28 hundred plants, In a grid pattern, were selected and two spears were removed from each plant and indexed for virus on C. qulnoa and with serologically specific electron microsopy (SSEM) for A V I (5). Isolates and Serology Isolates were established from indicator plants exhibiting symptoms typical of AV I or A V II (17). Asparagus virus I typically produces necrotic lesions on inoculated l e a v e s of C q u lnoa in 10 to 14 days and A V II produces chlorotic ringspots on inoculated leaves of C. quinoa in 5 to 10 days followed the development of a systemic mottle. During the spring of 1982, an Isolate was established from single, chlorotic lesions developing on C. qulnoa leaves in 8 days and designated Isolate A. Lesions were transferred sequentially three times on C. qulnoa to assure that symptom d e v e l o p m e n t was due to a s i n g l e virus. In the summer of 1984, an isolate was established from single local necrotic lesions developing on C. quinoa leaves after 12 days. This isolate was transferred three times as previously described and designated Isolate B. Plants were maintained in a growth chamber or greenhouse at 21 to 24 C with a 16 hour photoperiod. Michigan isolate A was used in agar gel double diffusion tests against antisera prepared against tobacco streak virus (strains BRN and HF), tomato rlngspot virus, tobacco rlngspot virus (blueberry isolate), tobacco rlngspot virus (tobacco isolate) [originally from G. Gooding], peach rosette mosaic 29 virus, A V I and A V II [ provided by G. Mink], antisera were kindly provided by D. Ramsdell. A l l other All double diffusion serology tests Included known positive controls for their respective virus and a healthy sap control. Serology plates were prepared by pouring 10 ml of autoclaved 0.8* agarose (w/v) containing 0.85* sodium chloride (w/v) and 0.1* sodium azide (w/v) into petri plates 90 mm in diameter. Wells were cut in agar immediately before use with Grafar auto-gel cutter [Grafar Corporation, Detroit, MI 48232]. Physical Properties of Isolates Isolates A and B were characterized by determining dilution end-point, thermal inactivation point and longevity in v itro. Dilution end-point was carried out on 5 g samples of infected C quinoa. Leaves were triturated in 0.01 M sodium phosphate buffer, layers of cheesecloth. pH 7.0, and strained through two Samples were diluted with glass distilled water to 10-5 and each dilution was rub-inoculated onto two C. qulnoa plants at the four to six leaf stage. The thermal inactivation point for each Isolate was determined by triturating 5 g of Infected C. qulnoa leaf tissue. The homogenate was strained through two layers of cheesecloth and 1-ml samples placed into test tubes. These samples were heated in a water bath for 10 min at temperatures ranging from 50 to 80 C at 5 d e gr e e increments, then q u i c k l y c o o l e d in ice water. qulnoa plants. Each sample was rub-inoculated onto two C. For longevity in v itro tests, 15 g samples of infected C. quinoa were triturated in glass distilled water 30 and the homogenates were maintained at 21 to 24 C. Sub­ samples of this preparation were rub-inoculated onto C. guinoa at i n t e r v a l s up to 14 days. P l a n t s used in the determination of physical properties were assessed for symptom development 5 to 14 days following inoculation. Determination of Host Range The host range for i s o l a t e s A and B were d e t e r m i n e d and compared with those reported by Uyeda (17). Partially purified preparations of isolate A were rub-inoculated onto single leaves of 12 plant species previously demonstrated to produce symptoms in response to mechanical inoculation with the AV II. The host range of isolate B was assessed by first triturating spear tips of Michigan asparagus plants known to be infected o n l y wi t h i s o l a t e B and the sap was d i l u t e d 1:2 (w/v) with 0.02 M sodium phosphate buffer, pH 7.0. Single leaves of 13 plant species were dusted lightly with carborundum (320 mesh) and rub-Inoculated with this preparation. Test plants were maintained in a greenhouse at 21 to 24 C and a 16 hour photoperiod and assessed for symptom development 5 to 14 days after inoculation. Plants suspected of being Infected were back inoculated to C. quinoa for confirmation. Virus Purification Isolate A, thought to be AV II, was purified from systemically infected C. qulnoa leaves using the method described by Brunt and Stace-Smith (2). procedures were carried out at 4 C. All purification One hundred g of tissue 31 was h o m o g enized in a War i n g blen d e r in 200 ml of 0.02 M sodium phosphate buffer, pH 6.5, containing 0.01 M MgCl and 0.1% 2-mercaptoethanol. The homogenate was strained through two layers of cheesecloth and adjusted to pH 5.2 with 10% citric acid (w/v) and stirred for 30 min. The homogenate was centrifuged at 10,000 rpm for 10 min in a Beckman No. 40 rotor, the supernatant was decanted and adjusted to pH 6.5 with 1 N NaOH. This solution was centrifuged at 28,000 rpm for 150 min in a Beckman No. 30 rotor and the pellet was resuspended overnight in 2 ml of 0.02 M sodium phosphate buffer, with 0.01 M MgCl and 0.1% 2-mercaptoethanol. low speed centrifugation, pH 6.5, After a second the supernatant was saved and the virus particles were pelleted by high speed centrifugation at 38,000 rpm for 90 min in a Beckman No. 40 rotor. A second round of low and high speed centrifugations was used to further purify the virus. Final purification was done using 0-30% linear-log sucrose density gradients which were centrifuged in a Beckman SW-41 rotor at 38,000 rpm for 90 min. Gradients were fractionated using an ISC0 density gradient fractionator and UV-analyzer [Instrumentation Specialties Co., Lincoln, NE 68505] and the sucrose fractions containing virus were diluted threefold with 0.01 M sodium phosphate buffer, pH 7.0, and centrifuged for 3 hours at 38,000 rpm in a Beckman No. 40 rotor. The pellet was resuspended overnight in 0.01 M sodium phosphate buffer, pH 7.0, and the concentration of the virus preparation was determined using the molar extinction coefficient value 2 80nm = 5.3. 32 Isolate B, suspected to be AV I, was purified from C. quinoa leaves using the methods of W. Howell (personal communication). at 4 C. All purification procedures were carried out Leaf tissue (20 g) was triturated in a Waring blender with 200 ml of 0.1 M sodium citrate buffer, pH 7.0, containing 0.01 M sodium ethylenediamine tetraacetate (EDTA). The homogenate was strained through two layers of cheesecloth and centrifuged at 6,000 rpm for 10 min in a Beckman No. 30 rotor. The supernatant was decanted and centrifuged for 90 min at 28,000 rpm in a Beckman No. 30 rotor and the pellet was resuspended overnight in 2 ml of 0.01 M sodium phosphate buffer, pH 7.0. An equal volume of chloroform was added, shaken vigorously for 10 min and the mixture centrifuged at 6,000 rpm for 10 min in a Beckman No. 30 rotor. The aqueous pha s e was layered onto a 5 ml cushion of 4% polyethene glycol (PEG) mol. wt. 6,000 (w/v) and 30* sucrose (w/v) containing 0.12 M sodium chloride in a Beckman SW-41 rotor tube and centrifuged at 28,000 rpm for 90 min. Each pellet was resuspended overnight in 2 ml 0.01 M sodium phosphate buffer, pH 7.0. Further purification was done using 0-30* linear-log sucrose density gradients as previously described, except that gradients were centrifuged at 15,000 rpm for 60 min. The concentration of the virus preparation was determined 0 1% using the molar extinction value Ei* z. = 2.8. 2 80nm 33 Serologically Specific Electron Microscopy The serologically specific electron microscopy (SSEM) or immunosorbent electron microscopy technique of Derrick (5) was used to test asparagus plants for the presence of AV I. Spear tips were t riturat e d in a s m a l l v o l u m e of 0.01 M sodium phosphate buffer, pH 7.0, and rub-inoculated onto leaves of two C. quinoa plants. Single necrotic local lesions developing after 10 days were excised and triturated in a small volume of 0.01 M phosphate buffer, pH 7.0. Formvar covered copper grids (200 mesh) were first carboncoated then treated for 1 hour at room temperature with a 1:10 or 1:100 d i l u t i o n of anti s e r u m to A V I [provided by G.I. Mink], AV I-J [provided by I. Fujisawa], or AV I-M. The grids were placed on a drop of the homogenate overnight at 4 C then rinsed with 0.01 M sodium phosphate buffer, 7.0. pH Degree of reaction in SSEM was determined by recording the number of vir u s p a r t i c l e s o b s e r v e d in a 5 min scan of the grid. Decoration of virus particles was carried out at room temperature using a 1:10 dilution of antiserum prepared against A V I-W, A V I-J or A V I-M. Grids we r e f l o a t e d specimen-side down for 1 hour then rinsed with 0.01 M sodium phosphate buffer, pH 7.0, for 3 min. Grids were stained with 2% phosphotungstlc acid (w/v), pH 7.0, or 2% ammonium molybdate (w/v), pH 7.0, and observed on a Philips 201 electron microscope. 34 Electron Microscopy Partially purified virus preparations of isolate A were first prepared by fixation in 1% glutaraldehye for 1 minute then placed onto grids prepared as before, and negatively stained with 2% ammonium molybdate (w/v), pH 7.0. Al l grids were examined with a Philips 201 electron microscope for the presence of virus particles. Antisera Production Antisera to the Michigan isolates A and B were prepared using the following procedure. The immunogens were prepared for injection by emulsifying 1.0 ml of 250 to 500 ug of the appropriate virus with 1.0 ml of Freund's complete or incomplete adjuvant [Difco Laboratories, Detroit, MI 48232]. Four-month old female New Zealand White rabbits received one intramuscular injection of immunogen in Freund's complete adjuvant followed by four intramuscular injections of immunogen in Freund's incomplete adjuvant at seven day intervals. Blood was collected from marginal ear veins beginning five days after the last injection and continued at weekly intervals for 1 month. Serum was separated from coagulated red blood cells by incubation at room temperature for 3 hours followed by overnight incubation at 4 C. Clear serum was pipetted off and centrifuged at 1,800 rpm for 15 min to remove cellular components and a few crystals of sodium azide were added prior to storage at 4 C or -20 C. 35 Gamma Globulin Purification Anti-A or anti-B gamma globulin was purified by the procedure described by Clark and Adams (3). Antiserum was diluted 1:10 (v/v) in distilled water and 10 ml of saturated ammonium sulfate was added drop-wise while stirring. After stirring 30 to 60 min, the mixture was centrifuged for 5 min at 6,000 rpm in an IEC model CL centrifuge [International Equipment Co., Needham Hts., MA 02194]. The precipitate was collected and dissolved in 2 ml half-strength PBS [0.01 M sodium phosphate buffer, pH 7.4, containing 0.8% sodium chloride (w/v), and 0.01% sodium azide (w/v)]. This gamma globulin fraction was dlalyzed three times against 500 ml h a l f - s t r e n g t h PBS at 4 C and passed through a 5 cm h i g h bed of DEAE (Whatman DE-23) cel l u l o s e in a 10 ml g l a s s pipette. Ha If-strength PBS was used to pre-equilibrate the column and elute the gamma globulin. Two-ml fractions were collected by monitoring spectrophotometrical ly at the A280 nm wavelenth for protein. The first protein fractions to elute were collected and adjusted to a 1 mg/ml concentration and stored at 4 C. ELISA Conditions Indirect enzyme-1 inked Immunosorbent assay (ELISA) was carried out using the method of Vo H e r and Bidwell (19). Samples were triturated in a 0.05 M carbonate buffer, pH 9.6, with 2% polyvinyl pyrrolidone (w/v) (PVP) mol. wt. 40,000 [Sigma Chemical Co., St. Louis, M0 63178] and 0.45% sodium diethydlthiocarbamate (w/v). Samples were diluted 36 1:10 or 1:50 (v/v) in the grinding buffer, filtered through two layers of cheesecloth and 200 ul of the dilutant added to the wells of flat bottomed polystyrene microtiter plates [Dynatech Laboratories Inc., Alexandria, VA 22314}. Plates were incubated o v e r n i g h t at 4 C and washed three times w i t h PBS containing 0.05% Tween 20 (v/v) (PBS-Tween). Purified gamma globulin to AV I or AV II was diluted 1:50 and 1:100 (v/v), respectively, with PBS-Tween and 200 ul was added to each well and Incubated at room temperature for 2 hours. Allcaline phosphatase conjugated with goat IgG prepared against whole molecule rabbit IgG [Sigma Chemical Co., St. Louis, M0 63178], diluted 1:1000 (v/v) in PBS- Tween, was added to rinsed p l a t e s in 200 ul a l i q u o t s and incubated at 37 C for 3 hours. Plates were then washed throughly three times with PBS-Tween and 200 ul of 1 mg/ml enzyme substrate, p-nltrophenyl phosphate [Sigma Chemical Co., St. Louis, M0 63178], dissolved in substrate buffer (10% diethanolamine (v/v), pH 9.8, in distilled water with 0.02% sodium azide (w/v)). Color change after 30 to 60 min was determined spectrophotometrically at A405 nm with a microELISA Minireader [Dynatech Laboratories, Alexandria V A 22314]. Tests were considered positive for virus if A405 nm v a l u e of a s a m p l e wel 1 was greater than the mea n A405 nm of the healthy control samples plus three standard deviations. 37 Results Field Survey The 1982 survey revealed widespread infection of commercial asparagus plantings in Oceana and Van Buren Counties by a single virus, isolate A, later identified as AV II (Table 2.1). This virus was present in 19 of 23 fields surveyed and most plantings were more than 50% infected. The incidence of infection with isolate A was greater for p l a n t i n g s 11 to 15 and 16 to 20 years o l d than for those 0 to 2 and 3 to 10 years of age. In the survey of asparagus seedling nurseries in Oceana County, four of five nurseries were infected with isolate A but generally at lower levels than commercial plantings (Table 2.2). No other viruses were detected in any asparagus planting during the 1982 growing season. No additional viruses were detected in commercial asparagus plantings during the 1983 survey (data not shown). In 1984, A V I-type local lesions we r e detected on C. qu l n o a rub-inoculated with spears collected from two commercial asparagus plantings in Oceana County and designated isolate B (data not shown). During the 1985 harvest season AV I was detected in five of ten newly surveyed plantings in Oceana County (Table 2.3). There was no correlation between the age of the planting and the incidence of AV I although the oldest field, planted in 1950, had the greatest percentage of plants (70%) infected by this virus. Two asparagus plantings immediately adjacent to Table 2.1. Percent asparagus virus II (AV II) infection as determined with Chencroodium guinea indicator plants for 12 asparagus fields in Oceana County and 11 fields in Van Buren County. z Oceana County Van Buren County Age of Plants (yrs) Fields Assayed (No.) Total Plants Assayed (No.) Incidence for Individual Field (%) Fields Assayed (No.) Total Plants (No.) Incidence for Individual Fields (%) Mean % Infection for all Fields 0-2 3 57 60, 11, 0 3 68 3-10 3 61 15, 3 62 60, 5, 70 25.0 11-15 3 60 50, 25, 71 3 60 75, 60, 80 60.2 16-20 2 41 55, 55 3 65 90, 70, 76 69.2 0, 0 0, 71, 67 34.8 zEach sample was ground in 0.01 M sodium phosphate buffer, pH 6.8, with 0.1% 2-mercaptoethanol added and rub-inoculated to separate indicator plants that had first been dusted with carborundum (320 mesh) Asparagus cultivars assayed were Mary Washington and Viking KB3. Samples were kept at 4 C after collection until processed 24 to 48 hours later. Indicator plants were observed for virus symptoms 5 to 20 days after inoculation. 39 Table 2.2. Percent asparagus virus II infection as determined with Chenopodium guinea indicator plants for five asparagus seedling nurseries in Oceana County, MI.Z Age of Plants (months) Fields Assayed (No.) Total Plants Tested (No.) Incidence for Individual Field (%) 12 2 24 16.6, 16.6 18 2 30 0, 41.6 24 1 21 33.3 zEach sample was ground in 0.01 M sodium phosphate buffer, pH 6.8, with 0.1% 2-mercaptoethanol added and rub-inoculated to separate indicator plants that had first been dusted with carborundum (320 mesh). Asparagus cultivars assayed were Mary Washington and Viking. Samples were kept at 4 C after collection until processed 24 to 48 hours later. Indicator plants were observed for virus symptoms 5 to 20 days after inoculation. 40 Table 2.3. Incidence of asparagus virus I (AV I) in commercial asparagus in Michigan as determined with Chenopodium quinoa indicator plants and serological ly specific electron microscopy. Presence of asparagus virus I y Cultivar Year Planted No. positives2/ No. tested Oceana County 1950 1955 1967 1969 1970 1975 1980 1980 1981 1982 Mary Washington Mary Washington Mary Washington Mary Washington Mary Washington Mary Washington Viking KB3 Viking KB3 Viking KB3 Viking KB3 18/25 0/25 16/25 2/25 0/25 3/25 0/25 0/25 8/25 0/25 (72%) ( 0%) (64%) ( 8%) ( 0%) (12%) ( 0%) ( 0%) (32%) ( 0%) 0/25 0/25 0/25 0/25 0/25 0/25 0/25 (0%) (0%) (0%) (0%) (0%) (0%) (0%) Van Buren County 1959 1960 1965 1969 1970 1975 1975 Mary Mary Mary Mary Mary Mary Mary Washington Washington Washington Washington Washington Washington Washington ^individual asparagus spear samples were indexed an Chenopodium quinoa by rub-inoculatian. zLocal necrotic lesions from C. quinoa leaves were checked for AV I particles using serologically specific electron microscopy. 41 the 35 year o l d fie l d had the next greatest Incidence of Infection by AV I and fields at greater distances were determined to contain even fewer plants infected with this virus. Asparagus virus I was not detected in any asparagus plantings surveyed in Van Buren County during the summer of 1985. Serology Sap from C. quinoa plants systemically infected with isolate A and diluted 1:1 produced a single preciptin line in agar gel double diffusion tests with antiserum prepared against A V II d i l u t e d 1:3 (v/v) but did not react w i t h other antisera tested (Figure 2.1). Sap from C. quinoa plants infected with a Washington isolate of AV II also produced a single precipitin line and no precipitin line was formed in tests with healthy sap. Physical Properties and Host Range The physical characteristics and host range of isolate A were the same as that of the Washington isolate of AV II. Isolate A had a dilution end-point of 10”3 to 10“4 , thermal i n a c t i v a t i o n point of 55 to 60 C and a l o n g e v i t y in v i t r o of 8 to 10 days. The general characteristics agree with the results obtained with the AV II isolate from Washington (17). The host range of is o l a t e A was d e t e r m i n e d to be the same as reported for AV II in Washington (18) (Table 2.4). Isolate B possessed physical characteristics and host range identical with those of the Washington isolate AV I (17). The isolate was determined to have a dilution end- 42 Figure 2.1. Ouchterlony gel double diffusion test. Center well was charged with asparagus virus II (AV II) antiserum (dilution = 1:1 or 1:2), peripheral wells were charged with asparagus virus II (Washington isolate) = A, Michigan isolate I = B, Michigan isolate II = C and D, asparagus virus I = E and sap front virus-free Chenopodium quinoa = F. 43 44 Table 2.4. Reactions of 12 plant species inoculated with a Michigan isolate of asparagus virus II (AV II). Plant species Symptoms Asparagus officinalis L. Latent systemic infection Beta vulgaris L. Local chlorotic ring lesions Chenopodium amaranticolor Coste & Reye Small local necrotic lesions C. murale L. Sunken local necrotic lesions No systemic infection C. quinoa Willd. Local chlorotic ring lesions Systemic mottle occasional slight necrosis Cucumis sativus L. Local chlorotic ring lesions Cucurbita pepo L. Systemic mottle Gotnphrena qlobosa L. Necrotic ringspot lesions Mottling of lower leaves Nicotiana tabaccum L. cv. Havana 423 Local chlorotic ring lesions N. clevlandii A. Gray Latent systemic infection Phaseolus vulgaris L. cv. Bountiful Small red local lesions Vigna unquiculata (L.) Walp. cv. California Blackeye Small red local lesions point of 10-3 to 10“4 , thermal inactivation point of 55 to 60 and a l o n g e v i t y in v i t r o of 1 to 2 days. The host range of B included only members of the Chenopodiaceae, C. quinoa and C. amaranticolor including (Table 2.5). Ultraviolet absorption profiles of purified preparations of isolate A which were sedimented in a 0-30 percent linearlog sucrose density gradient revealed two major peaks (Figure 2.2). Virus particles collected from each peaks were determined to infective when rub-inoculated onto quinoa, producing typical A V II-type symptoms 5 to 10 days after Inoculation. Serologically Specific Electron Microscopy Serologically specific electron microscopy studies indicated that isolate B was serologically related to the Washington Isolate of AV I. Formvar-covered, carbon-coated copper grids previously treated with a 1:10 dilution of antiserum to the Washington or Michigan isolates of AV I trapped 100 times more virus particles than did untreated grids. An average of 121 virus particles were observed in 5 min counts of grids pre-treated with antiserum prepared against the W a s h i n g t o n i s o l a t e of A V I. A n a v e r a g e of 10 particles were observed in 5 min counts of untreated grids. Virus particles of isolate B became decorated when they were treated with a 1:10 dilution of antiserum prepared against the Washington or Michigan isolates of AV I but TM V particles treated in the same manner were not decorated. Virus particles of isolate B treated with a 1:10 dilution of pre- 46 Table 2.5. Reactions of 13 plant species inoculated with a Michigan isolate of asparagus virus I (AV I). Plant species Symptoms Asparagus officinalis L. No reaction Beta vulgaris L. No reaction Chenopodium amaranticolor Coste & Reyes Local necrotic lesions C. murale L. Local necrotic lesions C. quinoa Willd. Local necrotic lesions Cucumis sativus L. No reaction Cucurbita peoo L. No reaction Goraphrena globosa L. No reaction Nicotiana tabaccum L. cv. Havana 423 No reaction N. clevlandii A. Gray No reaction Phaseolus vulgaris L. cv. Bountiful No reaction Tetragonia expansa Murr. No reaction Vigna unguiculata (L.) Walp. cv. California Blackeye NO reaction zIndicator plants not showing virus symptoms after 14 days were back inoculated to C. quinoa to test for latent infection. 47 Figure 2.2. Ultraviolet absorption profile of a Michigan isolate of asparagus virus II (AV II) sedimented in a 0-30 percent linear-log sucrose density gradient. The direction of sedimentation is from left to right. A Beckman SW-41 rotor was used at 38,000 rpm for 90 minutes at 4 C. 48 peakl .9 .8 254 nm .7 .6 peak 2 .5 .4 .3 .2 .1 .0 0 8 16 24 32 40 Depth below Meniscus (mm) 48 49 immune antiserum were not decorated (Figure 2.3). Electron Microscopy Isometric or quasi-isometric particles ranging in diameter from 23 to 30 nm (Figure 2.4) were observed when negatively stained purified virus preparations of isolate A were viewed on the electron microscope. shaped virus particles, Long flexous rod­ 700 to 880 nm in length (Figure 2.3) were readily visible in leaf dips prepared from single necrotic local lesions of C. quinoa leaves inoculated with isolate B. 50 Figure 2.3. Results of immunosorbent electron microscopy with a A & B. Michigan isolate of asparagus virus I (AV I) or C. tobacco mosaic virus (TMV). Formvar covered copper grids were carbon coated then treated overnight at 4 C with a 1/10 dilution of antiserum prepared against the Washington isolate or Japanese isolate of AV 1. A & C were decorated with the same antiserum for 1 hr at room temperature then stained with either 2% phosphotungstic acid or 2% ammonium molybdate. Scale bar = 100 nm. 51 52 Figure 2.4. Electron micrograph of Michigan isolate of asparagus virus II (AV II). Quasi-iscmetric virus particles were fixed with 195 g 1utara1hehyde for 10 minutes then negatively stained with 295 ammonium molybdate, pH 7.0. Scale bar = 50 nm. 53 54 Discussion Michigan isolates A and B appeared to be serologically related to AV II and AV I, respectively, described previously by Mink and Uyeda (14) in Washington. Isolate A reacted serologically only with antisera against the Washington isolate of AV II. Serological precipitin lines coalesced in agar and no spur formation was noted. Serologically specific electron microscopy indicated that isolate B is serologically related to the Washington isolate of AV I. Asparagus virus II was determined to be widespread in almost all Michigan commercial asparagus plantings surveyed (Table 2.1). Four of five seedling nurseries surveyed were infected with AV II but generally at a lower level than older production fields (Table 2.2). Though considerable variation in the incidence of AV II within age groups exists, the incidence of AV II was greater for asparagus plantings 11 to 20 years o l d than for those 0 to 10 years of age and s e e d l i n g nurseries generally had the lowest incidence of infection. Mean percent infection data indicate that younger fields had fewer plants infected with AV II than older fields. A sparagus v irus I was d e t e r m i n e d to be present in o n l y a few Michigan asparagus plantings in Oceana County and its incidence varied greatly (Table 2.3). No direct correlation exists between the age of a p l a n t i n g and its I ncidence of infection w i t h AV I. One 35 year o l d p l a n t i n g in O c e a n a C o u n t y was determ i n e d to be 70* Infected w i t h A V I and two immediately adjacent asparagus plantings were determined to be 33% and 63% infected w i t h A V I. F i e l d s at e v e n greater distances from the 35 year old planting had even lower incidences of AV I and the v i r u s was not d e t e c t e d in plantings located more than one mile away from the 35 year old planting. A relationship exists between the proximity of a p l a n t i n g to the 35 year o l d f i e l d in Oceana County and its percent infection with AV I. Asparagus virus I is a member of the P V Y group and is thought to be transm i t t e d by aphids in the nonpersistent manner (6, 7, 8, 13). The data suggest that the 35 year o l d p l a n t i n g is a sou r c e from w h i c h aphids have spread the virus to nearby asparagus plantings in Oceana County. 56 Literature Cited 1. Brunt, A. A., and Paludan, N. 1970. The serological relationship between "asparagus stunt" and tobacco streak virus. Phytopathol. Z. 69:277-282. 2. Brunt, A. A., and Stace-Smith, R. 1976. The occurrence of the black raspberry latent strain of tobacco streak virus in wild and cultivated Rubus species in British Columbia. Acta. Hortic. 66:71-76. 3. Clark, M. F., and Adams, A. N. 1977. C h a r a cteristics of the micro-plate method of enzyme-linked immunosorbent assay for the detection of plant viruses. J. Gen. Virol. 34:475-484. 4. Davis, R. F., and Garrison, S. 1985. First report of a virus present in New Jersey. Plant Dis. 66:628. 5. Derrick, K. S. 1973. Quantitative assay for plant viruses using serologically-specific electron microscopy. Virology 56:652-653. 6. Falloon, P. G. 1982. The need for asparagus breeding in New Zealand. New Zealand J. Exp. Agr. 10:101-109. 7. Falloon, P. G., Fallo o n , L. M., and Grogan, R. G. 1985. A survey of California asparagus for asparagus virus I (AVI), asparagus virus II (AVII) and tobacco streak virus (TSV). Plant Dis. (in press). 8. Fujisawa, I., Goto, T., Tsuchizaki, T., and Iizuka, N. 1983. Host range and some properties of asparagus virus I isolated from Asparagus officinalis in Japan. Ann. Phytopath. Soc. Japan 49:299-307. 9. Fujisawa, I., Goto, T., Tsuchizaki, T., and Iizuka, N. 1983. Some properties of asparagus virus II isolated from Asparagus officinalis in Japan. Ann. Phytopath. Soc. Japan 49:683-688. 10. Hartung, A. C., Evans, T. A., and Stephens, C. T. 1985. Occurrence of asparagus virus II in commercial asparagus fields in Michigan. Plant Dis. 69:501-504. 11. Hein, A. 1960. Uber das vorkommen einer virose an sparge1. Z. Planzenkr. Pflanzenpathol. Pflanzenschutz 67:217-219. 12. Hein, A. 1963. Viroses an spargel. Mitt. Biol. Bundesanst. Land-Forstwirtsch, Berlin-Dahlem 108:70-74 (Rev. Appl. M y c o l . 1964. 43:455). 57 13. Hein, A. 1969. Uber viruskrankheiten das spargels (Asparagus officinalis L.). Sparge 1 virus I. Z. Pflanzenkr. Pflanzenpathol. Pflanzenschutz 76:395-406. (Rev. Plant Pathol. 1970. 49:219). 14. Mink, G. I., and Uyeda, I. 1977. Three mechanically transmissible viruses isolated from asparagus in Washington. Plant Dis. Rep. 61:398-401. 15. Paludan, N. 1964. Virussygdomme hos Asparagus officinalis Mannedsovers. Platesygd. 407:11-16. 16. Posnette, A. F. 1969. Nematode transmitted viruses in asparagus. J. Hortic. Scl. 44:403-404. 17. Uyeda, I. 1978. Identification, characterization, and incidence of viruses isolated from asparagus. Ph.D. thesis, Washington State University. 115 pp. 18. Uyeda, I. and Mink, G. I. 1981. P r o p e r t i e s of asparagus virus II, a new member of th Ilarvirus group. Phytopathology 71:1264-1269. 19. Voller, A., and Bidwell, D. E. 1977. Enzyme immunoassays and their potential in diagnostic virology, pages 449-457 in, Comparative Diagnosis of Viral Diseases Vol. II. E. K u r s t a k and C. Kurstak, eds. 20. Weissenfels, M., and Schmelzer, K. 1976. Untersuchungen ueber das schadausmass durch viren am sparge1 (Asparagus officinalis L.). Arch. Phytopathol. Pflanzenschutz, Berlin. 12:67-73. 21. Yang, H. 1979. Early effects of viruses on the growth and productivity of asparagus plants. HortScience 14:734-735. CHAPTER III TRANSMISSION AND SPREAD OF ASPARAGUS VIRUSES WITHIN MICHIGAN ASPARAGUS Introduction Asparagus virus II (AV II) has been detected widely within most commercial asparagus plantings in Michigan and a spar agus virus I (AV I) has a l s o been d e t e r m i n e d to be present, but not widespread. A spa r a g u s vir u s I has b e e n c l a s s i f i e d as a memb e r of the potyvirus (PVY) group which are transmitted in a nonpersistent manner by a wide variety of aphids (5). Members of the potyvirus group typically have flexous filamentous particles mostly 730 to 790 nm in length and seed transmission is relatively rare (12). Some members of this group are mechanically transmissible to a moderately wide range of herbaceous hosts while others have relatively restricted host ranges (12). Asparagus virus I is widespread in asparagus plantings in Washington (19), California (8), New Jersey (4), and has been detected in a few Michigan plantings. Attempts to seed- transmit AV I within asparagus or rub-transmit the virus to healthy asparagus using sap or purified preparations of the 58 59 virus have been unsuccessful (9, 21). Asparagus virus II is a member of the ilarvirus group and its method of transmission within asparagus plantings is also unclear. Members of the ilarvirus group have isometric or quasi-isometric particles generally ranging in size from 22 to 35 nm and most are composed of two to three kinds of nucleoprotein components of different sizes (22). Many members of this group are known to be seed-transmitted and most are readily mechanically transmissible to a relatively wide range of herbaceous hosts (22). Asparagus virus II is widespread in asparagus plantings in Washington (19), and California (8), and has been detected at low l e v e l s in N e w Jersey. In Michigan, 50 to 70 percent of asparagus plants are Infected with AV II in most plantings 5 years or older (15). Asparagus virus II has been mechanically transmitted by some (10, 24) but not by others (21). The transmission of AV II by seed within asparagus is well documented (10, 16, 21, 22). A member of the Ilarvirus group, prunus necrotic ringspot virus (PNRSV), is pollen-borne in cherry and infects trees when they are pollinated by virus-carrying pollen (11). Virus particles have been localized on the outer wal l s of some pollen grains. Transmission electron microscopic examination of anthers from plants infected with bromegrass mosaic virus (BMV), southern bean mosaic virus (SBMV) or tobacco mosaic virus (TMV) showed that the exine of mature pollen was infested with virions and that homogenates of SBMV-infested pollen or dry TMV-infested pollen were 60 infectious when assayed on appropriate hosts (13). Hamilton et al (14) demonstrated the presence of PNRSV-antigens on the exine of bee and hand-collected sweet cherry p o l l e n using enzyme-linked immunosorbent assay (ELISA). When virus- containing pollen was washed the antigen was easily released and induced chlorotic local lesions when rub-inoculated onto Chenopodlum amarantlcolor and Cucumis sativa cv. Straight Eight. Scanning electron microscopic observations of pollen treated with latex-conjugated antibody specific for PNRSV revealed more latex beads bound to the surface of virusinfested than virus-free pollen. The objective of this work was to investigate the methods of t r a n s m i s s i o n for A V I and A V II and to b e g i n to assess the relative importance of the methods of transmission in the spread of these viruses within Michigan asparagus plantings. 61 MATERIALS AND METHODS Aphid Transmission of AV I The a b i l i t y of two aphid species to transmit A V I was evaluated In the greenhouse and growth chamber. Colonies of non-viruliferous green peach aphids (Myzus persicae Sulz.) and European asparagus aphids (Brachycolus asparaql Mord.). European asparagus aphids and green peach aphids were given an aquisition access period of 1 and 2 weeks, respectively, on asparagus plants infected with AV I, then transferred to 2-month old virus-free and AV II-infected asparagus seedlings. After 2 weeks all shoots were cut at ground level and plants transferred to aphid-free cages. Test plants were evaluated for the presence of AV I by rubinoculation to C. quinoa and SSEM after the second new spear had emerged. Mechanical Transmission of AV I Spears were collected from greenhouse-grown asparagus p l a n t s known to be infected o n l y with A V I and t riturated in a 0.02 M sodium phosphate buffer, mercaptoethanol. pH 7.0, containing O.lfc 2- This homogenate was rub-inoculated onto the stems of virus-free asparagus seedlings and leaves of C. quinoa that were first dusted with carborundum (320 mesh). Inoculated asparagus plants were evaluated for symptom development over a 2 month period then tested for A V I using rub-inoculation to quinoa. Necrotic lesions developing on leaves of C. quinoa were assessed for AV I particles using SSEM. 62 Mechanical Transmission of AV II to Asparagus A Michigan isolate of AV II was purified from systemically infected C. quinoa leaves using the method of Brunt and Stace-Smith (1) and further purified using three cycles of linear-log sucrose density gradients as previously described (15). A 1 mg/ml solution of purified AV II in 0.02 M sodium phosphate buffer, pH 7.0, was used to inoculate asparagus seedlings that had previously been screened for AV II using enzyme-linked immunosorbent assay (ELISA) following the procedures of Voller and Bidwell (23), and rub-inoculated to C. quinoa. Virus-free asparagus seedlings were maintained in the dark for 24 hours, then dusted with carborundum (320 mesh) and stems were rub-inoculated with purified virus or buffer using sterile cotton swabs. Plants were evaluated for symptom development over an 8 week period, after which young spears were harvested and tested for AV II using ELISA and rub-inoculation to C. quinoa. Mechanical transmission from asparagus to asparagus was assessed in a similar manner. Asparagus seedlings were first screened for AV II using ELISA and rub-inoculation to C. quinoa. Young spears from AV II-infected and virus-free asparagus plants were triturated with a mortar and pestle at 4 C and diluted 1:10 (w/v) in 0.02 M sodium phosphate buffer, pH 7.0, containing O.lfc 2-mercaptoethanol and triturated with a mortar and pestle at 4 C. These sap preparations were used to rub-inoculate virus-free asparagus seedlings either directly on the stem, as previously described, wounded tips of recently excised spears. or on the 63 Pollen Transmission of AV II in Asparagus Col lection of asparagus pollen During the winter of 1985, flowers were collected from greenhouse-grown, male asparagus seedlings previously indexed for the presence of AV II by ELISA and rub-inoculation to C. guinoa. Anthers were first examined under a dissection microscope and mature anthers, prior to dehiscence, were removed manually using fine forceps. The anthers were dried at room temperature for 48 hours and stored at 4 C. Pollen for individual experiments was prepared using the method of Cole et al (3). Dried anthers were swirled in a dry glass tube using a vortex tube agitator and pollen grains that adhered to the tube walls were collected. No evidence of damage was noted when suspensions of pollen were examined by light microscopy. Pollen was usually processed within 1 week of its collection. Tests for presence of Infective v iruses Pollen samples were tested for the presence of infective AV II using the methods of Hamilton et al (14). Samples were suspended in 5 ml of 0.01 M sodium phosphate buffer, pH 7.0, or in the same buffer amended with lfc polyvinyl pyrrolidone (v/v) (PVP) mol. wt. 40,000 [Sigma Chemical Co., St. Louis, M0 63178]. The tubes were shaken with a vortex tube agitator at m a x i m u m speed for 1 min. No e v i d e n c e of p o l l e n dam a g e was observed when suspensions were observed by light microscopy. P o l l e n grains were sedimented at 6,000 rpm for 5 to 10 min and a 2-ml sample was withdrawn with a sterile pipette. This 64 sample was rub-inoculated on single leaves of two C^ quinoa and the p l a n t s o b s e r v e d for 5 to 14 days for symptom development. Identification of AV II The identification of AV II in pollen samples or wash solutions was carried out by ELISA using the methods of Vol ler and Bidwe 11 (2 3) as m o d i f i e d by Hami 1 ton et al (14). Pollen was divided into equal test samples of 0.05 ml dry-packed volume and suspended in 0.01 M sodium phosphate buffer as previously described. Suspensions of intact pollen grains in grinding buffer added to duplicate wells of flat bottomed polystyrene microtiter plates [Dynatech Laboratories, Alexandria, VA 22314] and incubated overnight at 4 C. Plates were washed three times with phosphate buffered saline with 0.05% Tween 20 (v/v) (PBS-Tween). Gamma globulin specific for the Michigan isolate of AV II was purified using the procedure of Clark and Adams (2), d i l u t e d 1:50 (v/v) with PB S - T w e e n and 200 ul added to each well and incubated for 2 hours at room temperature. Plates were washed as before and 200 ill of alkaline phosphatase conjugated with goat IgG prepared against whole molecule rabbit IgG [Sigma Chemical Co., St. Louis, MO 63178], diluted 1:1000 (v/v) in PBS-Tween, was added. After 3 hours incubation at 37 C p l a t e s were again w a s h e d and 200 ill of 1 mg/ml enzyme substrate, p-nitrophenyl phosphate [Sigma Chemical Co., St. Louis, M0 63178], dissolved in substrate buffer (10% diethanolamine, pH 9.8, in distilled water with 65 0.02* sodium azide (w/v)) were added. Color change after 30 to 60 min was determined spectrophotometrical ly at A405 with a microELISA Minireader [Dynatech Laboratories, Alexandria, VA 22314]. A threshold value used for positive a reaction for each p l a t e was the mean A405 nm v a l u e of the h e a l t h y control plus three standard deviations. Pollen suspensions were incubated for 60 min at room temperature, centrifuged as previously described and a 200 ul sample of pollen wash solution was withdrawn and used as antigen in similar ELISA tests. Pollen samples were washed two more times in this same manner and resuspended in grinding buffer. No evidence of pollen damage or tapetal fragments was apparent when suspensions were examined by light microscopy. Washed pollen samples were divided in half and intact pollen used as antigen in ELISA. The remaining half of the sample was ground manually in a glass tissue homogenizer until all pollen grains were ruptured and the homogenate used as antigen as previously described. Detection AV II antigen on the surface of asparagus p o 1 1 en Protein A-coated latex-linked antisera (PALLAS) was used to d e termine if AV II anti g e n was present on the surf a c e of asparagus pollen using the methods of Querfurth and Paul (20) and Hamilton et al (14). A standard suspension of bacto- latex beads [0.81 urn, Difco Laboratories, Detroit, MI 48232] was first diluted with 14 volumes of 0.9* NaCL (w/v) solution then mixed 1:1 with a diluted protein A solution [Sigma Chemical Co., St. Louis, MO 63178], composed of 0.1 mg 66 protein A dissolved in 2.0 ml 0.1 M glycine buffer, pH 8.2. This s o l u t i o n was incubated for 2 to 4 hours at room temperature with occasional stirring then allowed to stand overnight at 4 C. The protein A-latex bead complexes were washed by centrifuging at 6000 rpm for 30 min and resuspended in 0.1 M glycine buffer, pH 8.2, with 0.02* PVP (v/v) added. The w a s h i n g step was repeated two more times and the final pellet resuspended in 2 ml 0.1 M glycine buffer, pH 8.2, with 0.05* sodium azide (w/v) [Sigma Chemical Co., St. Louis, M0 63178]. Purified gamma globulin specific for the Michigan isolate of AV II was conjugated with the protein A-latex bead complex. buffer, The globulin suspension (100 ug/ml in 0.1 M glycine pH 8.2) was mixed 1:1 with the protein A-latex bead complex, a l l o w e d to Incubate for 2 to 4 hours at ro o m temperature and washed as described previously. Pollen samples were dissolved in 0.1 M glycine buffer, pH 8.2, mixed 1:1 with protein A-latex bead complex and allowed to incubate for 1 hour at room temperature with occasional shaking. The pollen-protein A - 1atex-antibody c o m p l e x was c entrifuged at 6000 rpm for 5 m in and the supernatant was drawn off with a pipette. This wash step was repeated five times and the final pellet of pollen protein Alatex-antlbody complex was resuspended in a small volume of glycine buffer. The solution was pipetted onto the surface of aluminium stubs previously coated with adhesive and allowed to settle for 10 min. Excess fluid was drained by t o uching the edge of the drop w i t h a piece of f i l t e r paper 67 and the pollen was allowed to air dry overnight at room temperature. The pollen samples were gold-coated and examined with an ISI-Mini scanning electron microscope (Figure 3.1). To determine if protein A-latex bead-antibody complexes were bound specifically to AV II antigen, experiment was carried out. a "blocking" Pollen samples were pre-treated with a purified anti-AV II gamma globulin suspension (500 Mg/ml in 0.1 M glycine buffer, pH 8.2) or buffer alone for 1 hour at room temperature with gentle agitation. Samples were then washed three times as previously described then tested for the presence of surface antigen using PALLAS. F i e ld Indexing to determine the distribution and extent of A V I and AV 13^ infection in an asparagus p lanting A b l o c k of 100 p l a n t s in a 10 x 10 g ri d in a 4-year o l d asparagus planting in Oceana County was indexed in June of 1985 for the presence of AV I and AV II. A minimum of two spears were harvested from each plant and maintained at 4 C until indexing. Spear tips were either tested for AV II using the ELISA technique as previously described or triturated in 0.02 M sodium phosphate buffer, rub-inoculated onto C. guinoa. pH 7.0, and Inoculated plants were maintained in the greenhouse for 2 weeks and observed for symptom development. Single necrotic local lesions were excised triturated in a small volume of 0.01 M sodium phosphate buffer, using SSEM. pH 7.0, and tested for the presence of AV I Data from field indexing were evaluated for the non-randomness of AV I or AV II-infected plants using 68 Pollen Sample (in 0.1 M glycine buffer, pH 8.2) I Pol len Sample + IgG specific for AV II Pollen Sample Latex beads conjugated via protein A to IgG — specific for AV II ->-mix 1:1 I Incubate 1 hr at room temperature J Centrifugation (6K, 5 min) i Check for pollen integrity (5 times, Wash and Resuspend 1.0 ml 0.1 M glycine buffer) pH 8.2 Count Number Latex Beads Per Pollen Face on SEM Figure 3.1. Procedure for the localization of asparagus virus II (AV II) antigen on the surface of hand-collected pollen from greenhouse grown Asparagus officinalis using protein A-linked latex antiserum (PALLAS). 69 ordinary run analysis (Madden et al, 1982). Rows of plants we r e combined to form an ar b i t r a r y row with a l e n g t h of 100 plants. A row of plants was considered to have a non-random sequence of Infected and healthy plants if -ZM was greater than 1.64 (P = 0.05). 70 Results Aphid Transmission of AV I in Asparagus Asparagus virus I was transmitted to asparagus by green peach aphids (Myzus persicae) but not by European asparagus aphids (Brachycolus asparagi). Twenty-one of fourty asparagus plants became infected with AV I when fed on by viruliferous green peach aphids whereas none of the 30 asparagus plants fed on by European asparagus aphids became infected with the virus. Mechanical Transmission of AV I to Asparagus Asparagus virus 1 was not successfully mechanically transmitted to asparagus plants using sap prepared from spears known to be infected only with AV I (data not shown). Mechanical Transmission of AV II to Asparagus Manual inoculation of healthy asparagus seedlings with the sap prepared from spears of AV II-infected asparagus plants was an effective means of transmitting the virus (Table 3.1) with 75 to 80% of the test plants becoming infected. Rub-inoculation of sap to stems or to wounded stem tips was equally effective and infected plants sometimes exhibited a mild chlorosis. However, only 1 of 30 asparagus seedlings inoculated with concentrated (1 mg/ml) purified preparations of AV II became infected. 71 Table 3.1. Mechanical transmission of asparagus virus II (AV II) in Asparagus officinalis L. using sap or concentrated, purified virus preparations. Inoculum # of plants infectedY # of plants tested Method of Inoculation Purified2 Virus from C. quinoa Rub to stem 1/30 Rub to stem 16/20 ( 3.3%) Sap AV II-infected asparagus 15/20 (75.0%) Rub to stem 0/20 ( 0.0%) Rub to wounded stem tip 0/20 ( 0.0%) Rub to wounded stem tip Virus-free asparagus (80.0%) VAsparagus plants were indexed for AV II before and 2 months after virus inoculation by enzyme-linked immunosorbent assay and rubinoculation to C. quinoa. zCancentrated purified AV II (1 mg/ml) in 0.01 M sodium phosphate buffer, pH 7.0. 72 Pollen Transmission of AV II in Asparagus Association of AV II with asparagus pol len Pollen samples collected from asparagus plants known to be infected with AV II were determined to contain infective virus. In a preliminary study, pollen and pollen wash solutions from both samples of AV II-infected asparagus induced chlorotic local lesions in Chenopodium guinoa followed by systemic mottling (Table 3.2). Single lesions from these plants were transferred sequentially, on C. guinoa AV II. three times and symptoms induced were identical to those of Asparagus virus II antigen was determined by ELISA to be present in pollen and pollen wash solutions of pollen samples from AV II-infected plants (Table 3.2). No viruses were detected on pollen collected from virus-free asparagus seedlings. Asparagus virus II was determined to be an external contaminant of some asparagus pollen. In a second study, AV II antigen was again determined by ELISA to be present on pollen, in pollen wash solutions and on washed pollen collected from AV II-infected asparagus (Table 3.3). Washed and ruptured pollen from these samples did not contained detectable levels of AV II antigen. Intact pollen and p ollen wash solutions from these samples induced typical AV II symptoms when rub-inoculated onto guinoa. Asparagus virus II antigen was not detected on or within pollen collected from virus-free asparagus and no symptoms were produced on C. guinoa leaves rub-inoculated with these samples. 73 Table 3.2. Detection of antigens and infectious virions of asparagus virus II (AV II) an asparagus pollen and in pollen washes by enzyme-linked immunosorbent assay (ELISA) and rubinoculation. A405 nm values Sample Infectivity y Virus Contentw Pollenx Pollen Wash Pollen / Wash 1 AV II 0.23z 0.15 + / + 2 AV II 0.20 0.16 + / + 3 none 0.00 0.01 - / - 4 none 0.01 0.00 - / - "^Plants were previously indexed for the presence or absence of AV II using rub-inoculation to Chenopodium quinoa leaves and ELISA. ^Pollen was washed in 0.01 M phosphate buffer, pH 7.0, with 1% polyvinylpyrollidone for 60 min then centrifuged at 6,000 rpm for 6-10 min. Yvirus infectivity was determined by rub-inoculation of each sample onto the leaves of two Chenopodium ouinoa plants. zThe critical value for virus detection was the mean of the healthy control plus three standard deviations (A405 nm = 0.06). 74 Table 3.3. Detection of asparagus virus II (AV II) antigens on asparagus pollen, in pollen Mashes and within pollen by enzyme-linked immunosorbent assay (ELISA). A405 nm values Sample Virus Cantentw Pollenx Wash Washed Pollen Ruptured Pol leny 1 AV II 0.082 0.08 0.12 0.08 2 AV II 0.20 1.60 0.09 0.05 3 AV II 0.14 1.88 0.09 0.07 4 AV II 0.09 0.09 0.09 0.08 5 none 0.01 0.00 0.03 0.04 6 none 0.00 0.02 0.03 0.03 7 none 0.01 0.01 0.01 0.03 8 none 0.00 0.02 0.04 0.02 ’"^Plants were previously indexed for the presence or absence of AV II using rub-inoculation to Chenopodium quinoa and by ELISA. xPol len was washed in a 0.01 M phosphate buffer, pH 7.0, with lfc polyvinylpyrol 1idane for 60 minutes then centrifuged at 6,000 rpm for 5-10 minutes. Ypollen was ruptured using a ground glass tissue homogenizer. zThe critical value for virus detection were the mean of the healthy conrtrol plus three standard deviations (A405 nm = 0.06). 75 Local ization of AV II antigen In the exine of asparagus pol len Asparagus virus II antigen was localized in the exine of hand-collected asparagus pollen by PALLAS. Scanning electron microscopy revealed latex particles bound to the exine of p o llen collected from AV II-infected asparagus treated with anti-AV II-protein A-latex conjugate, but not to the same poll e n "blocked" by pre-treatment with anti-AV II IgG or p o llen collected from virus-free asparagus (Figure 3.2). An average of 11.6 latex particles per pollen face were bound to poll e n collected from AV II-infected asparagus and only 4.2 and 1.6 particles per face were observed on pollen collected from virus-free asparagus or "blocked" pollen, respectively (Table 3.4). and d i s t r i b u t i o n of A V I and AV II w i t h i n one a s p a r a gus planting The experimental block of Asparagus officinalis cultivar Viking KB3 spanned 10 rows of plants (Figure 3.3) and was located 100 m from the southeast corner of the field. Of the 100 plants indexed, 30* were determined to be virus-free, 28* were infected with A V I, 36* were infected wit h A V II and 5* we r e d o u b l y infected with both AV I and AV II. The p a t t e r n of distribution of AV I was determined by ordinary run analysis to be non-random (-Z^ = 1.79) whereas the distribution of AV II within that same field appeared to be rand o m (-Zw = 1.23). 76 Figure 3.2. Scanning electron microscopy of hand-col lected asparagus pollen after exposure to latex spheres (0.81 nm diameter) conjugated with asparagus virus II (AV II) antibodies. A) Pollen fran AV II-infected asparagus. B) Pollen from virus-free asparagus. C) Pollen from AV II-infected asparagus "blocked" with antiserum specific for AV II. 77 78 Table 3.4. Localization of asparagus virus II (AV II) antigen in the exine wall of hand-collected asparagus pollen using protein A-linked latex antiserum (PALLAS). Virus Content of Pollenw Number of latex beads/pollen face* + 3 .3^ AV II 11.6 none 4.2 + 1.4 AV II ("blocked"2) 1.6 + 1.4 'Virus content of pollen was determined by enzyme-linked immunosorbent assay (ELISA) and rub-inoculation to C. quinoa. xSamples were first gold coated then observed an an ISI mini scanning electron microscope. yValues indicate the mean of a minimum of 50 pollen faces plus or minus the standard errror. zPollen determined to contain AV II antigen in exine wall by ELISA were pre-treated with purified AV II specific IgG then treated with proteinA IgG-conjugated latex beads. 79 I oo oo oo o+ xo o+ o+ xo x+ oo oo oo o+ o+ oo xo o+ xo xo o+ oo o+ oo o+ oo o+ o+ o+ oo xo oo xo oo x+ xo o+ o+ oo o+ xo xo o+ oo o+ o+ oo o+ o+ o+ o+ xo o+ oo o+ o+ o+ x+ x+ xo xo oo o+ o+ oo o+ xo o+ oo xo xo xo oo oo o+ o+ xo oo xo xo xo oo o+ oo oo o+ xo o+ xo oo oo oo xo o+ xo xo xo xo oo x+ xo To 9 8 7 6 5 4 3 2 ~ ROW NUMBERS Figure 3.3. Distribution of asparagus virus I (AV I) and asparagus virus II (AV II) infected plants in a 4 year old asparagus planting of variety Viking KB3. oo = virus-free, xo = AV I-infected, of = AV II-infected and x+ = d o u b l y infected with AV I and AV II. 80 Discussion The results of these experiments demonstrated that AV II was readily transmissible from asparagus plant to asparagus plant by mechanical means and that the exine of p o l l e n grains from some virus-infected asparagus was contaminated with infectious virions of AV II. Asparagus virus I was determined in these studies to be transmissible in asparagus by the green peach aphids (M vzus persicae). Asparagus virus II has been demonstrated to spread within Michigan (15) and Washington (21, 22) asparagus plantings. It has been suggested that pollen surfaces contaminated with infectious plant viruses may serve as vehicles by which certain mechanically transmissible viruses may spread within some plant species (13). by necessity, This route of virus spread would, be more common and efficient in open-pollinated than in self-pollinated plants and could play a role in the seed transmission of some plant viruses. Asparagus is an open-pollinated plant and may become infected with AV II by mechanically means via wind-blown or bee-carried poll e n Infested with the virus. Honeybees have been observed routinely visiting asparagus flowers, especially in seed production blocks. Bees may mechanically Inoculate plant surfaces or floral parts with AV II during their visitations. This could result in the transmission of the virus to the female plant and/or to developing seed. In California knoves are usually used to sever asparagus spears at h a r v e s t and AV II may a l s o be spread by the knives. - 81 In most asparagus growing regions of the world, ferns are cut in late fall or early spring. This cultural practice may al s o be a means by w h i c h A V II is spread w i t h i n and between asparagus plantings. Contaminated blades could mechanically transmit AV II if asparagus ferns are not allowed to completely senesce prior to cutting. Falloon et al (8) suggested that the spread of AV II within one California seed production block has been a result of cutting green fern stalks with a machete or a tractor-mounted slasher, both methods providing a means of mechanical transmission of AV II. The random distribution of AV II within our Michigan asparagus planting, as determined by ordinary run analysis, is consistent with transmission by seed, mechanical means or the random m o v e m e n t of pol len by wind or bees. In asparagus in N e w Zealand, AV I has be e n reported to increase to 90 to 100 percent by the third year after planting (7) which was attributed to the possible transmission of the virus by aphids. The rate of spread of AV I in New Zealand asparagus plantings and non-random distribution of AV I within Michigan asparagus plantings is consistent with plant-to-plant transmission by aphids. Asparagus virus I was demonstrated in greenhouse studies by this author and others (17, 19) to be readily transmissible within asparagus by the green peach aphid and this is one possible method of spread for AV I within Michigan asparagus plantings. While the European asparagus aphid is widespread in most commercial growing regions of the U.S. it was not demonstrated to be a v e c t o r for the t r a n s m i s s i o n of A V I in 82 this study. It has been observed that asparagus plants Infected with A V I or AV II a l o n e exhibit a m i l d re d u c t i o n in g r o w t h and vigor as compared with virus-free plants (24). Asparagus infected with both AV I and A V II show e d the greatest reduction in growth and often died after 2 years in the field. It has been demonstrated (Chapter 4, this manuscript) that greenhouse-grown asparagus seedlings infected with AV II more readily succumb to Fusarium crown and root rot incited by Fusarium oxysporum f.sp. asparagi and F. monlliforme than virus-free seedlings. Asparagus seedlings doubly infected with both A V I and A V II bec a m e more d i s e a s e d when i n o c u l a t e d with F. oxysporum f.sp. asparagi than those seedlings infected with either AV I or AV II alone. For these reasons, it is imper a t i v e that one or both of these v i r u s e s be eradicated from asparagus plantings so that profitable yields can be maintained. Asparagus virus I is widespread in most asparagus-growing regions of the world and will probably be difficult to eradicate because of its potential transmission by aphids. A l t h o u g h A V II is a l s o widespread, it w o u l d be relatively easy to eliminate from breeding and seed production blocks through meristem tissue culturing (26). Furthermore, its mechanical spread within production plantings could be minimized by discontinuing the mechanical harvest of spears and by delaying the cutting of ferns until they have fully senesced. 83 Literature Cited 1. Brunt, A. A., and Stace-Smith, R. 1976. The occurrence of the black raspberry latent strain of tobacco streak virus in wild and cultivated Rubus species in British Columbia. Acta. Hortic. 66:71- 76. 2. Clark, M. F., and Adams, A. N. 1977. C h a r a c t e r i s t i c s of the micro-plate method of enzyme-linked immunosorbent assay for the detection of plant viruses. J. Gen. Virol. 34:475-484. 3. Cole, A., Mink, G. I., and Regev, S. 1982. L o c a t i o n of prunus necrotic ringspot virus on pollen grains from infected almond and cherry trees. Phytopathology 72:1542-1545. 4. Davis, R. F., and Garrison, S. virus present in New Jersey. 5. deBokx, J. A., and Huttinga, H. 1981. Potato v i r u s Y. No. 242 in: Descriptions of Plant Viruses. Commonwealth Mycological Institute, Kew, Surrey, England. 4 pp. 6. Derrick, K. S. 1973. Quantitative assay for plant viruses using serological ly-specific electron microscopy. Virology 56:652-653. 7. Falloon, P. G. 1982. The need for asparagus breeding in New Zealand. New Zealand J. Exp. Agr. 10:101-109. 8. Falloon, P. G., Falloon, L. M., and Grogan, R. G. 1985. A survey of California asparagus for asparagus virus I (AVI), asparagus virus II (AVII) and tobacco streak virus (TSV). Plant Dis. (in press). 9. Fujisawa, I., Goto, T., Tsuchizaki, T., and Iizuka, N. 1983. Host range and some properties of asparagus virus I isolated from Asparagus officinalis in Japan. Ann. Phytopath. Soc. Japan 49:299- 307. 1985. First report of a Plant Dis. 66:628. 10. Fujisawa, I., Goto, T., Tsuchizaki, T., and Iizuka, N. 1983. Some properties of asparagus virus II isolated from Asparagus offlcinalis in Japan. Ann. Phytopath. Soc. Japan 49:683-688. 11. George, J. A., and Davidson, T. R. 1963. P o l l e n transmission of necrotic ringspot and sour cherry yellows viruses from tree to tree. Can. J. Plant Sci. 43:276-288. 12. Gibbs, A., and Harrison, B. 1979. Plant Virology: The Principles. John Wiley and Sons, New York. 292 pp. 84 13. Hamilton, R. I., Leung, E., and Nichols, C. 1977. Surface contamination of pollen by plant viruses. Phytopathology 67:395-399. 14. Hamilton, R. I., Nichols, C., and Valentine, B. 1984. Survey for prunus necrotic ringspot and other viruses contaminating the exine of pollen collected by bees. Can. J. P l a n t Path. 6:196-199. 15. Hartung, A. C., Evans, T. A., and Stephens, C. T. 1985. Occurrence of asparagus virus II in commercial asparagus fields in Michigan. Plant Dis. 69:501-504. 16. Hein, A. 1963. Viroses am spargel. Mitt. Biol. Bundesanst. Land-Forstwirtsch, Berlin-Dahlem 108:70-74 (Rev. Appl. Mycol. 1964. 43:455). 17. Hein, A. 1969. Uber viruskrankheiten das spargels (Asparagus officinalis L.). Spargelvirus I. Z. Pflanzenkr. Pflanzenpathol. Pflanzenschutz 76:395406 (Rev. Plant Pathol. 1970. 49:219). 18. Madden, L. V., R a ymon d Louie, J. J., and Knoke, J. K. 1982. Evaluation of tests for randomness of infected plants. Phytopathology 72:195-198. 19. Mink, G. I., and Uyeda, I. 1977. Three mechanically transmissible viruses isolated from asparagus in Washington. Plant Dis. Rep. 61:398-401. 20. Querfurth, G., and Paul, H. L. 1979. Protein A-coated latex-linked antisera (PALLAS): New reagents for a sensitive test permitting the use of antisera unsuitable for the latex test. Phytopath. Z. 94:282-285. 21. Uyeda, I. 1978. Identification, characterization, and incidence of viruses isolated from asparagus. Ph.D. thesis, Washington State University. 115 pp. 22. Uyeda, I., and Mink, G. I. 1981. Properties of asparagus virus II, a new member of the Ilarvirus group. Phytopathology 71:1264-1269. 23. Voller, A., and Bidwell, D. E. 1977. Enzyme immunoassays and their potential in diagnostic virology. Pages 449-457 in. Comparative Diagnosis of Viral D i s e a s e s Vol. II. E. K u r s t a k and C. Kurstak, eds. 24. Weissenfels, M., and Schmelzer, K. 1976. Untersuchungen ueber das schadaumass durch viren an spargel (Asparagus officinalis L.). Arch. Phytopathol. Pflanzenschutz, Berlin. 12:67-73. 85 25. Yang, H. 1979. Early effects of viruses on the growth and productivity of asparagus plants. HortScience 14:734-735. 26. Yang, H.r and Clore, W. J. 1976. Obtaining virus-free plants of Asparagus officinalis L. by culturing shoot tips and apical meristems. HortScience 11:474-475. CHAPTER IV INTERACTION BETWEEN ASPARAGUS VIRUSES AND PATHOGENIC FUSARIUM SPECIES IN ASPARAGUS Introduction Declining asparagus yields in Michigan are attributed ultimately to crown and root rot incited by Fusarlum oxysporum f.sp. asparaqi and Fusarium monlliforme in stressed plants (11). It is the author's hypothesis that no single stress factor is responsible for this decline in vigor and longevity. Rather, several cultural, environmental or biological stresses on the asparagus plant may predispose it to infection by the Fusarium crown and root rotting organisms. A number of stress factors have been Identified in asparagus production including tillage (17), improper soil pH (A, Putnam, unpublished data), allelopathic compounds (10), defoliation caused by rust (M, Lacy, unpublished data), virus infection (21) and the over-harvesting of spears (18). The effects of over-harvesting on vigor and productivity of asparagus is well understood (18). Other research indicates that harvest stress may lead to a reduction in the ability of asparagus plants to wall off and lignify infection courts of pathogenic Fusarium species (R. Endo, personal communication). This could result in an increased incidence and severity of crown and root rot, thereby contributing to the problem of "asparagus decline". 86 87 Virus Infection has been demonstrated to negatively affect the vigor and productivity of asparagus. Asparagus plantings in Germany infected with cucumber mosaic virus, asparagus virus I or a spa r a g u s v i r u s II showed a 29 to 44% reduction in y i e l d and a 15 to 20% d e c r e a s e in number of spears and height of ferns (25). In Washington, asparagus plants infected with either asparagus virus I or asparagus virus II exhibited a mild reduction in vigor and productivity (21). Plants infected with both viruses showed severe decline and mortality in the second year in the field. Asparagus virus II has been detected widely within most commercial asparagus plantings in Michigan and asparagus vir u s I has al s o been dete r m i n e d to be present, but not widespread (Chapter 2, this manuscript). These viruses may constitute important biological stress factors leading to an increased incidence and severity of Fusarium crown and root rot in asparagus. A number of studies indicated that fungal infections of plant roots were increased when plants are also infected with one or more viruses (1, 5, 19, 24). Two possible mechanisms are proposed for the increased fungal root rot induced by virus infection. Either the inherent susceptibility of root tissue is increased by virus infection or increased leakage of nutrients from roots of virus-infected plants increases the inoculum potential of the fungi in the plant rhizosphere (2). Farley and Lockwood (7) demonstrated that pea plants became more diseased when inoculated with Fusarium solani 88 when they were first infected with any one of four different viruses. Beute and Lockwood (3), working with the same system, determined that the increased Fusarium root rot in peas infected wi t h bean y e l l o w mosaic v i r u s was not due to an increased susceptibility of root tissue but rather that virus infection increased the exudation of nutrients from roots thereby increasing the Inoculum potential of the pathogen within the rhizosphere. The roots of virus-infected pea plants released more electrolytes, carbohydrates, amino acids and nucleotides than did the roots of healthy plants. They attributed this increased exudation to an increase in the permeability of cell membranes of the root. The objective of this investigation was to evaluate the effect of infection by asparagus virus I and/or asparagus virus II on disease severity caused by the Fusarium crown and root rotting organisms in asparagus, and to investigate changes in root exudate composition and the ability of the root to lignify the area surrounding infection courts. 89 Materials and Methods Plant Material Preparation of seed All asparagus seed was first surface disinfested of Fusarium with a 10,000 mg/ml benomyl solution in acetone by placing on a orbital shaker overnight. Seeds were washed free of benomyl with 100 to 200 ml of acetone and air dried. Surface disinfested asparagus seeds were pregerminated on moistened sterile filter paper in petri dishes in the dark. When radicles had emerged, they were transferred to flats containing standard commercial potting mix and maintained in a greenhouse at 21 to 24 C and a 16 hour photoperiod for 2 months. Preparation of seedl lngs Virus-free and AV II-infected asparagus seedlings were prepared using a seedlot of cultivar Mary Washington 500 (MW 500) that had been determined to be approximately 50* infected with AV II [provided by G.I. Mink]. after germination, At six weeks seedlings were indexed for A V II using the enzyme-1 inked immunosorbent assay (ELISA) method of Voller and Bidwell (23). Asparagus seedlings (MW 500) either infected with AV I or d o u b l y infected with both AV I and A V II were pr e p a r e d from healthy and AV II-infected plants by aphid transmission. Green peach apids (Myzus persicae) were allowed to feed for 2 weeks on asparagus plants known to be infected only with AV I, then transferred to healthy or AV II-infected 6-week old 90 asparagus plants. An equal number or non-viruliferous green peach aphids were al lowed to feed on a s e c o n d group of healthy or AV II-infected asparagus seedlings as a control. After 2 weeks all stems were cut at soil level and plants were transferred to aphid-free cages. Plants were indexed for the pre s e n c e of A V I and A V II after the emergence of the second new shoot by rub-inoculating to C. quinoa. Plants were maintained in a greenhouse at 21 to 24 C and a 16 hour photoperiod for 5 to 14 days and observed for the development of typical AV I and AV II symptoms. Single necrotic lesions developing in 10 to 14 days were examined with serologically specific electron microscopy (SSEM) for AV I particles using the procedure of Derrick (6). Preparation of tissue culture c l ones Healthy and AV II-infected asparagus lines (cultivar Viking KB3) were prepared by culturing shoot tips and apical meristems from field grown asparagus plants known to be infected o n l y w i t h A V II usi n g the methods of Yan g and C l o r e (22). Explants of tissue were obtained from spears about 20 cm long and 2 cm in diameter. All subsequent procedures were carried out in a laminar flow hood. The surface of each spear was washed with water and then sterilized with 0.525* sodium hypochlorite solution for 10 min. The scales on the tip of each spear were r e m o v e d and meri s t e m s and shoot tips were excised under a binocular dissecting microscope with a knife made from a razor b l a d e chip a t t a c h e d to a woo d e n handle. 91 Two types of tissue explants were used. Apical domes less than 0.1 mm in height and free of leaf primordia were used for the production of virus-free plants and shoot tips greater than 2 mm in height with several leaf primordia for the production of AV II-infected plants. Isolated meristems and shoot tips were placed in Pyrex tubes (10 x 2.5 cm) on 10 ml of modified Murashige and Skoog's medium (MMS) (16) with 0.1 ppm naphthaleneacetic acid (NAA) and 0.1 ppm kinetin (6furfurylamino purine). The medium was adjusted to pH 5.7 with 1 N NaOH or 1 N HC1 and a u t o c l a v e d at 121 C and 20 psi for 15 min. Tubes containing meristems and shoot tips were placed in a growth chamber and maintained at 25 C with a 16 hour photoperiod. After 2 months the plantlets were transferred to 125-ml flasks containing 50 ml MMS medium with 0.1 ppm NAA and 0.3 ppm kinetin. One pi ant let was placed into each flask and after 2 to 3 months p l a n t l e t s wer e indexed for AV II using the procedure of Mink and Uyeda (13). A sample of the shoot tissue of each plantlet was triturated in a small volume of 0.01 M sodium phosphate buffer, mercaptethanol added. pH 7.0, with O.lfc 2- The homogenate was rubbed onto the surface of Chenopodium quinoa leaves that had previously been dusted with carborundum (320 mesh). Inoculated plants were maintained in a greenhouse at 21 to 24 C and a 16 hour photoperiod and assessed for symptom development 5 to 10 days after inoculation. 92 Source and Maintenance of Fungi Isolates of Fusarium oxysporum (Schelcht.) emend. and Snyder Hansen f.sp. asparagi Cohen and F. moniliforme (Sheldon) emend. Snyder and Hansen, originally Isolated from diseased asparagus, were kindly provided by M. Lacy, Michigan State University. Fungi were maintained in sterile soil at 4 C until use. Inoculation Procedures Preparation of colonized mi 1 let inoculum Soil containing F. oxysporum f.sp. asparagi and F. mo n l liforme was dusted onto potato dextrose agar (PDA) [Difco Laboratories, Detroit, Ml 48232] plates and grown for 14 to 21 days at 24 C. Millet colonized by Fusarium isolates was prepared using a modification of the method of Goth and Johnston (9). One hundred ml of glass distilled water was added to 200 g of m i l l e t in a f l a s k and a u t o c l a v e d for 1 hour at 20 psi on two consecutive days. After each autoclaving, millet was first allowed to cool and the flask was shaken vigorously until all large clumps of millet had been broken. Sterile millet was inoculated with one 4 mm diameter plug of PDA colonized by the appropriate Fusarium isolate and the flask was Incubated at room temperature for 10 to 14 days. Each flask was shaken vigorously daily to ensure an even distribution of the fungus within the inoculum. Inoculation of plants Two-month old asparagus seedlings or tissue cultured asparagus clones of comparable size were challenge inoculated 93 w i t h 8 g of infested or uninf e s t e d m i l l e t per 1000 g of pasteurized sandy loam soil. This concentration of inoculum was previously determined to provide a moderate level of disease pressure (T. Evans, unpublished data). Plants were maintained in a greenhouse at 21 to 24 C with a 16 hour photoperiod. Preparation of Conidial Suspensions Conidia were removed from the agar surface with glass distilled water and gentle agitation then filtered through two layers of sterile cheescloth to remove most hyphal fragments. Conidia were washed by centrifuging at 6,000 rpm for 15 min and resuspendin g the p e l l e t in 50 ml of s t e r i l e glass distilled water. This process was repeated three times and the final suspension was adjusted to the appropriate spore concentration. Evaluation of Inoculated Plants Disease severity of roots was evaluated 2 months after Fusarium inoculation on a scale of increasing severity from 1 to 5 where 1 = h e a l t h y plant, no v i s i b l e root lesions, no rotted roots or crown discoloration; 2 = few root lesions (15) and/or rotted roots, no crown discoloration, in number or shoots or roots; no reduction 3 = moderate number of root lesions (6-10) and/or rotted roots, no or slight crown discoloration, shoots reduced, feeder roots reduced; 4 = many root lesions (>10) and/or rotted roots, shoots reduced, crown discoloration, feeder roots sparse; 5 = many root lesions (>10) and/or rotted roots, crown discoloration, shoots 94 greatly reduced, feeder roots greatly reduced or absent or dead. Influence of Virus Infection on Root Rot Severity The effect of virus infection on subsequent infection by F. oxysporum f.sp. asparagi and F. mon i 1 iforme investigated in the greenhouse. was Two-month old healthy and AV II-infected asparagus seedlings or asparagus clones were challenge inoculated with Fusarium-infested millet as described above. The interaction between infection with AV I and/or AV II and subsequent infection by F. oxysporum f.sp. asparagi was evaluated using the procedures described above. Inoculated plants were maintained in the greenhouse at 21 to 24 C and 16 hour photoperio d and f e r t i l i z e d at 1 m o n t h wit h 100 ml of full strength Peter's 20-20-20 [W.R. Grace and Company, Fogelsvile, PA 18051]. Plants were evaluated for root rot severity 2 months after inoculation. Influence of Root Exudates From Healthy and Virus-Infected Asparagus on Root Rot Severity The influence of root exudates from healthy and virusinfected asparagus plants on root rot severity was investigated in the greenhouse. Five two-month old healthy or AV II-infected asparagus seedlings were grown in pasteurized sandy loam soil in 25.4 cm plas t i c pots and w a t e r e d twice daily with 100 ml of glass distilled water for 2 months. Leachates from these pots were allowed to drain onto the soil surface of pots containing virus-free asparagus plants inoculated with either F. oxysporum f.sp. or F. moniliforme- 95 colonized millet. After 2 months, plants that were watered with leachates were evaluated for disease severity using the procedure described above. Collection of Root Exudates From Healthy and VirusInfected Asparagus Healthy and AV-II infected clones were grown in liquid culture supported by nylon screens in staining dishes (10 x 8 x 6 cm). Four plantlets were grown in each dish with four replicates per treatment. Dishes were placed in autoclaved polypropylene bags [Bel-Art Products, Pequannock, NJ 07440] and maintained in a growth chamber at 25 C and 16 hour photoperiod. Plants were grown in full strength Hoagland's solution (4) for 6 days, washed with glass distilled water then transferred to 100 ml double distilled water for 24 hours for exudate collection. Plants were returned to fresh full strength Hoagland's solution and this pattern of exudate collection was continued for 4 weeks. All solutions containing root exudate samples were passed through 0.20 pm filters (Nalgene Sterilization Unit, Type S) [Nalgene Company, Rochester, NY 14602] and stored at 4 C. Solutions were concentrated 10-fold in a flash evaporator at 50 C and passed through 0.45 pm Millipore filters and stored in sterile tubes at 4 C or frozen. Chemical Analysis of Root Exudates From Healthy and VirusInfected Asparagus The conductivity of water culture solutions was determined with a standard conductivity bridge and the results were expressed in jumhos, the reciprocal of electrical 96 conductivity. Total carbohydrates of root exudates were determined using a modification of Dreywood's anthrone reagent (15). The anthrone reagent was prepared by dissolving 0.4 g of anthrone in 200 ml of 9.3 M H 2S04 . One ml of root exudate s a m p l e was mixed t h o r o u g h l y wi t h 2 ml of anthrone reagent then placed into a boiling water bath for 3 min. Samples were quickly cooled and the optical density at 620 nm measured in a Bausch & Lomb spectronic 20. A standard curve was prepared from 1 ml samples containing 10, 20, 40, 80, and 160 jig of glucose. Glucose content of root exudates was determined with an enzymatic colorimetric test (No. 115) [Sigma Chemical Co., St. Louis, M0, manufacturer's directions. 63178] using the Optical densities were read at 450 nm in a Bausch & Lomb Spectronic 20 and compared with a standard curve prepared using 1-ml samples containing 10, 20, 40, 80, and 160 ;ug of glucose. Total amino acids were determined using the ninhydrin method of Moore (14). A 2-ml sample of root exudate was mixed thoroughly with 1 ml of a ninhydrin reagent solution (No. N 1632) [Sigma Chemical Co., St. Louis, M0 heated in a boiling water bath for 15 min. 63178] and Samples were quickly cooled to below 30 C, vigorously shaken and optical density measure at 570 nm in a Bausch & Lomb spectronic 20. A standard curve was prepared from 2-ml samples containing 4, 8, 16 and 32 jug of glycine. Protein was determined with Folin Ciocalteu's Phenol Reagent (Micro-Protein Determination Kit) [Sigma Chemical Co., St. Louis, manufacturer's directions (12). M0 63178] using the Test samples were first 97 diluted with a 0.85* sodium chloride solution in distilled water (w/v) so that the final protein concentration was between 150 and 1000 jig/ml. A 2.2 ml aliquot of Biuret Reagent (No. 690-1) was added to 0.2 ml of diluted test solutions and to 0.2 ml of the sodium chloride solution, mixed thoroughly and allowed to stand at room temperature for 10 min. Then, 0.1 ml of Folin Ciocalteu's Phenol Reagent (No. 690-2) was added, mixed thoroughly and allowed to stand at room temperature for 30 min. Samples were transferred to a cuvet and their o p tical d e n s i t y at 725 nm d e t e r m i n e d in a Bausch & Lomb spectronic 20. A standard curve was prepared from 0.2 ml samples containing 5, 10, 20, 40 and 80 jog of bovine serum albumin. The Effect of Root Exudates From Healthy and Virus-Infected Asparagus on the Germination of Conidia and Germ Tube Growth The effect of diluted root exudates on the germination of conidia and subsequent germ tube growth for F. m o n i 1 iforme and F. oxysporum f.sp. asparagi was measured. Root exudates were diluted ten-fold by adding 0.1 ml sterile root exudate to a solution containing 0.1 g agar [Difco Laboratories, Detroit, MI 48232] in 9.9 ml of glass distilled water that had been p r e v i o u s l y a u t o c l a v e d at 121 C and 15 psi for 20 min and cooled to 55 C in a water bath. Washed conidia were pipetted onto the surface of water agar plates amended with root exudates from virus-infected and healthy asparagus, spread uniformally with the aid of a sterile glass rod and incubated at 24 C. Germination of conidia and subsequent germ tube growth was measured hourly using a substage 98 binocular dissecting microscope and ocular micrometer. A minimum of 25 spores were monitored on duplicate plates for each replication. The Effect of Virus Infection on Lignin Formation Within Asparagus Roots in Response to Infection by Fusarium spp. The effect of virus infection on the ability of asparagus seedling roots to lignify and wall off infection courts of F. oxysporum f.sp. asparagi and F. moni 1 iforme was evaluated in the greenhouse. Healthy and AV II-infected two- month old asparagus seedlings were challenge inoculated with F. oxysporum f.sp. asparagi and F. monil iforme as previously described. Two months after inoculation plants were evaluated for disease severity as before and root tissue with distinct lesions was excised. Root lesions were separated into two categories by size: 1 mm to 5 mm in length and those 5 mm to 10 mm in length. Root tissue with lesions was hand- sectioned using a razor blade and stained with 1% phloroglucinol in 50% HC1 (8) and rated for relative degree of lignification in the region surrounding the infection court. A m i nimum of 100 sect i o n s were e v a l u a t e d on a s c a l e of increasing 1 ignification from 1 to 5 for each treatment. 99 Results Influence of Virus Infection on Root Rot Severity Two month old asparagus seedlings or comparable tissue culture clones infected with AV II became significantly (P=0.05) more diseased than virus-free plants when challenge inoculated with F. oxysporum f.sp. asparagi (Table 4.1). The difference in disease severity was smaller between AV IIinfected and virus-free asparagus plants when challenged with F. moniliforme (Table 4.1). In a second experiment, asparagus seedlings infected with AV I or AV II were significantly (P=0.05) more diseased than virus-free seedlings when inoculated with F. oxysporum f.sp. asparagicolonized millet (Table 4.2). Asparagus seedlings doubly infected with AV I and AV II became significantly (P=0.05) more diseased than seedlings infected with either AV I or AV II alone. Influence of Leachates From Healthy and Virus-Infected Asparagus on Root Rot Severity Root rot severity in virus-free asparagus plants watered with leachates from AV II-infected asparagus was significantly (P=0.05) greater than that of virus-free asparagus watered with leacheates from virus-free asparagus (Figure 4.1). This difference in root rot severity was greater for plants challenge inoculated with F. oxysporum f.sp. asparagi than with F. moni11 forme. 100 Table 4.1. Disease severity of clones of Asparagus officinalis variety KB3 with and without asparagus virus II (AV II) when inoculated with Fusarium oxysporum f.sp. asparagi (FOA) or F. moniliforme (FM) colonized millet. Disease Rating^ Treatment Virus-free Virus--infected FOA 1.3 + 0.4Z 3.3 + 0.4 FM 2.3 + 0.3 3.0 ± 0.4 Control 1.0 + 0.0 1.4 + 0.4 ^Disease ratings are 1-5 with 1 = healthy plant, 2 = few root lesions and/or rotted roots, 3 = moderate number root lesions and/or rotted roots, 4 = many lesions and/or rotted roots, 5 = many root lesions and/or rotted roots or plant dead. 2Values are the mean for 10 plants plus or minus the standard error. Asterisks (* =) means for virus-infected plants differ (P=0.05) from corresponding virus-free plants according to LSD test. * * 101 Table 4.2. The effect of infection with asparagus virus I (AV I) or asparagus virus II (AV II) aiane or AV I and AV II together on disease severity in seedlings of Asparagus officinalis when inoculated with Fusarium oxysporum f.sp. asparagi (FOA) colonized millet. Disease Rating31 Treatment Controly FOA Virus-free 1.1 az 2.2 a AV I 1.2 a 3.2 b AV II 1.1 a 3.2 b 1.3 a 4.1 c AV I & AV II disease ratings are 1-5 with 1 = healthy plant, 2 = few root lesions and/or rotted roots, 3 = moderate number root lesions and/or rotted roots, 4 = many lesions and/or rotted roots, 5 * many root lesions and/or rotted roots or plant dead. ^Values are the mean for 10 plants. Within columns, numbers followed by a common letter are not significantly different (P=0.05) according to Duncan's multiple range test. 102 Figure 4.1. Effect of leachates from AV II-infected and virusfree asparagus on Fusarium crown and root rot. Virus-free asparagus plants inoculated with Fusarium oxysporum f.sp. asparagi (FOA), F. moniliforme (FM) or sterile millet received leachings from 5 AV IIinfected or healthy plants which were water with 50 ml glass distilled water twice daily over a 2 month period. Disease severity was determined on a scale of 1-5. X LU LlJ in < CONTROL 4 V IR U S -F R E E AV II-INFECTED ^ 103 Q 5 3 2 GO Q 1 • 0 C o n tro l FM FOA 104 Changes In Root Exudates of Virus-Infected Asparagus Clones Asparagus clones infected with A V II and grown in liquid culture released three times more electrolytes in a 24 hour period than did virus-free clones (Figure 4.2). separate experiments, In two AV II-infected asparagus clones grown in liquid culture released more glucose, total carbohydrates and amino acids than did virus-free asparagus clones (Figure 4.3). There were no differences in the amount of protein released by the roots of virus-infected and healthy asparagus plants. Each healthy asparagus plant released approximately 27 jag of carbohydrate (2.5 jug of this determined to be glucose) and less than 1 jug of amino acids per day. Asparagus plants infected with AV II each released about 50 ug of carbohydrate (10 jug of this determined to be glucose) and 8 jug of amino acids over the same period. There was a 100% increase in carbohydrate and a 800% increase in amino acids in the root exudates of virus-infected asparagus plants. The Effect of Root Exudates From Healthy and Virus-Infected Asparagus on the Germination of Conidia and Germ Tube Growth The influence of root exudates from healthy and virus infected asparagus on the germination of conidia and growth of germ tubes of F. oxysporum f.sp. and F. moniliforme was investigated. A significantly greater number of microconidia and macroconidia of F. oxysporum f.sp. asparagi and F. moniliforme germinated earlier on water agar amended with root exudates from virus-infected asparagus than on water agar amended with root exudates from healthy plants or water 105 Figure 4.2. Increased exudation of electrolytes by AV IIinfected asparagus clones in liquid culture relative to the exudation of virus-free clones. Values are the average release by 4 plantlets over 3 separate twenty-four hour periods. RELATIVE CONDUCTIVITY OF ROOT EXUDATES 20 30 50 60 70 80 90 100 (jumhos) 110 120 106 m m 107 Figure 4.3. Increased exudation of glucose, total carbohydrates, amino acids and proteins in AV II-infected asparagus clones grown in liquid culture relative to exudation of virus-free clones. Values are the average release by 4 plantlets over 3 separate twenty-four hour periods. EXUDATE ()jg/p!ant/day) o E o o to m 801 109 agar alone (Figure 4.4). Germ tubes from microconidia or macroconida of F. oxysporum f.sp. asparagi were significantly (P=0.05) longer after 12 hours on water agar amended with root exudates from virus-infected asparagus than on water agar amended with root exudates from healthy plants or unamended water agar (Table 4.3). Root exudates from virus- free virus-infected asparagus plants had an equal influence on the growth of germ tubes from conidia of F. m o n i liforme. Germ tubes of conidia grown on unamended water agar for 12 hours were the shortest. The Effect of Virus Infection on Lignin Formation Within Asparagus Roots in Response to Infection by Fusarium spp. Virus infection of asparagus seedlings signifcantly (P=0.05) reduced the amount of lignin produced in the areas of the root surrounding the infection courts of F. oxysporum f.sp.asparagi and F. m o n i Ilforme. Virus-free asparagus plants had a mean lignification rating of 3.5 to 4.0 whereas virus-infected plants had a mean rating of only 1.3 to 1.8 (Table 4.4). There were no apparent differences in the amount of lignin produced in response to infection by F. oxysporum f.sp. asparagi or F. moni1 iforme. 110 Figure 4.4. The effect of root exudates from healthy and virusinfected asparagus on the germination of conidia of Fusarium maniliforme and F. oxysporum f. sp. asparaqi. 111 o-o VIRUS-FREE B-B VIRUS-INFECTED 100 90 80 70 60 50 40 30 20 10 0 1 100 - 90 ' 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 10 2 0 21 22 2 3 2 4 25 7 8 9 10 11 12 13 14 15 16 17 18 10 20 21 2 2 2 3 2 4 25 o-o VIRUS-FREE B-B VIRUS—INFECTED 8070 6050403020 ' 10 - 01 2 3 4 5 6 TIME (hou rs) 112 Table 4.3. The effect of root exudates from healthy and AV IIinfected asparagus clones on the growth of Fusarium oxysporum f.sp. asparaqi (FOA) and F. monlliforme (BM) germ tubes. Germ Tube Length x (ocular micrometer units) Treatment FOA FM Root Exudates y Virus-free 0.94 b z 3.50 a AV II-infected 2.50 a 3.90 a Water Agar Alone 1.05 b 3.50 a PDA Alone 2.20 a 3.80 a xValues are the mean germ tube lengths at 12 hours for 50 spores per treatment. yExudates from plants grown in liquid culture were diluted ten-fold in water agar. zWithin columns, numbers followed by a common letter are not significantly different (P=0.05) according to Duncan's multiple range test. 113 Table 4.4. Effect of infection with asparagus virus II (AV II) on lignification within asparagus seedling roots in response to infection by Fusarium oxysporum f.sp. asparaqi (FOA) or F. maniliforme (FM). Relative degree of lignificatiany Virus Content 1 ran to 5 nan root lesions 5 mm to 10 mm root lesions none 3.7 + 0.8' 4.1 + 0.7 AV II 1.8 + 0.7 1.3 + 0.7’ none 3.6 + 0.5 4.0 + 0.5 AV II 1.3 + 0.4 1.5 + 0.6 FOA FM yHand-sectians of root lesions were first stained with 1% phloroglucinol in 50% HC1 and infection courts rated on a scale of 1 to 5 for the degree of lignification with 1 = no lignification; 2 = light lignification; 3 = moderate lignification 4 = heavy lignification and 5 = very heavy lignification. zValues indicate the mean of 100 sections plus or minus the standard error. Asterisks (* =) means for AV II-infected plants differ from corresponding virus-free plants according to LSD test. 114 Discussion Two-month old asparagus seedlings or comparable tissue culture clones infected with AV II became significantly more diseased than virus-free plants when inoculated with either Fusarlum oxysporum f.sp. asparaqi or F. m o n i l iforme. The virus-induced Increase in root rot severity observed in asparagus is due in part to an alteration in the composition of root exudates. The direct effect of root exudates on the severity of Fusarium crown and root rot was demonstrated in greenhouse experiments. Virus-free asparagus seedlings inoculated with F. oxysporum f.sp. asparaqi or F. moniliforme and watered with root exudates from virus-infected plants showed increased severity of root rot compared with those plants watered with root exudates from virus-free asparagus. Electrolytes, carbohydrates and amino acids were exuded in larger amounts from the roots of asparagus plants infected with A V II than from healthy plants. Leakage of electrolytes from plant cells is indicative of increased permeablitly of cell membranes (3) and the exudates of virus-infected plants contained three times more electrolytes than those of healthy plants. The quantity of carbohydrate exuded by roots of virusinfected asparagus plants growing in liquid culture was two times that of virus-free plants. Amino acids in the exudates of virus-infected plants was eight to ten times greater than that of virus-free plants. This large increase in the quantity of amino acids exuded by the roots of virus-infected 115 plants may play a critical role in increasing the severity of root rot seen in these plants. Root exudates from virus-infected asparagus plants may increase the inoculum potential in the rhizosphere of these root rot fungi thereby increasing the severity of disease in virus-infected asparagus. Soil-borne pathogens such as F. oxysporum f.sp. asparaqi and F. moniliforme attack their host in a sequence of steps. These steps include germination of spores, mycelial growth, penetration of host tissue and pathogensis. Each step is influenced by the nutrient environment. Nitrogen and carbon are clearly the most important nutrients in the soil and can greatly effect the pathogenic development of fungi. Toussoun et al (20) showed that the germination of conldia of F. solani f.sp. phaseoll was favored by glucose. Glucose also stimulated saprophytic mycelial growth but delayed the penetration of the bean hypocotyls and pathogenesis. On the other hand, colonization of bean hypocotyls and pathogenesis was favored by nitrogen. Similarly, Beute and Lockwood (3) observed that disease severity in F solani-infected peas was increased by the addition of amino acids and that glucose had no affect. Healthy asparagus roots have a propensity to ward off colonization by Fusarium prior to infection of the stele. reduced ability of virus-infected asparagus plants to w alloff and lignify infection courts of pathogenic Fusarium contributes to the increase in disease severity observed in these plants. Histological studies by Smith and Peterson (19) have revealed that wall appositions are produced by The 116 healthy asparagus roots in response to penetration by F. oxysporum f.sp. asparaqi. These apposition materials were produced in the region immediately basipetal of the root meristems and were composed of polysaccharides, acid mucopolysaccaride, callose and phenolic substances. The mechanism by which virus-infection leads to an increase in root rot appears to be two-fold. First, virus infection leads to an increased permeability of cell membranes of the root, nutrients, Secondly, resulting in an increased leakage of Including carbohydrates and amino acids. the roots of virus-infected asparagus have a reduced ability to synthesize lignin as barriers to infection by F. oxysporum f.sp. and asparaqi and F. moniliforme. 117 Literature Cited 1. Bateman, D. F. 1961. Synergism between cucumber mosaic virus and Rhizoctonia in relation to rhizoctonia damping-off of cucumber. (Abstr.) 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