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Detection of Ralstonia solanacearum in Irrigation Ponds and Semi-Aquatic Weeds, and Its Chemical Treatment in Water

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PAGE 1

DETECTION OF Ralstonia solanacearum IN IRRIGATION PONDS AND SEMIAQUATIC WEEDS, AND ITS CHEMICAL TREATMENT IN WATER By JASON CHRISTOPHER HONG A THESIS PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMEN T OF THE REQUIRMENTS FOR THE DEGREE OF MASTER OF SCIENCE UNIVERSITY OF FLORIDA 2005

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Copyright 2005 by Jason Christopher Hong

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ACKNOWLEDGMENTS I would like to thank my committee members, Drs. Timur M. Momol, Jeffrey B. Jones, Steve M. Olson, and Jerry A. Bartz, for their support and guidance through the entire research project and preparation of this manuscript. I especially would like to express my appreciation for Dr. Momols patience and guidance throughout this project, and Dr. Jones understanding and mentoring that he has shown. I would like to express my gratitude for the members of Dr. Momols lab for their support and help. I am grateful for Pingsheng Ji for teaching me the various techniques I would need to know for this project. I am grateful to Hank Dankers for showing me the location of the ponds and helping me with the work, and for Laura Ritchie for accompanying me on my visits to the ponds. I am also grateful to the members of Dr. Jones lab. I would like to express my gratitude for the instruction, aid, service, and advice I received from Ellen Dickstein and Jerry Minsavage. I would also like to thank friends that helped me throughout this project by providing long hours of service and encouragement. And, my deepest gratitude is expressed to my parents for their love and support. iii

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TABLE OF CONTENTS page ACKNOWLEDGMENTS .................................................................................................iii LIST OF TABLES ...............................................................................................................v LIST OF FIGURES ...........................................................................................................vi ABSTRACT ......................................................................................................................vii CHAPTER 1 INTRODUCTION........................................................................................................1 2 LITERATURE REVIEW...........................................................................................10 Detection of Ralstonia solanacearum in Irrigation Water and Solanum dulcamara.10 Methods for Controlling Ralstonia solanacearum.....................................................18 3 MATERIALS AND METHOD..................................................................................22 4 RESULTS...................................................................................................................30 5 DISCUSSION AND CONCLUSIONS......................................................................45 LIST OF REFERENCES...................................................................................................51 BIOGRAPHICAL SKETCH.............................................................................................57 iv

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LIST OF TABLES Table page 4.2 Incidence and bacterial concentration of R. solanacearum in water samples by direct isolation on modified SMSA agar from August 2003 to May 2005..............31 4.3 Correlation between water temperature and levels of R. solanacearum for ponds 1 and 2 from August 2003 to May 2005..................................................................32 4.4 Detection of R. solanacearum isolated from sterilized or nonsterilized roots and/or stems of semi-aquatic plants collected from ponds 1 and 2 by and direct plating onto modified SMSA agar in July 2004.......................................................33 4.5 Incidence R. solanacearum in surface sterilized and nonsterilized stems and roots of Polygonum pennsylvanicum and Hydrocotyle ranunculoides associated with irrigation pond number1 and 2, from July 2004 to May 2005.........................34 4.1 Populations (cfu/ml) of R. solanacearum in five irrigation ponds from August 2003 to May 2005....................................................................................................38 v

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LIST OF FIGURES Figure page 4.1 Sensitivity of modified SMSA for detecting R. solanacearum was determined by comparing a dilution series of known concentrations of the bacterium spread plated on modified SMSA and nutrient agar............................................................30 4.7 The effects of free chlorine and hydrogen peroxide at various concentrations on 10 8 cfu/ml of Ralstonia solanacearum.....................................................................36 4.8 The effects of free chlorine and hydrogen peroxide at various concentrations on 10 4 cfu/ml of Ralstonia solanacearum.....................................................................36 4.2 Detection of R. solanacearum colonies by direct isolation on modified SMSA medium during a survey of Pond 1 from August 2003 to August 2004..................40 4.3 Detection of R. solanacearum colonies by direct isolation on modified SMSA medium during a survey of Pond 2 from August 2003 to May 2005.......................41 4.4 Detection of R. solanacearum colonies by direct isolation on modified SMSA medium during a survey of Pond 3 from April 2004 to May 2005..........................42 4.5 Detection of R. solanacearum colonies by direct isolation on modified SMSA medium during a survey of Pond 4 from August 2003 to May 2005.......................43 4.6 Detection of R. solanacearum colonies by direct isolation on modified SMSA medium during a survey of Pond 5 from May 2004 to May 2005...........................44 vi

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Abstract of Thesis Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Master of Science DETECTION OF Ralstonia solanacearum IN IRRIGATION PONDS AND SEMI-AQUATIC WEEDS, AND ITS CHEMICAL TREATMENT IN WATER By Jason Christopher Hong August 2005 Chair: Timur M. Momol Cochair: Jeffrey B. Jones Major Department: Plant Pathology Bacterial wilt caused by Ralstonia solanecearum is one of the most destructive bacterial diseases in the tropical, sub-tropical, and temperate regions of the world, including the southern U.S. and Florida. It is causing serious yield losses on many crops such as tomato, potato, pepper, eggplant, tobacco, banana and geranium. It is estimated that the bacterium is pathogenic on several hundred plant species in over 50 families throughout the world. On tomato and other plants, R. solanacearum colonizes and blocks the vascular tissue resulting in wilting and eventual death. In August 2002, R. solanacearum (r1/b1) was detected in the irrigation pond water of tomato farms and ornamental nurseries in northern Florida. Research was initiated to sample the irrigation pond water for the presence of R. solanacearum. In addition to water sampling, various aquatic weeds were sampled to determine their relationship with the bacterium. The long term goal of this study was to improve our understanding of R. solanacearum in irrigation water and weed hosts (including semi-aquatic weeds) in Florida. The vii

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objectives of this research were to study the populations of R. solanacearum within irrigation ponds by analyses of water and semi-aquatic weeds and other weed samples, and to determine a method to reduce the bacterial populations before irrigating tomato fields in Florida. It was determined that temperature had a direct effect on the population of R. solanacearum. During the summer months as the temperature increased the population of the bacterium increased reaching populations as high as 5.6 x 10 4 cfu/ml. By December the bacterium was undetectable when spread plated on SMSA, even when concentrated by centrifugation. In May of the following year, the bacterium was re-detected. A survey of semi-aquatic weeds associated with the ponds for the detection of R. solanacearum indicated that the weeds that were infected with the bacterium were Hydrocotyle ranunculoides (common name dollar weed), and Polygonum pennsylvanicum (common name Pennsylvania Smut weed). Both H. ranunculoides and P. pennsylvanicum were latently infected, showing no signs of wilt. The bacterium was detected from the weeds by direct plating on modified SMSA in the summer months, but not in the winter months. A study also was performed to determine the effects of chlorine and hydrogen peroxide for possible control of the bacterium in irrigation water. Chlorine was effective at 2 mg/l at a bacterial concentration of 10 4 cfu/ml, while hydrogen peroxide was not effective. viii

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CHAPTER 1 INTRODUCTION Florida tomato producers are the primary providers for fresh market field-grown tomatoes (Lycopersicon esculentum Mill.) in the United States year around. The state accounts for 50% of the domestically produced tomatoes each year, and tomatoes provide 30.3% of the income of vegetables grown in Florida. During the 2003-2004 season, approximately 42,600 acres of fresh market tomatoes were harvested (Florida Tomato Committee, 2005). The value of the tomato crop during 2002, which exceeded more than 466 million dollars, accounted for 27% of all U.S. cash receipts for fresh market tomatoes (Florida Agriculture Statistical Services, 2005). Bacterial wilt caused by Ralstonia solanacearum (Smith) Yabuuchi et al., 1995, is one of the most destructive bacterial diseases in the tropical, sub-tropical, and temperate regions of the world, including the southern U.S. and Florida. It is causing serious yield losses on many crops such as tomato, potato, pepper, eggplant, tobacco, banana and geranium (Pelargonium) (Hayward, 1991). It is estimated that the bacterium is pathogenic on several hundred plant species in over 50 families throughout the world (Hayward, 1994). On tomato and other plants, R. solanacearum colonizes and blocks the vascular tissue resulting in wilting and eventual death. Thus the disease has been given the name bacterial wilt. Other common names of the disease have been used based on the location and plant host, such names as Granville Wilt on tobacco, southern wilt on geranium, brown rot on potato, and Moko disease on banana. Brown rot of potato caused by R. 1

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2 solanacearum has been estimated to affect about 3.75 million acres in 80 countries (except the U.S. and Canada) with global economic loss estimates currently more than $950 million per year (DEFRA, 2003). R. solanacearum is a Gram-negative rod, 0.5-1.5 m in length, with a single polar flagellum. The positive staining for poly--hydroxybutyrate granules with Sudan Black B or Nile Blue distinguishes R. solanacearum from Erwinia species. In addition, the staining appears heavily at the pole ends with carbol fuchsin. Agar colonies are initially smooth, shining and opalescent, but become brown with age (Lelliott and Stead, 1987). The bacterium gains entrance into the plant by way of injured roots and stem wounds or through stomata. Once inside the plant, the bacteria move in the vascular tissues. This process can be accelerated by high temperature. The speed of the bacteriums movement is dependent on the plant part that is colonized (Ono et al., 1984). The bacteria then colonize the primary xylem and secondary xylem tissues. The pathogen often destroys the pit membranes, and all parenchyma cells adjacent to vessels infested with bacteria are necrotic and can be colonized by the bacteria (Nakaho et al., 2000). The organism adheres by polar attraction to the cell surfaces and subsequently becomes localized at preferential sites of the mesophyll (Petrolini et al., 1986). Bacteria blocking the plants vessels, by production of exopolysaccharide, are the major cause of wilting. The disease is most severe at 24-35C. R. solanacearum (Race 1 and 2) is rarely detected in locations where the mean temperature for any winter month falls below 10C. Thus, there exists a distinct temperature requirement for optimum disease development and multiplication of the cells (Caruso et al., 2005). High soil moisture and periods of

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3 wet weather or rainy seasons are associated with elevated disease severity. Soil moisture is also one of the major factors affecting reproduction and survival of the pathogen (Hayward, 1991). Symptoms characteristic of bacterial wilt on most hosts are wilting, stunting and yellowing of the foliage. Symptoms can develop at almost any plant stage. However, the complete wilting and collapse occur mostly when young and susceptible plants become infected. The rate of disease development is heavily influenced by environmental factors. If environmental conditions are favorable, a young tomato plant will develop advanced symptoms of wilt within a few days (Aberdeen, 1945). Wilted leaves remain on the plant and maintain their green appearance for many days. In some cases, although true symptoms may not appear, stunting and dwarfing of the plant might develop as the plant matures. The vascular tissues of the stem show a brown discoloration and, if the stem is cut crosswise, drops of white or yellowish bacterial ooze may be visible in clear water (McCarter, 1991). R. solanacearum differs from other bacteria in that different strains can have a different host range, different locations of origination throughout the world, and different utilization of carbon sources. Due to this fact, classification of the bacterium is complex and based on the differing points. Previously in attempts to classify the bacterium, it has been placed in categories of groups, races, biovars, biotypes, sub-races and strains. R. solanacearum has been defined as a species complex by Prior and Fegan (2005). They suggested a new hierarchical classification scheme with these taxonomic levels: species, phylotype, sequevar, and clone.

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4 Buddenhagen et al. (1962) distinguished three races on the basis of pathogenicity. Race 1, affects tobacco, tomato, potato, eggplant, diploid bananas, and many other crops and weeds, with high a growth temperature optimum (35-37C). Race 2, affects triploid bananas (causing Moko disease) and Heliconia spp., with a high temperature optimum (35-37C). Race 3, affects mainly potato, tomato, and geranium, without a high virulence on other solanaceous crops, and with a lower temperature optimum (27C). Other hosts are weeds such as Solanum dulcamara, S. nigrum and S. ciereum and the composite weed Melampodium perfoliatum. Races 4 and 5 affect ginger and mulberry, respectively (Elphinstone, 2005). Hayward (1964) distinguished four biovars by their ability to produce acid from several disaccharides and sugar alcohols. These biovars however do not correlate with the races that Buddenhagen et al. (1962) had described. Only race 3, the cool temperature adapted race, is equivalent to biovar 2 (Hayward, 1983). Races and biovars have been classified into two main groups according to a restriction fragment length polymorphism (RFLP) analysis (Cook and Sequeira, 1988; 1994). Group 1 consists of the Asian stains of race 1, biovars 3, 4, and 5. Group 2 includes the American strains of race 1 biovar 1 (r1/b1) (most common strains in Florida), race 2 biovar 1, and race 3 biovar 2 (r3/b2). Since 1995, R. solanacearum r3/b2 has entered several states of the U.S. on various occasions in greenhouse grown geranium cuttings. The pathogen was traced back to infected geranium cuttings that were imported from Guatemala and Kenya (Williamson et al., 2002; Kim et al., 2003). R3/b2 of R. solanacearum is a listed select agent in the U.S. under the Agricultural Bioterrorism Protection Act of 2002 (Federal Register, December 13, 2002, Part V, Department of Agriculture, APHIS, 7 CFR Part

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5 331, 9 CFR Part 121) because it has the potential to be a severe threat to the potato industry and other crops. R. solanacearum is mainly disseminated in latently infected plant material as well as through surface water and soil. It survives in surface water and soil usually by establishing itself in host plants. In the U.S. and Florida, dispersal via surface irrigation water is not well understood and preliminary findings were reported recently (Hong et al., 2004). The sources of infections of R. solanacearum are contaminated soil, in plant materials, on farm equipment, and in irrigation or surface water. The bacterium survives in the soil for varying periods of time, and is able to persist between successive crops. The bacterium has been found to survive in sheltered sites such as plant debris and latently infected potato tubers, the deeper soil layers and in the rhizosphere of roots of weed hosts (Hayward, 1991). The range of weed hosts is extensive, but the significance of weed hosts depends on the environment and cropping systems. Some hosts are symptomless carriers. The bacterium has been isolated from irrigation water and proliferates when alternative aquatic weeds are present (EPPO, 2004). Irrigation water infested with R. solanacearum has caused several outbreaks on a number of crops (Elphinstone, 1998). Olsson in 1976 discovered a relationship between farms with infected potato tubers and the weed Solanum dulcamara, a common species of nightshade found in Sweden (Olsson, 1976). During the winter months when the temperatures were low R. solanacearum was undetectable in the irrigation water, but the bacterium could be detected in the xylem of the adventitious roots of the semi-aquatic weed growing in the

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6 water (Persson 1998). In a well documented case in England, potato brown rot occurrence was associated with effluent from the potato processing industry and municipal water purification plants that handled diseased potatoes. The bacterium was also found colonizing in the presence of S. dulcamara along the river banks (Buitrago, 2001 and EPPO, 2000). R. solanacearum is a serious obstacle to the culture of many solanaceous plants in both tropical and temperate regions. The greatest economic damage has been reported on potatoes, tobacco and tomatoes in the southeastern United States, Indonesia, Brazil, Colombia and South Africa. From 1966 to 1968 in the Philippines there were average losses of 15% in tomatoes, 10% in eggplant and Capsicum, and 2-5% in tobacco (Zehr, 1969). In Peru along the Amazon basin, about half of the banana plantations were affected and rapid spread of the pathogen threatened to destroy plantations throughout the Peruvian jungle (French and Sequeira, 1968). In India, occasionally total losses occur in tomato crops. Outbreaks in potato, which caused extensive losses, occurred in Israel (Volcani and Palti, 1960), and Greece (Zachos, 1957). Control Measures have been largely unsuccessful due to the nature of the organism, especially for race 1 with its broad host range, and race 3 with its ability to cause latent infection on potato tubers. Commercially available chemical control is not known. It has been noted previously that soil fumigants showed little or no effects (Murakoshi and Takahashi, 1984). However, it has been reported that broad spectrum soil fumigants (chloropicrin) will delay the initial disease onset (Ooshiro, 2004). Antibiotics such as streptomycin, ampicillin, tetracycline and penicillin were basically ineffective (Farag et al., 1982). In fact, Farag et al., reported an increase of disease when streptomycin was

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7 applied to potato crops in Egypt. Biological control has been investigated (Farag et al., 1986; Trigalet et al., 1994); yet more work needs to come forth. Resistant cultivars have been developed in various crops (Hayward, 1991). In crops such as tobacco and peanut, resistant cultivars have been very successful. Many resistant cultivars of tomato plants have been developed and are also successful in a particular environment. Yet, it has been difficult to develop cultivars which are resistant under conditions of high temperature and humidity in the lowland tropics (Hayward, 1991). Alternative disease management tactics were investigated with promising results, especially using Thymol (Pradhanag et al., 2003; Ji et al., 2005) as a biofumigant and acibenzolar-S-methyl as a plant activator on moderately resistant cultivars (Anith et al., 2004). Due to limited efficacy of current integrated management strategies, bacterial wilt continues to be economically important for many economically important crops in Florida, and many subtropical, tropical, and temperate areas of the world. Cultural practices, crop rotation and host resistance may provide limited control (Pradhanang el al, 2003). Tomato growers need to use an integrated approach to lower the impact of bacterial wilt on tomato production. These are current recommendations (Momol, 2005, unpublished): Preplant Choose resistant or moderately resistant cultivars, or graft susceptible cultivar onto resistant rootstock. Consider a preplant soil amendment or fumigation for infested fields against R. solanacearum and nematodes [i.e. Thymol (not commercially available) and Telone mixture]. Apply 2-3 years of rotation and cover crops for infested fields to reduce R. solanacearum, weeds and nematodes.

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8 Do not irrigate cover and rotation crops with R. solanacearum contaminated pond or surface water, avoid reinfestation. Use well drained and leveled fields, and do not use low-lying areas of the field. Raise soil pH to 7.5-7.6 and increase available calcium (liming). Consider using infested fields during cooler months for tomato production (i.e., spring season for north Florida). Production Exclude the pathogen by applying strict sanitation practices (pathogen free irrigation water, transplants, stakes, machinery, etc.). Chlorinate your irrigation water continuously if you are using surface water or R. solanacearum infested pond water. Irrigate based on water need, avoid over-irrigation. Apply Actigard (Syngenta) if you are using moderately resistant cultivars (i.e., FL 7514). After harvest Plow under crop residue immediately. Start with suitable rotation and cover crops (i.e., rye for winter, sudan-sorghum for summer in north Florida) to avoid weeds that support R. solanacearum populations. Potato growers are advised to avoid fields infested with the pathogen. Growers should not plant transplants in close proximity to fields where this problem has occurred. Infested fields should be rotated with a non-susceptible crop, because long term rotation might reduce the population of the bacterium. Growers should avoid moving equipment and soil from infested to non-infested fields and using irrigation water from ponds in close proximity to infested fields. Caution is given to excessive irrigation, because high soil moisture will induce high incidence of disease or buildup of population (Janse, 1996). In 2002 August, R. solanacearum (r1/b1) was detected in the irrigation pond water of tomato farms and ornamental nurseries in north Florida (Momol et al., unpublished).

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9 Most ponds in north Florida are interconnected by both aboveground and underground streams. In 2002, research was initiated to sample the irrigation pond water for the presence of R. solanacearum. In addition to water sampling, various aquatic and other weeds were sampled to determine their relationship with the bacteria. The long term goal of this study is to improve our understanding of R. solanacearum in relation to irrigation water and weed hosts (including aquatic weeds) in Florida. The objectives of this research were to study the populations of R. solanacearum within irrigation ponds by analyses of water and aquatic weeds and other weed samples, and to determine a method to reduce the bacterial populations before irrigating tomato fields in Florida.

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CHAPTER 2 LITERATURE REVIEW Bacterial wilt incited by R. solanacaerum, is a major bacterial disease on many crops in tropical, sub-tropical, and temperate regions of the world. Molecular aspects, including pathogen genetics, pathogenesis, host-plant interactions, molecular detection, and strain differentiation, are more studied than its epidemiology and ecology. Pathogen survival studies are very important in terms of understanding sources of inoculum and its long and short distance dispersal. Advances in semi-selective medium (SMSA, Engelbrecht 1994) and sensitive and specific molecular methods for R. solanacearum detection, improved data reliability for epidemiological and ecological studies. These are three major ways that R. solanacearum can survive: surface and irrigation water, plant material, and soil. R. solanacearum r3/b2 that causes brown rot on potatoes has been present in Europe since 1976 (Olsson, 1976). The description of race 3 survival in river water used for irrigation and its association with Solanum dulcamara (Elphinstone et al., 1998) was an important cornerstone in the understanding of its epidemiology and ecology. Detection of Ralstonia solanacearum in Irrigation Water and Solanum dulcamara Contaminated irrigation water has been the source of many outbreaks of bacterial wilt on a number of crops (Elphinstone et al., 1998). R. solanacearum can survive for long periods of time in water (Kelman, 1953) which has important implications for agricultural practices. 10

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11 The first report of R. solanacearum in Europe was made in Sweden. Over 30 years ago the bacterium was thought to be a warm-temperature pathogen. However, brown vascular rings were found in market potato tubers, cv. Bintje, during routine inspection in Sweden in 1972. The symptoms were first thought to be caused by Clavibacter michiganenis subspecies sepedonicus. Yet, after isolation of the Gram-negative, Sudan-positive bacterium which caused rapid wilting in tomato plants, a presumptive diagnosis indicated that the pathogen was R. solanacearum. Strains isolated from the tubers had optimum temperatures between 25 and 30C, which was lower than what was generally believed to be typical of R. solanacearum. The isolated strains caused severe wilting after 5 days in young tomato plants cv. Dansk Export and eggplant cv. Black Beauty (Olsson, 1973, 1977). Strains were typed as biovar 2 (Olsson, 1976). It was unexpected to find this pathogen as far north as the southern part of Sweden. Visual inspection of 400,000 tubers collected in 1973 from different fields and seed-potato lots, showed that brown rot symptoms were only present in tubers grown in the same field. Infected tubers were found in low frequency. The following year, in the same potato farm, but a different field brown rot was observed. The same year, a second brown rot outbreak occurred 80 km north of the first infection site in the Pinnn stream. On this farm, tubers from several fields were infected at a frequency of 5% in cv. Bintje and 2% in cv. Grata. A research study commenced to find the source of infection and a few facts were known. Seed potatoes from the same or from related seedlots, which had been used in other fields, produced healthy tubers. Large populations of the Colorado beetle (Lepinotarsa decemlineata) had invaded southern parts of Sweden in 1972, coming from

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12 the eastern parts of Europe. Beetles were present in the fields of the first outbreak, but not in the second. Thus, these two possible sources of the primary infection were ruled out. One important common fact for the two separate brown rot epidemics was that all infected potato tubers had been grown in fields irrigated with surface water from a nearby stream. Irrigation practices were beginning to be implemented in Sweden in the early 1970s (Hayward et al, 1997). Greenhouse experiments showed that tomato, eggplant, and wild Solanum species of the local flora, were found to be susceptible to the Swedish strains of R. solanacearum. These plants were irrigated with water from the streams used by fields of the infected potatoes. Tomato, eggplant and wild Solanum speices were planted in the fields where infected tubers had been growing. S. dulcamara was planted near the streams at the two separate outbreaks. None of the test plants became infected (Olsson, 1976). S. dulcamara was found growing near and in the Pinnn stream. In 1974 R. solanacearum was isolated from some of these plants. Isolation from the various parts of the plant showed a few cells were detected in the stem at the bottom of the plant. However, high numbers were found in the xylem of the thin adventitious roots growing in the water (Olsson, 1977, 1979). The pathogen was isolated the following year from the aquatic weed, which indicated that the bacterium was able to survive overwinter in S. dulcamara. The Pinnn stream was surveyed for infected S. dulcamara. The bacterium was isolated from plants growing 3 km downstream from where a potato processing plant had been until 1967. The processing plant had processed both domestic and imported

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13 potatoes, and the waste water was dumped directly into the stream. The stream led to two dams, in which a large population of infected S. dulcamara grew. Infected potato tubers had been produced in fields irrigated with water from these dams and downstream from the processing plant. In 1977 and 1978, water samples from streams close to infected fields were analyzed. Samples were filtered through a 0.45 m membrane to concentrate the bacteria. The different membranes were washed with sterile water. The suspensions were inoculated into 3 week old tomato plants. Isolations made from wilting plants resulted in R. solanacearum being detected from water collected at both locations (Olsson, 1979). High populations of the bacterium were found when the water was sampled near the bottom of the Pinnn dam, where S. dulcamara grew in abundance (Persson, 1998). Brown rot occurred in Sweden at two separate locations from 1972 through 1977. Infected crops were destroyed by industrial processing, in which tubers were destroyed by heat treatment at 145C. R. solanacearum was undetectable in samples from the remaining potato particles. Fields where the crop was contaminated were restricted from growing potatoes for the following 2 years. After the 2 years, were allowed to be grown again on infested farms potatoes, but only by using certified seed potatoes. It was recommended not to use the infected stream water for irrigation purposes. Analyses of water from streams near potato processing plants in southern and western parts of Sweden in 1979 and 1980 did not detect the bacterium. Since the reports from Sweden, other outbreaks of R. solanacearum have been reported in other European countries including Belgium, France, Italy, the Netherlands,

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14 Portugal, Spain, and the UK (Stead et al., 1996). For many cases these strains are not similar, and the source of infection is unknown (Janse et al, 1997). The pathogen is thought to have been spread in latently infected seed potatoes in the Netherlands in 1995, thus causing epidemic outbreaks on more than 90 seed and market potato farms (Elphinstone et al., 1997). The Netherlands reported brown rot of potato for the first time in 1992. The origin of the pathogen is assumed to come from uncertified seed. In the same year outbreaks in the UK and in Belgium were reported, yet no colonial link could be established (Janse et al, 1997). In 1995 a series of findings of brown rot in both seed and market tomatoes could be traced back to one grower of a local variety that was heavily infected. From this source, paths could be traced other infected lots. On the same infected lots where the local variety of potato could be found, a few other varieties were discovered to be infected, usually in latent form. Investigations of surface water near these farms showed the presence of the brown rot pathogen. Further studies indicated a link between water usage and diseased potatoes. An extensive survey of the pathogen in surface water and S. dulcamara was performed in 1996. About 14,000 samples were taken throughout the entire country. Five percent of the water samples were positive for R. solanacearum, which resulted in a ban for irrigating with these streams. The bacterium, similarly, was detected in S. dulcamara in which the roots were submerged in the water. In many cases the pathogen was detected in water in close proximity to the potato industrys purification plants. This brought about the possibility that the bacterium could escape the purification process.

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15 Moreover these findings suggest that probably all infections in the Netherlands originated from irrigation with contaminated surface water (Janse et al, 1997). Water samples from northern the Netherlands were taken in 1996 to determine pathogen population during different seasons. Samples were taken weekly in four rivers (i.e. one non-contaminated and three contaminated with the bacterium). The bacterium was never found in the non-infested river. In the heavily infested area the bacterium could be detected until ice formed. Directly after the ice melted the bacterium could be detected again, but at very low numbers (Janse et al, 1997). Outbreaks of brown rot lead to the development of detection methods of R. solanacearum in S. dulcamara and in irrigation water. In England, in 1992 and 1995, two outbreaks of brown rot occurred at different farms. The source of the infection in both outbreaks was contaminated irrigation water, in which, infected S. dulcamara plants were naturally growing (Stead et al., 1996). A detection process was established to better understand the nature of the pathogen. Four hundred twenty S. dulcamara plants were sampled from the various infected waterways. R. solanacearum was extracted from the stolon section and fibrous adventitious roots. The pathogen was detected by three methods; 1. indirect enzyme-linked immunosorbent assays performed on boiled plant samples; 2. polymerase chain reaction performed on sub-samples of the extract of the boiled samples; and 3. streaking concentrated extract onto modified semi-selective SMSA. Results varied from each test; therefore the most reliable detection was to employ the three methods simultaneously (Elphinstone et al., 1997). Over a four year period, S. dulcamara remained infected from year to year, and the viable pathogen could be isolated throughout the winter months.

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16 Only plants with roots submerged in water were found to be infected. Other aquatic weeds growing in contaminated streams were not infected. When volunteer potato plants were sampled, less than 1% infection of progeny tubers was detected and no infection was found in volunteers the following year. Thus it was concluded that the highest risk of dissemination of the pathogen was through establishment of the bacterium in aquatic S. dulcamara and the release into the waterways used for irrigation (Elphinstone et al., 1997). Upon a national survey conducted in the rivers throughout England and Wales, infected S. dulcamara plants were found in the Thames River, five of its tributaries, and the Witham River. Samples from the Thames River indicated several possible sources of entry of the pathogen into the water system. Included in the possible sources were outflows from sewage treatment plants where S. dulcamara was present. The potential for spread of the pathogen from domestic or industrial use of imported potatoes was a possibility and needed further investigation (Elphinstone et al., 1997). Water samples from the river water were analyzed to monitor R. solanacearum from a single sampling point over three years. The bacterium was only detected in the water where S. dulcamara was present. When necessary, samples were centrifuged to concentrate the pathogen population. Enriched samples were detected by PCR. The pathogen was only isolated during warm summer and fall months. Population levels were reduced when the temperature fell, water levels rose, and/or S. dulcamara died back (Elphinstone et al., 1997). In Egypt, the bacterium was found in both irrigation and drainage canals near infected potato fields (Farag, 1998). Contaminated irrigation water was reported as a

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17 source of infection in Kenya (Janse, 2005). Due to the bacterium having many diversified hosts, one crop contaminating irrigation water could essentially affect neighboring farms that grew other solanaceous crops. As the epidemiology and ecology of the pathogen are being studied, the more complex this organism is becoming. Studying the dynamics of the bacterial population in the water, a distinct relationship existed between the temperature of the water and the size of the population (Countinho, 2005). The aquatic weed Solanum dulcamara has been identified as a symptomless host (Olsson, 1977). Bacterial populations have been shown to overwinter in this weed, while the bacterium was undetectable in water (Perrson, 1998). In a Spanish river over a three year period there was a strong relationship between water temperature and detection of R. solanacearum. Collection of water began in the Tormes River after a confirmed outbreak of potato brown rot in the late fall of 1999. Following analysis of the irrigation water, it was determined that the water was contaminated with R. solanacearum. Samples were collected from five different locations along the river water and directly spread plated on SMSA. Presumptive colonies were confirmed using indirect immunofluorescence and double-antibody-sandwich indirect ELISA. Of a total of 65 samples analyzed over three years, R. solanacearum was isolated from 22 of them. The temperature range of the surface water that contained the bacterium varied from 9 to 20C. Most of the samples taken had an average temperature above 14C, with detection of the pathogen usually being unsuccessful in lower temperatures. Populations in the water ranged from 10 to 80 cfu/ml during the warmer months. However, by November the bacterium was non

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18 detectable. There was a significant positive correlation between the R. solanacearum populations and water temperatures for sampling sites (Caruso et al., 2005). Methods for Controlling Ralstonia solanacearum To avoid further dissemination of the pathogen, control measures must be designed and executed. For tomato production in Florida, latent infection is not a major concern, except seedlings have to be pathogen free. Current management recommendations were listed previously (see Introduction) for tomato producers. The Netherlands has developed guidelines to ensure quality of potatoes and to control the pathogen that causes brown rot (Janse et. al., 1997). Many other countries have followed these procedures or have made modifications to fit their needs. Testing seed for infections and possible latent infections of the pathogen. Survey testing of seed, market, and starch potatoes Prohibition of the use of contaminated surface water for irrigation and discouragement of its use in other areas. Eradication of S. dulcamara from infected waterways. Survey of contaminated production places where possible infected lots have been. Sample and analysis of imported potatoes for latent brown rot. Survey surface water areas surrounding contaminated surface water and purification plants. When an infected lot is found, infected potatoes will be destroyed, machinery disinfested, and all other potatoes and seeds on the farm are to be tested. On infected fields for five years no potato or tomato are to be grown, control of volunteers, in the first three years grass or cereals are grown, then in year six potatoes for seed or market from certified seed can be grown. Testing of all potatoes produced on the farm during this period and the following year. Initiation of national research program to facilitate detection of the pathogen, including serum production, evaluation of sure and rapid detection methods, and to study epidemiology in soil, water, and aquatic weeds.

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19 The Swedish Plant Protection Service carried out the first eradication program for S. dulcamara in the Pinnn stream. Both mechanical and chemical control measures were used between 1977 and 1981. The herbicide glyphosate, (trade name Roundup, Monsanto) was used for chemical treatments. S. dulcamara plants were treated manually with glyphosate and also removed mechanically at specific sites along a 30 km distance both up and downstream from the former potato processing plant. The herbicide was applied for 2 years at the two dams, where a large population of the weed existed (Persson, 1998). Following the eradication program in 1993 water samples were again collected from the two Pinn dams. R. solanacearum was not detected. However, soft rot bacteria Erwinia carotovora subsps carotovora and E. chrysanthemi were detected. E. c. caratovora is a well known pathogen found in surface water (Persson, 1998). Previously in the early 1970s, before the reports of brown rot, reports of wilted tomato plants in the market were made. The plants were irrigated using the same source. The production of the tomatoes had finished before a lab diagnosis could be made. It was speculated that R. solanacearum was probably the pathogen, yet later results allowed for the possibility that E. c. caratovora or E. chrysanthemi might have been the causal agent (Persson, 1998). In 1995 a visual inspection of the Pinn area showed that S. dulcamara was very rare. The eradication which started 17 years earlier has been successful. Ten plants were analyzed for possible infection; however, the bacterium was not isolated. In 1996 a survey of streams in the former infested areas was initiated. Water samples from irrigation waterways and wastewater from the potato industry were collected and plated on modified SMSA. Water samples were inoculated in tomato plants, and analyzed using

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20 polyclonal antiserum and ELISA. None of the tests detected the presence of R. solanacearum in the water. Tests were also carried out on seed potatoes. Analysis of the potatoes consisted of using a polyclonal antiserum. The pathogen was not detected in the seed potatoes (Persson, 1998). Based on these results it was assumed that R. solanacearum had been eradicated from Sweden. However, as it has been shown before the pathogen can be established in Swedens environment. Therefore, careful consideration must be taken for importing potatoes into the country. Although control of the bacterium has been possible in Sweden, conditions are not the same for the rest of Europe. S. dulcamara has proven to be a factor in the epidemiology of the pathogen, but other weed hosts need to be investigated. The sources of some waterways are areas of swampy conditions, thus complete eradication by mechanical and chemical means of S. dulcamara plants would be impossible (Hayward et al, 1997). Brown rot is now a major concern in Europe, and control methods are being implemented to hinder the dispersal of this pathogen. More information is required to fully understand the epidemiology of the pathogen in cool climates, including studies on the ecology of the pathogen and identification of survival sites. Studies on the disease have shown unique features of the epidemiology of the pathogen such as; its survival and the increase of populations in semi-aquatic S. dulcamara weeds, its distribution in rivers and waterways, and passage of the pathogen through industrial and domestic waste water. More information is needed on latently infected weed hosts and the pathogens method of

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21 entry. There exists a need for standardization of test methods so that equivalence is achieved across Europe and world-wide (Hayward et al, 1997).

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CHAPTER 3 MATERIALS AND METHOD Bacterial culture and inoculum preparation. R. solanacearum (race 1, biovar 1), tomato strain Rs5 (Pradhanang, 2001) and field strain SEF both isolated in Quincy, Florida (Gadsden County) were used in these studies. The bacterial pathogen was grown at 28 C either on modified semi selective medium SMSA (Engelbrecht 1994) or on casamino acid peptone glucose (CPG) agar (peptone 10 g, casamino acids 1 g, glycerol 2.5 ml, agar 15 g, deionized water 1 liter) for 60 h or in CPG broth on a shaker (100 RPM) for 60 h. Bacterial cells were suspended in sterile deionized water and the concentration of inoculum was estimated by measuring absorbance at 600nm. The viable bacterial population was determined following dilution plating on modified SMSA. Isolation of R. solanacearum from irrigation pond water. Water samples were collected at five different irrigation ponds used for tomato and ornamental plant production located in Gadsden County, Florida. The average air temperature for each month was collected by the weather station at Quincy, FL. Pond 1 was a retention pond heavily vegetated with various aquatic weeds, located at an ornamental nursery. In this pond, runoff water from the greenhouses was collected. Five samples were collected from the banks of this pond from August 2003 until August 2004. Pond 2 was located in the middle of a tomato growers field. Four samples were collected on the south end of the pond. Pond 2 had various kinds of aquatic weeds growing along the banks. Samples were taken from August 2003 to May 2005. Pond 3 was located on a different tomato farm and was the smallest of all the ponds. Pond 3 was dry until April 2004 and then 22

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23 samples commenced until May 2005. Weeds that grew in this pond were not similar to those found in ponds 1 and 2 because pond 3 was a field during the dry season. Pond 4 was located on the same farm as pond 3. Pond 4 was the largest pond of all the ponds, and was fed by local streams. This pond was surrounded by hills on all sides except on the west bank and was heavily shaded by the trees that grew on the hills. Four samples were collected from August 2003 through May 2005. Pond 5 was added in May 2004 and samples continued until May 2005. This pond was surrounded by hills on all sides and was located on the south side of a tomato farm. Two samples were collected on the west bank of this pond. Duckweed completely covered the surface of the pond all year around and the surrounding trees shaded the pond. Tomatoes infected with bacterial wilt were found on tomatoes on the northwest corner of the farm. Surface water samples were collected in the same manner for each pond. Samples were collected in close proximity to semi-aquatic weeds, namely Hydrocotyle ranunculoides and Polygonum pennsylvanicum because it was suspected that these weeds were symptomless hosts from a previous preliminary survey. Water samples were collected in a 7.6 liter metal bucket and transferred to sterile 50 ml Falcon Tubes. Non-diluted and ten-fold dilutions of each sample were spread plated on modified selective medium SMSA. R. solanacearum colonies are irregular shaped with a glossy wet appearance. The colonies appear white with a purple center in the beginning stages of growth, then marbling or swirls of purple and white colors appear at 60 h. The plates were incubated at 28 C for 60 h. Suspected R. solanacearum colonies were selected and transferred to SMSA. Presumptive colonies were confirmed by whole cell fatty acid methyl ester analysis (MIDI, 2001).

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24 Pathogenicity of the bacteria was determined by inoculating 10 to 14-day old Bonny Best tomato plants. Initially, root inoculations were performed by stabbing around the base of the plants to damage the roots. Following wounding, 10 ml of a bacterial suspension consisting of 10 8 cfu/ml were poured along the crown of tomato plants potted in 10-cm pots. The leaves would curl and droop within 4 to 5 days after inoculation. The inoculated plants were transferred to a growth room on 12 h light dark cycle with air temperature at 28 C. Because of inconsistent results, this method was replaced by the toothpick inoculation method. Upon using the toothpick method, symptoms developed quicker. Sterile toothpicks with the tips laden with bacteria were stabbed into the lower stems near the crown of the tomato plants. Each isolated colony was used to inoculate one plant. Plants stabbed with sterile toothpicks or toothpicks coated with Rs5 served as the negative and positive controls, respectively. Bacteria were re-isolated from the tomato plants that developed wilt symptoms and were stored in sterile 30% glycerol at -80 o C. During the winter months when the bacterium was undetectable, the collected water samples were concentrated by centrifugation. Thirty ml of the water samples were centrifuged at 10,000 rpm for 10 min at 28C. All but 100 l of the supernatant was discarded. The pellet was re-suspended in the remaining supernatant. Non-diluted and ten-fold dilutions of each sample were spread plated on modified selective medium SMSA. The plates were incubated at 28 C for 60 h. Suspected R. solanacearum colonies were selected, transferred to SMSA and characterized using the procedure previously outlined.

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25 For statistical analysis the data acquired from each sample were transformed using log transformation [z=log 10 (y+1)], and then the data were statistically analyzed using Speramans correlation coefficient by SAS for Windows program release 8.02 (Cary, NC). Monitoring of R. solanacearum in semi-aquatic and other weeds associated with irrigation ponds. Common aquatic weeds that were growing along the bank or submerged in the ponds were collected. Plants were identified by Karen Brown from the Center for Aquatic and Invasive Plants, University of Florida in Gainesville FL. Plant samples were analyzed for the presence of R. solanacearum. The plants were first washed under tap water to remove loose particles of sand and dirt. The roots were separated into three sections to sample the entire plant part; root hairs to the root tips, midsection, and from the midsection to where the root attaches to the stem. Stems were cut in three sections; the stolon, the crown, and before the first leaf. The root sections were combined and the stem section were combined and given the labels root and stem. The cuttings were separated into two groups and were either surface sterilized or untreated. Sterilizing consisted of soaking the cuttings in 70% ethyl alcohol for 5 min. Then the cuttings were rinsed in sterile water for 5 min. Each cutting was placed in separate clear 2 mil zipper plastic bags and mashed using a pestle and 1000 l of sterile water was added to the crushed plant residue. The plant residue was soaked for 20 min. The suspension was extracted using a pipette and stored in microfuge tubes. Each sample and a ten-fold dilution were spread plated onto SMSA. The plates were incubated for 60 h at 28 C. Colonies suspected of being R. solanacearum were selected and analyzed by

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26 whole cell fatty acid methyl esters analysis. Confirmed R. solanacearum colonies were tested for pathogenicity by the toothpick method and re-isolated as described previously. Bacteria were reisolated from tomato plants that developed wilt symptoms, and were stored in a sterile 30% glycerol solution at -80 o C. During the winter months when the bacterium was undetectable the collected plant samples were concentrated by centrifugation. Fifteen-hundred microliters of the macerated plant residue were centrifuged at 10,000 rpm for 10 min. at 28C. All of the supernatant was discarded. The pellet was re-suspended in 400 l of sterile water. Non-diluted and ten-fold dilutions of each sample were spread plated on modified SMSA. The plates were incubated at 28 C for 60 h. Suspected R. solanacearum colonies were selected and transferred to SMSA, and identified by whole cell fatty acid methyl esters analysis. Confirmed R. solanacearum colonies were tested for pathogenicity by the toothpick method and re-isolated and stored as described previously. Infested water treatment with chemicals. The minimal inhibitory concentration (MIC) for chlorine and hydrogen peroxide was determined for a known population of R. solanacearum. Ultra Clorox Bleach (The Clorox Co., Oakland, CA) was used and contains 6% active ingredient and 5.71% of free chlorine Cl 2 Hydrogen Peroxide (Diamond Products, Seffner, FL) was used and contained 3% active ingredient, but after the first trials Hydrogen Peroxide 30% (Fisher Scientific Co. LLC, Pittsburgh, PA). Laboratory strain Rs5 was grown in CPG medium as previously stated. The populations were adjusted to A 600 = 0.3 by using a spectrophotometer at 600 nm to achieve a bacterial concentration of approximately 10 8 cfu/ml. One hundred microliters of 10 8 cfu/ml in a sterile deionized water solution were added to a dilution series ranging from 2 mg/l to 10

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27 mg/l of the given chemical. Dilutions were made from a stock solution of the chemical diluted with sterile deionized water. Bacterial cells were exposed to the chemical for 1 min before being spread plated onto SMSA. Each dilution with the bacteria cell suspension was plated on two petri plates, and the experiment was repeated three times. The bacterial suspension at 10 8 cfu/ml was spread plated on SMSA to compare the effects the chemicals on R. solanacearum survival. A percentage of remaining cells was calculated by dividing the average colonies that survived after the chemical treatments against the untreated control. The protocol was modified to reduce error and to mimic what is found in the environment. Previously the stock solutions of the chemicals were made by adding minute amounts of the chemical, 166.3 l of chlorine and 3.33 l of hydrogen peroxide and were diluted in sterile tap water achieve a stock solution of 10 mg/l. Because small amounts of the chemicals were used, pipette error was possible. Thus, the chemicals were diluted with water at 1:10 for chlorine and 1:100 for H 2 O 2 before making the 10 mg/l stock solution. Thus the chemical could be measured more precisely. The bacterial suspension was diluted from 10 8 cfu/ml to 10 4 cfu/ml to simulate what may occur in the ponds. For both methods untreated bacterial suspensions of either controls 10 8 cfu/ml to 10 4 cfu/ml were spread plated. The control plates were compared to those of the dilution series to determine the percentage of recovered cells. Sensitivity of modified SMSA for recovery of R. solanacearum. Laboratory strain Rs5 was grown in CPG broth, and the suspension was adjusted to 10 8 cfu/ml following previous procedures. A dilution series of the bacterium was made ranging from 10 1 to 10 8 cfu/ml. The dilutions were spread plated on SMSA and on nutrient agar

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28 (NA) (0.8% nutrient broth [BBL, Becton Dickinson and Co., Cockeysville, MD] and 1.5% Bacto Agar [Difco, Becton Dickinson and Co., Sparks, MD]). The plates were incubated at 28 C for 60 h. Each dilution was spread plated on each of three plates, and the experiment was repeated three times. The bacterial colonies from each plate were counted and each dilution was averaged and the standard deviations were calculated. Colonization R. solanacearum on Hydrocotyle ranunculoides Two strains of R. solanacearum, laboratory strain Rs5 and SEF were grown in CPG broth at 28 C. The bacterial cells were adjusted to 10 8 cfu/ml according to procedures previously stated. The two strains were also grown on SMSA incubated at 28 C for 60 h to achieve colony growth. Bacteria grown on the plates were used for the toothpick method. Hydrocotyle ranunculoides plants were collected from pond 2 and transplanted in 10-cm pots in Terra-Lite agricultural mix (Scott Sierra Horticultural Products Co., Marysville, OH). Plants were grown in greenhouse conditions for 3 months. Once a week the plants were divided and propagated. Each week samples from the propagated plants were taken to detect the presence of R. solanacearum. Plants used for the experiment were a week old after propagation and then were inoculated strain Rs5 or SEF. The plants were inoculated by stabbing the crown of the stems with toothpicks and also by drenching the root zone with bacterial suspension. Sterile toothpicks were coated with the bacteria using either the lab strain Rs5 or SEF. Ten plants for each bacterial strain were inoculated using the toothpick method as described previously. Control plants were stabbed with sterile toothpicks in the same manner. Twenty additional plants were inoculated by the drench method. These plants were removed from their pots and the

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29 root system was broken up by hand before inoculation. A bacterial suspension consisting of 50 ml of 10 8 cfu/ml was poured into each pot. Ten plants were inoculated with strain Rs5 and another ten plants were inoculated with SEF. Every plant remained in its individual plastic saucer. After inoculation the plants were stored in greenhouse conditions. The saucer collected the water that was not taken up by the plant. The plants were watered daily by pouring the collected water from each saucer back into the corresponding plant; normal watering procedures were then followed. The roots and stems of the plants were analyzed for colonization of bacteria following the procedures outlined previously. A dilution series was spread plated on modified SMSA. This was replicated three times. The plates were incubated at 28 C for 60 h and then the number of colonies was counted. Two colonies of the recovered R. solanacearum from both of the different inoculation methods for both bacterial strains were selected for a pathogenicity test. Tomato plants were used for the pathogenicity test. The tomato plants were inoculated by toothpick method and observed for wilt symptoms. The pathogenicity test was performed for each replication.

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CHAPTER 4 RESULTS Sensitivity of modified SMSA for recovery of R. solanacearum. A dilution series of R. solanacearum ranging from 10 1 to 10 7 cfu/ml was spread plated on modified SMSA and on nutrient agar. The lowest concentration at which colonies formed on modified SMSA was at 10 3 cfu/ml. There was a 94% certainty that at least a single colony would form at 10 3 cfu/ml. At this dilution 17 of the 18 SMSA plates had at least one colony that formed. Indicating that 10 3 cfu/ml is the lowest concentration modified SMSA can detect R. solanacearum suspended in water. Nutrient agar was able to recover cells from every dilution as shown in Fig. 4.1. 05010015020025030012345678Dilution seriesAverage number of R. solanacearum (cfu/ml) Nutrient Agar SMSA Figure 4.1. Sensitivity of modified SMSA for detecting R. solanacearum was determined by comparing a dilution series of known concentrations of the bacterium spread plated on modified SMSA and nutrient agar. On many of the plates the colony count was greater than 300. These plates were only recorded up to 300. Bars represent standard errors. 30

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31 Monitoring of Ralstonia solanacearum in irrigation pond water. Samples were collected for the detection of R. solanacearum in irrigation ponds of tomato farms and ornamental nurseries in northern Florida from August 2003 to May 2005. Of the five ponds, the bacterium was detected consistently in two ponds, once in two others, and never in the fifth pond (Table 4.1). R. solanacearum was isolated from 13 of 78 samples analyzed (16.6%) (Table 4.2). Table 4.2. Incidence and bacterial concentration of R. solanacearum in water samples by direct isolation on modified SMSA agar from August 2003 to May 2005 Number of positive samples/total number of samples at temperatures greater than and equal to 17C or less than 17C Pond 17C <17C Concentration (cfu/ml) of bacterium a 1 5/13 0/13 4.2 x 10 2 3.9 x 10 4 2 5/20 1/20 2.9 x 10 2 5.6 x 10 4 3 1/13 0/13 2.4 x 10 2 4 1/20 0/20 3.7 x 10 2 5 0/12 0/12 0 total 12/78 1/78 a Data are from analyses of water samples and show ranges of concentrations. The bacterial concentration was directly related to the monthly average air temperature at which the samples were taken (Table 4.2). The surface water temperature ranged from 16.0 to 26.3C during the sampling periods when samples were positive for the presence of R. solanacearum. For the majority of the samples that were positive for the bacterium, the air temperature was above 23C. The detected concentration of the pathogen ranged from 2.4x10 2 to 5.6 x 10 4 cfu/ml. The lowest temperature at which the bacterium was recovered on modified SMSA was at 16C. As the temperature declined the population level also declined. During the winter months, December through March, when the average temperature high was less than 17C, the pathogen was undetectable with the methods used in this study (Figures 4.2-4.6). From November 2004 to March

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32 2005 the bacteria in each sample were concentrated by centrifugation and plated onto modified SMSA. The pathogen was not detected in concentrated samples. All R. solanacearum strains that were isolated from the water samples were pathogenic on tomato plants. Wilt symptoms developed within 4 to 5 days, while R. solanacearum Rs5 caused wilt symptoms on tomatoes within 7 to 9 days under growth room conditions. Statistical analysis of the data by Spearmans correlation coefficient (r s ) showed a significant positive correlation between the population levels of R. solanacearum and temperature for ponds 1 and 2. A regression model revealed that temperature had an effect on the density of R. solanacearum in the water (P < 0.001) (Table 4.3). Table 4.3. Correlation between water temperature and levels of R. solanacearum for ponds 1 and 2 from August 2003 to May 2005 Pond r s a P value 1 0.596 0.0034 2 0.626 0.0014 a Spearmans correlation coefficient. Monitoring of R. solanacearum in aquatic weeds associated with the irrigation pond water. A survey was performed on various aquatic weeds that grew in or in close proximity to the irrigation ponds for the presence of R. solanacearum. At ponds where the bacterium was found consistently (i.e., ponds 1 and 2) samples of similar weeds found at both ponds were collected and analyzed. Weeds ranged from the grass Tripsacum floridiana to water surface weeds that included Alternanthera philoxeroides (common name alligator weed), Lemnaceae spp (common name duckweed), Hydrocotyle ranunculoides (common name dollar weed), and Polygonum pennsylvanicum, and the tree Persea palustris (common name swamp bay) (Table 4.4).

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33 Table 4.4. Detection of R. solanacearum isolated from sterilized or nonsterilized roots and/or stems of semi-aquatic plants collected from ponds 1 and 2 by and direct plating onto modified SMSA agar in July 2004 Plants species (number of samples) Detection of R. solanacearum a Bacteria isolated from root or stem a Surface sterilized or nonsterilized a,b Alternanthera philoxeroides (2) Lemnaceae spp (5) Polygonum pennsylvanicum (1) + both both Polygonum pennsylvanicum (1) + both Sterilized Hydrocotyle ranunculoides (1) + both Sterilized Hydrocotyle ranunculoides (1) + both both Tripsacum floridiana (2) Persea palustris (1) a The sample roots and stems from each plant was divided in two groups. One half of the plant material were macerated in 1 ml sterile water and the other half was surface sterilized then macerated in 1 ml sterile water. The macerates were streaked on SMSA medium and plates were incubated at 28C. b R. solanacearum recovery from surface sterilized plant materials indicates latent infection. These plants harbored the bacteria without wilt. H. ranunculoides from the Apiaceae and P. pennsylvanicum from the Polygonaceae families were the only weeds from which R. solanacearum was isolated (Table 4.4). From July to October, R. solanacearum was detected by direct plating onto modified SMSA in both H. ranunculoides and P. pennsylvanicum. Although the bacterium was found on both the stem and root system of each plant, 56% of the positive results were found in the roots of each plant. On average more colonies developed from the root samples than from the stems. Sixty-one percent of the detected bacteria were from the surface sterilized samples. In growth chamber assays, R. solanacearum strains from H. ranunculoides and P. pennsylvanicum were pathogenic on tomato plants and wilt

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34 symptoms were observed in 4 to 5 days (Table 4.5). Control strain Rs5, previously isolated from tomato, caused wilt symptoms in 7 to 9 days. Table 4.5. Incidence R. solanacearum in surface sterilized and nonsterilized stems and roots of Polygonum pennsylvanicum and Hydrocotyle ranunculoides associated with irrigation pond number1 and 2, from July 2004 to May 2005 Dates sampled Plants Detection of R. solanacearum a Bacteria isolated from root or stem a Surface sterilized or nonsterilized a,b July 2004 Polygonum pennsylvanicum + both both Hydrocotyle ranunculoides + both both August 2004 Hydrocotyle ranunculoides + Both Sterilized September 2004 Polygonum pennsylvanicum + Both both Hydrocotyle ranunculoides + Roots Nonsterilized October 2004 Polygonum pennsylvanicum + Both Both Hydrocotyle ranunculoides + Both Both November 2004 Polygonum pennsylvanicum + c Roots Nonsterilized Hydrocotyle ranunculoides + c Roots Nonsterilized December 2004 Hydrocotyle ranunculoides c N/A N/A January 2005 Hydrocotyle ranunculoides c N/A N/A February 2005 Hydrocotyle ranunculoides c N/A N/A March 2005 Hydrocotyle ranunculoides c N/A N/A April 2005 Hydrocotyle ranunculoides c N/A N/A May 2005 Hydrocotyle ranunculoides + Both Nonsterilized Polygonum pennsylvanicum + Roots Nonsterilized a The sample roots and stems from each plant were divided in two groups. One half of the plant material were macerated in 1 ml sterile water and the other half was surface sterilized then macerated in 1 ml sterile water. The macerates were streaked on SMSA medium and plates were incubated at 28C. b R. solanacearum recovery from surface sterilized plant materials indicates latent infection. These plants harbored the bacteria without wilt. c Samples were concentrated by centrifugation.

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35 R. solanacearum was not detected during the winter months on either weed by direct plating. P. pennsylvanicum was not found in the ponds from December to May. The plant samples were concentrated during November 2004 through March 2005 by centrifugation. In November, the bacterium was detected only on the root samples at 2.7 x 10 3 cfu/ml and 1.8 x 10 5 cfu/ml after centrifugation for P. pennsylvanicum and H. ranunculoides, respectfully. Both root samples were untreated, while the untreated stem and the surfaced sterilized samples were negative for detection of the bacterium as shown in Table 4.5. The R. solanacearum population decreased for ponds 1 and 2 during the month of August. By this time growers had been spraying herbicides to control the aquatic weed population and in particular H. ranunculoides and P. pennsylvanicum. By September the weeds had recovered and were at the same density as in July. During the same time period the bacterial population recovered and the population was the same as for July as shown in Table 4.1. Effects of chemical treatments on R. solanacearum. Two different methods of chemical treatment gave similar results to a known concentration of R. solanacearum. In first method a bacterial concentration of 10 8 cfu/ml was used. At this concentration of bacteria the chlorine treatment resulted in a minimum inhibitory concentration (MIC) of 4.76 mg/l of free Cl 2 At this concentration, 100% of the bacterial cells were completely killed. The MIC for hydrogen peroxide at this bacterial concentration was not determined (Figure 4.7). The bacterial suspension was exposed to 300 mg/l of hydrogen peroxide (data not shown) and the chemical failed to inhibit colony growth.

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36 0%20%40%60%80%100%2.002.503.003.504.004.505.0010.00mg/lPercentage of Recovered Colonies Chlorine Hydrogen perioxide Figure 4.7. The effects of free chlorine and hydrogen peroxide at various concentrations on 10 8 cfu/ml of Ralstonia solanacearum. After exposure to the chemicals, the solution was spread plated on SMSA. The untreated bacterial suspension was spread plated and served as the control. The chemical treatments were compared to the untreated control to give the percentage of recovered cells 0%20%40%60%80%100%2.002.503.003.504.004.505.0010.00mg/lPercentage of Recovered Colonies Chlorine Hydrogen Perioxide Figure 4.8. The effects of free chlorine and hydrogen peroxide at various concentrations on 10 4 cfu/ml of Ralstonia solanacearum. After exposure to the chemicals, the suspension was spread plated on SMSA. The untreated bacterial suspension was spread plated and served as the control. The chemical treatments were compared to the untreated control to give the percentage of recovered cells. Samples from the collected irrigation water samples indicated that 10 4 cfu/ml would be the highest population level bacterium would reach in the pond water. The

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37 method was altered to resemble the sa me concentration found in ponds. At 10 4 cfu/ml, no R. solanacearum colonies developed at 2 l/l of ch lorine. Hydrogen peroxide again failed to inhibit colony growth even at this low concentration of R. solanacearum (Figure 4.8). Inoculation of H. ranunculoides with R. solanacearum H. ranunculoides plants were inoculated by two different methods and with two different strains of R. solanacearum Plants were inoculated by the toothpick or drench method. The strain SEF was isolated from pond 1 in 2004, while Rs5 was isolated from tomato in 2000. Colonization of the bacterium on this weed host has not been studi ed. Unexpectedly every plant that was inoculated by Rs5 died. Those plants inoculated by the toothpick method showed wilt symptoms within 5 to 6 days post inoculation. By day 8 all the plants died. While the majority of the pl ants inoculated by the drench method showed wilt symptoms by day 10, all the plants died fourteen days after inoculation. The uninoculated control plants fo r both inoculation techniques of the Rs5 strain developed wilt symptoms and died. However, none of the plants inoculated by SEF exhibited wilt symptoms.

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Table 4.1. Populations (cfu/ml) of R. solanacearum in five irrigation ponds from August 2003 to May 2005 38 Date samples taken Pond Aug, 2003 Sept, 2003 Oct, 2003 Nov, 2003 Dec, 2003 Jan, 2004 Feb, 2004 March, 2004 April, 2004 May, 2004 June, 2004 1 a e e e e e e e e 4.2 x 10 2 5.6 x 10 3 3.2 x 10 3 2 e e e e e e e e e e 3.5 x 10 2 3 b NS d NS d NS d NS d NS d NS d NS d NS d e e e 4 e e e e e e e e e e e 5 c NS d NS d NS d NS d NS d NS d NS d NS d NS d e e a Samples from pond 1 were discontinued after September 2004. b Pond 3 did not have water until April 2004. c Samples from pond 5 did not commence until May 2004. d NS stands for no sample was taken. e indicates no bacterium was detected.

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Table 4.1 continued 39 Date samples taken Pond July, 2004 Aug, 2004 Sept, 2004 Oct, 2004 Nov, 2004 Dec, 2004 Jan, 2005 Feb, 2005 March, 2005 April, 2005 May, 2005 1 a 3.9 x 10 4 3.9 x 10 3 NS d NS d NS d NS d NS d NS d NS d NS d NS d 2 3.6 x 10 3 6.8 x 10 3 9.3 x 10 3 5.6 x 10 4 2.9 x 10 2 e e e e e e 3 b e e e 2.4 x 10 2 e e e e e e e 4 e e e 3.7 x 10 2 e e e e e e e 5 c e e e e e e e e e e e a Samples from pond 1 were discontinued after August 2004. b Pond 3 did not have water until April 2004. c Samples from pond 5 did not commence until May 2004. d NS stands for no sample was taken. e indicates no bacterium was detected.

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051015202530Aug-03Sep-03Oct-03Nov-03Dec-03Jan-04Feb-04Mar-04Apr-04May-04Jun-04Jul-04Aug-04Temperature (C)00.511.522.533.544.55Log 10 cfu/ml irrigation wate r Temperature BacterialConcentration Figure 4.2. Detection of R. solanacearum colonies by direct isolation on modified SMSA medium during a survey of Pond 1 from August 2003 to August 2004. Surface water samples were collected at five different locations along the banks of the pond. The dates indicate the sampling months during the years 2003 through 2004. The populations of the bacterium (from 0 to 3.9 x 10 4 cfu/ml) and the temperatures at the time the samples were taken are indicated in the figure. 40

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051015202530Aug-03Sep-03Oct-03Nov-03Dec-03Jan-04Feb-04Mar-04Apr-04May-04Jun-04Jul-04Aug-04Sep-04Oct-04Nov-04Dec-04Jan-05Feb-05Mar-05Apr-05May-05Temperature (C)00.511.522.533.544.55Log 10 cfu/ml irrigation wate r Temperature BacerialConcentration Figure 4.3. Detection of R. solanacearum colonies by direct isolation on modified SMSA medium during a survey of Pond 2 from August 2003 to May 2005. Surface water samples were collected at five different locations along the banks of the pond. The dates indicate the sampling months during the years 2003 through 2005. The populations of the bacterium (from 0 to 5.6 x 10 4 cfu/ml) and the temperatures at the time the samples were taken are indicated in the figure 41

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051015202530Apr-04May-04Jun-04Jul-04Aug-04Sep-04Oct-04Nov-04Dec-04Jan-05Feb-05Mar-05Apr-05May-05Temperature (C)00.511.522.533.544.55Log 10 cfu/ml irrigation wate r Temperature BacerialConcentration Figure 4.4. Detection of R. solanacearum colonies by direct isolation on modified SMSA medium during a survey of Pond 3 from April 2004 to May 2005. Surface water samples were collected at five different locations along the banks of the pond. The dates indicate the sampling months during the years 2004 through 2005. The populations of the bacterium (from 0 to 2.4 x 10 2 cfu/ml) and the temperatures at the time the samples were taken are indicated in the figure. 42

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051015202530Aug-03Sep-03Oct-03Nov-03Dec-03Jan-04Feb-04Mar-04Apr-04May-04Jun-04Jul-04Aug-04Sep-04Oct-04Nov-04Dec-04Jan-05Feb-05Mar-05Apr-05May-05Temperature (C)00.511.522.533.544.55Log 10 cfu/ml irrigation wate r Temperature BacterialConcentration Figure 4.5. Detection of R. solanacearum colonies by direct isolation on modified SMSA medium during a survey of Pond 4 from August 2003 to May 2005. Surface water samples were collected at five different locations along the banks of the pond. The dates indicate the sampling months during the years 2003 through 2005. The populations of the bacterium (from 0 to 3.7 x 10 2 cfu/ml) and the temperatures at the time the samples were taken are indicated in the figure. 43

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051015202530May-04Jun-04Jul-04Aug-04Sep-04Oct-04Nov-04Dec-04Jan-05Feb-05Mar-05Apr-05May-05Temperature (C)00.511.522.533.544.55Log 10 CFU/ml irrigation wate r Temperature BacterialConcentration Figure 4.6. Detection of R. solanacearum colonies by direct isolation on modified SMSA medium during a survey of Pond 5 from May 2004 to May 2005. Surface water samples were collected at five different locations along the banks of the pond. The dates indicate the sampling months during the years 2004 through 2005. The populations of the bacterium were never detected from this pond and (0 cfu/ml )the temperatures at the time the samples were taken are indicated in the figure. 44

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CHAPTER 5 DISCUSSION AND CONCLUSIONS An investigation of the population densities of R. solanacearum over a two and a half year period of analyses at five different irrigation ponds on tomato farms and ornamental nurseries in northern Florida associated with bacterial wilt revealed that R. solanacearum was detected in pond water and in two weed species (H. ranunculoides from the Apiaceae and P. pennsylvanicum from the Polygonaceae families) associated with the pond environment. There was a correlation between detection of the pathogen and the recorded air temperature, confirming results reported in the United Kingdom (Elphinstone et al. 1998) and Spain (Caruso et al, 2005). Based on analyses performed in 2004, the first detection of R. solanacearum was in April, when the temperature was above 17C. The abundance of this bacterium was relatively higher (from 3.5 x 10 2 to 5.6 x 10 4 cfu/ml) from April to November, and then decreased dramatically until it was undetectable in December when temperatures were below 17C. The majority of reports of incidences of the disease in northern Florida have been during the fall harvest season, which correlates with density of the pathogens population in the pond environment. These results agree with previous studies in Spain, the United Kingdom, and the Netherlands, where the levels of R. solanacearum r3 b 2 in waterways were usually 80 cells/ml, dropping below the detection limit when exposed to cold temperatures (Caruso et al, 2005; Elphinstone, 1998; Janse, 1998). 45

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46 Bacterial populations varied from pond to pond. Descriptions of the five ponds are as follows; Pond 1 was a retention pond for an ornamental nursery. The waters source originated from the run off of the greenhouses and rain. The water was re-circulated into the greenhouse after going through a filtering process. R. solanacearum was detected as early as April 2004. This pond contained the highest concentrations of the bacterium, until sampling was discontinued. In June 2004, the producer removed all of the aquatic weeds and other weeds that surrounded the banks, and the population of R. solanacearum was reduced from 10 3 to 10 2 cfu/ml. However, by the next month the weeds grew back and population levels were as high as 3.9 x 10 4 cfu/ml. Plants from this pond were analyzed for colonization of R. solanacearum. The only plants that tested positive were H. ranunculoides and P. pennsylvanicum. This pond was not shaded by any trees, and vegetation growth was very abundant in and around the pond. The pond had an aeration system that kept the water in constant motion. Pond 2 was located in the middle of the tomato production fields. The source of the pond was tributary creeks that feed the pond. Samples were taken only on the south side of the pond due to the lack of access to the other sides. R. solanacearum was only found where H. ranunculoides and P. pennsylvanicum grew, which was only the southeast side of the pond. In June 2004, the grower sprayed an herbicide that killed most of the aquatic weeds. As a result populations were reduced, yet by July 2004 the plants grew back and bacterial populations were as high as 10 3 cfu/ml Pond 2 had tree coverage along the sides of the pond, but the trees did not provide much shade near the banks of the pond. Much vegetation grew in this pond along the banks of the pond.

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47 Pond 3 was a retention pond that was dry during the winter of 2003, but during 2004 was filled with water. Water was irrigated from this pond for the tomato farm across the street. None of the common aquatic weeds found at the other ponds were observed at this pond. R. solanacearum had been detected only once in sample taken in October 2004. Bacteria wilt of tomato had been a problem in the past for this particular farm; however, there were no reports of the disease during the tomato growing seasons in 2004. Pond 4 was located near Pond 3 and was surrounded by wooded hills. Pond 4 was the largest pond and was fed by a few local streams. Like pond 3, detection of R. solanacearum was only in October 2004. H. ranunculoides and P. pennsylvanicum were not observed at the areas where samples were taken. Bank areas of this pond were not heavily vegetated and sunlight was restricted due to the amount of trees in the area. Pond 5 was located at the far eastside of the tomato fields. At the far southwest corner of the field a few tomato plants had bacterial wilt. R. solanacearum was isolated from these plants. Water samples have been taken from this retention pond since May 2004; however, R. solanacearum was not isolated from the water. The surface of this pond was completely covered by duckweed (Lemnaceae spp), and was completely shaded by the surrounding trees. Further investigations would need to take place to understand why the pathogen was so prominent in ponds 1 and 2, yet not in 3-5. All five ponds had reports of bacterial wilt on each property and in all the ponds besides pond 5 R. solanacearum was detected. The main difference of each pond was the population of semi-aquatic weeds. In ponds 1 and 2, both H. ranunculoides and P. pennsylvanicum grew abundantly, while in the other

PAGE 56

48 ponds neither weed grew. Since both plants grow best in sunny conditions, a possible explanation for the lack of growth of the weeds could be due to the amount of sunlight that reaches these ponds. Understanding the difference between R. solanacearum positive ponds and negative ponds will aid us in controlling the pathogen. After a survey of several common semi-aquatic and other weeds, H. ranunculoides and P. pennsylvanicum were positive for the bacterium both in and on the surface of the stems and roots by direct plating an extraction of plant samples and water on SMSA. The bacterium was not detected by direct plating during the winter months when the temperature was below 17C. However, when the plant samples were concentrated by centrifugation, the bacterium was detected. To further understand the connection between these symptomless weed hosts and the bacterium further methods must be considered. Comparing both the plant sampling from the ponds and from the greenhouse experiment, it is assumed that the bacterium can colonize the surface of roots or stems, as well as colonize them internally. To confirm where colonization takes place, H. ranunculoides and P. pennsylvanicum would be inoculated with an R. solanacearum strain containing the green fluorescent gene (GFP). The bacterium could then be seen using a confocal microscope and the exact location of the bacterium could be determined. Confirmation of colonization of the bacterium on these weeds would benefit the growers to know which weeds to eradicate in order to help control to the pathogen population. The results of the chlorine and hydrogen peroxide indicate that the bacterium can be controlled by chlorine, however the hydrogen peroxide was ineffective. At 4.76 mg/l and 1.9 mg/l of free Cl 2 R. solanacearum can be eliminated at 10 8 cfu/ml, and 10 4 cfu/ml respectfully, however, both experiments were performed when the water and bacterial

PAGE 57

49 suspensions were stagnant. Chlorine would most likely be added to the water when or after it passes through the water pumping station. Water in this stage would be moving very rapidly, so tests would further need to be performed to find the correct MIC value for the water in motion. However, using chlorine as a means to control the population of the bacterium, one would need to be concerned by the negative effects of chlorine in the environment, the effects of a build-up of chloride in the soil, and its effects on tomato plants. Hydrogen peroxide was ineffective for each concentration of bacterial cells in this study. Hydrogen peroxide usually decomposes in the presence of numerous catalysts such as most metals, acids, or oxidizable organic materials. A small amount of stabilizer, usually acetanilide, is often added to it. Thus, without a stabilizer hydrogen peroxide would be inefficient, due to the amount of catalysts found in the pond water. Market products with the active ingredient being hydrogen peroxide were considered; however, after speaking with a representative from one of the manufacturers, he calculated that the volume needed to control the pathogen in irrigation water would result in expense of about $2,000 for 60 mg/l per acre foot of water. Between 50-60 mg/l is average for bacterial control for use of this product. SMSA can detect the presence of R. solanacearum suspended in water 94% of the time at 10 3 cfu/ml. For the purpose of the population study of this bacterium in the water, this concentration was sufficient, yet other methods could detect the lower concentration of the bacterium in low densities. Such method as enriching plus PCR and ELISA could be very effective in detecting the presence of the bacterium, but these techniques would require training, time and money.

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50 R. solanacearum was found to colonize H. ranunculoides by both inoculation methods. Results are being processed to determine where the bacterium colonizes on the plants. All the H. ranunculoides plants inoculated by the toothpick method with the laboratory strain Rs5 developed wilt symptoms within 5 to 6 days post inoculation. While those inoculated with the same strain using the drench method developed wilt symptoms 10 days after inoculation, the majority of the plants inoculated with Rs5 were unable to regenerate and no new growth was seen. None of the plants inoculated with SEF developed wilt symptoms. Rs5 was isolated from a tomato plant in November 2000, and SEF was isolated from a water sample from pond 1 in July 2004. It is assumed that the over the years bacterium has evolved to one which is able to coexist with the semi-aquatic weeds. Molecular analysis of the two different strains would need to be done to determine the differences. Contaminated irrigation water is a potential source of infection in north Florida and some of the semi-aquatic weeds act as a secondary host in which R. solanacearum could overwinter and multiply to further contaminate irrigation ponds. Tomato, geranium and hydrangea growers need to take into consideration these results, and use management tactics to avoid this source of inoculum.

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LIST OF REFERENCES Aberdeen, J. E. C. (1945). Diseases of tomatoes and their control. Queensland Agr. Jour. 63, 277-299. Anith, K. N., Momol, M. T, Kloepper, J. W., Marois, J. J., Olson, S. M., and Jones, J. B. (2004). Efficacy of plant growth-promoting rhizobacteria, acibenzolar-s-methyl, and soil amendment for integrated management of bacterial wilt on tomato. Plant Disease, 88(6), 669-673. Buddenhagen, I. W., Sequeira, L., and Kelman, A. (1962). Designation of races of Psuedomonas solanacearum. Phytopathology 52, 726. Buitrago Gallego, E. (2001). Impacto socio-econmico de la enfermedad del moko en plantaciones de pltano y banano en seis municipios del departamento del Quindo, Julio 1998-Diciembre 2000. In: Seminario Taller Manejo intergrado de sigatokas, moko y picudo negro del pltano en el eje cafetero. Armenia 24-25 Mayo, 2001. Caruso, P., Palomo, J. L., Bertolini, E., lvarez, B., Lpez, M., and Biosca E. (2005). Seasonal variation of Ralstonia solanacearum biovar 2 populations in a Spanish river: recovery of stressed cells at low temperatures. Applied and Environmental Microbiology 71(1) 140-148. Cook, D. R., and Sequeira, L. (1988). The use of restriction fragment length polymorphism (RFLP) analysis in taxonomy and diagnosis. ACIAR Bacterial Wilt Newsletter. No. 4, 4. Cook, D. R., and Sequeira, L. (1994). Strain differentiation of Pseudomonas solanacearum by molecular genetic methods. Bacterial Wilt: The Disease and Its Causative Agent, Pseudomonas solanacearum (Ed. by Hayward, A. C.; Hartman, G. L.), CAB International, Wallingford, UK. 77-93. Coutinho, T. A. (2005). Introduction and prospectus on the survival of Ralstonia solanacearum. Bacterial Wilt Disease and the Ralstonia solanacearum Species Complex. (Ed. by Allen, C., Prior, P., and Hayward, A. C.), The American Phytopathological Society, St. Paul, MN. 29-38. Department for Environment, Food and Rural Affairs (DEFRA) (2003). Potato brown rot: monitoring programme. Retrieved Jun. 24, 2005, from http://www.defra.gov.uk/planth/pbr1.htm 51

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52 Elphinstone, J. G., Stanford, H. M., and Stead, D. E. (1997). Detection of Ralstonia solanacearum in potato tubers, Solanum dulcamara and associated irrigation water. Bacterial Wilt disease. Molecular and ecological aspects. (Ed. by Prior, P., Allen, and Elphinstone, J. G.), Springer-Verlag, Heidelberg, Germany. 133-139. Elphinstone, J. G., Stanford, H. M., and Stead, D. E. (1998). Survival and transmission of Ralstonia solanacearum in aquatic plants of Solanum dulcamara and associated surface water in England. Bull. OEPP 28, 93-94 Elphinstone, J. G. (2005). The current bacterial wilt situation: a global overview. Bacterial Wilt Disease and the Ralstonia solanacearum Species Complex. (Ed. by Allen, C., Prior, P., and Hayward, A. C.), The American Phytopathological Society, St. Paul, MN. 9-28. Englebrecht, M. C., (1994). Modification of a semi-selective medium for the isolation and quantification of Pseudomonas solanacearum. Bacterial Wilt Newsletter 10, 2-5. European and Mediterranean Plant Protection Organization (EPPO) (2000). Data sheets on quarantine pests Ralstonia solanacearum. EPPO quarantine pest. CAB International, Wallingford, UK. European and Mediterranean Plant Protection Organization (EPPO) (2004). Optimised protocols for diagnosis of potato brown rot and bacterial wilt and detection of the causal bacterium (Ralstonia solanacearum) in infected plants, surface water, soil, industrial potato waste and sewage. CAB International, Wallingford, UK. Farag, N. S., Lashin, S. M., All-Abdel, R. S., Shatta, H. M., and Seif-Elyazal, A. M., (1982). Antibiotics and control of potato black leg and brown rot diseases. Agricultrual Research Review. 60, 149-166. Farag, N. S., Fawzi, F. G., El-Said, S. I. A., and Mikhail, M. S., (1986). Streptomycin in relation to potato brown rot control. Acta Phytopathologica et Entomolgical Hungarica. 21, 115-122. Farag, N. S., Stead. D. E., and Janse, J. D., (1998). Ralstonia (Pseudomonas) solanacearum race 3, biovar 2, detected in surface (irrigation) water in Egypt. J. Phytopathological 147, 485-487. Federal Register, December 13, 2002, Part V, Department of Agriculture, APHIS, 7 CFR Part 331, 9 CFR Part 121. Florida Agricultural Statistics Service. Vegetable Summary 2003-2004. Retrieved April 23, 2005, from http://www.nass.usda.gov/fl/rtoc0ho.htm Florida Tomato Committee. Florida Tomato Facts and Sizing. Retrieved April 23, 2005, from http://www.floridatomatoes.org/facts.html

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53 French, E. R., Sequeira, L. (1968). Bacterial wilt or moko of plantain in Peru. Fitopatologia 3, 27-38. Hayward A. C., (1964). Characteristic of Pseudomonas solanacearum. Journal of Applied Bacteriology. 27, 265-277. Hayward A. C., (1983). Pseudomonas solanacearum: bacterial wilt and moko disease. Plant Bacterial Disease. (ed. by Fahy, P. C., Persley, G. J.), Academic Press, Sydney, Australia. 129-135. Hayward A. C., (1991). Biology and epidemiology of bacterial wilt caused by Pseudomonas solanacearum. Annu. Rev. Phytopathol. 29, 65-87. Hayward A. C., (1994). The hosts of Pseudomonas solanacearum. Bacterial Wilt: the disease and its causative agent, Pseudomonas solanacearum. (ed. by Hayward, A. C., and Hartman, G. L.), CAB International, Wallingford, UK. 9-24. Hayward A. C., (1997). Round table on bacterial wilt (brown rot) of potato. Bacterial Wilt disease. Molecular and ecological aspects. (Ed. by Prior, P., Allen, and Elphinstone, J. G.), Springer-Verlag, Heidelberg, Germany. 420-430. Hong, J. C., Ji, P., Momol, M. T., Jones, J. B., Olson, S. M., Pradhanang, P., and Guven, K. (2004). Ralstonia solanacearum detection in tomato irrigation ponds and weeds. Presented at First International Symposium on Tomato Diseases and 19th Annual Tomato Disease Workshop, Orlando FL. Janse, J. D. (1996). Potato brown rot in western Europe history, present occurrence and some remarks on possible origin, epidemiology and control strategies. Bull. OEPP 26, 679-695. Janse, J. D. Araluppan, F. A. X., Schans, J., Wenneker, M., and Westerhuis, W. (1997). Experiences with bacterial brown rot Ralstonia solanacearum biovar 2, race 3 in the Netherlands. Bacterial Wilt disease. Molecular and ecological aspects. (Ed. by Prior, P., Allen, and Elphinstone, J. G.), Springer-Verlag, Heidelberg, Germany. 146-152. Janse, J. D., van den Beld, H. E., Elphinstone, J., Simpkins, S., Tjou-Tam-Sin, L. N. A., and van Vaerenbergh, J. (2005). Introduction to Europe of Ralstonia solanacearum biovar 2, race 3, in Pelargonium zonale cuttings from Kenya. Bacterial Wilt Disease and the Ralstonia solanacearum Species Complex. (Ed. by Allen, C., Prior, P., and Hayward, A. C.), The American Phytopathological Society, St. Paul, MN. 81-94. Ji, P., Momol, M. T., Olson, S. M. and Pradhanang, P. M., and Jones, J. B. (2005). Evaluation of thymol as biofumigant for control of bacterial wilt of tomato under field conditions. Plant Disease 89(5), 497-500.

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54 Kelman, A. (1953). The Bacterial Wilt caused by Pseudomonas solanacearum. A Literature Review and Bibliography. A N. C. State College Publication, Raleigh, N. C. Kim, S. H., Olson, R. N. and Schaad, N. (2002). Ralstonia solanacearum biovar 2, race 3 in geraniums imported from Guatemala to Pennsylvania in 1999. Plant Disease 92, 42. Lelliott, R. A., and Stead, D. E., (1987). Methods for the Diagnosis of Bacterial Diseases of Plants. Blackwell Scientific, Oxford, UK. McCarter, S. M. (1991). Bacterial wilt. Compendium of Tomato Disease. (Ed. by Jones, J. B., Jones, J. P., Stall, R. E., and Zitter, T. A.) American Phytopathological Society, St. Paul, MN. 28-29. MIDI, Inc. (2001). Identification of bacteria by gas chromatography of cellular fatty acids. Retrieved June 17, 2005, from http://www.midi-inc.com/media/pdfs/TechNote_101.pdf Momol, M. T., Pradhanang, P., Loper,. (2004). Bacterial wilt. Compendium of pepper disease. American Phytopathological Society, St. Paul, MN. Murakoshi, S., and Takahashi, M. (1984). Trials of some control of tomato wilt caused by Pseudomonas solanacearum. Bulletin of the Kanagawa Horticultural Experiment Station 31, 50-56. Nakaho, K., Hibino, H., and Miyagawa, H. (2000). Possible mechanisms limiting movement of Ralstonia solanacearum in resistant tomato tissues. Journal of Phytopathology-Phytopathologische Zeitschrift 148 (3) 181-190. Olsson, K. (1973). A new bacterial disease in potatoes in Sweden caused by Pseudomonas solanacearum. Vxtskyddsnotiser, 37 (5), 66-69 Olsson, K (1976). Experience of brown rot caused by Pseudomonas solanacearum in Sweeden. Bull. OPPE 6, 199-207. Olsson, K. (1977). Current news of poato brown rot. Vxtskyddsrapporter, Jordbruk, 8, 33-37. Olsson, K. (1979). Potato brown rot bacteria (Pseudomonas solanacearum) in the water of two streams. Vxtskyddsrapporter, Jordbruk 22, 170-182. Ono, K., Hara, H. and Akazawa, J. (1984). Ecological studies on the bacterial wilt, caused by Pseudomonas solanacearum. V. The movement of the pathogen in tobacco plants. Bulletin of the Okayama Tobacco Experiment Station 43, 41-46.

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55 Ooshiro, A., Takaesu, K., Natsume, M., Taba, S., Nasu, K., Uehara, M., and Muramoto, Y. (2004). Identification and use of a wild plant with antimicrobial activity against Ralstonia solanacearum, the cause of bacterial wilt of potato. Weed Biology and Management 4, 187-194. Persson, P. (1998). Successful eradication of Ralstonia solanacearum from Sweden. Bull OEPP/EPPO 28, 113-119. Petrolini, B., Quaroni, S., and Saracchi, M. (1986). Scanning electron microscopy investigations on the relationships between bacteria and plant tissues. II. Investigations on the initial process of Pseudomonas solanacearum pathogenesis. Rivista di Patologia Vegetale 22, 100-115. Pradhanang, P. M., Momol, M. T., Olson, S. M., and Jones, J. B. (2003). Effects of plant essential oils on Ralstonia solanacearum population density and bacterial wilt incidence in tomato. Plant Disease 87(4), 423-427. Pradhanang, P.M., and Momol, M.T. (2001). Survival of Ralstonia solanacearum in soil under irrigated rice culture and aquatic weeds. Journal of Phytopathology 149, 707-711. Prior, P., and Fegan, M. (2005). Diversity and molecular detection of Ralstonia solanacearum race 2 strain by multiplex PCR. Bacterial Wilt Disease and the Ralstonia solanacearum Species Complex. (Ed. by Allen, C., Prior, P., and Hayward, A. C.), The American Phytopathological Society, St. Paul, MN. 405-414. Stead, D. E., Elphinstone, J. G., and Pemberton, A. W. (1996). Potato brown rot in Europe. Proceedings, Brighton Crop Protection Conference Pest and Diseases 1996, 1145-1152 Trigalet A, Frey P, and Trigalet-Demery, D. (1994). Biological control of bacterial wilt caused by Pseudomonas solanacearum: State of the art and understanding. Pages 225-233 in : Bacterial Wilt: The Disease and Its Causative Agent, Pseudomonas solanacearum. (A.C. Hayward and G.L. Hartman, eds,) CAB International, Wallingford, Oxon, UK. Volcani, Z.; Palti, J. (1960). Pseudomonas solanacearum in Israel. Plant Disease Reporter 44, 448-449. Williamson, L., Nakoho, K., Hudelson, B. and Allen, C. (2002). Ralstonia solanacearum race 3, biovar 2 strains isolated from geranium are pathogenic on potato. Plant Dis. 86, 987-991. Yabuuchi, E., Kosak, Y., Yano, I., Hotta, H., and Nishiuchi, Y. (1995). Transfer of two Burkholderia and an Alcaligenes species to Ralstonia gen. nov. Proposal of Ralstonia picketti (Ralston, Palleroni and Doudoroff 1973) comb. nov., Ralstonia solanacearum (Smith 1896) comb. nov. and Ralstonia eutropha (Davis 1969) comb. nov. Microbiol. Immunol. 39, 897-904.

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56 Zachos, D.G. (1957). The brown rot of potatoes in Greece. Annales de l'Institut Phytopathologique Benaki, New Series 1, 115-117. Zehr, E.I. (1969). Studies of the distribution and economic importance of Pseudomonas solanacearum in certain crops in the Philippines. Philippine Agriculturist 53, 218-223.

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BIOGRAPHICAL SKETCH Jason Hong was born in Logan, Utah, in 1978, but grew up in Wooster, Ohio. For the last two years of high school he was dual enrolled at the local community college and graduated with an Associate of Science the same year he graduated from high school. In 1997, he served a two year mission for The Church of Jesus Christ of Later-Day Saints in Santiago, Chile. In 2003, he graduated from The Ohio State University with a degree of Bachelor of Science in microbiology. During the summers of 2002 and 2003 he had internships with Dr. Jackson at AgriPhi (now Omnilytics) and was exposed to plant pathology, bacteriophage and research. He attended the graduate program of the University of Florida, College of Agricultural and Life Sciences, Department of Plant Pathology, from August 2003 to August 2005. He conducted a research project to monitor the population of Ralstonia solanacearum in irrigation pond water, aquatic weeds, and developed methods of control of the bacterium in the water under the guidance of Drs. Timur M. Momol, Jeffery B. Jones, Steve M. Olson, and Jerry A. Bartz. 57


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Material Information

Title: Detection of Ralstonia solanacearum in Irrigation Ponds and Semi-Aquatic Weeds, and Its Chemical Treatment in Water
Physical Description: Mixed Material
Copyright Date: 2008

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Source Institution: University of Florida
Holding Location: University of Florida
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DETECTION OF Ralstonia solanacearum IN IRRIGATION PONDS AND SEMI-
AQUATIC WEEDS, AND ITS CHEMICAL TREATMENT IN WATER















By

JASON CHRISTOPHER HONG


A THESIS PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF
FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE
DEGREE OF MASTER OF SCIENCE

UNIVERSITY OF FLORIDA


2005





























Copyright 2005

by

Jason Christopher Hong















ACKNOWLEDGMENTS

I would like to thank my committee members, Drs. Timur M. Momol, Jeffrey B.

Jones, Steve M. Olson, and Jerry A. Bartz, for their support and guidance through the

entire research project and preparation of this manuscript. I especially would like to

express my appreciation for Dr. Momol's patience and guidance throughout this project,

and Dr. Jones' understanding and mentoring that he has shown.

I would like to express my gratitude for the members of Dr. Momol's lab for their

support and help. I am grateful for Pingsheng Ji for teaching me the various techniques I

would need to know for this project. I am grateful to Hank Dankers for showing me the

location of the ponds and helping me with the work, and for Laura Ritchie for

accompanying me on my visits to the ponds. I am also grateful to the members of Dr.

Jones' lab. I would like to express my gratitude for the instruction, aid, service, and

advice I received from Ellen Dickstein and Jerry Minsavage.

I would also like to thank friends that helped me throughout this project by

providing long hours of service and encouragement. And, my deepest gratitude is

expressed to my parents for their love and support.
















TABLE OF CONTENTS



A C K N O W L E D G M E N T S ................................................................................................. iii

LIST OF TABLES ..................... .......... ....................v

LIST OF FIGURES ..................................... vi

ABSTRACT................................. .............. vii

CHAPTER

1 INTRODUCTION ................... ............................ ......... .. .......... 1

2 L ITE R A TU R E R E V IE W ...................................................................................... 10

Detection of Ralstonia solanacearum in Irrigation Water and Solanum dulcamara .10
Methods for Controlling Ralstonia solanacearum .......................................18

3 MATERIALS AND METHOD.............................................22

4 RESULTS ........................... .................... .........30

5 DISCUSSION AND CONCLUSIONS ........................... ....... ...............45

LIST OF REFEREN CES .............................................................................. .................. 51

BIOGRAPHICAL SKETCH .................................................. ............... 57
















LIST OF TABLES


Table page

4.2 Incidence and bacterial concentration of R. solanacearum in water samples by
direct isolation on modified SMSA agar from August 2003 to May 2005...........31

4.3 Correlation between water temperature and levels of R. solanacearum for ponds
1 and 2 from August 2003 to May 2005 ..........................................32

4.4 Detection of R. solanacearum isolated from sterilized or nonsterilized roots
and/or stems of semi-aquatic plants collected from ponds 1 and 2 by and direct
plating onto modified SM SA agar in July 2004....................................................33

4.5 Incidence R. solanacearum in surface sterilized and nonsterilized stems and
roots of Polygonum pennsylvanicum and Hydrocotyle ranunculoides associated
with irrigation pond number and 2, from July 2004 to May 2005 ......................34

4.1 Populations (cfu/ml) of R. solanacearum in five irrigation ponds from August
2003 to M ay 2005 ......................................... ...... .. ...... 38
















LIST OF FIGURES


Figure page

4.1 Sensitivity of modified SMSA for detecting R. solanacearum was determined by
comparing a dilution series of known concentrations of the bacterium spread
plated on modified SMSA and nutrient agar................................ ..............30

4.7 The effects of free chlorine and hydrogen peroxide at various concentrations on
108 cfu/ml of Ralstonia solanacearum ....................................................................36

4.8 The effects of free chlorine and hydrogen peroxide at various concentrations on
104 cfu/m l of Ralstonia solanacearum .......................................... ............... 36

4.2 Detection of R. solanacearum colonies by direct isolation on modified SMSA
medium during a survey of Pond 1 from August 2003 to August 2004. ................40

4.3 Detection of R. solanacearum colonies by direct isolation on modified SMSA
medium during a survey of Pond 2 from August 2003 to May 2005.....................41

4.4 Detection of R. solanacearum colonies by direct isolation on modified SMSA
medium during a survey of Pond 3 from April 2004 to May 2005..........................42

4.5 Detection of R. solanacearum colonies by direct isolation on modified SMSA
medium during a survey of Pond 4 from August 2003 to May 2005..................43

4.6 Detection of R. solanacearum colonies by direct isolation on modified SMSA
medium during a survey of Pond 5 from May 2004 to May 2005 ...........................44
















Abstract of Thesis Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Master of Science

DETECTION OF Ralstonia solanacearum IN IRRIGATION PONDS AND SEMI-
AQUATIC WEEDS, AND ITS CHEMICAL TREATMENT IN WATER

By

Jason Christopher Hong

August 2005

Chair: Timur M. Momol
Cochair: Jeffrey B. Jones
Major Department: Plant Pathology

Bacterial wilt caused by Ralstonia solanecearum is one of the most destructive

bacterial diseases in the tropical, sub-tropical, and temperate regions of the world,

including the southern U.S. and Florida. It is causing serious yield losses on many crops

such as tomato, potato, pepper, eggplant, tobacco, banana and geranium. It is estimated

that the bacterium is pathogenic on several hundred plant species in over 50 families

throughout the world. On tomato and other plants, R. solanacearum colonizes and blocks

the vascular tissue resulting in wilting and eventual death. In August 2002, R.

solanacearum (rl/bl) was detected in the irrigation pond water of tomato farms and

ornamental nurseries in northern Florida. Research was initiated to sample the irrigation

pond water for the presence of R. solanacearum. In addition to water sampling, various

aquatic weeds were sampled to determine their relationship with the bacterium. The long

term goal of this study was to improve our understanding of R. solanacearum in

irrigation water and weed hosts (including semi-aquatic weeds) in Florida. The









objectives of this research were to study the populations of R. solanacearum within

irrigation ponds by analyses of water and semi-aquatic weeds and other weed samples,

and to determine a method to reduce the bacterial populations before irrigating tomato

fields in Florida.

It was determined that temperature had a direct effect on the population of R.

solanacearum. During the summer months as the temperature increased the population

of the bacterium increased reaching populations as high as 5.6 x 104 cfu/ml. By

December the bacterium was undetectable when spread plated on SMSA, even when

concentrated by centrifugation. In May of the following year, the bacterium was re-

detected.

A survey of semi-aquatic weeds associated with the ponds for the detection of R.

solanacearum indicated that the weeds that were infected with the bacterium were

Hydrocotyle ranunculoides (common name dollar weed), and Polygonum

pennsylvanicum (common name Pennsylvania Smut weed). Both H. ranunculoides and

P. pennsylvanicum were latently infected, showing no signs of wilt. The bacterium was

detected from the weeds by direct plating on modified SMSA in the summer months, but

not in the winter months.

A study also was performed to determine the effects of chlorine and hydrogen

peroxide for possible control of the bacterium in irrigation water. Chlorine was effective

at 2 mg/l at a bacterial concentration of 104 cfu/ml, while hydrogen peroxide was not

effective.














CHAPTER 1
INTRODUCTION

Florida tomato producers are the primary providers for fresh market field-grown

tomatoes (Lycopersicon esculentum Mill.) in the United States year around. The state

accounts for 50% of the domestically produced tomatoes each year, and tomatoes provide

30.3% of the income of vegetables grown in Florida. During the 2003-2004 season,

approximately 42,600 acres of fresh market tomatoes were harvested (Florida Tomato

Committee, 2005). The value of the tomato crop during 2002, which exceeded more

than 466 million dollars, accounted for 27% of all U.S. cash receipts for fresh market

tomatoes (Florida Agriculture Statistical Services, 2005).

Bacterial wilt caused by Ralstonia solanacearum (Smith) Yabuuchi et al., 1995, is

one of the most destructive bacterial diseases in the tropical, sub-tropical, and temperate

regions of the world, including the southern U.S. and Florida. It is causing serious yield

losses on many crops such as tomato, potato, pepper, eggplant, tobacco, banana and

geranium (Pelargonium) (Hayward, 1991). It is estimated that the bacterium is

pathogenic on several hundred plant species in over 50 families throughout the world

(Hayward, 1994).

On tomato and other plants, R. solanacearum colonizes and blocks the vascular

tissue resulting in wilting and eventual death. Thus the disease has been given the name

bacterial wilt. Other common names of the disease have been used based on the location

and plant host, such names as Granville Wilt on tobacco, southern wilt on geranium,

brown rot on potato, and Moko disease on banana. Brown rot of potato caused by R.









solanacearum has been estimated to affect about 3.75 million acres in 80 countries

(except the U.S. and Canada) with global economic loss estimates currently more than

$950 million per year (DEFRA, 2003).

R. solanacearum is a Gram-negative rod, 0.5-1.5 Crm in length, with a single polar

flagellum. The positive staining for poly-8-hydroxybutyrate granules with Sudan Black

B or Nile Blue distinguishes R. solanacearum from Erwinia species. In addition, the

staining appears heavily at the pole ends with carbol fuchsin. Agar colonies are initially

smooth, shining and opalescent, but become brown with age (Lelliott and Stead, 1987).

The bacterium gains entrance into the plant by way of injured roots and stem

wounds or through stomata. Once inside the plant, the bacteria move in the vascular

tissues. This process can be accelerated by high temperature. The speed of the

bacterium's movement is dependent on the plant part that is colonized (Ono et al., 1984).

The bacteria then colonize the primary xylem and secondary xylem tissues. The

pathogen often destroys the pit membranes, and all parenchyma cells adjacent to vessels

infested with bacteria are necrotic and can be colonized by the bacteria (Nakaho et al.,

2000). The organism adheres by polar attraction to the cell surfaces and subsequently

becomes localized at preferential sites of the mesophyll (Petrolini et al., 1986). Bacteria

blocking the plant's vessels, by production of exopolysaccharide, are the major cause of

wilting.

The disease is most severe at 24-350C. R. solanacearum (Race 1 and 2) is rarely

detected in locations where the mean temperature for any winter month falls below 100C.

Thus, there exists a distinct temperature requirement for optimum disease development

and multiplication of the cells (Caruso et al., 2005). High soil moisture and periods of









wet weather or rainy seasons are associated with elevated disease severity. Soil moisture

is also one of the major factors affecting reproduction and survival of the pathogen

(Hayward, 1991).

Symptoms characteristic of bacterial wilt on most hosts are wilting, stunting and

yellowing of the foliage. Symptoms can develop at almost any plant stage. However, the

complete wilting and collapse occur mostly when young and susceptible plants become

infected. The rate of disease development is heavily influenced by environmental factors.

If environmental conditions are favorable, a young tomato plant will develop advanced

symptoms of wilt within a few days (Aberdeen, 1945). Wilted leaves remain on the plant

and maintain their green appearance for many days. In some cases, although true

symptoms may not appear, stunting and dwarfing of the plant might develop as the plant

matures. The vascular tissues of the stem show a brown discoloration and, if the stem is

cut crosswise, drops of white or yellowish bacterial ooze may be visible in clear water

(McCarter, 1991).

R. solanacearum differs from other bacteria in that different strains can have a

different host range, different locations of origination throughout the world, and different

utilization of carbon sources. Due to this fact, classification of the bacterium is complex

and based on the differing points. Previously in attempts to classify the bacterium, it has

been placed in categories of groups, races, biovars, biotypes, sub-races and strains. R.

solanacearum has been defined as a "species complex" by Prior and Fegan (2005). They

suggested a new hierarchical classification scheme with these taxonomic levels: species,

phylotype, sequevar, and clone.









Buddenhagen et al. (1962) distinguished three races on the basis of pathogenicity.

Race 1, affects tobacco, tomato, potato, eggplant, diploid bananas, and many other crops

and weeds, with high a growth temperature optimum (35-370C). Race 2, affects triploid

bananas (causing Moko disease) and Heliconia spp., with a high temperature optimum

(35-370C). Race 3, affects mainly potato, tomato, and geranium, without a high

virulence on other solanaceous crops, and with a lower temperature optimum (270C).

Other hosts are weeds such as Solanum dulcamara, S. nigrum and S. ciereum and the

composite weed Melampodium perfoliatum. Races 4 and 5 affect ginger and mulberry,

respectively (Elphinstone, 2005).

Hayward (1964) distinguished four biovars by their ability to produce acid from

several disaccharides and sugar alcohols. These biovars however do not correlate with

the races that Buddenhagen et al. (1962) had described. Only race 3, the cool

temperature adapted race, is equivalent to biovar 2 (Hayward, 1983). Races and biovars

have been classified into two main groups according to a restriction fragment length

polymorphism (RFLP) analysis (Cook and Sequeira, 1988; 1994). Group 1 consists of

the Asian stains of race 1, biovars 3, 4, and 5. Group 2 includes the American strains of

race 1 biovar 1 (rl/bl) (most common strains in Florida), race 2 biovar 1, and race 3

biovar 2 (r3/b2). Since 1995, R. solanacearum r3/b2 has entered several states of the

U.S. on various occasions in greenhouse grown geranium cuttings. The pathogen was

traced back to infected geranium cuttings that were imported from Guatemala and Kenya

(Williamson et al., 2002; Kim et al., 2003). R3/b2 of R. solanacearum is a listed "select

agent" in the U.S. under the Agricultural Bioterrorism Protection Act of 2002 (Federal

Register, December 13, 2002, Part V, Department of Agriculture, APHIS, 7 CFR Part









331, 9 CFR Part 121) because it has the potential to be a severe threat to the potato

industry and other crops.

R. solanacearum is mainly disseminated in latently infected plant material as well

as through surface water and soil. It survives in surface water and soil usually by

establishing itself in host plants. In the U.S. and Florida, dispersal via surface irrigation

water is not well understood and preliminary findings were reported recently (Hong et

al., 2004).

The sources of infections of R. solanacearum are contaminated soil, in plant

materials, on farm equipment, and in irrigation or surface water. The bacterium survives

in the soil for varying periods of time, and is able to persist between successive crops.

The bacterium has been found to survive in sheltered sites such as plant debris and

latently infected potato tubers, the deeper soil layers and in the rhizosphere of roots of

weed hosts (Hayward, 1991). The range of weed hosts is extensive, but the significance

of weed hosts depends on the environment and cropping systems. Some hosts are

symptomless carriers.

The bacterium has been isolated from irrigation water and proliferates when

alternative aquatic weeds are present (EPPO, 2004). Irrigation water infested with R.

solanacearum has caused several outbreaks on a number of crops (Elphinstone, 1998).

Olsson in 1976 discovered a relationship between farms with infected potato tubers and

the weed Solanum dulcamara, a common species of nightshade found in Sweden

(Olsson, 1976). During the winter months when the temperatures were low R.

solanacearum was undetectable in the irrigation water, but the bacterium could be

detected in the xylem of the adventitious roots of the semi-aquatic weed growing in the









water (Persson 1998). In a well documented case in England, potato brown rot

occurrence was associated with effluent from the potato processing industry and

municipal water purification plants that handled diseased potatoes. The bacterium was

also found colonizing in the presence of S. dulcamara along the river banks (Buitrago,

2001 and EPPO, 2000).

R. solanacearum is a serious obstacle to the culture of many solanaceous plants in

both tropical and temperate regions. The greatest economic damage has been reported on

potatoes, tobacco and tomatoes in the southeastern United States, Indonesia, Brazil,

Colombia and South Africa. From 1966 to 1968 in the Philippines there were average

losses of 15% in tomatoes, 10% in eggplant and Capsicum, and 2-5% in tobacco (Zehr,

1969). In Peru along the Amazon basin, about half of the banana plantations were

affected and rapid spread of the pathogen threatened to destroy plantations throughout the

Peruvian jungle (French and Sequeira, 1968). In India, occasionally total losses occur in

tomato crops. Outbreaks in potato, which caused extensive losses, occurred in Israel

(Volcani and Palti, 1960), and Greece (Zachos, 1957).

Control Measures have been largely unsuccessful due to the nature of the organism,

especially for race 1 with its broad host range, and race 3 with its ability to cause latent

infection on potato tubers. Commercially available chemical control is not known. It has

been noted previously that soil fumigants showed little or no effects (Murakoshi and

Takahashi, 1984). However, it has been reported that broad spectrum soil fumigants

chloropicrinn) will delay the initial disease onset (Ooshiro, 2004). Antibiotics such as

streptomycin, ampicillin, tetracycline and penicillin were basically ineffective (Farag et

al., 1982). In fact, Farag et al., reported an increase of disease when streptomycin was









applied to potato crops in Egypt. Biological control has been investigated (Farag et al.,

1986; Trigalet et al., 1994); yet more work needs to come forth. Resistant cultivars have

been developed in various crops (Hayward, 1991). In crops such as tobacco and peanut,

resistant cultivars have been very successful. Many resistant cultivars of tomato plants

have been developed and are also successful in a particular environment. Yet, it has been

difficult to develop cultivars which are resistant under conditions of high temperature and

humidity in the lowland tropics (Hayward, 1991). Alternative disease management

tactics were investigated with promising results, especially using Thymol (Pradhanag et

al., 2003; Ji et al., 2005) as a biofumigant and acibenzolar-S-methyl as a plant activator

on moderately resistant cultivars (Anith et al., 2004).

Due to limited efficacy of current integrated management strategies, bacterial wilt

continues to be economically important for many economically important crops in

Florida, and many subtropical, tropical, and temperate areas of the world. Cultural

practices, crop rotation and host resistance may provide limited control (Pradhanang el al,

2003).

Tomato growers need to use an integrated approach to lower the impact of bacterial

wilt on tomato production. These are current recommendations (Momol, 2005,

unpublished):

Preplant
* Choose resistant or moderately resistant cultivars, or graft susceptible cultivar onto
resistant rootstock.

* Consider a preplant soil amendment or fumigation for infested fields against R.
solanacearum and nematodes [i.e. Thymol (not commercially available) and
Telone mixture].

* Apply 2-3 years of rotation and cover crops for infested fields to reduce R.
solanacearum, weeds and nematodes.









* Do not irrigate cover and rotation crops with R. solanacearum contaminated pond
or surface water, avoid reinfestation.

* Use well drained and leveled fields, and do not use low-lying areas of the field.

* Raise soil pH to 7.5-7.6 and increase available calcium limingg).

* Consider using infested fields during cooler months for tomato production (i.e.,
spring season for north Florida).

Production
* Exclude the pathogen by applying strict sanitation practices (pathogen free
irrigation water, transplants, stakes, machinery, etc.).

* Chlorinate your irrigation water continuously if you are using surface water or R.
solanacearum infested pond water.

* Irrigate based on water need, avoid over-irrigation.

* Apply Actigard (Syngenta) if you are using moderately resistant cultivars (i.e., FL
7514).

After harvest
* Plow under crop residue immediately.

* Start with suitable rotation and cover crops (i.e., rye for winter, sudan-sorghum for
summer in north Florida) to avoid weeds that support R. solanacearum populations.

Potato growers are advised to avoid fields infested with the pathogen. Growers

should not plant transplants in close proximity to fields where this problem has occurred.

Infested fields should be rotated with a non-susceptible crop, because long term rotation

might reduce the population of the bacterium. Growers should avoid moving equipment

and soil from infested to non-infested fields and using irrigation water from ponds in

close proximity to infested fields. Caution is given to excessive irrigation, because high

soil moisture will induce high incidence of disease or buildup of population (Janse,

1996).

In 2002 August, R. solanacearum (rl/bl) was detected in the irrigation pond water

of tomato farms and ornamental nurseries in north Florida (Momol et al., unpublished).









Most ponds in north Florida are interconnected by both aboveground and underground

streams. In 2002, research was initiated to sample the irrigation pond water for the

presence of R. solanacearum. In addition to water sampling, various aquatic and other

weeds were sampled to determine their relationship with the bacteria. The long term goal

of this study is to improve our understanding of R. solanacearum in relation to irrigation

water and weed hosts (including aquatic weeds) in Florida. The objectives of this

research were to study the populations of R. solanacearum within irrigation ponds by

analyses of water and aquatic weeds and other weed samples, and to determine a method

to reduce the bacterial populations before irrigating tomato fields in Florida.














CHAPTER 2
LITERATURE REVIEW

Bacterial wilt incited by R. solanacaerum, is a major bacterial disease on many

crops in tropical, sub-tropical, and temperate regions of the world. Molecular aspects,

including pathogen genetics, pathogenesis, host-plant interactions, molecular detection,

and strain differentiation, are more studied than its epidemiology and ecology. Pathogen

survival studies are very important in terms of understanding sources of inoculum and its

long and short distance dispersal. Advances in semi-selective medium (SMSA,

Engelbrecht 1994) and sensitive and specific molecular methods for R. solanacearum

detection, improved data reliability for epidemiological and ecological studies. These are

three major ways that R. solanacearum can survive: surface and irrigation water, plant

material, and soil.

R. solanacearum r3/b2 that causes brown rot on potatoes has been present in

Europe since 1976 (Olsson, 1976). The description of race 3 survival in river water used

for irrigation and its association with Solanum dulcamara (Elphinstone et al., 1998) was

an important cornerstone in the understanding of its epidemiology and ecology.

Detection of Ralstonia solanacearum in Irrigation Water and Solanum dulcamara

Contaminated irrigation water has been the source of many outbreaks of bacterial

wilt on a number of crops (Elphinstone et al., 1998). R. solanacearum can survive for

long periods of time in water (Kelman, 1953) which has important implications for

agricultural practices.









The first report of R. solanacearum in Europe was made in Sweden. Over 30 years

ago the bacterium was thought to be a warm-temperature pathogen. However, brown

vascular rings were found in market potato tubers, cv. Bintje, during routine inspection in

Sweden in 1972. The symptoms were first thought to be caused by Clavibacter

michiganenis subspecies sepedonicus. Yet, after isolation of the Gram-negative, Sudan-

positive bacterium which caused rapid wilting in tomato plants, a presumptive diagnosis

indicated that the pathogen was R. solanacearum. Strains isolated from the tubers had

optimum temperatures between 25 and 300C, which was lower than what was generally

believed to be typical of R. solanacearum. The isolated strains caused severe wilting

after 5 days in young tomato plants cv. Dansk Export and eggplant cv. Black Beauty

(Olsson, 1973, 1977). Strains were typed as biovar 2 (Olsson, 1976).

It was unexpected to find this pathogen as far north as the southern part of Sweden.

Visual inspection of 400,000 tubers collected in 1973 from different fields and seed-

potato lots, showed that brown rot symptoms were only present in tubers grown in the

same field. Infected tubers were found in low frequency. The following year, in the

same potato farm, but a different field brown rot was observed. The same year, a second

brown rot outbreak occurred 80 km north of the first infection site in the Pinndn stream.

On this farm, tubers from several fields were infected at a frequency of 5% in cv. Bintje

and 2% in cv. Grata.

A research study commenced to find the source of infection and a few facts were

known. Seed potatoes from the same or from related seedlots, which had been used in

other fields, produced healthy tubers. Large populations of the Colorado beetle

(Lepinotarsa decemlineata) had invaded southern parts of Sweden in 1972, coming from









the eastern parts of Europe. Beetles were present in the fields of the first outbreak, but

not in the second. Thus, these two possible sources of the primary infection were ruled

out.

One important common fact for the two separate brown rot epidemics was that all

infected potato tubers had been grown in fields irrigated with surface water from a nearby

stream. Irrigation practices were beginning to be implemented in Sweden in the early

1970's (Hayward et al, 1997). Greenhouse experiments showed that tomato, eggplant,

and wild Solanum species of the local flora, were found to be susceptible to the Swedish

strains of R. solanacearum. These plants were irrigated with water from the streams used

by fields of the infected potatoes. Tomato, eggplant and wild Solanum speices were

planted in the fields where infected tubers had been growing. S. dulcamara was planted

near the streams at the two separate outbreaks. None of the test plants became infected

(Olsson, 1976).

S. dulcamara was found growing near and in the Pinnan stream. In 1974 R.

solanacearum was isolated from some of these plants. Isolation from the various parts of

the plant showed a few cells were detected in the stem at the bottom of the plant.

However, high numbers were found in the xylem of the thin adventitious roots growing

in the water (Olsson, 1977, 1979). The pathogen was isolated the following year from

the aquatic weed, which indicated that the bacterium was able to survive overwinter in S.

dulcamara.

The Pinnan stream was surveyed for infected S. dulcamara. The bacterium was

isolated from plants growing 3 km downstream from where a potato processing plant had

been until 1967. The processing plant had processed both domestic and imported









potatoes, and the waste water was dumped directly into the stream. The stream led to two

dams, in which a large population of infected S. dulcamara grew. Infected potato tubers

had been produced in fields irrigated with water from these dams and downstream from

the processing plant.

In 1977 and 1978, water samples from streams close to infected fields were

analyzed. Samples were filtered through a 0.45 [im membrane to concentrate the

bacteria. The different membranes were washed with sterile water. The suspensions

were inoculated into 3 week old tomato plants. Isolations made from wilting plants

resulted in R. solanacearum being detected from water collected at both locations

(Olsson, 1979). High populations of the bacterium were found when the water was

sampled near the bottom of the Pinndn dam, where S. dulcamara grew in abundance

(Persson, 1998).

Brown rot occurred in Sweden at two separate locations from 1972 through 1977.

Infected crops were destroyed by industrial processing, in which tubers were destroyed

by heat treatment at 1450C. R. solanacearum was undetectable in samples from the

remaining potato particles. Fields where the crop was contaminated were restricted from

growing potatoes for the following 2 years. After the 2 years, were allowed to be grown

again on infested farms potatoes, but only by using certified seed potatoes. It was

recommended not to use the infected stream water for irrigation purposes. Analyses of

water from streams near potato processing plants in southern and western parts of

Sweden in 1979 and 1980 did not detect the bacterium.

Since the reports from Sweden, other outbreaks of R. solanacearum have been

reported in other European countries including Belgium, France, Italy, the Netherlands,









Portugal, Spain, and the UK (Stead et al., 1996). For many cases these strains are not

similar, and the source of infection is unknown (Janse et al, 1997). The pathogen is

thought to have been spread in latently infected seed potatoes in the Netherlands in 1995,

thus causing epidemic outbreaks on more than 90 seed and market potato farms

(Elphinstone et al., 1997).

The Netherlands reported brown rot of potato for the first time in 1992. The origin

of the pathogen is assumed to come from uncertified seed. In the same year outbreaks in

the UK and in Belgium were reported, yet no colonial link could be established (Janse et

al, 1997). In 1995 a series of findings of brown rot in both seed and market tomatoes

could be traced back to one grower of a local variety that was heavily infected. From this

source, paths could be traced other infected lots. On the same infected lots where the

local variety of potato could be found, a few other varieties were discovered to be

infected, usually in latent form. Investigations of surface water near these farms showed

the presence of the brown rot pathogen. Further studies indicated a link between water

usage and diseased potatoes.

An extensive survey of the pathogen in surface water and S. dulcamara was

performed in 1996. About 14,000 samples were taken throughout the entire country.

Five percent of the water samples were positive for R. solanacearum, which resulted in a

ban for irrigating with these streams. The bacterium, similarly, was detected in S.

dulcamara in which the roots were submerged in the water. In many cases the pathogen

was detected in water in close proximity to the potato industry's purification plants. This

brought about the possibility that the bacterium could escape the purification process.









Moreover these findings suggest that probably all infections in the Netherlands originated

from irrigation with contaminated surface water (Janse et al, 1997).

Water samples from northern the Netherlands were taken in 1996 to determine

pathogen population during different seasons. Samples were taken weekly in four rivers

(i.e. one non-contaminated and three contaminated with the bacterium). The bacterium

was never found in the non-infested river. In the heavily infested area the bacterium

could be detected until ice formed. Directly after the ice melted the bacterium could be

detected again, but at very low numbers (Janse et al, 1997).

Outbreaks of brown rot lead to the development of detection methods of R.

solanacearum in S. dulcamara and in irrigation water. In England, in 1992 and 1995,

two outbreaks of brown rot occurred at different farms. The source of the infection in

both outbreaks was contaminated irrigation water, in which, infected S. dulcamara plants

were naturally growing (Stead et al., 1996). A detection process was established to better

understand the nature of the pathogen.

Four hundred twenty S. dulcamara plants were sampled from the various infected

waterways. R. solanacearum was extracted from the stolon section and fibrous

adventitious roots. The pathogen was detected by three methods; 1. indirect enzyme-

linked immunosorbent assays performed on boiled plant samples; 2. polymerase chain

reaction performed on sub-samples of the extract of the boiled samples; and 3. streaking

concentrated extract onto modified semi-selective SMSA. Results varied from each test;

therefore the most reliable detection was to employ the three methods simultaneously

(Elphinstone et al., 1997). Over a four year period, S. dulcamara remained infected from

year to year, and the viable pathogen could be isolated throughout the winter months.









Only plants with roots submerged in water were found to be infected. Other aquatic

weeds growing in contaminated streams were not infected. When volunteer potato plants

were sampled, less than 1% infection of progeny tubers was detected and no infection

was found in volunteers the following year. Thus it was concluded that the highest risk

of dissemination of the pathogen was through establishment of the bacterium in aquatic S.

dulcamara and the release into the waterways used for irrigation (Elphinstone et al.,

1997).

Upon a national survey conducted in the rivers throughout England and Wales,

infected S. dulcamara plants were found in the Thames River, five of its tributaries, and

the Witham River. Samples from the Thames River indicated several possible sources of

entry of the pathogen into the water system. Included in the possible sources were

outflows from sewage treatment plants where S. dulcamara was present. The potential

for spread of the pathogen from domestic or industrial use of imported potatoes was a

possibility and needed further investigation (Elphinstone et al., 1997).

Water samples from the river water were analyzed to monitor R. solanacearum

from a single sampling point over three years. The bacterium was only detected in the

water where S. dulcamara was present. When necessary, samples were centrifuged to

concentrate the pathogen population. Enriched samples were detected by PCR. The

pathogen was only isolated during warm summer and fall months. Population levels

were reduced when the temperature fell, water levels rose, and/or S. dulcamara died back

(Elphinstone et al., 1997).

In Egypt, the bacterium was found in both irrigation and drainage canals near

infected potato fields (Farag, 1998). Contaminated irrigation water was reported as a









source of infection in Kenya (Janse, 2005). Due to the bacterium having many

diversified hosts, one crop contaminating irrigation water could essentially affect

neighboring farms that grew other solanaceous crops.

As the epidemiology and ecology of the pathogen are being studied, the more

complex this organism is becoming. Studying the dynamics of the bacterial population in

the water, a distinct relationship existed between the temperature of the water and the size

of the population (Countinho, 2005). The aquatic weed Solanum dulcamara has been

identified as a symptomless host (Olsson, 1977). Bacterial populations have been shown

to overwinter in this weed, while the bacterium was undetectable in water (Perrson,

1998).

In a Spanish river over a three year period there was a strong relationship between

water temperature and detection of R. solanacearum. Collection of water began in the

Tormes River after a confirmed outbreak of potato brown rot in the late fall of 1999.

Following analysis of the irrigation water, it was determined that the water was

contaminated with R. solanacearum. Samples were collected from five different

locations along the river water and directly spread plated on SMSA. Presumptive

colonies were confirmed using indirect immunofluorescence and double-antibody-

sandwich indirect ELISA. Of a total of 65 samples analyzed over three years, R.

solanacearum was isolated from 22 of them. The temperature range of the surface water

that contained the bacterium varied from 9 to 200C. Most of the samples taken had an

average temperature above 140C, with detection of the pathogen usually being

unsuccessful in lower temperatures. Populations in the water ranged from 10 to 80

cfu/ml during the warmer months. However, by November the bacterium was non-









detectable. There was a significant positive correlation between the R. solanacearum

populations and water temperatures for sampling sites (Caruso et al., 2005).

Methods for Controlling Ralstonia solanacearum

To avoid further dissemination of the pathogen, control measures must be designed

and executed. For tomato production in Florida, latent infection is not a major concern,

except seedlings have to be pathogen free. Current management recommendations were

listed previously (see Introduction) for tomato producers.

The Netherlands has developed guidelines to ensure quality of potatoes and to

control the pathogen that causes brown rot (Janse et. al., 1997). Many other countries

have followed these procedures or have made modifications to fit their needs.

* Testing seed for infections and possible latent infections of the pathogen.

* Survey testing of seed, market, and starch potatoes

* Prohibition of the use of contaminated surface water for irrigation and
discouragement of its use in other areas.

* Eradication of S. dulcamara from infected waterways.

* Survey of contaminated production places where possible infected lots have been.

* Sample and analysis of imported potatoes for latent brown rot.

* Survey surface water areas surrounding contaminated surface water and
purification plants.

* When an infected lot is found, infected potatoes will be destroyed, machinery
disinfested, and all other potatoes and seeds on the farm are to be tested. On
infected fields for five years no potato or tomato are to be grown, control of
volunteers, in the first three years grass or cereals are grown, then in year six
potatoes for seed or market from certified seed can be grown. Testing of all
potatoes produced on the farm during this period and the following year.

* Initiation of national research program to facilitate detection of the pathogen,
including serum production, evaluation of sure and rapid detection methods, and to
study epidemiology in soil, water, and aquatic weeds.









The Swedish Plant Protection Service carried out the first eradication program for

S. dulcamara in the Pinndn stream. Both mechanical and chemical control measures

were used between 1977 and 1981. The herbicide glyphosate, (trade name Roundup,

Monsanto) was used for chemical treatments. S. dulcamara plants were treated manually

with glyphosate and also removed mechanically at specific sites along a 30 km distance

both up and downstream from the former potato processing plant. The herbicide was

applied for 2 years at the two dams, where a large population of the weed existed

(Persson, 1998).

Following the eradication program in 1993 water samples were again collected

from the two Pinna dams. R. solanacearum was not detected. However, soft rot bacteria

Erwinia carotovora subsps carotovora and E. c l1i)iywh,,w/in were detected. E. c.

caratovora is a well known pathogen found in surface water (Persson, 1998). Previously

in the early 1970's, before the reports of brown rot, reports of wilted tomato plants in the

market were made. The plants were irrigated using the same source. The production of

the tomatoes had finished before a lab diagnosis could be made. It was speculated that R.

solanacearum was probably the pathogen, yet later results allowed for the possibility that

E. c. caratovora or E. c hyii//lJweini might have been the causal agent (Persson, 1998).

In 1995 a visual inspection of the Pinna area showed that S. dulcamara was very

rare. The eradication which started 17 years earlier has been successful. Ten plants were

analyzed for possible infection; however, the bacterium was not isolated. In 1996 a

survey of streams in the former infested areas was initiated. Water samples from

irrigation waterways and wastewater from the potato industry were collected and plated

on modified SMSA. Water samples were inoculated in tomato plants, and analyzed using









polyclonal antiserum and ELISA. None of the tests detected the presence of R.

solanacearum in the water. Tests were also carried out on seed potatoes. Analysis of the

potatoes consisted of using a polyclonal antiserum. The pathogen was not detected in the

seed potatoes (Persson, 1998).

Based on these results it was assumed that R. solanacearum had been eradicated

from Sweden. However, as it has been shown before the pathogen can be established in

Sweden's environment. Therefore, careful consideration must be taken for importing

potatoes into the country.

Although control of the bacterium has been possible in Sweden, conditions are not

the same for the rest of Europe. S. dulcamara has proven to be a factor in the

epidemiology of the pathogen, but other weed hosts need to be investigated. The sources

of some waterways are areas of swampy conditions, thus complete eradication by

mechanical and chemical means of S. dulcamara plants would be impossible (Hayward et

al, 1997).

Brown rot is now a major concern in Europe, and control methods are being

implemented to hinder the dispersal of this pathogen. More information is required to

fully understand the epidemiology of the pathogen in cool climates, including studies on

the ecology of the pathogen and identification of survival sites. Studies on the disease

have shown unique features of the epidemiology of the pathogen such as; its survival and

the increase of populations in semi-aquatic S. dulcamara weeds, its distribution in rivers

and waterways, and passage of the pathogen through industrial and domestic waste water.

More information is needed on latently infected weed hosts and the pathogen's method of






21


entry. There exists a need for standardization of test methods so that equivalence is

achieved across Europe and world-wide (Hayward et al, 1997).














CHAPTER 3
MATERIALS AND METHOD

Bacterial culture and inoculum preparation. R. solanacearum (race 1, biovar 1),

tomato strain Rs5 (Pradhanang, 2001) and field strain SEF both isolated in Quincy,

Florida (Gadsden County) were used in these studies. The bacterial pathogen was grown

at 280 C either on modified semi selective medium SMSA (Engelbrecht 1994) or on

casamino acid peptone glucose (CPG) agar (peptone 10 g, casamino acids 1 g, glycerol

2.5 ml, agar 15 g, deionized water 1 liter) for 60 h or in CPG broth on a shaker (100

RPM) for 60 h. Bacterial cells were suspended in sterile deionized water and the

concentration of inoculum was estimated by measuring absorbance at 600nm. The viable

bacterial population was determined following dilution plating on modified SMSA.

Isolation of R. solanacearum from irrigation pond water. Water samples were

collected at five different irrigation ponds used for tomato and ornamental plant

production located in Gadsden County, Florida. The average air temperature for each

month was collected by the weather station at Quincy, FL. Pond 1 was a retention pond

heavily vegetated with various aquatic weeds, located at an ornamental nursery. In this

pond, runoff water from the greenhouses was collected. Five samples were collected

from the banks of this pond from August 2003 until August 2004. Pond 2 was located in

the middle of a tomato grower's field. Four samples were collected on the south end of

the pond. Pond 2 had various kinds of aquatic weeds growing along the banks. Samples

were taken from August 2003 to May 2005. Pond 3 was located on a different tomato

farm and was the smallest of all the ponds. Pond 3 was dry until April 2004 and then









samples commenced until May 2005. Weeds that grew in this pond were not similar to

those found in ponds 1 and 2 because pond 3 was a field during the dry season. Pond 4

was located on the same farm as pond 3. Pond 4 was the largest pond of all the ponds,

and was fed by local streams. This pond was surrounded by hills on all sides except on

the west bank and was heavily shaded by the trees that grew on the hills. Four samples

were collected from August 2003 through May 2005. Pond 5 was added in May 2004

and samples continued until May 2005. This pond was surrounded by hills on all sides

and was located on the south side of a tomato farm. Two samples were collected on the

west bank of this pond. Duckweed completely covered the surface of the pond all year

around and the surrounding trees shaded the pond. Tomatoes infected with bacterial wilt

were found on tomatoes on the northwest corner of the farm.

Surface water samples were collected in the same manner for each pond. Samples

were collected in close proximity to semi-aquatic weeds, namely Hydrocotyle

ranunculoides and Polygonum pennsylvanicum because it was suspected that these weeds

were symptomless hosts from a previous preliminary survey. Water samples were

collected in a 7.6 liter metal bucket and transferred to sterile 50 ml Falcon Tubes. Non-

diluted and ten-fold dilutions of each sample were spread plated on modified selective

medium SMSA. R. solanacearum colonies are irregular shaped with a glossy wet

appearance. The colonies appear white with a purple center in the beginning stages of

growth, then marbling or swirls of purple and white colors appear at 60 h. The plates

were incubated at 28 C for 60 h. Suspected R. solanacearum colonies were selected and

transferred to SMSA. Presumptive colonies were confirmed by whole cell fatty acid

methyl ester analysis (MIDI, 2001).









Pathogenicity of the bacteria was determined by inoculating 10 to 14-day old

Bonny Best tomato plants. Initially, root inoculations were performed by stabbing

around the base of the plants to damage the roots. Following wounding, 10 ml of a

bacterial suspension consisting of 108 cfu/ml were poured along the crown of tomato

plants potted in 10-cm pots. The leaves would curl and droop within 4 to 5 days after

inoculation. The inoculated plants were transferred to a growth room on 12 h light dark

cycle with air temperature at 28 C. Because of inconsistent results, this method was

replaced by the toothpick inoculation method. Upon using the toothpick method,

symptoms developed quicker. Sterile toothpicks with the tips laden with bacteria were

stabbed into the lower stems near the crown of the tomato plants. Each isolated colony

was used to inoculate one plant. Plants stabbed with sterile toothpicks or toothpicks

coated with Rs5 served as the negative and positive controls, respectively. Bacteria were

re-isolated from the tomato plants that developed wilt symptoms and were stored in

sterile 30% glycerol at -800C.

During the winter months when the bacterium was undetectable, the collected water

samples were concentrated by centrifugation. Thirty ml of the water samples were

centrifuged at 10,000 rpm for 10 min at 280C. All but 100 pl of the supernatant was

discarded. The pellet was re-suspended in the remaining supernatant. Non-diluted and

ten-fold dilutions of each sample were spread plated on modified selective medium

SMSA. The plates were incubated at 28 C for 60 h. Suspected R. solanacearum

colonies were selected, transferred to SMSA and characterized using the procedure

previously outlined.









For statistical analysis the data acquired from each sample were transformed using

log transformation [z=log,, (y+l)], and then the data were statistically analyzed using

Speraman's correlation coefficient by SAS for Windows program release 8.02 (Cary,

NC).

Monitoring of R. solanacearum in semi-aquatic and other weeds associated

with irrigation ponds. Common aquatic weeds that were growing along the bank or

submerged in the ponds were collected. Plants were identified by Karen Brown from the

Center for Aquatic and Invasive Plants, University of Florida in Gainesville FL. Plant

samples were analyzed for the presence of R. solanacearum. The plants were first

washed under tap water to remove loose particles of sand and dirt. The roots were

separated into three sections to sample the entire plant part; root hairs to the root tips,

midsection, and from the midsection to where the root attaches to the stem. Stems were

cut in three sections; the stolon, the crown, and before the first leaf. The root sections

were combined and the stem section were combined and given the labels "root" and

"stem".

The cuttings were separated into two groups and were either surface sterilized or

untreated. Sterilizing consisted of soaking the cuttings in 70% ethyl alcohol for 5 min.

Then the cuttings were rinsed in sterile water for 5 min. Each cutting was placed in

separate clear 2 mil zipper plastic bags and mashed using a pestle and 1000 [.l of sterile

water was added to the crushed plant residue. The plant residue was soaked for 20 min.

The suspension was extracted using a pipette and stored in microfuge tubes. Each sample

and a ten-fold dilution were spread plated onto SMSA. The plates were incubated for 60

h at 28 C. Colonies suspected of being R. solanacearum were selected and analyzed by









whole cell fatty acid methyl esters analysis. Confirmed R. solanacearum colonies were

tested for pathogenicity by the toothpick method and re-isolated as described previously.

Bacteria were reisolated from tomato plants that developed wilt symptoms, and were

stored in a sterile 30% glycerol solution at -800C.

During the winter months when the bacterium was undetectable the collected plant

samples were concentrated by centrifugation. Fifteen-hundred microliters of the

macerated plant residue were centrifuged at 10,000 rpm for 10 min. at 280C. All of the

supernatant was discarded. The pellet was re-suspended in 400 .l of sterile water. Non-

diluted and ten-fold dilutions of each sample were spread plated on modified SMSA.

The plates were incubated at 28 C for 60 h. Suspected R. solanacearum colonies were

selected and transferred to SMSA, and identified by whole cell fatty acid methyl esters

analysis. Confirmed R. solanacearum colonies were tested for pathogenicity by the

toothpick method and re-isolated and stored as described previously.

Infested water treatment with chemicals. The minimal inhibitory concentration

(MIC) for chlorine and hydrogen peroxide was determined for a known population of R.

solanacearum. Ultra Clorox Bleach (The Clorox Co., Oakland, CA) was used and

contains 6% active ingredient and 5.71% of free chlorine Cl2. Hydrogen Peroxide

(Diamond Products, Seffner, FL) was used and contained 3% active ingredient, but after

the first trials Hydrogen Peroxide 30% (Fisher Scientific Co. LLC, Pittsburgh, PA).

Laboratory strain Rs5 was grown in CPG medium as previously stated. The populations

were adjusted to A600= 0.3 by using a spectrophotometer at 600 nm to achieve a bacterial

concentration of approximately 108 cfu/ml. One hundred microliters of 108 cfu/ml in a

sterile deionized water solution were added to a dilution series ranging from 2 mg/l to 10









mg/1 of the given chemical. Dilutions were made from a stock solution of the chemical

diluted with sterile deionized water. Bacterial cells were exposed to the chemical for 1

min before being spread plated onto SMSA. Each dilution with the bacteria cell

suspension was plated on two petri plates, and the experiment was repeated three times.

The bacterial suspension at 108 cfu/ml was spread plated on SMSA to compare the effects

the chemicals on R. solanacearum survival. A percentage of remaining cells was

calculated by dividing the average colonies that survived after the chemical treatments

against the untreated control.

The protocol was modified to reduce error and to mimic what is found in the

environment. Previously the stock solutions of the chemicals were made by adding

minute amounts of the chemical, 166.3 Cl of chlorine and 3.33 pl of hydrogen peroxide

and were diluted in sterile tap water achieve a stock solution of 10 mg/l. Because small

amounts of the chemicals were used, pipette error was possible. Thus, the chemicals

were diluted with water at 1:10 for chlorine and 1:100 for H202 before making the 10

mg/l stock solution. Thus the chemical could be measured more precisely. The bacterial

suspension was diluted from 108 cfu/ml to 104 cfu/ml to simulate what may occur in the

ponds. For both methods untreated bacterial suspensions of either controls 108 cfu/ml to

104 cfu/ml were spread plated. The control plates were compared to those of the dilution

series to determine the percentage of recovered cells.

Sensitivity of modified SMSA for recovery of R. solanacearum. Laboratory

strain Rs5 was grown in CPG broth, and the suspension was adjusted to 108 cfu/ml

following previous procedures. A dilution series of the bacterium was made ranging

from 101 to 108 cfu/ml. The dilutions were spread plated on SMSA and on nutrient agar









(NA) (0.8% nutrient broth [BBL, Becton Dickinson and Co., Cockeysville, MD] and

1.5% Bacto Agar [Difco, Becton Dickinson and Co., Sparks, MD]). The plates were

incubated at 28 C for 60 h. Each dilution was spread plated on each of three plates, and

the experiment was repeated three times. The bacterial colonies from each plate were

counted and each dilution was averaged and the standard deviations were calculated.

Colonization R. solanacearum on Hydrocotyle ranunculoides. Two strains of R.

solanacearum, laboratory strain Rs5 and SEF were grown in CPG broth at 28 C. The

bacterial cells were adjusted to 108 cfu/ml according to procedures previously stated. The

two strains were also grown on SMSA incubated at 28 C for 60 h to achieve colony

growth. Bacteria grown on the plates were used for the toothpick method.

Hydrocotyle ranunculoides plants were collected from pond 2 and transplanted in

10-cm pots in Terra-Lite agricultural mix (Scott Sierra Horticultural Products Co.,

Marysville, OH). Plants were grown in greenhouse conditions for 3 months. Once a

week the plants were divided and propagated. Each week samples from the propagated

plants were taken to detect the presence of R. solanacearum. Plants used for the

experiment were a week old after propagation and then were inoculated strain Rs5 or

SEF.

The plants were inoculated by stabbing the crown of the stems with toothpicks and

also by drenching the root zone with bacterial suspension. Sterile toothpicks were coated

with the bacteria using either the lab strain Rs5 or SEF. Ten plants for each bacterial

strain were inoculated using the toothpick method as described previously. Control plants

were stabbed with sterile toothpicks in the same manner. Twenty additional plants were

inoculated by the drench method. These plants were removed from their pots and the









root system was broken up by hand before inoculation. A bacterial suspension consisting

of 50 ml of 108 cfu/ml was poured into each pot. Ten plants were inoculated with strain

Rs5 and another ten plants were inoculated with SEF. Every plant remained in its

individual plastic saucer. After inoculation the plants were stored in greenhouse

conditions. The saucer collected the water that was not taken up by the plant. The plants

were watered daily by pouring the collected water from each saucer back into the

corresponding plant; normal watering procedures were then followed.

The roots and stems of the plants were analyzed for colonization of bacteria

following the procedures outlined previously. A dilution series was spread plated on

modified SMSA. This was replicated three times. The plates were incubated at 28 'C for

60 h and then the number of colonies was counted.

Two colonies of the recovered R. solanacearum from both of the different

inoculation methods for both bacterial strains were selected for a pathogenicity test.

Tomato plants were used for the pathogenicity test. The tomato plants were inoculated

by toothpick method and observed for wilt symptoms. The pathogenicity test was

performed for each replication.














CHAPTER 4
RESULTS

Sensitivity of modified SMSA for recovery of R. solanacearum. A dilution

series of R. solanacearum ranging from 101 to 107 cfu/ml was spread plated on modified

SMSA and on nutrient agar. The lowest concentration at which colonies formed on

modified SMSA was at 103 cfu/ml. There was a 94% certainty that at least a single

colony would form at 103 cfu/ml. At this dilution 17 of the 18 SMSA plates had at least

one colony that formed. Indicating that 103 cfu/ml is the lowest concentration modified

SMSA can detect R. solanacearum suspended in water. Nutrient agar was able to recover

cells from every dilution as shown in Fig. 4.1.


300

SE 250 -
'4-
200 -
E 0 H Nutrient Agar
8 150 -

0g 1 -

0


1 2 3 4 5 6 7 8
Dilution series

Figure 4.1. Sensitivity of modified SMSA for detecting R. solanacearum was determined
by comparing a dilution series of known concentrations of the bacterium
spread plated on modified SMSA and nutrient agar. On many of the plates the
colony count was greater than 300. These plates were only recorded up to
300. Bars represent standard errors.









Monitoring of Ralstonia solanacearum in irrigation pond water. Samples were

collected for the detection of R. solanacearum in irrigation ponds of tomato farms and

ornamental nurseries in northern Florida from August 2003 to May 2005. Of the five

ponds, the bacterium was detected consistently in two ponds, once in two others, and

never in the fifth pond (Table 4.1). R. solanacearum was isolated from 13 of 78 samples

analyzed (16.6%) (Table 4.2).

Table 4.2. Incidence and bacterial concentration of R. solanacearum in water samples by
direct isolation on modified SMSA agar from August 2003 to May 2005
Number of positive samples/total number
of samples at temperatures greater than and Concentration
equal to 170C or less than 170C (cfu/ml) of
Pond >170C <170C bacterium a
1 5/13 0/13 4.2 x 10 3.9 x 10
2 5/20 1/20 2.9 x 102 5.6 x 104
3 1/13 0/13 2.4 x 102
4 1/20 0/20 3.7x 102
5 0/12 0/12 0
total 12/78 1/78
a Data are from analyses of water samples and show ranges of concentrations.

The bacterial concentration was directly related to the monthly average air

temperature at which the samples were taken (Table 4.2). The surface water temperature

ranged from 16.0 to 26.30C during the sampling periods when samples were positive for

the presence of R. solanacearum. For the majority of the samples that were positive for

the bacterium, the air temperature was above 230C. The detected concentration of the

pathogen ranged from 2.4x102 to 5.6 x 104 cfu/ml. The lowest temperature at which the

bacterium was recovered on modified SMSA was at 160C. As the temperature declined

the population level also declined. During the winter months, December through March,

when the average temperature high was less than 170C, the pathogen was undetectable

with the methods used in this study (Figures 4.2-4.6). From November 2004 to March









2005 the bacteria in each sample were concentrated by centrifugation and plated onto

modified SMSA. The pathogen was not detected in concentrated samples. All R.

solanacearum strains that were isolated from the water samples were pathogenic on

tomato plants. Wilt symptoms developed within 4 to 5 days, while R. solanacearum Rs5

caused wilt symptoms on tomatoes within 7 to 9 days under growth room conditions.

Statistical analysis of the data by Spearman's correlation coefficient (r,) showed a

significant positive correlation between the population levels of R. solanacearum and

temperature for ponds 1 and 2. A regression model revealed that temperature had an

effect on the density of R. solanacearum in the water (P < 0.001) (Table 4.3).

Table 4.3. Correlation between water temperature and levels of R. solanacearum for
ponds 1 and 2 from August 2003 to May 2005
Pond rsaP value
1 0.596 0.0034
2 0.626 0.0014
a Spearman's correlation coefficient.

Monitoring of R. solanacearum in aquatic weeds associated with the irrigation

pond water. A survey was performed on various aquatic weeds that grew in or in close

proximity to the irrigation ponds for the presence of R. solanacearum. At ponds where

the bacterium was found consistently (i.e., ponds 1 and 2) samples of similar weeds

found at both ponds were collected and analyzed. Weeds ranged from the grass

Tripsacumfloridiana to water surface weeds that included Alternantheraphiloxeroides

(common name alligator weed), Lemnaceae spp (common name duckweed), Hydrocotyle

ranunculoides (common name dollar weed), and Polygonum pennsylvanicum, and the

tree Perseapalustris (common name swamp bay) (Table 4.4).









Table 4.4. Detection of R. solanacearum isolated from sterilized or nonsterilized roots
and/or stems of semi-aquatic plants collected from ponds 1 and 2 by and
direct plating onto modified SMSA agar in July 2004
Plants species (number Detection of R. Bacteria isolated Surface sterilized or
of samples) solanacearuma from root or stem nonsterilizedab
Alternanthera
philoxeroides (2)
Lemnaceae spp (5)

Polygonum + both both
pennsylvanicum (1)
Polygonum + both Sterilized
pennsylvanicum (1)
Hydrocotyle + both Sterilized
ranunculoides (1)
Hydrocotyle + both both
ranunculoides (1)
Tripsacumfloridiana (2)

Persea palustris (1)

a The sample roots and stems from each plant was divided in two groups. One half of the plant material
were macerated in 1 ml sterile water and the other half was surface sterilized then macerated in 1 ml sterile
water. The macerates were streaked on SMSA medium and plates were incubated at 280C.
b R. solanacearum recovery from surface sterilized plant materials indicates latent infection. These plants
harbored the bacteria without wilt.

H. ranunculoides from the Apiaceae and P. pennsylvanicum from the Polygonaceae

families were the only weeds from which R. solanacearum was isolated (Table 4.4).

From July to October, R. solanacearum was detected by direct plating onto modified

SMSA in both H. ranunculoides and P. pennsylvanicum. Although the bacterium was

found on both the stem and root system of each plant, 56% of the positive results were

found in the roots of each plant. On average more colonies developed from the root

samples than from the stems. Sixty-one percent of the detected bacteria were from the

surface sterilized samples. In growth chamber assays, R. solanacearum strains from H.

ranunculoides and P. pennsylvanicum were pathogenic on tomato plants and wilt









symptoms were observed in 4 to 5 days (Table 4.5). Control strain Rs5, previously

isolated from tomato, caused wilt symptoms in 7 to 9 days.

Table 4.5. Incidence R. solanacearum in surface sterilized and nonsterilized stems and
roots of Polygonum pennsylvanicum and Hydrocotyle ranunculoides
associated with irrigation pond number and 2, from July 2004 to May 2005
Dates sampled Plants Detection of R. Bacteria isolated Surface sterilized
solanacearuma from root or stem or nonsterilizeda,b


July 2004


August 2004

September 2004



October 2004



November 2004



December 2004

January 2005

February 2005

March 2005

April 2005


May 2005


Polygonum
pennsylvanicum
Hydrocotyle
ranunculoides
Hydrocotyle
ranunculoides
Polygonum
pennsylvanicum
Hydrocotyle
ranunculoides
Polygonum
pennsylvanicum
Hydrocotyle
ranunculoides
Polygonum
pennsylvanicum
Hydrocotyle
ranunculoides
Hydrocotyle
ranunculoides
Hydrocotyle
ranunculoides
Hydrocotyle
ranunculoides
Hydrocotyle
ranunculoides
Hydrocotyle
ranunculoides
Hydrocotyle
ranunculoides
Polygonum
vennsvlvanicum


both

both

Both

Both

Roots

Both

Both

Roots

Roots

N/A

N/A

N/A

N/A

N/A

Both

Roots


both


both

Sterilized

both

Nonsterilized

Both

Both

Nonsterilized

Nonsterilized

N/A

N/A


N/A

N/A

N/A


Nonsterilized

Nonsterilized


a The sample roots and stems from each plant were divided in two groups. One half of the plant material
were macerated in 1 ml sterile water and the other half was surface sterilized then macerated in 1 ml sterile
water. The macerates were streaked on SMSA medium and plates were incubated at 280C.
b R. solanacearum recovery from surface sterilized plant materials indicates latent infection. These plants
harbored the bacteria without wilt.
' Samples were concentrated by centrifugation.









R. solanacearum was not detected during the winter months on either weed by

direct plating. P. pennsylvanicum was not found in the ponds from December to May.

The plant samples were concentrated during November 2004 through March 2005 by

centrifugation. In November, the bacterium was detected only on the root samples at 2.7

x 103 cfu/ml and 1.8 x 105 cfu/ml after centrifugation for P. pennsylvanicum and H.

ranunculoides, respectfully. Both root samples were untreated, while the untreated stem

and the surfaced sterilized samples were negative for detection of the bacterium as shown

in Table 4.5.

The R. solanacearum population decreased for ponds 1 and 2 during the month of

August. By this time growers had been spraying herbicides to control the aquatic weed

population and in particular H. ranunculoides and P. pennsylvanicum. By September the

weeds had recovered and were at the same density as in July. During the same time

period the bacterial population recovered and the population was the same as for July as

shown in Table 4.1.

Effects of chemical treatments on R. solanacearum. Two different methods of

chemical treatment gave similar results to a known concentration of R. solanacearum. In

first method a bacterial concentration of 108 cfu/ml was used. At this concentration of

bacteria the chlorine treatment resulted in a minimum inhibitory concentration (MIC) of

4.76 mg/l of free Cl2. At this concentration, 100% of the bacterial cells were completely

killed. The MIC for hydrogen peroxide at this bacterial concentration was not

determined (Figure 4.7). The bacterial suspension was exposed to 300 mg/l of hydrogen

peroxide (data not shown) and the chemical failed to inhibit colony growth.










100% {
80%
60%

40%
20%
0%


-- Chlorine
-- Hydrogen perioxide


mg/I

Figure 4.7. The effects of free chlorine and hydrogen peroxide at various concentrations
on 108 cfu/ml of Ralstonia solanacearum. After exposure to the chemicals,
the solution was spread plated on SMSA. The untreated bacterial suspension
was spread plated and served as the control. The chemical treatments were
compared to the untreated control to give the percentage of recovered cells


100% 1

80%

60%

40%

20%

0%


-*- Chlorine
-a- Hydrogen Perioxide


mg/I

Figure 4.8. The effects of free chlorine and hydrogen peroxide at various concentrations
on 104 cfu/ml of Ralstonia solanacearum. After exposure to the chemicals,
the suspension was spread plated on SMSA. The untreated bacterial
suspension was spread plated and served as the control. The chemical
treatments were compared to the untreated control to give the percentage of
recovered cells.

Samples from the collected irrigation water samples indicated that 104 cfu/ml

would be the highest population level bacterium would reach in the pond water. The









method was altered to resemble the same concentration found in ponds. At 104 cfu/ml,

no R. solanacearum colonies developed at 2 pl/1 of chlorine. Hydrogen peroxide again

failed to inhibit colony growth even at this low concentration of R. solanacearum (Figure

4.8).

Inoculation of H. ranunculoides with R. solanacearum. H. ranunculoides plants

were inoculated by two different methods and with two different strains of R.

solanacearum. Plants were inoculated by the toothpick or drench method. The strain

SEF was isolated from pond 1 in 2004, while Rs5 was isolated from tomato in 2000.

Colonization of the bacterium on this weed host has not been studied. Unexpectedly

every plant that was inoculated by Rs5 died. Those plants inoculated by the toothpick

method showed wilt symptoms within 5 to 6 days post inoculation. By day 8 all the

plants died. While the majority of the plants inoculated by the drench method showed

wilt symptoms by day 10, all the plants died fourteen days after inoculation. The

uninoculated control plants for both inoculation techniques of the Rs5 strain developed

wilt symptoms and died. However, none of the plants inoculated by SEF exhibited wilt

symptoms.










Table 4.1. Populations (cfu/ml) of R. solanacearum in five irrigation ponds from August 2003 to May 2005
Date
samples


taken
Pond Aug,
2003
la e


Sept,
2003
e


Oct,
2003
e


Nov,
2003
e


Dec,
2003
e


Jan,
2004
e


Feb,
2004
e


March, April,
2004 2004
e 4.2 x 102


-e 3.5 x102


3b NSd


5c NSd


NSd

e

NSd


NSd

e

NSd


NSd

e

NSd


NSd

e

NSd


NSd

e

NSd


NSd

e

NSd


a Samples from pond 1 were discontinued after September 2004.
b Pond 3 did not have water until April 2004.
' Samples from pond 5 did not commence until May 2004.
d NS stands for no sample was taken.
e indicates no bacterium was detected.


May,
2004
5.6x 103


June,
2004
3.2 x 103


NSd

e

NSd


NSd


I










Table 4.1 continued
Date
samples
taken
Pond July,
2004
la 3.9 x 104


2 3.6 x 103 6.8 x 103 9.3 x 103


5.6 x 104 2.9 x 102


e 2.4 x102

e 3.7 x102


a Samples from pond 1 were discontinued after August 2004.
b Pond 3 did not have water until April 2004.
' Samples from pond 5 did not commence until May 2004.
d NS stands for no sample was taken.
e indicates no bacterium was detected.


Aug,
2004
3.9x 103


Sept,
2004
NSd


Oct,
2004
NSd


Nov,
2004
NSd


Dec,
2004
NSd


Jan,
2005
NSd


Feb,
2005
NSd


March,
2005
NSd


April,
2005
NSd


May,
2005
NSd


e e

e e

e e

e e


e e

e e

e e

e e














30 5
4.5
25 4
4 3:
3.5 o
20 2
T3 2
S1 Temperature
1 15 2.5
S2 -- Bacterial
E 10 1 Concentration
1.5


0.5
0 0
Aug-03 Sep-03 Oct-03 Nov-03 Dec-03 Jan-04 Feb-04 Mar-04 Apr-04 May-04 Jun-04 Jul-04 Aug-04

Figure 4.2. Detection of R. solanacearum colonies by direct isolation on modified SMSA medium during a survey of Pond 1 from
August 2003 to August 2004. Surface water samples were collected at five different locations along the banks of the pond.
The dates indicate the sampling months during the years 2003 through 2004. The populations of the bacterium (from 0 to
3.9 x 104 cfu/ml) and the temperatures at the time the samples were taken are indicated in the figure.













5
4.5
4
3.5 2
3 .2
2.5 _
2
1.5 t
1 '
0.5 3


Temperature

-*- Bacerial
Concentration


Aug- Sep- Oct- Nov- Dec- Jan- Feb- Mar- Apr- May- Jun- Jul- Aug- Sep- Oct- Nov- Dec- Jan- Feb- Mar- Apr- May-
03 03 03 03 03 04 04 04 04 04 04 04 04 04 04 04 04 05 05 05 05 05


Figure 4.3. Detection of R. solanacearum colonies by direct isolation on modified SMSA medium during a survey of Pond 2 from
August 2003 to May 2005. Surface water samples were collected at five different locations along the banks of the pond.
The dates indicate the sampling months during the years 2003 through 2005. The populations of the bacterium (from 0 to
5.6 x 104 cfu/ml) and the temperatures at the time the samples were taken are indicated in the figure


30

25

S-20

1 15
C
E 10

5

0















25 4

3.5 o
20
WII I l3 _
2 Temperature
15 2.5
wE
0 2 -4- Bacerial
E 10-
S1.5 L Concentration
5- 0
5 1
1 10.5 -j
0 -- 0
Apr-04 May-04 Jun-04 Jul-04 Aug-04 Sep-04 Oct-04 Nov-04 Dec-04 Jan-05 Feb-05 Mar-05 Apr-05 May-05

Figure 4.4. Detection of R. solanacearum colonies by direct isolation on modified SMSA medium during a survey of Pond 3 from
April 2004 to May 2005. Surface water samples were collected at five different locations along the banks of the pond. The
dates indicate the sampling months during the years 2004 through 2005. The populations of the bacterium (from 0 to 2.4 x
102 cfu/ml) and the temperatures at the time the samples were taken are indicated in the figure.













5
4.5
4 $
3.5 |
3 .2
2.5
2
1.5 5
10
0.5 3


Temperature

--- Bacterial
Concentration


Aug- Sep- Oct- Nov- Dec- Jan- Feb- Mar- Apr- May- Jun- Jul- Aug- Sep- Oct- Nov- Dec- Jan- Feb- Mar- Apr- May-
03 03 03 03 03 04 04 04 04 04 04 04 04 04 04 04 04 05 05 05 05 05

Figure 4.5. Detection of R. solanacearum colonies by direct isolation on modified SMSA medium during a survey of Pond 4 from
August 2003 to May 2005. Surface water samples were collected at five different locations along the banks of the pond.
The dates indicate the sampling months during the years 2003 through 2005. The populations of the bacterium (from 0 to
3.7 x 102 cfu/ml) and the temperatures at the time the samples were taken are indicated in the figure.


30

25

- 20

1i 15

E 10

5
5















30 5
4.5 2
25-
4
3.5
20
w3 .2
May 2004 to May 2005. Surface water samples were collected at five different locations along the banks of Temperature
wE
dates 2 the bacteria Bacterial
1.5detected from this pond and (0 )the temperatures at the time the samples were taken are indicated Concentration

0.5 3
0- 0
May-04 Jun-04 Jul-04 Aug-04 Sep-04 Oct-04 Nov-04 Dec-04 Jan-05 Feb-05 Mar-05 Apr-05 May-05

Figure 4.6. Detection of R. solanacearum colonies by direct isolation on modified SMSA medium during a survey of Pond 5 from
May 2004 to May 2005. Surface water samples were collected at five different locations along the banks of the pond. The
dates indicate the sampling months during the years 2004 through 2005. The populations of the bacterium were never
detected from this pond and (0 cfu/ml )the temperatures at the time the samples were taken are indicated in the figure.














CHAPTER 5
DISCUSSION AND CONCLUSIONS

An investigation of the population densities of R. solanacearum over a two and a

half year period of analyses at five different irrigation ponds on tomato farms and

ornamental nurseries in northern Florida associated with bacterial wilt revealed that R.

solanacearum was detected in pond water and in two weed species (H. ranunculoides

from the Apiaceae and P. pennsylvanicum from the Polygonaceae families) associated

with the pond environment.

There was a correlation between detection of the pathogen and the recorded air

temperature, confirming results reported in the United Kingdom (Elphinstone et al. 1998)

and Spain (Caruso et al, 2005). Based on analyses performed in 2004, the first detection

of R. solanacearum was in April, when the temperature was above 170C. The abundance

of this bacterium was relatively higher (from 3.5 x 102 to 5.6 x 104 cfu/ml) from April to

November, and then decreased dramatically until it was undetectable in December when

temperatures were below 170C. The majority of reports of incidences of the disease in

northern Florida have been during the fall harvest season, which correlates with density

of the pathogen's population in the pond environment. These results agree with previous

studies in Spain, the United Kingdom, and the Netherlands, where the levels of R.

solanacearum r3 b 2 in waterways were usually 80 cells/ml, dropping below the detection

limit when exposed to cold temperatures (Caruso et al, 2005; Elphinstone, 1998; Janse,

1998).









Bacterial populations varied from pond to pond. Descriptions of the five ponds are

as follows;

Pond 1 was a retention pond for an ornamental nursery. The water's source

originated from the run off of the greenhouses and rain. The water was re-circulated into

the greenhouse after going through a filtering process. R. solanacearum was detected as

early as April 2004. This pond contained the highest concentrations of the bacterium,

until sampling was discontinued. In June 2004, the producer removed all of the aquatic

weeds and other weeds that surrounded the banks, and the population of R. solanacearum

was reduced from 103 to 102 cfu/ml. However, by the next month the weeds grew back

and population levels were as high as 3.9 x 104 cfu/ml. Plants from this pond were

analyzed for colonization of R. solanacearum. The only plants that tested positive were

H. ranunculoides and P. pennsylvanicum. This pond was not shaded by any trees, and

vegetation growth was very abundant in and around the pond. The pond had an aeration

system that kept the water in constant motion.

Pond 2 was located in the middle of the tomato production fields. The source of

the pond was tributary creeks that feed the pond. Samples were taken only on the south

side of the pond due to the lack of access to the other sides. R. solanacearum was only

found where H. ranunculoides and P. pennsylvanicum grew, which was only the

southeast side of the pond. In June 2004, the grower sprayed an herbicide that killed

most of the aquatic weeds. As a result populations were reduced, yet by July 2004 the

plants grew back and bacterial populations were as high as 103 cfu/ml. Pond 2 had tree

coverage along the sides of the pond, but the trees did not provide much shade near the

banks of the pond. Much vegetation grew in this pond along the banks of the pond.









Pond 3 was a retention pond that was dry during the winter of 2003, but during

2004 was filled with water. Water was irrigated from this pond for the tomato farm

across the street. None of the common aquatic weeds found at the other ponds were

observed at this pond. R. solanacearum had been detected only once in sample taken in

October 2004. Bacteria wilt of tomato had been a problem in the past for this particular

farm; however, there were no reports of the disease during the tomato growing seasons in

2004.

Pond 4 was located near Pond 3 and was surrounded by wooded hills. Pond 4 was

the largest pond and was fed by a few local streams. Like pond 3, detection of R.

solanacearum was only in October 2004. H. ranunculoides and P. pennsylvanicum were

not observed at the areas where samples were taken. Bank areas of this pond were not

heavily vegetated and sunlight was restricted due to the amount of trees in the area.

Pond 5 was located at the far eastside of the tomato fields. At the far southwest

corner of the field a few tomato plants had bacterial wilt. R. solanacearum was isolated

from these plants. Water samples have been taken from this retention pond since May

2004; however, R. solanacearum was not isolated from the water. The surface of this

pond was completely covered by duckweed (Lemnaceae spp), and was completely

shaded by the surrounding trees.

Further investigations would need to take place to understand why the pathogen

was so prominent in ponds 1 and 2, yet not in 3-5. All five ponds had reports of bacterial

wilt on each property and in all the ponds besides pond 5 R. solanacearum was detected.

The main difference of each pond was the population of semi-aquatic weeds. In ponds 1

and 2, both H. ranunculoides and P. pennsylvanicum grew abundantly, while in the other









ponds neither weed grew. Since both plants grow best in sunny conditions, a possible

explanation for the lack of growth of the weeds could be due to the amount of sunlight

that reaches these ponds. Understanding the difference between R. solanacearum

positive ponds and negative ponds will aid us in controlling the pathogen.

After a survey of several common semi-aquatic and other weeds, H. ranunculoides

and P. pennsylvanicum were positive for the bacterium both in and on the surface of the

stems and roots by direct plating an extraction of plant samples and water on SMSA. The

bacterium was not detected by direct plating during the winter months when the

temperature was below 170C. However, when the plant samples were concentrated by

centrifugation, the bacterium was detected. To further understand the connection

between these symptomless weed hosts and the bacterium further methods must be

considered. Comparing both the plant sampling from the ponds and from the greenhouse

experiment, it is assumed that the bacterium can colonize the surface of roots or stems, as

well as colonize them internally. To confirm where colonization takes place, H.

ranunculoides and P. pennsylvanicum would be inoculated with an R. solanacearum

strain containing the green fluorescent gene (GFP). The bacterium could then be seen

using a confocal microscope and the exact location of the bacterium could be determined.

Confirmation of colonization of the bacterium on these weeds would benefit the growers

to know which weeds to eradicate in order to help control to the pathogen population.

The results of the chlorine and hydrogen peroxide indicate that the bacterium can

be controlled by chlorine, however the hydrogen peroxide was ineffective. At 4.76 mg/l

and 1.9 mg/l of free Cl2, R. solanacearum can be eliminated at 108 cfu/ml, and 104 cfu/ml

respectfully, however, both experiments were performed when the water and bacterial









suspensions were stagnant. Chlorine would most likely be added to the water when or

after it passes through the water pumping station. Water in this stage would be moving

very rapidly, so tests would further need to be performed to find the correct MIC value

for the water in motion. However, using chlorine as a means to control the population of

the bacterium, one would need to be concerned by the negative effects of chlorine in the

environment, the effects of a build-up of chloride in the soil, and its effects on tomato

plants. Hydrogen peroxide was ineffective for each concentration of bacterial cells in

this study. Hydrogen peroxide usually decomposes in the presence of numerous catalysts

such as most metals, acids, or oxidizable organic materials. A small amount of stabilizer,

usually acetanilide, is often added to it. Thus, without a stabilizer hydrogen peroxide

would be inefficient, due to the amount of catalysts found in the pond water. Market

products with the active ingredient being hydrogen peroxide were considered; however,

after speaking with a representative from one of the manufacturers, he calculated that the

volume needed to control the pathogen in irrigation water would result in expense of

about $2,000 for 60 mg/l per acre foot of water. Between 50-60 mg/l is average for

bacterial control for use of this product.

SMSA can detect the presence of R. solanacearum suspended in water 94% of the

time at 103 cfu/ml. For the purpose of the population study of this bacterium in the water,

this concentration was sufficient, yet other methods could detect the lower concentration

of the bacterium in low densities. Such method as enriching plus PCR and ELISA could

be very effective in detecting the presence of the bacterium, but these techniques would

require training, time and money.









R. solanacearum was found to colonize H. ranunculoides by both inoculation

methods. Results are being processed to determine where the bacterium colonizes on the

plants. All the H. ranunculoides plants inoculated by the toothpick method with the

laboratory strain Rs5 developed wilt symptoms within 5 to 6 days post inoculation.

While those inoculated with the same strain using the drench method developed wilt

symptoms 10 days after inoculation, the majority of the plants inoculated with Rs5 were

unable to regenerate and no new growth was seen. None of the plants inoculated with

SEF developed wilt symptoms. Rs5 was isolated from a tomato plant in November 2000,

and SEF was isolated from a water sample from pond 1 in July 2004. It is assumed that

the over the years bacterium has evolved to one which is able to coexist with the semi-

aquatic weeds. Molecular analysis of the two different strains would need to be done to

determine the differences.

Contaminated irrigation water is a potential source of infection in north Florida and

some of the semi-aquatic weeds act as a secondary host in which R. solanacearum could

overwinter and multiply to further contaminate irrigation ponds. Tomato, geranium and

hydrangea growers need to take into consideration these results, and use management

tactics to avoid this source of inoculum.
















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BIOGRAPHICAL SKETCH

Jason Hong was born in Logan, Utah, in 1978, but grew up in Wooster, Ohio. For

the last two years of high school he was dual enrolled at the local community college and

graduated with an Associate of Science the same year he graduated from high school. In

1997, he served a two year mission for The Church of Jesus Christ of Later-Day Saints in

Santiago, Chile. In 2003, he graduated from The Ohio State University with a degree of

Bachelor of Science in microbiology. During the summers of 2002 and 2003 he had

internships with Dr. Jackson at AgriPhi (now Omnilytics) and was exposed to plant

pathology, bacteriophage and research. He attended the graduate program of the

University of Florida, College of Agricultural and Life Sciences, Department of Plant

Pathology, from August 2003 to August 2005. He conducted a research project to

monitor the population of Ralstonia solanacearum in irrigation pond water, aquatic

weeds, and developed methods of control of the bacterium in the water under the

guidance of Drs. Timur M. Momol, Jeffery B. Jones, Steve M. Olson, and Jerry A. Bartz.