In this book, the current status of various aspects of integrated disease management in fruits, vegetable, ornamentals, cereals, pulses, oilseeds, medicinal and forest plants etc.
Major focus is on the integrated disease management in horticultural crops. Emphasis has been given to the use of non-chemical methods like cultural practices, soil solarization, plant growth promoting microorganisms, organic amendments, botanicals and biocontrol agents.
It is hoped that the book will serve as an important guide to the plant pathologists, horticulturists, nematologists, microbiologists, mushroom scientists, breeders and students. This manual describes in detail a variety of protocols for determining the properties and identity of a virus and its behavior in infected plants.
A Springer Lab Manual. New additions to the second edition include five new topic and exercise chapters on soilborne pathogens, molecular tools, biocontrol, and plant-fungal interactions, information on in vitro pathology, an appendix on plant pathology careers, and how to use and care for the microscope.
An accompanying cd-rom contains figures from the text as well as supplemental full-color photos and PowerPoint slides. Unique Learning Tools Retaining the informal style of the previous edition, this volume begins each topic with a concept box to highlight important ideas. Several laboratory exercises support each topic and cater to a wide range of skill sets from basic to complex.
Procedure boxes for the experimental exercises give detailed outlines and comments on the experiments, step by step instruction, anticipated results, and thought provoking questions. Case studies of specific diseases and processes are presented as a bulleted list supplying essential information at a glance.
Comprehensive Coverage Divided into six primary parts, this valuable reference introduces basic concepts of plant pathology with historical perspectives, fundamental ideas of disease, and disease relationships with the environment. It details various disease-causing organisms including viruses, prokaryotic organisms, plant parasitic nematodes, fungi, plant parasitic seed plants, and other biotic and abiotic diseases.
Exploring various plant-pathogen interactions including treatments of molecular attack strategies, extracellular enzymes, host defenses, and disruption of plant function, the book presents the basic ideas of epidemiology, control strategies, and disease diagnosis.
Pyricularia oryzae Rhynchosporium secalis Rosellinia necatrix Sclerotinia spp Sclerotium spp Scolicotrichum graminis Seiridium spp Selenophoma donacis Septoria spp Sordaria spp Sphaeropsis ulmicola Stagonospora spp Stemphylium spp Stigmina carpophila Thanatephorus cucumeris Thielaviopsis basicola Typhula idahoensis Ustilaginoidea virens Venturia inaequalis Verticillium spp Bacterial Pathogens Periodic Transfer Mineral Oil Storage in Water Storage in Silica Gel Storage in Soil Preservation in Liquid Nitrogen Lyophilization and Vacuum Drying Soilborne Pathogens Aspergillus flavus Hymenula cerealis Laetisaria arvalis Mucor piriformis Ophiostoma wageneri Phialophora gregata Plasmodiophora brassicae Phoma terrestris Sclerotinia minor Spongospora subterranea Talaromyces flavus I 00 Detection and Estimation of Inoculum in the Air Glass Slides and Cylindrical Rods Volumetric Spore Traps Schenck's Sampler Panzer's Spore Sampler Grainger's Trap Davis and Sechler's Sampler Miniature Spore Trap Manually Operated Portable Spore Trap Czabator and: Scott's Sampler The Morris Spore Trap Slit Samplers Cyclone Collector High Throughput Jet Trap Muslin Cloth Spore Trap Biological Spore Detectors Detection of Pathogens Associated with Seeds Examination of Dry Seeds Examination of Seed Washings Blotter Test Agar Test Seedling Symptom Test Use of Light in Detecting Seedbome Fungi Evaulation of Routine Seed Testing Methods Plant Inoculation Test Detection of Bacterial Pathogens Morton's Method Pope's Method Kavanagh and Mumford Method Edinburgh Method Factors Influencing Disease Establishment Root Inoculation Soil Infestation Hydroponic Method Root-Dip Method Cut and Dip Method Root-Stabbing Method Inoculation of Large Woody Roots Root Inoculation Without Disturbing the Roots In Vitro Methods The Slant-Board Technique Seed Inoculation Other Methods Stem Inoculation Leaf Inoculation Quantitative Inoculations Inoculation of Cereal Inflorescences Inoculation with Bacteria Inoculation of Detached Plant Parts Field Inoculation Methods Some Special Hints Inoculation with Toothpicks Seedling Box Homemade Atomizer Isolation and Enumeration of Soil Organisms Dilution Plate Soil Plate Immersion Tubes, Plates and Slides Paper Strip Baits Isolation of Fungal Hyphae Soil Washing Isolations from the Rhizosphere Isolation of Specific Organism Groups Cellulolytic Fungi Paraffinolytic Fungi Observation of Soil Microorganisms In Situ Direct Microscopy Vital Staining and Auorescent Microscopy Staining with Acridine Orange Staining with Auorescein Diacetate Staining with Mg-ANS Staining with Europium Chelate and Fluorescent Brightener Staining with Auorescein Isothiocyanate Staining with Rose Bengal Staining with Ethidium Bromide Infra-red Photography Soil Sectioning Resin Embedding Gelatin Embedding Buried and Immersion Slides Membrane Filter Partial Purification of Soil Bacteria and Fungi Observation of Rhizosphere Microorganisms Physiological Processes as an Estimate of Biological Acitivi.
Dehydrogenase Assay Urease Assay Phosphatase Assay Extraction of ATP from Soil.. Soil Biomass Determination by Soil Fumigation Spore Germination in Soil and Soil Fungistasis Indirect Methods Agar Disk Cellophane Film Membrane Filters Agar Slide Technique Direct Methods Survival of Plant Pathogens in Soil Mycelial Growth in Soil.
Collection of Root and Seed Exudates Collection of Root Exudates Determination of Root Exudation Zone Collection of Seed Exudates Collection of Spore Exudates Artificial Rhizosphere or Spermosphere Spore Germination Test Glass Slides Depression Slides Dry-Drop Technique Agar Film Method Spore Germination Test in Shaker Flasks Tests Based on Growth of Organisms in Vitro Paper Disk Plate Method Glass Slide Method Transfer-Bioassay Techniques Filter Paper Transfer Bioassay Cellophane-Transfer Bioassay Laboratory Testing for Soil Fungicides Greenhouse Testing of Foliar Fungicides Planning Field Experiments Seed Treatment with Chemicals Collection and Testing of Soil Samples for Antagonists Soil Tube and Culture Plates Tests Based on Pathogen Infectivity Isolation of Antibiotic-Producing Organisms Testing Antibiotic Production in Culture Antibiotics in Culture Filtrates Cell-Free Culture Filtrates Filter Paper Disk Method Well-in-Agar Method Assay in Liquid Medium Antibiotic Production in Soil Isolation and Testing of Lytic Organisms Isolation and Testing of Mycoparasitic Organisms Spore Perforating Amoebae Isolation of Antagonists from Leaves Testing Antagonists for Biological Control Glass or Plastic Tubes Seedling Bioassay Chamber Seed Treatment Testing Antagonists for Pathogen Survival in Soil Mounting and Staining of Fungi for Microscopic Examiniation Glycerin Jelly Shear's or Patterson's Fluid Hoyer's Medium Polyvinyl Alcohol Mountant Wittmann's Direct Mounting Medium Glycerin Jelly-Methyl Green Orseillin in BB and Crystal Violet Lactophenol-Acid Fuchsin Lactophenol-Cotton Blue Lactophenol-Aniline Blue-Acid Fuchsin Phenol-Dye Mixtures Tannic Acid-Basic Fuchsin Trypan Blue Isaac's Staining Mountant Malachite Green-Acid Fuchsin Stain Staining Fungi in Agar Medium Vital Staining Aniline Blue Geimsa-HCl Stain Iron-Acetocarmine Stain Propiono-Carmine Stain Safranin Toluidine Blue Fluorescent Staining Acridine Orange Mithramycin Areolic Acid Hoechst Auramine-0 and Acrinol Gram's Stain Staining of Flagella Leifson's Stain Silver Impregnation Rhodes' Stain Gray's Stain Cell Wall Staining Staining of Spores of Streptomycotina Fungus Cultivating Making Permanent Mounts Agar-Colony Mounts Dried Reference Cultures Staining of Freehand Sections Staining of Paraffin Sections Pianeze Illb Margolena Stain Methylene Blue-Erythrosin Stain Periodic 'ACid-Schiff Reagent Safranin-0 and Fast Green Stain Silver Nitrate-Bromophenol Stain Flemming's Triple Stain Conant's Quadruple Stain Iron Hematoxylin Harris' Hematoxylin Differential Staining of Wood Tissues Cartwright's Method Gram and Jorgensen's Method Pearce's Method Staining of Bacteria in Host Tissues Polychromatic Staining with Toluidine Blue Staining with Iodine Fumes Preparation of Whole Mounts Chloral Hydrate Fluorescent Staining of Whole Mounts Staining of Vascular System in Whole Mounts Making Plastic Prints of Leaf Surfaces Cultivation and Sporulation Media Media for Isolation of Actinomycotina from Soil..
Media for Isolation of Bacteria from Soil Media for Isolation of Fungi from Soil All require sterile conditions. Sterilization of apparatus and working areas involves the inactivation or physical elimination of all living cells and infective agents from the environment. It does not include the destruction or elimination of constitutive enzymes, metabolic by-products, or removal of dead cells. Sterilization is achieved by exposing materials to lethal agents which may be chemical, physical, or ionic in nature or, in the case of liquids, physical elimination of cells or infective agents from the medium.
Selection of a method depends on the desired efficiency, its applicability, toxicity, ease of use, availability and cost, and effect on the properties of the object to be sterilized.
Several publications review the theoretical aspects of sterilization methods. High temperature can be attained by using either dry or moist heat. The mechanisms of cell destruction by heat were reviewed. The action of dry heat is an oxidation process resulting from heat conduction from the contaminated object and not from the hot air surrounding it.
Thus, the entire object must be heated to a temperature for a sufficient length of time to destroy contaminants. Dry heat requires higher temperatures for longer duration than moist heat for sterilization because heat conduction by the former is slower than the latter.
Many bacteria in a desiccated vegetative state or as spores can survive dry heat at high temperatures. Hot air ovens equipped with a thermostat and heated either by electricity or burning gas are used for dry heat sterilization. To determine whether or not a uniform temperature occurs throughout the load, thermometers should be placed at different sites within the chamber.
The time required for sterilization is inversely proportional to temperature. The air in the oven heats faster than the objects to be sterilized; therefore, the duration of the heat treatment should be increased by 1.
Objects, such as glass culture plates, should be placed in sealable metal or other heat-resistant containers to prevent recontamination during cooling, transport, or storage. The objects can be wrapped in heavy paper if metallic containers are not available. However, paper may leave organic residue and become brittle and charred. Calibrated glassware should not be sterilized with dry heat since the expansion and contraction can cause changes in the graduations.
Objects with tight-fitting joints or plugs should be separated during hot air sterilization; otherwise, they may break. Sterilization chambers whether using dry or wet heat, should be loaded in such a way as to provide ample space between items allowing for air circulation and to avoid breakage. Containers plugged with cotton, plastic, or rubber stoppers should be sterilized at lower temperatures for longer times.
Slip-on metal caps can be substituted for cotton for culture tubes. Rubber-stoppered or screw-cap bottles, flasks, or culture tubes should be sterilized with moist heat. After sterilization, the oven and its contents should be allowed to reach ambient temperature before opening to prevent breakage and recontamination by cool air rushing into the chamber.
Sterilized material may remain in the oven until used or stored in a dry area free of air currents, but should be used within a short time and not stored for long periods.
MOIST HEAT Moist heat is usually provided by saturated steam under pressure in an autoclave or pressure cooker, and is the most reliable method of sterilization for most materials.
It is not suitable for materials damaged by moisture or high temperature, or culture media containing compounds hydrolyzed or reactive with other ingredients at high temperature. Moist heat has advantages over dry heat in that conduction is rapid and the temperature required for sterilization is lower and the duration of exposure is shorter.
Materials to be sterilized should be in contact with the saturated steam for the recommended time and temperature. The process is usually carried out in an autoclave7 or a kitchen-type pressure cooker equipped with pressure gauges, thermometer, automatic pressure control valves, and exhaust valves.
Autoclaves may be nonjacketed Figure or jacketed Figure In jacketed types, the duration for heating is less than in the nonjacketed types, moisture does not condense on objects, and the steam is "dry", i. Steam is supplied either from a central source or is generated within the autoclave or pressure cooker by electric or gas heating. Pressure cookers and autoclaves are available in a variety of sizes and models follow operationing instructions provided by the manufacturer.
The temperature and length of time for sterilization with steam are different from that of dry heat. Thiel et al. From Cruickshank, A. Livingston, Edinburgh, With permission. These temperatures are attained at 1. All of the air must be removed from within the chamber before closing the exhaust valve.
The effect of air removal on temperature is summarized Table 2. Sterilization begins after the load has reached the desired temperature.
The preheating times required for various liquid volumes are 2 min for loosely packed culture tubes containing 10 ml; 5 min if tightly packed; 5 min for flasks or bottles containing ml plugged with cotton or loosely screwed caps loosely packed; I 0 to 15 min for ml, 15 to 20 min for 1 l; and 20 to 25 min for 2 I.
If flasks or bottles are stacked or layered, the preheating time should be increased 5 to 10 min. It is not desirable to autoclave large and small volumes at the same time because of the different preheating and autoclaving times. The effect may be harmful or beneficial, but no medium should be exposed to more heat than necessary. The medium pH usually is changed by 0. Glucose and amino acids may react to form compounds inhibitory to microorganisms.
Additives, such as antibiotics, hormones, vitamins, and other compounds may be destroyed by heating and therefore should be sterilized by filtration or other means and added after autoclaving the medium.
Remember that when liquids are mixed a dilution factor must be considered. Schematic diagram of a steam jacketed autoclave with automatic gravity discharge of air and condensate, and system for drying by vacuum and intake of filtered air: A, chamber discharge and vacuum valve; B, venturi tube; C, steam to chamber valve; D, chamber pressure gauge; E, jacket safety valve and pressure gauge; F, air intake and filter; G, chamber door, H, thermometer; I, pressure regulator; J, steam supply valve; K, drain; L, vapor trap, M, chamber steam trap; N, jacket steam trap; 0, perforated tray; P, baffle; Q, discharge channel; R, discharge to atmosphere.
Livingstone, Edinburgh, Table 1. All vents, exhaust values, and safety valves as well as the chamber should be kept clean. Use nonabsorbent cotton for plugs, which should be loose enough to allow for access of steam and air exhaust during decompression. If screw-cap containers are used, treatment time may have to be increased 5 min over cotton-plugged containers.
Always check the effect of heat on an object or material to be autoclaved before beginning the process. Containers should be no more than one- half to two-thirds full. All air should be replaced before closing the exhaust valve. Exhaust of steam after autoclaving should be slow to prevent blowing of stoppers and boiling of liquids.
Tempered metal can be heated to "red hot" and remains sterile as long as it is hot. Glass objects are passed through the flame several times and should not be placed immediately on a cool surface or they will crack. Advantages are that the process can be carried out at low temperatures and relative humidity; objects can be sterilized in their containers since most gases will diffuse out of most containers with time; the process can be carried out using simple equipment such as plastic or rubber bags, or metal or plastic drums.
Major disadvantages are that a longer time is required for sterilization over that of heat, materials used are flammable and highly toxic, and the cost is higher than heat. If the gas is highly reactive, it may combine with organic matter. Some gases used are ethylene oxide, formaldehyde, propylene oxide, methyl bromide, ozone, and Bpropiolactone.
The first three are alkylating agents. The mechanism of gas sterilization was summarized. However, it is highly explosive when mixed with air, toxic at low concentrations, and a direct-contact skin irritant. Flammability is eliminated when mixed with inert gases.
This mixture is available in metal cylinders containing 15 to 30 kg of gas as a liquid, in to g cans to be used with needle valves and a can holder, or in glass bottles. When in glass, the mixture must be stored refrigerated since the boiling point of ethylene oxide is l0. Sterilization requires a minimum of 3 hr; the time at a fixed temperature is inversely proportional to gas concentration.
Hygroscopic compounds cannot be sterilized with ethylene oxide. However, it can be used to sterilize aqueous solutions. C may change its composition by reaction with certain compounds. It is used generally as a surface sterilant although a thin film of organic matter can restrict its activity. It is less effective than ethylene oxide and has less penetrating power. For laboratory studies sclerotia produced in onion bulbs simulate field conditions. Select pathogen-free mature onions bulbs.
Trim the basal plate of dry roots, taking care not to remove the plate. Remove the outer dry scales. Using a sterile scalpel cut four flaps in the fleshy scales and place a 5-mm disk from an actively growing culture on medium 81 beneath each flap and seal with tape.
Remove the tape and reincubate for 4 days or when white mycelium is visible around the cuts in the bulb. Keep 'the mixture moist but not waterlogged. Harvest sclerotia by w'et sieving, when they are mature, generally after 6 to 8 wk. Place the sclerotia in polyester fabric bags and bury in field soil until required.
Place the mixture in glass jars or Erlenmeyer flasks, autoclave for 90 min, then seed with a PDA culture disk and incubate for 6 wk at room temperature. Air dry it in the laboratory for 48 hr. Separate the sclerotia by screening through two layers of cheesecloth. In both cases collect sclerotia by wet sieving. The sclerotia produced in soil are physiologically and structurally different from those produced on agar media. Synchronous formation of sclerotia can be induced by 1 removal of aerial mycelium with a scalpel; 2 growing the fungus under a cover glass which is later removed; or 3 pouring a layer of agar over the colony.
In all cases hyphae that emerge from the submerged phase produce sclerotia synchronously. In liquid cultures, sclerotia production is induced by pouring a to hr shake culture into a culture plate and incubating. Seiridium spp. Dis infested seeds inoculated with a conidial suspension in 0.
Septaria spp. Sporulation inS. However, on medium 50 seeded in the similar manner, the quantity of spores increases. Placing a sterile filter paper disk over medium 50 and seeding by the same method further increases spore production. Sordaria spp. Perithecia of S. However, distributing sterile wheat kernels on PDA increases growth rate and spore production. Colonized kernels provide a source of inoculum for greenhouse and field inoculations.
Interaction between the medium and temperature may be significant. Stagnospora spp. For S. To harvest conidia, flood the plates with 30 ml of water and scrape to remove air bubbles; in 30 min conidia are discharged in the water. A film of water on the medium helps smearing. If conidia of S. Limited but sufficient sporulation occurs. Crush pycnidia in sterile water and spread the suspension over medium 94 in culture plates or test tubes.
Subculturing must be by spores. Stemphylium spp. Grow on medium ; pigmentation does not occur. Medium is useful for sporulation of S. Thanatephorus cucumeris Grow on a mixture of 1 g rice straws each 2. Thielaviopsis basicola For chlamydospore production, grow on medium 38 or for 3 to 4 wk, then flood the culture with distilled water and scrape the surface with a scalpel. Combine and mix the suspension in a high speed blender and filter through a jlm screen under suction. Collect chlamydospores from the screen.
Grow the fungus on PDA slants for 8 to 10 wk, wash several times with tap water to remove endoconidia, then scrape the agar surface with a scalpel to remove chlamydospores, and pour into a jlm sieve.
Wash the contents of the sieve under running tap water long enough to remove any endoconidia and mycelial fragments. Grow the fungus on agar slants for 3 to 4 wk, wash to remove endoconidia, and homogenize at high speed in a blender for 1 to 2 min. The suspension is freed from agar by repeated washing and slow-speed centrifugation.
Chlamydospores are separated from endoconidia and mycelial fragments by sieving as described previously or resuspending the pellet in a small amount of water, pouring into large culture plates and drying overnight. Chlamydospore chains project upward and can be removed with a fine brush.
Homogenize the culture mats in ml of sterile water at high speed for 3 min. Atomize the suspension into 4. Fungal population is checked by plating on a selective medium. Prepare the medium by mixing ml of wheat grain with ml of hot water and autoclave for 50 min. Shake the medium periodically for even distribution of the fungus and to prevent caking. Sclerotia are mature in about 30 days. Ustilaginoidea virens Cover the surface of medium in culture plates with a sterile butter-paper disk and.
Place a portion of a surface sterilized smut ball in the center. To produce smut balls in culture, place several surface-sterilized rice flowers, collected just before bursting, together with chlamydospores on medium and incubate under the same conditions. Venturia inaequalis Grow it on medium 4 for perithecia formation.
Perithecia are induced ori medium if amended with apple-leaf decoction from medium 4. To prepare a decoction, boil 20 g of plant material in ml water, filter through cheesecloth and add the filtrate to ml of the medium Autoclave for I hr in 5 ml of glass distilled water, then place in test tubes containing l ml of medium 5 and autoclave twice for 45 min at a hr interval.
Conidia are removed with a soft brush and collected in water after 18 to 20 days. Other methods used are to cut filter paper in a rectangle and roll into a cylinder held in place with stainless steel clips. Place the cylinder in a culture tube and heat sterilize. Seed the paper with a conidial or mycelium suspension prepared in sterile glass distilled water sufficient to cover the paper; shake and rotate for even distribution. Filtering is essential for high yields of uniform conidia.
Aseptically homogenize cultures in distilled water, mix equal volumes of the two suspensions and spread a portion over medium 81 made in pear-leaf decoction.
Pseudoperithecia initials fonn in 2 to 3 months and mature in another 2 to 3 months. Verticillium spp. Grow V. After 3 days irradiate with NUV for 15 min or with blue light to nm for 60 min and incubate in the dark. Irradiation with NUV for 30 min or more suppresses conidiation. Seed the middle of the plates with a culture disk and after 10 to 15 days in the dark, remove the cellophane with sclerotia and homogenize in a small amount of sterile distilled water, then atomize onto a small amount of soil and mix.
After air drying, sieve the mixture to break up large particles and use to infest soil in the field or greenhouse. Separate the conidia by shaking on a rotary shaker for 1 hr and then filter through two or three layers of cheesecloth. The filtrate contains the conidia; or culture the fungus on g autoclaved moist wheat grain in Erlenmeyer flasks until all grains arc covered with mycelium; conidia are removed by washing with ml of water and filtering through cheesecloth 45 CULTURE OF PATHOGENS Mixed inoculum conidia, mycelium, sclerotia is prepared on maize meal: sand mixture which is used directly for soil infestation.
Xanthomonas campestris pv. Baker, R. Bouchereau, P. Garrell, S. Cambridge, Garrett, S. Press, Cambridge, Isaac, I. Phillips, D. Toussoun, T. Snyder, W. Hansen, H. Phytopathology, 37, , Ostazeski, S. Zentmyer, G. Satyanarayana, K. Plant Pathol. Goth, H. Slade, S. Leach, C. Marsh, P. Harrison, J. Singh, R. Bhama, K. Aragaki, M. Leach, J. Strandberg, J. Child, J. Sacks, L. Boddy, L. Eichenmuller, J. Suryanarayanan, T. Lukens, R. Kilpatrick, R. McDonald, W. Srinivasan, M.
Schroeder, H. Dhiman, J. Gilbertson, R. Papavizas, G. Haglund, W. Bhalla, H. Yang, C. Mitchell, J. Cunningham, J. Carmen, L. Schneider, C. Sugar Beet Techno!. Ghafoor, A. Humaydan, H. Shahin, E. Senior, D. Billotle, J. Seances Acad. Plant Sci. McRae, C. Dimitrijeivc, B. Miusov, LN. Nauki Alma Ata , 11, 86, Ludwig, R. Plant Dis. Survey, 42, , Douglas, D. Dorozhkin, N. Zhu, Z. Mycologica Sinica, 4, , Engelhard, A. Paul, , Latch, G. Amy, D. Reifschneider, F.
Hodges, C. VanderMeer, Q. Ellerbrock, L. Clark, C. Laha, S. Faretra, F. Last, F. Presly, A. Murakishi, H.
Lewis, R. Nagel, C. Starkey, T. Smith, D. Calpouzos, L. Jauch, C. Aires, 3, 80, Canova, A. Ruppel, E. Mew, I. Khandar, R. Diachun, S.
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