Downy Mildew Disease of Crucifers: Biology, Ecology and Disease

Govind Singh Saharan · Naresh Mehta Prabhu Dayal Meena

Downy Mildew Disease of Crucifers: Biology, Ecology and Disease Management Downy Mildew Disease of Crucifers: Biology, Ecology and Disease Management Govind Singh Saharan • Naresh Mehta Prabhu Dayal Meena

Downy Mildew Disease of Crucifers: Biology, Ecology and Disease Management Govind Singh Saharan Naresh Mehta Department of Plant Pathology Department of Plant Pathology CCS Haryana Agricultural University CCS Haryana Agricultural University Hisar, Haryana, India Hisar, Haryana, India

Prabhu Dayal Meena ICAR-Directorate of Rapeseed-Mustard Research Bharatpur, Rajasthan, India

ISBN 978-981-10-7499-8 ISBN 978-981-10-7500-1 (eBook) https://doi.org/10.1007/978-981-10-7500-1

Library of Congress Control Number: 2017961551

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This Springer imprint is published by Springer Nature The registered company is Springer Nature Singapore Pte Ltd. The registered company address is: 152 Beach Road, #21-01/04 Gateway East, Singapore 189721, Singapore Foreword

Crucifers are very important and widespread crops grown worldwide in cool temperate, continental, and subtropical regions. Agriculturally economic crops include Brassica oil-yielding crops, Brassica vegetable crops, fodder crops, horticultural crops, and several weeds including Arabidopsis – a model plant for genomic studies in this modern era. Cruciferous Brassica vegetables are a major source of vitamins, fiber, minerals, and proteins in the human diet, while Brassica oilseeds are a major source of quality vegetable oil, and cake for animal feed. The demand for Brassica vegetables, and oilseeds is increasing annually globally. Crucifers are damaged by several biotic, and abiotic stresses under environmental conditions wherever these crops are grown. Downy mildew in crucifers is the third major biotic stress that causes severe annual yield losses of this crop. This book, Downy Mildew Disease of Crucifers: Biology, Ecology and Disease Management, is an important, timely contribution to our knowledge about Hyaloperonospora parasitica (Gaum.) Goker, the most destructive pathogens of crucifers globally. In spite of our access to current information through the Internet, an encyclopae- dic book is required where students, researchers, teachers, and industrialists can get well-documented updated information at their desk. This book is arranged into 16 different chapters with proper headings, and subheadings; illustrations such as pho-

v vi Foreword tographs, graphs, figures, and tables; and references to stimulate better comprehension on the disease. It provides a much-needed background on the disease, and current information with insight knowledge on future priorities, areas of research, and methodologies, making it a central reference for use by the ‘Brassicalogists’ across the world regarding investigations on the pathogen, and its host. I congratulate the authors Drs. G. S. Saharan, Naresh Mehta and P. D. Meena for timely bringing their lifelong professional interest, and expertise into a comprehensive treatise. This book has been designed to be the most useful resource with a wide range of logically organized, and easily accessable information, and is a very important contribution in the series on Sclerotinia diseases, white rust disease, and Alternaria diseases of crucifers authored by the aforementioned scientists and published by Springer.

New Delhi T. Mohapatra 11th September, 2017 Preface

Since the first comprehensive treatise publication in the form of aMonograph on Downy Mildew of Crucifers (Saharan, Verma, Nashaat, 1997), lots of information have been generated, and published on downy mildew of crucifers in the form of research papers, reviews, book chapters, scientific popular or extension articles, and news for farmers. This book entitled Downy Mildew Disease of Crucifers: Biology, Ecology, and Disease Management encompasses all the available published information for the access to researchers, teachers, students, extension experts, and other end users like industrialists, and farmers to comprehend the third most widespread, and devastating disease of crucifers. The disease is known to cause severe yield losses quantitatively and qualitatively in cruciferous crops grown all over the world. The major cruciferous crops challenged by the disease are Brassica crops grown for high-quality edible (rapeseed-mustard, canola, and other rape), and industrial (Crambe) oils, forage Brassicas, common vegetable crops (cabbage, cauliflower, radish, kohlrabi, broccoli, Brussels sprouts, kales, and other Brassicas), ornamental plants (wall flower, stocks), and several weeds including Arabidopsis, a model plant for studying genomics, and molecular genetics through Arabidopsis- Hyaloperonospora pathosystem. The information has been arranged in 16 chapters with several headings and subheadings in numerical series. Each chapter includes references for original consultation by the readers. The chapter-wise arrangement includes the introduction to the book narrating development in the downy mildew research of crucifers over time comprising of present status of disease, and pathogen; the disease and its distribution, symptomatology, host range, yield losses, and disease assessment; the pathogen and its taxonomy, morphology, phylogeny, variability, sporulation, survival, and perpetuation; spore germination, infection, pathogenesis, seed infection, disease cycle, epidemiology, forecasting, and fine structures; biochemical, histological, genetic, and molecular mechanisms of host resistance, including cloning and mapping of R genes; sources of resistance, disease resistance breeding strategies, and genetics of host-parasite interactions; disease management through cultural, chemical, biological, and host resistance methods, and integrated approach; and standardized reproducible techniques. Chapter 15 offers suggestions for future priority areas of research. To track the information given in different

vii viii Preface chapters, subject index has been included in Chapter 16. The text has been vividly illustrated with photographs, graphs, figures, histograms, tables, coloured plates, and flowcharts for effective, and stimulating easy comprehension by the readers. We believe that this book will be immensely useful to the researchers, teachers, extension specialists, students, industrialists, farmers, and all others who are inter- ested to grow healthy, and profitable cruciferous crops all over the world. Any suggestions by the readers are always a source of inspiration for the authors. Any shortcomings, lacunae, and flaws in the book are responsibility of ours.

Hisar, Haryana, India G.S. Saharan Naresh Mehta Bharatpur, Rajasthan, India P.D. Meena Acknowledgements

Authors are highly grateful to the following persons/scientists/publishers/societies/ journals/institutes/websites, and all others whose valuable materials such as photographs (macroscopic, microscopic, electron micrographs, scanning electron micrographs), drawings, figures, histograms, graphs, tables, flow charts, etc., have been used through reproduction in the present document. The address of the author(s)/ source(s) from where material has been adapted can be obtained from the reference which is cited in the reference section of the book. The authors are also sincerely thankful to all the scientists/publishers/journals/institutes/societies, and websites whose materials have been reproduced in one or the other form in this manuscript but forget to acknowledge their name(s) inadvertently. A. Persons/Scientists/Authors Allen RL, Asada Y, Awasthi RP, Babber S, Bahcevandziev K, Baxter L, Bergot M, Cabral A, Cao H, Chakraborty S, Channon AG, Chattopadhyay C, Choi YJ, Chou CK, Coelho PS, Constantinescu O, Cooke RC, Crute IR, Damgaard C,

ix x Acknowledgements

Daniels MJ, Dangl JL, Davison EM, Dean C, Dickinson CH, Fatehi J, Felton MW, Garcia-Blazquez G, Goker M, Gorden PL, Greenhalgh JR, Hartman H, Hayter JBR, Heran A, Hidaka H, Holt III BF, Holub EB, Jang P, Jensen BD, Jones JD, Kamoun S, Kluczewski SM, Koch E, Kolte SJ, Kruger W, Lebeda A, Leung H, Li X, Lin CY, Lister BJ, Lucas JA, Mims CW Mansfield JW, Matsumoto I, Mauch-Mani B, Mitchell SE, Monteirio AA, Moss NA, Nagatani T, Nashaat NI, Parker JE, Ploch S, Proctor R, Rehmany AP, Richardson EA, Acknowledgements xi

Riethmuller A, Ryan EW, Safeeulla KM, Sakamoto K, Sangeetha CG, Schwin FJ, Schlaich NL, Sharma SR, Shaw CG, Sherriff C, Shin HD, Singh D, Shiraishi M, Siddaramaiah AL, Singh RB, Singh RN, Slusarenko AJ, Sohi HS, Soylu EM, Soylu S, Spring O, Staskawicz BJ, Sutton JC, Szabo V, Tham FY, Thomas L, Thornton JD, Thines Macro, U N, Valerio L, Verma PR, Vicente JG, Vishwanath, Voglmayr H, Walker JC, Wang TM, Williams PH, Wilson ZA, Xiao D, Yerkes WD, B. Journals African Journal of Agricultural Research African Journal of Biotechnology Annals of Applied Biology xii Acknowledgements

Annals of Botany Annals of Phytopathological Society, Japan Annual Review of Phytopathology Australian Journal of Agricultural Research Brassica Botany Brazilian Phytopathological Society Canadian Journal of Botany Canadian Journal of Plant Pathology Canadian Journal of Plant Science Cell Crop Protection Cruciferae Newsletter Euphytica European Journal of Plant Pathology Evolution Fungal Diversity Fungal Genetics & Biology Genes, Genomics, Genetics Horticulture Research Indian Phytopathology Indian Journal of Agricultural Sciences International Journal of Agriculture, Environment & Biotechnology Japan Journal of Botany Journals of Agricultural Research Journal of Applied Microbiology Journal of Biological Chemistry Journal of General Plant Pathology Journal of Indian Botanical Society Journal of Mycology and Plant Pathology Journal of Oilseed Brassica Journal of Phytopathology Journal of Plant Disease and Protection Molecular Microbiology Molecular Plant Microbe Interaction Molecular Plant Pathology Molecules Mycologia Mycological Progress Mycological Research Mycopathologia Nature New Pathologist Nova Hedwigia Nucleic Acids Research Acknowledgements xiii

Oikos Physiological and Molecular Plant Pathology Physiological Plant Pathology Phytopathology Phytochemistry Phyton Plant Breeding Plant Disease Plant Disease Research Plant Journal Plant Molecular Biology Plant Pathology PLoS Pathogens Proceedings of the National Academy of Sciences USA Proceedings of the Indian Academy of Sciences, Plant Science Review of Plant Pathology Science Seed Science & Technology Journal Sydowia Tests of Agrochemicals The Canadian Journal of Plant Pathology The Plant Cell The Plant Pathology Journal Transaction of British Mycological Society Tropical Plant Pathology Theoretical and Applied Genetics C. Websites http://diwww.epfl.ch/ wstamatak/ index-Dateien/publications/GCB2006_Poster. pdf; http://prodes.toulouse.fr/multialign/multialign.html http://brassicadb.org/brad http://www.bio-rad.com http://frodo.wi.mit.edu/primer3 http://en.cellfood.com.cn/culture.aspx http://prgdb.cbm.fvg.it http://cals.arizona.edu/PLP/pryorlab/alternaria.html http://nt.ars-grin.gov/fungaldatabases/ http://www.mycobank.org http://www.marinespecies.org http://www.elsevier.com/locate/yfgbi http://www.sciencedirect.com http://www.ipcc.ch xiv Acknowledgements

D. Publishers/Press Academic Press, London CABI, London, UK CRC Press, Boca Raton, Florida, USA, CSIRO Publishing Elsevier Scientific Publisher, Oxford John Wiley & Sons, Inc. Narosa Publishing House, New Delhi Springer, Netherlands/ New York Taylor & Francis Group Scientific Publications, Jodhpur, India Indus Publication Company, New Delhi, India American Oil Chemists Society Press Kluwer Academic Press, Dordrecht, Netherlands Wiley Blackwell Publication, Hoboken, New Jersey, USA E. Institutions/ Societies All Indian Coordinated Research Project, ICAR, New Delhi Agriculture and Agri-Food Canada, Saskatoon Research Centre, Saskatoon, Canada American Phytopathological Society, USA Canadian Phytopathological Society CCS Haryana Agricultural University, Hisar, India Directorate of Rapeseed-Mustard Research, Bharatpur, India Indian Council of Agricultural Research, India Indian Phytopathological Society, New Delhi Indian Society of Mycology and Plant Pathology Indian Society of Plant Pathologist, Ludhiana International Development Research Centre, Ottawa, Ontario, Canada Korean Society of Plant Pathology Mycological Society of America Pryor Laboratory, University of Arizona School of Plant Sciences, USA Society for Rapeseed-Mustard Research The American Phytopathological Society The Australasian Plant Pathology Society The British Society for Plant Pathology The Korean Society of Plant Pathology University of Wisconsin, Madison GCIRC F. Databases MycoBank, International mycological Association Systematic Mycology and Microbiology Laboratory Fungal Database, U.S. Department of Agriculture Contents

1 Introduction...... 1 1.1 Crops and Their Distribution...... 1 1.2 The Disease and Pathogen...... 3 1.3 The Downy Mildew of Crucifers...... 6 1.4 The Pathogen/Causal Organism of Downy Mildew of Crucifers...... 12 1.5 Taxonomy and Classification of Downy Mildew Pathogen...... 13 1.6 Current Generic Status of Downy Mildew of Crucifers...... 14 1.7 Species Concepts in Crucifer’s Downy Mildew...... 14 1.8 Broad and Narrow Species Concepts...... 15 1.9 Use of Molecular Data for Downy Mildew Species Concept...... 16 1.10 Hyaloperonospora Species on Crucifers...... 17 1.11 Strategies to Breed Downy Mildew Resistance Cultivars of Crucifers...... 18 1.11.1 Identification and Utilization of Receptor-Like Kinases Involved in Plant Immunity...... 18 1.11.2 Identification and Utilization of R Genes Involved in ETI...... 19 1.11.3 The Utilization of Quantitative Trait Loci (QTLs)...... 19 1.11.4 Screening and Utilization of Recessive Gene-Mediated Broad-Spectrum Resistance...... 19 1.11.5 Engineering Broad-Spectrum Resistance Through Biotechnology...... 20 1.11.6 Designation and Nomenclature of Downy Mildew Resistance Genes (R Genes) and Isolates (Races/Pathotypes)...... 20 1.12 Importance of Hyaloperonospora arabidopsidis in Molecular Plant Pathology...... 23 1.13 Impact of Climate Change on the Diseases of Crucifers...... 24 References...... 28

xv xvi Contents

2 The Disease: Downy Mildew...... 35 2.1 Introduction...... 35 2.2 Geographical Distribution...... 35 2.3 Economic Importance...... 40 2.3.1 Brassica Oilseeds...... 41 2.3.2 Brassica Vegetables...... 42 2.4 Host Range...... 43 2.5 Symptoms...... 43 2.5.1 Brassica Oilseeds...... 43 2.5.2 Brassica Vegetables...... 52 2.5.3 Broccoli...... 54 2.5.4 Wallflower (Cheiranthus)...... 54 2.5.5 Stock (Matthiola)...... 54 2.5.6 Rocket (Eruca sativa)...... 55 2.5.7 Cruciferous Weed (Arabidopsis thaliana)...... 55 2.6 Disease Assessment...... 56 References...... 59 3 The Pathogen: Hyaloperonospora parasitica (Gaum.) Goker [H. brassicae (Gaum.) Goker]...... 67 3.1 Introduction...... 67 3.2 Taxonomy and Morphology...... 69 3.2.1 Phylogenetic Analyses...... 75 3.2.2 The Species Concept of Downy Mildew...... 81 3.2.3 Relationship of Peronospora with Hyaloperonospora and Perofascia...... 84 3.2.4 Major Species Clusters in Hyaloperonospora...... 84 3.2.5 Hyaloperonospora arabidopsidis on Arabidopsis...... 86 3.3 Reproduction and Reproductive Structures...... 87 3.3.1 Asexual Phase...... 87 3.3.2 Conidiophores and Conidia...... 88 3.3.3 Sexual Phase...... 88 References...... 89 4 Electron Microscopy and Ultrastructures...... 93 4.1 Introduction...... 93 4.2 Host Penetration...... 93 4.3 Haustorium Development...... 97 4.4 The Host-Pathogen Interface...... 98 4.5 Ultrastructural Features of Intercellular Hyphae, Haustorium, and Host Cell...... 109 4.6 Conidiophore Development...... 111 4.6.1 Conidiophore Primordial...... 111 4.6.2 Unbranched Conidiophores...... 111 4.6.3 Production of Branches...... 111 Contents xvii

4.6.4 Development of Conidia...... 112 4.6.5 Formation of a Cross Wall...... 112 4.6.6 Conidiophore Growth...... 112 4.6.7 Conidial Formation...... 114 4.6.8 Host Response...... 115 4.6.9 Cytology and Genetics...... 116 References...... 125 5 Physiologic Specialization (Pathogenic Variability)...... 127 5.1 Introduction...... 127 5.2 Pathogenic Variability...... 128 5.2.1 DNA Fingerprinting of H. parasitica...... 138 5.2.2 Identification of Host Differentials and Nomenclature of Pathotypes...... 140 5.3 Heterothallism and Homothallism...... 142 5.4 Hybridization of Hyaloperonospora Isolates...... 143 References...... 143 6 Perpetuation and Survival of Pathogen...... 147 6.1 Introduction...... 147 6.2 Mycelium...... 147 6.3 Conidia...... 147 6.4 Oospores...... 148 6.5 Seed Infection...... 149 6.6 Axenic Culture...... 149 6.7 Conidial Discharge...... 149 6.8 Conidial Germination...... 151 6.9 Oospore Germination...... 154 References...... 155 7 Infection and Pathogenesis...... 157 7.1 Introduction...... 157 7.2 The Process of Infection...... 157 7.2.1 Light Microscopic Observation of Infection Process...... 159 7.3 Nature and Mechanism of Pathogenesis...... 165 References...... 174 8 Disease Cycle...... 175 8.1 Introduction...... 175 8.2 General Disease Cycle on Cruciferous Crops...... 175 8.3 Disease Cycle on Arabidopsis...... 177 8.4 Factors Affecting Disease Cycle...... 180 References...... 180 xviii Contents

9 Epidemiology and Forecasting...... 183 9.1 Introduction...... 183 9.2 Disease Development in Relation to Temperature, Humidity, Rainfall, and Leaf Wetness...... 183 9.3 Disease Development in Relation to Planting Time...... 192 9.4 Disease Development in Relation to Host Nutrition...... 192 9.5 Disease Interaction with Insecticidal Sprays...... 196 9.6 Disease Prediction Models...... 196 References...... 197 10 Association or Mixed Infection of Downy Mildew and White Rust Disease Complex...... 199 10.1 Introduction...... 199 10.2 Symptoms...... 200 10.3 Yield Losses...... 201 10.4 Pathogenesis...... 202 10.5 Histopathology...... 204 10.6 Epidemiology...... 208 10.7 Disease Forecasting...... 209 10.8 Altered Phenotypic Expression of Downy Mildew...... 211 References...... 211 11 Biochemistry of the Host Pathogen Interaction...... 215 11.1 Introduction...... 215 11.2 Metabolic Changes...... 215 11.3 Role of Natural Biochemical Compounds...... 220 11.4 Biochemistry of Disease Resistance...... 220 References...... 223 12 Host Resistance...... 225 12.1 Introduction...... 225 12.2 Mechanism of Host Resistance...... 226 12.3 Host-Pathogen Recognition System...... 231 12.4 Systemic Acquired Resistance...... 231 12.4.1 Expression of Systemic Acquired Resistance...... 232 12.5 Genetics of Host-Pathogen Relationship...... 233 12.5.1 Seedling and Adult Plant Resistance to Downy Mildew...... 240 12.5.2 Inheritance of Partial Resistance to Downy Mildew...... 241 12.5.3 Molecular Basis of Downy Mildew Resistance...... 242 12.5.4 Mutation Approach to Identify Resistance Genes...... 243 12.5.5 Genetics of Multiple Disease Resistance...... 245 12.5.6 Disease Resistance Increases Competitive Ability of Host Plants...... 247 12.5.7 Expression of Age-Related Resistance (ARR) to Downy Mildew...... 247 Contents xix

12.5.8 Different Requirements for Disease Resistance Genes..... 249 12.5.9 Differential Expression of Downy Mildew Resistance Genes...... 250 12.5.10 Cloning of Major Resistance Genes...... 252 12.5.11 Mapping of Downy Mildew Resistance Genes...... 256 12.5.12 Resistance Gene-Mediated Signal Transduction...... 257 12.6 Biochemical Basis of Resistance...... 260 12.6.1 Role of Phytoalexins in Resistance to Downy Mildew...... 261 12.6.2 Lignification of Host Cells...... 262 12.7 Sources of Resistance...... 264 12.8 Breeding for Disease Resistance...... 267 12.8.1 Strategies to Breed Downy Mildew Resistance Cultivars of Crucifers...... 267 12.8.2 Designation and Nomenclature of Resistance Genes...... 267 12.9 Mechanisms and Application of Gene Silencing Techniques to Downy Mildew of Crucifers...... 268 12.9.1 Stable Versus Transient Gene Silencing...... 270 12.10 Development of Resistance to Fungicides...... 271 References...... 271 13 Disease Management...... 285 13.1 Introduction...... 285 13.2 Cultural Practices...... 285 13.3 Seed Treatment...... 286 13.4 Soil Treatment...... 287 13.5 Compost Treatment...... 288 13.6 Foliar Spray of Fungicides...... 288 13.6.1 Brassica Vegetables...... 289 13.6.2 Brassica Oilseeds...... 294 13.7 Biological Control...... 299 13.7.1 Plant Extracts as Fungitoxicant...... 299 13.7.2 Antagonists for Biocontrol...... 299 13.8 Host Resistance...... 301 13.9 Fungicide Resistance...... 301 13.10 Integrated Disease Management...... 301 References...... 304 14 Techniques...... 309 14.1 Introduction...... 309 14.2 Culturing of Hyaloperonospora parasitica...... 309 14.3 Maintenance of H. parasitica Isolates and Production of Inoculum...... 310 14.4 Germplasm Screening and Evaluation...... 311 14.5 Preservation of Hyaloperonospora parasitica...... 313 14.6 Artificial Inoculation of Excised Cotyledons...... 314 xx Contents

14.7 Propagation of Hyaloperonospora parasitica on Cotyledons or True Leaves of Japanese Radish Seedlings...... 314 14.8 Laboratory Tests of Fungicides...... 315 14.9 Fungicide Resistance Assay...... 316 14.10 Measuring Systemic Infection of the Downy Mildew Pathogen.... 316 14.11 Methods of Breeding for Multiple Disease Resistance...... 317 14.12 Heterothallism and Homothallism...... 320 14.13 Seed-Borne Nature of H. parasitica...... 320 14.14 Conidial Germination...... 321 14.15 Sporulation...... 322 14.16 Discharge of Conidia...... 322 14.17 DNA Fingerprinting of Hyaloperonospora parasitica...... 322 14.17.1 DNA Isolation...... 323 14.18 Molecular Marker for Identification of H. parasitica...... 324 14.19 Leaf Disc Test to Assess Resistance...... 328 14.20 Use of Rooted Leaves for Screening Brassica Germplasm...... 329 14.21 Artificial Inoculation Technique Under Growth Chamber (Williams 1985)...... 330 14.22 Microscopic Studies...... 332 14.23 Light and Transmission Electron Microscopy (TEM)...... 334 14.24 C DNA-AFLP Analysis to Reveal Gene Expression...... 335 14.25 Thawing and Revival of Inoculum...... 337 14.26 Obtaining New Isolates from Dried Leaf Tissue Containing Oospores...... 338 References...... 338 15 Future Strategies and Priorities of Downy Mildew DiseaseManagement...... 343 15.1 Introduction...... 343 15.2 Disease Epidemiology...... 343 15.3 Physiological Specialization...... 344 15.4 Genetics of Resistance...... 344 15.5 Molecular Plant Pathology/Genomics/Genetic Engineering...... 345 15.6 Biochemical Aspects of Resistance...... 346 15.7 Disease Management...... 346 15.8 Phylogenetic Relationship and Co-evolution of Cruciferous Hosts and Downy Mildew Pathogen...... 347

Index...... 349 Authors

G.S. Saharan, ex-professor and head of the Department of Plant Pathology, retired from the active service in 2002. He did his B.Sc. (1965), and M.Sc. (1967) in agriculture from S.K.N. College of Agriculture, Jobner, University of Udaipur, and Ph.D. (1977) from Himachal Pradesh University, Palampur, India. He served as lecturer (1967–1976) and assistant professor (1976–1980) at HPKVV, Palampur, and as associate professor (1980–1988), professor (1988–2002), and professor and head (2002) at the Department of Plant Pathology at CCS Haryana Agricultural University, Hisar. Dr. Saharan has been a visiting professor at the Department of Plant Sciences, University of Alberta, Edmonton, Canada (1991 and 1994); Agriculture and Agri- Food Canada, Saskatoon Research Station, Saskatoon, Canada (1991, 1994, 1997); and Rothamsted Research, IACR, Harpenden, UK (1994 and 1997). Dr. Saharan has more than 250 research publications in journals of national and international repute. He has been editor of books, i.e. Diseases of Oilseed Crops, Annual Review of Plant Pathology, Phytopathological Techniques, Plant Pathology at a Glance, and Plant Pathological Research Problems and Progress, and author of books Diseases of Oilseed Crops (in Hindi), Sclerotinia Diseases of Crop Plants: Biology, Ecology and Disease Management, White Rust of Crucifers: Biology, Ecology and Management, and Alternaria Diseases of Crucifers: Biology, Ecology and Disease Management. He has authored monographs on white rust, Alternaria blight, and downy mildew diseases of rapeseed-mustard, including 5 bibliographies and 40 review articles in books. He is on the panel of experts of State Agricultural Universities, Indian Council of Agricultural Research, Central Scientific and Industrial Research, University Grants Commission, and Department of Biotechnology in India. He has contributed immensely in the preparation and release of Crop Protection Compendium (2002), CAB International, UK. He has guided three M.Sc. and eight Ph.D. students who are well-established scientists at different universities and research organizations in India and abroad.

xxi xxii Authors

Dr. Saharan has conducted research in diverse fields of plant pathology including standardization of artificial inoculation techniques, identification of sources of resistance, determination of pathogenic variability, genetics of host-parasite interaction, epidemiology, and management of several diseases. He has been president (North Zone) of the Indian Phytopathological Society (2001), editor-in-chief of the Journal of Mycology and Plant Pathology (1999–2000) and the Journal of Oilseed Brassica (2012 to date), and president of the Indian Society of Mycology and Plant Pathology (2009) and has played a major role in the organization of global and Asian congress by the leading phytopathological societies of India. He has been a member of QRT, ICAR, New Delhi, for the soybean (2010) and rapeseed-mustard (2015). Dr. Saharan has been awarded with Y. L. Nene Outstanding Plant Pathology Teacher Award (2015) by the Indian Society of Mycology and Plant Pathology, Udaipur, India. He has been bestowed with Lifetime Achievement Award (2017) for his outstanding research leadership and expertise in oilseed Brassica research by the Society for Rapeseed-Mustard Research, Bharatpur, India.

Naresh Mehta did his B.Sc. in agriculture (Hons.) in 1978 and M.Sc. in plant pathology in 1980 from Haryana Agricultural University, Hisar. He received his Ph.D. degree in plant pathology in 1993 from CCS HAU, Hisar, under the guidance of Dr. G. S. Saharan. During his study, he attained first position in elective plant protection at bachelor’s degree level. Dr. Mehta is the recipient of Excel Industries Ltd. Bombay, India, Award as research fellowship for master’s degree programme and ‘Senior Research Fellowship’ (SRF) Award to pursue Ph.D. programme by the Council of Scientific and Industrial Research (CSIR), New Delhi. He was awarded Ms. Manju Utereja Memorial Gold Medal for best Ph.D. thesis (1993–1994). Dr. Mehta joined as assistant scientist (plant pathology) in 1981, scientist/assoc. professor in 1994, and professor in 2002 at CCS, HAU, Hisar. He was co-principal investigator in the scheme ‘Pathogenic variability and epidemiology of Alternaria brassicae’ funded by ICAR, New Delhi. He has been teaching plant pathology courses to undergraduate and postgraduate students. He has guided five M.Sc. (plant pathology) and two Ph.D. students, and he is a member of many students’ advisory committees. He is the recipient of Best Poster Paper Award for the year 2005 by INSOPP and Indian Phytopathological Soc. (NZ). His students have been awarded P.R. Verma M.Sc. Student Thesis Award for the year 2009 by Indian Soc. of Mycology and Plant Pathology and M.J. Narasimhan Academic Awards (NZ) by Indian Phytopathological Society, New Delhi, for the year 2010. He has conducted research in diverse fields of plant pathology covering pathogenic variability, genetics of host-pathogen interaction, epidemiological studies, identification of resistant sources, biochemical/genetic basis for resistance, residual analysis of fungicides, and disease management. Authors xxiii

Dr. Mehta was a member of Expert Committee, UGC, New Delhi, for 12 B status for the Gandhigram Rural Institute-Deemed University, Gandhigram, Tamil Nadu. Dr. Mehta is one of the editors of the book “Diseases of Oilseed Crops” and one of the authors of books Sclerotinia Diseases of Crop Plants: Biology, Ecology and Disease Management and Alternaria Diseases of Crucifers: Biology, Ecology and Disease Management. He has published more than 100 research papers in the journals of national and international repute. In addition, 10 review articles, 20 book chapters, 10 practical manuals, 25 lead lectures in the conferences, 91 research paper presentations in the conferences, 35 popular articles, and 13 radio/TV talks are to his credit. Dr. Mehta has been admitted as fellow of Indian Phytopathological Society (FPSI), New Delhi, Indian Society of Plant Pathologist (FINSOPP), Ludhiana, and Indian Society of Mycology and Plant Pathology (FISMPP), Udaipur. He has been on the editorial board of Indian Phytopathological Society (2012–2013, 2017– 2019), councillor (North Zone) of ISMPP (2005, 2011), a member of editorial board (2012–2014), and editor-in-chief (2014). He is also a member of Editorial Board of Indian Society of Plant Pathologist, Ludhiana, 2017–2018. Dr. Mehta has been a visiting scientist to University of Alberta, Edmonton, Canada, in 1999 as a FAO fellow and presented a research paper in the 8th International Congress of Plant Pathology at Christchurch, New Zealand, 2002. He was invited to deliver lectures in the 9th International Congress of Plant Pathology, at Torino, Italy, 2008, and in the 5th International Conference on Plant Pathology on the theme ‘Plant Pathology in the Globalized Era’, New Delhi, 2009. In 2012, he has delivered lead lecture in the 3rd Global Conference on the theme ‘Plant Pathology for Food Security’ with several lead lectures in the national conferences held from time to time.

P.D. Meena is working as Principal Scientist (Plant Pathology) at the Directorate of Rapeseed-Mustard Research (Indian Council of Agricultural Research), Bharatpur-321 303 Rajasthan, India. He started his career in Indian Council of Agricultural Research in 1989 as a senior technical assistant at Central Soil & Water Conservation Research & Training Institute, Dehradun. He obtained his B.Sc. (Ag.) (1987) from the University of Rajasthan, Jaipur, and M.Sc. (Ag.) in plant pathology (1997) from Rajasthan Agricultural University, Bikaner, and Ph.D. in botany (2005) from the University of Rajasthan, Jaipur. He has developed garlic bulb aqueous extract (2 % w/v) as botanical product for control of Sclerotinia rot and Alternaria blight diseases of mustard. He has identified white rust resistance genotypes, viz. NRCDR 515, NRCDR-02, NRCHB-506, NRCHB-101, and NRCDR-601 of Brassica juncea and NRCYS-05-2 of B. rapa ssp. yellow sarson, and developed weather-based forecasting models for rapeseed-mustard diseases. He has published more than 50 research papers, reviews, and book chapters in xxiv Authors international and national reputed refereed journals, and he is editor of the book Principles of Plant Breeding. He has been a member of monitoring team for All India Coordinated Research Project on Rapeseed-Mustard in 2004–2010. He has been honoured with Fellowship of Indian Society of Mycology and Plant Pathology and fellow of Plant Protection Association of India and also awarded with Dr. P.R. Kumar Outstanding Brassica Scientist Award in 2011 by the Society for Rapeseed-Mustard Research. He has served as councillor of Plant Protection Association of India, the founder secretary of the Society for Rapeseed-Mustard Research since 2008, a member of the editorial board (2010–2011), and managing editor (2012–2013) for the Journal of Oilseed Brassica. Dr. Meena has been a principal investigator (PI) and co-PI for ICAR Outreach Programme on Diagnosis and Management of Leaf Spot Diseases in Field and Horticultural Crops (2009–2013), ICAR Network Project on Transgenics in Crops (Functional Genomics Component for Alternaria and Drought), and National Network for Management of Alternaria Blight in Brassica juncea and Vegetable Crops (2004–2008). He is the recipient of the Best Paper Presentation Award (2005), Appreciation Certificate from Rajasthan Jankalyan Van Avum Paryavarn Vikas Sansthan, Jaipur (2006), and K.S. Bilgrami Best Paper Presentation Award (2010). He undertook 3 months of research attachment training (2007) at the Rothamsted Research, Harpenden, UK, under Indo-UK collaborative research on oilseed brassica crops. He has supervised nine M.Sc. students and co-supervised one Ph.D. student. Abbreviations

@ At the rate of < Less than > greater than μ Micron μEm−2S−1 Micro Einsteins per square metre per second μg Microgram μgml−1 Microgram per millilitre μh−1 Per microhenries μm Micrometre μmol g−1 Micromole per milligram °C Degree Celsius a.i. Active ingredient a.m. Ante meridiem; before noon AB Alternaria blight Ab Alternaria brassicae Ac Albugo candida AFLP Amplified fragment length polymorphism AICRP-RM All India Coordinated Research Project on Rapeseed-Mustard AIP Aminoindan phosphonic acid AN Autogenic necrosis Ao Angstrom Ar Aphanomyces raphani At Agrobacterium tumefaciens AUDPC Area under the disease progress curve AY Aster yellows B (N) Belonolaimus (nematode) BABA β-Amino-butyric acid BAC term BAC Bacterial artificial chromosomes Bc Botrytis cinerea BMY(V) Beet western yellows virus

xxv xxvi Abbreviations bp Base pair BrS Bremer decay indices BS Black speck BTH Benzothiadiazole C.D. Critical difference CAD Cinnamyl alcohol dehydrogenase CaM Calmodulin CAM (V) Cauliflower mosaic virus Cb Cercospora brassicicola cDNA Complementary DNA cDNA-RFLP cDNA-restriction fragment length polymorphism CHP Conserved hypothetical protein cm Centimetre cv. Cultivar D (N) Dolichodorus (nematode) dai Days after inoculation DAS Days after sowing DI Disease index DM Downy mildew dsRNA Double-stranded RNA Ec Erwinia carotovora Ec Erysiphe cruciferarum EDS1 Enhanced disease susceptibility ER Endoplasmic reticulum ER1 Exoribonuclease F Fusarium Fo Fusarium oxysporum G Gummosis g Gram g/kg−1 Gram per kilogram GP Genetic and physiological disorder h Hour Hs (N) Heterodera schachtii (nematode) h.p.i. Hour post inoculation ha−1 Per hectare Hp Hyaloperonospora parasitica HP Hyaloperonospora pathotypes HR Hypersensitive cell death response ICS Isochorismate synthase IP Interaction phenotypes ITS Internal transcribed spacer K Potash kb Kilobyte KCl Potassium chloride lbs Pound Abbreviations xxvii

LIF Lignification-inducing factor Lm Leptosphaeria maculans LSD Least significant difference LSU Large subunit M(N) Meloidogyne (nematode) Max Maximum Mb Mycosphaerella brassicicola Mg Magnesium min Minute Min Minimum miRNA Micro RNA ML Maximum likelihood ml Millilitre mm Millimetre MM (V) Mustard mosaic virus MP Maximum parsimony MRC Major recognition complexes mRNA Messenger RNA N Nitrogen NBS-LRR Nucleotide-binding site-leucine-rich repeat NDR1 Non-race-specific disease resistance NiF term NIF Nucleus-inducing factor No Nigrospora oryzae Oa Orobanche aegyptiaca OH-PAS Hydroxyphenyl-aminosulphinyl acetic acid dimethyl ester oz/acre Ounce per acre PLO Phytoplasma P Phosphorus P(N) Paratylenchus (nematode) p.m. Post meridiem; after noon PA(N) Paratylenchus (nematode) PAD Protrusion array devise PAL Phenylalanine ammonia-lyase PAMP Pathogen-associated molecular pattern PAZ Protein domain – Piwi, Argonaute, and Zwille Pb Plasmodiophora brassicae PBrS Partitioned Bremer support Pc Pseudocercosporella capsellae PCC Photosynthetic carbon cycle PCR Polymerase chain reaction PDS Percentage disease severity PEG Polyethylene glycol pH Potential of hydrogen ion Pm Phytophthora megasperma xxviii Abbreviations

Pmm Pseudomonas marginalis pv. marginalis Po Phymatotrichum omnivorum ppm Parts per million PR Pathogenesis related Psm Pseudomonas syringae pv. maculicola PT Pythium PTGS Post-transcriptional gene silencing Pyb Pyrenopeziza brassicae QRT-PCR Quantitative real-time polymerase chain reaction QTL Quantitative trait locus r Correlation coefficient R.H. Relative humidity RAPD Random amplification of polymorphic DNA rDNA Ribosomal ribonucleic acid RdRP RNA-directed RNA polymerase RFLP Restriction fragment length polymorphism Ribosomal DNA rDNA RISC RNA-induced silencing complex RITS RNA-induced transcriptional silencing RM(V) Radish mosaic virus RNA Ribonucleic acid ROS Reactive oxygen species Rs Rhizoctonia solani Rst Rhizopus stolonifer SA Salicylic acid SAR Systemic acquired resistance SCT Sexual compatibility type SDW Sterile distilled water siRNA Short interfering RNAs sq ft Square feet Sr Sclerotium rolfsii Ss Sclerotinia sclerotiorum Sts Streptomyces scabies T Temperature T.S. Transverse section TAS Trans-acting siRNA TB(P) Tip burn TDFs Transcript-derived fragments TEM Transmission electron microscopy TGS Transcriptional gene silencing TIR Toll, interlcukin-1, resistance TUM(V) Turnip mosaic virus Tu MV Turnip mosaic virus USA United States of America UK United Kingdom Abbreviations xxix v/v Volume by volume Va Verticillium albo-atrum var Variety Vd Verticillium dahliae w Watt w/v Weight by volume WP Wettable powder WR White rust Xca Xanthomonas campestris pv. armoraciae Xcc Xanthomonas campestris pv. campestris yr Year Ψ Atmospheric water potentials List of Figures

Fig. 1.1 U’s triangle showing species relationship among different Brassica species (UN 1935) �� 3 Fig. 2.1 Disease assessment (1–9) on leaves of rapeseed-mustard �� 58 Fig. 3.1 Phylogeny of the Peronosporales (Shaw 1981) �� 73 Fig. 3.2 Lower half of the phylogenetic tree inferred from the complete dataset with RAxML under a GTRMIX nucleotide substitution model approximation and rooted with Perofascia. Branch lengths are scaled in terms of the expected number of substitutions per site. Numbers above branches represent BS values above 50 % from ML (left) and MP (right) bootstrapping. Labels for Hyaloperonospora (H.) and Perofascia (P.) specimens indicate DNA isolation number and host species; if sequences were taken from GenBank, accession numbers are given. Thick vertical bars and adjacent names show the proposed species names. Asterisks mark binomials proposed in the present study. In the case of uncertainty regarding species boundaries, the bars are drawn in light grey. Thin vertical bars and adjacent numbers indicate clades apparently above species level as described in the text. See Fig. 3.3 for the upper half of this tree (Voglmayr 2003) �� 76 Fig. 3.3 Upper half of the phylogenetic tree depicted in Fig. 3.2, including clades 4–6. For a description, see Fig. 3.2 (Voglmayr 2003) �� 77 Fig. 3.4 Strict consensus of the 404 most parsimonious trees (length, 2066 bp) inferred from the reduced dataset that contains only specimens from which both ITS and LSU sequences could be obtained. Numbers below branches represent RAxML/GTRCAT (left)

xxxi xxxii List of Figures

and MP (right) BS values above 50 %. Numbers above branches (except terminal ones) are partitioned Bremer support values; the partitions examined were (from left to right) ITS1, 5.8S, ITS2, and LSU rDNA. Numbers above terminal branches represent their average lengths as inferred with DELTRAN optimization, as implemented in PAUP from the same partitions. Specimen labels are as in Figs. 3.2 and 3.3; affiliation of specimens to the clades used in these figures is indicated by vertical bars and numbers on the right side. The following symbols are used to indicate the suggested species boundaries according to Goker et al. (2004), as well as in the present text, and whether these are in accordance with Gaumann’s (1918, 1923, 1926) taxonomy: (-) in agreement with Gaumann; (!) not in agreement, with Gaumann’s species being paraphyletic, and including additional hosts; (x) not in agreement, Gaumann’s species polyphyletic; (0) host not examined by Gaumann; (?) type host not included in our sample. Asterisks point to molecular uncertainty with respect to species boundaries (Voglmayr 2003) �� 79 Fig. 3.5 Maximum likelihood phylogenetic tree inferred with RAxML from concatenated internal spacer region (ITS) and large ribosomal subunit (LSU) rDNA sequences. The dataset represents a subset of the one analysed by Goker et al. (2009). Technical details on the inference of this tree as well on the files used and on the origin of the sequences are provided in Goker et al. (2009). Note that the recognition of the clade numbers is based on the extended sampling used in Goker et al. (2009), whereas this figure only shows the specimens for which both ITS and LSU rDNA could be amplified. The numbers above the branches are bootstrap support values equal to or larger than 60% from 100 replicates. Abbreviations: H., Hyaloperonospora; P., Perofascia (Thines et al. 2009b) �� 85 Fig. 4.1 Increase in length of five individual conidiophores growing in the humidity chamber; br, time at which branching commenced; sp., spore formation (Davison 1968c) �� 115 Fig. 4.2 Increase in length of conidiophores A, B, and C. br, formation of primary branch; sp., spore formation (Davison 1968c) �� 116 Fig. 4.3 Increase in volume of conidiophores A, B, and C. sp., spore formation (Davison 1968c) �� 117 Fig. 4.4 Increase in branch length and apical diameter during spore formation. I branch length, b apical diameter (Davison 1968c) �� 117 List of Figures xxxiii

Fig. 4.5 The distribution of (a) nuclei, (b) RNA, and (c) mitochondria in the developing conidiophores of Hyaloperonospora parasitica (Davison 1968c) �� 122 Fig. 4.6 Migration of nuclei (n) into the nucleate spores of Hyaloperonospora parasitica (Davison 1968c) �� 123 Fig. 4.7 The distribution of (a) lipid material, (b) protein, and (c) insoluble carbohydrates in the developing conidiophores of Hyaloperonospora parasitica (Davison 1968c) �� 124 Fig. 6.1 (a) Pattern of Hyaloperonospora parasitica conidia discharge from infested Chinese cabbage plants and (b) temperature and humidity on 3 fine days in November 1978 (Lin 1981) �� 150 Fig. 6.2 The mechanism of Hyaloperonospora parasitica conidia discharge. (a) Conidiophores in damp air with attached conidia; (b) and (c) changes in conidiophores on exposure to dry air and (d) recovery on return to damp condition (Lin 1981) �� 151 Fig. 6.3 The effect of temperature and relative humidity on the germination of conidia of Hyaloperonospora parasitica (Lin 1981) �� 152 Fig. 6.4 Effect of hot water treatment on the germination of (a) conidia of Hyaloperonospora parasitica and (b) seeds of three Chinese cabbage cultivars. Conidia were held at each temperature for 15, 30, and 45 min, whereas seeds were held for 30 min only (Lin 1981) �� 153 Fig. 8.1 Diagramatic life cycle of Hyaloperonospora parasitica causing downy mildew of crucifers (Lucas et al. 1995) �� 176 Fig. 8.2 Disease cycle of downy mildew of crucifers (Saharan et al. 2005) �� 178 Fig. 8.3 Life cycle of Hyaloperonospora parasitica. (a) Infections arise initially from oospores germinating in the soil. (b) Plants are colonized by a coenocytic, intercellularly growing mycelium which swells to fit the intercellular spaces, giving it an irregular appearance. The hyphae put out pear-shaped feeding organs called haustoria into host cells. After a variable period of growth (1–2 weeks), conidiophores, bearing asexual, spherical hyaline conidiospores (c), grow out of stomata. (d) On germination, conidia initiate new rounds of infection. (e–g) Oospores are formed concurrently with asexual spores. (e) The female sexual organs, oogonia, contain an oosphere that is fertilized via a fertilization tube growing through its outer wall from the male antheridium. (f) The fertilized oosphere develops into a mature oospore. (g) Oospores are very profuse in infected leaves (Mauch-Mani and Slusarenko 1994) �� 179 xxxiv List of Figures

Fig. 9.1 The relationship of host, pathogen, and environment in the interaction phenotype of downy mildew of crucifers �� 184 Fig. 9.2 Effect of time and temperature on germination of conidia of Hyaloperonospora parasitica (Felton and Walker 1946) �� 185 Fig. 9.3 Effect of temperature upon penetration and development of haustoria of Hyaloperonospora parasitica (Felton and Walker 1946) �� 186 Fig. 9.4 Effect of five different temperatures on the initial sporulation of Hyaloperonospora parasitica at high humidity and upon initial appearance of symptoms at low and at high humidity (Felton and Walker 1946) �� 187 Fig. 9.5 Graphic summary of infection by and development of Hyaloperonospora parasitica on cabbage plants grown in sand culture supplied with various nutrient solutions (Felton and Walker 1946) �� 188 Fig. 9.6 Progression of downy mildew (Hyaloperonospora parasitica) of mustard (Brassica juncea) in relation to temperature (AUDPC) (Mehta et al. 1995) �� 189 Fig. 9.7 Effect of leaf wetness duration on the development of downy mildew (Hyaloperonospora parasitica) infection on mustard (Brassica juncea) cultivar RH-30 at 20 °C (Mehta et al. 1995) �� 191 Fig. 9.8 Effect of leaf wetness duration on the development of downy mildew (Hyaloperonospora parasitica) on mustard (Brassica juncea) seedlings of cultivar RH-30 at 15 °C (Mehta et al. 1995) �� 191 Fig. 9.9 Weather factors associated with occurrence (A) and no occurrence (B) periods of stag head phase of white rust (Albugo candida) and downy mildew (Hyaloperonospora parasitica) on mustard (Brassica juncea) in crop

seasons Y1 (1976–1977), Y2 (1977–1978), Y3 (1978–1979), Y4 (1979–1980), Y5 (1980–1981), Y6 (1981–1982), and Y7 (1982–1983). Symbol Y represents the number of crop seasons covering the period from 1977–1978 to 1982–1983 under no occurrence periods of stag heads (B) (Kolte et al. 1986) �� 193

Fig. 11.1 Rates of O2 uptake of infected and uninfected cotyledons at various times after inoculation: (●-●), infected; (○-○), uninfected; A, visible signs of sporulation (Thornton and Cooke 1974) �� 216

Fig. 11.2 Chlorophylla plus chlorophyllb, content of infected, and uninfected cotyledons at various times after inoculation: (●-●), infected; (○-○), uninfected (Thornton and Cooke 1974) �� 216 List of Figures xxxv

Fig. 11.3 Carbohydrate content of the alcohol-soluble fraction of infected and uninfected cotyledons at various times after inoculation with Hyaloperonospora parasitica: ■ = infected; □ = uninfected; T trace (indicating that the peak height of the TMS derivative was indeterminable at an attenuation of 20 × 103 (Thornton and Cooke 1974) �� 217 Fig. 11.4 Principal carbohydrates of the alcohol-soluble fraction of sporangia from infected cotyledons and control washings, 7 days after inoculation with Hyaloperonospora parasitica. ■ = infected; □ = uninfected (Thornton and Cooke 1974) �� 218 Fig. 11.5 Conductivity changes of de-ionized glass-distilled water containing samples of uninfected cotyledons (...) and cotyledons infected (−) by Hyaloperonospora parasitica isolate from cauliflower ○( ) and oilseed rape (–) (Kluczewski and Lucas 1982). Each point represents the mean of four replicates �� 218 Fig. 11.6 β-glucosidase activity in extracts of control cotyledons (...) and cotyledons infected (−) by either cauliflower ○( ) or oilseed rape (−) isolate of Hyaloperonospora parasitica (Kluczewski and Lucas 1982) �� 219 Fig. 11.7 Acid ribonuclease activity in extracts of control cotyledons (...) and cotyledons infected (−) by Hyaloperonospora parasitica isolate from cauliflower ○( ) and oilseed rape (−) (Kluczewski and Lucas 1982) �� 219 Fig. 11.8 Peroxidase activity in extracts of control cotyledons (...) and cotyledons infected by either cauliflower ○( ) or oilseed rape (−) isolate of Hyaloperonospora parasitica (−) (Kluczewski and Lucas 1982) �� 220 Fig. 11.9 Simplified scheme of the biosynthesis of the defence-related compounds camalexin, salicylic acid, and lignin in Arabidopsis. Chorismate is the first branch point, since camalexin arises via tryptophan, while salicylic acid is synthesized via isochorismate and phenylalanine, and lignins arise via phenylalanine. Chorismate is converted into isochorismate by isochorismate synthase (ICS). Phenylalanine ammonia-lyase (PAL) converts phenylalanine into cinnamic acid and is specifically inhibited by aminoindan phosphonic acid (AIP). From cinnamic acid, the pathway branch to produce salicylic acid or, via cinnamaldehydes and monolignols, lignin. The conversion from cinnamaldehydes to monolignols by cinnamyl alcohol dehydrogenase (CAD) is inhibited by hydroxyphenyl-aminosulphinyl acetic acid dimethyl ester (OH-PAS) (Slusarenko and Schlaich 2003) �� 222 xxxvi List of Figures

Fig. 12.1 Relationship between mycelial development and host-cell necrosis estimated as granulation and browning of cells in (a) cauliflower and (b) oilseed rape inoculated with Hyaloperonospora parasitica isolates from cauliflower (CI) and oilseed rape (Rl). □-CI mycelial growth index; ■–CI necrotic cell index; □-Rl necrotic cell index. Bars indicate + standard deviation (Kluczewski and Lucas 1982) �� 228 Fig. 12.2 Time course of sporulation of Hyaloperonospora parasitica isolate from cauliflower (o) and oilseed rape ■( ) on cauliflower (- -) and oilseed rape (…). Bars indicate + standard deviation (Kluczewski and Lucas 1982) �� 229 Fig. 12.3 Linkage map of Arabidopsis chromosome 4 showing location of RPP-5, relative to cosmid (g), λ (m) RFLP markers, and RAPD (OP) markers, based on the segregation analysis of La-er x Col-0 Rls. (a) Mapping data derived from segregation analysis of RFLP markers on 100Rls and RAPD markers on 50Rls; (b) map position of RPP-5 relative to closely linked markers, from the analysis of 289 Rls (Parker et al. 1993) �� 244 Fig. 12.4 The Bayesian posterior distribution of the competition coefficient of two A. thaliana genotypes, Nd-1 (susceptible) and C24 (resistant), competing against each other. Solid line: without pathogens. Dashed line: with pathogens. Percentiles (2.5%, 50%, and 97.5%) in the posterior distributions: cC24, without pathogen (−0.23, 0.01, 0.31), with pathogen (0.40, 0.92, 1.60); cNd-1, without pathogen (0.99, 1.52, 2.19), with pathogen (1.28, 1.76, 2.34). The Bayesian posterior distribution of a parameter provides information on our degree of beliefs in the different possible values of the parameter. The mode of the posterior distribution corresponds to the maximum likelihood value, and increasing variance of the distribution corresponds to an increasing degree of uncertainty about the true value of the parameter (increasing experimental variation) (Damgaard and Jensen 2002) �� 248 Fig. 12.5 Distribution of the mapped RPP genes along the five chromosomes of Arabidopsis thaliana. To the left: a numerical list of the known 27 RPP genes with their chromosomal or MRC location given, where known. RPP3 is not mapped yet; thus, the Arabidopsis ecotype and the H. parasitica isolate are given. Underlined RPP genes have been renamed once. RPP genes with an asterisk have been cloned. To the right: graphical representation of the five List of Figures xxxvii

Arabidopsis chromosomes with chromosome 1 to the left and chromosome five to the right. The centromeres are shown as black boxes. Mapping markers are given to the left of the chromosome. The region of a MRC is indicated with a black bracket, and the locus of a specific RPP gene is shown with a black arrow head to the right of the chromosome (Slusarenko and Schlaich 2003) �� 253 Fig. 12.6 Schematic representation of RPP-conditioned signal transduction. H. arabidopsidis isolates are shown above the corresponding RPPs that recognize them below the isolates (hyphenated with the Arabidopsis accession from which they were cloned). RPP1 from Ws (RPP1-Ws) is dependent on PAD4 and EDS1. RPP 2A+B from Col-0 is quantitatively influenced by the resistance protein RPS5 that recognizes the bacterial pathogen Pseudomonas and requires SGT1b, PAD4, and EDS1. RPP4 from Col-0 is particularly sensitive to changes in resistance signalling. Thus, mutations in many defence-related genes lower the defence responses to EMOY and EMWA. RPP5 from Ler requires RAR1 and SGT1b as well as PAD4 and EDS1. Furthermore, RPP5 was shown to be dependent on LURP1. RPP8 from Ler is an exception, because it encodes a CC-NB-LRR-type resistance protein, yet is dependent not only on NDR1 and RIN4 (like other CC-NB-LRRs) but also on PAD4 and EDS1. No downstream signalling components of RPP13-Nd have so far been reported. Please note: Signalling by the TIR-NB-LRR-type RPP genes RPP1-Ws, RPP2A+B-Col-0, RPP4-Col-0, and RPP5-L-er was dependent on PAD4 and EDS1, which have a positive interlocked feedback regulation with salicylic acid (SA) [Slusarenko and Schlaich 2003; Yoshioka et al. (2006); Zhang et al. (2005); Zhang and Li (2005); Knoth and Eulgem (2008); Knoth et al. (2007)] (Schlaich and Slusarenko 2009) �� 260 Fig. 12.7 The UV absorption spectra of the diseased parenchyma cell wall (a), the vessel wall (b), and the healthy parenchyma cell wall (c) of the Japanese radish root (Asada and Matsumoto 1972) �� 263 Fig. 12.8 UV absorption spectra of the authentic compounds (a) and the degradation products (b) obtained from the extraction of paper chromatograms. I, P-hydroxybenzaldehyde; II, vanillin; III, syringaldehyde (Asada and Matsumoto 1972) �� 263 Fig. 12.9 Suggested pathway of lignin biosynthesis in healthy (full lines) and diseased (broken lines) plants. Px, Py, Pz: peroxidase isoenzymes x,y,z (Asada and Matsumoto 1972) �� 264 xxxviii List of Figures

Fig. 14.1 Rating scale for downy mildew (Hyaloperonospora parasitica) interaction phenotypes on Chinese cabbage (Williams and Leung 1981) �� 313 Fig. 14.2 Location of inoculum placement of eight pathogens in multiple disease screening of seedling Chinese cabbage. Pb = Plasmodiophora brassicae, Ec = Erwinia carotovora, Ac = Albugo candida, Pl = Phoma lingam, Ab = Alternaria brassicae, Hp = Hyaloperonospora parasitica, Xc = Xanthomonas campestris, and TuMV = Turnip mosaic virus (Williams and Leung 1981) �� 318 Fig. 14.3 Sequence for individual and multiple disease resistance screening in Chinese cabbage (Williams and Leung 1981) �� 319 Fig. 14.4 Inoculation of leaves with conidial suspension �� 330 Fig. 14.5 Inoculated leaves in the dew chamber �� 330 Fig. 14.6 Plants inoculation technique with fungicides �� 331 Fig. 14.7 Multiple inoculation methods �� 332 Fig. 14.8 Time course and environmental regime for multiple disease resistance screening of crucifer seedlings �� 333 List of Tables

Table 1.1 Commonly cultivated crucifers and Brassica species susceptible to downy mildew disease �� 2 Table 1.2 Biotic and abiotic stresses of crucifers (Saharan 1984, 1992; Kolte 1985; Williams 1985) �� 4 Table 1.3 Significant historical developments in the downy mildew research of crucifers �� 7 Table 1.4 Comparison of some ordinal, family, and generic classifications of downy mildews, white blister/rusts, and relatives �� 13 Table 1.5 Resistance genes (R genes) identified in crucifers (A. thaliana) against downy mildew (H. arabidopsidis) isolates (pathotypes) �� 22 Table 2.1 World records of Hyaloperonospora parasitica on crucifers (Saharan et al. 1997 updated) �� 36 Table 2.2 Percent avoidable loss in seed yield, 1000 seed weight, and percent oil content in different commercial varieties of mustard due to Alternaria blight, white rust, and downy mildew for 1996–1997 to 1998–1999 (pooled data) (Singh and Singh 2005) �� 42 Table 2.3 Reaction of seedling cotyledons of members of the Cruciferae to inoculation with Brassica and Raphanus forms of H. parasitica (Dickinson and Greenhalgh 1977) �� 44 Table 2.4 Performance of the Brassica form of H. parasitica on cultivars of B. oleracea (Dickinson and Greenhalgh 1977) �� 45 Table 2.5 Host species of H. parasitica (Channon 1981; Saharan et al. 1997 up dated) �� 47

xxxix xl List of Tables

Table 2.6 Downy mildew interaction-phenotype classes (Table 2.6) used for cotyledon and leaf-disc evaluation (Monterio et al. 2005) is presented below �� 50 Table 3.1 Measurements of Hyaloperonospora conidia on crucifers �� 71 Table 3.2 Measurements of Peronospora conidia on Chenopodiaceae �� 72 Table 5.1 Host differentials of Hyaloperonospora parasitica (Mehta and Saharan 1994) �� 130 Table 5.2 Response of seventeen Brassica species to nine isolates of Hyaloperonospora parasitica (Mehta and Saharan 1994) �� 131 Table 5.3 Conidial size of Hyaloperonospora parasitica isolates derived from eleven Brassica species (Mehta and Saharan 1994) �� 132 Table 5.4 Percent conidial germination of Hyaloperonospora parasitica isolates at 18°C (Mehta and Saharan 1994) �� 132 Table 5.5 Sources of seed for accessions of Brassica juncea, arranged in five groups according to the response of their seedlings at the cotyledon stage to Hyaloperonospora parasitica and one accession of B. napus (Nashaat and Awasthi 1995) �� 133 Table 5.6 Differential virulence of Hyaloperonospora parasitica isolates from B. campestris on six host lines (Moss et al. 1988, 1991) �� 134 Table 5.7 Brassica oleracea standard host differentials to classify pathotypes of Hyaloperonospora brassicae (H. parasitica) (Coelho et al. 2012) �� 135 Table 5.8 Mean interaction phenotypes in seedlings following cotyledon inoculations for different combinations of eight Brassica oleracea lines and thirteen European isolates of Hyaloperonospora brassicae (H. parasitica) (Coelho et al. 2012) �� 136 Table 5.9 Origin of the Hyaloperonospora brassicae (H. parasitica) isolates collected from field samples on different crop types of Brassica oleracea (Coelho et al. 2012) �� 137 Table 5.10 Interaction-phenotype scores used to evaluate the response of Brassica oleracea cotyledons and relative amount of sporulation following inoculation with Hyaloperonospora brassicae (H. parasitica) (Coelho et al. 2012) �� 137 Table 5.11 Isolates used for RAPD analysis (Tham et al. 1994) �� 139 Table 5.12 Identification of pathotypes of Hyaloperonospora parasitica and H. arabidopsidis �� 141 Table 6.1 Percentage of seed infection by Hyaloperonospora parasitica in Raphanus sativus �� 148 Table 6.2 Percentage of seedling infection by Hyaloperonospora parasitica and seed transmission in Raphanus sativus �� 148 List of Tables xli

Table 6.3 An analysis of sporulation of Hyaloperonospora parasitica on cabbage cotyledons at two temperatures and in free water or at atmospheric water potentials of or −30 bars �� 154 Table 9.1 Effect of temperature on infection by Hyaloperonospora parasitica and disease development on mustard seedlings (cv. RH-30) (Mehta et al. 1995) �� 189 Table 9.2 Effect of leaf wetness duration on infection by Hyaloperonospora parasitica and disease development on mustard seedlings (cv. RH-30) at 20 °C (Mehta et al. 1995) �� 190 Table 9.3 Effect of leaf wetness duration on infection by Hyaloperonospora parasitica and disease development on mustard seedlings (cv. RH-30) at 15 °C (Mehta et al. 1995) �� 190 Table 9.4 Prediction equations for the progress of downy mildew and white rust complex of rapeseed-mustard using different combinations of weather factors (Kolte et al. 1986) �� 192 Table 9.5 Effect of weather factors on development and severity of white rust, downy mildew, and Alternaria blight (Sangeetha and Siddaramaiah 2007) �� 194 Table 9.6 Correlation value between disease index of white rust, downy mildew, and Alternaria blight of Indian mustard with environmental factors (Sangeetha and Siddaramaiah 2007) �� 195 Table 9.7 Effect of planting time on the severity of downy mildew and white rust complex of mustard (Saharan 1984) �� 195 Table 9.8 Influence of planting dates on stag head incidence and severity of white rust and downy mildew of rapeseed and mustard in three rabi crop seasons starting from 1977–1978 to 1979–1980 (Kolte et al. 1986) �� 196 Table 10.1 Effect of downy mildew infection on yield, yield components, and oil contents of different cultivars of Indian mustard (Brassica juncea) (Meena et al. 2014) �� 203 Table 10.2 Estimation of economic loss due to downy mildew in Brassica juncea during 2010–2011 crop season in India (Meena et al. 2014) �� 204 Table 10.3 Interaction between Albugo candida and Hyaloperonospora parasitica during pathogenesis of B. juncea (Mehta et al. 1995) �� 204 Table 10.4 Effect of planting dates on the development of stag head due to downy mildew and white rust disease complex in Indian mustard cv. RH-30 during 1991–1992 crop season (Mehta and Saharan 1998) �� 208 xlii List of Tables

Table 10.5 Effect of planting dates on the development of stag head due to downy mildew and white rust disease complex in Indian mustard cv. RH-30 during 1992–1993 crop season (Mehta and Saharan 1998) �� 209 Table 10.6 Prediction equation for progression of white rust and downy mildew complex in relation to environmental factors during 1991–1992 and 1992–1993 crop seasons (Mehta and Saharan 1998) �� 210 Table 10.7 Correlation coefficient between white rust-downy mildew disease complex and meteorological parameters (Mehta and Saharan 1998) �� 210 Table 12.1 Expected sequence of events leading to hypersensitive reaction expression in crucifers to H. parasitica infection (Lebeda and Schwinn 1994) �� 231 Table 12.2 Inheritance of resistance in cauliflower to H. parasitica using Palermo Green model (Moss et al. 1988) �� 234 Table 12.3 Response of groups A, B, C, D, and E of Brassica juncea accessions and of one accession of B. napus at the cotyledon stage to infection with four isolates of Hyaloperonospora parasitica (Nashaat and Awasthi 1995) �� 235 Table 12.4 Examples of a successful selection for putative homozygous resistance response to Hyaloperonospora parasitica from a heterogeneous starting population of Brassica juncea at the cotyledon stage (Nashaat and Awasthi 1995) �� 235 Table 12.5 Reaction to Hyaloperonospora parasitica isolate P003

of F1, F2, and back cross F1 (BC1F1) progeny from crosses involving spring Brassica napus accessions RES-26 and Callypso (Nashaat et al. 1997) �� 236

Table 12.6 Reaction to Hyaloperonospora parasitica isolates of F1, F2, and back cross F1 (BC1F1) progeny from crosses involving Brassica juncea accessions RESBJ-190 and RESBJ-200 (Nashaat et al. 2004) �� 237 Table 12.7 Inheritance of resistance in crucifers to H. parasitica �� 239 Table 12.8 Recognition specificities of RPP genes (for each RPP gene the MRC or chromosomal location, the formerly assigned number (where applicable), the ecotype, and the isolate(s) recognized are given (Slusarenko and Schlaich 2003) �� 254 Table 12.9 Contribution of the various signalling components to the resistance reaction mediated by the various RPP resistance proteins (to be read from top to bottom for each RPP column) (Slusarenko and Schlaich 2003) �� 258 Table 12.10 Amounts of degradation products by alkaline nitrobenzene oxidation of the isolated lignin (Asada and Matsumoto 1972) �� 262 List of Tables xliii

Table 12.11 Elemental compositions and empirical formulae of the isolated lignins and the related compounds (Asada and Matsumoto 1972) �� 262 Table 12.12 Sources of resistance to Hyaloperonospora parasitica (Saharan et al. 1997 updated) �� 265 Table 12.13 Responses of phenylamide sensitive and insensitive isolates of H. parasitica to phenylamide fungicides (Moss et al. 1988) �� 271 Table 13.1 Effect of planting dates on the severity of white rust and downy mildew of Indian mustard cv. Varuna (Saharan 1992b) �� 286 Table 13.2 Efficacy, economics, and spray schedule of fungicides against downy mildew of mustard (Mehta et al. 1996) �� 287 Table 13.3 Efficacy of fungicidal treatments on the severity of downy mildew of cauliflower (Ryan 1977) �� 288 Table 13.4 Fungicides found effective against downy mildew of crucifers (Saharan et al. 1997 updated) �� 290 Table 13.5 Efficacy of fungicidal sprays on downy mildew of radish (Sharma and Sohi 1982) �� 294 Table 13.6 Efficacy of fungicidal treatments on the downy mildew of mustard in India (Saharan 1984, 1992a) �� 295 Table 13.7 Efficacy and spray schedule of fungicides against downy mildew of mustard during 1991–1992 and 1992–1993 crop seasons (Mehta et al. 1996) �� 296 Table 13.8 Comparative yield increase and cost-benefit ratio of fungicides used against downy mildew of mustard (Mehta et al. 1996) �� 297 Table 13.9 Efficacy of fungicides against stag head of mustard due to combined infection of white rust and downy mildew (Mehta et al. 1996) �� 297 Table 13.10 Persistence of metalaxyl in mustard foliage after seed treatment (Mehta 1993) �� 298 Table 13.11 Persistence of metalaxyl in foliage of mustard after foliar application (Mehta 1993) �� 298 Table 13.12 Persistence of metalaxyl in mustard foliage after seed treatment and foliar sprays (Mehta 1993) �� 298 Table 13.13 Safe period and residue half-life values of metalaxyl in mustard (Mehta 1993) �� 299 Table 13.14 Translocation of metalaxyl residues into mustard seed following different treatments at harvest (Mehta 1993) �� 299 Table 13.15 Effect of fungicidal treatments on percent disease intensity, seed yield, and avoidable yield loss for 1996–1997 to 1998–1999 (Pooled data) (Singh and Singh 2005) �� 300 xliv List of Tables

Table 13.16 Integrated disease management module (seed treatment, spray schedule, and fertilizer doses for the control of DM, WR, AB) and its significance in achieving higher yield of mustard during 1999–2000 to 2001–2002a (Kolte 2005) �� 303 Table 13.17 Some micronutrients as possible inducer for multiple disease resistance in rapeseed-mustard (Kolte 2005) �� 304 List of Plates

Plate 2.1 (a) Downy mildew growth on cotyledon leaves of rapeseed-mustard; (b) yellowish flecks on the upper surface of the leaf of mustard; (c) downy mildew growth on the mustard leaf; (d) initial growth of downy mildew on stag head; (e) inflorescence showing conidial growth of Hyaloperonospora; (f) close-up view of conidial growth on stag head �� 51 Plate 2.2 (a) Initial symptoms of downy mildew on cabbage leaf; (b) abaxial side of cabbage leaf showing initial symptom, (c) adaxial surface of cabbage leaf showing downy growth; (d)- close-up of leaf showing conidial growth of Hyaloperonospora; (e) drying of the leaf due to advance stage of downy mildew �� 53 Plate 4.1 (a) Electron micrograph of TS of epidermal cells of cabbage cotyledon at 6 h after inoculation showing appressorium (ap) and penetrating hypha of Hyaloperonospora parasitica in between the anticlinal walls (j) of host epidermal cells. In one of the cells, a haustorium was formed but the section only shows part of sheath(s). The penetration was cut obliquely, and part of the hyphal wall (arrow pointed) is shown, × 8200; (b) photomicrograph of whole mount of a cleared cabbage cotyledon at 6 h after inoculation showing appressorium (ap) formation predominantly at the junction line of epidermal cells × 313; (c) photomicrograph CTS of cabbage cotyledon at 6 h after inoculation showing penetration as in A. × 500; (d) photomicrograph of whole mount of a cleared cotyledon showing intercellular hypha and haustorium completely ensheathed × 840 (Chou 1970) �� 94

xlv xlvi List of Plates

Plate 4.2 (a) Electron micrograph of TS of epidermal cells of cabbage cotyledons at 6 h after inoculation showing intercellular hyphae at various stages of penetration to the outside of host epidermis. Arrow points at the spearhead-like thickening of hyphal tip, × 5400; (b) electron micrograph of part of outgrowing hypha in between two host epidermal cells showing dieback of hyphal tip and walling-off (arrow pointed) of apparently intact cytoplasm, × 18000 (Chou 1970) �� 95 Plate 4.3 (a) Electron micrograph of TS of epidermal cells of cabbage cotyledon at 6 h after inoculation. The penetration region was cut medially through showing the appressorium which is almost empty with cytoplasm migrating into the penetrating hypha, × 14400; (b) electron micrograph of TS of appressorium and part of host epidermal cells showing the mucilaginous sheath of the appressorium. Membranous boundary of the mucilaginous sheath is shown by arrow, × 6000; (c) electron micrograph of section of a haustorium initial in host epidermal cell. Pan of the neck of a fully grown haustorium is shown by its side. Note the hyphal wall is continuous with wall of the intercellular hypha at this stage, × 13800; (d) electron micrograph of a section of intercellular hypha and host epidermal cell showing pan of host wall in contact with hypha is swollen and partially eroded (arrow), × 17700 (Chou 1970) �� 96 Plate 4.4 (a) Electron micrograph of a section of a haustorium initial in host mesophyll cell (section slightly oblique to the penetration zone), × 16500; (b) electron micrograph of a section of a very young haustorium in host epidermal cell showing breakdown of host cytoplasm into large number of vesicles, × 18000 (Chou 1970) �� 99 Plate 4.5 Electron micrograph of a section of a haustorium in host mesophyll cell at 6 h after inoculation, x 1 2000 (Chou 1970) �� 100 Plate 4.6 Electron micrograph of a section of a haustorium in host mesophyll cell at 6 h after inoculation, showing the saclike sheath and numerous vesicles (arrow pointed) and intra-vacuolar vesicles (pointed out by double arrow) in the sheath matrix, × 8580 (Chou 1970) �� 101 Plate 4.7 (a) Electron micrograph of a section of part of haustorium neck and sheath, × 24600; (b) electron micrograph of a section of part of haustorium neck and sheath showing numerous vesicles (arrow pointed) and dense granules in the sheath matrix (Smx) and the dentate extensions List of Plates xlvii

(pointed out by double arrow) of the dense zone (z) of haustorium wall, × 33000; (c) electron micrograph of a section of the interface between haustorium and host cytoplasm showing a dense vesicle (arrow pointed) like the secretary body, × 33000; (d) electron micrograph of a section of haustorium sheath showing incorporation of host cytoplasm (arrow pointed) in the sheath matrix and numerous membrane-bounded vesicles both in host cytoplasm and the sheath matrix, × 33000 (Chou 1970) �� 102 Plate 4.8 (a) Electron micrograph of a section of haustorium in host mesophyll cell showing the vacuoles or pro-vacuoles possibly in the process of fusion with each other and also with the sheath (arrow), × 7200; (b) electron micrograph of a section of the interface between haustorium and host cytoplasm showing vesiculation of the host plasmalemma, × 48000; (c) electron micrograph of a section of haustorium in host mesophyll cell showing fusion of vacuoles in host cytoplasm and sheath formation, × 7200; (d) electron micrograph of a section of interface between haustorium and host cytoplasm showing the structure of outer dense zone of haustorium wall distinguished into two well-defined layers (z1) and (z2), × 49500; (e) electron micrograph of a section of intercellular hyphae showing the hyphal wall also exhibiting a dense outer layer composed of z1 and z2 × 33000 (Chou 1970) �� 103 Plate 4.9 (a) Electron micrograph of a section of haustorium in epidermal cell at 6 h after inoculation showing the typical fine structure of haustorium at this stage. Ring formation in mitochondria pointed out by arrow, × 13200; (b) electron micrograph section of haustorium in epidermal cell 45 h after inoculation, × 24000 (Chou 1970) �� 104 Plate 4.10 (a) Electron micrograph of a section of the interface between haustorium and host cytoplasm of mesophyll cell showing sphaerosome-like bodies (arrow pointed) in host cytoplasm, × 33000; (b and c) electron micrograph of sections of young penetrating hyphae, (b) showing complicated membrane system of unknown nature and (c) showing intra-vacuolar membrane systems, × 55200 and 36000, respectively (Chou 1970) �� 105 Plate 4.11 (a) Tangential section of the dense zone of haustorium neck showing foldings of host plasmalemma (arrow pointed) forming tubular extensions and incorporation of numerous dense granules (d), × 33000; (b) electron micrograph section of haustorium in host epidermal cell showing xlviii List of Plates

lomasome, × 55200; (c) electron micrograph of a section of haustorium in host epidermal cell showing pinocytotic vesicles formed from host plasmalemma and abundant porous substance (arrow pointed) at the host-parasite interface, × 73200; (d) electron micrograph section of interface between dense zone of haustorium and host cytoplasm showing the deposition of porous substance (arrow pointed), × 48000 (Chou 1970) �� 106 Plate 4.12 Schematic representation of a haustorium of Hyaloperonospora arabidopsidis in an Arabidopsis host cell (based on electron micrographs from Mims et al. 2004). C collar of host material (including callose), CC host cell cytoplasm, CV host cell vacuole, CW host cell wall, EHMa electron-dense extra-haustorial matrix, EHM extra-haustorial membrane, G Golgi body, H haustorium, HM hyphal membrane, HW hyphal wall, Ne neck region, PM host plasma membrane, HW hyphal wall, ICH intercellular hypha, L electron-dense lipid vesicle, M mitochondrion, N nucleus, Ne constricted neck region, P plastid, PM invaginated host cell plasma membrane, T tonoplast membrane of the host cell vacuole, V vacuoles in the haustorium, Ves vesicles either fusing with or budding off from the extra-haustorial matrix. �� 107 Plate 4.13 Ultrastructural features of the structures produced by Hyaloperonospora parasitica in susceptible accessions of Arabidopsis. Sections were taken from samples 3 (a–d), 5 (e), and 7 (f) days after inoculation. (a) Median section through a penetration site showing an intercellular hypha from which two haustoria penetrate two different host mesophyll cells of Ws-eds1. Note the presence of nucleus, lipid bodies, mitochondria, and large vacuoles in the intercellular hyphae. (b) Median section through a penetration point and two haustorial bodies in a host mesophyll cell of Ws-eds1. Note that the old haustorium (with haustorial neck) contains organelles such as mitochondria and nucleus. (c) Median section through a penetration point and haustorium in a mesophyll cell of Oy-0. Callose deposition (*) occurred at the penetration point around the proximal region of the haustorial neck. Note the presence of mitochondria and small and large vacuoles in the haustorium. The wall of the intercellular hypha is at its thickest where it penetrates the host cell wall to form the haustorial neck. The extra-haustorial matrix (arrows) is present around the haustorium. (d) Median section through a haustorium within the mesophyll cell of Ws-eds1. The cytoplasm List of Plates xlix

of the intercellular hypha and haustorium contains mitochondria, lipid bodies, and small and large vacuoles. The host mesophyll cell appears unaffected by the presence of the haustorium as organelles are well preserved. (e) and (f) Callose deposition in Oy-0 (e) and Ws-eds1 (f). Callose deposition stained lightly around the haustorium shown in (e) but densely around the haustorium and in the cell wall (*) shown in (f). Note that contents of the haustorium and infection hypha appear normal during the early stage of infection (e) as organelles are clearly distinguished in the host cytoplasm. The contents of the haustorium and the infection hypha (arrow) became necrotic at the late stage of infection (f). All bars ¼ 3 lm. H haustorium; IH intercellular hypha; IS intercellular space; m mitochondrion; n nucleus; ca callose; Cv cell vacuole (Soylu and Soylu 2003) �� 108 Plate 4.14 (a) Section of wax-embedded material showing a hyphal branch growing towards a stoma; (b) section of wax-embedded material illustrating two conidiophore primordia, one of which is beginning to grow; (c) stained and macerated preparation of an unbranched conidiophore; (d) stained and macerated preparation of a branched conidiophore; (e) a branched conidiophore with small spores in a stained and macerated preparation; (f) very young spores; (g) mature spores; (h) mature spores delimited by a cross wall (arrow); (1-L) frames from the cine film illustrating the development of conidiophores A, B, and C; (i) incubation time 3 h 30 min.; (j) incubation time 3 h 50 min.; (k) incubation time 4 h 10 min.; and (l) incubation time 4 h 30 min. A–H scale line is 10 μm; I–L scale line is 100 μm (Davison 1968b) �� 113 Plate 4.15 Continued development of conidiophores: (a) incubation time 4 h 50 min.; (b) incubation time 5 h 10 min.; (c) incubation time 5 h 30 min.; (d) incubation time 5 h 50 min.; (e) incubation time 6 h 10 min.; (f) incubation time 6 h 30 min.; (g) incubation time 6 h 50 min.; and (h) incubation time 7 h 30 min. Scale line is 100 μm (Davison 1968b) �� 114 Plate 4.16 Electron micrograph of conidia, germ tubes, and initial period of Hyaloperonospora parasitica invasion on Japanese radish leaves. (a) Mature conidium. The surface is rough, with wart-shaped structure; (b) separation of a mature conidium from its conidiophore; l List of Plates

(c) an appressorium above a stoma and a penetration peg into the stomatal cavity; (d) enlargement of C. Wrinkly structures on an appressorium in the initial period of formation; (e) an appressorium over a stoma 48 h after germination; (f) enlargement of E. Slight degeneration of the epidermal cells where the appressorium is in contact with the stomatal guard cells; (g) cuticular invasion. Germ tube growing from the side of a spore; (h) enlargement of G. The appressorium is quite contracted (Shiraishi et al. 1975) �� 118 Plate 4.17 Electron micrograph of initial period of Hyaloperonospora parasitica invasion on Japanese radish leaves. (a) Invasion through a junction between a stomatal guard cell and an auxiliary cell; (b) cuticular invasion of an auxiliary cell. The germ tube is quite extended, but invasion does not depend on a stoma being present; (c) enlargement of B. The viscous substance used by the appressorium to adhere to the epidermal cell wall is not very visible; (d) enlargement of C. The germ tube and appressorium are clearly contracted, and circular traces of where the penetration peg has entered can be seen in the epidermal cell wall; (e) cuticular invasion with a long germ tube; (f) cuticular invasion through a short germ tube. Although the conidium is adjacent to a stoma, germination has occurred from the conidium wall on the side away from the stoma, and cuticular invasion is taking place (Shiraishi et al. 1975) �� 119 Plate 4.18 Electron micrograph showing development of conidiophores and conidia of Hyaloperonospora parasitica on Japanese radish leaves. (a) Conidiophores invariably grow out of stomata, sometimes two at a time; (b) a conidiophore branching during the initial stage of new growth; (c) surface of a conidiophore during the initial stage of new growth, with a wavy structure; (d) an extended conidiophore with appearance of a crimp at the base; (e) initial stage of conidium formation. The tips of the conidiophore swell, forming conidia. The conidiophores and conidia have similar surface structures; (f) clusters of conidia that have matured and begun to take on a tuft-like shape (Shiraishi et al. 1975) �� 120 Plate 4.19 Electron micrographs showing conidiophores and conidia of Hyaloperonospora parasitica on Japanese radish leaves. (a) Conidiophores without conidia. The area at the top right is a relatively young diseased area, and exfoliation List of Plates li

of epidermal cell wax and cuticular material can be seen; (b) diseased area with advanced signs of disease. Wrinkles have appeared in the epidermis of the diseased area, and open stomata can be seen; (c) diseased area with many developed conidiophores; (d) diseased area with advanced symptoms of disease. A crimp in the base of the conidiophore is visible. Yeast-shaped fungi are also present; (e) stoma in a healthy area. It is formed of two stomatal guard cells and several auxiliary cells; (f) the base of the conidiophore is crimped, perhaps due to mechanical force exerted by the stoma. Wrinkles on the surface of the host are clearly visible (Shiraishi et al. 1975) �� 121 Plate 5.1 Random amplified polymorphic DNA (RAPD) from 16 isolates of crucifer downy mildew (Hyaloperonospora parasitica), lanes 2–13 are isolates from oilseed rape Brassica napus, and lanes 14–17 are isolates from cauliflower B. oleracea (Tham et al. 1994) �� 138 Plate 7.1 Growth of crucifer downy mildew, Hyaloperonospora parasitica, in cotyledon tissues of Brassica. (a) Intercellular hyphae forming club-shaped intracellular haustoria in host cells, stained with trypan blue (x 250); (b) fluorescence micrograph of similar preparation, stained with aniline blue, showing bright collars, presumed to callose-like material of host origin, at sites of haustorial penetration (x 280); (c) electron micrograph of intercellular hypha (I) and haustorium (H) in host cell (HC). A second haustorium can be seen in the same cell (x4200) and (d) cell wall encasement surrounding developing haustorium at site of attempted penetration. Such host cell responses are commonly seen during development of the pathogen in partially resistant hosts (x 10,500) (Lucas et al. (1995) �� 159 Plate 7.2 Infection process and vegetative growth of Hyaloperonospora parasitica in Arabidopsis thaliana strain Weiningen (Koch and Slusarenko 1990). All samples were taken at 18 h after inoculation unless otherwise stated. (a) A germinated conidium (c) with an appressorium (a) on the leaf surface; both structures are devoid of cytoplasm. Bar= 10 μm. (b) A penetration hypha (ph) at the point of entry between anticlinal walls (aw) of two epidermal cells. Note the opposition of material adjacent to the site of penetration (arrows). Bar= 10 μm. (c) The same infection site as in (E) but focused through to the epidermal cells. The penetration hypha (ph) lii List of Plates

has expanded, and the first haustorium (h) has been formed in one of the epidermal cells. Note the material localized at the haustorial neck (arrows). aw, anticlinal walls. Bar= 10 μm. (d) Germinated conidia (c) on the leaf surface. In the cases shown, the appressoria (arrows) were produced directly from conidia without the formation of a germ tube. Appressoria are positioned over anticlinal walls (aw). Bar= 10 μm. (e) Simultaneous formation of haustoria (arrows) in both epidermal cells. aw, anticlinal walls. Bar= 10 μm. (f) Encasements (arrows) in epidermal cells surrounding haustorial initials. aw, anticlinal walls. Bar= 10 μm. (g) Formation of multiple haustoria (arrows) in mesophyll cells. The penetration hypha (ph) is arrowed. Bar = 10 μm. (h) Branched intercellular hyphae (ih) with numerous haustoria in mesophyll cells. A trichome (t) on the leaf surface can be seen clearly. Sample taken 3 days after inoculation. Bar = 100 μm. (i) Fully expanded haustoria (h) in mesophyll cells. Sample taken 3 days after inoculation. Bar = 10 μm. (j) Normal and encased (arrows) haustoria (h). The encasement is deposited around the haustorial neck and body. Sample taken 7 days after inoculation. Bar= 10 μm �� 160 Plate 7.3 Infection process and vegetative growth of Hyaloperonospora parasitica in Arabidopsis thaliana strain Weiningen (Koch and Slusarenko 1990). (a) Hypersensitive reaction. Two necrotic epidermal cells adjacent to the penetration hypha (ph). Because of collapse of the epidermal cells, the outline of underlying mesophyll cells is visible (arrows). Bar= 10 μm. (b) Hypersensitive necrosis of a single epidermal cell after penetration of the hypha through the anticlinal wall. Letters a and c refer to appressorium and conidium, respectively. Bar= 1 μm. (c) and (d) Penetration hypha (ph) growing between palisade cells at two different depths of focus. Haustoria are not formed. The cytoplasm in the two cells in contact with the hypha differs in appearance from that of the surrounding cells. Note the reduction in diameter of the hypha towards the tip. Bar= 10 μm. (E) to (H) Development of the fungus at a single penetration site, documented by varying the depth of focus. Note that no haustoria are present. (e) Conidium (c), germ tube (gt), and appressorium (a). Bar= 10 μm. (f) The penetration hypha (ph) can be seen growing between the anticlinal epidermal cell walls. The cell contents adjacent to the penetration hypha appear granular (arrows). Bar= 10μm. (g) and (h) Growth List of Plates liii

of the penetration hypha (ph) in the mesophyll. Material has been deposited by cells at the point of contact with the fungus (arrows). Bar= 10 μm �� 163 Plate 7.4 Asexual, and sexual reproductive structures of Hyaloperonospora parasitica in, and on tissues of Arabidopsis thaliana strain Weiningen (Koch and Slusarenko 1990). Samples were taken 7 days after inoculation. (a) Conidiophore (cp) emerging from the leaf surface; the conidia are partly discharged. Conidiophore and trichome (t) are similar in length. Bar = 45 μm. (b) The base of a conidiophore (cp) and several discharged conidia lying on the leaf surface. Note the constriction of the conidiophore in the stomatal opening (arrow). Conidia have a smooth to slightly verrucose surface. Bar = 5 μm. (c) A conidiophore initial (ci) growing out of a stoma and branching. Two conidiophore initials are apparently growing out of the neighbouring stoma (arrows). Bar =10 μm. (d) An oogonium (o) with a paragynous antheridium (an) can be seen in the mesophyll. h and ih, haustorium and intercellular hypha, respectively. Bar= 25 μm. (e) An oogonium (o) with an antheridium (an) attached and mature oospores. The different structural layers of the mature oospores are clearly visible; h, haustorium; ih, intercellular hypha; osp, oospore; osw, oospore wall; ow, oogonial wall; pe, periplasm. Bar = 25 μm �� 164 Plate 7.5 Sporulation of Hyaloperonospora parasitica on leaves of Arabidopsis strain Weiningen (viewed under a Stereo Microscope). A lawn of conidiophores is present on the leaves (thick arrows); on petioles the conidiophores are formed singly (thin arrows) (Koch and Slusarenko (1990) �� 166 Plate 7.6 Ultrastructural features of the compatible interaction between the Emoy2 isolate and susceptible accession Oy-0 sampled 3 (a–c) or 5 dai (d). (a–c) Median sections through penetration points showing haustoria connected to large intercellular hyphae. Note the presence of nucleus, lipid bodies, mitochondria, small and large vacuoles in the intercellular hyphae, and haustoria. Cell wall appositions (asterisks) occurred at the penetration points. The wall of the intercellular hypha is at its thickest where it penetrates the host cell wall to form the haustorial neck. The host mesophyll cell appears unaffected by the presence of the haustorium as organelles are well preserved. (d) Shows the formation of haustorial ensheathment (arrow) liv List of Plates

during a compatible interaction. Bars 2 mm; ch chloroplast, Cv cell vacuole, H haustorium, IS intercellular space, IH intercellular hyphae, l lipid body, m mitochondrion, n nucleus, Pv pathogen vacuole (Soylu et al. (2004) �� 167 Plate 7.7 Incompatible interaction between Emoy2 and the resistant accession Ler-0, 1 (a and b), 2 (c), and 3 dai (d). (a) Penetration of host tissue between anticlinal walls of two epidermal cells. Note that epidermal cells contain electron-dense cytoplasm and distorted organelles, but the associated pathogen penetration peg (pp) remains intact. (b) A necrotic mesophyll cell with shrunken electron-dense cytoplasm (arrows). The haustorial body is totally necrotic. (c) Collapsed epidermal cells and extensive vacuolation (asterisks) with wall alterations in an adjacent mesophyll cell. (d) Typical necrotic mesophyll cell underlying a necrotic epidermal cell (arrow). Note that both the epidermal and mesophyll cell contain several distorted host organelles. Bars 2 mm; EC epidermal cell, H haustorium, IH intercellular hyphae, IS intercellular space, MC mesophyll cell, pp penetration peg (Soylu et al. (2004) �� 168 Plate 7.8 Incompatible interaction between Emoy 2 and the resistant accession Ws-0, 2 (a), 3 (b and c), and 5 dai (d). (a) A haustorium in a dead mesophyll cells. The plant plasma membrane (arrow) has dislocated from the plant cell wall, and the penetrated cell exhibits organelle disruption. (b) Shows a necrotic intercellular hypha and haustorium. Note that both host and pathogen contain electron-dense cytoplasm in which organelles are hard to distinguish. (c) A necrotic mesophyll cell containing a heavily encased haustorium. Both haustorium and connected hypha are necrotic. (d) Shows penetrated and nearby mesophyll cells. Note the upper necrotic mesophyll cell contains several distorted chloroplasts, electron-dense cytoplasm, and darkly stained wall apposition (arrows) along the cell wall. The cytoplasm of the adjacent cell is severely disorganized containing misaligned chloroplasts and nucleus. A callose-containing deposit (asterisk) is also present along the cell wall. Bars 2 mm; H haustorium, IH intercellular hyphae, IS intercellular space, MC mesophyll cell, ch chloroplast, n nucleus (Soylu et al. (2004) �� 170 Plate 7.9 Pathogen development and host cell responses during the intermediate interaction between Emoy-2 and the accession Col-0. (a–c) Sections through List of Plates lv

the penetration point of the haustorium showing accumulation of cell wall apposition 2 (a) and 3 dai (b and c). Note that cell wall appositions (arrows) develop at penetration points (a), spread along the plant cell wall (b), and gradually extend around the haustoria (c). (d–f) Show penetration of mesophyll cells associated with cell disorganization 5 (d) and 7 dai (e and f). In (d), the host plasma membrane (arrow) has retracted from the cell wall, and penetration is associated with vacuolation and cytoplasmic disorganizations as characterized by the accumulation of electron-dense deposits (asterisks) along the tonoplast. (e) Shows a very rare infection site at which an apparently viable haustorium is located within a collapsed mesophyll cell. The tonoplast of the penetrated cell has ruptured, and organelles have dispersed into the central vacuole. In (f), the penetrated cell, intercellular hypha, and haustorium are necrotic as illustrated by the accumulation of amorphous material in their cytoplasm. Bars, (a) 1 mm; (b)–(f) 2 mm; ch chloroplast, Cv cell vacuole, H haustorium, IS intercellular space, IH intercellular hyphae, m mitochondrion, n nucleus (Soylu et al. (2004) �� 171 Plate 7.10 Immunogold localization of callose during compatible and incompatible interactions between Emoy2 and the Arabidopsis accessions Oy-0 (a, and b), Col-0 (c), and Ws-0 (d). In (a) and (b) note that labelling is confined to the pathogen cell wall within the intercellular hypha (large arrows), around the haustorial body (small arrows) and collar (*) at the site of penetration, 3 dai. In (c), callose is detected in the material (arrows) ensheathing haustoria, and within the cell wall, 3 dai. In (d) very dense labelling is found within the cell wall apposition (arrow) in a cell adjacent to a necrotic mesophyll cell, 5 dai. Bars, (a) 2 mm, (b)–(d) 1 mm; Cv cell vacuole, H haustorium, IH intercellular hyphae, MC mesophyll cell (Soylu et al. (2004) �� 173 Plate 10.1 White rust pustules are surrounded by conidial mass of Hyaloperonospora �� 201 Plate 10.2 White rust-infected stag head with initial growth of downy mildew �� 201 Plate 10.3 Conidial growth of Hyaloperonospora covered the white rust pustules in stag head �� 202 lvi List of Plates

Plate 10.4 (a) Transverse section of leaf 3 days post-inoculation AC-HP or HP-AC showing mycelium in the intercellular |spaces. UEP upper epidermis layer, LEP lower epidermis layer, F fungus GMS × 66 × 8 approx. (Mehta et al. 1995). (b) Transverse section of mustard leaf 6 days post-inoculation (AC-HP) showing mycelium in the intercellular spaces. UEP upper epidermis layer, LEP lower epidermis layer, F fungus GMS × 66 × 8 approx. (c) Transverse section of mustard leaf 6 days post-inoculation (AC alone) showing mycelium in the intercellular spaces and developing white rust pustules. UEP, upper epidermis layer; F fungus, M mesophyll cells, WRP white rust pustules GMS × 66 × 8 approx. (d) Transverse section of mustard leaf 9 days post-inoculation (AC alone) showing fungal mycelium and white rust pustules with sporangia/sporangiophores on abaxial surface. UEP upper epidermis layer, F fungal mycelium, M mesophyll cells, SP sporangia/sporangiophores GMS × 66 × 8 approx. (e) Transverse section of DM-infected mustard inflorescence depicting fungal mycelium and haustoria in the cortical cells and pith. EP epidermis layer, F fungus, CR cortex cells, X xylary vessels, Ph phloem, P pith, HA haustoria; GMS × 66 × 8 approx. (f) Transverse section of white rust-infected mustard inflorescence showing fungal mycelium, haustoria, and oospores in the cortical, xylary vessels. OS oospores, P pith, HA haustoria, F fungus; GMS × 66 × 8 �� 206 Plate 12.1 Hyaloperonospora parasitica brassicae race 2. Entry of germ tube of the conidium, through (a) an epidermal cell and (b) a stoma. Mycelium in tissue of (c) the susceptible Chinese rape host and (d) the immune radish host. Legend: Sp, conidium; Ap, appressorium; IH, infection hypha; My, mycelium; Ha, haustorium; Sh, sheath; Ep, epidermis; St, stoma; Sp, spongy mesophyll tissue; and DC, dead host cells (Wang 1949) �� 227 Plate 12.2 Cotyledon tissue 4 days after inoculation with Hyaloperonospora parasitica cauliflower isolate stained with trypan blue and cleared in chloral hydrate. (a) Intercellular hyphae in oilseed rape cultivar Primor showing left, developing haustoria (arrows) in host mesophyll cells close behind the hypha apex, and right, necrosis of penetrated host cells in older regions of a hypha x 400; (b) intercellular hyphae in cauliflower cultivar VSAG forming abundant intracellular haustoria. Note absence of host-cell necrosis, x 400 (Kluczewski and Lucas 1982) �� 230 Chapter 1 Introduction

1.1 Crops and Their Distribution

Crucifers occupy prominent place in world’s agrarian economy as vegetables, oilseeds, feed, and fodder, green manure, and condiment. Oilseed Brassica, also known by their trade name of rapeseed-mustard, include Brassica napus, B. juncea, B. carinata, and three ecotypes of B. rapa (B. rapa var. brown sarson, B. rapa var. yellow sarson, B. rapa var. toria). Major crucifers susceptible to downy mildew disease grown all over the world are given in Table 1.1 with their botanical name, common name, and usages. Global production of oilseed Brassica crops exceeded 63.76 mt, making them the second most valuable source of vegetable oil in the world. The leading oilseed Brassica producers in the world are the European Union, China, Canada, and India (USDA 2015). Different forms of oilseed Brassica are cultivated throughout the world. Winter-type B. napus predominates in Europe, parts of China, and Eastern USA, while spring-type B. napus is cultivated in Canada, Australia, and China. Spring forms of B. rapa are now mainly grown in the Indian subcontinent. Winter-type B. rapa has largely been replaced by higher-yielding winter-type B. napus and spring crops in its traditional production zones. Only winter type of B. juncea is cultivated in the Indian subcontinent and has now been actively considered as an option in drier areas of Canada, Australia, and even Northern USA. In India, B. juncea predominates and is grown on an over 80% of the area under rapeseed-mustard crops. The goal of developing canola forms has been accomplished for B. rapa, B. napus, and B. juncea but remains as an important objective in B. carinata. Almost all rapeseed produced in Australia, Canada, and Europe, and to a very large extent in China, is now canola. The cultivation of canola rapeseed-mustard has just begun in India (Chauhan et al. 2010). Crucifer vegetable forms an important group of vegetable crops of the world. These include a wide array of crops that span numerous genera and species in the family Brassicaceae. However, cole crops belonging to B. oleracea, viz. cauliflower, cabbage, broccoli,

© Springer Nature Singapore Pte Ltd. 2017 1 G. S. Saharan et al., Downy Mildew Disease of Crucifers: Biology, Ecology and Disease Management, https://doi.org/10.1007/978-981-10-7500-1_1 2 1 Introduction

Table 1.1 Commonly cultivated crucifers and Brassica species susceptible to downy mildew disease Botanical name Common name Usages B. nigra Black mustard Condiment (seed), vegetable fodder (leaves) B. oleracea B. oleracea var. acephala Kale Vegetable (head) B. oleracea var. capitata Cabbage Vegetable (head) B. oleracea var. sabauda Savoy cabbage Vegetable (head) B. oleracea var. gemmifera Brussels sprouts Vegetable, fodder (stem) B. oleracea var. botrytis Cauliflower Vegetable (inflorescence) B. oleracea var. gongylodes Kohlrabi Vegetable, fodder (stem) B. oleracea var. italic Broccoli Vegetable (inflorescence) B. oleracea var. fruticosa Branching bush kale Fodder (leaves) B. oleracea var. alboglabra Chinese kale Vegetable (stem, leaves) B. rapa B. rapa subsp. oleifera Turnip rape Oilseed B. rapa var. brown sarson Brown sarson Oilseed B. rapa var. yellow sarson Yellow sarson Oilseed B. rapa var. toria Toria Oilseed B. rapa subsp. rapifera Turnip Fodder, vegetable (root) B. rapa subsp. chinensis Bok choy Vegetable (leaves) B. rapa subsp. pekinensis Chinese cabbage Vegetable, fodder (head) B. rapa subsp. nipposinica – Vegetable (leaves) B. rapa subsp. parachinensis – Vegetable (leaves) B. carinata Ethiopian mustard Vegetable, oilseed B. juncea Mustard Oilseed, vegetable B. napus B. napus subsp. oleifera Rapeseed Oilseed B. napus subsp. rapifera Rutabaga, swede Fodder Eruca sativa Rocket, taramira Oilseed, fodder(leaves) Raphanus sativus Radish Vegetable, fodder Raphanus raphanistrum Wild radish Fodder

Brussels sprouts, kohlrabi, and kale, are most susceptible to downy mildew disease. As these crops are grown in a wide array of climate and cropping systems, these require general or specific adaptation to specific situations. Varieties with varying maturity duration are required to escape frost (Canada) or late-season drought (Southern Australia) or to fit in multiple cropping sequences (India, China). Breeding programmes are also concerned with the cultivar suitability for existing or emerging management practices, e.g. herbicide resistance or mechanical harvesting, resistance to pod shattering, etc. (Kumar et al. 2015). Brassica species relationship has been given in Fig. 1.1. Crucifers are confronted with several biotic and 1.2 The Disease and Pathogen 3

Black Mustard

B. nigra 2n=16 BB

Ethiopian mustardIndian mustard B. carinata B. juncea 2n=34 2n=36 BBCC AABB

B. oleracea B. napus B. rapa 2n=18 2n=38 2n=20 CC AACC AA

Wild cabbage Oilseed rape/canolaTurnip/field mustard

Fig. 1.1 U’s triangle showing species relationship among different Brassica species (UN 1935) abiotic stresses (Table 1.2). Among biotic stresses (diseases), downy mildew ranks the third most widespread devastating disease all over the world (Kolte 1985; Saharan et al. 1997, 2005).

1.2 The Disease and Pathogen

The term ‘mildew’ was first used in the USA to denote a wide group of parasitic fungi with little in common except their appearance as a white or lightly coloured delicate outgrowths caused by the proliferation, and fructification of mycelium on the surface of green, and necrotic plant tissues. Downy mildew quickly adapted to European conditions when vine mildew was introduced from North America. Downy mildew or members of the family Peronosporaceae are a distinctive group of obligate plant pathogens classified within the Mastigomycotina in the Oomycete order Peronosporales. In the family Cruciferae, about 50 genera and more than 100 different species are susceptible to infection by downy mildew pathogen. Originally Gaumann (1918), on the basis of conidial measurements and cross- inoculation tests, recognized 52 species of Peronospora on crucifer hosts. Later studies by Yerkes and Shaw (1959) concluded that there are no reliable morphological 4 1 Introduction

Table 1.2 Biotic and abiotic stresses of crucifers (Saharan 1984, 1992; Kolte 1985; Williams 1985) Sr. No. Common name Symbola Pathogen or cause 1. Alternaria disease, black Ab Alternaria spp. (A. brassicae (Berk.) Sacc.; A. spot, leaf stem or pod spots, brassicicola (Schw.) Wiltsh.; A. raphani Groves and blight & Skolko; A. alternata 2. Anthracnose Ch Colletotrichum higginsianum Sacc. 3. Aster yellows AY Mycoplasma-like organism or phytoplasma organism (MLO) 4. Autogenic necrosis AN(G) Genetic disorder 5. Bacterial leaf spot Psm Pseudomonas syringae pv. maculicola (McCulloch) Young et al. 6. Bacterial soft rot, Erwinia Ec Erwinia carotovora pv. carotovora (Jones) stalk rot Bergey et al. 7. Bacterial soft rot, Pmm Pseudomonas marginalis pv. marginalia Pseudomonas rot (Brown) Stevens 8. Black leg and phoma root rot Lm Leptosphaeria maculans (Desm.) Ces. & de not. (anamorph: Phoma lingam (Tode: Fr.) Desm. 9. Black mould rot Rst Rhizopus stolonifer (Ehr.: Fr.) Vuill. 10. Black root Ar Aphanomyces raphani Kendrick 11. Black rot Xcc Xanthomonas campestris pv. campestris (Pammel) Dowson 12. Black speck BS Physiological disorder 13. Bottom rot, damping off, Rs Rhizoctonia solani Kuhn (teleomorph: head rot, seedling root rot, Thanatephorus cucumeris (Frank) Donk.) wire stem, basal stem rot 14. Broomrape Oa Orobanche aegyptiaca Pers. 15. Cercospora leaf spot Cb Cercospora brassicicola Henn. 16. Club root Pb Plasmodiophora brassicae Wor. 17. Crown gall At Agrobacterium tumefaciens (Smith & Townsend) Conn. 18. Damping off, Fusarium F Fusarium spp. 19. Damping off, Pythium PT Pythium spp. 20. Downy mildew, stag head Hp Hyaloperonosporaparasitica (Gaum.) Gokar 21. Grey mould Bc Botrytis cinerea Pers.: Fr. (teleomorph: Botryotinia fuckeliana (de Bary) Whetz. 22. Gummosis G Physiological disorder 23. Light leaf spot Pyb Pyrenopeziza brassicae Sut. & Rawl (continued) 1.2 The Disease and Pathogen 5

Table 1.2 (continued) Sr. No. Common name Symbola Pathogen or cause 24. Nematode (a) Awl D(N) Dolichodorus spp. (b) Cyst Hs(N) Heterodera schachtii Schmidt Heterodera cruciferae Franklin (c) Pin PA(N) Paratylenchus spp. (d) Root knot M(N) Meloidogyne spp. (e) Root lesion P(N) Paratylenchus pratensis (de Man) Filipjev Paratylenchus spp. (f) Sting B(N) Belonolaimus spp. 25. Phyllody PLO Phytoplasma 26. Pod malformation GP Genetical and physiological disorder 27. Powdery mildew Ec Erysiphe cruciferarum Opiz. ex. Junell 28. Ring spot Mb Mycosphaerella brassicicola (Duby) Lindau 29. Root rot Phymatotrichum Po Phymatotrichum omnivorum (Shear) Dug. 30. Root rot Phytophthora Pm Phytophthora megasperma Drechs. 31. Scab Sts Streptomyces scabies (Thaxter) Waksman & Henrici 32. Sclerotinia stem rot Watery Ss Sclerotinia sclerotiorum (Lib.) de Bary soft rot 33. Southern blight, root rot Sr Sclerotinia rolfsii Sacc. (teleomorph: Athelia rolfsii (Curzi) Tu & Kimbrough) 34. Stem blight No Nigrospora oryzae (Berk. & Brown) Peteb 35. Tip burn TB(P) Calcium deficiency 36. Verticillium wilt Vd,Va Verticillium dahliae Kleb. V. albo-atrum Reinke & Berth. 37. Virus diseases (a) Cauliflower mosaic CAM Cauliflower mosaic virus (V) (b) Mustard mosaic MM Mustard mosaic virus (c) Radish mosaic RM(V) Radish mosaic virus (d) Turnip mosaic TUM(V) Turnip mosaic virus (e) Yellows BMY(V) Beet western yellows virus 38. White rust, stag head Ac Albugo candida (Pers. Ex. Lev.) Kuntze (Hyaloperonospora sp. commonly present in stag head phase) 39. White leaf spot, grey stem Pc Pseudocercosporella capsellae (Ell. & Ev.) Deighton 40. Xanthomonas leaf spot Xca Xanthomonas campestris pv. armoraciae (Mc Culloch) Dye 41. Yellows Fo Fusarium oxysporum Schlecht. spp. 42. Chlorosis LC Loss of chlorophyll; S, Mg, N, P, K deficiency aPathogen symbol: G genetic disorder, P physiological disorder, N nematode, V virus 6 1 Introduction criteria for distinguishing Peronospora isolates from different host species, and all collections of downy mildew from the Cruciferae are currently grouped in the single aggregate species P. parasitica (Pers. ex Fr.) Fr. Constantinescu (1989) later proposed a new genus, Paraperonospora, to accommodate several species of Peronospora pathogenic on hosts in the family Compositae. Constantinescu and Fatehi (2002) splitted Peronospora into three separate genera, Peronospora s. str., Hyaloperonospora, and Perofascia, on the basis of morphology, ITS 1, ITS 2, and 5.8 S rDNA sequence analysis. The latest revisions are based on molecular phylogenetic methods. Interestingly, the latter two genera were found to be almost entirely restricted to a single host family, Brassicaceae. Within Hyaloperonospora, only six species were accepted because of the differences in morphology of conidia and conidiophores. This concept is similar to that of Yerkes and Shaw (1959) and de Bary (1863). There are two different patterns of host colonization: systemic and localized. Systemic infection is characterized by colonization of leaves, stems, and sometimes roots, mostly through the infection of the seedlings by primary inoculum. The symptoms vary from chlorotic discolouration to stunting and distortion of the whole plant. Localized infections are characterized by the occurrence of lesions on leaves, surrounded by a conspicuous characteristic white ‘down’ on the abaxial surface (Lucas and Sherriff 1988).

1.3 The Downy Mildew of Crucifers

The term mildew was first used in the USA to denote fungal growth on the surface of green and necrotic plant tissues. The word downy mildew came from the vine mildew introduced from North America which was quickly adapted by researchers of different countries. Persoon (1796) was the first person who described downy mildew of crucifers on Capsella bursa-pastoris. All isolates obtained from cruciferous hosts were ascribed to Peronospora parasitica (Pers. ex. Fr.) Fr. However, Gaumann (1918) named isolates of Peronospora affecting plants of Brassica species as P. brassicae Gaum. After several arguments and controversies among mycologists and taxonomists and following an extensive biometric study, over 80 species names were reduced to one synonym, and a single species, i.e. P. parasitica, was recognized on cruciferous hosts (Dickinson and Greenhalgh 1977; Hiura and Kanegae 1934; Waterhouse 1973; Yerkes and Shaw 1959). Based on molecular and morphological features, the large genus Peronospora was segregated by Constantinescu and Fatehi (2002) into two genera, i.e. Hyaloperonospora and Perofascia. Based on molecular, morphological, ITS1, ITS2, and 5.8 S rDNA sequence analysis and phylogenetic investigations, Hyaloperonospora was demonstrated to be the pathogen on cruciferous hosts with specificity of H. parasitica to Capsella bursa-pastoris, H. brassicae to Brassicaceae, and H. arabidopsidis to Arabidopsis thaliana (Choi et al. 2003; Goker et al. 2003, 2004, 2007, 2009a, Voglmayr 2003). Since, the first report of downy mildew disease on crucifers in 1796, periodic milestones set, and significant historical development in the downy mildew research have been outlined in Table 1.3. It is quite clear from the 1.3 The Downy Mildew of Crucifers 7 (continued) Milestones in discovery/first report of downy mildew report of downy Milestones in discovery/first (Persoon, 1796; Corda, 1837; Gaumann, 1918) reported for the first time on Capsella bursa-pastoris as a disease of crucifers was mildew Downy on crucifers were recognized (Gaumann, 1918). On the basis of conidial measurements and cross-inoculation tests, 52 species Peronospora Scand. 193, 1849 Veg. Sum. Fr. (Pers. Ex. Fr.) parasitica identified as P. was Pathogen and (Yorkes Fr. (Pers. Ex. Fr.) parasitica species P. from the crucifers were grouped in single aggregate mildew All collections of downy 1959) Shaw, 1918; (Butler, oilseeds and vegetables infections in Brassica mildew losses ranging from 50 to 60 percent were estimated due downy Yield et al., 1998; et al., 1997; Paul 1998; Davis 1992; Saharan, 1984, Koike, Achar, 1985; 1979; Kolte, 1976; Bains and Jhooty, Vasileva, Mahajan and Gill, 1993; Singh Singh, 2005) 1985; Saharan et al., 1997; 1958; Kolte, Vasudeva, 1918; on crucifers crops were described in details (Butler, mildew Symptoms of downy and 1963; Chou, 1970; Slusarenko 1952; Jafar, Weber, Sherf and Macnab, 1986; Ramsay Smith, 1961; Natti et al., 1956; Gram Schlaich, 2003; Larren et al., 2006) 1987; Ebrahini et al., disease assessment scales to assess leaf and stag head infection were suggested (Natti et al., 1967; Sadowaski, Different 1985; Kruger, Williams, 1994; Saharan, 1992; 1980; Nashaat and Rawlinson, 1976; Dickinson and Greenhalgh, 1977; Knight Furber, and Laing, 1992; Jensen et al., 1999; Monterio 2005; Coelho 2012) 1991; Brophy 1980; 1918; Fraymouth, 1956; Holliday, were described in details (Butler, structures of H. peronospora Reproduction and reproductive 1900) Wager, 1946, Channon, 1981; Preece et al., 1967; Walker, Asada, 1990; Chu, 1935; Ohguchi and and genetics were cytology, conidia, and conidiophore development, host-pathogen interface, Host penetration, haustorium development, 1968; Shiraishu, 1975; fine structures (Fraymouth, 1956; Chou, 1970; Davison, to reveal studied through light and electron microscopy 2003) and Soylu, Sansome and Sansome, 1974; Ehrlich Ehrlich, 1966; Soylu 1920; Kabel, 1921; Gaumann, pathogen on crucifers were determined (Gardener, mildew and specificity in the downy variability Pathogenic 1944; Felton and Wang, 1934; Thung, 1926; Lucas et al., 1988, 1994; Uknes 1992; Nashaat 1995; Hiura and Kanegae, 1926; 1963; Chang et al., 1964; Semb, 1969; Dickinson and Greenhalgh, 1977; Bains 1980; Dzhanuzakov, 1946; Natti, 1958; Knox-Davis, walker, 1994; et al., 1996; Moss 1991, 1994; Silue Nashaat and Rawlinson, 1983; Mehta and Saharan, 1994; Masheva and Jhooty, et al., 1996; Lackie and Ohguchi, 1998; Silve Yashida and Lucas, 1990, 1994; Hill et al., 1988; Sequeira Monteiro, 1996; Sherriff et al., 1998; Coelho 2012) Vishunavat et al., 2000; 1996; Rehmany reproduction in the pathogen (De Bruyn, 1937; Mc Meekin, 1960; Kluczewski for sexual Heterothallism and homothallism both were observed and Lucas, 1989; Sequeira Monteiro, 1996) and Lucas, 1983; Sherriff Significant historical developments in the downy mildew research of crucifers mildew in the downy Significant historical developments Period 1796–1918 1918 1959 1918–2005 1918–2006 1967–2012 1918–1990 1956–2003 1920–2012 1937–1996 Table 1.3 Table 8 1 Introduction Milestones in discovery/first report of downy mildew report of downy Milestones in discovery/first and perpetuation through mycelium, conidia, oospores; germination dispersal of conidia oospores were observed Survival 1970, 1981; Le beau, 1945; McMeekin, 1960; Jang and Safeeulla, (Gaumann, 1926; Chang et al., 1963; Jang and Safeeulla, 1990; Krober, Asada, 1989; Lin and Liang, 1974; Shao et al., 1990; Pinckard, 1942; Lin, 1981; Hartman 1993; Ohguchi and and Kolte, Vishunavat 1990; et al., 1998; Guo and Ohguchi, 1996) 1993; Paul 1998; Karuna and Kolte, Achar, 1995, 1998; Badul and Achar, et al., 1983; 1918; Preece et al., 1967; described (Butler, crucifers was pathogen in different mildew The process of infection and pathogenesis downy 1946; Jonsson, 1966; Le Beau, 1945; Chang 1992; Chu, 1935; Felton and walker, Achar, 1969; Chou, 1970; Shiraishi et al., 1975; Kroher, 2003) and Soylu, 1990; Soylu and Slusarenko, and Lucas, 1982; Singh et al., 1980; Koch et al., 1963; Jang and Safeeulla, 1990; Kluczewski (Le Beau, 1945; Chang et al., 1963; Shiraishi 1975; Lucas explained on cruciferous hosts was mildew of downy Disease cycle 1995; Saharan et al., 2005; Holub 1994) and leaf wetness on infection disease rainfall, initiated with the influence of temperature, humidity, was mildew Epidemiology of downy and Leung, 1981; Jonssen 1966; Williams 1972; Nashaat, 1997; Chou, 1970; Nakov, Walker,1946; (Chu, 1935; Eddins, 1943; development Alonso, Alaonso and 1998; Achar, Vladimirskaya et al., 1975; et al., 1986; 1979; Mehta et al., 1995; Kolte D’Ercole, 1975; Bains and Jhooty, 1995; Sangeetha and Siddaramaiah, 2007; Banerjee et al., 2010) et al., 1986; Mehta, 1993; recorded (Saharan, 1984; Kolte of disease in relation to planting time and host nutrition was The development 1946; Hammarlund, 1931; Petraitiene and Brazauskiene, 2005; and walker, 1935; Butler and Jones, 1949; Falton Townsend, 1928; Quanjer, 2004; Jiang and Caldwell, 2015) Verret, Sochting and et al., 1986; Mehta and Saharan, 1998) (Kolte prediction models were developed mildew Downy Wiese, 1918; recorded (Butler, with white rust especially at leaf and stag head stage was mildew infection of downy The association and mixed 1987; Mehta Verma, 1992; Choudhary and Verma, 1978, 1985; Chaurasia et al., 1982; Saharan and 1927; Boning, 1936;Bains and Jhooty, et al., 1995; Singh 2002; Saharan 2014) studied in relation to metabolic changes and role of natural biochemical compounds host Biochemistry of host pathogen interaction was et al., 1994; Ausubel, 1989; Delaney and and Lucas, 1982; Singh et al., 1980; Davis 1974; Kluezewski resistance (Thorntan and Cooke, and Schlaich, 2003) Mansfield, 2000; Scheideler et al., 2002; Slusarenko 1949; viz. pre-penetration and postinfection (Wang, levels, studied at various was mildew downy Mechanism of host resistance against Asada, 1991; Saharan et al., 1997) and Lucas, 1992; Ohguchi Kluezewski and Hammerschmidt, 1993; Lebeda model system (Davis - Hyaloperonospora studied using Arabidopsis Host-pathogen recognition system was and Schwinn, 1994; Holub et al., Reignault 1996; Joos 1996) (continued) Period 1926–1998 1918–2003 1945–2005 1935–2010 1928–2015 1986–1998 1918–2014 1974–2003 1949–1997 1993–1996 Table 1.3 Table 1.3 The Downy Mildew of Crucifers 9 (continued) Milestones in discovery/first report of downy mildew report of downy Milestones in discovery/first et al., 1996; plants treated with chemical inducers (Uknes et al., 1992; Lawton demonstrated in Arabidopsis Systemic acquired resistance was 1996) et al., 1992; Cao Hui 1998; Mauch–Mani and Slusarenko, et al., 1997; Clarke Zimmerli et al., 2000; Bowling by dominant genes (Natti et al., 1967; Hoser-Krause governed of host-pathogen relationship studies indicated that host resistance was Growth et al., 1991, 1995; Niu 1983; Bennet and Blancard, 1987; Lucas 1988; Nashaat 1996,1997, 2004; Je nsen 1999; et al., 2012) Vicente and Montario, 1996; Reignoult et al., 1996; Joos Dickson and Petzoldt, Carvalho Williams, 1983; Dickson and Petzoldt, 1996; Leung identified (Leung and was mildew or partial resistance to downy Quantitative 1983; Jensen et al., 1999) Williams, 1994; et al., 1989; Nashaat and Rawlinson, demonstrated (Greenhalgh and Mitchell, 1976; Rawlinson Biochemical basis of resistance was Daughty et al., 1995; Menard 1999; Glazebrook 1997) resistance (Asada and Matsumoto, 1969, 1972; Ohguchi et al., demonstrated to provide was Role of lignin formation in the host cell walls Asada, 1984; Matsumoto, 1994) Asada et al., 1975; Matsumoto and Asada, 1975; Matsumoto et al., 1978; 1974; Ohguchi and were identified in cruciferous crops which had been used and are being for breeding downy mildew Sources of resistance to downy and biotechnological techniques (Jonsson, 1966; Bonnet Blancard, 1987; Lucas et al., 1988; Through conventional resistance cvs. mildew 1995; Nashaat et al., 1997; Silue 1996; Saharan Ebrahimi 1976; Awasthi, 1994; Nashaat and Nashaat and Rawlinson, Greenhalgh and Mitcheli, 1976; Dickinson, 1975) disease management (cultural, chemical, biological resistance) including integrated strategies of crucifers, several mildew manage downy To 1985; and Bolton, 1996; Kolte, 1960; Schmidt, Sherf and Macnab,1986;Downy 1958; Conroy, Vasudeva, 1918; were suggested (Butler, White et al., 1984; Mehta and Nelson, 1977; Crute, 1984; 1957; Pauls Saharan, 1984,1992; Saharan et al., 1997, 2005; Kupryanova, 1967; Ryan et al., 1984; and Griffin, Whitewell 1987; Chann, 1981; Channon et al., 1970; Wafford, and 1996; Ryan, 1977; Chiu, 1959; Davies Thompson, 1959; Nicolas and Ark and et al., 1983; Sharma and Sohi, 1982; Mehta, 1993; Yang and Laing, 1992; McKay et al., Brophy 1940; Crute et al., 1985; Silue 1996; Shao 1991) Aggery, pathogen (Choi et al., 2003, 2005, mildew and molecular data were used to circumscribe species concept of downy investigations Phylogenetic et al., 2006; Landa Voglmayr et al., 2004, 2007; Scott 2004; Cunnington, 2006; Spring 2003; Goker Voglonayr, 2006, 2007; et al., 2007; Garcia-Blazguez 2008) on Capsella bursa- parasitica pathogen species identified and established were Hyaloperonospora mildew On cruciferous hosts, the downy 2002; Choi et al., 2003; thaliana (Constantinescu and Fatehi, on Arabidopsis on Brassicaceae, and H. arabidopsidis pastoris , H. brassicae 2003) Voglmaur, et al., 2003, 2004, 2007; Goker Period 1992–1998 1967–2012 1983–1999 1976–1999 1972–1994 1966–1997 1918–2005 2003–2008 2002–2007 10 1 Introduction

was cloned, encoding a protein et al., 2000). RPP13 was HpB ) (Rehmany ) that correspond with different cloned DM resistance ) that correspond with different , and ATR13 ATR8 , ATR5 , ATR4 ( HpA ) or Brassica provide evidence for highly conserved regulators (also found in monocots) and the likely regulators for highly conserved evidence PBS2 provide Milestones in discovery/first report of downy mildew report of downy Milestones in discovery/first described as a thaliana (At) was of DM resistance in Arabidopsis Natural variation Era of genomics and molecular genetics/molecular plant pathology. borne from an oospore in a seedling of was The DM isolate Emoy2 1990). and Slusarenko, (Koch model for molecular genetic investigation the first reference genome of downy and provide be used to establish genetics in the organism This isolate would thaliana ‘Columbia’. Arabidopsis notion of disease resistance being conferred by single R genes can parasites (Holub, 2006). Mutation of NDR1 demonstrates that H.H. Flor’s mildew This established a precedent for pathogens. diverse common links in the signalling of defence against actually be a multigenic process and involve (Century et al., 1995, 1997). Systemic laboratory experiments thaliana in comparative using oomycete and bacterial pathogens of Arabidopsis et al., 1995). Mutation of EDS1 acid (Lawton found to require salicylic thaliana was acquired resistance to bacterial disease and DM in Arabidopsis This gene and barriers (non-host resistance) to biotrophic parasites can be amenable mutation and genetic analysis. demonstrates that species level et al., 1996; proteins (Parker thaliana , were found to encode lipase-like which is also typically required for DM resistance in Arabidopsis PAD4, was cloned and found toThe first DM resistance gene RPP5 2004). et al.,1999; Jirage et al., 1999; Holub and Cooper, Glazebrook et al., 1997; Falk et al., 1997). Major R-gene clusters proteins (Parker receptor-like TIR-NBS-LRR class of cytoplasmic described encode a member of the previously thaliana populations combination of recombinant inbred Arabidopsis thaliana using a powerful on four chromosomes of Arabidopsis were revealed class of molecular a powerful homologues provide 1997). R-like probes to map RPP loci (Holub and Beynon, and DM isolates as physiological thaliana and in crops such as lettuce potato (Aarts et al., 1998a, b; Botella oomycete resistance genes in Arabidopsis for map-based new markers in DM resistance genes (TIR-NBS-LRR subclass) that differ locus RPP1 contains several The multicopy et al., 1997; Speulman 1998). proteins (Aarts et al., 1998a, b; Eulgem regulatory confer defence via different they vary in how specificity (Botella et al., 1998). DM resistance genes acid dependent and independent) defence responses (McDowell (salicylic A single DM resistance gene ( RPP7 ) can confer accumulative et al., 2004). distinct from thaliana appear to be phylogenetically et al., 2002; Eulgem 2007). DM isolates collected from Arabidopsis Tor et al., 2000; ( isolates: referred to hence as subsp. Arabidopsidis Brassica Arabidopsis thaliana (Bittner- gene in Arabidopsis of a receptor-like benchmark for allelic diversification the most extreme homologous to RPP8 and providing HpA enables geneticAn outcross of established in the At-HpA pathosystem. paradigm was Eddy et al., 2000; Rose 2004). ‘Gene-for-gene’ , ( ATR1 At-recognizable effectors independent for five evidence Hyaloperonospora parasitica renamed as Hyaloperonospora ) was parasitica Peronospora The DM parasite of crucifer species (previously genes (Gunn et al., 2002). involvement of proteolysis in defence signalling (Austin et al., 2002; Muskett et al., 2002; Tor et al., 2002; Tornero et al., 2002; Warren et al., 1999). A. Warren et al., 2002; Tornero et al., 2002; Tor et al., 2002; of proteolysis in defence signalling (Austin et al., 2002; Muskett involvement ) was ATR13 ( thaliana recognized effector The first Arabidopsis and Jensen, 2002). launched (Damgaard thaliana -oomycete molecular ecology was among UK isolates of of sequence variation a high degree cloned from HpA isolate Maks9 and found to encode a small secreted protein that exhibits resistance (DMR) mutants were described and launched the genetic analyses of induced mildew HpA (Allen et al., 2004). Enhanced downy of R genes has been expression Damme et al., 2005; Holub, 2006). Differential thaliana (Van accessibility for oomycete parasites in Arabidopsis (Xiao et al., 2016) downregulated and 8 are TDFs are upregulated been identified of which 121 TDFs have identified (Xiao et al., 2016). 129 (Constantinescu and Fatehi, 2002). SGT1b and RAR1 / (Constantinescu and Fatehi, (continued) Period 1990 to date Table 1.3 Table been arranged in chronological order after analysing the published research of crucifers have mildew in the downy The significant historical developments chapters in respective Reference details are given literature so far. 1.3 The Downy Mildew of Crucifers 11 milestones set by Gaumann (1918) on various aspects/areas of research which were further pursued and strengthened to comprehend the downy mildews on cruciferous crops all over the world by various researchers. The validation of Gaumann’s research by others on crucifer downy mildew warrants that Gaumann may be considered as father of crucifers downy mildew disease. The third largest group of downy mildews, which is mostly restricted to one plant family, is the brassicolous downy mildews (BDM), even though a few species of this group are parasitic to other plant families, such as Capparaceae, Resedaceae, Limnanthaceae, Cistaceae, and Zygophyllaceae (Constantinescu and Fatehi 2002; Goker et al. 2009b). The BDM contain about 100 known species (Constantinescu 1991), but there are several species level clades which have not yet been formally described (Goker et al. 2009b). Importantly, it also includes an economically relevant but yet undescribed species occurring on arugula (or rocket; Eruca sativa). Without quarantine regulations in effect, it is fast-spreading throughout the world (Goker et al. 2009b; Koike 1998). The BDM contain two genera, Hyaloperonospora and Perofascia, of which the monotypic genus Perofascia is characterized by sporangiophores that often intertwine, thick-walled oospores, and hyphal haustoria. The genus seems to be restricted to the Brassicaceae tribe Lepidieae (Constantinescu and Fatehi 2002). On plants belonging to the same tribe, but also on plants of about 20 other tribes of Brassicaceae, the genus Hyaloperonospora can be found. Hyaloperonospora is characterized by treelike sporangiophores, comparatively thin-walled oospores, and globose to lobate haustoria. Like Perofascia, this genus was segregated from Peronospora only after molecular phylogenetic analyses were able to prove distinctiveness from Peronospora (Constantinescu and Fatehi 2002; Riethmuller et al. 2002; Voglmayr 2003), rendering them the first two downy mildew genera described with the aid of molecular phylogenies. The genus Hyaloperonospora is the third largest genus of downy mildews, containing more than 100 species, which can infect economically important Brassicaceae crops, e.g. horseradish (Armoracia rusticana), mustard greens (Brassica juncea), rapeseed (B. napus), cabbage (B. oleracea), Chinese cabbage (B. rapa), arugula (Eruca sativa), wasabi (Eutrema japonicum), watercress (Nasturtium officinale), and radish (Raphanus sativus). The downy mildew of Arabidopsis thaliana and Hyaloperonospora arabidopsidis (Goker et al. 2009b) has become a model organism to dissect plant pathogen interactions (Baxter et al. 2010; Coates and Beynon 2010). The BDM have been monographed by Gaumann (1918), and this almost 100-year-old work is still the most important source of reference for this group, together with the list of downy mildew names at that time placed in Peronospora by Constantinescu (1991). Since this last monograph, only few new species of this genus have been published (Voglmayr et al. 2014a), despite the fact that numerous new species-level clades have been found in phylogenetic analyses (Goker et al. 2009b; Thines and Choi 2016). The downy mildew of crucifers is a very widely destructive disease all over the world wherever cruciferous plant species, cultivated or wild, are available (Table 1.1). The disease is very devastating causing yield losses in oil-yielding Brassica crops and cruciferous vegetable crops ranging from 50 to 100% depending 12 1 Introduction upon amount of pathogen inoculum present in the soil or near vicinity of host, favourable environmental conditions present for infection, and development, cultural practices adopted, and disease management practices followed. The information generated by the dedicated research workers so far on downy mildew of crucifers have been arranged in the present book in 16 chapters with appropriate headings and subheadings in numerical order. The present manuscript on downy mildew of crucifers deals with the various aspects, viz. the disease and its synonymous; geographical distributions and symptomatology on oil-yielding Brassica crops, cruciferous vegetables, and other economically important host plants; host ranges; yield losses and disease assessment scales/procedures; the pathogen, its taxonomy, and classification, phylogeny, and pathogenic variability; pathogen morphology, sporulation, perpetuation, and germination; host-parasite interaction; seed infection, process of infection, and pathogenesis; disease cycle; epidemiology and forecasting; fine structures observed though light and electron microscopy; biochemical changes during host-parasite interaction; host resistance, its sources, and genetical, biochemical, morphological, and histological mechanism of inheritance; molecular aspects dealing with cloning, mapping, and identification of resistance genes; disease management practices like cultural, chemical, biological, and integrated control; and standardized techniques on all aspects have been included. Chapter 15 deals with the future priorities of research on downy mildew of cruciferous crops for resolving unanswered questions by the researchers in the time to come. In the last Chapter, 16, subject index has been given to facilitate the readers to search desired information included in different chapters of the book.

1.4 The Pathogen/Causal Organism of Downy Mildew of Crucifers

In the literature published up to 2002, the pathogen/causal organism of downy mildew of cruciferous crops has been referred as Peronospora parasitica (Pers. Ex. Fr.) Fr under a very large genus Peronospora infecting large number of diverse kinds of plant families. In the present book, to keep the coherence of text, the pathogen has been referred as Hyaloperonospora parasitica (Gaum.) Goker. Based on molecular, morphological, and phylogenetical studies, taxonomists and mycologists have established three different species of downy mildew pathogen on crucifers, viz. Hyaloperonospora parasitica (Gaum.) Goker on Capsella bursa-pastoris, H. brassicae (Gaum.) Goker on Brassicaceae, and H. arabidopsidis (Gaum.) Goker on Arabidopsis thaliana. 1.5 Taxonomy and Classification of Downy Mildew Pathogen 13

1.5 Taxonomy and Classification of Downy Mildew Pathogen

The downy mildews are very large group of obligate parasite infecting the hosts of very diverse families of plants. The downy mildews (Peronosporaceae) in the traditional sense are morphologically diverse group which is mainly united by obligate parasitism in combination with more or less complex conidio- or sporangiophores with determinate growth. Even after recent molecular and phylogenetic analysis, it is still uncertain whether downy mildews are monophyletic or stem from different groups. The comparison of downy mildews order, family, and generic level by renowned taxonomists/mycologists is given in Table 1.4 including downy mildew

Table 1.4 Comparison of some ordinal, family, and generic classifications of downy mildews, white blister/rusts, and relatives Riethmuller et al. Goker et al. (2007), Waterhouse (1973) Kirk et al. (2001) (2002) Thines and Spring (2005) Peronosporales Peronosporales (no order name) Peronosporales Peronosporaceae Peronosporaceae Peronosporaceae Peronosporaceae Basidiophora Basidiophora Basidiophora Basidiophora Bremia Benua (Benua) Benua Bremiella Bremia Bremia Bremia Peronospora Bremiella Paraperonospora Graminivora Plasmopara Paraperonospora Peronophythora Hyaloperonospora Pseudoperonospora Peronospora (Peronosclerospora) Paraperonospora Sclerospora Plasmopara Peronospora Perofascia Albuginaceae Pseudoperonospora Phytophthora Peronosclerospora Albugo Albuginaceae Peronospora Peronospora Pythiaceae Albugo Plasmopara Peronospora Phytophthora Pythiales Pseudoperonospora Plasmopara Pythiogeton Pythiaceae Sclerospora Plasmoverna Pythium Halophytophthora Albuginaceae Protobremia Sclerophthora Peronophythora Albugo Pseudoperonospora Trachysphaera Phytophthora Pythiaceae Sclerospora Pythium Lagenidium Viennotia Trachysphaera Pythium (family not formally classified) Pythiogetonaceae (Pythiogeton) Phytophthora Pythiogeton (Trachysphaera) Albuginales Sclerosporales (Sclerophthora) Albuginaceae Sclerosporaceae Albugo Peronosclerospora Pustula Sclerospora Wilsoniana Verrucalvaceae Sclerophthora 14 1 Introduction of crucifers. The taxonomic classification ofHyaloperonospora parasitica causing downy mildew of crucifers is as follows:

Kingdom Mycota Subkingdom Eumycota Division Mastigomycotina Class Oomycete Order Peronosporales Family Peronosporaceae Genus Hyaloperonospora parasitica (Gaum.) Goker (Peronospora parasitica) H. brassicae (Gaum.) Goker on Brassicaceae (Peronospora brassicae) H. arabidopsidis (Gaum.) Goker on Arabidopsis thaliana H. arabidopsidis (Gaum.) Goker, Rieth:, Voglmayr, Weiss and Oberw [as Arabidopsis]. Mycol. Prog. 3(2): 89 (2004). Synonymy Peronospora arabidopsidis Gaum. (1918)

1.6 Current Generic Status of Downy Mildew of Crucifers

Generic concepts in downy mildews were (and still are) mainly based on conidio−/sporangiophore morphology in combination with conidial/sporangial morphology. Dichotomous versus monopodial branching of conidio−/sporangiophore, shape of the terminal branches, and presence of conidia or sporangia were the primary features used for genus classification. However, interpretation of these morphological features was not always unequivocal and dependent on the observer’s vision, which sometimes resulted in conflicting generic concepts and delimitation. With the availability of molecular phylogenies, it soon became apparent that current generic classification and circumscription contained numerous problems and had to be adapted if standards of phylogenetic classification were applied. Based on molecular and morphological features, the genera Hyaloperonospora and Perofascia were segregated from the large genus Peronospora (Constantinescu and Faheti 2002).

1.7 Species Concepts in Crucifer’s Downy Mildew

The species concept is probably the most controversial issue in downy mildew systematics, partly due to experimental difficulties to test it and partly due to its pro- found implications for researchers outside the systematic research community. In 1.8 Broad and Narrow Species Concepts 15 downy mildews, several species concepts were applied, which resulted in highly different numbers of accepted species depending on the criteria used. The main problem in species delimitation in downy mildews is that there are numerous indica- tions that, due to their obligate parasitism, they often have narrow host ranges and, therefore, represent genetically distinct species. On the other hand, host specificity is not always paralleled by morphological distinctness. Therefore, if morphology is used as a primary criterion for species definition, only a few species can be defined and accepted in many lineages, resulting in genetically heterogeneous species. Historically, two approaches were commonly followed, which were both mainly based on host ranges: the splitting approach of Gaumann (1918, 1923) versus the lumping approach of Yerkes and Shaw (1959). Gaumann (1918, 1923) advocated a narrow species concept in Peronospora, based on his results of cross-inoculation studies and minute morphological differences. Each species was usually assumed to be confined to one host genus or even a few host species (one host-one species concept). Conversely, Yerkes and Shaw (1959) argued that host specificity is neither sufficient nor suitable for the recognition of a species without clear-cut morphological differences. As a consequence, the numerous Peronospora species were recognized on Brassicaceae and Chenopodiaceae each into a single species (Peronospora parasitica and P. farinosa, respectively), resulting in a wide one host family-one species concept. Both the splitting and the lumping approach have sincere shortcomings. Using the narrow species concept, identification of morphologically similar species is often difficult or impossible without correct identification of the host. In addition, high host specificity has rarely been conclusively demonstrated, weakening the primary underlying assumption of the narrow species concept. In a wide species concept, there is the problem that genetically distinct or even unrelated entities may be classified in the same species, raising incorrect assumptions on biology and host ranges. This is especially problematic if host jumps are common and parasitism on the same host family has evolved multiple times, resulting in polyphyletic species. However, due to its easier applicability, the approach to classify all accessions of a given host family within a single species was widely followed by phytopathologists and molecular biologists.

1.8 Broad and Narrow Species Concepts

With respect to species concepts in downy mildews and in particular in the genus Peronospora, there have long been two conflicting views – the narrow species concept, advocated by Gaumann (1918, 1923) and Gustavsson (1959), and the broad species concept advocated by de Bary (1863) and Yerkes and Shaw (1959). While the narrow species concept was followed by most taxonomists, the broad species concept, which ascribed the host specificity of some downy mildew pathogens to specialized forms (formae speciales) of the same species, was mostly followed by applied plant pathologists. This schism made the sequencing of the Arabidopsis 16 1 Introduction downy mildew seems to be of direct importance to plant pathologists working on economically important Brassicaceae crops. However, due to the fact that the downy mildew species are only distantly related, even though the genome of Hyaloperonospora arabidopsidis has provided important insights into downy mildew evolution (Baxter et al. 2010), the findings are difficult to translate into applications in Brassica crops. In general, molecular phylogenetic analyses have provided solid evidence for a high degree of specialization for most downy mildew species, also including the genera Hyaloperonospora (Goker et al. 2009b; Voglmayr et al. 2014a) and Peronospora (Belbahri et al. 2005; Choi et al. 2007b, 2008, 2009, 2010, 2015; Thines et al. 2009; Voglmayr et al. 2014b). There are only rare exceptions – a few downy mildew species seem to have broad host ranges (Kenneth 1981; Runge et al. 2011). However, in line with molecular phylogenetics, infection trials have shown that downy mildews are generally highly host specific (Byford 1967; Gaumann 1918, 1923; Lebeda and Syrovatko 1988; Sherriff and Lucas 1990). Thus, it seems reasonable to treat emerging downy mildew diseases on new hosts as separate species – especially with respect to quarantine – until infection trials and phylogenetic investigations have revealed whether or not they are highly host specific. Had this been done in the past, it might have been possible to restrict pandemic downy mildew agents, such as Peronospora belbahrii from sweet basil, Peronospora somniferi from opium poppy, Peronospora salvia officinalis from sages, and Plasmopara obducens s.l. from cultivated Impatiens species, preventing high yield losses around the world (Thines and Choi 2016).

1.9 Use of Molecular Data for Downy Mildew Species Concept

Recently, molecular phylogenetic investigations have enabled the evaluation of the species problem using new perspectives and have led to the shift from a morphological to a phylogenetic species concept. A biological species concept directly addressing mating barriers has never been applied to downy mildews due to sincere methodological difficulties, and it is unlikely that these can be overcome. Therefore, reproductive isolation can only be indirectly assessed, e.g. by genetic distance of sequence data. The impact of molecular data is manifold: (1) numerous additional characters are available for recognition and distinction; (2) presence and amount of reproductive isolation can be assessed; (3) presence and amount of genetic distances provide indirect but strong evidence for host specificity and host ranges; (4) molecular data are less variable and prone to subjective interpretation than morphological data; (5) molecular data provide a sound basis for species identification even if morphological data are missing or incomplete; and (6) pathotypes or races, the basic entities for experiments in applied sciences, can be properly attributed to a species, and their phylogenetic relationships can be assessed. Therefore, in the absence of sound morphological characters, the species concept is increasingly based on 1.10 Hyaloperonospora Species on Crucifers 17 molecular evidence of reproductive isolation, which is a general tendency within mycology. Consequently, morphologically similar cryptic species are often recognized as distinct species if reproductive isolation and genetic distinctness can be demonstrated. However, evaluation of species boundaries by molecular data requires thorough sampling throughout the distribution area to assess genetic variability as well as reproductive isolation, and at best several molecular markers should be used for corroboration of species boundaries. Due to easy amplification and variability, the ITS rDNA region has been used in most investigations addressing the species concept in downy mildews and white blister rusts (Choi et al. 2003, 2005, 2006, 2007a, b, c, d; Voglmayr 2003; Goker et al. 2004; Scott et al. 2004; Cunnington 2006; Spring et al. 2006; Voglmayr et al. 2006; Landa et al. 2007; Garcia Blazquez et al. 2008). However, the mitochondrial COX2 region may also be a promising candidate to resolve species boundaries and for species identification (Choi et al. 2006, 2007d). Interestingly, the current evidence from molecular phylogenetic investigations often supports a narrow species concept as advocated by Gaumann (1918, 1923), although there are sometimes marked differences in detail.

1.10 Hyaloperonospora Species on Crucifers

According to Constantinescu and Faheti (2002), about 140 species names were published attributable to this genus. In their separation of Hyaloperonospora from Peronospora, Constantinescu and Faheti (2002) recognized only six morphologically distinct species. The accessions from most hosts of Brassicaceae were placed in Hyaloperonospora parasitica. However, subsequent molecular phylogenetic investigations demonstrated that the latter was a paraphyletic assemblage with respect to the other five Hyaloperonospora species and that many more species should be accepted based on the high genetic distances observed (Choi et al. 2003; Goker et al. 2003, 2004; Voglmayr 2003). Usually, these genetically distinct entities deserving species rank have a narrow host range and are confined to host genera or even species; however, in some cases, accessions from the same host do not form a monophylum (Armoracia rusticana; Goker et al. 2004). Therefore, it is problematic when species are determined solely on host association, as this is often but not always conclusive. The case study of Hyaloperonospora is also relevant for investigations at the molecular level of plant-pathogen interactions, as numerous studies are performed with the plant model organism Arabidopsis thaliana and its Hyaloperonospora parasite. The parasite is usually named H. parasitica, but it is genetically quite distinct from H. parasitica sensu stricto which is confined to Capsella bursa-pastoris (Goker et al. 2004); therefore, the name H. arabidopsidis should be used for the Arabidopsis parasite. 18 1 Introduction

1.11 Strategies to Breed Downy Mildew Resistance Cultivars of Crucifers

The incompatible interaction between host and pathogen results into agriculturally important resistant phenotypes. Thus, any strategies that can contribute to the incompatible interaction are potentially useful in plant disease resistance breeding. On the one hand, incompatible interaction (R) can be converted into compatible interaction (S) in the case that host loses the related genes in immunity or pathogen evolves new virulence effectors (genes), especially in the gene-for-gene interaction of race-specific resistance in crucifer host-patho (H. parasitica) system. On the other hand, compatible interaction (S) can be converted into incompatible interaction (R) when the host or the pathogen loses the function of certain genes that are essential for pathogenesis or plant/host gains novel resistance genes. Incompatible interaction due to the loss of function mutation in a certain host gene is often of high value for disease resistance breeding programmes because the resistance is usually durable and non-race specific. Based on current information generated and understanding on the molecular mechanisms of crucifers, H. parasitica interactions following strategies for developing downy mildew resistance cvs. of crucifers may be adopted.

1.11.1 Identification and Utilization of Receptor-Like Kinases Involved in Plant Immunity

The first layer of immunity, termed pathogen-associated molecular pattern (PAMP)- triggered immunity (PTI), is initiated upon the recognition of PAMP’s by plant pattern recognition receptors (PRR) at the cell surface (Dodds and Rathjen 2010); PTI is often phenotypically reflected by basal resistance that is able to prevent infection by diverse potentially pathogenic pathogen (Catanzarite et al. 2010). The second layer of immunity is triggered upon the recognition of specific effector proteins from invading pathogens by host immune receptor proteins traditionally called resistance (R) proteins. Because a R protein specifically recognizes one or a few pathogen effectors (and the recognized effectors are termed avirulence factors or Avr), effector-triggered immunity (ETI) typically endows hosts with race-specific resistance, also known as R gene-mediated resistance against well-adopted pathogen carrying the recognized Avr genes. PTI is highly conserved at or above the species level. ETI is often polymorphic within a particular plant species with some cvs. Being resistant and others being susceptible. Due to its importance and genetic ame- nability, ETI has been extensively studied, and R genes have been widely exploited in crop production. However, large-scale deployment of elite cultivars carrying an R gene (i.e. monoculture) imposes higher selection pressure on the pathogen carrying the cognate Avr gene to survive, resulting in the modification or depletion of the recognized Avr gene or generation of novel effector gene that can escape the 1.11 Strategies to Breed Downy Mildew Resistance Cultivars of Crucifers 19 recognition of the old R gene. This R-Avr interactive co-evolution explains why many R cvs. in field lose their resistance in a relatively short period of time. In this regard, exploiting new knowledge on the molecular mechanisms of PTI and durable and broad-spectrum resistance has become important in disease resistance breeding programme of crucifers host – downy mildew pathosystem (Jones and Dangl 2006; Li et al. 2013). Thus, screening and employment of receptor-like proteins involved in plant immunity appear to be very promising strategies for creating crop cultivars or germplasm with broad-spectrum and durable resistance in cruciferous crops.

1.11.2 Identification and Utilization of R Genes Involved in ETI

Most of the characterized R proteins involved in ETI belong to NB-LRR family and are extensively exploited in crop breeding and production. Some R genes contribute to broad-spectrum resistance because they confer resistance against a large part of strains of a pathogen. Identification of novel broad-spectrum resistance R genes involved in ETI and using molecular markers for such genes can highly improve selection efficiency in breeding programme for disease resistance. Whole-genome sequencing of plant pathogens in generating an increasing list of effector proteins that can facilitate the identification of new R genes in crop plants or in their wild relatives is essential. A strategy of ‘pyramiding’ R genes can be exploited in which several R genes each of which recognizes a specific range of strains of a pathogen are introduced into a single plant via marker-assisted selection (Xiao et al. 2008; Dangl and Jones 2001).

1.11.3 The Utilization of Quantitative Trait Loci (QTLs)

Resistance-associated QTLs provide abundant resources for disease resistance breeding because they generally render non-race-specific and durable resistance. Some resistance-related QTL genes can be directly used in breeding and production of cruciferous crops.

1.11.4 Screening and Utilization of Recessive Gene-Mediated Broad-Spectrum Resistance

In host-pathogen interaction, there are some host genes whose functions are required for the pathogenesis of certain pathogens. When mutation occurs in such a gene, host containing the mutant gene usually confers broad-spectrum or non-race- specific resistance to that pathogen. It is possible to screen for resistance mutants 20 1 Introduction from mutagenized susceptible plant populations under conditions favourable for pathogenesis of certain diseases (Li et al. 2013). The mutated genes can be used in the investigation of the underlying mechanisms and breeding programme. Identification of such genes can be a novel approach in crucifiers.

1.11.5 Engineering Broad-Spectrum Resistance Through Biotechnology

The mechanism of host-pathogen interaction has been more thoroughly investigated in model host plant Arabidopsis thaliana and then in other crucifers. NPR1 is a key regulator of the expression of pathogenesis-related (PR) genes; PR1, PR2, and PR5 are of disease resistance response termed as systemic acquired resistance (SAR) in Arabidopsis. Ectopic expression of NPR1 leads to broad-spectrum resistance or enhanced resistance against pathogens. The resistance obtained due to NPR1 – or NPR1 homologous – overexpression is usually associated with faster and greater expression of the PR genes (Xiao 2012; Lacombe et al. 2010; Cao et al. 1997, 1998; Mukhtar et al. 2009). Another potential resource of broad-spectrum resistance is RPW 8.2 that confers salicylic acid-dependent resistance to a wide range of pathogens at the host pathogen interfacial membrane in Arabidopsis. RPW 8.2 enhances the callose deposition and induces the H2O2 accumulation in the invaded cell to limit the invasion of the pathogen (Wang et al. 2007, 2010; Collier et al. 2011). RNA interference (RNAi)-based host-induced gene silence (HIGS) is another promising biotechnology to create resistance in cvs. by knockdown of either the host or pathogen-originated virulence-related regulators. The principle of this strategy is to express a small RNA in planta that can target genes of pathogen to suppress the virulence (Li et al. 2013). Mathematical modelling that combines ecological parameters (to explain spatial and temporal changes in population) with evolutionary genetics (natural selection acting on multiple loci on interacting species) represents an enormous challenge for breeding downy mildew resistance cvs. of crucifers.

1.11.6 Designation and Nomenclature of Downy Mildew Resistance Genes (R Genes) and Isolates (Races/ Pathotypes)

Breeders have interogressed disease resistance (R) genes from both cultivated and wild cruciferous plants in their efforts to produce more resistant varieties. Even so, new races of downy mildew pathogen regularly evolve through sexual reproduction of the pathogen that can overcome individual R genes. As per the term ‘gene-for- gene’ hypothesis, a plant to exhibit resistance (incompatibility) to a pathogen, a R gene must be present in the plant, and a corresponding avirulence (AVR) gene must 1.11 Strategies to Breed Downy Mildew Resistance Cultivars of Crucifers 21 be present in the pathogen. An absence of either leads to disease (compatibility). This led to the hypothesis (elicitor/receptor model) that R genes encode receptors that enable plant to detect the ingress of pathogens whose avirulence genes cause them to produce the corresponding legends. Thus, R-gene products might be expected to have two functions: first, molecular recognition, and second, activation of plant defence upon recognition. Historically, and as per convention, R genes have been designated in different host-pathogen interactions on the basis of name of a disease and/or a pathogen/host. To narrate some of the R genes, viz. Sr for stem rust, Lr for leaf rust, Yr for yellow rust, Pm for powdery mildew resistance of wheat, and WRR for white rust resistance of crucifers, are based on the name of the diseases caused in respective hosts. Resistant genes (R-genes) Hm1 confers resistance to maize leaf blight (Helminthosporium maydis, Cochliobolus carbonum), Xa 21 confers resistance to Xanthomonas of rice, Cf9, Cf2 confers resistance to Colletotrichum fulvum of tomato, RPP1, RPP5 confers resistance to Peronospora parasitica (H. parasitica) of crucifers are based on the names of the pathogens. R genes ATR1 and ATR13 conferring resistance to downy mildew pathogen (H. arabidopsidis) of Arabidopsis thaliana are based on the name of hosts generic and specific names (AT). In the past specificity loci (R genes) of A. thaliana were named as RPP loci (abbreviation of recognition of P. parasitica or else recognized by P. parasitica (AVR)) and were numbered consecutively (i.e. RPP1, RPP2, etc.). Specificity loci (AVR genes) of P. parasitica had been named ATR loci (abbreviation of A. thaliana recognized or else A. thaliana recognition) and were numbered the same as the corresponding RPP locus (R genes). This nomenclature is descriptive of an interaction regardless of which partner is actively recognizing the other. Ideally new loci (R gene) should be named strictly on the basis of genetic recombination. Unfortunate consequence of change of pathogen name from P. parasitica to H. parasitica to H. arabidopsidis is no longer intuitively connected with the downy mildew pathogen from its current name to recognize P. parasitica (RPP) gene designation. Such changes in the names of host (Sisymbrium thalianum (L.) Grey to Arabidopsis thaliana (L.) Heynh) and pathogen (P. arabidopsidis Gaum. to P. parasitica (Pers. ex. Fr.) Fr. to H. parasitica to H. arabidopsidis (Gaum.) Goker) may be confusing and irritating for students and researchers, but it is inevitable in this modern era of molecular genetics and phylogenetic analysis of living beings. However, there is a need to develop and adopt a standardized system and procedure for the designation of R genes. On the basis of host and pathogen (both) recognition which can reflect both in their interaction phenotype, i.e. ATHA1 and ATHA2 for recognition of R genes by H. arabidopsidis from A. thaliana; ATAC 1, ATAC 2, etc. for recognition of R genes by A. candida from A. thaliana; and BJHP 1 and BJHP 2 for recognition of R genes by H. parasitica from B. juncea. The downy mildew resistance genes (R genes) recognized by H. parasitica isolates (pathotypes) from A. thaliana accessions are given in Table 1.5. Similarly, for other crucifers, a uniform system may be adopted, viz. BNHP for B. napus-H. parasitica, BOHP for B. oleracea-H. parasitica, and BRHP for B. rapa-H. parasitica interaction phenotypes (R genes). Designation and nomenclature of pathogenic isolates (races/pathotypes) have gone through evolutionary process and methods. (1) Initially, physiologic races were 22 1 Introduction

Table 1.5 Resistance genes (R genes) identified in crucifers (A. thaliana) against downy mildew (H. arabidopsidis) isolates (pathotypes) Arabidopsis accessions R genes Downy mildew isolates (pathotypes) Col-0 (Columbia) RPP-4 EMOY 2, EMWA 1 Col-0 RPP 2 CALA-2 WS-0 (Wassilewskija) RPP 1A,B EMOY 2 Ler-0 (Landsberg erecta) RPP 5, RPP 8 EMOY 2, NOCO 2, EMWA 1 Ws-0 RPP 1A CALA 2 Ws-0 RPP 1A, B, C NOCO 2 Nd-1 (Niederzenz) RPP 13 (ATR 13Nd) MAKS 9, ASWA, EDCO, EMCO, GOCO Ws-0 RPP 1 (ATR 1 Ws B) MAKS 9 Rld 2 (Reschew) RPP 11 WELA 1 Col-0 RPP 6 WELA Col-0 RPP 7 HIKS Nd-0 RPP 25 AHCO Ler-0 RPP 27 HIKS Ws-0 RPP 10 NOCO, EMOY, MAKS, COLA Ws-0 RPP 14 NOCO, EMOY, MAKS Nd-1 RPP 26 WACO Nd-1 RPP 16 ASWA Nd-1 RPP 17 EMCO Ws-0 RPP 12 WELA Col-0 RPP 18 HIND Ler-0 RPP 23 GOWA Ler-0 RPP 21 MADI, MAKS Ler-0 RPP 22 ASWA Ler-0 RPP 24 EDCO Oy-0 (Oystese) RPP 3 CALA Wei-0 (Weiningen) RPP 9 HIKS Cola-0 RPP 19 HIND 4 Cola-0 RPP 20 WAND Cola-0 RPP 28 HIND 2 Tightly linked genes RPP 1- Ws A CALA, EMOY, HIKS, MAKS, NOCO RPP 1- Ws B RPP 1-Ws C generally designated as numbers or letters in an arbitrary manner in order of their discovery, i.e. Puccinia spp., Melampsora lini, Albugo candida, Peronospora parasitica, etc. (2) An improvement over the use of arbitrary numbers or letters was Black’s nomenclature in which the races were designated on the basis of their virulence on particular genes for resistance, i.e. an isolate of Phytophthora infestans attacking a potato cv. carrying the R genes, R1, was designated as race 1, the one attacking R4 as race 4, and an isolate attacking both R1 and R4 as race 1 and 4. The race which was avirulent on all the genes for resistance was designated as race 0. (3) Virulence formulae were proposed to designate races of stem, and leaf rust of wheat 1.12 Importance of Hyaloperonospora arabidopsidis in Molecular Plant Pathology 23 virulent or avirulent on particular genes for resistance, e.g. the formula 6,7,10/5,8,9, 9a, 11 for a race of P. graminis tritici, indicates that the race is virulent on Sr6, Sr7, and Sr10 but avirulent on Sr5, Sr8, Sr9 a, and Sr11. (4) A very complicated method was proposed by Habgood using binary and decanary values. (5) A virulence analysis method was suggested using mobile nurseries in case of powdery mildew of barley. Like with any other host-pathosystem, the designation and nomenclature of downy mildew of crucifers pathogenic isolates/races/pathotypes have not been standardized at International level. No standard method and procedure has been adopted. Each researcher has used his own vision and system to name the pathogenic isolates collected from different locations/countries from cruciferous host species/ varieties/accessions (Table 5.12). However, a naming system for the isolates of H. arabidopsidis from A. thaliana was introduced by Dangl et al. (1992), Holub et al. (1994), and Slusarenko and Schlaich (2003) on the basis of geographical location and ecotypes infected. As, for example, an isolate collected from suburb of Zurich called Weiningen and virulent on (among others) the ecotype Landsberg erecta was named WELA using the first two letters of the location where the isolate was found (WE), combined with the first two letters of susceptible ecotype (LA). Thus, NOCO was found in Norwich and is virulent on Columbia, EMWA at East Malling, and is virulent on Wassilewskija. The isolate EM (East Malling, UK), CA (Canterbury, UK); WE (Weiningen), CH, and NO (Norwich, UK) and the susceptible host line used for the isolates, third and fourth letters OY=OY-0, LA=LA-er, ND=Nd-0, CO=Col-0, etc have been used. New isolates collected from the same location and maintained on the same host genotypes were distinguished by a number (e.g. EMOY1 and EMOY 2). However, the system and procedure of naming of an isolate should reflect both host-pathogen interactions to recognize avirulence gene (AVR gene) of the pathogen along with R gene of the host, i.e. HPBJ 1, HPBJ 2, etc., indicating H. parasitica isolate/pathotype recognized R genes 1 and 2 from B. juncea after interaction of isolate (pathotype) HPBJ1 and HPBJ2. Like international code of botanical nomenclature for naming an organism, a pattern or system of designation and nomenclature of R genes and AVR genes should be developed at International level with code of conduct so that researchers can compare/confirm and validate each other’s results. It will avoid unnecessary repetition and confusion among Brassica scientists.

1.12 Importance of Hyaloperonospora arabidopsidis in Molecular Plant Pathology

Hyaloperonospora arabidopsidis is a prominent pathogen in natural populations of Arabidopsis thaliana (Coates and Beynon 2010; Holub 2008). As such, it was adopted in the 1980s as one of the two pathogens of Arabidopsis, together with the bacterium Pseudomonas syringae (Koch and Slusarenko 1990). The Top 10 ranking of H. arabidopsidis reflects the subsequent success of the Arabidopsis-H. arabidopsidis pathosystem. H. arabidopsidis was initially utilized as a ‘physiological probe’ of the Arabidopsis immune system (Holub et al. 1994). This research led to the cloning of the first plant diseaseR genes against an oomycete, better understanding 24 1 Introduction of the evolutionary dynamics of R genes, the definition of broadly important immune system regulators, the identification of downy mildew-resistant mutants, and genetic definition of the complexity of the plant immune signalling network (Coates and Beynon 2010; Lapin and Van den Ackerveken 2013; Slusarenko and Schlaich 2003). On the pathogen side, research is hampered by the lack of protocols for culture and genetic transformation, established techniques with other oomycetes such as P. infestans. However, work in the early 2000s led to the development of genetic maps and DNA libraries that enabled the discovery of the first avirulence effector (Allen et al. 2004) and later to the R X LR effector family (Rehmany et al. 2005). Genome sequencing of H. arabidopsidis isolate Emoy2, completed in 2010, unveiled 134 predicted RXLR effectors and other components of the H. arabidopsidis secretome (Baxter et al. 2010). Notably, this report also revealed important genomic signatures of obligate biotrophic that have evolved convergently in other obligate oomycete and fungal lineages (McDowell 2011). Protein interaction assays have shown that H. arabidopsidis effectors target highly interconnected host machinery, helping to define a representative plant-pathogen interaction network (Mukhtar et al. 2011). In addition, several high-throughput functional studies have investigated effector subcellular localizations, suppression of immune responses, molecular targets, and cognate immune receptors (Cabral et al. 2011, 2012; Caillaud et al. 2011; Fabro et al. 2011). Future studies with the H. arabidopsidis experimental system will include (i) direct or Agrobacterium-mediated transformation for genetic manipulation required for the molecular analysis of downy mildew pathogenicity; (ii) the establishment of the temporal hierarchy of effectors during penetration, colonization, and sporulation, which may serve as a blueprint for a better understanding of the molecular basis of biotrophy; (iii) the role of genetic recombination and epigenetic on the emergence of new effectors; (iv) the development of tools to understand how plant- originated molecules regulate pathogen response; and (v) the relevance of interspecies transfer of small RNAs. These investigations on H. arabidopsidis will continue to provide new insights into the molecular mechanisms of downy mildew pathogenicity and contribute to comparative and functional analysis of (obligate) biotrophic oomycete and fungal pathogens (Kamoun et al. 2015).

1.13 Impact of Climate Change on the Diseases of Crucifers

Climate change has become a household topic of discussion with more scientists getting involved in scientific research on the aspect, while politicians are trying to derive mileage from the paradigm. The last decade of the twentieth century and the beginning of the twenty-first century have been the warmest period in the entire global instrumental temperature record. Climate change is defined as any long-term significant change in the ‘average weather’ that a given region experiences, or in other words, it is the shift in the average statistics of weather for long term at a specific time for a specific region. Average weather may include temperature, 1.13 Impact of Climate Change on the Diseases of Crucifers 25 precipitation, and wind patterns. It involves changes in the variability or average state of the atmosphere over durations ranging from decades to millions of years. These changes can be caused by dynamic process on earth, external forces including variations in sunlight intensity, and human activities. Climate change in the usage of the Intergovernmental Panel on Climate Change (IPCC) refers to a change in the state of the climate that can be identified (e.g. using statistical tests) by changes in the mean and/or the variability of its properties that persists for an extended period, typically for decades or longer. It refers to any change in climate over time, whether due to natural variability or as a result of human activity (IPCC 2007).

Increased emission of carbon dioxide (CO2) and other greenhouse gases, predominantly methane (CH4) and nitrous oxide (N2O), has been ascribed as the main agents causing increase in global temperature. The second assessment report (AR2) of IPCC indicated that the increase of greenhouse gas concentrations leads to an additional warming of the atmosphere and the earth’s surface. Concentration of CO2 has increased from about 280 to almost 360 ppmv since preindustrial time, CH4 from 700 to 1720 ppmv, and N2O from about 275 to about 310 ppmv. This development is ascribed to the magnitude of human intervention mostly in terms of fossil- fuel use, change in land-use pattern, and agriculture. Global mean surface temperature has increased by 0.3–0.6 °C since the late nineteenth century, a change that is unlikely to be entirely natural in origin. The temperature increase is widespread over the globe and is greater at higher northern latitudes (http://www.ipcc. ch). According to IPCC, cold days and cold nights have become less frequent and hot days, hot nights, and heat waves more common. Rising temperature also affect the pattern of precipitation. Changes in rainfall pattern have already been noticed. The IPCC reports that the frequency of heavy precipitation has increased over most land areas, which is consistent with warming and increase of atmospheric water vapour. Based on the trends since 1900, precipitation significantly increased in eastern parts of North and South America, northern Europe, and northern and central Asia whereas declined in the Sahel, the Mediterranean, Southern Africa, and parts of southern Asia. Globally, the area affected by drought has increased since the 1970s. Effect of climate change on agriculture or more precisely on insect pests and diseases of agricultural crops is multidimensional. Magnitude of this impact could vary with the type of species and their growth patterns. With the change in the temperature and rainfall pattern, the natural vegetation over a region is facing a new phase of competition for survival. The fittest species are more likely to dominate in the changing pattern of climate. It may be assumed that the vegetation tolerating high temperature and salinity and having high CO2-use efficiency could fair better than other species. Any change in the managed vegetation system, i.e. agriculture and forestry, will directly affect the socio-economic implications of the regions involved. IPCC in its report of 1995 predicted that a double increase in the CO2 level will increase yield by 30% in several crops. The elevated production could be off-set partly or entirely by the insect pest, pathogens, or weeds. It is, therefore, important to consider all the biotic components under the changing pattern of climate. There is also thought about shorter winters, which may affect the oil yields of the rapeseed- mustard crops. 26 1 Introduction

World over research on effect of climate change on pests and diseases of crops is inadequate (Huda et al. 2005). In India, there is limited effort in this area for any insect pest or disease of any crop (Subba Rao et al. 2007; Chattopadhyay and Huda 2009). However, at the genomic level, advances in technologies for the high- throughput analysis of gene expression have made it possible to begin discriminat- ing responses to different biotic and abiotic stresses and potential trade-offs in responses. At the scale of the individual plant, enough experiments have been performed to begin synthesizing the effects of climate variables on infection rates, though pathosystem-specific characteristics make such synthesis challenging. At the population level, the adaptive potential of plant and pathogen populations may prove to be one of the most important predictors of magnitude of effects of climate change. Ecologists are now addressing the role of plant disease in ecosystem processes and the challenge of scaling up from individual infection probabilities to epidemics and broader impacts (Garrett et al. 2006). Swaminathan (1986) indicated that the number of diseases on the same crops was much higher in tropics than under temperate conditions to indicate how rising temperatures could impact occurrence of plant diseases on agricultural crops. Presently, most of the work related to climate change vis-à-vis plant diseases is going on in rice (blast, bacterial leaf blight), wheat (Puccinia, Septoria), and horticultural (Meloidogyne) crops. The trend indicates that severity of majority of diseases is found to be higher with elevated CO2 levels (Chakraborty et al. 2008), an off-shoot of climate change. It is also being opined that climate change could lead to a changed profile (variants) of pathogen, insect pest (‘climate change can activate “sleeper” pathogens, while others may cease to be of economic importance’ – Bergot et al. 2004). The facultative pathogens with broad host range may survive better. There is also possibility of broadening of host range of the facultative pathogens. The need for further work in this area has been highlighted in adaptation experiments using twice-ambient CO2, which increased the aggressiveness (Chakraborty and Datta 2003) and fecundity (Chakraborty et al. 2000) of Colletotrichum gloeosporioides, which causes anthracnose of tropical legumes. Elevated CO2 may modify pathogen aggressiveness and/or host susceptibility and affect the initial establishment of the pathogen, especially fungi, on the host (Coakley et al. 1999; Plessl et al. 2005; Matros et al. 2006). In most examples, host resistance has increased, possibly due to changes in host morphology, physiology, and composition. Increased fecundity and growth of some fungal pathogens under elevated CO2 have also been reported (Hibberd et al. 1996; Coakley et al. 1999; Chakraborty et al. 2000). However, it has been reported that greater plant canopy size, especially in combination with humidity, and increased host abundance can increase pathogen load (Manning and Tiedemann 1995; Chakraborty and Datta 2003; Mitchell et al. 2003; Pangga et al. 2004). Sporulation by the pathogenic fungi could be 15–20-folds higher, leading to massive increase in the pathogen (Mitchell et al. 2003). New strains may develop, with adaptation occurring faster, and their evolution may get accelerated (Coakley et al. 1999). Among the 27 diseases examined under elevated CO2 levels, 13 caused higher crop losses than expected. Ten of the diseases had a reduced impact, and four had the same effect as they do now (NSW DPI 2007). Low solar radiation and short-day periodicity could 1.13 Impact of Climate Change on the Diseases of Crucifers 27 result in higher infections by Fusarium, Sclerotinia, and Verticillium (Nagarajan and Muralidharan 1995). Root rot is an emerging threat for rapeseed-mustard production system, recently reported from the farmers’ field in some pockets of the country (Meena et al. 2010), which was initially identified as stand-alone and along with bacterial or fungal incidence or in combinations (Erwinia carotovora pv. carotovora, Fusarium, Rhizoctonia solani, and Sclerotium rolfsii). Keeping in view the fact that some isolates of Alternaria brassicae sporulated at 35 °C and several isolates had increased fecundity under higher RH, it seems that as per recent changes towards warmer and humid winters, being in line with current projections for future climate change (Waugh et al. 2003), existence of such isolates could pose more danger to the oilseed Brassica due to Alternaria blight in times to come. The immense variation available among 20 representative isolates of A. brassicae also indicates their ability to adapt to varied climatic situations (Meena et al. 2012). Powdery mildew (Erysiphe cruciferarum) disease in oilseed Brassica was mostly occurring in Gujarat state barring stray incidences elsewhere, and the appearance of the disease used to occur from late January onwards in other parts of the country. However, in recent times, the disease has been found to be occurring in other oilseed Brassica growing states, viz. Haryana, Central UP, MP, parts of Rajasthan, Jharkhand, and Bihar, with the disease making its appearance even in December possibly due to shortening of cold spell during the crop period. Bihar hairy caterpil- lar (Spilarctia obliqua) surprisingly on mustard has been noted to be on the rise. Oilseeds Brassica have been affected a lot by the painted bug (Bagrada cruciferarum) in the Western and by saw fly (Athalia proxima) in the Eastern India. Presently, the Indian Meteorological Department (IMD, GOI) and the National Centre for Medium Range Weather Forecasting (NCMRWF) in coordination with scientists from other agencies as ICAR, etc. are regularly issuing location-specific weather forecast and agrometeorological advisory as per different climatic conditions and cropping systems. The Indian Council of Agricultural Research (ICAR) recently launched National Initiative on Climate Resilient Agriculture (NICRA) in February 2011 to boost research on the impact of climate change and its mitigation at national level. The project aims to enhance resilience of Indian agriculture to climate change, climate vulnerability through strategic research, and technology demonstration. Research on adaptation and mitigation covers crops, livestock, fish- eries, and natural resource management. It also demonstrates site-specifictechnology packages on farmers’ fields for adapting to current climate risks. This will certainly enhance the capacity of scientists and other stakeholders in climate-resilient agricultural research and its application (http://www.icar.org.in). The mitigation of the adverse effect of climate is challenging. Acquaintances between pragmatic and modelling studies could prop up swift advancement in perception and prediction of climate change effects (Chattopadhyay et al. 2011). 28 1 Introduction

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2.1 Introduction

It is commonly known as ‘mildew’, ‘mould’, ‘false oidium or mildew’, mildiou des crucifers (French), ‘Falscher-Mehltau’: Kreuzbluetler (German), downy mildew (English), downy mildew of crucifers (English), mildiu des las cruciferas (Spanish), and more commonly by the name of downy mildew. The disease is caused by the fungus Hyaloperonospora parasitica (Gaum.) Goker. The nomenclature of the disease is based on symptoms on the infected leaves with downy fungal growth. The upper surface of affected young and older leaves have ill-defined, irregular, pale yellow necrotic lesions, whereas the lower surface is covered by white-grey mycelium. The cotyledons and older leaves may be killed prematurely when single lesions coalesce to form large blotches. Attacked pods may be covered with angular brown lesions, or under high humidity, a sparse white-greyish mycelium may develop. Severe attacks may lead to premature ripening. Geographical distribution, economic importance, symptoms on various hosts, host range, and disease assessment are discussed below.

2.2 Geographical Distribution

Downy mildew on cultivated Brassica species, and other cruciferous host species, is prevalent in widely separated localities in numerous countries throughout the world (Channon 1981; Verma et al. 1994; Saharan et al. 1997). World records of H. parasitica causing downy mildew disease on crucifers are given in Table 2.1. The names of various hosts in this table are as reported in the original papers.

© Springer Nature Singapore Pte Ltd. 2017 35 G. S. Saharan et al., Downy Mildew Disease of Crucifers: Biology, Ecology and Disease Management, https://doi.org/10.1007/978-981-10-7500-1_2 36 2 The Disease: Downy Mildew

Table 2.1 World records of Hyaloperonospora parasitica on crucifers (Saharan et al. 1997 updated) Recording Location year Host References Argentina 1939 Cabbage, radish, swede Lindquist (1946) Argentina 2006 Eruca sativa Larran et al. (2006) Australia 1924 Cauliflower, cabbage Samuel (1925) Austria 1969 Cabbage Glaeser (1970) 1987 Radish, Chinese cabbage Bedlan (1987) 1989 Cabbage Bedlan (1989) Bavaria 1936 Horseradish Boning (1936) Bermuda 1939 Stock Waterston (1940) Bhutan Crucifers Herb (IMI) Borneo 1962 Chinese cabbage Anonymous (1962) Brazil 1943 Cabbage and broccoli Viegas and Teixeira (1943) Britain (UK) 1948 Capsella bursa-pastoris Foister (1948) 1959 Broccoli, Brussels sprouts, cabbage, Moore (1959) cauliflower, kale, kohlrabi, marrow-stem kale, rape, turnips, radish, horseradish, swede, stock, wallflower, watercress Brunei 1981 Crucifers Channon (1981) Bulgaria 1979 Turnip Khristov (1979) Canada 1944 Cauliflower Jones (1944) 1961 Rape, crucifers Downey and Bolton (1961) 1993 Cruciferous vegetables Cerkauskas et al. (1998) Chile 1960 Crucifers Mujica and Vergara (1960) China 1925 Rape Porter (1926) 1957 Cabbage Pai (1957) Costa Rica 1967 Crucifers McGuire and Crandall (1967) Cuba 1973 Crucifers Fernandez (1973) Cypress 1981 Crucifers Channon (1981) Czechoslovakia 1968 Crucifers Rydl (1968) Denmark 1924 Crucifers Gram and Rostrup (1924) 1949 Stock Anonymous (1949) Dominica 1972 Crucifers Anonymous (1972a) Egypt 1996 Cakile maritima Baka (1996) (continued) 2.2 Geographical Distribution 37

Table 2.1 (continued) Recording Location year Host References Ethiopia 1981 Crucifers Channon (1981) Fiji 1969 Chinese cabbage Anonymous (1969) Finland 1981 Crucifers Channon (1981) France 1943 B. napus, turnip, Camelina sativa, Sinapis Darpoux (1945) alba Germany 1938 Colza Klemm (1938) 1939 Rape Raabe (1939) 1955 Cabbage Neumann (1955) 1995 Camelina sativa, false flex Foller et al. (1998) 2003 Arabidopsis thaliana Slusarenko and Schlaich (2003) Greece 1981 Crucifers Channon (1981) Guatemala 1950 Brassica spp. Muller (1950) Haiti 1972 Crucifers Anonymous (1972a) Holland 1924 Cabbage Thung (1926a) Hong Kong 1962 B. alboglabra, Chinese kale Johnston (1963) Hungary 1957 Stock Lehoczky (1957) Iberia (Spain) 1924 B. napus, B. oleracea Gonzalez (1924) India 1918 Brassica spp. Butler (1918) 1940 B. campestris, B. napus, radish, Eruca Thind (1942) sativa, Maledmia africana, Sisymbrium iris 1968 Capsella bursa-pastoris Rao (1968) 1976 Cardamine impatiens Sharma and Munjal (1977) 1980 Kohlrabi, kale Puttoo and Choudhary (1984) 1981 B. pekinensis Karwasara and Saharan (1982) 1982 Cabbage Gupta and Choudhary (1987) 1995 Cress Rakholiya et al. (1998) Iran 1989 Radish Etebarian (1989) 1993 Kohlrabi, red cabbage, turnip Sharifuabe and Nekoei (1995) Iraq 1981 Crucifers Channon (1981) Ireland 1970 Cauliflower McKee (1971) Israel 1953 Cabbage, cauliflower Peleg (1953) Italy 1961 Crucifers Ciferri (1961) Jamaica 1967 B. oleracea Leather (1967) (continued) 38 2 The Disease: Downy Mildew

Table 2.1 (continued) Recording Location year Host References Japan 1934 Brassica spp., crucifers Hiura and Kanegae (1934) Kampuchea 1969 Crucifers Soonthronpoet (1969) Kenya 1957 Kale Anonymous (1957) Korea 1972 Crucifers Anonymous (1972b) 1981 Chinese cabbage So et al. (1981) Libya 1981 Crucifers Channon (1981) Ludlow 1929 Swedes Preston (1929) Malawi 1972 Crucifers Peregrine and Siddiqi (1972) Malaysia 1949 B. rapa McIntosh (1951) Malta 1981 Crucifers Channon (1981) Mauritius 1950 Cabbage Orian (1951) Mexico 1983 Rapeseed Ponce and Mendoza (1983) Montpellier 1941 Stock Kuhnholtz and Gastuad (1943) Moravia 1928 Radish Baudys (1928) Morocco 1981 Crucifers Channon (1981) Mozambique 1948 Crucifers, cabbage De Carvalho (1948) Nepal 1966 Crucifers Bhatt (1966) Netherlands 1926 Crucifers, cabbage Thung (1926b) New South 1938 Cauliflower, mustard, kohlrabi, turnip Anonymous Wales (1938) (Australia) 1955 Cabbage Anonymous (1955) 1959 Brussels sprouts Anonymous (1960b) 1966 Stock Bertus (1968) New Zealand 1963 Crucifers Channon (1981) Norway 1969 Cabbage, Chinese cabbage, kohlrabi, kale, Semb (1969) red cabbage, rape, turnip, radish Palestine 1935 Cauliflower Rayss (1938) and Chorin (1946) Panama 1967 Crucifers McGuire and Crandall (1967) Pakistan 1969 Brassica, crucifers Perwaiz et al. (1969) (continued) 2.2 Geographical Distribution 39

Table 2.1 (continued) Recording Location year Host References Papua New 1981 Crucifers Channon (1981) Guinea Peru 1994 Lepidium meyenii Icochea et al. (1994) Philippines 1925 B. juncea, B. pekinensis Ocfemia (1925) Poland 1970 Crucifers Zarzycka (1970) 1990 Radish Madej and Majchrowicz (1998) Portugal 1953 Cabbage Da Costa and Da Camara (1953) Puerto Rico 1972 Crucifers Channon (1981) Queensland 1948 Coronopus didymus Langdon (1948) Romania 1930 B. napus, B. nigra, Capsella Savulescu and Rayss (1930) 1948 Wallflower Savulescu (1948) Russia 1989 Radish Timina et al. (1989) Sabah 1962 Crucifers Anonymous (1962) Samoa 1975 Crucifers Firman (1975) Sarawak - Crucifers Herb (IMI) Saxony 1927 Wallflower, stocks Wiese (1927) South Africa 1934 Cabbage, cauliflower, turnips, radish, Dippenaar (1934) kohlrabi Spain 1924 B. napus, B. oleracea Gonzalez (1924) 1991 Sisymbrium Sinobas and Dias (1995) Sri Lanka 1932 Crucifers Park (1932) Sweden 1931 Radish Hammarlund (1931) 1944 Colza, white mustard Bjorling (1944) 1952 Camelina sativa Borg (1952) Switzerland 1923 Brassica spp., crucifers Gaumann (1923) 1990 Arabidopsis thaliana Koch and Slusarenko (1990) Taiwan 1961 Crucifers Lo (1961) Tanzania 1981 Crucifers Channon (1981) Thailand 1962 Crucifers Chandrasrikul (1962) Trinidad and 1922 Cabbage Stell (1922) Tobago Turkey 1981 Crucifers Channon (1981) (continued) 40 2 The Disease: Downy Mildew

Table 2.1 (continued) Recording Location year Host References Uganda 1981 Crucifers Channon (1981) UK 1996 Crucifers, Cheiranthus cheiri Herb. (IMI), Fox (1996 ) USA 1883 Brassica spp., crucifers Farlow (1883) 1889 Sisymbrium spp., Lepidium Swingle (1890) 1903 Cauliflower Schrenk (1905) 1918 Turnip Gardner (1920) 1923 Cabbage Harter and Zones (1923) 1927 Watercress Davis (1929) 1932 Cabbage, crucifers, Brassica spp. Weber (1932) 1940 Horseradish Kadow and Anderson (1940) 1942 Cabbage Snyder and Baker (1943) 1954 Radish Thompson and Decker (1955) 1960 Brassica spp., crucifers Anonymous (1960a) 1998 Arugula (Eruca spp.) Koike (1998) Uruguay 1955 Crucifers Koch and Boasse (1955) USSR 1955 Cabbage Pimenova and Maslennikov (1955) Venezuela 1981 Crucifers Channon (1981) Victoria 1996 Crucifers, Brassica Smith and Price (1996) Vietnam 1966 Crucifers My (1966) Yugoslavia 1954 Cabbage Sutic and Klijajic (1954) 1961 Horseradish Macek (1961) Zimbabwe - Crucifers Herb. (IMI)

2.3 Economic Importance

The economic importance of Hyaloperonospora parasitica (downy mildew) has been adequately documented over the years. This pathogen, alone or in combination with Albugo candida (white rust), is responsible for causing severe losses in yield of several temperate and tropical Brassicaceae crops, particularly rapeseed and mustard. Yield loss due to downy mildew infection alone is very difficult to estimate, since in most cases it is always associated with white rust at inflorescence stage and with viral infection at seedling stage. 2.3 Economic Importance 41

2.3.1 Brassica Oilseeds

Hypertrophied host tissues termed as stag heads are often observed in association with a mixed infection of A. candida and H. parasitica particularly at the flowering stage. Yield losses in B. rapa var. Toria (Toria) due to such combined infections is estimated to be about 34%, when the average length of individual hypertrophied racemes is 10 cm (Kolte 1985). The combined infection with both pathogens on B. juncea may cause 37–47% and 17–32% reduction in silique formation and seed production, respectively (Bains and Jhooty 1979). Others have reported 23–55% yield loss in the same host species due to the mixed infection with both pathogens (Saharan 1984, 1992a). Kolte (1985) suggested the following formula for estimating the yield loss due to infection with white rust or downy mildew alone or for combined infections:

 BC×  QA=− ×100  A  where: Q = percentage yield A = average actual or expected yield of a healthy plant B = average or expected yield from the affected raceme, which is equal to the actual average yield from the corresponding length of the healthy raceme C = number of affected racemes per plant In California, USA, downy mildew of arugula (Eruca sativa) caused by H. parasitica was so severe that at times it reduced the crop quality to the extent that it could not be harvested (Koike, 1998). In the UK, the presence of weeds in oilseed rape was positively correlated with downy mildew (H. parasitica) severity (Davies et al. 1997). In Germany, in autumn, 1996, downy mildew of winter oilseed rape was so severe that at some location large areas were totally destroyed at the cotyledon stage (Paul et al. 1998). In India, Mahajan and Gill (1993) indicated the direct effect on gross weight of cauliflower curd by the downy mildew disease severity. Disease rating was negatively correlated with all the yield components. Singh and Singh (2005) estimated losses caused by Alternaria blight, white rust, and downy mildew infection of mustard under protected and unprotected conditions. Highest avoidable losses due to combined effects of three diseases in seed yield (34.7%), seed test weight (13.1%), and oil content (4.2%) were recorded when crop was sown after seed treatment with Apron SD-35 at 6 g/kg−1 followed by three sprays of mancozeb 75 WP (0.2%) at 15-day interval (Table 2.2). 42 2 The Disease: Downy Mildew

Table 2.2 Percent avoidable loss in seed yield, 1000 seed weight, and percent oil content in different commercial varieties of mustard due to Alternaria blight, white rust, and downy mildew for 1996–1997 to 1998–1999 (pooled data) (Singh and Singh 2005) Yield (Kg ha−1) 1000 seed weight (g) Oil content (%) % % % Variety UP P Mean loss UP P Mean loss UP P Mean loss NDR 8501 1080.0 1688.9 1384.4 36.0 4.9 5.7 5.3 13.6 39.6 41.0 40.3 3.3 Varuna 966.7 1733.3 1350.0 44.2 4.1 4.9 4.5 14.9 39.4 41.6 40.5 5.1 Kranti 1031.1 1528.9 1280.0 32.6 4.1 4.6 4.3 11.3 38.6 39.9 39.2 3.3 Krishna 973.3 1466.7 1220.0 36.1 4.1 4.8 4.5 14.2 38.0 39.8 38.9 4.7 Rohini 973.9 1413.3 1193.6 33.6 4.0 4.7 4.3 13.3 39.0 40.6 39.8 4.0 Vardan 960.0 1301.3 1130.6 26.0 4.0 4.6 4.3 11.6 37.4 39.2 38.3 4.6 Mean 997.5 1522.1 – 34.7 4.2 4.9 – 13.1 38.7 40.3 – 4.2 SEm± Main plot 38.67 0.02 0.06 (Treat.) Sub plot 66.93 0.04 0.11 (Variety) Treat. x 94.67 0.05 0.15 Variety C.D. (P < 0.05) Main plot 111.99 0.06 0.18 (Treat.) Sub plot 193.33 0.11 0.32 (Variety) Treat. x 275.99 0.16 0.45 Variety UP unprotected; P protected

2.3.2 Brassica Vegetables

During 1911–1912, downy mildew infection in cabbage near Lahore, Pakistan, caused more than 50% yield loss (Butler 1918). Vasileva (1976) reported that under favourable conditions, H. parasitica may infect up to 50–60% of cabbage seeds and reduces yield by 16–20%. Downy mildew disease can significantly affect the yield and developmental characters of radish (Achar 1992). Variables affected are the size and weight of silique, number of silique/plant, number of seeds/silique, and weight of seeds. Seed yield loss can be as high as 58%. Infection also adversely affects the size and weight of roots. Disease loss assessment has been estimated according to the following equation:

Mean yield of healthy plants − Mean yield of diiseased plants Yield loss()% =  ×100 Mean yield of healthy plants   2.5 Symptoms 43

2.4 Host Range

Few detailed studies have been made to determine the extent of the host ranges affected by downy mildew. In earlier work these fungi were inoculated on matured host tissues and then scored for the presence or absence of disease symptoms. The results suggested that downy mildew fungi had a very restricted host range (Gaumann 1918). Hyaloperonospora on crucifers was originally examined with this assumption in mind. As a result, a large number of species were created, mostly based on their occurrence on a particular crucifer genus. This process continued until Yerkes and Shaw (1959) called attention to the remarkable morphological similarity of the Hyaloperonospora species which attack crucifers. Following an extensive biometric study, they reduced over 80 species names to synonymy with H. parasitica (Pers. ex Fr.) Fr. More work, involving examination of the reactions of crucifer’s seedlings to infection, has led to an even wider host range being established for H. parasitica (Foster 1947; Davison 1967; Mc Meekin 1969). Wide variation can be encountered in the reaction of seedlings of different crucifer species to the isolates of Hyaloperonospora from Brassica and Raphanus (Tables 2.3 and 2.4), but the pathogen can grow well enough to sporulate on several species, apart from the original host (Dickinson and Greenhalgh 1977). Hyaloperonospora parasitica can infect a wide range of Brassica and other cruciferous species. Gaumann (1923) listed over 80 cruciferous species as susceptible to infection by the numerous species of Hyaloperonospora which are now regarded as all being H. parasitica. Among the common hosts of economic importance are rapeseed-mustard, cabbage, Chinese cabbage, cauliflower, broccoli, Brussels sprouts, marrow, stem kale, kohlrabi, turnip, turnip rape, swede, oilseed rape (canola), mustard, radish, horseradish, collards, rutabaga, watercress, stock, and wallflower (Channon 1981; Verma et al. 1994; Saharan et al. 1997; Nashaat 1997) (Table 2.5). Apart from these, the inven- tory of hosts reported to be infected by H. parasitica are given in Table 2.1.

2.5 Symptoms

Downy mildew (H. parasitica) is the most frequently recorded disease on horticultural and agricultural members of the genus Brassica. The disease mainly affects young plants that may, in severe cases, be stunted or killed. Infection at later stages results in the debilitation and reduction in performance and quality of the host plant.

2.5.1 Brassica Oilseeds

Rapeseed-Mustard The disease appears on all above-ground plant parts, but its symptoms are usually more conspicuous on leaves, stems, and inflorescences. At the seedling stage on cotyledons (Plate 2.1a), and the first few true leaves, small 44 2 The Disease: Downy Mildew

Table 2.3 Reaction of seedling cotyledons of members of the Cruciferae to inoculation with Brassica and Raphanus forms of H. parasitica (Dickinson and Greenhalgh 1977) Pathogen Brassica Host form Raphanus form Brassica oleracea L. subsp. oleracea L. Wild cabbage 3/4 3 B. oleracea L. Cultivated brassicas 4 3 B. nigra (L.) Koch Black mustard 1 1 B. juncea (L.) Czern Brown mustard 3/4 2 B. rapa L. Turnip 3 3 B. pekinensis (Lour.) Rupr. Chinese cabbage 3 2 Sinapis alba L. White mustard 2 2 Raphanus raphanistrum L. Wild radish 2 4 R. maritimus Sm. Sea radish 2 4 R. sativa L. Cultivated radish 2 4 Crambe maritima L. Seakale 1 1 Cakile maritima Scop. Sea rocket 1 1 Lepidium sativum L. Garden cress 1 1 Isatis tinctoria L. Woad 2/4 3 Iberis amara L. Wild candytuft 3/4 3 I. umbellata L. Garden candytuft 3/4 3 I. sempervirens L. Perennial candytuft 3 3 Thlaspi arvense L. Field pennycress 2 3 T. rotundifolium (L.) Gaudin – 2 3 Aethionema grandiflora R. Br. – 1 1 Capsella bursa-pastoris (L.) Medic. Shepherd’s purse 1 1 Lunaria annua L. Honesty 2 1 Alyssum saxatile L. Golden alyssum 2 2 A. maritimum (L.) Lam. Sweet Alison 2 2 Draba pyrenaica L. – 1 1 Arabis alpina L. Alpine rock cress 2 1 A. caerulea (All.) Haenke – 1 1 Rorippa nasturtium-aquaticum (L.) Hayek Watercress 1 1 Aubrieta deltoidea (L.) DC. Aubretia 1 1 Matthiola incana L. (R. Br.) Stock 3 2 M. bicornis (Sibth. & Sm.) DC Night-scented stock 2 2 Malcolmia maritime (L.) R. Br. Virginia stock 1/2 1/2 Hesperis matronalis L. Dame’s violet 2 1 Cheiranthus cheiri L. Wallflower 3 2 Camelina sativa (L.) Crantz Gold of pleasure 1 1 (Reactions were scored: 1 = no symptoms; 2 = necrotic flecking; 3 = extensive necrosis + slight sporulation; 4 = heavy sporulation; 1/2, etc., indicates intermediate reactions) 2.5 Symptoms 45

Table 2.4 Performance of the Brassica form of H. parasitica on cultivars of B. oleracea (Dickinson and Greenhalgh 1977) Mycelial Occurrence of pathogen (seedlings development in positive %) Sporulation cotyledons (grid Main Lateral Host variety intensitya squares covered %) Cotyledons Hypocotyl root root var. capitata L. (cabbage) cv. Red +++ 76 100 100 90 25 Drumhead cv. Savoy +++ 33 100 95 0 0 Drumhead cv. Flower of ++ 31 100 100 0 0 Spring cv. Standby ++ 47 100 100 25 0 cv. Greyhound ++ 26 100 95 5 0 cv. Harbinder ++ 30 100 100 45 0 cv. Primo ++ 31 100 100 35 0 cv. January + 11 100 60 0 0 King var. botrytis L. (cauliflower/broccoli) cv. Veitch’s +++ 63 100 100 50 0 Self Protecting cv. Roscoff +++ 56 100 100 5 0 Early cv. Calabrese +++ 53 100 100 40 5 cv. All the Year ++ 50 100 100 15 0 Round cv. Veitch’s ++ 45 100 100 25 5 Autumn Giant cv. June ++ 23 100 100 0 0 cv. Snowball ++ 50 100 100 0 0 cv. Majestic ++ 24 100 95 0 0 var. gemmifera Zenker (Brussels sprouts) cv. Masterman ++ 33 100 100 10 0 cv. British ++ 37 100 100 5 0 All-rounder cv. Cambridge ++ 51 100 100 15 5 No.5 cv. Jade Cross ++ 18 100 100 0 0 cv. Exhibition ++ 21 100 90 0 0 cv. Cambridge ++ 34 100 75 0 0 No.1 var. gongylodes L. (kohlrabi) cv. Green ++ 16 100 95 0 0 Vienna cv. Purple ++ 20 100 100 0 0 (continued) 46 2 The Disease: Downy Mildew

Table 2.4 (continued) Mycelial Occurrence of pathogen (seedlings development in positive %) Sporulation cotyledons (grid Main Lateral Host variety intensitya squares covered %) Cotyledons Hypocotyl root root var. acephala D.C. (Brecole) cv. Tall Green ++ 42 100 95 5 0 cv. ++ 38 100 90 0 0 1000-headed subsp. oleracea +++ 60 100 100 60 10 (wild cabbage) a+ = sparse sporulation; ++ = moderate sporulation; +++ = heavy sporulation angular translucent light green lesions appear. These lesions later enlarge and develop into greyish white, irregular necrotic patches on the upper surface of the leaf (Plate 2.1b), while downy fungal growth appears on the undersurface (Plate 2.1c). In a severe attack, diseased leaves dry up and shrivel. On cotyledons, necrotic lesions are more pronounced in B. rapa, whereas on true leaves, lesions are conspicuous on B. juncea. Symptoms of mixed infection of downy mildew and white rust are common on leaves and inflorescence of B. rapa and B. juncea. On leaves, downy growth of the fungus appears in or around the white rust pustules (Plate 10.1). On malformed inflorescences, sporulation of the downy mildew fungus is predominant in the form of white granular conidia and conidiophores (Plate 2.1d) (Saharan 1992a). According to Butler (1918), owing to very frequent co-existence of white rust, and downy mildew, it is not easy to separate their effects, but white rust produces the greatest deformities in the stem and flowers (Awasthi et al. 1995, 1997). Stem swellings may be limited often with abrupt bending of the stalk, or swelling may be several inches long (Plate 2.1d). The axis of the inflorescence is equally susceptible to deformity. The leaves and flowers are not often swollen, except for the young ovary, which may be transformed to a twisted body about 2 or 3 inches in length (Plate 2.1e). More often, the floral buds are atrophied with all the parts (i.e. sepals, petals, stamens, and pistil) being shrunken and almost colourless. If the attack is late, the buds/silique may be partly normal, partly deformed, or atrophied, and a single bud may similarly be affected in part only. Greyish white growth of conidia of pathogen is quite visible at the later stage of the crop growth on stag head (Plate 2.1f). There is never any trace of the violet colour produced in downy mildew infections which often occur with white rust. Systemically mixed infected plants with H. parasitica and A. candida have stunted and thickened growth of the whole plant which bears profuse sporulation of both pathogens. Hypertrophy of the affected cells, which is mainly attributed to infection with A. candida, causes thickening of the stem and inflorescence. The hypertrophied tissues tend to attract infection by H. parasitica because their relative susceptibility to this pathogen is much higher than normal tissues (Awasthi et al. 1995, 1997). The pith of the stem has more hypertrophied tissue than the cortex. The 2.5 Symptoms 47

Table 2.5 Host species of H. parasitica (Channon 1981; Saharan et al. 1997 up dated) Scientific name English name Reference Alliaria petiolata Garlic mustard weed Ramsfjell (1960) and Jorstad (1964) Alyssum Weed Arabidopsis spp. Weed Koch and Slusarenko (1990) A. arenosa Sand rock crass Ramsfjell (1960) and Jorstad (1964) A. petraea Northern rock cress Ramsfjell (1960) and Jorstad (1964) A. thaliana Thale cress Ramsfjell (1960) and Jorstad (1964) Armoracia rusticana Horseradish Moore (1959) Arabis spp. Rock cress Anonymous (1960a, b) Arabis glabra (Turritis Tower mustard glabra) A. hirsuta Hairy rock cress Ramsfjell (1960) and Jorstad (1964) Aubretia spp. Aubretia Moore (1959) Brassica alba White mustard Anonymous (1960a) B. arvensis Wild mustard Anonymous (1960a) B. alboglabra Chinese kale Johnston (1963) B. chinensis Chinese cabbage Hiura and Kanegae (1934) B. juncea Mustard Gaumann (1926) B. kaber White mustard Anonymous (1960a) B. hirta White mustard Anonymous (1960a) B. fruticulosa Gaumann (1926) B. napus Rape Moore (1959) B. napus subsp. napus Oilseed rape Ramsfjell (1960) and Jorstad (1964) B. napus var. Swedes Moore (1959) napobrassica B. nigra Black mustard Gaumann (1926) B. rapa ssp. campestris Turnip rape Ramsfjell (1960) and Jorstad (1964) B. oleracea var. Marrow stem kale Moore (1959) acephala B. oleracea var. botrytis Cauliflower Moore (1959) and Ramsey (1935) B. oleracea var. capitata Cabbage Moore (1959) and Ramsey (1935) B. oleracea var. Moore (1959) and caulorapa Ramsey (1935) (continued) 48 2 The Disease: Downy Mildew

Table 2.5 (continued) Scientific name English name Reference B. oleracea var. Brussels sprouts Moore (1959) and Thung gemmifera (1926a, b) B. oleracea var. Kohlrabi Johnston (1963) gongylodes B. oleracea var. italica Broccoli Ramsfjell (1960) and Jorstad (1964) B. oleracea var. Kale Ramsfjell (1960) and sabellica Jorstad (1964) B. pekinensis Chinese cabbage Chang et al. (1963) and Ocfemia (1925) B. campestris var. Swedes, turnip Moore (1959) rapifera B. campestris var. Yellow sarson Saharan (1992a) yellow sarson B. campestris var. Brown sarson Saharan (1992a) brown sarson B. campestris var. toria Toria Saharan (1992a) B. tournefortii Gaumann (1926) Barbarea Anonymous (1960a) Barbarea stricta Winter cress, yellow rocket Ramsfjell (1960) and Jorstad (1964) B. vulgaris Bitter cress, rocket cress, herb barbara, Ramsfjell (1960) and yellow rocket cress, wound rocket Jorstad (1964) Berteroa incana Hairy alyssum Bunias orientalis Hill mustard, Turkish rocket Ramsfjell (1960) and Jorstad (1964) Chenopodium album Bathu Saharan (1996) Camelina sativa Darpoux (1945) Cheiranthus allioni De Bruyn (1935) C. cheiri Wallflower Moore (1959) and Wiess (1927) Capsella bursa-pastoris Shepherd’s purse Farlow (1883) Cardamine amara Large bitter cress Ramsfjell (1960) and Jorstad (1964) C. bulbifera Coralroot bitter cress Ramsfjell (1960) and Jorstad (1964) C. flexuosa Woodland bitter cress Ramsfjell (1960) and Jorstad (1964) C. impatiens Narrow leaf bitter cress Sharma and Munjal (1977) C. pratensis Cuckoo flower, lady’s smock Ramsfjell (1960) and Jorstad (1964) C. rhomboidea Farlow (1883) Coronopus didymus Langdon (1948) (continued) 2.5 Symptoms 49

Table 2.5 (continued) Scientific name English name Reference C. squamatus Dias and Da Camara (1953) Crambe maritima Crambe Moore (1959) Dentaria spp. Anonymous (1960a) D. laciniata Farlow (1883) Descurainia spp. Anonymous (1960a) D. saphia Bixweed Ramsfjell (1960) and Jorstad (1964) Draba spp. Anonymous (1960a) D. caroliniana Farlow (1883) D. globella Smooth draba Ramsfjell (1960) and Jorstad (1964) D. verna Shadflower, nailwort Ramsfjell (1960) and Jorstad (1964) Diplotaxis tenuifolia Wild rocket Ramsfjell (1960) and Jorstad (1964) Eruca sativa Taramira, rocket Gaumann (1926) E. vesicaria Euphorbia Erysimum Wormseed mustard, treacle mustard Anonymous (1960a) cheiranthoides E. strictum Tall wormseed, wallflower Ramsfjell (1960) and Jorstad (1964) Hesperis spp. Anonymous (1960a) H. matronalis Dame’s rocket Iberis amara Candytuft Anonymous (1960a) Lepidium campestre Lepidium sativum Garden cress Anonymous (1960a) L. intermedium Swingle (1890) L. graminifolium Nicolas and Aggery (1940) L. virginicum Farlow (1883) Lobularia spp. Koniga Anonymous (1960a) Lobularia maritima Sweet alyssum Ramsfjell (1960) and Jorstad (1964) Malcolmia africana Thind (1942) Matthiola incana Stock Moore (1959) and Wiese (1927) M. incana var. annua Tenweeks stock Ramsfjell (1960) and Jorstad (1964) Noccaea caerulescens Alpine pennycress Ramsfjell (1960) and Jorstad (1964) Nasturtium officinale Watercress Moore (1959) (continued) 50 2 The Disease: Downy Mildew

Table 2.5 (continued) Scientific name English name Reference Raphanus sativus Radish Moore (1959) R. raphanistrum Wild radish Gaumann (1926) R. raphanistrum ssp. Wild radish Ramsfjell (1960) and raphanistrum Jorstad (1964) Radicula Davis (1929) nasturtium-aquaticum Rorippa Anonymous (1960a) R. islandica Northern marsh yellowcress Ramsfjell (1960) and Jorstad (1964) Rumex Sinapis alba White mustard Gaumann (1926) S. arvensis Wild mustard Gaumann (1926) Sisymbrium altissimum Tumble mustard Ramsfjell (1960) and Jorstad (1964) Sisymbrium officinale Hedge mustard Anonymous (1960a) S. irio Tumbling weed Thind (1942) Thlaspi arvense Field penny cress

Table 2.6 Downy mildew interaction-phenotype classes used for cotyledon and leaf-disc evaluation (Monterio et al. 2005) are presented below Class Interaction phenotype 0 No host reaction, no sporulation 1 Light host necrosis localized on the upper cotyledon/leaf disc surface, no sporulation 2 Diffuse host necrosis localized on the upper cotyledon/leaf disc surface, no sporulation 3 Host necrosis localized on the upper cotyledon/leaf disc surface, weak sporulation (five conidiophores), localized on the lower cotyledon/leaf disc surface, confined to the point of infection 4 Host necrosis localized on the upper cotyledon/leaf disc surface, heavy sporulation localized on the lower, cotyledon/leaf disc surface confined to point of infection 5 No necroses on the upper surface, sparse to moderate sporulation dispersed over the whole cotyledon/leaf disc surface 6 No necroses on the upper surface, abundant, and dense sporulation dispersed over the whole cotyledon/leaf disc surface affected inflorescence either bears no silique, or produces abnormal silique, which are often curled without seeds. In the initial stages, an affected inflorescence does not show typical symptoms of infection such as the presence of oospores and downy growth on the surface. But at the later stages, conidial fungal growth of downy mildew and sporangiophore blisters of white rust occur on the surface of affected tissue, and formation of oospores takes place in the tissue as it dries. Necrotic lesions bearing downy growth of the fungus may also be observed on well-developed silique (Awasthi et al. 1995, 1997; Kolte 1985; Saharan 1992a; Vasudeva 1958). 2.5 Symptoms 51

The internal changes due to infection by Hyaloperonospora differ from those caused by Albugo in many respects. With Hyaloperonospora infections, the palisade cells of the leaf are not changed. In the deeper layers of the cortex, endoderm, and pericycle, new cell layers may be formed by normal cell division. Only the cells around the vascular bundles become enlarged and thin-walled; the rest of the interfascicular sclerenchyma remains unaltered. There is no interfascicular cambium in these host plants. The cambium of the vascular bundles remains active, whereas the 52 2 The Disease: Downy Mildew xylem and phloem vessels become enlarged and separated by radial bands of parenchyma when hyphae have penetrated. There are no accessory bundles. In general, the effect on the cells seems to be more destructive than in Albugo, the chlorophyll content is diminished, and the cell contents are more rapidly used up. There is no tendency for chlorophyll accumulation in unusual places as with white rust. In general, the effect of downy mildew on the cell seems to be more destructive than white rust (Butler 1918). The protoplast of epidermal cells responds differently to haustorial development than protoplast of the mesophyll cells (Chou 1970). The epidermal cells result in a severe disruption of the protoplast. The central vacuoles contract and probably undergo fragmentation, the plasmalemma is broken down or detached from the wall, and numerous vesicles are formed from it. The cytoplasm is either dislocated and aggregated into a vacuolated blob or completely dispersed to the extent that its identity cannot be discerned. Consequently, haustoria in epidermal cells are not, in most cases, surrounded by a clearly defined layer of host cytoplasm. Haustoria formation in a mesophyll cell causes less disruption. The host cytoplasm is merely invigilated by the invading haustorium, while the tonoplast and plasmalemma apparently remain intact.

2.5.2 Brassica Vegetables

Plants can be infected at any time (Sherf and Macnab 1986). In seedbeds, the cotyledons and first leaves are invaded. The adaxial surface of the leaf bears small, pale- yellow, angular spots, which may grow together to form irregular brown patches (Plate 2.2a–b). On the abaxial surface, the corresponding areas are covered with a light-grey fungus formed by multi-branched conidiophores bearing conidia (Plate 2.2c). Young leaves and cotyledons may drop off as they become yellow. Older leaves usually persist and show the downy growth of the pathogen (Plate 2.2d), and affected areas enlarge becoming papery and tan coloured (Plate 2.2e). Severe infection may cause the death of the whole leaves. Minute necrotic flecks covering the leaf surface may often form resembling peppery leaf spot caused by bacteria. When the fungus enters the stalk at the leaf base of an old head of cabbage, a greyish black discolouration of the stalk occurs (Ramsay and Smith 1961). In some storage with lots of cabbage, this discolouration has been found extending up through the stalk to the innermost bud leaves. On cabbage heads, the pathogen may cause numerous sunken black spots, varying in size from minute dots to an inch or more in diameter (Sherf and Macnab 1986). A similar blackening occurs on cauliflower curds (Chorin 1946). The infection is evident as brown to black streaks in the vascular system of the upper portion of the main stalk and branches leading to the florets. The fleshy roots of turnips and radishes have an internal irregular region of discolouration extending from the root crown downward or beginning on the side at soil level. The flesh is brown to black or shows net necrosis. In advanced stages the skin can be roughened by minute cracks, and the root can split open (Sherf and Macnab 1986). 2.5 Symptoms 53

Plate 2.2 (a) Initial symptoms of downy mildew on cabbage leaf; (b) abaxial side of cabbage leaf showing initial symptom, (c) adaxial surface of cabbage leaf showing downy growth; (d)- close-up of leaf showing conidial growth of Hyaloperonospora; (e) drying of the leaf due to advance stage of downy mildew

According to Butler (1918), the fungus is visible as a thin, greyish white, downy growth, occurring in scattered patches on the undersurfaces of the leaves in cabbage, cauliflower, and turnip and on the leaves, stem, and inflorescence in radishes. 54 2 The Disease: Downy Mildew

The upper surface of the leaf is marked by white spots corresponding to the downy growth below. In severe attacks, the spots may be so crowded that the leaf dries up, shrivels, and tears easily. In seedlings, the whole undersurface may be evenly covered, and total infection of the young inflorescence is also found. Occasionally the roots of radish and Swedish turnip are attacked in Europe. The tissues blacken and rot near the surface, oospores occur within the tissues, and conidiophores form if exposed to the air.

2.5.3 Broccoli

Downy mildew appears first on the lower leaves of broccoli plants (Natti et al. 1956). Leaf infection may occur soon after the plants are set in the field or may take place later in the season. Older leaves appear to be more susceptible than newly developed leaves. When the surface of the foliage is wet, the downy white mycelium of the fungus is readily observed on the undersurface of the leaves. The first symptoms of leaf infection are small water-soaked spots surrounded by a halo of light green tissue on the undersurface of the leaf. Under conditions favourable for development of infection, the spots enlarge to form indefinite yellow areas. Later, the tissues within these infected areas collapse and become light brown and parchment-like. The mildew lesions vary in size and shape. The largest lesions usually are bounded by leaf veins. The initial spots of infection may also remain localized. The tissues of the spot collapse to form a small brown lesion. Systemic infections are usually confined to the upper portion of the main stalk and to the branches leading to the florets of the head. Infected tissues develop brown to black netted lesions and in others as long strands of discoloured tissues. In some plants systemic invasion of the head can be detected by diffuse blue to purple areas on the stalk and branches of the head.

2.5.4 Wallflower (Cheiranthus)

On the diseased plants, the upper surface of the leaves shows pale yellowish patches, while the corresponding parts of the undersurface are covered with a greyish or white fungal growth (Gram and Weber 1952). The infected stems and flowers are swollen and often twisted. Diseased flower buds do not develop.

2.5.5 Stock (Matthiola)

The disease is more common on young plants before they are transplanted but may also appear later, especially on crop grown indoors (Gram and Weber 1952). On the upper surface of the leaves, there are pale spots, while on the corresponding parts of 2.5 Symptoms 55 the undersurface is a whitish layer of the fungus. Stalks and flower heads may also be attacked. The diseased parts show various kinds of distortion. According to Jafar (1963), the disease appears as light green areas on the upper surface of leaves. The corresponding undersurface is chlorotic with white growth of fungal conidia and conidiophores. Infected areas turn yellow and become necrotic leading to premature leaf fall. The flowers of infected plants frequently fail to open and often die. Fructification appears on cotyledons, and seedlings may be killed.

2.5.6 Rocket (Eruca sativa)

The disease affects newly expanded leaves of plants when they are 15–20 days old causing a major reduction in quality. Adaxial surfaces of the leaves show small irregular, dark brown to black speckling. Abaxial leaf surfaces usually support the white fungal growth. Older, fully expanded foliage is unaffected. Hyaline sporangiophores emerge from stomata and branch dichotomously ending the slender curved tips. Sporangia are slightly brown, and ovoid, and measure 22.5–27.5 μm (average 25) long x 17.5–20.0 μm (average 18) wide (Larran et al. 2006).

2.5.7 Cruciferous Weed (Arabidopsis thaliana)

The symptoms are first apparent to the naked eye as a carpet or ‘down’ of conidiophores covering the upper and lower surfaces of leaves and petioles. Isolates infecting A. thaliana have so far proven to be non-pathogenic on other crucifers tested but exist in a clear gene-for-gene relationship with different ecotypes (Slusarenko and Schlaich 2003). Macroscopic symptoms of downy mildew on mature plants of A. thaliana are discrete, pale green patches on rosette leaves which eventually turn chlorotic when relative humidity is high (95%), and often before other symptoms are evident, the fungus sporulate on the lower leaf surface producing a forest of sporangiophores. The sporangiophores are dichotomously branched structures about the size of a trichome, emerging through stomata and producing conidiosporangia at the branch tips (Koch and Slusarenko 1990). The density of sporulation is variable, but a downy white mat covering a substantial area of leaf is common. On bolting plants, sporulation can often be observed over considerable lengths of the flowering stem. In wild population, small seedlings also become infected. Such seedlings have paler cotyledons and, because of their diminutive size, require a practiced eye to observe the sporangiophores (Dangl et al. 1992). 56 2 The Disease: Downy Mildew

2.6 Disease Assessment

Different scales have been used for classifying leaf infection by downy mildew pathogen. (a) Natti et al. (1967) and Sadowski (1987) used scales ranging from 0 to 5, where: 0 = No symptoms 1 = Spots, necrotic flecks or streaks, but no sporulation 2 = Spots, necrotic flecks or streaks, with sparse sporulation confined to necrotic tissue 3 = Systemic infection and sporulation in increasing degree 4 = Systemic infection and sporulation in increasing degree 5 = Systemic infection and sporulation in increase degree Plants with ratings up to 2 are considered resistant. (b) Ebrahimi et al. (1976) has rated downy mildew resistance in B. juncea lines on a scale of 1 to 5 where, 1 = indicates no sporulation, 2 = very sparse sporulation, and 5 = heavy sporulation. (c) A similar scoring scale of 1 to 4 with slight modification has been used by Dickinson and Greenhalgh (1977). (d) The use of a 0–9 scale has been suggested by several workers (Knight and Furber 1980; Nashaat and Rawlinson 1994; Saharan 1992b, 1996; Williams 1985). It is described as follows: 0 = no symptoms or signs of H. parasitica 1 = very minute to larger scattered necrotic flecks under the inoculum drop, no or small amounts of necrosis on the lower cotyledon surface, no sporulation. 3 = very sparse sporulation, one to a few conidiophores on the upper or lower surfaces, necrotic flecking often present, tissue necrosis present. 5 = sparse scattered sporulation on either or both cotyledon surfaces, tissue necrosis 7 = abundant to heavy sporulation mainly on lower surfaces, light to scattered sporulation on upper surfaces; tissue necrosis and chlorosis may be present. 9 = abundant sporulation; leaf or cotyledon collapsed A disease index (DI) was calculated using the formula:

9 DI = ∑()ixjn/ i=0 where n = total plants, i = infection phenotype class, and j = number of plants per class. Genotypes are categorized as resistant (0 to 1), partially resistant (3 to 5), and susceptible (7 to 9). 2.6 Disease Assessment 57

(e) Interaction-phenotype scores of 0–6 have been used by Coelho et al. (2012) to evaluate the response of B. oleracea cotyledons and relative amount of sporulation following inoculation with H. brassicae is as follows:

IP scores Interaction phenotype 0 No host reaction, no sporulation 1 Light host necrosis localized on upper cotyledon surface, no sporulation 2 Heavy host necrosis localized on the upper cotyledon surface, no sporulation 3 Host necrosis localized on the upper cotyledon surface, weak sporulation (five conidiophores) localized on the lower cotyledon surface confined to the point of infection 4 Host necrosis localized on the upper cotyledon surface, heavy sporulation localized on the lower cotyledon surface confined to the point of infection 5 No necrosis on the upper surface, sparse to moderate sporulation dispersed over the whole cotyledon surface 6 No necrosis on the upper surface, abundant to dense sporulation dispersed over the whole cotyledon surface

(f) Kruger (1991) suggested the use of 1–9 scale in the form of diagrams to estimate disease on leaves of oilseed rape (Fig. 2.1); scores 3, 5, 7, and 9, respectively, represent 7, 27, 65, and 100 percent of the leaf area infected. If larger leaves are concerned, those in Fig. 2.1 should be enlarged by two to five times to get a comparable shape and size to the leaves found in the field. (g) Brophy and Laing (1992) assessed disease severity using an image analyser to determine logarithmic rating scales of percentage leaf area infected for both cotyledons and primary leaves of cabbage. They found that the maximum area infected can be 100% in cotyledons but in primary leaves it rarely exceeds 25%. In order to integrate the two components, transformation of the data is necessary. Percentage disease severity (PDS), expressed as a function of cotyledon and primary leaf infection, is calculated using the formula PDS = (C + x P)/2, where C is the percentage cotyledon area infected with a maximum value of 100%, x is the inverse of the maximum measured percentage primary leaf area infected, and P is the percentage primary leaf area infected of treated plants. (h) Disease assessment scales for partial resistance: Disease attack was mainly assessed 6 days after inoculation and was assessed visually on the cotyledons by a sporulation score, modified from Natti et al. (1967) and Williams (1985). This score was based on the area with visible sporulation (score 0–3), combined with the sporulation intensity (score 4–5). 0 = no sporulation, 1 = sporulation confined to less than four conidiophores at the inoculation site, 2 = sporulation on the inoculation site, 3 = sporulation area greater than the inoculation site, but less than the entire cotyledon, 4 = moderate sporulation over the entire cotyledon, 5 = profuse sporulation over the entire cotyledon. The conidium production on the seedlings was also estimated. Cotyledons from ten seedlings were excised and shaken for 30 s in 10 ml of electrolyte Isoton II (Coulter Euro 58 2 The Disease: Downy Mildew

Fig. 2.1 Disease assessment (1–9) on leaves of rapeseed-mustard

Diagnostics GmbH, Krefeld, Germany). The electrolyte inhibited conidium germination, thereby preserving them for counting. Eight independent haemocytometer counts, representing 1.6mm2 each, were carried out per sample and used for calculation of the conidium production per seedling (Jensen et al. 1999). (i) Revised rating scale (0–9) of AICRP Rapeseed-Mustard scientist group meeting of 2011, rating scale for AB, WR, and DM as follows.

Rating score Leaf area covered (%) Disease reaction 0 No symptoms Immune (I) 1 > 5 Highly resistant (HR) 3 5–10 Resistant (R) 5 11–25 Moderately resistant (MR) References 59

Rating score Leaf area covered (%) Disease reaction 7 26–50 Susceptible (S) 9 < 50 Highly susceptible (HS)

Revised rating scale (0–9) of AICRP Rapeseed-Mustard group meeting of 2011 for scoring disease severity and disease reaction of downy mildew, white rust, and Alternaria blight of rapeseed mustard. Formula for calculating disease severity:

()NNN−×10+−()21× +−()33× +−()NN45× +−()5× 76+−()N × 9 Average severity score = No. of leaf samples

(j) Downy mildew leaf resistance has been evaluated using following 0–6 scoring scale (Monterio et al. 2005) as follows: (k) Holub et al. (1994) used a scale to describe phenotypic and genotypic characterization of interaction between isolates of H. parasitica and accessions of A. thaliana as follows: 1 EH, early heavy sporulation; 2. FDL, minute necrotic flecks, delayed, and low sporulation; 3 FN, minute necrotic flecks and no sporulation; and 4. PN, necrotic pits and no sporulation. Interaction phenotype was characterized using an assessment of host and pathogen characteristics: the emergence of sporangiophores of H. parasitica as early (E) 3 days after inoculation (dai) or delayed (D) >4 dai; the intensity of asexual sporulation as heavy (H; >20 sporangiophores per cotyledons), moderate (M, 10–20 sporangiophores), light (L,<10 sporangiophores), rare (R, <5 sporangiophores on < 10% of inoculated seedlings), or none (N); and the type of response by A. thaliana as minute necrotic flecks (F), evident 7 dai, flecks clearly visible 3 dai which form necrotic cavities (C) 7 dai, or necrotic pits (P) observed as early as 3 dai and often expanding until much of cotyledon is necrotic 7 dai.

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3.1 Introduction

Downy mildew of crucifers is caused by an obligate pathogen, Hyaloperonospora parasitica (Gaum.) Goker synonymy Peronospora parasitica (Pers. ex. Fr.) Fr., Sum. Veg. Scand. 493, 1849. Extensive synonymy is given by Yerkes and Shaw (1959). Sometimes it is referred to as Peronospora brassicae synonymy H. brassicae (Gaum.) Goker specific to family Brassicaceae. On Arabidopsis, currently it is known as H. arabidopsidis (Gaum.) Goker, Riethm., Voglmayr. Weiss & Oberw. [as Arabidopsis], Mycol. Prog. 3(2): 89 (2004), synonymy P. arabidopsidis Gaum. (1918). The genus Peronospora s. lat. is one of the largest genera of downy mildews (Peronosporales, Peronosporomycetes, phylum Heterokonta). It can be regarded as the most highly evolved and successful genus of its order in terms of both biodiversity and distribution. The members of this group parasitize mainly temperate, her- baceous host genera from a broad range of dicotyledonous families and even one monocotyledonous family (Alliaceae) (Dick 1988, 2001a). In addition, several species are economically important pathogens of crop and ornamental plants (Spencer 1981), and the genus Peronospora comprises about 75 to 460 species (Dick 2001b), depending on the species definition applied. According to Hall (1996), definition and circumscription of species entities is a major problem in Peronospora, and unfortunately, no modern taxonomic treatment is available, and this has caused much confusion concerning species definition and delimitation. In the concept of Gaumann (1918, 1923), excessive splitting was applied according to the principle of one host-one parasite species, postulating a pronounced host specificity. It was decided for distinction of different species even if the morphological differences observed were minimal, based on evidence that some strains investigated by cross-inoculation experiments were strictly host specific. However, many of Gaumann’s taxa were described from just a few or even single collections, neglecting the possibility of variability in features. In addition, the proposed host specificity has been confirmed by meaningful cross-infection

© Springer Nature Singapore Pte Ltd. 2017 67 G. S. Saharan et al., Downy Mildew Disease of Crucifers: Biology, Ecology and Disease Management, https://doi.org/10.1007/978-981-10-7500-1_3 68 3 The Pathogen: Hyaloperonospora parasitica (Gaum.) Goker [H. brassicae (Gaum.… only in a few experiments. Therefore, Gaumann’s species often lack a clear circumscription, and identification is often impossible if the host species is unknown. As a consequence, some authors applied excessive species lumping, accepting only one or a few species within one host family (Yerkes and Shaw 1959). However, this approach neglects morphological and genetic distances between isolates from different hosts belonging to the same family, and unrelated species may be recorded under the same name. This is undesirable not only for theoretical reasons but also for applied plant pathology where clear species definition is a prerequisite for appropriate species determination and subsequent control mechanisms, quarantine restrictions, and risk perception of the pathogens. Recently, phylogenetic analyses of DNA sequences have received increasing attention in Peronosporales, opening a new era in the investigation of their phylogenetic relationships. Analysing the internal transcribed spacer (ITS) region, Cooke et al. (2000, 2002) showed that Peronospora s. str. is embedded within a paraphyletic genus Phytophthora. In the analysis of Constantinescu and Faheti (2002), also using ITS sequences, the genus Peronospora s. lat. appeared as paraphyletic and segregated the two genera Hyaloperonospora and Perofascia from Peronospora based on molecular and morphological evidence. Similar results were obtained by Riethmuller et al. (2002) using large subunit ribosomal DNA (LSU rDNA) sequences. Both genera Hyaloperonospora and Perofascia almost exclusively infect hosts in the Brassicaceae and related families. The genera were segregated from Peronospora due to pronounced morphological and molecular differences (Constantinescu and Faheti 2002). In the analyses using Bayesian inference, Perofascia has consistently been shown to be the sister genus of Hyaloperonospora with 100% posterior probability, which is consistent with the MP tree of Constantinescu and Faheti (2002). Gaumann published many species in this group, which was questioned by Yerkes and Shaw (1959) who accepted only one species on the Brassicaceae. After morphological investigations, Constantinescu and Faheti (2002) accepted eight distinct species in Hyaloperonospora: three parasitic on Brassicaceae (H. niessleana, H. lunariae, H. lepidii-sativi), one on Limnanthaceae (H. floerkeae), and one on Zygophyllaceae (H. tribulina). One species, H. parasitica s. str., was assumed to have a wide host range based on morphological evidence. In the sense of Constantinescu and Faheti (2002), H. parasitica s. str. contains isolates from diverse species of the Brassicaceae and also of other closely related families such as the Resedaceae and Capparidaceae and, remarkably, even the unrelated Cistaceae. Although only few collections regarded as contaxic by Constantinescu and Faheti (2002) have been included in the study of Voglmayr (2003), the molecular data do not favour this as a tenable solution. H. parasitica from Capsella bursa-pastoris, the type host, has a 302 bp insertion in the ITS2 region which is lacking in all other species of the group. Peronospora lepidii-sativi, assumed to be conspecific with H. parasitica by Constantinescu and Faheti (2002), has a 343 bp insertion in the ITS2, which, however, apparently is not homologous with that of H. parasitica s. str. In addition, the genetic distances between collections from different hosts are usually very high within the clade (3.9–23.8%), evidence for presence of genetically dis- 3.2 Taxonomy and Morphology 69 tinct taxa. This is consistent with the results of Rehmany et al. (2000), who also found high genetic distances between collections from different hosts. Therefore, H. parasitica in the sense of Constantinescu and Faheti (2002) should represent an assemblage of sibling species with little morphological differences. The species regarded as morphologically distinct by Constantinescu and Faheti (2002) are embedded within the different collections of H. parasitica s. lat., rendering it highly paraphyletic (Voglmayr 2003). Downy mildews and white blister rusts are members of the class Oomycetes (Peronosporomycetes), a comparatively small lineage with estimates of <1000 species (Kirk et al. 2001). Due to their morphological, physiological, and ecological similarities to fungi, the Oomycetes are traditionally treated within mycology; however, ultrastructural, biochemical, and molecular phylogenetic data confirm that they are not related to true fungi (kingdom Fungi), but belong to the kingdom Chromista (Straminipila) which also contains the chromistan (heterokont) algae (Dick 2001a, 2002; Kirk et al. 2001). Downy mildews are an important group of obligate biotrophic plant parasites, which have a great economic impact on numerous crops (Plasmopara viticola on Vitis vinifera, Pseudoperonospora humuli on Humulus lupulus, Peronospora tabacina on Nicotiana spp., etc.). With the commercial large-scale production of some crops, and ornamental plants, some downy mildew diseases have recently become of major concern, such as downy mildew of Ocimum basilicum (basil; Belbahri et al. 2005), Eruca sativa (rocket; Larran et al. 2006), or Rubus spp. (arctic bramble and boysenberry; Hukkanen et al. 2006). In several of these cases, classification of the causal agent is still unclear, which demonstrates the ignorance of biodiversity of downy mildews. With the rapid development of new research techniques in applied and theoretical plant pathology, the interest in phylogeny of downy mildews has increased over the last few years. The availability of a substantial number of additional new characters less prone to subjective interpretations in general led to a change of paradigms in classification, as phylogenetic hypotheses could be vigorously tested for the first time, which led to a shift from a phonetic to a phylogenetic classification. In addition, phylogenetic analyses of DNA sequence data also enabled a re-evaluation of morphological features in an evolutionary background and a reinvestigation of species boundaries and host specificity. Significant progress has been made towards a phylogenetic classification; however, several important questions are still unresolved (Voglmayr 2008).

3.2 Taxonomy and Morphology

Taxonomy and morphology are very essential to understand and communicate the relationship of different species in the fields of biology, ecology, genetics, mycology, physiology, and plant pathology. A framework is required that roughly expresses the phylogenetic relationship of taxa examined. Although this can vary in different groups of organisms, members of one species are more closely related to each other than distinct species within a genus or distinct genera within a family. 70 3 The Pathogen: Hyaloperonospora parasitica (Gaum.) Goker [H. brassicae (Gaum.…

The earliest reference of downy mildew on crucifers is by Persoon (1796) who ascribed the cause of the disease on Thlaspi bursa-pastoris (Capsella bursa- pastoris) to the fungus Botrytis parasitica Pers. In 1849, Fries (Gaumann 1918) transferred the fungus to the genus Peronospora which had been established in 1837 by Corda in his description of Peronospora ramicis (Corda 1837). At that time, all isolates obtained from cruciferous hosts were ascribed to P. parasitica (Pers. ex. Fr.) Fr. The taxonomy of downy mildew of Brassica has undergone a number of revisions since Corda (1837) originally coined the genus Peronospora. Originally, de Bary (1863) applied a broad species concept and, with few exceptions, considered all Peronospora accessions infecting a specific host family as single species. However, Gaumann (1918) named isolates of Peronospora affecting Brassica species as P. brassicae Gaum. He considered that the various isolates obtained from different hosts should be classified as separate entities and on this basis recognized 52 species of Peronospora. The conclusions were based largely upon conidial dimensions and the results of cross-inoculation tests. The value of conidial dimension as a taxonomic criterion has since been questioned because size may vary according to environmental conditions (Thung 1926). Yerkes and Shaw (1959) reported remarkable morphological similarity of Peronospora species which attack crucifers. Measurements of conidia (Tables 3.1 and 3.2), an inability to associate conidiophores types with particular host genera, and the uniformity of oospores led Yerkes and Shaw (1959) to conclude that there is no reliable morphological basis to distinguish different species of Peronospora affecting the crucifers. Following an extensive biometric study, over 80 species names were reduced to one synonym, and a single species, P. parasitica, has been recognized on cruciferae hosts (Dickinson and Greenhalgh 1977; Hiura and Kanegae 1934; Waterhouse 1973; Yerkes and Shaw 1959). However, in view of the apparent differences in the anther- idial structure in the isolates of Peronospora on Capsella bursa-pastoris (Wager 1889), and on B. oleracea (Mc Meekin 1960), the merits of some separate speciation must not be ruled out. For the first time, the phylogeny of the Peronosporales (Fig. 3.1) has been shown by Shaw (1981). Species definitions for plant pathogens have considerable practical impact for measures such as plant protection or biological control and are also important for comparative studies involving model organisms. However, in many groups, the delimitation of species is a notoriously difficult taxonomic problem. This is particularly evident in the obligate biotrophic downy mildew genera (Peronosporaceae, Peronosporales, Oomycetes), which display a considerable diversity with respect to genetic distances and host plants, but are, for the most part, morphologically rather uniform. The recently established genus Hyaloperonospora is of particular biological interest because it shows an impressive radiation on virtually a single host family, Brassicaceae, and it contains the downy mildew parasite, Arabidopsis thaliana, of importance as a model organism. Based on the most comprehensive molecular sampling of specimens from a downy mildew genus to date, including various collections from different host species, and geographic locations, Goker et al. (2009) investigated the phylogenetic relationships of Hyaloperonospora by molecular analysis of the nuclear ribosomal ITS and LSU sequences. Phylogenetic trees were 3.2 Taxonomy and Morphology 71 120 (2) 100 (1) 1200 (13) 100 (1) 1000 (1) 100 (1) 1000 (1) 1200 (2) 140 (3) 220 (3) 101 (1) 140 (3) 100 (1) 100 (1) 100 (1) 1000 (1) 220 (3) 100 (1) 1000 (1) 100 (1) 100 (1) 100 (1) 1000 (1) 100 (1) No. of spores + no. of collections 15.3–25.3 15.7–26.8 11.9–23.8 14.2–22.6 12.0–24.0 11.9–18.4 14.0–24.0 11.0–24.0 11.5–21.8 11.9–20.7 12.8–20.8 12.3–21.1 12.6–16.9 14.5–21.4 11.5–20.7 14.0–22.0 11.1–19.2 11.5–16.1 9.0–21.0 11.5–18.0 11.5–17.2 9.6–15.3 8.0–20.0 12.6–19.9 Range 20.14 19.70 18.30 17.41 17.66 17.37 14.48 Max. (19.8) (19.7) (17.1) (18.7) (13.3) (15.3) (18.4) (18.3) (16.5) (16.6) (17.1) (16.2) (14.6) (17.0) (16.0) (18.2) (13.5) (14.2) (15.8) (14.3) (14.8) Grand (11.9) (13.9) (13.6) 19.44 15.86 18.21 15.69 15.68 15.33 13.02 Min. Width ( μ ) Width Means 1.48 1.44 1.20 1.31 1.25 1.25 1.37 Max. (1.50) (1.47) (1.26) (1.36) (1.77) (1.50) (1.35) (1.18) (1.29) (1.19) (1.22) (1.35) (1.16) (1.25) (1.10) (1.21) (1.25) (1.12) (1.31) (1.28) (1.30) (1.32) (1.16) (1.24) Grand 1.46 1.26 1.15 1.26 1.21 1.17 1.23 Min. Quotient (length/ width) Means 23.0–38.3 19.1–38.3 18.0–29.5 17.0–32.0 19.9–33.3 19.0–37.0 14.6–36.8 12.0–35.0 15.3–27.6 16.0–27.2 14.6–26.4 15.3–23.8 15.7–23.7 15.3–27.2 14.0–26.0 14.6–26.8 13.4–23.0 11.0–24.0 15.3–24.9 14.2–23.0 13.4–22.6 11.9–19.9 11.0–23.0 10.3–16.1 Range 29.38 28.40 21.90 22.87 21.37 20.29 18.11 Max. (29.5) (29.2) (23.5) (26.4) (27.1) (27.5) (23.1) (21.4) (21.2) (20.3) (20.2) (19.6) (19.8) (20.0) (20.0) (19.5) (17.7) (17.6) (18.8) (18.9) (17.6) (15.7) (16.1) (16.9) Grand 28.94 20.79 20.87 20.23 19.53 18.32 16.99 Length ( μ ) Means Min. conidia on crucifers Measurements of Hyalo peronospora L. Lepidium virginicum Raphanus sativus L. Sisymbrium canescens Sisymbrium canescens Nutt. Lepidium virginicum Lepidium apetalum Willd. (L.) Medic Capsella bursa-pastoris Capsella bursa-pastoris Nasturtium officinale Dentaria laciniata Muhl Cardamine pensylvanica Muhl. Cardamine (L.) Koch nigra Brassica Brassica arvensis (L.) Ktze Brassica Raphanus sativus Walt. caroliniana Draba (Schreb.) BSP. (Schreb.) bulbosa Cardamine Draba caroliniana Draba L. Rorippa parviflora Cardamine parviflora L . parviflora Cardamine R.Br. Nasturtium officinale Sisymbrium altissimum L. Arabis hirsuta Arabis (Muhl.) Poir laevigata Arabis (L.) Scop hirsuta Arabis Host species Yerkes and Shaw ( 1959 ) and Shaw Yerkes Table 3.1 Table 72 3 The Pathogen: Hyaloperonospora parasitica (Gaum.) Goker [H. brassicae (Gaum.… 500(1) 500(1) 500(1) 500(1) 375(15) No. of spores + no. of collections 25(1) 100(1) 750(30) 125(5) 500(1) 25(1) 25(1) 150(6) 825(33) 12.8–25.6 16.0–31.0 8.0–24.0 12.8–28.8 13.7–23.7 Range 15.0–23.7 16.2–26.2 15.0–25.0 16.2–23.7 12.2–27.2 16.2–26.2 17.5–21.2 15.3–25.7 14.6–27.5 20.9 Max. 20.5 20.7 21.3 22.2 23.2 (19.9) (22.9) (17.1) (21.6) (18.9) Grand (19.3) (20.1) (19.6) (20.7) (19.0) (22.2) (19.6) (19.9) (21.0) 17.1 Min. Width ( μ ) Width Means 19.1 18.1 20.5 17.1 17.8 1.36 Max. 1.40 1.47 1.33 1.50 1.6 (1.2) (1.1) (1.5) (1.1) (1.3) Grand (1.4) (1.3) (1.4) (1.3) (1.5) (1.3) (1.5) (1.4) (1.4) 1.25 Min. Quotient (length/width) Means 1.30 1.26 1.28 1.30 1.31 17.6–32.0 17.0–34.0 17.6–32.0 16.0–32.0 17.5–32.5 Range 20.0–33.7 20.0–35.0 20.0–37.5 22.5–33.7 17.2–40.2 25.0–35.0 26.2–36.5 16.9–40.2 20.0–40.0 27.69 Max. 28.72 28.99 27.67 33.44 33.5 (24.3) (26.0) (25.0) (24.9) (24.8) Grand (26.7) (26.6) (26.9) (26.9) (29.1) (29.1) (28.6) (27.5) (29.7) 21.5 Min. Length ( μ ) Means 24.9 24.4 26.2 23.9 23.8 conidia on Chenopodiaceae Measurements of Peronospora Beta vulgaris L. Spinacia oleracea Host species Chenopodium spp. C. bonus-henricus C. hybridum Chenopodium murale L. Spinacia oleracea C. leptophyllum Nutt. C. bonus-henricus L. C. hybridum L. L. C. album C. album C. murale C. gigantospermum Aeller Table 3.2 Table Yerkes and Shaw ( 1959 ) and Shaw Yerkes 3.2 Taxonomy and Morphology 73

Fig. 3.1 Phylogeny of the Peronosporales (Shaw 1981) inferred with ML and MP from the combined dataset; partitioned Bremer support (PBrS) was used to assess potential conflict between data partitions. As in other downy mildew groups, the molecular data clearly corroborate earlier results that too supported the use of narrow species delimitations and host ranges as taxonomic markers. With few exceptions, suggested species boundaries are supported without conflict between different data partitions. The results indicate that a combination of molecular and host features is a reliable means to discriminate downy mildew species for which morphological differences are unknown (Goker et al. 2009). Constantinescu and Faheti (2002) presented molecular and morphological evidence to split Peronospora into three separate genera, Peronospora s. str., Hyaloperonospora, and Perofascia, and the latter two genera were found to be entirely restricted to a single host family, Brassicaceae. Within Hyaloperonospora, only six species were accepted because of the differences in morphology of conidia and conidiophores, and only two accessions (H. niessleana and H. parasitica s. lat. from Thlaspi arvense) were included in their molecular phylogenetic analyses. According to Constantinescu and Faheti (2002), H. parasitica comprises the vast majority of Peronospora infecting Brassicaceae; their concept is similar to that of 74 3 The Pathogen: Hyaloperonospora parasitica (Gaum.) Goker [H. brassicae (Gaum.…

Yerkes and Shaw (1959) and de Bary (1863). Constantinescu and Faheti (2002) proposed to divide Peronospora into three genera, which were confirmed by the molecular studies conducted by Choi et al. (2003), Goker et al. (2003), and Voglmayr (2003). These authors found sequence differences within H. parasitica as large as those between the different Hyaloperonospora species that were morphologically distinguished by Constantinescu and Faheti (2002) and concluded that H. parasitica s. l. should be split up. This result has been confirmed by recent multigene analyses (http://diwww.epfl.ch/wstamatak/index-Dateien/publications/GCB2006_Poster. pdf; Goker et al. 2007, which used several loci but only a relatively small sample of Hyaloperonospora specimens. It turned out that Brassicaceae have had a unique role in DM evolution because the family has apparently been colonized only once, but now is infected by a considerable number of parasitic species. Most probably, Hyaloperonospora was also subject to the highest diversification of any DM genus relative to the number of species of its preferred host family (Goker et al. 2007). Goker et al. (2004) obtained Hyaloperonospora nu-r DNA ITS sequences from a variety of host plants. Molecular results clearly supported the concept of narrowly confined species that were highly specialized to a few closely related or even a single host species and that the morphology-based species circumscription applied by Constantinescu and Faheti (2002) does not meet the criteria of phylogenetic classification (Hennig 1965; Meier and Willmann 2002) because it does not result in monophyletic groups (Goker 2006). Recent molecular phylogenetic investigations show that much narrower species concepts are appropriate not only for Hyaloperonospora (Choi et al. 2003; Goker et al. 2003, 2004) but also for most other DM genera (Voglmayr 2003; Voglmayr et al. 2004, 2006; Garcıa Blazquez et al. 2008; Voglmayr and Constantinescu 2008). This approach to DM taxonomy also implies but in many cases, the separation of taxon, cannot be distinguished by morphological characteristics. This may lead to radical changes in both taxonomic methodology, and taxonomic results, but still this is to be accepted. Goker et al. (2004) included a larger number of collections, as well as sequences from GenBank that were submitted by other workers (Choi et al. 2003) which were critically analysed by Goker et al. (2007) for phylogenetics of Hyaloperonospora. In several studies, the nuITS rDNA has been proven to be a good choice for molecular phylogenetic analysis on the subgeneric level in DMs (Choi et al. 2003, 2005, 2007; Voglmayr 2003; Garcıa Blazquez et al. 2008) and their relatives (Matsumoto et al. 1999; Cooke et al. 2000; Forster et al. 2000; Levesque and De Cock 2004). Additionally, Goker et al. (2009) obtained partial nu LSU rDNA sequences, including the D1, D2, and D3 regions, for the majority of the collections examined. Data from the nu LSU rDNA have so far not been used on the infra-generic level in DMs, but are promising because they are relatively diverse in Hyaloperonospora (Riethmuller et al. 2002; Goker et al. 2003). Phylogenetic analyses of the combined dataset were conducted as an empirical test of whether the species concepts suggested earlier remain stable when more character data are assembled. Further, the relative contribution of the different data partitions was estimated using partitioned Bremer support values (PBrS; Baker and 3.2 Taxonomy and Morphology 75

De Salle 1997; Baker et al. 1998). Therefore, to assess whether branches are unam- biguously supported by the different partitions, this is of particular interest with respect to those branches that seem to separate Hyaloperonospora species. Further, PBrS summary statistics are used to identify the partition that is most informative phylogenetically in Hyaloperonospora, hence the most efficient to use for molecular identification.

3.2.1 Phylogenetic Analyses

According to Goker et al. (2009), the best log likelihood value obtained with RA x ML on the complete dataset was 14885.742; the alpha parameter of the gamma distribution was estimated as 0.293. The best tree is shown in Figs. 3.2 and 3.3 along with the ML BS values. Heuristic MP analysis of the complete dataset yielded 156 most parsimonious trees of length 2225, which were obtained in 182 of the 1 K replicates; the RI was 0.926. The strict consensus tree of the best trees was topologically similar to the ML tree. MP BS values, host names, and assignments of Hyaloperonospora specimens to species are also indicated on the ML tree (Figs. 3.2 and 3.3). Most of the groups were characterized by similar ML and MP support values. Using Perofascia as an out-group, Hyaloperonospora was highly supported as monophyletic by a BS of 100 % (Fig. 3.2). Even though the sequences were sampled from a number of different Lepidium host species, P. lepidii appears as a single and genetically rather uniform species. Within Hyaloperonospora, six diverse clades were recognized comprising more than one species supported with high BS and a number of isolated species. Clade 1 is the sister group to a clade containing the remaining five clades plus all single-species lineages with low (65 %) and high (98 %) BS in ML and MP analysis, respectively (Fig. 3.2). The resolution of the backbone connecting these remaining groups is low, in contrast to the usually highly supported terminal nodes. Clade 1 comprises six clearly distinct lineages that are parasitic on different European and East Asian Cardamine or Rorippa species, European Barbarea vulgaris, or Arabis soyeri. Except for the latter two, these host genera belong to Cardaminineae. Clade 2 comprises collections from European Draba verna and East Asian Draba nemorosa in two distinct clusters, the genetic distance between which is considerable. H. arabidisalpinae separates before clade 2 and H. niessleana after clade 2 on the same grade in the tree, but without support. H. isatidis, H. thlaspeos- arvensis, and the parasite of Biscutella auriculata appear in close relationship to clade 3, but also without support. Clade 3 comprises parasites of a rather diverse assemblage of hosts, such as Reseda, which is one of the few host genera apart from Brassicaceae. The basal split within clade 3 between the East Asian Sisymbrium luteum pathogen and the other specimens is moderately to highly supported (87 to 95 % BS). Some of the subclades of clade 3 are not easy to delineate; it is not clearly evident whether the parasite of Arabis turrita also belongs to H. arabidopsidis or whether H. thlaspeosperfoliati Fig. 3.2 Lower half of the phylogenetic tree inferred from the complete dataset with RAxML under a GTRMIX nucleotide substitution model approximation and rooted with Perofascia. Branch lengths are scaled in terms of the expected number of substitutions per site. Numbers above branches represent BS values above 50 % from ML (left) and MP (right) bootstrapping. Labels for Hyaloperonospora (H.) and Perofascia (P. ) specimens indicate DNA isolation number and host species; if sequences were taken from GenBank, accession numbers are given. Thick vertical bars and adjacent names show the proposed species names. Asterisks mark binomials proposed in the present study. In the case of uncertainty regarding species boundaries, the bars are drawn in light grey. Thin vertical bars and adjacent numbers indicate clades apparently above species level as described in the text. See Fig. 3.3 for the upper half of this tree (Voglmayr 2003) 3.2 Taxonomy and Morphology 77

Fig. 3.3 Upper half of the phylogenetic tree depicted in Fig. 3.2, including clades 4–6. For a description, see Fig. 3.2 (Voglmayr 2003) 78 3 The Pathogen: Hyaloperonospora parasitica (Gaum.) Goker [H. brassicae (Gaum.… includes the specimens from Noccaea caerulescens. Isolated lineages (Fig. 3.3) that are highly supported as monophyletic, but have unresolved interrelationships, are represented by H. cheiranthi, H. hesperidis, H. nesliae, H. camelinae, and H. parasitica, as well as by the pathogens of Lepidium draba, Descurainia sophia, Iberis sempervirens, and Helianthemum (Cistaceae). Clade 4 comprises of H. galligena and H. berteroae, whereas clade 5 consists of two highly distinct lineages of Sisymbrium parasites. The last clade, clade 6, contains a particularly large number of specimens, which is divided into two subclades: specimens that are mainly parasitic on Cardamine (Cardamine and its close relatives, such as Nasturtium) and specimens that are mainly parasitic on Brassica and its close relatives (Brassicaceae; Diplotaxis, Eruca, Erucastrum, Raphanus, Sinapis). This subdivision is strongly supported by both ML and MP analyses. Within the former subclade, the distinction between samples from North America (specimen 21-01), East Asia (GenBank sequences), and Europe is obvious. Besides parasites of Brassicaceae, the second subclade also contains downy mildews of Armoracia, Lobularia, Lunaria, Sisymbrium, and Teesdalia, as well as of Tribulus (Zygophyllaceae). Parasites of Diplotaxis and Sinapis are present within at least two lineages. Both Sinapis and Brassica downy mildews seem to also infect Armoracia. H. lunariae cannot clearly be separated from the downy mildew on Erucastrum nasturtiifolium. A tree search with RAxML was performed on the reduced dataset comprising specimens from which both ITS and LSU could be sequenced. This resulted in 13776.841 as the highest log likelihood value and an alpha parameter estimate of 0.278. The corresponding ML BS values are indicated on the MP consensus tree (Fig. 3.4). A heuristic MP search resulted in 404 most parsimonious trees of length 2066 and a RI of 0.904. MP BS and PBrS values are indicated on the strict consensus of these trees (Fig. 3.4). The symbols on the branches indicate the species boundaries according to Goker et al. (2004) and whether these agree with species delimitations which were suggested by Gaumann (1918, 1923). The total Bremer support (BrS) value obtained from the combined partitions was 1124, i.e. 0.48 per character and 1.60 per informative character. The corresponding PBrS values (total PBrS/total PBrS per character/total PBrS per informative character) were ITS1, 153.41/0.76/1.42; 5.8S, 59.86/0.36/2.22; ITS2, 537.39/0.59/1.61; and LSU, 373.33/0.34/1.61. If the large ITS2 insertions do not significantly contribute to the support for that partition, the total BrS values are 1.13 per ITS2 character and 2.13 per informative ITS2 character. The total BrS values from separate analyses amounted to ITS1, 196/0.98/0.81; 5.8S, 28/0.17/1.04; ITS2, 444/0.49/1.33 (without insertions, 444/0.93/1.76); and LSU, 379/0.35/1.63. The sum of the total support values from the four separate analyses thus amounted to 1047, which is 77 less than the total BrS obtained in combined analysis (Goker et al. 2009). Fig. 3.4 Strict consensus of the 404 most parsimonious trees (length, 2066 bp) inferred from the reduced dataset that contains only specimens from which both ITS and LSU sequences could be obtained. Numbers below branches represent RAxML/GTRCAT (left) and MP (right) BS values above 50 %. Numbers above branches (except terminal ones) are partitioned Bremer support values; the partitions examined were (from left to right) ITS1, 5.8S, ITS2, and LSU rDNA. Numbers above terminal branches represent their average lengths as inferred with DELTRAN optimization, as implemented in PAUP from the same partitions. Specimen labels are as in Figs. 3.2 and 3.3; affiliation of specimens to the clades used in these figures is indicated by vertical bars and numbers on the right side. The following symbols are used to indicate the suggested species boundaries according to Goker et al. (2004), as well as in the present text, and whether these are in accordance with Gaumann’s (1918, 1923, 1926) taxonomy: (-) in agreement with Gaumann; (!) not in agreement, with Gaumann’s species being paraphyletic, and including additional hosts; (x) not in agreement, Gaumann’s species polyphyletic; (0) host not examined by Gaumann; (?) type host not included in our sample. Asterisks point to molecular uncertainty with respect to species boundaries (Voglmayr 2003) 80 3 The Pathogen: Hyaloperonospora parasitica (Gaum.) Goker [H. brassicae (Gaum.…

3.2.1.1 Taxonomy

Hyaloperonospora lobulariae (Ubriszy & Voros) Goker, Voglmayr, & Oberwinkler, comb. nov. Myco Bank No.: MB512921 Basionym: Peronospora lobulariae Ubriszy & Voros, Acta Phytopath. Acad. Sci. Hung. 1: 147 (1966) Hyaloperonospora rorippae-islandicae (Gaumann) Goker, Voglmayr & Oberwinkler, comb. nov. Myco Bank No.: MB512922 Basionym: Peronospora rorippae-islandicae Gaumann, Beihefte zum Botanischen Centralblatt 35: 527 (1918) Hyaloperonospora sisymbrii-sophiae (Gaumann) Goker, Voglmayr & Oberwinkler, comb. nov. Myco Bank No.: MB512923 Basionym: Peronospora sisymbrii-sophiae Gaumann, Beihefte zum Botanischen Centralblatt 35: 529 (1918) Hyaloperonospora species listed, neither in Constantinescu and Faheti (2002) nor in Goker et al. (2003, 2004, 2009), the use of ‘Hyaloperonospora parasitica s. lat.’ is appropriate to recombine Peronospora binomials into Hyaloperonospora if a species is genetically distinct from all other Hyaloperonospora species, and this can be proven. In this manner, superfluous synonyms and naming conflicts can be avoided. Accordingly, it is safe to recombine both P. sisymbrii-sophiae and P. lobulariae. Goker et al. (2009) commented on the complicated taxonomy of the main host from which P. rorippae-islandicae takes its name. For a long time, the diploid Rorippa islandica and the tetraploid R. palustris, which are morphologically similar, were considered synonymous, until Jonsell (1968) finally separated them taxonomically (Bleeker et al. 2002). Jonsell (1968) was able to show that R. islandica has a disjunctive euatlantic-arctic-alpine distribution and is a comparatively rare relic species found in undisturbed habitats. In contrast, the tetraploid R. palustris is a distinctly weedy species, which is common all over Europe. Bleeker et al. (2002) were able to show that R. islandica and R. palustris are not closely related despite their highly similar morphology. Gaumann (1918) described the hosts for ‘Peronospora roripae-islandicae’ as ‘Habitat Roripam islandicam (Meder) [a slip of the pen for ‘Oeder’] Schinz et Thellung et Roripam silvestrem (L.) Besser’ and also cited the host of one collection as ‘Roripa islandica [Oeder] Schinz et Thellung als Nasturtium palustre, Dahlem bei Berlin’. Therefore, it is evident that the main host listed in Gaumann (1918) is not R. islandica s. str., but R. palustris (Nasturtium palustre being a synonym for R. palustris) (Jonsell 1968). Rorippa islandica occurs neither in Germany nor the Czech Republic (Valentine and Jonsell 1993), which is where the specimens cited for R. islandica by Gaumann (1918) were collected. No Hyaloperonospora specimens from R. islandica s. str. have ever been collected. The hosts of the Korean specimens given as R. islandica (Choi et al. 2003; Fig. 3.2) should be reinvestigated, which are unlikely to represent 3.2 Taxonomy and Morphology 81

R. islandica s. str., which is confined to Europe and western Russia; instead, these are likely to belong to R. palustris, which is widespread also in East Asia (Taiyan et al. 2001).

3.2.2 The Species Concept of Downy Mildew

The findings of Goker (2006), Goker et al. (2004, 2007), and Garcıa Blazquez et al. (2008) advocated that downy mildew collections with distinct DNA sequences and hosts should be regarded as different species even if they are morphologically indistinguishable from each other. When applying this concept to the reclassification of Gaumann’s binomials, Goker et al. (2003, 2004) nevertheless assumed that, even though they created many more Hyaloperonospora binomials than Constantinescu and Faheti (2002), superfluous synonyms and other problems with nomenclature were avoided because taxonomical consequences were restricted to collections present in the molecular tree and genetic distances were taken into account. The greatly extended dataset assembled in the study represents an empirical test case for the species concept. To assess whether this concept has met its goals, now focus should be on the taxonomic arrangements suggested earlier (Goker et al. 2003, 2004), which apparently need revision. For instance, H. erophilae is polyphyletic, as two distinct lineages are found on its host, Draba verna. This is a good example of the importance of using molecular examination to detect cryptic species of downy mildews that share a similar morphology and the same host plant. For another case study, refer to Voglmayr et al. (2006). Future research is needed to clarify which of the lineages has to be assigned to an emended H. erophilae. However, this does not disprove the species concept of Goker et al. (2003, 2004) because no superfluous synonyms have been created. Goker et al. (2004) have already found that H. brassicae is non-monophyletic, includes parasites of Armoracia rusticana, and is split into three different lineages corresponding to the formae speciales described by Gaumann (1926). Goker et al. (2009) revealed an additional clade found on Sinapis arvensis (specimen 27-04). As formae speciales are not valid botanical taxon names, new species descriptions will have to be provided. Research still needs to clarify whether morphological differences between these lineages exist. Adding a type sequence to the new species descriptions would be advisable, as was done by Lutz et al. (2005) for Microbotryum saponariae. As no superfluous names have been created, the need to revise H. brassicae is in accordance with Goker et al. (2003, 2004) who did not draw taxonomic consequences for nomenclatural reasons. A similar situation is found in Peronospora sisymbrii-officinalis sensu Gaumann (1918) as parasites of Sisymbrium officinale and S. irio do not cluster together. The European Cardamine parasites ascribed to P. dentariae by Gaumann (1918, 1923) also present a nomenclatural problem, because they are not monophyletic and because the type host, C. heptaphylla, has not been sequenced so far. These examples illustrate that the species concept of Goker et al. (2003, 2004) represents a conservative approach to downy mildew taxonomy. Parasites of hosts not mentioned 82 3 The Pathogen: Hyaloperonospora parasitica (Gaum.) Goker [H. brassicae (Gaum.… by Gaumann and his followers have to be treated with particular caution regarding nomenclature. Goker et al. (2009) dataset includes a couple of Hyaloperonospora specimens from new hosts, like parasite of arugula (Eruca sativa). The economic importance of this host has increased considerably in recent years, and thus its downy mildew attracted the attention of plant pathologists in various parts of the world (Koike 1998; Satou et al. 2004; Larran et al. 2006). Interestingly, molecular data indicate that this parasite is a new species which has long been reported from a number of locations. Finally, in a few cases, molecular phylogenies do not allow species boundaries to be recognized with certainty. For instance, Noccaea and Microthlaspi parasites are not easy to separate from each other if specimen 36-01 is included (Figs. 3.2 and 3.3), which may be caused by the lack of its nu LSU rDNA sequence. Likewise, the Arabis turrita pathogen is nested within H. arabidopsidis in Fig. 3.2. However, these uncertainties have been taken into account by Goker et al. (2004), as neither the Peronospora binomial of the Noccaea nor that of the Arabis turrita parasite have been transferred to Hyaloperonospora. Moreover, genetic distances between the Arabidopsis thaliana and the Arabis turrita parasites are evident (Figs. 3.2 and 3.4); thus, the observed pattern may just reflect the lack of molecular synapomorphies in H. arabidopsidis (Goker et al. 2009). Although genetic isolation is still frequently regarded as the main criteria for separating species, it is not universally applicable and probably should not be applied exclusively, even in the case of sexually reproducing species. Moreover, in the case of Peronospora and Hyaloperonospora, the high host specificity (Gaumann 1918, 1923) strongly suggests genetic isolation (Goker et al. 2004). If these DMs were not (at least to some degree) genetically isolated, it would be hard to explain the distinct host specificities; conversely, if they did not occur on the same hosts, it is not evident how genetic exchange may have occurred in the field. Because the supposed species boundaries based on host specificity are in agreement with the phylogenetic analyses, this obviously non-random distribution of the host species across the terminal taxa in Goker et al. (2009) trees can only be explained by genetic isolation. Biogeography plays a minor role here, because collections from the same hosts cluster together on the trees, regardless whether the hosts were sourced from East Asia (Choi et al. 2003) or Europe, and they displayed only small genetic differences (Figs. 3.2 and 3.3). By contrast, isolates from Central Europe which occur on distinct hosts are for the most part phylogenetically quite divergent. The above listed examples have convincingly shown that based on a broader sample of both specimens, and molecular loci, reconfirms the species concept suggested in earlier publications (Goker et al. 2003, 2004). The vast majority of Hyaloperonospora species delimitations are corroborated, pointing to the stability of the taxonomy, whereas cases of disagreement do not affect the species concept itself, but rather indicate the necessity of future taxonomic work in this group of organisms. Nomenclatural issues have been treated conservatively to avoid the introduction of superfluous taxon names. Researchers have to assess whether a sufficient number of collections and characters per collection have been examined to draw taxonomic conclusions irrespective of the kind of data examined. Thus, one should not refrain from taxonomic 3.2 Taxonomy and Morphology 83 proposals until certain kinds of characters such as morphological characters become accessible. Rather, it may be much less cautious for both taxonomists and practitio- ners to apply the same taxon name to organisms that are clearly distinct biologically. Plant pathologists have frequently been misled by the application of species delimitations that are too broad (Gustavsson 1959). Further, delimiting species based on molecular data and host features alone is justified and does not imply that future work about morphology or other nonmo- lecular characteristics is unnecessary. A thorough and comprehensive examination of Hyaloperonospora morphology may well be able to identify taxonomically useful new characteristics. However, it is highly unlikely that a sufficient number of morphological characters and character states can be determined to separate all species that are classified within the same taxa based on molecular characteristics and hosts. Second, even if a sufficient number of morphological characters were found, it is unlikely that they would contradict the classification based on sequences and hosts. Finally, if disagreement occurred, it would be unlikely that the phenotypic classification would be preferred. Recent studies on downy mildew morphology mostly used molecular data to assess which morphological characters were suitable for taxonomy, and not vice versa (Goker et al. 2003, 2007, 2009; Voglmayr et al. 2004). It is now widely accepted that for many groups, such as bacteria (Schloss and Handelsman 2004), microbial eukaryotes (Moreira and Lopez-Garcia 2002), deep- sea organisms (Sogin et al. 2006), nematodes (Floyd et al. 2002), mites (Ben-David et al. 2007), and Glomeromycota (Husband et al. 2002), which are morphologically indistinguishable or cannot be cultivated, and are beyond the scope of experimental studies, genetic data is often not only the sole source of information but is also sufficient to delimit species. In the case of the DM, the unambiguous support from recent molecular studies for narrow species concepts (Choi et al. 2003; Garcıa Blazquez et al. 2008; Goker et al. 2003, 2004, 2009; Voglmayr 2003; Voglmayr et al. 2004, 2006) also implies that the separation of taxa that are indistinguishable by morphological features has to be accepted. These studies also showed that the vast majority of the species can be interpreted biologically with ease regarding host specialization, a conclusion that is confirmed by Goker et al. (2009), which is based on the most comprehensive molecular sampling for a downy mildew genus to date. Because similar observations have been made in other obligate biotrophic fungi, such as Microbotryales (Lutz et al. 2005; Kemler et al. 2006), Exobasidiales (Begerow et al. 2002a), Entyloma (Begerow et al. 2002b), and ‘true’ smut fungi (Stoll et al. 2003), a combination of molecular and host characters is likely to be of general applicability for these parasites. Future studies will thus not benefit from a return to a focus on morphology, but from making the combined species concept more operational, possibly including an algorithmic integration of sequence clustering (Floyd et al. 2002) and host information (Goker et al. 2009). 84 3 The Pathogen: Hyaloperonospora parasitica (Gaum.) Goker [H. brassicae (Gaum.…

3.2.3 Relationship of Peronospora with Hyaloperonospora and Perofascia

The genera Hyaloperonospora and Perofascia were introduced by Constantinescu (Constantinescu and Faheti 2002 to accommodate the noncoloured downy mildews that had previously been assigned to Peronospora. Perofascia is monotypic and so far only known from Lepidium; it differs from Hyaloperonospora in having hyphal haustoria and curved ultimate branchlets, in which the spiral curving is often less pronounced than in Hyaloperonospora (Constantinescu and Faheti 2002; Thines 2006). Depending on the species concept applied, the genus Hyaloperonospora encompasses only one (Yerkes and Shaw 1959), very few (Constantinescu and Faheti 2002), or based on a narrow species concept (Gaumann 1923; Gustavsson 1959), roughly 100 species. Most of these species are parasitic to Brassicaceae, with only a few exceptions in Zygophyllaceae, Cistaceae, Resedaceae, Capparaceae, and Cleomaceae. Hyaloperonospora can be distinguished from Peronospora on the basis of three main morphological characteristics. First, Hyaloperonospora has uncoloured conidia, whereas in Peronospora, the conidia are with very few exceptions greyish to brownish. Second, haustoria in Peronospora are usually hyphal, whereas in Hyaloperonospora, haustoria are globose to largely lobate when mature. Third, the ultimate branchlets are straight to curve in Peronospora, whereas they are often spiral in Hyaloperonospora. Phylogenetically, Hyaloperonospora and Peronospora are very distinct, and it seems more likely that Hyaloperonospora is the sister genus to the graminicolous downy mildews (Goker et al. 2007) or the downy mildews with pyriform haustoria (Thines 2007) than Hyaloperonospora being the sister genus to Peronospora. As the genus Hyaloperonospora can be easily distinguished from Peronospora both morphologically and phylogenetically, the generic name Hyaloperonospora should be applied for the species included in what Yerkes and Shaw (1959) classified as Peronospora parasitica.

3.2.4 Major Species Clusters in Hyaloperonospora

Molecular phylogenies (Goker et al. 2004, 2009) so far conducted have revealed many distinct lineages in Hyaloperonospora, which mostly represent single species although the backbone of the tree could not be well resolved. However, six clades could so far be identified, which are highly supported, and harbour several distinct species from different hosts (Fig. 3.5). These are clade 1, mainly on Cardamine, and its close relatives; clade 2, on Draba species; clade 3, on a variety of host species including A. thaliana; clade 4, on Aurinia and Berteroa; clade 5, on different Sisymbrium species; and clade 6, on a variety of host genera but with a focus on Brassica and its relatives (wild Brassicaceae) in one subclade and on Cardamine and its relatives (subtribe Cardamininae) in the other subclade. Hyaloperonospora Fig. 3.5 Maximum likelihood phylogenetic tree inferred with RAxML from concatenated internal spacer region (ITS) and large ribosomal subunit (LSU) rDNA sequences. The dataset represents a subset of the one analysed by Goker et al. (2009). Technical details on the inference of this tree as well on the files used and on the origin of the sequences are provided in Goker et al. (2009). Note that the recognition of the clade numbers is based on the extended sampling used in Goker et al. (2009), whereas this figure only shows the specimens for which both ITS and LSU rDNA could be amplified. The numbers above the branches are bootstrap support values equal to or larger than 60% from 100 replicates. Abbreviations: H., Hyaloperonospora; P. , Perofascia (Thines et al. 2009b) 86 3 The Pathogen: Hyaloperonospora parasitica (Gaum.) Goker [H. brassicae (Gaum.… brassicae, which includes the economically important pathogens of Brassica, Raphanus, and Sinapis, is a polyphyletic construct, as specimens from these three host genera are phylogenetically distinct with maximum bootstrap support. The most noteworthy species in clade 6, which also includes H. brassicae, is Hyaloperonospora tribulina. As Hyaloperonospora is primarily parasitic to Brassicaceae (i.e. the most early diverging lineages are parasites of Brassicaceae), the occurrence of Hyaloperonospora on a Zygophyllaceae is the result of a host jump from Brassicaceae to Zygophyllaceae. From the molecular genetic perspec- tive, Hyaloperonospora clade 3 is probably the most interesting because, although it does not include economically important species, this clade includes a parasite of the most intensely studied model plant A. thaliana. Also, Arabidopsis arenosa is among the hosts of clade 3. The investigation of the species closely related to Hyaloperonospora arabidopsidis offers the unique possibility to identify key pathogenicity genes involved in host jumps and pathogen establishment on a specific host. Thus, the general patterns of how plant pathogen evolution and adaptation to specific hosts take place might be revealed. Similar to clade 6, a host jump across host families occurred within the clade 3, with Hyaloperonospora crispula s.l., which is parasitic to R. lutea and Reseda luteola of the Resedaceae.

3.2.5 Hyaloperonospora arabidopsidis on Arabidopsis

In the widespread application of the broad species concept advocated by Yerkes and Shaw (1959), it was commonly believed that Hyaloperonospora on Arabidopsis was the same species as Hyaloperonospora on Brassica and Hyaloperonospora on C. bursa-pastoris, the type host of H. parasitica sensu stricto. This led to confusion and misconceptions over the taxonomy of the Arabidopsis-infecting downy mildew. Research on the genetics of the downy mildew pathogen of Arabidopsis has been justified by, among other arguments, the claims that this pathogen belongs to the same species as economically important lineages that infect Brassica, Raphanus, and Sinapis crops. In addition, Arabidopsis is the most extensively studied model plant for molecular studies, this claim is based on the approach of Yerkes and Shaw (1959). However, it was soon realized that the pathogens from different hosts (e.g. Arabidopsis, Brassica, and Sinapis) were not compatible regarding host range (Tor et al. 1994; Satou and Fukumoto 1996). Therefore, different ‘races’ of Hyaloperonospora on various hosts were named, assuming these would merely represent specialized forms of the same species. In the light of molecular phylogenetic reconstructions (Rehmany et al. 2000; Riethmuller et al. 2002; Voglmayr 2003; Choi et al. 2003; Goker et al. 2004, 2009), this perception needs to change, as these studies revealed that the genus Hyaloperonospora contains a multitude of distinct species, which have partly gone through a long independent evolution. Considering this, the problems in relating results regarding pathogenicity genes and avirulence genes obtained from H. arabidopsidis to H. brassicae become understandable. 3.3 Reproduction and Reproductive Structures 87

Because of the high genetic divergence of H. brassicae and H. arabidopsidis, it is not surprising that pathogenicity genes, which are in common for H. arabidopsidis and H. brassicae, are hard to find. Hyaloperonospora arabidopsidis is the most intensely studied obligate biotrophic pathogen because the host genome was the first plant genome available, whereas H. brassicae is still not well studied in many respects. For H. parasitica – a species so far known only from C. bursa-pastoris – only very few data are available. As this pathogen neither occurs on an economically important nor a genetically well-studied host, it could be assumed that this species will never attract considerable attention. However, it should be borne in mind that C. bursa-pastoris is probably the most widely distributed host for a Hyaloperonospora species and that H. parasitica is among the most often observed downy mildew diseases in natural populations in Europe and therefore is well suited for investigations on ecology and population structure of a downy mildew species (Thines et al. 2009a, b).

3.3 Reproduction and Reproductive Structures

The general morphology and infection cycle of H. parasitica is similar to that of other members of the family Peronosporaceae.

3.3.1 Asexual Phase

Myceliumand haustoria: The mycelium is hyaline and coenocytic. It grows intercellularly in the host tissues and produces haustoria to penetrate the host cells. The haustoria are large, lobed, elongated, or club shaped (Butler 1918; Fraymouth 1956; Holliday 1980). They branch extensively and can nearly fill the entire cell. In the leaf of Japanese radish, the mycelia turn and twist irregularly in the intercellular spaces of the spongy parenchyma and usually develop only one haustorium for each host cell (Ohguchi and Asada 1990). However, in root tissues where parenchyma cells are large, and much closer together, the mycelia are smooth, and one to several haustoria are formed in the infected host cell. Mycelial growth patterns in petioles and hypocotyls are similar to those in root tissue. Prior to haustorium formation, a leaflike structure is formed from the intercellular mycelium in the narrow spaces between root parenchyma cells. It is flat and 6μ m thick and covers the surface of host cell. The leaflike structure forms various types of haustoria, ranging from 0 to 25 numbers in one cell. In turnip and radish roots, the haustoria are initially spherical to pyriform, but later become cylindrical or clavate, and often dichotomously or trichotomously branched (Chu 1935). In cabbage, some haustoria are large irregular vesicles, while others are bilobed and regular in shape. In cauliflower, they are single, globose, and uniform in size. Variations in shape and size of haustoria of H. 88 3 The Pathogen: Hyaloperonospora parasitica (Gaum.) Goker [H. brassicae (Gaum.… parasitica occur in hosts other than Brassica spp., such as Matthiola incana, Cheiranthus cheiri, Capsella bursa-pastoris, Diplotaxis muralis, and Rhynchosynapis manensis (Fraymouth 1956). Penetration of the haustorial branch occurs through a hole 1–2 μm diameter in the cell wall which may form a collar-like structure around the base of the primordial haustorium. As the haustorium enlarges, invigilation of host plasmalemma occurs, and a sheath, possibly of callose, forms around the intru- sive organ (Fraymouth 1956). Moderately high temperatures between 20 and 240C favour the most rapid development of the haustoria (Felton and Walker 1946).

3.3.2 Conidiophores and Conidia

After vegetative growth of the mycelium, erect conidiophores singly or in groups emerge vertically through stomata on the abaxial surface of the host leaves during a period of darkness. The conidiophores are hyaline and measure 200–300 μm. Conidiophores are uniform with a flattened base and stout main axis. At 80C, the rate of elongation reaches 100–200 μm h−1, and the whole process from emergence to spore formation takes approximately 4–6 h (Davison 1968). They are dichotomously branched, six to eight times, tips bifurcate, branching acute, and slightly thickened above each fork. The terminal branches are long, slender, and pointed and end in a single conidium. The sterigmata are slender and acutely pointed (Butler 1918; Holliday 1980; Channon 1981). The conidia are hyaline, broadly elliptic, or nearly globose, measure 24–27 × 15–20 μm, and are delimited from sterigmata by cross walls at maturity. A single conidium is borne at the tip of each branch and is deciduous (Butler 1918; Holliday 1980). Detachment of conidia is possibly caused by hygroscopic twisting of the conidiophores which in turn is related to fluctuations in humidity (Pinckard 1942). Conidia germinate in free water by a lateral germ tube, not by zoospores. Infection occurs both by direct penetration of the epidermis and through stomata (Butler 1918). In cauliflower leaves, conidia form appressoria in the junction areas between the anticlinal walls of adjoining epidermal cells (Preece et al. 1967).

3.3.3 Sexual Phase

Sexual organs, gametogenesis fertilization, and oospore formation: During sexual reproduction, H. parasitica forms spherical oogonia and paragynous antheridia. Oogonia are pale yellow, irregularly round, and swollen into crest-like folds (Butler 1918; Holliday 1980). Antheridia are tendril-like and are produced on separate hyphae. Wager (1889, 1900) observed that the protoplasm of the oogonium becomes differentiated into a central vacuolated ooplasm and a peripheral multinucleate granular periplasm. A receptive thin-walled papilla forms on the oogonium at the point of contact with the antheridium. A fertilizing tube grows from the antheridium 3.3 Reproduction and Reproductive Structures 89 through the receptive papilla towards a ‘central body’ in the ooplasm, to discharge a single ‘male’ nucleus. Meanwhile, a single ‘female’ nucleus detaches itself from the periplasm and also migrates towards the central body. The two nuclei fuse and initiate the uninucleate oospore. During ripening of the oospore, the periplasm is deposited on its wall as an exosporial layer. The oospores are formed in the host tissues at late stage of sporulation. They have also been found in the cavity of the ovary on hyphae emerging between the cells of the inner epidermis of the carpels. The oospore lies inside, almost filling the cavity. The mature oospore is thick- walled, yellow-brown, and globose or spherical and measures 30–40 μm in diameter (Butler 1918; Holliday 1980). Oospore formation is favoured by conditions which induce senescence of the host tissues such as a deficiency of N, P, or K (Mc Meekin 1960). Germination of oospores is by a germ tube (Butler 1918).

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4.1 Introduction

Electron microscopy in association with physiological, biochemical, and genetical studies has provided information which helps in understanding the complex host- parasite relationship of this disease. With the use of light electron microscopy, and transmission electron microscopy, fine structures of H. parasitica have been observed during the process of host penetration, haustorium development, the host- pathogen interface, conidia and conidiophore formation and development, host response and cytology, and genetics of the pathogen.

4.2 Host Penetration

Penetration and haustorial formation in epidermal cells begin 6 h after inoculation (Plate 4.1b, c) (Chou 1970). Appressoria, which look like swollen discs 7–10 μm, across form at the junction of epidermal cells (Plate 4.1b). At this stage, the appressoria and haustoria appear densely granulated, as the spores empty their contents during the process of germination and infection. The penetrating hyphae lay in between the anticlinal cell walls of the two epidermal cells (Plate 4.1c) along with the formation of one or two haustoria, reaching to the adjacent mesophyll cells. After 45 h, intercellular hyphae ramify through more cells and reach the opposite epidermis. At this stage, the haustoria appear broad and conspicuous, reaching 20 μm in length. The intercellular hyphae are about 7 μm, across. Sometimes a sheath can be observed enveloping a fully grown haustorium (Plate 4.1d). The earliest detectable stage of penetration is the formation of penetration hyphae which are as long as the vertical depth of the entire epidermal cell. The thick wall of the appressorium, continuous with the wall of the penetration hypha (Plates 4.1a and 4.3a), has only a thin peripheral layer of cytoplasm. The cell contents migrate into

© Springer Nature Singapore Pte Ltd. 2017 93 G. S. Saharan et al., Downy Mildew Disease of Crucifers: Biology, Ecology and Disease Management, https://doi.org/10.1007/978-981-10-7500-1_4 94 4 Electron Microscopy and Ultrastructures

Plate 4.1 (a) Electron micrograph of TS of epidermal cells of cabbage cotyledon at 6 h after inoculation showing appressorium (ap) and penetrating hypha of Hyalo peronospora parasitica in between the anticlinal walls (j) of host epidermal cells. In one of the cells, a haustorium was formed but the section only shows part of sheath(s). The penetration was cut obliquely, and part of the hyphal wall (arrow pointed) is shown, × 8200; (b) photomicrograph of whole mount of a cleared cabbage cotyledon at 6 h after inoculation showing appressorium (ap) formation predominantly at the junction line of epidermal cells × 313; (c) photomicrograph CTS of cabbage cotyledon at 6 h after inoculation showing penetration as in A. × 500; (d) photomicrograph of whole mount of a cleared cotyledon showing intercellular hypha and haustorium completely ensheathed × 840 (Chou 1970) 4.2 Host Penetration 95

the newly formed penetration hypha. In some cases, the appressorium can be seen to be embedded in an electron-dense, vacuolate material (Plate 4.3b) appearing to be a mucilaginous sheath. This sheath is bound by an outer membrane which adheres to the cuticle of the host epidermis except in the penetration region where it is slightly separated (Plate 4.3b). The penetrating hyphae wedge into the middle lamella between the anticlinal walls of two epidermis cells. The hole in the wall through which the fungus penetrates is 4–5 μm across. After entering the host, the hypha expands to a diameter of 7–8 μm. There is no clearing zone or dissolution of wall material in the immediate vicinity of the penetrating hypha. The penetrating hypha is always seen to be embedded in a moderately electron-dense matrix of the middle lamella (Plates 4.1a and 4.3a). The cuticle breaks and fits closely around the penetrating hypha. No sign of swelling or change in electron density of the cuticle can be detected in the immediate vicinity of the penetration zone (Chou 1970). After 80 h, hyphal growth develops conidiophores which may be seen coming out from the epidermal cells (Chou 1970). The intercellular hyphae appear to aggre- 96 4 Electron Microscopy and Ultrastructures

Plate 4.3 (a) Electron micrograph of TS of epidermal cells of cabbage cotyledon at 6 h after inoculation. The penetration region was cut medially through showing the appressorium which is almost empty with cytoplasm migrating into the penetrating hypha, × 14400; (b) electron micrograph of TS of appressorium and part of host epidermal cells showing the mucilaginous sheath of the appressorium. Membranous boundary of the mucilaginous sheath is shown by arrow, × 6000; (c) electron micrograph of section of a haustorium initial in host epidermal cell. Pan of the neck of a fully grown haustorium is shown by its side. Note the hyphal wall is continuous with wall of the intercellular hypha at this stage, × 13800; (d) electron micrograph of a section of intercellular hypha and host epidermal cell showing pan of host wall in contact with hypha is swollen and partially eroded (arrow), × 17700 (Chou 1970) 4.3 Haustorium Development 97 gate beneath the epidermis and grow either through stomata or in between two epidermal cells to the outside of the host tissue (Plate 4.2a). An electron-dense spearhead-like thickening of the hyphal tip is observed to wedge in between two guard cells. This thickening may give rigidity to the hyphal tip for penetration. The hyphae are cemented to each other and also to the host cell walls by an amorphous, moderately electron-dense material, presumably of a mucilaginous nature (Plate 4.2a, b). Hyphae penetrating through the junction of epidermal cells invariably show a dieback of the tip (Plate 4.2a). A new wall is laid down round the remaining living cytoplasm, while a new growing tip is organized to carry on further growth (Plate 4.2b). Large numbers of lomasomes appear around the newly formed walls, and numerous dense vesicles approximately 500–1000 Å in diameter are concentrated in the walled-off cytoplasm.

4.3 Haustorium Development

Host penetration by haustoria of the Peronosporales is usually by boring a narrow canal at the point of contact between the hypha and the host cell wall (Fraymouth 1956). However, according to Chou (1970), it is not possible to find the stage at which the walls of both host and pathogen are perforated prior to haustorial initiation. Localized swelling of the host wall (3X original) is observed in the area of hyphal contact. The swollen area is about 1.5–2 μm long and shows a clearer fibrillar structure, with a partially eroded area (Plate 4.3d). The dimension of the swollen region coincides closely with the size of the hole in the host wall made by the haustorium. These observations strongly suggest that the breach of host wall during haustorium initiation is achieved at least partly by chemical means. A dome-shaped protuberance, about 1 μm in diameter, is formed by the bulging of the wall of intercellular hypha into the lumen of host cell (Plates 4.3c and 4.4a). The host wall is perforated at this stage, and the wall of the haustorium primordium is continuous with that of the intercellular hypha (Plate 4.3c). The haustorium initial is completely enclosed in a mound-like sheath quite distinct from the host wall in structure as well as density. The perforation made by the invading haustorium measures 1–2 μm across. The perforated host wall in most cases remains smooth, but a slight unfolding of the wall to form a short collar-like structure is sometimes observed. The external part of the wall of the haustorial initial consists of a very electron-dense layer varying in thickness from 0.1 to 0.2 μm and exhibiting an undulating surface bounded externally by a thickened membrane (Plates 4.3c and 4.4a) which is presumably the invaginated plasmalemma of the host. The primordial haustoria are filled with homogeneous groundplasm packed with ribosomes. Lomasomes are the only organelles present at this stage. The growth and differentiation of the primordial haustorium is in the form of an elongated neck and expanding head (Plate 4.4b). The sheath seems to burst apart, remaining as a collar-like structure around the neck region (Plate 4.4b). 98 4 Electron Microscopy and Ultrastructures

In a young haustorium, the contents are invariably dense with a high population of ribosomes, a profuse system of endoplasmic reticulum, and relatively few vacuoles (Chou 1970). The dictyosomes occur more frequently, and the mitochondria are strikingly irregular (Plates 4.5, 4.6 and 4.9b). The same pattern of these structures is also present in young penetration hyphae. A complicated membrane system of unknown nature and origin is always present (Plate. 4.10b, c). One type consists of a complicated system of tubules and vesicles enclosed by a unit membrane. The inter-tubular spaces do not contain ribosomes. This organelle looks like a lomasome except that there is no apparent connection with plasmalemma. Another type consists of whorls of closely packed membranes formed in vacuoles (Plate 4.10c). Generally the lomasomes are more or less hemispherical to saucer shaped, about 0.2–0.3 μm in the longer diameter, but occasionally they can extend to 2–3 μm in diameter (Plate 4.6). The tubules and vesicles of lomasomes range from 15 to 80 μm in diameter. The nuclei are about 3–3.5 μm in diameter. As many as three sections of nuclei are observed in one haustorium section (Plate 4.9a). The nuclei envelope consists of a double membrane interrupted by pores. The envelope is very similar in form to the endoplasmic reticulum, and connections between these two are often observed. The endoplasmic reticulum is mainly of the smooth type (Plates 4.9a and 4.6) enlarged in part to form cisternae of various forms. The mitochondria are large (1–2 μm in diameter), usually elongated dumb bell shaped, or irregularly branched (Plates 4.6, 4.9, 4.1a and 4.3d). Those in old haustoria are roundish with a much less dense matrix (Plate 4.9b).

4.4 The Host-Pathogen Interface

The external surface of the haustorial walls always appears to consist of very electron-dense layer (Plates 4.4b, 4.5, 4.6, 4.9a, 4.10a and 4.11c, d) which is well developed at the earliest stage of haustorial development (Chou 1970). The outer region of the hyphal wall can be further differentiated into a well-defined, very dense, and thin outer boundary, about 50–100 Å thick, and an inner less dense zone of rather obscure lateral limit (Plate 4.8e). The hyphal wall thus appears to be a three-layered structure. The zone of apposition of the haustorial wall consists of a well-defined, very dense, and thin outer layer approximately 50–100 Å thick and a broad, slightly less dense inner zone without a well-defined boundary (Plates 4.8d). Chou (1970) proposed that the zone of apposition should be termed as an outer and inner dense zone being both an integral part of the haustorial wall. The surface of the haustorium neck is covered by a dense layer much thicker than that of the rest of the haustorium. Its surface always appears to be deeply roughened with numerous vesicular and tubular extensions (Plates 4.6, 4.7b and 4.11b). Dense granular bodies can sometimes be seen lodged between the surface of the dense layer and the invaginated host plasmalemma and in both of the matrix of the dense layer and of the tubular extension (Plates 4.7a and 4.11b). There is a frequent occurrence of a porous substance of uniform pore diameter (about 200 Å) covering the entire haustorium 4.4 The Host-Pathogen Interface 99

Plate 4.4 (a) Electron micrograph of a section of a haustorium initial in host mesophyll cell (section slightly oblique to the penetration zone), × 16500; (b) electron micrograph of a section of a very young haustorium in host epidermal cell showing breakdown of host cytoplasm into large number of vesicles, × 18000 (Chou 1970)

surface. The host plasmalemma covering the haustorium surface is often masked due to the accumulation of this substance (Plate 4.11a, d). As soon as the haustorium penetrates the host, the haustorium becomes covered with a layer, moulded to its shape, and produced by the host protoplast (Fraymouth 1956). This layer is named ‘The Sheath’ and appears to be composed of modified cellulose and callose. A sudden increase in the growth rate of the fungus often causes the sheath to burst, remaining as a collar around the base. A morphologically analogous structure enveloping a haustorium initial which has penetrated the host wall (Plates 4.3c and 4.4a) has been detected in the cabbage – Hyaloperonospora system by Chou (1970). In mature haustoria which have differentiated into a neck and head, the sheath remains as a collar-like structure at the base (Plates 4.5 and 4.6) although completely ensheathed mature haustoria are sometimes observed under the light microscope (Plate 4.1d). 100 4 Electron Microscopy and Ultrastructures

Plate 4.5 Electron micrograph of a section of a haustorium in host mesophyll cell at 6 h after inoculation, x 1 2000 (Chou 1970)

Electron microscope observation revealed that the sheath is a saclike structure sometimes flattened to a narrow strip (Plate 4.5), but in most cases dilated to a broadly conical shape and quite distinct in texture and electron density from the host wall. The sheath is bounded by a unit membrane which is generally presumed to be the host plasmalemma. No membranous structure can be detected along the sheath/ host wall interface, though the two can be clearly distinguished by their difference in electron density and texture. The sheath matrix is electron transparent, while the host wall is moderately electron dense and often exhibits a fibrillar structure (Plate 4.2a). The sheath matrix is always permeated by large numbers of blurred electron- dense granules and dense vesicles with single or double membranes. These vesicles appear to be of host origin, as they are also found in the adjacent host cytoplasm (Plates 4.7b, d and 4.6). The sheath matrix is also interspersed with host cytoplasm (Plate 4.7d) which occurs in isolated packets or as an extension of the adjoining host 4.4 The Host-Pathogen Interface 101

Plate 4.6 Electron micrograph of a section of a haustorium in host mesophyll cell at 6 h after inoculation, showing the saclike sheath and numerous vesicles (arrow pointed) and intra-vacuolar vesicles (pointed out by double arrow) in the sheath matrix, × 8580 (Chou 1970) cytoplasm. The permeation of vesicles into the sheath matrix and the extension of host cytoplasm within it suggest a liquid or semi-liquid state of the sheath matrix (Fraymouth 1956; Chou 1970). During penetration, the host cytoplasm adjoining the sheath increases markedly in amount and comes to contain a large number of vacuoles (Plates 4.5, 4.6 and 4.8a, c) (Chou 1970). At an early stage of haustorium development, coalescence of these vacuoles with the sheath can be seen. Intrusion of intra-vacuolar vesicles in the host cytoplasm and in the sheath can be seen (Plates 4.6 and 4.7a, c). The ultrastructure of the interface between H. arabidopsidis and A. thaliana has very nicely been described by Mims et al. (2004). The diagrammatic representation 102 4 Electron Microscopy and Ultrastructures

Plate 4.7 (a) Electron micrograph of a section of part of haustorium neck and sheath, × 24600; (b) electron micrograph of a section of part of haustorium neck and sheath showing numerous vesicles (arrow pointed) and dense granules in the sheath matrix (Smx) and the dentate extensions (pointed out by double arrow) of the dense zone (z) of haustorium wall, × 33000; (c) electron micrograph of a section of the interface between haustorium and host cytoplasm showing a dense vesicle (arrow pointed) like the secretary body, × 33000; (d) electron micrograph of a section of haustorium sheath showing incorporation of host cytoplasm (arrow pointed) in the sheath matrix and numerous membrane-bounded vesicles both in host cytoplasm and the sheath matrix, × 33000 (Chou 1970) 4.4 The Host-Pathogen Interface 103

Plate 4.8 (a) Electron micrograph of a section of haustorium in host mesophyll cell showing the vacuoles or pro-vacuoles possibly in the process of fusion with each other and also with the sheath (arrow), × 7200; (b) electron micrograph of a section of the interface between haustorium and host cytoplasm showing vesiculation of the host plasmalemma, × 48000; (c) electron micrograph of a section of haustorium in host mesophyll cell showing fusion of vacuoles in host cytoplasm and sheath formation, × 7200; (d) electron micrograph of a section of interface between haustorium and host cytoplasm showing the structure of outer dense zone of haustorium wall distinguished into two well-defined layers (z1) and (z2), × 49500; e( ) electron micrograph of a section of intercellular hyphae showing the hyphal wall also exhibiting a dense outer layer composed of z1 and z2 × 33000 (Chou 1970) 104 4 Electron Microscopy and Ultrastructures

Plate 4.9 (a) Electron micrograph of a section of haustorium in epidermal cell at 6 h after inoculation showing the typical fine structure of haustorium at this stage. Ring formation in mitochondria pointed out by arrow, × 13200; (b) electron micrograph section of haustorium in epidermal cell 45 h after inoculation, × 24000 (Chou 1970) 4.4 The Host-Pathogen Interface 105

Plate 4.10 (a) Electron micrograph of a section of the interface between haustorium and host cytoplasm of mesophyll cell showing sphaerosome-like bodies (arrow pointed) in host cytoplasm, × 33000; (b and c) electron micrograph of sections of young penetrating hyphae, (b) showing complicated membrane system of unknown nature and (c) showing intra-vacuolar membrane systems, × 55200 and 36000, respectively (Chou 1970) 106 4 Electron Microscopy and Ultrastructures

Plate 4.11 (a) Tangential section of the dense zone of haustorium neck showing foldings of host plasmalemma (arrow pointed) forming tubular extensions and incorporation of numerous dense granules (d), × 33000; (b) electron micrograph section of haustorium in host epidermal cell showing lomasome, × 55200; (c) electron micrograph of a section of haustorium in host epidermal cell showing pinocytotic vesicles formed from host plasmalemma and abundant porous substance (arrow pointed) at the host-parasite interface, × 73200; (d) electron micrograph section of interface between dense zone of haustorium and host cytoplasm showing the deposition of porous substance (arrow pointed), × 48000 (Chou 1970) 4.4 The Host-Pathogen Interface 107

Plate 4.12 Schematic representation of a haustorium of Hyaloperonospora arabidopsidis in an Arabidopsis host cell (based on electron micrographs from Mims et al. 2004). C collar of host material (including callose), CC host cell cytoplasm, CV host cell vacuole, CW host cell wall, EHMa electron-dense extra-haustorial matrix, EHM extra-haustorial membrane, G Golgi body, H haustorium, HM hyphal membrane, HW hyphal wall, Ne neck region, PM host plasma membrane, HW hyphal wall, ICH intercellular hypha, L electron-dense lipid vesicle, M mitochondrion, N nucleus, Ne constricted neck region, P plastid, PM invaginated host cell plasma membrane, T tonoplast membrane of the host cell vacuole, V vacuoles in the haustorium, Ves vesicles either fusing with or budding off from the extra-haustorial matrix. in Plate 4.12 revealed the vesicles (Ves) at the interface of the extra-haustorial matrix (EHMa) and the host cell cytoplasm (CC). It could be interpreted that these vesicles are budding off from, or coalescing with, the extra-haustorial membrane or both. Thus, these vesicles may represent a major mechanism and site of information/nutrient exchange between host and pathogen. The collar of host cell material could be decorated with immunogold particles labelled with a monoclonal antibody recognizing β-1,3-glucan epitopes showing that it is at least partially composed of callose. The host cell plasma membrane (PM) could be clearly observed to be invaginated by the haustorium, becoming the EHM. Other interesting features of the haustorium are the presence of multiple vacuoles and several nuclei. Golgi bodies, which are absent in true fungi but present in oomycete cells, were also clearly visible (Schlaich and Slusarenko 2009). 108 4 Electron Microscopy and Ultrastructures

Plate 4.13 Ultrastructural features of the structures produced by Hyaloperonospora parasitica in susceptible accessions of Arabidopsis. Sections were taken from samples 3 (a–d), 5 (e), and 7 (f) days after inoculation. (a) Median section through a penetration site showing an intercellular hypha from which two haustoria penetrate two different host mesophyll cells of Ws-eds1. Note the presence of nucleus, lipid bodies, mitochondria, and large vacuoles in the intercellular hyphae. (b) Median section through a penetration point and two haustorial bodies in a host mesophyll cell of Ws-eds1. Note that the old haustorium (with haustorial neck) contains organelles such as mitochondria and nucleus. (c) Median section through a penetration point and haustorium in a mesophyll cell of Oy-0. Callose deposition (*) occurred at the penetration point around the proximal region of the haustorial neck. Note the presence of mitochondria and small and large vacuoles in the haustorium. The wall of the intercellular hypha is at its thickest where it penetrates the host cell wall to form the haustorial neck. The extra-haustorial matrix (arrows) is present around the haustorium. (d) Median section through a haustorium within the mesophyll cell of Ws-eds1. The cytoplasm of the intercellular hypha and haustorium contains mitochondria, lipid bodies, and small and large vacuoles. The host mesophyll cell appears unaffected by the presence of the haustorium as organelles are well preserved. (e) and (f) Callose deposition in Oy-0 (e) and Ws-eds1 (f). Callose 4.5 Ultrastructural Features of Intercellular Hyphae, Haustorium, and Host Cell 109

4.5 Ultrastructural Features of Intercellular Hyphae, Haustorium, and Host Cell

Observation of infected tissue under transmission electron microscopy (TEM) revealed the coenocytic hyphae, with an average diameter of 7.2 μm, ramified, and spread intercellularly throughout the host tissue (Plate 4.13a). The cytoplasm of intercellular hyphae was bounded by the pathogen plasma membrane and contained many ribosomes, endoplasmic reticulum (ER), lipid bodies, mitochondria, and nuclei. Small and large vacuoles containing moderately electron-dense material occupied a significant volume of the hyphae (Plate 4.13a–d). Further growth of hyphae within intercellular spaces and penetration of individual host mesophyll cells have been resulted in the formation of haustoria. Intracellular haustoria were large and lobate with an average diameter of 5.4– 7.1 μm. A typical mature haustorium is differentiated into a very short neck and head (Plate 4.13c, d). The haustorial neck is constricted at the penetration site, ranging in width from approximately 0.4 to 0.95 lm. Callose-like deposits were frequently observed at sites of penetration around the proximal region of the haustorial neck (Plate 4.13c). Median sections through penetration sites revealed that the pathogen wall was continuous from the intercellular hypha along the entire haustorial body (Plate 4.13c, d). The cell wall consisted of a single layer that was thicker around the intercellular hyphae (0.34 μm) compared with that around the haustorial body (0.18 μm). The pathogen plasma membrane was continuous from the intercellular hypha throughout the haustorium and was tightly pressed to the haustorial wall. Haustoria in infected cells were surrounded by an invagination of the host plasma membrane, the extra-haustorial membrane (EHM). EHM is separated from the pathogen wall around the haustorial body by an electron-dense extra-haustorial matrix (Plate 4.13c). The extra-haustorial matrix is a non-cytoplasmic part of haustorium. This region was variable in width up to approximately 0.3 μm thick and was located around the haustoria and the neck region (Plate 4.13c). The cytoplasm of the haustorium contained organelles, which were similar to that of the intercellular hyphae. Large vacuoles were common in hyphae and older haustoria (Plate 4.13a– e). The haustorial body contained more mitochondria than other parts of pathogen (Plate 4.13a). One nucleus with dense heterochromatic regions was observed in haustoria but is not a constant feature (Plate 4.13b). During the early stages of infection, one to two haustoria were often observed in single mesophyll cells (Plate 4.13a, b), but as many as four to five haustorial profiles were found within a single cell at 5 dai. Organelles within the penetrated host cell

Plate 4.13 (continued) deposition stained lightly around the haustorium shown in (e) but densely around the haustorium and in the cell wall (*) shown in (f). Note that contents of the haustorium and infection hypha appear normal during the early stage of infection (e) as organelles are clearly distinguished in the host cytoplasm. The contents of the haustorium and the infection hypha (arrow) became necrotic at the late stage of infection (f). All bars ¼ 3 lm. H haustorium; IH intercellular hypha; IS intercellular space; m mitochondrion; n nucleus; ca callose; Cv cell vacuole (Soylu and Soylu 2003) 110 4 Electron Microscopy and Ultrastructures did not appear to be affected following formation of haustoria (Plate 4.13d). The region around and between the haustoria contained an abundance of cytoplasm with many vesicles, plastids, mitochondria, and microbodies. Occasionally, ensheathed healthy haustoria were observed at infection sites. In general, the cytoplasm of the ensheathed haustorium and associated intercellular hypha were apparently normal at 5 dai (Plate 4.13e). By 7 dai, however, ensheathed haustoria and associated intercellular hypha became necrotic as evident by their misshapen, irregular appearance, and dense contents (Plate 4.13f) (Soylu and Soylu 2003). Certain ultrastructural features of the haustorial apparatus of H. parasitica on susceptible Arabidopsis plants do not resemble those of H. parasitica on cabbage (Chou 1970) and other relevant Peronospora species such as Peronospora pisi in pea plants (Hickey and Coffey 1977). Ultrastructural features of the H. parasitica haustorium clearly distinguish the pathogen from the downy mildews in other crops. Unlike other downy mildews, which produce simple, cylindrical haustoria with a narrow cylindrical long neck, the complete structural unit of a haustorium produced by H. parasitica consists of a lobe-shaped haustorial head and extra-haustorial matrix in the absence of an apparent haustorial neck. Features of coenocytic intercellular hyphae were similar to those of other downy mildew pathogens. The cytoplasm of intercellular hyphae contained typical organelles characteristic of the pathogen. Large vacuoles containing moderately electron-dense material occupied a significant volume of the hyphae. The pathogen cell wall consisted of single layer of consistent thickness around the intercellular hyphae except at the penetration points where it was frequently thicker. As described for other downy mildews (Chou 1970; Hickey and Coffey 1977), the haustorial body contained a normal comple- ment of organelles, usually including one to two nuclei. Observation of several median sections of haustoria revealed that the hyphal wall appears to be continuous from the intercellular hypha at the penetration sites, then through the host wall, and finally around the haustorial body. Median sections through penetration sites revealed the presence of cell wall depositions (papillae) at the site of penetration. Aniline blue fluorescence confirmed the presence of callose- like material in these depositions. Callose-like deposits or papillae are frequently reported at sites of penetration by other species of Peronosporales (Hickey and Coffey 1977; Sargent 1981; Woods et al. 1988; Cohen et al. 1989; Enkerli et al. 1997). Deposition of papillae adjacent to the penetrating haustoria represents a nonspecific host cytoplasmic response to both mechanical and pathogenic injury (Ingram et al. 1976). Among obligate fungal pathogens, the invagination of the plasma membrane by a haustorium has been termed the EHM. In H. parasitica, the haustorium was also surrounded by an EHM. The EHM has been shown to differ from the host plasma membrane in a number of aspects (Spencer-Phillips and Gay 1981; Roberts et al. 1993). However, at penetration sites, a clear continuity between the membrane binding the dense staining matrix and the plasma membrane was not always definitely established. The EHM is separated from the fungal structures in the haustorium by a gel-like substance termed the extra-haustorial matrix. The extra-haustorial matrix completely surrounded the haustorium of H. parasitica (Soylu and Soylu 2003). 4.6 Conidiophore Development 111

4.6 Conidiophore Development

Conidiophore development of H. parasitica can be divided into five stages (Davison, Davison 1968a, b, c).

4.6.1 Conidiophore Primordial

The emergence of H. parasitica from the host cotyledons during sporulation can be seen as a densely stained region beneath the host stomata. In the sub-stomatal space, a hyphal branch, about 5 μm in diameter, grows towards the stoma and then between the guard cells (Plate 4.14a). When the tip of this hypha is about level with the top of the guard cells, it becomes more rounded and completely blocks the stomatal pore (Plate 4.14b). It is referred to as conidiophore primordium. According to Shiraishi et al. (1975), a contracted image is found in the region when the conidiophores develop.

4.6.2 Unbranched Conidiophores

It is the earliest stage of conidiophore development visible on the surface of the host and can be seen about 4 h after the cotyledons are placed in a moist, dark environment. From the primordia, unbranched conidiophores may develop immediately, or a narrow wall surrounding a ‘blowout’ forms at the apex (Plate 4.14b). The basal constriction surmounted by a bulge which is observed in older conidiophores is probably the result of the ‘blowout’ formation. Developing conidiophores are more or less cylindrical at this stage (Plate 4.14c), approximately 10–12 μm in diameter and of varying length.

4.6.3 Production of Branches

When the conidiophores reach about two third of its eventual height, branches are formed one at a time, just behind the conidiophore apex (Plate 4.14d). Secondary and tertiary branches are also formed which are narrower than the primary ones. The conidiophore axis is also narrower at the apex, with decrease in branch diameter being proportional to the increase in branch length. The ultimate branches are very slender, often about 1 μm in diameter, and usually curved. Branches form at a projected angle of 55–85° with the major axis, and the number produced is approximately proportional to the conidiophore height. 112 4 Electron Microscopy and Ultrastructures

4.6.4 Development of Conidia

Young conidia are formed about 2 h after initiation of branch production. Initially, conidia are spherical, but as they increase in size, they become ellipsoidal (Plate 4.14e). Conidia produced on a single conidiophore are of uniform size, but conidia borne by different conidiophores frequently vary in size.

4.6.5 Formation of a Cross Wall

Conidia are delimited by a cross wall about 2 h after the beginning of conidial formation, when they reach about 15 × 20 μm in size (Plate 4.14h). However, the cross walls are observed only occasionally since spores are usually detached before cross wall formation.

4.6.6 Conidiophore Growth

The increase in conidiophore length shows an initial slow period of elongation, just after the fungus has emerged from the host leaf followed by a rapid increase (Plate 4.15; Figs. 4.1 and 4.2). During branch formation, increase in length is slightly slower and less regular, while just before spore formation, conidiophore elongation slows down and almost ceases. Once formed, the spores enlarge rapidly, but increase in conidiophore length is only by the enlargement of the apical spore. The most rapid rate of elongation of conidiophores is 100–200 μm/h. As the branches usually begin about two third of the way up the final length of the conidiophore stalk, late conidiophores are usually shorter, are less profusely branched, and bear fewer spores than the conidiophores formed earlier. Although increase in volume may be approximately linear during the growth of unbranched conidiophores, and branch production, there is a decrease in the rate of volume increase just before spore formation. This is followed by a massive increase in volume just after spore formation, when the total conidiophore volume may be more than quadrupled (Fig. 4.3) depending on the number of spores produced. The inflation of the branch apex is a gradual process which occurs without any interruption (Fig. 4.4) in branch elongation (Davison 1968b). 4.6 Conidiophore Development 113

Plate 4.14 (a) Section of wax-embedded material showing a hyphal branch growing towards a stoma; (b) section of wax-embedded material illustrating two conidiophore primordia, one of which is beginning to grow; (c) stained and macerated preparation of an unbranched conidiophore; (d) stained and macerated preparation of a branched conidiophore; (e) a branched conidiophore with small spores in a stained and macerated preparation; (f) very young spores; (g) mature spores; (h) mature spores delimited by a cross wall (arrow); (1-L) frames from the cine film illustrating the development of conidiophores A, B, and C; (i) incubation time 3 h 30 min.; (j) incubation time 3 h 50 min.; (k) incubation time 4 h 10 min.; and (l) incubation time 4 h 30 min. A–H scale line is 10 μm; I–L scale line is 100 μm (Davison 1968b) 114 4 Electron Microscopy and Ultrastructures

Plate 4.15 Continued development of conidiophores: (a) incubation time 4 h 50 min.; (b) incubation time 5 h 10 min.; (c) incubation time 5 h 30 min.; (d) incubation time 5 h 50 min.; (e) incubation time 6 h 10 min.; (f) incubation time 6 h 30 min.; (g) incubation time 6 h 50 min.; and (h) incubation time 7 h 30 min. Scale line is 100 μm (Davison 1968b)

4.6.7 Conidial Formation

Surface ultrastructure of conidia, germ tubes, appressoria, and conidiophores of H. parasitica infecting Japanese radish has been observed by Shiraishi et al. (1974) through scanning electron microscopy (Plates 4.16, 4.17, 4.18 and 4.19). Mature conidia are approximately 7 μm in width and 10 μm in length. Conidia are formed directly from the swelling tips of the conidiophores, and they have the same surface structure as the conidiophores. Old conidia have many wartlike structures, although the mature conidiophores have a smooth surface (Plates 4.17 and 4.19). 4.6 Conidiophore Development 115

sp 500

400

300 br br sp br Height in µ

br 200

100

0 2 3456789 Incubation time in hours

Fig. 4.1 Increase in length of five individual conidiophores growing in the humidity chamber; br, time at which branching commenced; sp., spore formation (Davison 1968c)

4.6.8 Host Response

The host protoplast of epidermal and mesophyll cells responds differently to infection by the downy mildew pathogen (Chou 1970). The epidermal cells in most cases respond vigorously to infection resulting in a severe disruption of the protoplast. The central vacuoles contract and undergo fragmentation. The plasmalemma is broken down or detached from the wall, and numerous vesicles are formed from it. The cytoplasm, which originally appeared as a thin peripheral coating of the wall, is either dislocated and aggregated into a vacuolated blob or completely dispersed to the extent that its identity cannot be discerned. Apparently intact mitochondria and chloroplasts appear to be set free from the groundplasm. Consequently, haustoria in epidermal cells are not surrounded by a clearly defined layer of host cytoplasm. Haustoria formation in a mesophyll cell is less disruptive. The host cytoplasm is invaginated by the invading haustorium, while the tonoplast and plasmalemma remain intact. 116 4 Electron Microscopy and Ultrastructures

500 sp br br

br B sp 400 br br br C br sp br br br br br br br 300 br br br

br br Height in m br 200 br

100

0 2.5 4.0 4.5 5.0 5.5 6.0 6.57.0 7.58.0 Incubation time in hours

Fig. 4.2 Increase in length of conidiophores A, B, and C. br, formation of primary branch; sp., spore formation (Davison 1968c)

4.6.9 Cytology and Genetics

The haploid chromosome number of H. parasitica is n = 18–20 and it is a tetraploid (Sansome and Sansome 1974). Nuclei, mitochondria, lipid material, protein, and RNA in the intercellular mycelia and haustoria of H. parasitica are uniformly distributed (Davison 1968a). Insoluble carbohydrate material has been detected in the fungal cell wall. Callose sheaths are occasionally seen partially surrounding the haustoria. A distinct plasmalemma, porate nuclei, tubular endoplasmic reticulum, 4.6 Conidiophore Development 117

A 14

12 B

10 4

x10 C 3 8

6 Volume in µ

4 sp

2 sp

0 3.5 4.0 4.5 5.0 5.56.0 6.57.0 7.5 Incubation time in hours

Fig. 4.3 Increase in volume of conidiophores A, B, and C. sp., spore formation (Davison 1968c)

20 A

µ 10 B

0 01020304050 Time in minutes

Fig. 4.4 Increase in branch length and apical diameter during spore formation. I branch length, b apical diameter (Davison 1968c) 118 4 Electron Microscopy and Ultrastructures

Plate 4.16 Electron micrograph of conidia, germ tubes, and initial period of Hyaloperonospora parasitica invasion on Japanese radish leaves. (a) Mature conidium. The surface is rough, with wart-shaped structure; (b) separation of a mature conidium from its conidiophore; (c) an appressorium above a stoma and a penetration peg into the stomatal cavity; (d) enlargement of C. Wrinkly structures on an appressorium in the initial period of formation; (e) an appressorium over a stoma 48 h after germination; (f) enlargement of E. Slight degeneration of the epidermal cells where the appressorium is in contact with the stomatal guard cells; (g) cuticular invasion. Germ tube growing from the side of a spore; (h) enlargement of G. The appressorium is quite contracted (Shiraishi et al. 1975) 4.6 Conidiophore Development 119

Plate 4.17 Electron micrograph of initial period of Hyaloperonospora parasitica invasion on Japanese radish leaves. (a) Invasion through a junction between a stomatal guard cell and an auxiliary cell; (b) cuticular invasion of an auxiliary cell. The germ tube is quite extended, but invasion does not depend on a stoma being present; (c) enlargement of B. The viscous substance used by the appressorium to adhere to the epidermal cell wall is not very visible; (d) enlargement of C. The germ tube and appressorium are clearly contracted, and circular traces of where the penetration peg has entered can be seen in the epidermal cell wall; (e) cuticular invasion with a long germ tube; (f) cuticular invasion through a short germ tube. Although the conidium is adjacent to a stoma, germination has occurred from the conidium wall on the side away from the stoma, and cuticular invasion is taking place (Shiraishi et al. 1975) 120 4 Electron Microscopy and Ultrastructures

Plate 4.18 Electron micrograph showing development of conidiophores and conidia of Hyaloperonospora parasitica on Japanese radish leaves. (a) Conidiophores invariably grow out of stomata, sometimes two at a time; (b) a conidiophore branching during the initial stage of new growth; (c) surface of a conidiophore during the initial stage of new growth, with a wavy structure; (d) an extended conidiophore with appearance of a crimp at the base; (e) initial stage of conidium formation. The tips of the conidiophore swell, forming conidia. The conidiophores and conidia have similar surface structures; (f) clusters of conidia that have matured and begun to take on a tuft-like shape (Shiraishi et al. 1975) 4.6 Conidiophore Development 121

Plate 4.19 Electron micrographs showing conidiophores and conidia of Hyaloperonospora parasitica on Japanese radish leaves. (a) Conidiophores without conidia. The area at the top right is a relatively young diseased area, and exfoliation of epidermal cell wax and cuticular material can be seen; (b) diseased area with advanced signs of disease. Wrinkles have appeared in the epidermis of the diseased area, and open stomata can be seen; (c) diseased area with many developed conidiophores; (d) diseased area with advanced symptoms of disease. A crimp in the base of the conidiophore is visible. Yeast-shaped fungi are also present; (e) stoma in a healthy area. It is formed of two stomatal guard cells and several auxiliary cells; (f) the base of the conidiophore is crimped, perhaps due to mechanical force exerted by the stoma. Wrinkles on the surface of the host are clearly visible (Shiraishi et al. 1975) 122 4 Electron Microscopy and Ultrastructures

Fig. 4.5 The distribution of (a) nuclei, (b) RNA, and (c) mitochondria in the developing conidiophores of Hyaloperonospora parasitica (Davison 1968c) 4.6 Conidiophore Development 123 mitochondria with tubular cristae, Golgi dictyosomes, and lipid bodies are present within the protoplast (Ehrlich and Ehrlich 1966). The distribution of organelles, storage products, and other substances within the developing conidiophores of H. parasitica is very different from the distribution observed within the intercellular mycelium (Davison 1968c). In developing conidiophores, the nuclei, mitochondria, protein, and lipid material are more or less uniformly distributed at first, but gradually shift into the conidia so that by maturity all these substances have relocated, leaving the conidiophore stalk and branches almost completely empty (Plate 4.19; Figs. 4.5, 4.6 and 4.7). Glycogen has not been detected within conidiophores or conidia of H. parasitica. Trehalose and either glucose or mannose are identified in the conidiophores and conidia of H. parasitica but sugar alcohols are absent.

Fig. 4.6 Migration of nuclei (n) into the nucleate spores of Hyaloperonospora parasitica (Davison 1968c) 124 4 Electron Microscopy and Ultrastructures

Fig. 4.7 The distribution of (a) lipid material, (b) protein, and (c) insoluble carbohydrates in the developing conidiophores of Hyaloperonospora parasitica (Davison 1968c) References 125

References

Chou CK (1970) An electron-microscope study of host penetration and early stages of haustorium formation of Peronospora parasitica (Fr.) Tul. on cabbage cotyledons. Ann Bot 34:189–204 Cohen Y, Eyal H, Hanania J, Malik Z (1989) Ultrastructure of Pseudoperonospora cubensis in muskmelon genotype susceptible and resistant to downy mildew. Physiol Mol Plant Pathol 34:27–40 Davison EM (1968a) Cytochemistry and ultrastructure of hyphae and haustoria of Peronospora parasitica (Pers. ex Fr.) Fr. Ann Bot 32:613–621 Davison EM (1968b) Development of sporangiophores of Peronospora parasitica (Pers. ex Fr.) Fr. Ann Bot 32:623–631 Davison EM (1968c) The distribution of substances in sporangiophores of Peronospora parasitica (Pers. ex Fr.) Fr. Ann Bot 32:633–647 Ehrlich MA, Ehrlich HG (1966) Ultrastructure of the hyphae and haustoria of Phytophthora infestans and hyphae of Peronospora parasitica. Can J Bot 44:1495–1504 Enkerli K, Hahn MG, Mims CW (1997) Ultrastructure of compatible and incompatible interactions of soybean roots infected with the plant pathogenic oomycete Phytophthora sojae. Can J Bot 75:1493–1508 Fraymouth J (1956) Haustoria of the Peronosporales. Trans Br Mycol Soc 39:79–107 Hickey EL, Coffey MD (1977) A fine-structural study of the pea downy mildew fungus Peronospora pisi in its host Pisum sativum. Can J Bot 55:2845–2858 Ingram DS, Sargent JA, Tommerup IC (1976) Structural aspects of infection by biotrophic fungi. In: Friend J, Threlfall DR (eds) Biochemical aspects of pant- and parasite relationships. Academic, London, pp 43–78 Mims CW, Richardson EA, Holt BF III, Jl D (2004) Ultrastructure of the host-pathogen interface in Arabidopsis thaliana leaves infected by the downy mildew Hyaloperonospora parasitica. Can J Bot 82:1001–1008 Roberts AM, Mackie AJ, Hathaway V, Callow JA, Green JR (1993) Molecular differentiation in the extrahaustorial membrane of pea powdery mildew haustoria at early and late stages of development. Physiol Mol Plant Pathol 43:147–160 Sansome E, Sansome FW (1974) Cytology and life history of Peronospora parasitica on Capsella bursa-pastoris and of Albugo candida on C. bursa-pastoris and on Lunaria annua. Trans Br Mycol Soc 62:323–332 Sargent JA (1981) The fine structure of the downy mildews. In: Spencer DM (ed) The downy mildews. Academic, London, pp 183–236 Schlaich NL, Slusarenko A (2009) Downy mildew of Arabidopsis caused by Hyaloperonospora arabidopsidis (formerly Hyaloperonospora parasitica) (Chapter 13). In: Kurt L, Kamoun S (eds) Oomycete genetics and genomics: diversity, interactions and research tools. Wiley, Hoboken, pp 263–285 Shiraishi MK, Sakamoto S, Asada Y (1974) An electron microscope study of Japanese radish leaves infected with Peronospora parasitica. Mem Coll Agric Ehime Univ 19:137–168 Shiraishi M, Sakamoto K, Asada Y, Nagatani T, Hidaka H (1975) A scanning electron microscopic observation on the surface of Japanese radish leaves infected by Peronospora parasitica (Fr.) Fr. Ann Phytopathol Soc Japan 41:24–32 Soylu EM, Soylu S (2003) Light and electron microscopy of the compatible interaction between Arabidopsis and the downy mildew pathogen Peronospora parasitica. J Phytopathol 151:300–306 Spencer-Phillips PTN, Gay JL (1981) Domains of ATPase in plasma membranes and transport through infected plant cells. New Phytol 89:393–400 Woods AM, Didehvar F, Gay JL, Mansfield JW (1988) Modification of the host plasmalemma in haustorial infections of Lactuca sativa by Bremia lactucae. Physiol Mol Plant Pathol 33:299–310 Chapter 5 Physiologic Specialization (Pathogenic Variability)

5.1 Introduction

Specificity in the downy mildew fungus on crucifers is very complex since it occurs on a wide range of wild hosts as well as agricultural and horticultural species. However, there has been little sustained effort to introduce resistance to the disease, and hence there has been less selection pressure exerted on the pathogen population than is the case with many other obligate parasites. Further impetus has been added by the exponential growth in research on the wild crucifer Arabidopsis thaliana, as a host for Hyaloperonospora parasitica and serving as a model system for genetic and molecular analysis (Uknes et al. 1992). Discontinuities in the host range of isolates from different host genera and species suggest that the fungus may exist as a series of pathotypes adapted to each host of origin, although some cross-infections may occur. There is also growing evidence that within host species, specificity may be determined by genotype-specific interactions consistent with a gene for gene recognition system (Lucas et al. 1988, 1994; Nashaat and Awasthi 1995). Specificity might therefore be expressed at several levels including family, genus, species, and cultivar or accession. In view of the close cytogenetic relationship between the major Brassica species, coupled with the strongly outbreeding nature of several of these, some overlap in the host range of species-adapted isolates is perhaps predictable. At the generic level, pathogenic specialization has been observed by several workers all around the world. Gardner (1920)) and Kobel (1921) suggested that H. parasitica is highly specialized and seldom occurs in the same biological form on more than one crucifer. An isolate of H. parasitica obtained from turnip is able to infect seedlings of turnip but not rutabaga or radish (Gardner 1920). In Holland disease on cabbages is classified in two groups, both representing distinct biological forms of the fungus. The first is characterized by short, ellipsoid conidia and the second by larger, elongated conidia with protuberant apices. The average dimen-

© Springer Nature Singapore Pte Ltd. 2017 127 G. S. Saharan et al., Downy Mildew Disease of Crucifers: Biology, Ecology and Disease Management, https://doi.org/10.1007/978-981-10-7500-1_5 128 5 Physiologic Specialization (Pathogenic Variability) sions of the later group are 32.51 × 25.66 μm and that of the former 26.67 × 23.13 μm, the corresponding ratios of length to breadth being 1.26 and 1.11, respectively (Thung 1926a). However, Gaumann (1926) subdivided H. parasitica (= H. brassicae) into three biological strains, namely, 1. f. sp. brassicae, the chief hosts of which are B. oleracea, B. napus, B. rapa, B. nigra, B. juncea, B. tournefortii, and B. fruticulosa, but can also cause some infection on Sinapis arvensis, S. alba, Raphanus raphanistrum, R. sativus, and Eruca sativa; 2. f. sp. sinapidis, the principal hosts of which are S. arvensis and S. alba, but are also able to produce sub-infections on all the above-mentioned species of Brassica (except B. rapa and B. juncea) and Raphanus, with occasional conidiophore formation on B. oleracea; and 3. f. sp. raphani, the chief hosts of which are R. raphanistrum and R. sativus, but can also produce sub-infections on all the above-mentioned species of Brassica (except B. fruticulosa), as well as on S. arvensis and S. alba, with occasional conidiophore formation on B. oleracea and B. napus. The downy mildew on radish does not attack cabbage (B. oleracea var. bullata and capitata) and is slightly pathogenic on Chinese cabbage (B. pekinensis, B. chinensis), rape (B. campestris), and mustard (B. juncea) (Hiura and Kanegae 1934). Conversely, the form derived from B. pekinensis does not infect radish but is allied to one on B. chinensis and rape. The forms derived from B. pekinensis, B. chinensis, and rape are mutually pathogenic on one another (Hiura and Kanegae 1934).

5.2 Pathogenic Variability

The identification of pathogenic variability seems to be initiated in 1944 with the observation of Wang (1944) who classified the reaction of the hosts into four categories: 1. susceptible with normal symptoms; 2. resistant, showing large necrotic spots; 3. para-immune, showing slightly visible necrotic dots; and 4. immune, with no visible symptoms. Three pathotypes of H. parasitica were differentiated: H. parasitica Brassicae on Brassica, H. parasitica Raphani on Raphanus, and H. parasitica Capsellae on Capsella. The three pathotypes were not mutually compatible with each other’s host. Six forms of H. parasitica Brassicae were differentiated by their reaction to B. chinensis, B. oleracea, B. juncea, and B. napobrassica. Wang (1944) prepared a dichotomous key to physiological forms from China as follows: A. Capsella bursa-pastoris, immune B. Raphanus sativus, immune or para-immune variety Brassicae C. B. oleracea, resistant or para-immune D. B. chinensis, susceptible E. B. juncea (Meitan Dav Yu Tsai), susceptible Ph.fm. 1 EE. B. juncea (Meitan Dav Yu Tsai,) resistant F. B. napobrassica, immune Ph. fm. 2 FF. B. napobrassica, resistant Ph. fm. 3 DD. B. chinensis, resistant E. B. juncea (Dav Ching Tsai), susceptible Ph. fm. 4 5.2 Pathogenic Variability 129

EE. B. juncea (Dav Ching Tsai), resistant Ph. fm. 5 CC. B. oleracea, susceptible Ph. fm. 6 BB. R. sativus, susceptible variety Raphani AA. C. bursa-pastoris, susceptible variety capsellae Felton and Walker (1946) and Natti (1958) differentiated the races of H. parasitica found on R. sativus and B. oleracea on the basis of their host specificity. Morris and Knox-Davies (1980) also indicated distinct races of H. parasitica on B. oleracea and R. raphanistrum based on host specificity. H. parasitica f. brassicae on cabbage, f. rapae on turnip, f. rapifera on B. rapa, f. rapifera, f. napi on rape, f. raphani on radish, and f. sinapsidis on Sinapis alba have been distinguished as special forms of H. parasitica from Leningrad though all are similar morphologically (Dzhanuzakov 1963). Three vars. of H. parasitica have been differentiated in 35 samples of downy mildew from B. pekinensis and other crucifers, namely, f. sp. brassicae on Brassica, f. sp. raphani on Raphanus, and f. sp. capsellae on Capsella (Chang et al. 1964). Hyaloperonospora parasitica f. sp. brassicae exists in at least three different sub- forms (pekinensis, oleracea, and juncea). Isolates from B. pekinensis, B. chinensis, and turnip were classified in the same group and can attack all three hosts but did not infect Capsella bursa-pastoris, radish, cabbage, Chinese mustard, and B. juncea var. multiceps. B. juncea var. megarrhiza expressed various reactions to these isolates. Isolates from Chinese mustard, B. juncea var. megarrhiza and B. juncea var. multiceps, were limited to these hosts, except that those from Chinese mustard did not infect some vars. of B. pekinensis, B. chinensis, and turnip. Radish isolates are of two types, one infects only radish, the other vars. of B. pekinensis, cabbage, and turnips as well as radish. Isolates from cabbage and C. bursa-pastoris are host specific. In Norway, cross-inoculation experiments with downy mildew from cabbage, turnip rape, and radish indicated the occurrence of different races on cabbage and radish (Semb 1969). According to Natti et al. (1967), the predominant physiologic race of H. parasitica pathogenic to broccoli and other types of B. oleracea grown commercially in New York were race 1 and race 2. The later race was pathogenic to plants resistant to race 1. Dickinson and Greenhalgh (1977) observed a wide variation in the reaction of seedlings of different crucifers’ species to isolates of Hyaloperonospora derived from Brassica and Raphanus species (Table 2.3). Masheva et al. (1996a) found that the pathogenicity of two isolates of H. parasitica (one from Plovdiv and one from Goma Oriahovitsa, Bulgaria) towards ten hybrids of cabbage, broccoli, and cauliflower differed in pathogenicity and host specificity. In India, H. parasitica isolates from different hosts vary in host range. Isolates from Brassica, Raphanus, Eruca, and Sisymbrium are not cross-infective (Bains and Jhooty 1983). Mehta and Saharan (1994) tested the host range of 9 isolates of H. parasitica collected from the leaves and stag heads of 6 host species on 17 host differentials (Tables 5.1 and 5.2). Isolates from Brassica oilseeds infected all species, except B. alba, whereas isolates from cauliflower leaves did not infect B. carinata, B. alba, B. nigra, B. chinensis, B. pekinensis, and B. napus (Table 5.2). There was no significant difference among the conidial size of the isolates collected from 130 5 Physiologic Specialization (Pathogenic Variability)

Table 5.1 Host differentials of Hyaloperonospora parasitica (Mehta and Saharan 1994) Common name Botanical name Cultivar Indian mustard (raya) Brassica juncea RH-30 Toria Brassica campestris var. toria TH-68 Yellow sarson B. campestris var. yellow sarson YSPb-24 Brown sarson B. campestris var. brown sarson BSH-1 Ethiopian mustard B. carinata HC-1 White mustard B. alba Local Black mustard B. nigra Local Chinese mustard B. chinensis Local Chinese mustard B. pekinensis Local Rapeseed B. napus GSL-1 Wild turnip B. tournefortii Local Cabbage B. oleracea var. capitata Pride of India Cauliflower B. oleracea var. botrytis Snowball-16 Turnip B. rapa White Purple Top Knol khol B. caulorapa Early White Vienna Taramira Eruca sativa Local Radish Raphanus sativus HR-1 leaves and stag heads, but significant differences were observed among these groups (Table 5.3). The isolates were classified into two distinct pathotypes, one from cauliflower and the other from oilseeds Brassica. There was no significant difference between the isolates in percentages of spore germination (Table 5.4). Specific populations of H. parasitica differing in pathogenesis and host specificity were reported from Bulgaria (Masheva et al. 1996a, b). The populations formed at a lower temperature were more aggressive on cabbage heads. In the UK, differential host resistance in relation to pathogenic variation of isolates derived from the same host species was identified in B. rapa (Moss et al. 1991; Silve et al. 1996), B. napus (Nashaat and Rawlinson 1994), B. juncea (Nashaat and Awasthi 1995), and B. oleracea (Silve et al. 1996). Isolates from different Brassica species found to be most virulent on their species of origin were nevertheless able to grow to less extent on other Brassica species (Sherriff and Lucas 1990). Nashaat and Awasthi (1995) identified five groups of B. juncea accessions with differential resistance to UK isolates Rl and P003 derived from oilseed rape (B. napus ssp. oleifera) and Indian isolates IP01 and IP02 derived from mustard (B. juncea) (Table 5.5). All B. juncea accessions were resistant to isolates from B. napus, but at the same time, B. napus cv. Arian was resistant to isolates from B. juncea. Twenty-one differential responses to H. parasitica isolates from B. oleracea and two from B. rapa were identified. Of the seven isolates tested, four were from crops of cauliflower in France, two from oilseed rape in the UK, and one was from mustard in India. All Raphanus sativus accessions were resistant to all seven isolates (Silve et al. 1996). 5.2 Pathogenic Variability 131 Inflorescence Inoculum B. nigra + (4) + (4) + (4) + (4) + (4) – + (4) + (4) + (4) + (4) + (4) + (4) + (4) + (4) + (4) + (4) + (4) Leaf inoculum Cauliflower + (4) + (4) + (4) + (4) – – – – – – + (4) + (4) + (4) + (4) + (4) + (4) + (4) Inflorescence inoculum + (4) + (4) + (4) + (4) + (4) – + (4) + (4) + (4) + (4) + (4) + (4) + (4) + (4) + (4) + (4) + (4) Leaf inoculum + (4) Raya + (4) + (4) + (4) + (4) – + (4) + (4) + (4) + (4) + (4) + (4) + (4) + (4) + (4) + (4) + (4) Inflorescence inoculum + (4) + (4) + (4) + (4) + (4) – + (4) + (4) + (4) + (4) + (4) + (4) + (4) + (4) + (4) + (4) + (4) Leaf inoculum Brown sarson Brown + (4) + (4) + (4) + (4) + (6) – + (4) + (4) + (4) + (4) + (6) + (4) + (4) + (4) + (4) + (4) + (4) Inflorescence inoculum + (4) + (4) + (4) + (4) + (4) – + (4) + (4) + (4) + (4) + (4) + (4) + (6) + (4) + (4) + (4) + (4) Leaf inoculum isolates/reactions Sources of H. parasitica sarson Yellow + (4) + (4) + (4) + (4) + (6) – + (6) + (6) + (6) + (4) + (6) + (4) + (4) + (4) + (4) + (4) + (4) leaf inoculum Toria + (4) + (6) + (4) + (4) + (6) – + (6) + (6) + (6) + (4) + (6) + (4) + (4) + (4) + (4) + (4) + (4) (Mehta and Saharan 1994 ) parasitica species to nine isolates of Hyaloperonospora Brassica Response of seventeen Brassica campestris Brassica toria var. Differential hosts Differential var. B. campestris var. sarson yellow var. B. campestris var. sarson brown Brassica juncea Brassica B. carinata B. alba B. nigra B. chinensis B. pekinensis B. napus B. tournefortii Eruca sativa Raphanus sativus var. var. B. oleracea capitata var. var. B. oleracea botrytis B. rapa B. caulorapa ; 1, leaf inoculum; 2, inflorescence inoculum (), incubation period in days; +, infection; –, no B, Brassica Table 5.2 Table 132 5 Physiologic Specialization (Pathogenic Variability)

Table 5.3 Conidial size of Hyaloperonospora parasitica isolates derived from eleven Brassica species (Mehta and Saharan 1994) Conidial dimensions (μ) H. parasitica Range Average Sources of isolates isolatesa Length Width Length Width

Brassica campestris var. toria T1 19.50– 16.62– 25.93 19.30 29.25 27.30

B. campestris var. yellow YS1 21.45– 19.50– 25.35 21.74 sarson 29.25 24.37

B. campestris var. yellow YS 2 19.50– 19.50– 26.81 23.39 sarson 29.25 24.37

B. campestris var. brown BS 1 21.45– 19.00– 25.35 21.45 sarson 29.25 24.37

B. campestris var. brown BS2 21.93– 19.50– 26.56 23.64 sarson 29.25 26.81

Brassica juncea R1 20.47– 19.50– 25.64 21.84 29.25 26.32

Brassica juncea R2 19.50– 19.50– 27.05 22.90 29.25 26.81

Brassica oleracea var. C1 19.50– 17.55– 23.30 20.76 botrytis 29.25 24.37

B. tournefortii BT2 19.50– 16.57– 23.59 20.96 24.37 19.50

Raphanus sativus RS2 19.50– 18.52– 22.03 19.69 24.37 21.45

B. nigra BN2 21.93– 19.50– 26.81 22.90 29.25 26.81 LSD (P< 0.05) – – 2.43 1.78 asource of inoculum; 1, leaves inoculum; 2, hypertrophied inflorescence

Table 5.4 Percent conidial germination of Hyaloperonospora parasitica isolates at 18°C (Mehta and Saharan 1994) Cauliflower Toria Brown sarson Yellow sarson Raya Incubation Leaf Leaf Leaf Leaf Leaf period inoculum inoculum inoculum inoculum inoculum 0.0 – – – – – 0.5 0 0 0 0 0 1.0 0 0 0 0 10.57 1.5 12.20 13.73 15.72 11.87 32.46 2.0 36.50 33.25 45.87 39.31 52.68 3.0 66.30 63.25 69.12 61.06 69.12 4.0 84.25 83.86 80.99 81.92 88.85 5.0 86.02 84.91 82.61 84.16 88.85 6.0 86.51 (68.72) 87.36 (70.11) 82.90 (65.80) 85.42 (67.85) 89.19 (73.20) LSD (P< 0.05) NS 5.2 Pathogenic Variability 133

Table 5.5 Sources of seed for accessions of Brassica juncea, arranged in five groups according to the response of their seedlings at the cotyledon stage to Hyaloperonospora parasitica and one accession of B. napus (Nashaat and Awasthi 1995) Brassica junceaa Seed source Brassica juncea Seed source Group A Group C RES-BJ01 ( Kranti) (India) Ecotype/BGRC 34263 FAL RES-BJ02 ( Krishna) (India) BGRC 34283 FAL RES-BJ03 ( Varuna) (India) BGRC 34291 FAL RES-BJ04 ( BGRC 34253) (FAL) BGRC 34781 FAL BGRC 34789 FAL BGRC 46069 FAL Group B Change Yang Huang Jie HAU BGRC 46071 FAL BGRC 324294 FAL PPBJ-1 India Skorosjelka-2 Group C BGRC 34275 FAL Aurea/BGRC 28602 FAL Stephniacka Blaze/BGRC 30288 FAL BGRC 34274 FAL Burgonde/BGRC 30289 FAL Stoke/BGRC 51764 FAL Commercial brown Ag Cda Yi Men Feng Wei Zi HAU Cutlass Ag Cda Zaria/BGRC 16254 FAL Ecotype BGRC 34255 FAL Group D Group E Hatano/BGRC 22527 FAL BGRC 34282 FAL BGRC 34239 FAL Landrace/BGRC 46323 FAL B. napus Larja/BGRC 34273 FAL Ariana Semundo Line/BGRC 34295 FAL aRES-BJ01 to RES-BJ04 lines selected from seedling population of accessions in parenthesis

Differential resistance of rapid cycling and commercial genotypes of B. rapa were also reported from the UK (Moss et al. 1988). A range of differential host responses were characterized by four homologous isolates (Table 5.6). Pathogen isolates were also characterized in relation to their sexual compatibility type and response to phenylamide fungicides (Moss et al. 1988). Eleven isolates of H. parasitica tested on rapid cycling populations of B. rapa (aa, CrGc-1-1), B. nigra (bb, CrGc-2-1), B. oleracea (cc, CrGc-3-1), B. juncea (aabb CrGc-4-1), B. napus (aacc CrGc-5-1), B. carinata (bbccCrGc-6-1), and R. sativus (rr CrGc-7-1) indicated specificity towards particular genotypes within each rapid cycling population (Hill et al. 1988). Variation in the response of different host lines to H. parasitica has also been detected within wild crucifer species such as shepherd’s purse (Capsella bursa-pastoris) and Arabidopsis thaliana (Lucas et al. 1994). Moss et al. (1994) attempted to cross fungal isolates originating from different Brassica spp. by co-inoculating host lines previously identified as susceptible to these 134 5 Physiologic Specialization (Pathogenic Variability)

Table 5.6 Differential virulence of Hyaloperonospora parasitica isolates from B. campestris on six host lines (Moss et al. 1988, 1991) Reaction of H. parasitica isolates Host lines P007 P008 P013 P014 CA88014*a + + + + JADE PAGODA + – + + CA87063* – – + + SNOWBALL – – + – CA87068* – – – + CA87065* – – – – +, susceptible; –, resistant; a, universally susceptible; *, rapid cycling lines isolates. A proportion of oospore progeny recovered from these crosses appeared to be hybrids and had reduced virulence on both hosts of origin. The differential resistance to H. parasitica identified in Brassica spp. can be used for future studies of the genetics of the host-pathogen interaction and for breeding for disease resistance. The pathogenicity of four isolates of H. parasitica obtained from Japanese radish, rape, broccoli, and shepherd’s purse (Capsella bursa-pastoris) was compared by observing infected cotyledons. These isolates eventually sporulated on host plant cotyledons (compatible combinations), but formed only small necrotic lesions on non-host cotyledons (incompatible combinations). Sheath encasement of haustoria and host cell death frequently occurred in the incompatible combinations. The C. bursa-pastoris isolate could only infect the host plant with haustorium formation in non-hosts inhibited mainly by papillae. Cytochemical observation showed that these papillae contained callose and polyphenol compounds but not lignin (Yoshida and Ohguchi 1998). Silve et al. (1996) identified 21 differential responses to H. parasitica isolates from B. oleracea and 2 from B. rapa. All Raphanus sativus accessions were resistant to all seven isolates. Accessions for which seedling populations exhibited a heterogeneous reaction to some isolates were classified in a separate category. However, Mehta and Saharan (1994) classified 9 isolates ofH. parasitica (collected from Brassica species) into 2 distinct pathotypes, 1 from cauliflower, and the other from oil-yielding Brassica species on the basis of their reaction to a set of 17 host differentials. Variation has been found among interaction phenotypes when a range of A. thaliana (At) and Brassica accessions are inoculated with different isolates of H. parasitica (Hp). In Brassica, genetic characterization of this variation was rudimentary, and only one allele at a single locus associated with isolate-specific resistance toH. parasitica in B. napus has been defined. In contrast, and considering only those phenotypes that can be discriminated readily among segregating individuals, alleles controlling genotype-specific variation for response to these pathogens have been identified at 21 loci (RPP and RAC) in the At genome. Most of these loci now have been mapped with varying degrees of resolution and occur in several linkage groups in four of the five At chromosomes (Leckie et al. 1996). 5.2 Pathogenic Variability 135

Eight B. oleracea lines were evaluated for their phenotypic reaction to the thirteen European isolates of H. brassicae (Table 5.7) by Coelho et al. (2012). The host lines exhibiting a similar phenotypic pattern to different isolates were clustered into six classes (A–F) according to their response to the isolates (Table 5.8). The control lines cv. Senna were highly susceptible with no visible host response to all of the pathogen isolates tested (designated class A) mean IP≥5.9, showing a consistent susceptible response (Tables 5.9 and 5.10). The two class F lines (PCA 20.14 and EBH 527) were resistant or moderately resistant to all thirteen isolates (mean IP≤3.0). The remaining five lines exhibited resistance to at least one, but not all of the isolates including a single line in class B (EBH508) which exhibited resistance to three isolates (Hp006, Hp 539, and Hp 704), and were highly susceptible to the remaining isolates: a class C line (EBH 502) which was resistant at least to some degree to seven isolates, a class D line (EBH 525) which was fully resistant to eleven isolates, and two class E lines (PC10 and PC11) which were resistant to all of the isolates except for high susceptibility to the single FP 06 isolates (from France). Six different pathotypes of H. brassicae were identified by classifying the mean IP as either being compatible (mean IP ≥4.5), incompatible (mean IP <2.5), or intermediate (Table 5.8). Isolates showing the same virulence pattern on the differential set of host lines were grouped and considered to belong to the same pathotype. Pathotypes 1, 4, and 6 were represented by a single isolate, Hp006, Hp806, and FP06, respectively. Other pathotypes were represented by more than one isolate including two examples of pathotype 2 (Hp539 and Hp704), five of pathotype 3 (Hp 530, Hp533, Hp535, Hp717, and Hp710), and three of pathotype 5 (Hp541, Hp702, and Hp801). The three referred cases with multiple isolates included examples from different countries. For example, pathotype 3 includes isolates from Italy and two from diverse locations in Portugal (Odemira and Azores Island) and the UK (Coelho et al. 2012). Tham et al. (1994) used RAPD analysis to compare 16 isolates of H. parasitica from two different Brassica species, B. napus and B. oleracea. Two out of twenty random primers screened gave reproducible band patterns capable of dis- criminating between the different host-adopted isolates (Plate 5.1).

Table 5.7 Brassica oleracea standard host differentials to classify pathotypes of Hyaloperonospora brassicae (H. parasitica) (Coelho et al. 2012) Host line (code) Crop type Description Origin EBH502 (JV06092) Cabbage (subsp. capitata) Double haploid* UK

EBH508 (JV06096) Calabrese (subsp. botrytis) S4 inbred* UK

EBH525 (JV06085) Borecole (subsp. acephala) S4 inbred* UK EBH527 (JV06103) Romanesco cauliflower (subsp. botrytis) Double haploid* UK

PC11 (OL87098) Broccoli (subsp. italica) S4 inbred Portugal

PC10 (OL87123) Broccoli (subsp. italica) S4 inbred Portugal

PCA20.14 Couve de Corte (subsp. tronchuda) S4 inbred Portugal

Senna (GK97362) Rapid cycling brassica S4 inbred UK *Lines derived from crosses between rapid-cycling and the indicated ‘crop-type’ brassicas 136 5 Physiologic Specialization (Pathogenic Variability) 1.7 1.8 1.5 1.5 1.7 3.7 5.1 6.0 Mean 1.0±0.0 1.5±0.2 6.0±0.0 5.9±0.1 5.8±0.1 5.8±0.1 6.0±0.0 6.0±0.0 FP06 PT 6 3.0±0.2 2.0±1.0 1.5±0.2 1.4±0.2 1.8±0.3 5.9±0.1 6.0±0.0 6.0±0.0 801 UK 1.4±0.2 1.0±0.0 1.0±0.0 1.0±0.0 1.0±0.0 5.9±0.1 6.0±0.0 5.9±0.1 702 UK 1.1±0.1 2.9±0.3 1.0±0.0 1.0±0.0 1.0±0.0 5.0±0.2 6.0±0.0 6.0±0.0 541 PT 5 2.9±0.4 2.8±0.5 2.1±0.3 3.1±0.3 5.0±0.3 3.8±0.1 6.0±0.0 6.0±0.0 806 UK 4 2.7±0.4 2.3±0.3 1.3±0.2 1.0±0.0 1.1±0.1 2.4±0.2 6.0±0.0 6.0±0.0 710 UK 1.8±0.3 1.2±0.2 1.0±0.0 1.0±0.0 1.0±0.0 2.8±0.4 5.5±0.3 6.0±0.0 717 UK 1.1±0.1 1.1±0.1 1.0±0.0 1.0±0.0 1.0±0.0 3.0±0.3 6.0±0.0 6.0±0.0 717 PT 5.10 ). Means and standard errors were calculated from a total of 18 plants (6 x 3 1.1±0.1 1.4±0.1 1.0±0.0 1.0±0.0 1.0±0.0 3.3±0.2 6.0±0.0 6.0±0.0 535 PT 1.3±0.2 1.9±0.3 1.0±0.0 1.0±0.0 1.0±0.0 1.1±0.1 5.5±0.1 6.0±0.0 533 PT 1.1±0.1 1.4±0.1 1.0±0.0 1.0±0.0 1.0±0.0 1.6±0.2 5.6±0.1 6.0±0.0 530 PT 3 2.8±0.3 2.1±0.3 1.0±0.0 1.0±0.0 1.0±0.0 4.9±0.2 2.8±0.3 6.0±0.0 704 UK 1.3±0.2 2.4±0.3 1.0±0.0 1.1±0.1 1.0±0.0 4.9±0.2 1.0±0.0 6.0±0.0 539 2 PT ≠ isolates brassicae ± S.E. for Hyaloperonospora ˠ 1.1±0.1 1.8±0.2 1.3±0.1 1.0±0.0 1.0±0.0 1.2±0.1 2.4±0.3 PT 6.0±0.0 006 IP 1 lines and thirteen oleracea combinations of eight Brassica cotyledon inoculations for different Mean interaction phenotypes in seedlings following EBH527 PCA 20.14 PC11 PC10 EBH525 EBH502 EBH508 Senna Host line Pathotype F F E E D C B A Class replicates) for each combination of host line and pathogen isolate. Combinations with IP<2.5 are incompatible; 2.5 ≤ IP < 4.5 moderately incompatible/ weakly compatible, and IP ≥ 4.5 are compatible or the UK, respectively either in Portugal collected from an experiment ≠ PT or UK indicates whether the data for each isolate was Interaction phenotypes (IP) were assessed using seven scores (Table ˠ Interaction phenotypes (IP) were assessed using seven Table 5.8 Table ) (Coelho et al. 2012 ( H. parasitica brassicae European isolates of Hyaloperonospora 5.2 Pathogenic Variability 137

Table 5.9 Origin of the Hyaloperonospora brassicae (H. parasitica) isolates collected from field samples on different crop types of Brassica oleracea (Coelho et al. 2012) Geographic origin country Country where it Isolates Original host Descriptiona (region) Yearb was tested Hp702 Calabrese SSI UK (Cambridge shire) 2001 UK Hp704 Unknown SSI UK (Lancashire) 2001 UK Hp710 Cauliflower SSI UK (Lincolnshire) 2001 UK Hp801 Cauliflower Field isolate UK (Somerset) 2007 UK Hp806 Unknown Field isolate UK(Lancashire) 2008 UK Hp717 Calabrese SSI UK (Lincolnshire) 2001 UK and Portugal Hp530 Mixed SSI Italy (Sicilia)c 1998 Portugal Hp533 Portuguese SSI Portugal (Odemira) 2006 Portugal cabbage Hp535 Portuguese SSI Portugal (Acores) 2006 Portugal cabbage Hp539 Broccoli SSI Portugal (Batalha) 2007 Portugal Hp541 Broccoli Field isolate Portugal (Batalha) 2008 Portugal FP06 Cauliflower SSI France (Yonne)d 2002 Portugal Hp006 B. oleracea SSI UK (Kirton) 1983 Portugal aSingle-sporangiospore-derived isolate bDate of entry in the UK or Portuguese collection cObtained from Dr. F. Branca (University di Catania, Italy) dObtained from Dr. D. Silve (Prince de Bretagne Biotechnie, France)

Table 5.10 Interaction-phenotype scores used to evaluate the response of Brassica oleracea cotyledons and relative amount of sporulation following inoculation with Hyaloperonospora brassicae (H. parasitica) (Coelho et al. 2012) IP scores Interaction phenotype 0 No host, no sporulation 1 Light host necrosis localized on the upper cotyledon surface, no sporulation 2 Heavy host necrosis localized on the upper cotyledon surface, no sporulation 3 Host necrosis localized on the upper cotyledon surface, weak sporulation (five conidiophores) localized on the lower cotyledon surface confined to the point of infection 4 Host necrosis localized on the upper cotyledon surface, heavy sporulation localized on the lower cotyledon surface confined to the point of infection 5 No necrosis on the upper surface, sparse to moderate sporulation dispersed over the whole cotyledon surface 6 No necrosis on the upper surface, abundant, and dense sporulation dispersed over the whole cotyledon surface 138 5 Physiologic Specialization (Pathogenic Variability)

LANES 2 - 13 14 - 17

3.0

2.0 1.6

1.0

0.5

Plate 5.1 Random amplified polymorphic DNA (RAPD) from 16 isolates of crucifer downy mildew (Hyaloperonospora parasitica), lanes 2–13 are isolates from oilseed rape Brassica napus, and lanes 14–17 are isolates from cauliflower B. oleracea (Tham et al. 1994)

5.2.1 DNA Fingerprinting of H. parasitica

Different pathotypes can be distinguished on the basis of host specificity (Sherriff and Lucas 1990), but such tests are time-consuming and may not reveal the full extent of variations present. Other phenotypic markers such as sexual compatibility type (Sherriff and Lucas 1989a, b) or fungicide sensitivity (Crute et al. 1985) can be assessed, but they provide limited information for epidemiological studies or genetic analysis. There is need, therefore, for alternative molecular markers (Michelmore and Hulbert 1987) to further define variations in the pathogen. Randomly amplified polymorphic DNAs (RAPDs) have been proposed as genetic markers that overcome many of the technical limitations of restriction fragment length polymorphism (RFLP) analysis (Williams et al. 1990; Welsh and McClelland 1990). RAPD scan can be used in the constructions of linkage maps (Williams et al. 1990; Reiter et al. 1992), in the identification of strains, and varieties by genomic fingerprinting (Welsh and McClelland 1990; Goodwin and Annis 1991; Hu and Quiros 1991; Schafer and Wostemeyer 1992; Kresovich et al. 1992; Klein-Lankhorst et al. 1991; Koller et al. 1993; Stiles et al. 1993) and following the cloning of amplified fragments may also serve as conventional RFLP probes. In the 5.2 Pathogenic Variability 139 study carried out by Tham et al. (1994), the potential use of RAPDs as a source of genetic markers in H. parasitica was evaluated. Several isolates from different host species were compared to determine whether reproducible banding patterns correlated with host specificity. As H. parasitica can only be cultured on living plant tissues, particular attention was paid to possible artefacts arising through contamination of sample DNA from plant or other microbial sources. Random amplified polymorphic DNA (RAPD) fingerprints were generated from a range of H. parasitica isolates from different Brassica species (Table 5.11). Reaction conditions, in particular DNA template, primer, and Mg2+ concentrations, were optimized to ensure that amplifications were reproducible. Possible artefacts arising through host plant DNA were assessed by including such DNA in control reactions. Confirmation that diagnostic RAPD bands were generated from fungal DNA was also obtained by southern hybridization of a RAPD band to genomic fungal DNA. By screening 20 decamer primers, 2 were found to detect sufficient genetic variation to allow complete differentiation between pathotypes. These results illustrate the potential value of RAPDs for detecting polymorphisms between isolates of a non-culturable plant pathogenic fungus. Rehmany et al. (2000)

Table 5.11 Isolates used for RAPD analysis (Tham et al. 1994) Isolate Source Year of Geographic Mating Response to code host isolation origin typea metalaxylb P001 B. napus 1982 Leicestershire Homothallic S P1072 B. napus 1992 Hertfordshire Homothallic nt P1118 B. napus 1993 Leicestershire Homothallic nt P1119 B. napus 1993 Leicestershire Homothallic nt P1130 B. napus 1993 Nottinghamshire Homothallic nt P1121 B. napus 1993 Essex Homothallic nt P1122 B. napus 1993 Hertfordshire Homothallic nt P003c B. napus 1983 – P1 S P004c B. napus 1983 – P1 S P033c B. napus 1983 – P2 S P1100c B. napus 1992 – P2 nt P1105c B. napus 1992 – P1 nt P005 B. 1977 Tyne and wear P1 S oleracea P006 B. 1983 Lincolnshire P2 R oleracea P1091 B. 1991 Lincolnshire nt nt oleracea P1092 B. 1991 Lincolnshire nt nt oleracea aMating defined as in Sherriff and Lucas (1989b) bS, sensitive; R, resistant; nt, not tested cSingle-spore oospores progeny from self of isolates P001, P003, and P033 segregated for virulence to the oilseed rape (B. napus var. oleiferas) cv. Cresor 140 5 Physiologic Specialization (Pathogenic Variability)

compared downy mildew isolates from A. thaliana and B. oleracea using AFLP and ITS analysis dividing the isolates into five groups that correlated with taxonomic species and in most of the cases with host origin.

5.2.2 Identification of Host Differentials and Nomenclature of Pathotypes

Physiologic specialization/pathogenic variability have long been recorded in H. parasitica infecting cruciferous plants all over the world (Gardner 1920; Kobel 1921; Thung 1926b; Gaumann 1926; Hiura and Kanegae 1934; Wang 1944). However, nomenclature and designation of H. parasitica pathotypes/races were initiated by Natti et al. (1967) based on differential interaction of isolates on cruciferous hosts, viz. broccoli and B. oleracea. Natti et al. (1967) identified and designated race 1 and race 2 of H. parasitica showing differential reactions on these two hosts. Later, several reports have come from different countries on the host specificity of H. parasitica isolates collected from numerous cruciferous hosts (Felton and Walker 1946; Natti 1958; Dzhanuzakov 1963; Chang et al. 1964; Semb 1969; Dickinson and Greenhalgh 1977; Morris and Knox-Davies 1980; Hill et al. 1988; Sherriff and Lucas 1990; Moss et al. 1991; Mehta and Saharan 1994; Lucas et al. 1994; Tham et al. 1994; Silve et al. 1996; Masheva et al. 1996b; Yoshida and Ohguchi 1998). The methodology and host species/varieties/lines used in these studies were not comparable and reproducible in the absence of non-maintenance of isolates and the use of international standard set of host differential. It is difficult to compare the results and determine whether distinct pathotypes are the same or different. Mehta and Saharan (1994) designated distinct pathotypes, one from cauliflower and other from oilseed Brassica on the basis of differential interaction of 9 isolates on a set of 17 host cruciferous host differential set (Table 5.12). Nashaat and Awasthi (1995) identified five groups of B. juncea (Table 5.5) accessions with differential resistance to UK isolates R1 and P003 derived from oilseed rape (B. napus ssp. oleifera) and Indian isolates IP01 and IP02 derived from mustard (B. juncea). Coelho et al. (2012) assembled set of genetically uniform lines of B. oleracea (doubled haploid or self- pollinated inbred lines) to use as host differential set for the characterization of pathotypic variation of H. parasitica (H. brassicae) within a European sample of the pathogen collected from various crop types of B. oleracea. Six pathotypes (HP1–HP6) were distinguished indicating a potential use of the host differentials for monitoring frequencies in H. parasitica populations (Table 5.8). A naming system for the isolates of H. arabidopsidis was introduced by Dangl et al. (1992), Holub et al. (1994), and Slusarenko and Schlaich (2003) on the basis of geographical location and ecotypes infected (AHCO, ASWA, CALA, EDCO, EMWA, etc.). For detailed description, see chapter 1, section 1.11.6. However, still there is a need to standardize a set of host differentials (isogenic lines) and a system of nomenclature and designation of pathotypes of H. parasitica at international level. It is 5.2 Pathogenic Variability 141

Table 5.12 Identification of pathotypes of Hyaloperonospora parasitica and H. arabidopsidis International Designated pathotype Country primary host References Isolate 1 USA Turnip Gardner (1920) Biological forms 2 Holland Cabbage Thung (1926b) Biological strains 3 Germany Crucifers Gaumann (1926) Biological forms Japan Crucifers Hiura and Kanegae (1934) Pathotypes 3 China Brassica, Wang (1944) Raphanus, Capsella Races USA Raphanus, B. Felton and Walker (1946), oleracea and Natti (1958) Races South B. oleracea, Morris and Knox-Davies Africa Raphanus (1980) Special forms Leningrad Crucifers Dzhanuzakov (1963) Three forms China Crucifers Chang et al. (1964) Races Norway Cabbage, radish Semb (1969) Races 1 and 2 USA (New Broccoli, B. Natti et al. (1967) York) oleracea Isolates India Crucifers Bains and Jhooty (1983) Pathotypes 2 (9 isolates) India Brassica, Mehta and Saharan (1994) cauliflower Isolates P007, P008, P013, P014 UK B. rapa (B. Moss et al. (1991), and campestris) Silve et al. (1996) Isolates UK B. napus Nashaat and Rawlinson (1994) IP01 and IP02 UK B. juncea Nashaat and Awasthi (1995) R1 and P003 UK B. napus Nashaat and Awasthi (1995) 7 isolates UK, Cauliflower (4) Silve et al. (1996) France, Rape (2), mustard India (1) 11 isolates USA Crucifer rapid Hill et al. (1988) cycling population 2 isolates Bulgaria Plovdiv, Goma Masheva et al. (1996a, b) Oriahovitsa 4 isolates Japan Japanese radish, Yoshida and Ohguchi rape, broccoli, (1998) shepherd’s purse 21 isolates UK Brassica oleracea Silve et al. (1996) 2 isolates UK B. rapa Silve et al. (1996) 6 pathotypes HP1–HP6 Portugal, B. oleracea Coelho et al. (2012) UK 5 groups of isolates through DNA UK A. thaliana, B. Tham et al. (1994) fingerprinting (P001–P005) oleracea (continued) 142 5 Physiologic Specialization (Pathogenic Variability)

Table 5.12 (continued) International Designated pathotype Country primary host References AHCO, ASWA, CALA 2, CALA, Germany, Arabidopsis Dangl et al. (1992), Holub CAND 3, EDCO, EMCO, EMOY, UK accessions Col-0, et al. (1994), Slusarenko EMOY 2, EMWA 1, GOCO, Ler-0, Oy-0, and Schlaich (2003), and GOWA, HIKS1, HIKS, HIND, Nd-1, Ws-0, Schlaich and Slusarenko HIND 2, HIND 4, MADI, MAKS, Wei-0 (2009) MAKS9, NOCO, NOCO 2, NOKS 1, WAND, WELA, WELA1, WELA 3 essential to know whether pathotypes of H. parasitica reported so far from different countries are the same or different (Table 5.12). It will be very helpful to identify resistant genes and their combinations to breed resistant cvs. and their durability extent under different agroecological situations.

5.3 Heterothallism and Homothallism

Induction of the sexual process in several downy mildew species requires the presence of two strains of opposite mating type. Such heterothallic behaviour has been reported for H. parasitica (De Bruyn 1937; McMeekin 1960; Kluczewski and Lucas 1983; Sherriff and Lucas 1989b; Sequeira and Monteiro 1996). Homothallic forms of H. parasitica have also been observed (De Bruyn 1937; Sherriff and Lucas 1989b; Sequeira and Monteiro 1996). No relationship has been observed between geographic origin and mating types. Both heterothallic and homothallic isolates have been isolated from the same fields (Sequeira and Monteiro 1996). The two forms of sexual reproduction are very important for the maintenance and evolution of the fungal strains and epidemiology of the disease. Two mating type heterothallic isolates of B. oleracea and B. campestris designated as P1 and P2 have been identified. Isolates from oilseed rape B. napus have been found to be uniformly homothallic and remained self-fertile even after months of laboratory subculture (Sherriff and Lucas 1989b). In a cytogenetic study of heterothallic and homothallic isolates of H. parasitica at metaphase 1 of meiosis, a ring of four chromosomes is found (Sherriff and Lucas 1989a). This ring is interpreted as a reciprocal translocation complex between chromosomes carrying the mating-type alleles. In homothallic isolates a fifth chromosome is associated with the ring of four. The self-fertility of these isolates may therefore be due to the presence of a third mating-type allele on the fifth chromosome, a condition known as secondary homothallism. The determination of sexual compatibility type (SCT) of an unknown isolate can be achieved by mixing conidia in a 1:1 ratio with isolates of known SCT and inoculating to a common compatible host. In heterothallic isolates, oospores may form in combination with isolates of opposite SCT. Isoenzyme markers are particularly important in discrimi- nating between self and true hybrid progeny (Moss et al. 1988). References 143

5.4 Hybridization of Hyaloperonospora Isolates

Moss et al. (1994) produced viable oospore of H. parasitica under laboratory conditions and described recovered isolates (referred as sexual progeny) from these oospore populations. Oospores were produced when isolates of opposite sexual compatibility type, specialized to the same or different Brassica species, were grown together in seedling cotyledons of a host line capable of supporting growth of both isolates. Recovery of sexual progeny from oospore populations produced from two out of four pairing between isolates, is specialized in the same host species (homologous pairings) and proved to be relatively easy. On the basis of their characterization with respect to virulence, response to phenylamide fungicides, sexual compatibility type, and isoenzyme polymorphisms, there was evidence that the sexual progeny from these homologous pairing could be of hybrid origin. For the first time in a member of the Peronosporaceae, it provided possibility to recover and successfully characterize a new sexual progeny from pairing between isolates specialized to different host species (heterologous pairing). However, the majority of such isolates sporulated weakly and, as consequence, proved difficult to maintain and were lost. Nevertheless, it is concluded that some evidence for the hybrid nature of progeny from heterologous pairing was obtained.

References

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Goodwin PH, Annis SL (1991) Rapid identification of genetic variation and pathotype of Leptosphaeria maculans by random amplified polymorphic DNA assay. Appl Environ Microbiol 57(2482):2486 Hill CB, Crute IR, Sherrife C, Williams PH (1988) Specificity of Albugo candida and Peronospora parasitica pathotypes towards rapid-cycling Brassicas. Cruciferae NewsLett 13:112–113 Hiura M, Kanegae H (1934) Studies on the downy mildews of cruciferous vegetables in Japan. Trans Sapp Nat His Soc 13:125–133 Holub EB, Beynon JL, Crute IR (1994) Phenotypic and genotypic characterization of interactions between isolates of Peronospora parasitica and accessions of Arabidopsis thaliana. Mol Plant- Microbe Interact 7:223–239 Hu J, Quiros CF (1991) Identification of broccoli and cauliflower cultivars with RAPD markers. Plant Cell Rep 10:505–511 Klein-Lankhorst RM, Vermunt A, Weide R, Liharska T, ZabeI P (1991) Isolation of molecular markers for tomato (L. esculentum) using random amplified polymorphic DNA (RAPD). Theor Appl Genet 83:108–114 Kluczewski SM, Lucas JA (1983) Host infection and oospore formation by Peronospora parasitica in agricultural and horticultural Brassica species. Trans Br Mycol Soc 81:591–596 Kobel F (1921) The problem of host selection by parasitic fungi. Naturwissen schaftliche Wochenschrift 36:113–118 Koller B, Lehmann A, McDermott JM, Gessler C (1993) Identification of apple cultivars using RAPD markers. Theor Appl Genet 85:901–904 Kresovich S, Williams JGK, McFerson JR, Routman EJ, Schaal BA (1992) Characterization of genetic identities and relationships of Brassica oIeracea L. via a random amplified polymorphic DNA assay. Theor Appl Genet 85:190–196 Leckie D, Astley D, Crute IR, Ellis PR, Pink DAC, Boukema I, Monteiro AA, Dias S (1996) The location and exploitation of genes for pest and disease resistance in European gene bank collections of horticultural Brassicas. Acta Hortic 407:95–101 Lucas JA, Crute IR, Sherriff C, Gordon PL (1988) The identification of a gene for race-specific resistance to Peronospora parasitica (downy mildew) in Brassica napus var. oleifera (oilseed rape). Plant Pathol 37:538–545 Lucas JA, Hayter JBR, Crute IR (1994) The downy mildews: host specificity and pathogenesis. In: Kohmoto K (ed) Pathogenesis and host specificity in plant diseases, vol Vol. 2. Elsevier Science Ltd, Oxford Masheva S, Antonova G, Bahariev D (1996a) Seasonal variability in the Peronospora parasitica (Pers.) Fr. population. Cruciferae NewsLett 18:120 Masheva S, Antonova G, Bahariev D (1996b) Pathogenicity of two isolates Peronospora parasitica with different district origin. Cruciferae NewsLett 18:118 McMeekin D (1960) The role of the oospores of Peronospora parasitica from cabbage and radish. Phytopathology 50:93–97 Mehta N, Saharan GS (1994) Morphological and pathological variations in Peronospora parasitica infecting Brassica species. Indian Phytopath 47:153–158 Michelmore RW, Hulbert SH (1987) Molecular markers for genetic analysis of phytopathogenic fungi. Annu Rev Phytopathol 25:383–404 Morris MJ, Knox-Davies PS (1980) Raphanus raphanistrum as a weed host of pathogens of cultivated cruciferae in the Western Cape province of South Africa. Phytophylactica 12:53–55 Moss NA, Crute IR, Lucas JA, Gordon PL (1988) Requirements for analysis of host species specificity in Peronospora parasitica (downy mildew). Cruciferae NewsLett 13:114–116 Moss NA, Lucas JA, Crute IR (1991) Evidence for differential response to isolates of Peronospora parasitica (downy mildew) in Brassica rapa. Test Agro Chem. cv. 12. Ann Appl Biol 118:96–97 Moss NA, Crute IR, Lucas JA (1994) Laboratory production of oospores of Peronospora parasitica (crucifer downy mildew) and the recovery and characterization of sexual progeny from crosses between isolates with different host specificity. Plant Pathol 43:713–725 Nashaat NI, Awasthi RP (1995) Evidence for differential resistance to Peronospora parasitica (downy mildew) in accessions of Brassica juncea (mustard) at the cotyledon stage. J Phytopathol 143:157–159 References 145

Nashaat NI, Rawlinson CJ (1994) The response of oilseed rape (Brassica napus ssp. oleifera) accessions with different glucosinolate and erucic acid contents to four isolates of Peronospora parasitica (downy mildew) and the identification of new sources of resistance. Plant Pathol 43:278–285 Natti JJ (1958) Resistance of broccoli and other crucifers to downy mildew. Plant Dis Rep 42:656–662 Natti JJ, Dickson MH, Atkin JD (1967) Resistance of Brassica oleracea varieties to downy mildew. Phytopathology 57:144–147 Rehmany AP, Lynn JR, Tor M, Holub EB, Beynon JL (2000) A comparison of Peronospora parasitica (downy mildew) isolates from Arabidopsis thaliana and Brassica oleracea using amplified fragment length polymorphism and internal transcribed spacer 1 sequence analyses. Fungal Genet Biol 30:95–103 Reiter RS, Williams JGK, Feldmann KA, Rafalski JA, Tingey SV, Scolnik PA (1992) Global and local genome mapping in Arabidopsis thaliana by using recombinant inbred lines and random amplified polymorphic DNAs. Proc Natl Acad Sci U S A 89:1477–1481 Schafer C, Wostemeyer J (1992) Random primer dependent PCR differentiates aggressive from non-aggressive isolates of the oilseed rape pathogen Phoma lingam (Leptosphaeria maculans). J Phytopathol 136:124–136 Schlaich NL, Slusarenko A (2009) Downy mildew of Arabidopsis caused by Hyaloperonospora arabidopsidis (formerly Hyaloperonospora parasitica). In: Kurt L, Kamoun S (eds) Oomycete genetics and genomics: diversity, interactions and research tools, Chapter 13. Wiley, Hoboken, pp 263–285 Semb L (1969) Cabbage downy mildew. Jord Avling 12:32–35 Sequeira P, Monteiro A (1996) Heterothallism and homothallism in Portuguese isolates of Peronospora parasitica (Pers. ex Fr.) Fr. Curciferae NewsLett 18:126–127 Sherriff C, Lucas JA (1989a) Cytogenetic study of heterothallic and homothallic isolates of Peronospora parasitica. Mycol Res 92:302–310 Sherriff C, Lucas JA (1989b) Heterothallism and homothallism in Peronospora parasitica. Mycol Res 92:311–316 Sherriff C, Lucas JA (1990) The host range of isolates of downy mildew, Peronospora parasitica from Brassica crop species. Plant Pathol 39:77–91 Silve D, Nashaat NI, Tirilly Y (1996) Differential responses of Brassica oleracea and B. rapa accessions to seven isolates of Peronospora parasitica at the cotyledon stage. Plant Dis 80:142–144 Slusarenko AJ, Schlaich NL (2003) Downy mildew of Arabidopsis thaliana caused by Hyaloperonospora parasitica (formerly Peronospora parasitica). Mol Plant Pathol 4:159–170 Stiles JI, Lemme C, Sondur S, Morshidi MB, Manshardt R (1993) Using randomly amplified polymorphic DNA for evaluating genetic relationships among papaya cultivars. Theor Appl Genet 85:697–701 Tham FY, Lucas JA, Wilson ZA (1994) DNA fingerprinting ofPeronospora parasitica, a biotrophic fungal pathogen of crucifers. Theor Appl Genet 88:490–496 Thung TH (1926a) Observations on Peronospora parasitica on cabbage. Tijdschr Over Plantenziekten 32:161–179 Thung TH (1926b) Peronospora parasitica (Pers.) De by attacking cabbage heads. Phytopathology 16:365–366 Uknes S, Mauch-Mani B, Moyer M, Potter S, Williams S, Dincher S, Chandler D, Slusarenko A, Ward E, Ryals J (1992) Acquired resistance in Arabidopsis. Plant Cell 4:645–656 Wang CM (1944) Physiological specialization in Peronospora parasitica and reaction of hosts. Chin J Sci Agri 1:249–257 Welsh J, McClelland M (1990) Fingerprinting genomes using PCR with arbitrary primers. Nucleic Acids Res 18:7213–7218 Williams JGK, Kubelik AR, Livak KJ, Rafalski JA, Tingey SV (1990) DNA polymorphisms amplified by arbitrary primers are useful as genetic markers. Nucleic Acids Res 18:6531–6535 Yoshida K, Ohguchi T (1998) Suppression of haustorium formation of Peronospora parasitica in heat-treated Japanese radish [Raphanus sativus] root tissues. Ann Phytopathol Soc Japan 64:307–314 Chapter 6 Perpetuation and Survival of Pathogen

6.1 Introduction

Oospores formed in malformed and senesced host tissues constitute an important means of survival of H. parasitica over periods of unfavourable conditions (Gaumann 1926; Kolte 1985; Saharan et al. 1997, 2005). It is also known to survive through mycelium and conidia (Jang and Safeeula 1990b; Krober 1970; McMeekin 1969; Vishunavat and Kolte 1993) for a short period of time. It is also known as oversummering and overwintering of the pathogen under unfavourable environmental conditions.

6.2 Mycelium

The presence of H. parasitica mycelium in the seed coat of Chinese cabbage has been recorded by Chang et al. (1963). According to Jang and Safeeulla (1990c), the presence of mycelium in the pericarp and embryo of radish seeds varies from 0.1% to 12.5% (Tables 6.1 and 6.2). The coenocytic branched mycelium is clearly visible in the intercellular space of the pericarp. In the embryonic tissues, the mycelium is comparatively thin. The percentage of seeds with viable mycelium is directly correlated with the percentage of embryo infection.

6.3 Conidia

Conidia of H. parasitica on cabbage survive longer under cool, dry conditions (Krober 1981). Relative humidity is more important than temperature. In the field conidia can survive on detached leaves of kohlrabi for 10 days during warm days.

© Springer Nature Singapore Pte Ltd. 2017 147 G. S. Saharan et al., Downy Mildew Disease of Crucifers: Biology, Ecology and Disease Management, https://doi.org/10.1007/978-981-10-7500-1_6 148 6 Perpetuation and Survival of Pathogen

Table 6.1 Percentage of seed infection by Hyaloperonospora parasitica in Raphanus sativus Seed showing infection (%) Cultivar Place of collection Pericarp Endosperm Embryo Japanese Mysore Seed Multiplication Farm 12.8 0 12.5 white Arka Nishant Indian Council of Agricultural Research Station, 0.5 0 0.5 Bangalore Pusa Desi Bangalore Seed Health Testing Station 0.2 0 0.3 Pusa Reshmi Bangalore Seed Health Testing Station 0 0 0.1 Jang and Safeeulla (1990c)

Table 6.2 Percentage of seedling infection by Hyaloperonospora parasitica and seed transmission in Raphanus sativus Seed Infection (%) Cultivar Seedling infection (%) Pericarp Embryo Japanese white 14.0 13.5 12.8 Arka Nishant 1.5 0.5 0.4 Pusa Desi 1.0 0.1 0.2 Pusa Reshmi 1.0 0.0 0.1 Jang and Safeeulla (1990c)

When buried in dry soil, conidia can survive for 110 days. The survival period is greatly reduced to a maximum of 22 days if the soil is moist. Marked reduction in survival has also been observed after storage in both dry and moist soils during the summer. The conidial viability is longest, up to 130 days, when the spores are stored in air-dried soil at a constant temperature of 5 °C (Krober 1970). Conidia kept at −25 °C and relatively dry on leaf discs (air dried at 20 °C) maintain a relatively high rate of germination after 1 year or longer.

6.4 Oospores

Oospores formation is abundant in the infected tissues of all crucifers, and they form primary source of survival for the pathogen (LeBeau 1945; McMeekin 1960; Chang et al. 1963; Kolte 1985). In radish and rapeseed-mustard, there is abundant production of oospores in infected leaf tissues, on the seed surface, pericarp, and embryo of seeds (Jang and Safeeulla 1990c; Vishunavat and Kolte 1993). However, in rapeseed and mustard, seed transmission is low and may be nonsystemic, ranging from 0.4% to 0.9% in the seedlings grown from infected seeds (Vishunavat and Kolte 1993). In radish seed transmission to the extent of 14% was observed by Jang and Safeeulla (1990d). 6.7 Conidial Discharge 149

6.5 Seed Infection

Hyaloperonospora parasitica infection levels in the cabbage seed coat and endosperm varied between 15% and 70%, while in the embryo percentage infection varied between 5% and 65%. Germinating oospores, intact oospores, mycelia, and conidia of H. parasitica were observed in the severely infected macerated host tissues (Badul and Achar 1998). A tissue culture technique developed by Achar (1995) is quicker and more reliable than the seedling symptom test for determining the viability of mycelium. Vishunavat and Kolte (1993) found that oospores of H. parasitica occurred on the seed surface and in the hypodermis of seed coat tissue in sarson, toria, and Indian mustard. The fungus was seed transmitted in a nonsystemic manner in sarson and toria at rates of 0.9 and 0.4%, respectively, but not in Indian mustard. Infected seedlings showed downy growth of the fungus on the lower surface of cotyledonary leaves and up to two or three successive true leaves. Further growth of such seedlings appeared normal with no hypertrophy of the inflorescence, and seeds collected from such plants showed no infection.

6.6 Axenic Culture

Hyaloperonospora parasitica hyphae grow on water agar from the infected tissues and form haustorium-like structures (Ohguchi and Asada 1989). The growth is greater on the modified Knop medium with many haustorium-like structures and conidiophores being formed on this medium. If cod liver oil or minerals are added to this medium, then branched hyphae are formed. The fungus does not grow on Japanese radish root homogenate medium but grows well on the dialysed homogenate medium. The decoction of residuum of the root homogenate and the sap in the intercellular spaces of the root tissues also stimulate the growth of the fungus. In the decoction medium, the growth of the hypha is vigorous, and the formation of conidiophores is stimulated. In the sap medium, the formation of a haustorium-like structure is promoted (Ohguchi and Asada 1989).

6.7 Conidial Discharge

The maximum conidial discharge of H. parasitica from kohlrabi leaves is between 5 and 6 a.m. (258 conidia/cm2) (Lin and Liang 1974). The conidial discharge decreases greatly from 12 noon to 8 p.m. If infected leaves are covered with plastic bag during the night, then the production and discharge of conidia decreases drasti- cally, and the disease index is half that of uncovered seedlings. The discharge of 150 6 Perpetuation and Survival of Pathogen conidia from diseased leaves of Chinese cabbage shows a periodic cycle each day (Fig. 6.1) (Lin 1981). Conidial release increases steadily after 2 a.m. each day and reaches a peak around 6–8 a.m. Conidial discharge decreases rapidly after 8 a.m. Few conidia can be detected from noon to 10 p.m. The discharge of conidia is favoured by temperatures below 18 °C and RH above 75% (Fig. 6.1). If the conidia are ready to be released from (Fig. 6.2) conidiophores but the RH suddenly decreases, then the branched conidiophores become dry, and the twirling movement of the drying conidiophores may flick the spores in to the air and discharge abundant conidia from the diseased leaf. According to Shao et al. (1990), conidia release during favourable temperature and RH conditions is three times higher in the morning than in the afternoon. Dispersal of conidia of H. parasitica on Lepidium virginicum begins with incipi- ent desiccation and concludes with hygroscopic distortion of the aerial fructifica- tions (Pinckard 1942). Several complete twists occur in the portion of tall conidiophores extending up to the first branch, with a lesser number between each successively shorter branch. With the progress of drying, a twisting and binding motion is imparted to the sterigma-like structure on which the conidia are borne. If the process of desiccation stops, the twisting motion also stops. However, if humidity increases, the rotation reverses itself. Under conditions of delicate moisture bal- ance, the breath of an observer is sufficient to induce the above-mentioned movements. The outcome of the movement is the release of mature conidia. By slowly decreasing the vapour pressure, a point is reached when abscission occur, and the conidia are forcibly released with the stimulus for the requisite energy being

300 a 2 200

100 No conidio / cm

0 80 30 b 75 70 20 65 60 Temperature (°C ) Relative humidily (% ) 10 55 04 81216200 48121620 04812 16 20 I Day II Day III Day Time of day (hr)

Fig. 6.1 (a) Pattern of Hyaloperonospora parasitica conidia discharge from infested Chinese cabbage plants and (b) temperature and humidity on 3 fine days in November 1978 (Lin 1981) 6.8 Conidial Germination 151

Fig. 6.2 The mechanism of Hyaloperonospora parasitica conidia discharge. (a) Conidiophores in damp air with attached conidia; (b) and (c) changes in conidiophores on exposure to dry air and (d) recovery on return to damp condition (Lin 1981) derived from differential stresses set up within the sterigmata. The mechanical action of wind, and rain, during periods of atmospheric saturation does not appear to contribute significantly to dispersal of conidia.

6.8 Conidial Germination

For germination of conidia collected from Chinese cabbage, 8–20 °C is favourable with an optimum range of 12–16 °C (Fig. 6.3) (Lin 1981). Germ tubes usually grow normally and extensively at these temperatures. The germination rate of conidia is low, and the germ tubes show limited and malformed growth at temperatures below 8 °C and above 20 °C. Conidia fail to germinate at extreme temperatures even after a long period of incubation. Conidial germination usually increases after treatment with hot water of up to 42 °C (Fig. 6.4). Germination of conidia from Chinese cabbage (B. pekinensis) was optimum at 15–20 °C and was stimulated by light (Shao et al. 1990). Maximum conidial germination of H. parasitica from B. juncea was 78–92% at 20 °C temperature (Singh 1997). However, Achar (1998) from B. oleracea obtained 80–98% and 70–80% conidial germination between 15 and 25 °C at 100% relative humidity after 12 and 6 h incubation period, respectively; optimum temperature for germ tube growth was 20 °C. According to Paul et al. (1998), optimal conidial germination from B. napus required temperature of 5–15 °C and relative humidity of 90–98%. 152 6 Perpetuation and Survival of Pathogen

Fig. 6.3 The effect of 40 temperature and relative 95 humidity on the 36 germination of conidia of Hyaloperonospora 81 parasitica (Lin 1981) 32

28 Relative Humidity (%)

24 55 20

16 Temperature (°C) 32 12

0 0102030405060 Germination (%)

This was also the optimal infection temperature, whereas sporulation only occurred at a relative humidity of 98%. Under Germany conditions, the main occurrence of conidia in the air was found at 09.00 and 12.00 h, whereas the lowest incidence of conidia in the air was recorded between 21.00 and 06.00 h. Conidial germination and germ tube elongation of H. parasitica were promoted by Ca (NO3) (a compound of Knop’s solution), homogenate of Capsella bursa- pastoris leaves, and dialysed homogenate. Five of the 6 fractions of the dialysed homogenate separated on a Sephadex G − 100 column were also affective. The effect was increased by addition of MgSO4 to the dialysed homogenate, but not by Ca (NO3)2 or NH4NO3 (Guo and Ohguchi 1996). H. parasitica sporulate on intact cabbage seedlings when incubated at 13 °C or 18 °C in the presence of free water or to atmospheric water potentials (Ψ) of or −30 ± 10 bars (Table 6.3) (Hartman et al. 1983). The pathogen fails to sporulate at these temperatures when the atmospheric Ψ is −60, −90, or −120 bars. More conidia are produced at 13 °C (1466–2265 conidia/45 mm2 cotyledons) than at 18 °C (821–1042 conidia/45 mm2cotyledon). Conidia germinate in the presence of free water but do not germinate when exposed to atmospheric Ψ of 0, −30, −60, or 6.8 Conidial Germination 153

60 a

50 15 minutes

40 30 minutes

20 45 minutes Conidia Germination (%)

100 b

Kin-Po 80

San-Fon

40 Seed Germination (%) Cheng-Pao 20

26 30 34 38 42 46 50 54 58 60 Water Temperature (°C)

Fig. 6.4 Effect of hot water treatment on the germination of (a) conidia of Hyaloperonospora parasitica and (b) seeds of three Chinese cabbage cultivars. Conidia were held at each temperature for 15, 30, and 45 min, whereas seeds were held for 30 min only (Lin 1981) 154 6 Perpetuation and Survival of Pathogen

Table 6.3 An analysis of sporulation of Hyaloperonospora parasitica on cabbage cotyledons at two temperatures and in free water or at atmospheric water potentials of or −30 bars Numbers of sporangia/45 mm2 cotyledona and integers b assigned for contrasts of these numbers in the following treatments c 13 °C 13 °C 13 °C 18 °C 18 °C 18 °C FW Ψ = 0 Ψ = 30 FW Ψ = 0 Ψ = 30 Contrasts numbers 1978e 1466 2265 1042 821 855 F Testsd 1 −1 −1 −1 1 1 1 P = 0.01 2 0 −1 0 1 0 0 NS 3 0 0 0 1 −1 0 NS 4 0 −1 1 0 0 0 P = 0.01 5 0 0 0 0 −1 1 NS 6 −1 1 0 0 0 0 P = 0.05 7 −1 0 1 0 0 0 NS Hartman et al. (1983) aNumbers of sporangia were contrasted using non-orthogonal coefficients, and the differences were assessed by an F test. F tests are approximate for non-orthogonal contrasts bTreatments assigned positive integers were contrasted with those assigned negative integers cTreatments are identified according to temperatures and free water (FW) or atmospheric water potentials (Ψ in − bars) at germination dSignificance level or non-significance (NS) for results of the F test eThis value is the summation of mean numbers of sporangia/mm2 for each of 45 pairs of cotyledons

−90 bars for 24 h. The level of atmospheric Ψ and the presence or absence of free water during sporulation exert preconditioning effects on the ability of conidia to germinate. Conidia collected from B. campestris (B. rapa) (toria, brown sarson, yellow sarson) and B. oleracea (cauliflower) leaves germinate after 1.5 h at 18 °C, whereas conidia derived from B. juncea germinate after 1 h (Mehta and Saharan, 1994). Germination increases as the incubation period is increased. For instance, more than 80% conidia germinate after 4 h (Table 5.4).

6.9 Oospore Germination

The germination of oospores of H. parasitica infecting radish is dependent on temperature, light, pH of the medium, and age of oospores (Jang and Safeeulla 1990a). The optimum temperature for germination is 23 °C. Drying and chilling of oospores has no marked effect on germination. At a pH of 7.5, germination is 42% but at a pH of 4.5 only 1% oospores germinate. Oospore germination also increases with age (Jang and Safeeulla 1990a). References 155

References

Achar PN (1995) Tissue culture techniques to determine the viability of Peronospora parasitica in Brassica oleracea. J Phytopathol 143:647–649 Achar PN (1998) Effects of temperature on germination of Peronospora parasitica conidia and infection of Brassica oleracea. J Phytopathol 146:137–141 Badul J, Achar PN (1998) Screening of Brassica oleracea var. capitata cultivars against Peronospora parasitica. Seed Sci Tech 26:385–391 Chang IH, Xu RF, Chiu WF (1963) On the primary sources of infection of the downy mildew of Chinese cabbage caused by Peronospora parasitica (Pers.) Fr. and the limited systemic infection of seedlings. Acta Phytopath Sin 6:153–162 Gaumann E (1926) On the specialization of downy mildew (Peronospora brassicae Gaum.) on cabbage and related species. Landw Jahrb der Schewiz 40:463–468 Guo LX, Ohguchi T (1996) Effects of salts and components shepherd’s purse leaf on conidial germination of the downy mildew fungus. Mem Coll Agric Ehime Univ 40:279–287 Hartman H, Sutton JC, Proctor R (1983) Effects of atmospheric water potentials, free water and temperature on production and germination of sporangia in Peronospora parasitica. Can J Plant Pathol 5:70–74 Jang P, Safeeulla KM (1990a) Factors affecting germination of Peronospora parasitica in radish. Proc Indian Acad Sci Plant Sci 100:113–116 Jang P, Safeeulla KM (1990b) Production and viability of Peronospora parasitica in radish. Proc Indian Acad Sci Plant Sci 100:117–122 Jang P, Safeeulla KM (1990c) Seed-borne nature of Peronospora parasitica in Raphanus sativus. Proc Indian Acad Sci Plant Sci 100:225–258 Jang P, Safeeulla KM (1990d) Modes of entry, establishment and seed transmission of Peronospora parasitica in radish. Proc Indian Acad Sci Plant Sci 100:369–373 Kolte SJ (1985) Diseases of annual edible oilseed crops, vol. 2 Rapeseed mustard and sesame diseases. CRC Press, Boca Raton Krober H (1970) Longevity of conidia of several Peronospora species under different environmental conditions. Phytopathol Z 69:64–70 Krober H (1981) Germination and longevity of conidia of some Peronosporaceae kept under different conditions. Zeitsch Pflanzenk Pflanzenschutz 88:510–517 LeBeau FJ (1945) Systemic invasion of cabbage seedlings by the downy mildew fungus. Aust J Agric Res 71:453–463 Lin CY (1981) Studies on downy mildew of Chinese cabbage caused by Peronospora parasitica. In: Chinese cabbage. Proceedings of the First International Symposium Asian Vegetable Research Development Center, Shanhua, Taiwan, pp 105–112 Lin CY, Liang TC (1974) The discharge of conidia of Peronospora brassicae from kohlrabi and the polyethylene cover as a protection to seedlings against infection of this pathogen. Bull Agric Res 3:41–48 McMeekin D (1960) The role of the oospores of Peronospora parasitica from cabbage and radish. Phytopathology 50:93–97 McMeekin D (1969) Other hosts for Peronospora parasitica from cabbage and radish. Phytopathology 59:693–696 Mehta N, Saharan GS (1994) Morphological and pathological variations in Peronospora parasitica infecting Brassica species. Indian Phytopath 47:153–158 Ohguchi T, Asada Y (1989) Behavior of downy mildew fungus of Japanese radish on various media. II. Culture from the infected tissue. Mem Coll Agric Ehime Univ 33:201–209 Paul VH, Klodt-Bussmann E, Dapprich PD, Capelli C, Tewari JP (1998) Results on preservation, epidemiology, and aggressiveness of Peronospora parasitica and results with regard to the disease resistance of the pathogen on Brassica napus. Bull OILB/SROP 2:49–56 Pinckard JA (1942) The mechanism of spore dispersal in Peronospora tabacina and certain other downy mildew fungi. Phytopathology 32:505–511 156 6 Perpetuation and Survival of Pathogen

Saharan GS, Verma PR, Nashaat NI (1997) Monograph on downy mildew of crucifers. Technical Bulletin 1997–01. Research Branch Agriculture and Agri-Food Canada, 197p Saharan GS, Mehta Naresh, Sangwan MS (2005) Fungal diseases of rapeseed-mustard. In: Diseases of oilseed crops. Indus Publishing, New Delhi, pp 15–86 Shao XH, Chen QQ, Zhang FQ (1990) Research on relation between some ecological factors and Peronospora parasitica (Pers.) Fr. on Chinese cabbage. Acta Agric Shanghai 6:78–81 Singh HV (1997) Effect of temperature on sporangial germination of Peronospora parasitica and Albugo candida from Brassica juncea cv. Varuna. Plant Dis Res 12:192–194 Vishunavat K, Kolte SJ (1993) Brassica seed infection with Peronospora parasitica (Pers. ex. Fr.) Fr. and its transmission through seed. Indian J Mycol Plant Path 23:247–249 Chapter 7 Infection and Pathogenesis

7.1 Introduction

In crucifers, the downy mildew infection may be either general or local. In the former case, all or most of the leaves and inflorescence (which may be hypertrophied as a result of pre-infection with Albugo candida) may bear conidiophores. Although some parts (especially the stem) may show no external injury, microscopic examination shows that the mycelia are in the tissues (Butler 1918). Generalized infection is restricted to young tissues, and this is why seedlings show completely infected leaves. Localized infection also occurs in young tissues, especially those still in active division. In the hypertrophy caused by Albugo (Awasthi et al. 1997), the cells of the epidermis and cortex are dividing and may readily give entrance to Hyaloperonospora. Young inflorescence may wholly or partly be infected, while normal tissues of older stems and leaves below the initial site of infection may remain free.

7.2 The Process of Infection

The conidia of H. parasitica on surface of a susceptible host form germ tubes when environmental conditions are favourable from which appressorium develop (Preece et al. 1967). In cauliflower, appressoria are found at the junction of the anticlinal walls of the epidermal cells. The contents of the conidium pass into the appressorium from which an infection hypha develops (Chou 1970). Penetration is usually direct and only occasionally through a stoma (Shiraishi et al. 1975). It breaks a hole, 4–5 um in dia., through the cuticle, and after entering the host, the hypha expands to a diameter of 7–8 μm. The fungus grows initially in the region of the middle lamella between the anticlinal walls of the epidermal cells. Penetration between adjacent epidermal cells rather than via stomata had been earlier reported by Chu (1935). The

© Springer Nature Singapore Pte Ltd. 2017 157 G. S. Saharan et al., Downy Mildew Disease of Crucifers: Biology, Ecology and Disease Management, https://doi.org/10.1007/978-981-10-7500-1_7 158 7 Infection and Pathogenesis infection hypha continues its growth between the cells of the host tissues branching in all directions, and varying in diameter, and forms according to the size and shape of the intercellular spaces (Chou 1970). A single conidium of H. parasitica is sufficient to infect kohlrabi (Krober1969 ) and radish (Achar 1992). The disease intensity and rate of infection increase as the number of spores in the inoculum increases. Commensurately, more conidia are required to produce comparable responses in older plants which are less susceptible to infection than young plants (Krober 1969). Disease intensity increases with increasing inoculum concentrations up to 30,000 conidia/ml of water, but further increases in inoculum have no significant effect on the host (Achar 1992). The rate of spore germination and host penetration are markedly affected by temperature. At 15 °C, conidia germinate in 4–6 h, appressoria form in 12 h, and penetration occurs in 18–24 h (Chu 1935). Felton and Walker (1946), however, reported that on cabbage, germination of conidia and the subsequent penetration of the host take place most rapidly at 8–12 °C and 16 °C, respectively. Jonsson (1966) found that development of the disease on winter rape is also favoured by temperatures of 8–16 °C. In contrast, Chou (1970) noted that at 20–25 °C, infection occurs within 6 h of the conidium deposition on the surface of the host cotyledon. Haustoria develop from the infection hyphae in the epidermis as well as in those of the inner tissues (Chu 1935). The typical symptoms of infection by H. parasitica begin to appear 2 days after inoculation at 15 °C, and a day or two, later the formation of conidiophores and conidia is initiated. The haustoria in turnip and radish roots are at first spherical to pyriforms, becoming cylindrical or clavate, often dichotomously or trichotomously branched, the maximum dimensions of an unbranched haustorium in this situation being 18 × 25 μm compared with only 11 × 8 μm in the leaves. They are usually spherical and bilobate to trilobate and 57 × 14 μm in the stem of B. chinensis, where they are cylindrical or clavate and sometimes dichotomous. Some haustoria are surrounded by a sheath of variable extent from a collar around the neck to a third or half the length of the organ itself. The few full-grown haustoria found completely enveloped in vigorous roots inoculated with the fungus are probably incapable of functioning (Plate 7.1). The systemic invasion of the hypocotyls and cotyledons of cabbage seedlings may take place from the soil contaminated with oospores (LeBeau 1945). Further spread of the pathogen is by dissemination of conidia released from conidiophores formed on the cotyledons and hypocotyls (Chang et al. 1963). The pathogen can also enter directly through the inflorescence axis (Jang and Safeeulla 1990). The infection through the stigma and ovary wall results in embryonal infection. Pathogenesis in a susceptible combination is accompanied by large increases in electrolyte leakage and increased activity of the enzymes, β-glucosidase, ribonuclease, and peroxidase (Kluczewski and Lucas 1982). The large increase in β-glucosidase originates from the pathogen, and the enhanced ribonuclease activity is due to a new post-infectional form of the enzyme. Infected B. juncea produce cellulase, indo-PMG, and endo-PG (Singh et al. 1980). 7.2 The Process of Infection 159

Plate 7.1 Growth of crucifer downy mildew, Hyaloperonospora parasitica, in cotyledon tissues of Brassica. (a) Intercellular hyphae forming club-shaped intracellular haustoria in host cells, stained with trypan blue (x 250); (b) fluorescence micrograph of similar preparation, stained with aniline blue, showing bright collars, presumed to callose-like material of host origin, at sites of haustorial penetration (x 280); (c) electron micrograph of intercellular hypha (I) and haustorium (H) in host cell (HC). A second haustorium can be seen in the same cell (x4200) and (d) cell wall encasement surrounding developing haustorium at site of attempted penetration. Such host cell responses are commonly seen during development of the pathogen in partially resistant hosts (x 10,500) (Lucas et al. (1995)

7.2.1 Light Microscopic Observation of Infection Process

Light microscopic observations of whole-leaf mounts 18 h after inoculation of Arabidopsis strain Weiningen revealed that development of the fungus had advanced beyond the initial stages of infection, i.e. haustoria were already present. The majority of conidia were deposited over or in close proximity to anticlinal walls of adjoining epidermal cells. Conidia had germinated and formed appressoria, as shown in Plate 7.2a. Appressoria were, in most cases, produced directly from conidia; 160 7 Infection and Pathogenesis

Plate 7.2 Infection process and vegetative growth of Hyaloperonospora parasitica in Arabidopsis thaliana strain Weiningen (Koch and Slusarenko 1990). All samples were taken at 18 h after inoculation unless otherwise stated. (a) A germinated conidium (c) with an appressorium (a) on the leaf 7.2 The Process of Infection 161 however, well-developed germ tubes several spore diameters in length were also observed. Conidia and appressoria were empty by this time (Plate 7.2a), indicating that the cytoplasm had moved into penetration hyphae that extended from the appressorium and penetrated between the anticlinal walls of adjoining epidermal cells (Plate 7.2a–d). Material apparently deposited by the host in response to the ingress of the pathogen was observed in epidermal cells adjacent to the site of penetration (Plate 7.2b). Necrosis of epidermal cells in contact with the penetrating hypha was not observed in the Weiningen strain of Arabidopsis. Often, the first haustorium was inserted into one of the adjoining epidermal cells (Plate 7.2c). One haustorium in each of the epidermal cells neighbouring the penetrating hypha was also observed frequently (Plate 7.2e). In a few instances, haustorial bodies in epidermal cells were not fully expanded but were encapsulated with a material that had apparently been deposited by the host as a defence mechanism (Plate 7.2f). Growth of the penetrating hypha was, however, unaffected, and normal haustoria were present in the underlying mesophyll cells. This phenomenon may be related to observations by Chou (1970), who concluded from ultrastructural studies that epidermal cells of cabbage responded much more vigorously to infection with H. parasitica than did mesophyll cells. Sometimes the hypha passed between anticlinal walls without forming haustoria in the epidermis. In this case, single or multiple haustoria were produced upon contact of the penetrating hypha with cells of the mesophyll (Plate 7.2g). Three days after inoculation, extensive growth of the coenocytic, intercellular mycelium was evident on Arabidopsis strain Weiningen (Plate 7.2h). Hyphal strands were multiply branched, and haustoria were present in abundance. Necrosis of host cells or signs of retarded growth of the fungus were absent, indicating a high degree of compatibility between host and parasite. Young haustoria were pyriform (Plate 7.2e, g) and later became somewhat heart shaped (Plate 7.2h, i) to multiple lobed. Collar-like structures were often present around haustorial necks. At the time of sporulation, several haustoria were observed that were encased by host cell material. The encasements appeared as thick capsules surrounding the haustorial neck

Plate 7.2 (continued) surface; both structures are devoid of cytoplasm. Bar= 10 μm. (b) A penetration hypha (ph) at the point of entry between anticlinal walls (aw) of two epidermal cells. Note the opposition of material adjacent to the site of penetration (arrows). Bar= 10 μm. (c) The same infection site as in (E) but focused through to the epidermal cells. The penetration hypha (ph) has expanded, and the first haustorium. (h) has been formed in one of the epidermal cells. Note the material localized at the haustorial neck (arrows). aw, anticlinal walls. Bar= 10 μm. (d) Germinated conidia (c) on the leaf surface. In the cases shown, the appressoria (arrows) were produced directly from conidia without the formation of a germ tube. Appressoria are positioned over anticlinal walls (aw). Bar= 10 μm. (e) Simultaneous formation of haustoria (arrows) in both epidermal cells. aw, anticlinal walls. Bar= 10 μm. (f) Encasements (arrows) in epidermal cells surrounding haustorial initials. aw, anticlinal walls. Bar= 10 μm. (g) Formation of multiple haustoria (arrows) in mesophyll cells. The penetration hypha (ph) is arrowed. Bar = 10 μm. (h) Branched intercellular hyphae (ih) with numerous haustoria in mesophyll cells. A trichome (t) on the leaf surface can be seen clearly. Sample taken 3 days after inoculation. Bar = 100 μm. (i) Fully expanded haustoria (h) in mesophyll cells. Sample taken 3 days after inoculation. Bar = 10 μm. (j) Normal and encased (arrows) haustoria (h). The encasement is deposited around the haustorial neck and body. Sample taken 7 days after inoculation. Bar= 10 μm 162 7 Infection and Pathogenesis and body (Plate 7.2j). Encased haustoria had generally reached almost full size. Apparently, this host cell response occurred slowly and was possibly related to the ageing of the haustoria. Observation of tissues of Arabidopsis strain RLD 18 h after inoculation showed that one or both of the epidermal cells adjacent to the penetration hypha had reacted hypersensitively to the pathogen (Plate 7.3a, b). A hypersensitive response (HR), i.e. a rapid, localized necrosis of host cells, indicates a high degree of incompatibility between the host and pathogen. However, the fungus was able to penetrate through to the mesophyll where, although in a few cases branching was observed, apical growth of the hypha quickly ceased. Near the growing tip, the diameter of the hypha was markedly reduced compared with its size in the initial stages of ingress (Plate 7.3c, d). At 48 h after inoculation, approximately three to five deep blue-staining mesophyll cells were present below penetration sites and adjacent to hyphae. Although the stain penetrates fungal cells quite easily, it is usually excluded from healthy plant cells. Penetration of the stain into host cells is indicative of membrane damage. The stain has been used to highlight hypersensitivity responding host cells in incompatible interactions of plants with fungi (Keogh et al. 1980) and bacteria (Slusarenko and Longland 1986). Apart from the pronounced HR in strain RLD, the initial stages of infection, i.e. germination of conidia, formation of appressoria (Plate 7.3b, e), and penetration of hyphae between anticlinal walls of epidermal cells (Plate 7.3a, f), all occurred in the same manner as described for infection of Arabidopsis strain Weiningen. Plate 7.3f, b shows conidial germination and appressorium formation with and without germ tube production, respectively. In addition to the HR, apposition of host material in mesophyll cells at the point of contact with the penetrating hypha was observed (Plate 7.3g, h). Occasionally hyphae in contact with host cells formed a knob-like outgrowth, possibly in an attempt to produce haustoria. Fully developed haustoria, however, were never observed. In samples stained 48 h after inoculation, intercellular hyphae were very rarely seen. They appeared thin and devoid of cytoplasm and had in no case advanced beyond the immediate vicinity of the site of penetration (Koch and Slusarenko 1990). Sporulation and oospore formation on Arabidopsis strain Weiningen was recorded 7 days post-inoculation; both asexual and sexual reproduction of H. parasitica were observed in infected leaves of Arabidopsis strain Weiningen. Conidiophore primordial developed from the apices of comparatively very broad hyphae in sub-stomatal cavities. Plate 7.4a shows that conidiophores were similar in

Plate 7.3 (continued) Thecytoplasm in the two cells in contact with the hypha differs in appearance from that of the surrounding cells. Note the reduction in diameter of the hypha towards the tip. Bar= 10 μm. (E) to (H) Development of the fungus at a single penetration site, documented by varying the depth of focus. Note that no haustoria are present. (e) Conidium (c), germ tube (gt), and appressorium (a). Bar= 10 μm. (f) The penetration hypha (ph) can be seen growing between the anticlinal epidermal cell walls. The cell contents adjacent to the penetration hypha appear granular (arrows). Bar= 10μm. (g) and (h) Growth of the penetration hypha (ph) in the mesophyll. Material has been deposited by cells at the point of contact with the fungus (arrows). Bar= 10 μm 7.2 The Process of Infection 163

Plate 7.3 Infection process and vegetative growth of Hyaloperonospora parasitica in Arabidopsis thaliana strain Weiningen (Koch and Slusarenko 1990). (a) Hypersensitive reaction. Two necrotic epidermal cells adjacent to the penetration hypha (ph). Because of collapse of the epidermal cells, the outline of underlying mesophyll cells is visible (arrows). Bar= 10 μm. (b) Hypersensitive necrosis of a single epidermal cell after penetration of the hypha through the anticlinal wall. Letters a and c refer to appressorium and conidium, respectively. Bar= 1 μm. (c) and (d) Penetration hypha (ph) growing between palisade cells at two different depths of focus. Haustoria are not formed. 164 7 Infection and Pathogenesis

Plate 7.4 Asexual, and sexual reproductive structures of Hyaloperonospora parasitica in, and on tissues of Arabidopsis thaliana strain Weiningen (Koch and Slusarenko 1990). Samples were taken 7 days after inoculation. (a) Conidiophore (cp) emerging from the leaf surface; the conidia are 7.3 Nature and Mechanism of Pathogenesis 165 height to the leaf trichomes. Conidiophores showed a marked constriction in the region of the stoma (Plate 7.4b), and two conidiophores were often seen emerging from a single stoma. After growth out of the stomata (Plate 7.4c), conidiophores expanded, elongated, and quickly adopted a tree-like shape. Vesicles appeared at the end of each branch and expanded to form conidia. Mature conidia had a smooth to slightly verrucose surface (Plate 7.4b). Sexual reproduction started with the inter- twining of hyphae, which then differentiated to form oogonia and antheridia. Antheridia were of the paragynous type and appeared firmly appressed to oogonia (Plate 7.4d). Mature oospores were present in great number by 8 days after inoculation (Plate 7.4e). Their formation apparently coincided with the onset of sporulation (Koch and Slusarenko 1990). Before the onset of sporulation, infected leaves remained macroscopically free of disease symptoms. Sporulation of the fungus occurred 6 days after inoculation, after incubation of plants in the dark in a moist chamber for approximately 16 h. Conidiophores grew singly from stems and leaf petioles. Thick tufts of conidiophores were observed on the abaxial and adaxial side of leaves, as seen in Plate 7.5. Heavily infected plants died within 1–3 days. No sporulation or macroscopically visible symptoms were induced on inoculated plants of Arabidopsis strain RLD (Koch and Slusarenko 1990).

7.3 Nature and Mechanism of Pathogenesis

Pathogenesis of H. parasitica during compatible (S), intermediate, and incompatible (R) host interaction has been explained using TEM. Through transmission electron microscopy, ultrastructural characterization of pathogen development and host responses during compatible (susceptible) and incompatible (resistant) interactions between A. thaliana and H. parasitica (H. arabidopsidis) have been described in details by Soylu et al. (2004). In compatible interaction(s), the first haustoria were produced in epidermal cells, and intercellularly hyphae subsequently grew rapidly into the underlying mesophyll cells. During the early stages of infection, one to two haustoria were often observed in single mesophyll cells, but as many as four to five haustorial profiles were observed within a single host mesophyll cell at 5 days after inoculation (dai). The region around and between the haustoria contained an

Plate 7.4 (continued) partly discharged. Conidiophore and trichome (t) are similar in length. Bar = 45 μm. (b) The base of a conidiophore (cp) and several discharged conidia lying on the leaf surface. Note the constriction of the conidiophore in the stomatal opening (arrow). Conidia have a smooth to slightly verrucose surface. Bar = 5 μm. (c) A conidiophore initial (ci) growing out of a stoma and branching. Two conidiophore initials are apparently growing out of the neighbouring stoma (arrows). Bar =10 μm. (d) An oogonium (o) with a paragynous antheridium (an) can be seen in the mesophyll. h and ih, haustorium and intercellular hypha, respectively. Bar= 25 μm. (e) An oogonium (o) with an antheridium (an) attached and mature oospores. The different structural layers of the mature oospores are clearly visible; h, haustorium; ih, intercellular hypha; osp, oospore; osw, oospore wall; ow, oogonial wall; pe, periplasm. Bar = 25 μm 166 7 Infection and Pathogenesis

Plate 7.5 Sporulation of Hyaloperonospora parasitica on leaves of Arabidopsis strain Weiningen (viewed under a Stereo Microscope). A lawn of conidiophores is present on the leaves (thick arrows); on petioles the conidiophores are formed singly (thin arrows) (Koch and Slusarenko (1990)

abundance of cytoplasm with nucleus, many vesicles, chloroplasts, mitochondria, and microbodies (Plate 7.6a–c). Cell wall appositions were frequently found at sites of penetration around the base or neck of the haustorium (collar). Appositions were typically uniformly electron translucent but occasionally contained electron-dense vesicles (Plate 7.6a). The enormously localized deposits usually formed at the inner surface of the host cell wall without disruption of the host plasma membrane at site of penetration. By 5 days after inoculation, hyphae were found as far as 15 cell layers within the cotyledon tissue, and neither host cells nor hyphae of the pathogen appeared necrotic. By this time, accumulation of cell wall appositions was occasionally found to extend around haustoria (Plate 7.6d). In general, the cytoplasm of the ensheathed haustorium and associated intercellular hypha were apparently normal. During incompatible interaction (R) with accessions Ler-0 (minute necrotic flecks and no sporulation) and Ws-0 (necrotic pits and no sporulation), disorganization of the host cytoplasm in penetrated cells was observed. The reaction of Ler-0 to isolate EMoy 2 (minute necrotic flecks and no sporulation) was the more rapid and striking. Reactions in both accessions were characterized by the disorganization of the host cytoplasm in penetrated cells. The reaction of Ler-0 to Emoy2 (the FN phenotype) was the more rapid and striking. Epidermal cells appeared to undergo the HR soon after penetration through anticlinal walls (Plate 7.7a, b). Despite epidermal cell collapse 1 dai, invading hyphae remained alive with clearly distinguish- able organelles (Plate 7.7a). Large aggregations of plant cytoplasm, wall appositions, and electron-dense deposits were found in some penetrated epidermal cells (Plate 7.7b). Cell wall appositions were also frequently observed in adjacent epidermal cells, although no penetration was observed in sections. Numerous amorphous electron-dense vesicles were present in the cell wall appositions formed during the incompatible interactions. By 2 dai, many penetrated and neighbouring epidermal cells appeared disorganized with disintegrated cytoplasm showing numerous abnor- 7.3 Nature and Mechanism of Pathogenesis 167

Plate 7.6 Ultrastructural features of the compatible interaction between the Emoy2 isolate and susceptible accession Oy-0 sampled 3 (a–c) or 5 dai (d). (a–c) Median sections through penetration points showing haustoria connected to large intercellular hyphae. Note the presence of nucleus, lipid bodies, mitochondria, small and large vacuoles in the intercellular hyphae, and haustoria. Cell wall appositions (asterisks) occurred at the penetration points. The wall of the intercellular hypha is at its thickest where it penetrates the host cell wall to form the haustorial neck. The host mesophyll cell appears unaffected by the presence of the haustorium as organelles are well preserved. (d) Shows the formation of haustorial ensheathment (arrow) during a compatible interaction. Bars 2 mm; ch chloroplast, Cv cell vacuole, H haustorium, IS intercellular space, IH intercellular hyphae, l lipid body, m mitochondrion, n nucleus, Pv pathogen vacuole (Soylu et al. (2004) 168 7 Infection and Pathogenesis

Plate 7.7 Incompatible interaction between Emoy2 and the resistant accession Ler-0, 1 (a and b), 2 (c), and 3 dai (d). (a) Penetration of host tissue between anticlinal walls of two epidermal cells. Note that epidermal cells contain electron-dense cytoplasm and distorted organelles, but the associated pathogen penetration peg (pp) remains intact. (b) A necrotic mesophyll cell with shrunken electron-dense cytoplasm (arrows). The haustorial body is totally necrotic. (c) Collapsed epidermal cells and extensive vacuolation (asterisks) with wall alterations in an adjacent mesophyll cell. (d) Typical necrotic mesophyll cell underlying a necrotic epidermal cell (arrow). Note that both the epidermal and mesophyll cell contain several distorted host organelles. Bars 2 mm; EC epidermal cell, H haustorium, IH intercellular hyphae, IS intercellular space, MC mesophyll cell, pp penetration peg (Soylu et al. (2004) 7.3 Nature and Mechanism of Pathogenesis 169 mal host organelles such as mitochondria, chloroplasts, and endoplasmic reticulum (ER). As indicated from light microscopy, the growth of EMoy-2 was restricted close to the site of penetration in Ler-0. Deposition of callose (confirmed by immunogold labelling) was frequently seen in both necrotic and adjacent epidermal cell walls and also in the underlying mesophyll cells (Plate 7.7c). Cell wall appositions appeared thinner in these mesophyll cells than those in epidermal cells. Extensive vacuolation was frequently observed in mesophyll cells below the dead epidermal cells (Plate 7.7c). By 3 dai, a common reaction in penetrated epidermal cells and underlying mesophyll cells was the increasing electron density of cytoplasm and loss of organellar structure (Plate 7.7d). The resistant reaction in Ws-0, exhibiting the more extensive PN phenotype (necrotic pits and no sporulation) was different from that observed on Ler-0. Growth of the pathogen was not restricted to that reached during the first dai. As intercellular hyphae extended, the cells containing haustoria close to the penetration site (including the first penetrated epidermal cells) collapsed, often without extensive deposition of callose containing ensheathments (Plate 7.8a). Three days after inoculation, penetrated cells at the edge of the advancing lesion had collapsed and often contained dense appositions lining their cell walls and surrounding haustoria (Plate 7.8b–d). Most intercellular hyphae observed within the lesion were necrotic at this time. Necrosis of mesophyll cells had often extended beyond the outer limits of spread of hyphae 5 dai. The responding cells were characterized by the formation of extensive wall appositions, cytoplasmic vacuolation, and the progressive degeneration of organelles (Plate 7.8d). However, during intermediate interaction with Col-o (minute necrotic flecks, delayed, and low sporulation) in cotyledons of Col-0, during the first 2 dai, many infection sites were found to contain haustoria in host cells that appeared similar to those observed during compatible interactions (Plate 7.9a). By 3 dai, unsheathing materials developed around many haustoria, and penetrated cells often appeared necrotic (Plate 7.9b, c). Observation of several infection sites showed that ensheathment progressed from the mesophyll cell penetration point and gradually extended along the haustorial neck and around the haustorial body, finally to surround the haustorium completely. Despite such depositions, at certain sites hyphae continued to spread intercellularly, albeit more slowly than in fully susceptible tissue, and formed numerous haustoria. Sections taken from cotyledons showing sparse or no sporulation, 5 and 7 dai, revealed differential reactions in mesophyll cells. The most recently penetrated cells were similar to those observed during compatible interactions with at most small cell wall appositions at the penetration site. In other cells, haustoria were ensheathed. The first penetrated cell showed ensheathment and also various stages of necrosis indicative of the HR. Progressive vacuolation, increasing electron density of cytoplasm, and loss of organelles were also observed within haustoria in mesophyll cells undergoing the HR and also associated intercellular hyphae (Plate 7.9d, e). By 7 dai, both invading hyphae and haustoria gradually became disorganized and finally necrotic as illustrated by the accumulation of amorphous electron-dense material in their degenerated cytoplasm in which organelles were hardly discernible (Plate 7.9f). 170 7 Infection and Pathogenesis

Plate 7.8 Incompatible interaction between Emoy 2 and the resistant accession Ws-0, 2 (a), 3 (b and c), and 5 dai (d). (a) A haustorium in a dead mesophyll cells. The plant plasma membrane (arrow) has dislocated from the plant cell wall, and the penetrated cell exhibits organelle disruption. (b) Shows a necrotic intercellular hypha and haustorium. Note that both host and pathogen contain electron-dense cytoplasm in which organelles are hard to distinguish. (c) A necrotic mesophyll cell containing a heavily encased haustorium. Both haustorium and connected hypha are necrotic. (d) Shows penetrated and nearby mesophyll cells. Note the upper necrotic mesophyll cell contains several distorted chloroplasts, electron-dense cytoplasm, and darkly stained wall apposition (arrows) along the cell wall. The cytoplasm of the adjacent cell is severely disorganized containing misaligned chloroplasts and nucleus. A callose-containing deposit (asterisk) is also present along the cell wall. Bars 2 mm; H haustorium, IH intercellular hyphae, IS intercellular space, MC mesophyll cell, ch chloroplast, n nucleus (Soylu et al. (2004) 7.3 Nature and Mechanism of Pathogenesis 171

Plate 7.9 Pathogen development and host cell responses during the intermediate interaction between Emoy-2 and the accession Col-0. (a–c) Sections through the penetration point of the haustorium showing accumulation of cell wall apposition 2 (a) and 3 dai (b and c). Note that cell wall appositions (arrows) develop at penetration points (a), spread along the plant cell wall (b), and gradually extend around the haustoria (c). (d–f) Show penetration of mesophyll cells associated with cell disorganization 5 (d) and 7 dai (e and f). In (d), the host plasma membrane (arrow) has retracted from the cell wall, and penetration is associated with vacuolation and cytoplasmic disorganizations as characterized by the accumulation of electron-dense deposits (asterisks) along the tonoplast. (e) Shows a very rare infection site at which an apparently viable haustorium is located within a collapsed mesophyll cell. The tonoplast of the penetrated cell has ruptured, and organelles 172 7 Infection and Pathogenesis

Plate 7.9 (continued) have dispersed into the central vacuole. In (f), the penetrated cell, intercellular hypha, and haustorium are necrotic as illustrated by the accumulation of amorphous material in their cytoplasm. Bars, (a) 1 mm; (b)–(f) 2 mm; ch chloroplast, Cv cell vacuole, H haustorium, IS intercellular space, IH intercellular hyphae, m mitochondrion, n nucleus (Soylu et al. (2004)

Immunocytochemical localizations of callose have been observed in susceptible host-pathogen interaction at infection sites. Immunogold labelling using a commercially available antibody against branched β-1,3-linked glucan confirmed the presence of callose at infection sites. The examination of numerous sections from susceptible tissue revealed that callose labelling was consistently detected in cell walls within intercellular hyphae, around the haustorial body, and in cell wall appositions at sites of penetration. Some labelling was also observed in membrane- like structures found in the cytoplasm of intercellular hyphae. No labelling was observed on the extra haustorial matrix or in any part of the host cell including cell wall, cytoplasm, and organelles (Plate 7.10a, b). Accumulations of callose during incompatible interactions are illustrated in Plate 7.10c, d. Sites of labelling corre- sponded to electron translucent regions of deposits and ensheathing materials. The large papillae which formed in living mesophyll cells adjacent to dead cells were intensely labelled, and some gold particles were also observed in necrotic cells (Plate 7.10d). In conclusion, the resistance of Arabidopsis accessions harbouring the resistance genes to Emoy2 carrying corresponding Avr genes is characterized by the induction of a wide range of defence mechanisms including the encasement of haustoria with callose-containing material and the HR in invaded and neighbouring cells. Limited pathogen development is clearly associated with the HR in FN and PN phenotypes. The contribution of callose deposition per se to plant defence, particularly in FDL phenotype, should be addressed using the approach of reverse genetics. 7.3 Nature and Mechanism of Pathogenesis 173

Plate 7.10 Immunogold localization of callose during compatible and incompatible interactions between Emoy2 and the Arabidopsis accessions Oy-0 (a, and b), Col-0 (c), and Ws-0 (d). In (a) and (b) note that labelling is confined to the pathogen cell wall within the intercellular hypha (large arrows), around the haustorial body (small arrows) and collar (*) at the site of penetration, 3 dai. In (c), callose is detected in the material (arrows) ensheathing haustoria, and within the cell wall, 3 dai. In (d) very dense labelling is found within the cell wall apposition (arrow) in a cell adjacent to a necrotic mesophyll cell, 5 dai. Bars, (a) 2 mm, (b)–(d) 1 mm; Cv cell vacuole, H haustorium, IH intercellular hyphae, MC mesophyll cell (Soylu et al. (2004) 174 7 Infection and Pathogenesis

References

Achar PN (1992) Dose-response relationship of Peronospora parasitica in Raphanus sativus. Phyton (Buenos Aires) 53:89–94 Awasthi RP, Nashaat NI, Heran A, Kolte SJ, Singh US (1997) The effect of Albugo candida on the resistance to Peronospora parasitica and vice versa in rapeseed-mustard. ISHS Symposium on Brassicas, Tenth Crucifer Genetics Workshop, 23–27 Sept 1997, Rennes, France Butler EJ (1918) Fungi and diseases in plant. Thaker Spink & Co., Calcutta, pp 297–300 Chang IH, Xu RF, Chiu WF (1963) On the primary sources of infection of the downy mildew of Chinese cabbage caused by Peronospora parasitica (Pers.) Fr. and the limited systemic infection of seedlings. Acta Phytopathol Sin 6:153–162 Chou CK (1970) An electron-microscope study of host penetration and early stages of haustorium formation of Peronospora parasitica (Fr.) Tul. on cabbage cotyledons. Ann Bot 34:189–204 Chu HT (1935) Notes on the penetration phenomena and haustorium formation of Peronospora brassicae Gaum. Ann Phytopathol Soc Japan 2:150–157 Felton MW, Walker JC (1946) Environmental factors affecting downy mildew of cabbage. J Agril Res 72:69–81 Jang P, Safeeulla KM (1990) Modes of entry, establishment and seed transmission of Peronospora parasitica in radish. Proc Indian Acad Sci Plant Sci 100:369–373 Jonsson R (1966) Peronospora on oil yielding Brassicas. Methods for testing resistance in winter rape and their results. Sver Utsadestor Tidskr 76:54–62 Keogh RC, Deverall BJ, McLeod S (1980) Comparison of histological and physiological responses to Phakopsora pachyrhizi in resistant and susceptible soybean. Trans Br Mycol Soc 74:329–333 Kluczewski SM, Lucas JA (1982) Development and physiology of infection by the downy mildew fungus Peronospora parasitica (Pers. ex Fr.) Fr. in susceptible and resistant Brassica species. Plant Pathol 31:373–389 Koch E, Slusarenko AJ (1990) Arabidopsis is susceptible to infection by a downy mildew fungus. Plant Cell 2:437–445 Krober H (1969) Degree of infestation of cabbage turnip by Peronospora parasitica (Pers.) Fr. and of tobacco by Peronospora tabacina Adam as related to concentration of conidia. Phytopath Z 66:180–187 LeBeau FJ (1945) Systemic invasion of cabbage seedlings by the downy mildew fungus. J Agric Res 71:453–463 Lucas JA, Hayter JBR, Crute IR (1995) The downy mildews: host specificity and pathogensis. In: Kohmoto K (ed) Pathogensis and host specificity in plant diseases, vol Vol. 2. Elsevier Science Ltd, Oxford Preece TF, Barnes G, Bayley JM (1967) Junctions between epidermal cells as sites of appresorium formation by plant pathogenic fungi. Plant Pathol 16:117–118 Shiraishi M, Sakomoto K, Asada Y, Nagatani T, Hidaka H (1975) A scanning electron microscopic observation on the surface of Japanese radish leaves infected by Peronospora parasitica (Fr.) Fr. Ann Phytopathol Soc Japan 41:24–32 Singh SB, Singh DV, Bains BS (1980) In vivo cellulase and pectinase production by Albugo candida and Peronospora parasitica. Indian Phytopath 33:370–371 Slusarenko AJ, Longland AC (1986) Changes in gene activity during the expression of the hypersensitive response in Phaseolus vulgaris cv. Red Mexican to an avirulent race 1 isolate of Pseudomonas syringae pv. phaseolicola. Physiol Mol Plant Pathol 29:79–94 Soylu EM, Soylu S, Mansfield JW (2004) Ultra-structural characterization of pathogen development and host responses during compatible and incompatible interactions between Arabidopsis thaliana and Peronospora parasitica. Physiol Mol Plant Pathol 65:67–78 Chapter 8 Disease Cycle

8.1 Introduction

Downy mildew of crucifer’s pathogen is an obligate parasite and is specific to the kind of host it attacks, possibly because it has co-evolved in the nature along with the kind of hosts available. The pathogen requires certain specific nutrients which are produced or become available during pathogenesis to proliferate and continue in life cycle. In every infectious disease, there is a series in succession of one another and lead to the development and perpetuation of the disease and the pathogen. The chain of events is called a disease cycle. A disease cycle sometimes corresponds fairly closely to the life cycle of the pathogen, but it refers primarily to the appearance, development, and perpetuation of the disease rather than the pathogen. The disease cycle involves the changes in the plant, and the plant symptoms as well as those in the pathogen, and spans periods within a growing season and from one growing season to the next. The main events in a disease cycle include inoculation (landing of pathogen propagules on the plant in the nature or otherwise), penetration, infection, growth, and reproduction of the pathogen, dissemination of the pathogen, and overwintering or oversummering of the pathogen. Downy mildew of crucifers is a polycyclic disease under natural conditions. The disease cycle has been explained in the sections 1, general on crucifers, and 2 specific on Arabidopsis as under.

8.2 General Disease Cycle on Cruciferous Crops

Downy mildew of crucifers is essentially a disease of foliar and other aerial plant tissues. The fungus survives as oospores in A. candida-induced malformed inflorescence and senesced host tissues, as conidia on leaves, and inflorescence, as latent systemic mycelium in seeds or infected plant debris. Infections are favoured by temperatures between 10 and 15 °C and by high atmospheric humidity following

© Springer Nature Singapore Pte Ltd. 2017 175 G. S. Saharan et al., Downy Mildew Disease of Crucifers: Biology, Ecology and Disease Management, https://doi.org/10.1007/978-981-10-7500-1_8 176 8 Disease Cycle rain or heavy dew. The conidia produce germ tubes which penetrate anticlinal cell walls often on the lower surface of the leaves. The penetration is usually direct but occasionally also occurs through a stoma (Shiraishi et al. 1975). Primary infection from soil-borne oospores has been obtained (LeBeau 1945; Chang et al. 1963). Transmission by infected seed is possible but its importance has not been well documented. Further spread of the pathogen is by dissemination of conidia released from conidiophores formed on the cotyledons or hypocotyls. The true leaves are usually infected through wind-borne conidia, resulting in spread of the disease through secondary infection. The pathogen dispersal over short distances in water droplets can also occur. Although there is no exact information on the relationship between leaf and floral infection under natural conditions, most inflorescence infections, as in Albugo candida, probably result from secondary spread of the pathogen rather than systemic infection. The diagrammatic life cycle of the disease developed by Lucas et al. (1995) is given in Fig. 8.1. The disease is typical of cool, moist conditions, but empirical data for optimal field conditions for disease development are lacking. Nevertheless, a general rule of thumb for other downy mildews on Brassicas is that disease in the field is most problematic between 10 and 15 °C and high humidity. However, it should be borne in mind that different developmental stages of the pathogen might show different optima, as is the case for other cruciferous downy mildews. Thus, Felton and Walker (1946) showed that the germination of conidia was most rapid at 8–12 °C, whereas penetration was optimal at 16 °C and haustoria formation at 20–24 °C. For laboratory-grown Arabidopsis, the routine conditions employed for infection and conidiophore production are 16–18 °C and 100%

Fig. 8.1 Diagramatic life cycle of Hyaloperonospora parasitica causing downy mildew of crucifers (Lucas et al. 1995) 8.3 Disease Cycle on Arabidopsis 177

R.H. Sporulation occurs mainly at night, and spores are disseminated during the morning as drying conidiophores twist violently and fling conidia into the air. If you want a good infectious preparation of conidia, it is better to be early; otherwise, many of the spores will already have been lost from the conidiophores on the leaves. A detailed diagrammatic representation of downy mildew on cruciferous crops has been given by Saharan et al. (2005) (Fig. 8.2). The pathogen perpetuates in the soil through oospores which are formed in abundance in the malformed tissues of the infected plants. Seeds may get contaminated with plant trash containing oospores during threshing operation. Infection originates when such seeds are sown after getting suitable temperature and relative humidity. Oospores formed in malformed and senesced host tissues constitute an important means of survival of H. parasitica over periods of unfavourable conditions. It is also known to survive through mycelium and conidia (Fig. 8.2) for a short period depending on environmental conditions in an area. The presence of H. parasitica mycelium in the seed coat of Chinese cabbage has been recorded by Chang et al. (1963). According to Jang and Safeeulla (1990), the presence of mycelium in the pericarp and embryo of radish seeds varies from 0.1 to 12.5%. The coenocytic branched mycelium is clearly visible in the intercellular space of the pericarp. In the embryonic tissues, the mycelium is comparatively thin. The percentage of seeds with viable mycelium is directly correlated with the percentage of embryo infection. Conidia of H. parasitica on cabbage survive longer under cool and dry conditions. Relative humidity is more important than temperature. In the field, conidia can survive on detached leaves of kohlrabi for 10 days during warm days. When buried in dry soil, conidia can survive for 110 days. The survival period is greatly reduced to a maximum of 22 days if the soil is moist. Marked reduction in survival has also been observed after storage in both dry and moist soils during the summer. The conidial viability is longest, up to 130 days when the spores are stored in air-dried soil at a constant temperature of 5 °C. Conidia kept at −25 °C and relatively dry on leaf disc (air dried at 20 °C) maintain a relatively high rate of germination after 1 year or longer. Oospore formation is abundant in the infected tissues of all crucifers which form primary source of the pathogen. In radish and rapeseed-mustard, there is abundant production of oospores in infected leaf tissues, on the seed surface, pericarp, and embryo of seeds. However, in rapeseed and mustard, seed transmission is low and may be nonsystemic, ranging from 0.4 to 0.9 percent in the seedlings grown from infected seeds.

8.3 Disease Cycle on Arabidopsis

The disease cycle of H. parasitica on Arabidopsis has been described by Slusarenko and Schlaich (2003). In spring, new infections of Arabidopsis plants occur via oospores which have overwintered in leaf debris in the soil. Successive rounds of infection on leaves and cotyledons occur via conidia. In contrast to several H. parasitica isolates infecting other Brassica, H. parasitica infecting Arabidopsis is 178 8 Disease Cycle

Fig. 8.2 Disease cycle of downy mildew of crucifers (Saharan et al. 2005) 8.3 Disease Cycle on Arabidopsis 179

Fig. 8.3 Life cycle of Hyaloperonospora parasitica. (a) Infections arise initially from oospores germinating in the soil. (b) Plants are colonized by a coenocytic, intercellularly growing mycelium which swells to fit the intercellular spaces, giving it an irregular appearance. The hyphae put out pear-shaped feeding organs called haustoria into host cells. After a variable period of growth (1–2 weeks), conidiophores, bearing asexual, spherical hyaline conidiospores (c), grow out of stomata. (d) On germination, conidia initiate new rounds of infection. (e–g) Oospores are formed concurrently with asexual spores. (e) The female sexual organs, oogonia, contain an oosphere that is fertilized via a fertilization tube growing through its outer wall from the male antheridium. (f) The fertilized oosphere develops into a mature oospore. (g) Oospores are very profuse in infected leaves (Mauch-Mani and Slusarenko 1994) homothallic. Oospores are usually found in the leaves of Arabidopsis a week or so after infection of true or cotyledon leaves by conidia of a single isolate. The disease cycle is shown in Fig. 8.3. Infection cycle begins via oospore germination and infecting roots. Infection arises after a conidium germinates to either directly produce an appressorium or after making a short germ tube, generally within 6 h of coming into contact with the leaf. A penetration hypha grows from the underside of the appressorium, and this penetrates the leaf at the anticlinal juncture of two epidermal cells (Fig. 8.3, Koch and Slusarenko 1990). Rarely, an appressorium forms over a stomata, and the penetration hypha grows into the leaf directly through the stoma. Haustoria are often budded off into the epidermal cells as the penetration hypha grows down between them, and further haustoria are produced into mesophyll cells as the hyphae make intercellular growth. Conidiophores are produced from conidiophore initials which grow out of the stomatas (Koch and Slusarenko 1990; Mauch-Mani and 180 8 Disease Cycle

Slusarenko 1994). The highest conidiophore density is generally observed where the density of stomata is greatest, i.e. on the underside of the leaves. Incompatible combinations of different Arabidopsis/Hyaloperonospora genotypes can show a range of resistance phenotypes. Thus, leaves of RLD show a rapid and complete cessation of growth associated with the death of one or a few plant cells (i.e. a typical HR) against WELA, whereas in Col-0, WELA makes more growth, and the HR extends well into the mesophyll. In some genotype combinations, for example, Columbia with EMOY, necrosis appears behind the advancing hyphal tip (Holub et al. 1994) giving rise to a trailing necrosis phenotype which had previously been noted in some circumstances in susceptible tissue conditioned to systemic acquired resistance (Uknes et al. 1992). In a compatible combination, the degree and rate of colonization and the quantity of conidiophore production also vary in a genotype interaction-specific manner (Holub et al. 1994).

8.4 Factors Affecting Disease Cycle

There are numbers of biotic and abiotic factors which may be congenial and/or detrimental for the life cycle of the pathogen influencing disease cycle. The environmental factors, viz. temperature, relative humidity, wind velocity, duration, and intensity of rainfall, and dew period have great effect on the pathogen survival, infection, multiplication, and dissemination to serve as primary and secondary source of inoculum. Secondary spread of the pathogen is also influenced by these factors. The cultural factors like time of planting, nutritional conditions (amount of N, P, K, and micronutrients applied), cultural operations, cultivars grown, crop rotation, continuous cropping of crucifers, plant protection measures adopted, etc. have an enormous bearing on the life cycle of downy mildew of crucifers. The biotic factors related to the host-like closeness to the primary source of inoculum, age of host and tissues for infection, duration of availability of young tissues for infection, higher plant population (close spacing), duration of leaf wetness, absence of wax on leaf surface, susceptibility of host tissues, virulence spectrum of the pathogen, host vulnerability conditions created by Albugo, and mustard mosaic virus are favourable factors for polycyclic diseases like downy mildew of crucifers.

References

Chang IH, Xu RF, Chiu WF (1963) On the primary sources of infection of the downy mildew of Chinese cabbage caused by Peronospora parasitica (Pers.) Fr. and the limited systemic infection of seedlings. Acta Phytopathol Sin 6:153–162 Felton MW, Walker JC (1946) Environmental factors affecting downy mildew of cabbage. Aust J Agric Res 72:69–81 References 181

Holub EB, Beynon JL, Crute IR (1994) Phenotypic and genotypic characterization of interactions between isolates of Peronospora parasitica and accessions of Arabidopsis thaliana. Mol Plant Microbiol Interact 7:223–239 Jang P, Safeeulla KM (1990) Modes of entry, establishment and seed transmission of Peronospora parasitica in radish. Proc Indian Acad Sci, Plant Sci 100:369–373 Koch E, Slusarenko AJ (1990) Arabidopsis is susceptible to infection by a downy mildew fungus. Plant Cell 2:437–445 LeBeau FJ (1945) Systemic invasion of cabbage seedlings by the downy mildew fungus. Aust J Agric Res 71:453–463 Lucas JA, Hayter JBR, Crute IR (1995) The downy mildews: host specificity and pathogenesis. In: Kohmoto K, Singh US, Singh RP (eds) Pathogenesis and host specificity in plant diseases: histopathological, biochemical, genetic and molecular bases, Eucaryotes, vol II. Pergamon, Oxford, pp 217–238 Mauch-Mani B, Slusarenko AJ (1994) Downy mildew of Arabidopsis thaliana. In: Bowman J (ed) Arabidopsis—an Atlas of morphology and development. Springer, New York, pp 414–417 Saharan GS, Naresh M, Sangwan MS (2005) Diseases of oilseed crops. Indus Publication Co., New Delhi. 643p Shiraishi M, Sakamoto K, Asada Y, Nagatani T, Hidaka H (1975) A scanning electron microscopic observation on the surface of Japanese radish leaves infected by Peronospora parasitica (Fr.) Fr. Ann Phytopathol Soc Japan 41:24–32 Slusarenko AJ, Schlaich NL (2003) Downy mildew of Arabidopsis thaliana caused by Hyaloperonospora parasitica (formerly Peronospora parasitica). Mol Plant Pathol 4:159–170 Uknes S, Mauch-Mani B, Moyer M, Potter S, Williams S, Dincher S, Chandler D, Slusarenko A, Ward E, Ryals J (1992) Acquired resistance in Arabidopsis. Plant Cell 4:645–656 Chapter 9 Epidemiology and Forecasting

9.1 Introduction

In epidemics of downy mildews, the pathogen population starts from a low level of initial inoculum which then increases exponentially through successive cycles on the host during the growing season. Therefore, downy mildew of crucifers is a compound interest disease. The seasonal increase of the pathogen population has been investigated much more thoroughly than that of the initial inoculum. Information has been generated on the multiplication phase of the disease which relates to the sequence of events in the life of the pathogen on its host, which are infection, colonization, and sporulation. The studies on forecasting of crucifers downy mildew disease is limited to prediction models.

9.2 Disease Development in Relation to Temperature, Humidity, Rainfall, and Leaf Wetness

The relationship of host-pathogen-environment interaction in case of downy mildew of crucifers is a complex phenomenon which determines the rate of disease development (Fig. 9.1). Among the major environmental factors which markedly influence the development of downy mildew is air temperature and relative humidity Environmental factors effect:Leaf wetness. The rate of spore germination and host penetration is affected by temperature variations. Chu (1935) found that at 15 °C conidia germinate in 4–6 h, appressoria form in 12 h, and penetration occurs in 18–24 h. According to Eddins (1943), the downy mildew of cabbage is most destructive when the temperature ranges between 10 °C and 15 °C and when the plants remain wet until mid-morning for 4 consecutive days. However, Felton and Walker (1946) reported that on cabbage, germination of the conidia (Fig. 9.2) and subsequent penetration of the host take place most

© Springer Nature Singapore Pte Ltd. 2017 183 G. S. Saharan et al., Downy Mildew Disease of Crucifers: Biology, Ecology and Disease Management, https://doi.org/10.1007/978-981-10-7500-1_9 184 9 Epidemiology and Forecasting

Fig. 9.1 The relationship of host, pathogen, and environment in the interaction phenotype of downy mildew of crucifers rapidly at 8–12 °C and 16 °C, respectively. Formation of haustoria and growth of the fungus in the host tissues are most rapid at 20–24 °C (Fig. 9.3). Symptoms develop quickly at 24 °C, but sporulation and reinfection is limited at 24 °C and 28 °C. The lower temperature of 16 °C results in slower growth of both the host and the pathogen, less damage, more prolific sporulation, more reinfection, and, consequently, more profuse disease development. The severity of the disease at 10–15 °C seems to be the effect of temperature upon production of inoculum, spore germination, and infection (Figs. 9.4 and 9.5). However, according to Saharan et al. (1997), a temperature of 15 °C seems to be the most favourable for epidemic development as this favours slower growth of both host and pathogen resulting in less drastic damage and hence more profuse disease development. The temperature range for maximum infection of seedlings of a highly susceptible cabbage cv. and subsequent disease development in vitro was 15–25 °C, and 90–100% infection was achieved after 48 h of incubation. At less than 15 °C and 26–30 °C temperature, infection percentage was decreased to 40–50% and 35–40%, respectively. No disease incidence was recorded at temperature above 35 °C. Penetration of cotyledons by germ tube was mostly via stomata and occasionally directly through the cuticle (Achar 1998). In Spain, downy mildew disease intensity on cole crops was positively correlated with RH and negatively with mean temperature and number of hours of daily insolation. A low number of hours of insolation for several consecutive days combined with cool temperature appeared to be determining factors in disease development (Sinobas Alonso and Diaz Alonso 1995). 9.2 Disease Development in Relation to Temperature, Humidity, Rainfall, and Leaf… 185

100 HOURS 16

90 8

6 80

60 4½

30 SPORES GERMINATING (PERCENT)

1½ 3 0 4812 16 20 24 TEMPERATURE (°C)

Fig. 9.2 Effect of time and temperature on germination of conidia of Hyaloperonospora parasitica (Felton and Walker 1946)

By contrast, Chou (1970) noted 20–25 °C, and Nakov (1972) found 15–20 °C as the most favourable temperature for infection. In temperate coastal regions of Madison, Wisconsin, USA, where Chinese cabbage is grown from late summer through the winter and spring, downy mildew thrives during periods of frequent rains and high humidity. There is an 8–12 h requirement of 100% RH for the production and dissemination of its airborne conidia. Once inside the Chinese cabbage, hyphae spread through the leaves, petioles, and stems, first feeding on the cells without apparent injury then suddenly causing yellowing, collapse, and death of the tissues. Conidiophores and conidia are produced primarily on the lower side of the leaves (Williams and Leung 1981). On Brassica oilseeds, H. parasitica is favoured by temperatures of 8–16 °C, moist air, and weak light (Jonsson 1966; D'Ercole 1975). According to Bains and Jhooty (1979), a 17 °C temperature and 51 mm rainfall result in low infection of 186 9 Epidemiology and Forecasting

8 PRODUCTION OF HAUSTORIA COLLAPSE OF SPORE PENETRATION FORMATION OF 7 APPRESSORIA APPEARANCE OF GERM TUBES

4 TIME ( HOURS )

0 4 8121620 24 TEMPERATURE (°C)

Fig. 9.3 Effect of temperature upon penetration and development of haustoria of Hyaloperonospora parasitica (Felton and Walker 1946) mustard in contrast to high infection at 14 °C temperature and 152 mm rainfall during the crop season. In a subsequent study, 15–20 °C were the best temperatures for infection and development of downy mildew. At this temperature regime, infection occurs within 24 h of inoculation (Table 9.1, Fig. 9.6). The infection frequency is reduced at 25 °C temperature with no infection observed at 30 °C temperature (Table 9.1, Fig. 9.6). The maximum area under disease progress curve occurs at 20 °C temperature (AUDPC-60.54%, Fig. 9.6). Leaf wetness duration of 4–6 h at 20 °C temperature and for 6–8 h at 15 °C temperature is essential for severe infection and disease development on mustard (Tables 9.2 and 9.3 Figs. 9.7 and 9.8). 9.2 Disease Development in Relation to Temperature, Humidity, Rainfall, and Leaf… 187

132

INITIAL SPORULATION

APPEARANCE OF SYMPTOMS 120 HIGH HUMIDITY

APPEARANCE OF SYMPTOMS LOW HUMIDITY

108

72 TIME AFTER INOCULATION ( HOURS )

40 12 16 19 20 26 TEMPERATURE (°C)

Fig. 9.4 Effect of five different temperatures on the initial sporulation of Hyaloperonospora parasitica at high humidity and upon initial appearance of symptoms at low and at high humidity (Felton and Walker 1946)

The infection frequency, and disease development increases significantly with the increase in duration of leaf wetness (Mehta et al. 1995). According to Kolte et al. (1986), sunshine has a significant negative correlation, whereas total rainfall has a significant positive correlation with A. candida-induced stag head development on rapeseed-mustard (Table 9.4, Fig. 9.9). The data in Table 9.4 revealed that prediction equations for the development of staghead incidence due to downy mildew and white rust disease complex were up to 68%, whereas staghead severity was up to 62% when all the weather factors were taken into consideration. A reduced period of sunlight (2–6 h/d) and rainfall of up to 161 mm during the flowering period favours severe occurrence of the stag heads. Maximum temperature was positively correlated with disease index of white rust, downy mildew, and Alternaria blight. Maximum temperature from 26 to 290 C and average relative humidity of more than 65% (Tables 9.5 and 9.6) favoured the 188 9 Epidemiology and Forecasting

Fig. 9.5 Graphic summary of infection by and development of Hyaloperonospora parasitica on cabbage plants grown in sand culture supplied with various nutrient solutions (Felton and Walker 1946) development of all the three diseases of B. juncea (Sangeetha and Siddaramaiah 2007). Under West Bengal conditions, downy mildew of mustard was at its peak when maximum temperature was >27 °C, minimum temperature >10 °C, and relative humidity around 100% in the morning and 50% in the afternoon (Banerjee et al. 2010) In Ukraine and Russia, downy mildew of white cabbage is more severe with abundant rain (75–100 mm/10 year) and a 14–15 h of day light (Vladimirskaya et al. 1975). 9.2 Disease Development in Relation to Temperature, Humidity, Rainfall, and Leaf… 189

Table 9.1 Effect of temperature on infection by Hyaloperonospora parasitica and disease development on mustard seedlings (cv. RH-30) (Mehta et al. 1995) Percent disease incidence after inoculation (h) Temp. (°C) 24 48 72 96 Mean AUDPCa 10 0.00 (1.8) 54.9 (47.5) 64.192 70.5 (54.5) 47.2 (40.1) 43.8 (53.5) 15 2.91 (6.7) 58.5 (50.3) 73.9 (60.4) 75.6 (62.4) 52.7 (44.9) 45.9 20 34.31 78.3 (62.5) 87. 6 (70.0) 90.3 (75.0) 72.6 (60.8) 60.5 (35.6) 25 0.00 (1.8) 5.1 (9.3) 14.3 (20.9) 19.7 (25.8) 9.8 (14.3) 14.9 30 0.00 (1.8) 0.0 (1.8) 0.0 (1.8) 0.0 (1.8) 0.00 (1.8) 1.8 Mean 9.40 (9.5) 489.0 (34.2) 60.0 (41.5) 64.0 (44.5) Correlation 0.08 0.78 0.82 0.80 coefficient (r) LSD (0.05); temperature (T) = 3.55; observation (O) = 3.55; temp. x observation (TxO) = 7.11 Figures in the parentheses are angular transformed values after adding 0.1 aArea under disease progression curve

Fig. 9.6 Progression of 100 downy mildew D ×+ ×+ (Hyaloperonospora ×+ + + I [60.54] parasitica) of mustard S + E (Brassica juncea) in A ×+ relation to temperature S

(AUDPC) (Mehta et al. E 1995) I N [45.86] C [14.89] I D 10 E N C +

E [43.83] L O G [1.81] × ××

1 24 48 72 96 PER CENT DISEASE INCIDENCE RECORDED AFTER INOCULATION (h) 10° C + 15° C ×+ 20° C 25° C × 30° C 190 9 Epidemiology and Forecasting

Table 9.2 Effect of leaf wetness duration on infection by Hyaloperonospora parasitica and disease development on mustard seedlings (cv. RH-30) at 20 °C (Mehta et al. 1995) Percent disease incidence after inoculation (h) Leaf wetness duration (h) 24 48 72 96 Mean 0 0.0 (1.8) 0.0 (1.8) 0.0 (1.8) 7.1 (15.4) 1.8 (5.2) 2 0.0 (1.8) 0.0 (1.8) 5.2 (13.3) 8.5 (16.6) 3.4 (8.4) 4 0.0 (1.8) 3.1 (8.9) 7.8 (16.1) 29.0 (32.5) 10.0 (14.8) 6 0.0 (1.8) 6.8 (15.1) 22.1 (27.9) 29.4 (32.6) 14.6 (19.3) 8 0.0 (1.8) 21.5 (27.0) 35.8 (36.0) 35.3 (38.7) 24.2 (25.8) 10 0.0 (1.8) 24.6 (29.7) 45.4 (42.4) 44.4 (41.9) 28.6 (28.9) 12 0.0 (1.8) 18.6 (25.3) 53.0 (46.8) 62.3 (52.2) 33.5 (34.1) 14 0.0 (1.8) 25.3 (30.2) 56.5 (48.8) 61.9 (52.0) 35.9 (33.2) 16 0.0 (1.8) 27.9 (31.8) 56.5 (48.8) 62.5 (52.3) 36.7 (33.7) 18 5.0 (8.8) 26.8 (31.2) 59.9 (50.8) 68.3 (55.9) 40.0 (36.7) 20 9.2 (15.2) 34.4 (35.5) 61.6 (51.8) 68.6 (56.1) 43.4 (39.6) 22 26.3 (30.4) 46.1 (42.8) 61.7 (58.0) 69.2 (56.0) 50.8 (45.3) 24 33.0 (35.0) 53.2 (46.9) 67.8 (56.0) 72.8 (58.8) 56.7 (49.2) Mean 5.6 (8.12) 22.2 (25.2) 41.0 (37.9) 47.9 (48.1) 29.2 (28.6) Correlation coefficient (r) 0.72 0.95 0.95 0.94 LSD (0.05); observation (O) = 2.54; leaf wetness duration (W) = 4.59; observation x leaf wetness (O x W) = 9.19 Figures in the parentheses are angular transformed values after adding 0.1

Table 9.3 Effect of leaf wetness duration on infection by Hyaloperonospora parasitica and disease development on mustard seedlings (cv. RH-30) at 15 °C (Mehta et al. 1995) Percent disease incidence after inoculation (h) Leaf wetness duration (h) 24 48 72 96 Mean 0 0.0 (1.8) 0.0 (1.8) 0.0 (1.8) 0.0 (1.8) 0.0 (1.8) 2 0.0 (1.8) 0.0 (1.8) 0.0 (1.8) 3.8 (9.8) 0.9 (3.8) 4 0.0 (1.8) 0.0 (1.8) 0.0 (1.8) 10.0 (18.2) 2.5 (5.9) 6 0.0 (1.8) 1.7 (5.6) 1.7 (5.6) 20.8 (26.9) 8.5 (10.0) 8 0.0 (1.8) 34.0 (35.7) 47.1 (43.4) 47.1 (43.4) 32.1 (31.1) 10 0.0 (1.8) 49.3 (44.7) 56.4 (48.7) 56.4 (48.7) 40.513 (36.0) 12 0. 0 (1.8) 46.8 (43.3) 58.1 (49.7) 58.5 (50.0) 40.5 (36.2) 14 0.0 (1.8) 53.8 (47.2) 62.7 (52.4) 62.8 (52.5) 44.8 (38.0) 16 0.0 (1.8) 54.0 (47.4) 63.3 (52.8) 63.3 (52.8) 45.2 (38.7) 18 0.0 (1.8) 57.9 (49.6) 64.6 (53.5) 67.0 (54.9) 47.4 (39.5) 20 0.0 (1.8) 65.8 (54.3) 67.5 (55.3) 67.5 (55.3) 50.2 (41.7) 22 0.0 (1.8) 67.5 (55.3) 68.1 (55.7) 68.9 (56.2) 51.1 (42.3) 24 0.0 (1.8) 70.1 (57.1) 70.1 (57.1) 70.1 (57.1) 52.6 (46.5) Mean 0.0 (1.8) 38.5 (34.3) 43.0 (38.5) 45.9 (42.3) 31.9 (29.2) Correlation coefficient (r) 0.00 0.91 0.91 0.93 LSD (0.05); observation (O) = 1.38; leaf wetness duration (W) = 2.49; observation × leaf wetness (O × W) =4.99 Figures in the parentheses are angular transformed values after adding 0.1 100

+× +×+× +× +×+× +× +× + + D +× + I + + + + S +× + E + A S +×

+ E +× I N 10 C + I D E N C

E L O G +× +

1 0612 18 24 LEAF WETNESS DURATION (H) 24 h4+ 8 h7+× 2 h96 h

Fig. 9.7 Effect of leaf wetness duration on the development of downy mildew (Hyaloperonospora parasitica) infection on mustard (Brassica juncea) cultivar RH-30 at 20 °C (Mehta et al. 1995)

100

+× +× +× +× +× +× +× + + +× +++ D +× + + I + S E A S

E I N C 10 I D E N +× C

E L O G +×+×

1 0612 18 24 LEAF WETNESS DURATION (H) 24 h4+ 8 h7+× 2 h96 h

Fig. 9.8 Effect of leaf wetness duration on the development of downy mildew (Hyaloperonospora parasitica) on mustard (Brassica juncea) seedlings of cultivar RH-30 at 15 °C (Mehta et al. 1995) 192 9 Epidemiology and Forecasting

Table 9.4 Prediction equations for the progress of downy mildew and white rust complex of rapeseed-mustard using different combinations of weather factors (Kolte et al. 1986)

2* Equations bO X1 X2 X3 X4 X5 X6 R Stag head +16.925 +0.019 −0.132 −0.086 +0.158 +0.030 −1.469 0.6849 incidence (Y1) (%) Stag head severity +86.169 −1.241 −0.129 −0.503 +0.054 +0.472 −2.125 0.6283 (Y2) (%)

X1 = mean maximum temperature X2 = mean minimum temperature

X3 = mean relative humidity X4 = total rainfall (mm)

X5 = total rainy days X6 = mean bright sunshine period (h/day) *Significant at 5% level

9.3 Disease Development in Relation to Planting Time

In India, infection of mustard foliage starts by the end of October (cotyledon stage) and progresses up to November (Tables 9.7 and 9.8). The severity of downy mildew and white rust disease complex in relation to planting times has been studied (Saharan 1984) at three different locations, viz., Hisar, Kanpur, and Pantnagar. The results revealed that disease complex intensity increases with the delay in date of planting from October to December during 1978 at Hisar. Similar trend was also observed at Kanpur and Pantnagar (Table 9.7). It is evident from the observations (Table 9.8) that staghead incidence and severity was maximum on yellow sarson and toria during 1977–1978, whereas during 1978–1979 and 1979–1980, it was not consistent. It indicates that probably environmental conditions were not favourable in these years. The crop planted after mid-November may not contract downy mildew. However, downy mildew growth as a mixed infection with white rust on floral parts can be seen up to March (Saharan 1984; Kolte et al. 1986; Mehta 1993). In Lithuania, the severity of downy mildew of winter oilseed rape was higher when sown in the beginning of August in comparison to end of August sown crop (Petraitiene and Brazauskiene 2005).

9.4 Disease Development in Relation to Host Nutrition

Hyaloperonospora parasitica is severe on cauliflower plants which suffer from potash deficiency, while plants with a sufficient quantity of potash are only slightly attacked (Quanjer 1928). Cabbage plants grown in soil fertilized with less potash and more phosphorus are more prone to downy mildew than cabbages grown in unfertilized soil (Townsend 1935). However, according to Butler and Jones (1949), there is no consistent effect of fertilizers on the development of downy mildew of Brassicas. Felton and Walker (1946) found no direct relationship between mildew incidence and any excess or deficiency of nitrogen, phosphorus, or potash. On 9.4 Disease Development in Relation to Host Nutrition 193

Fig. 9.9 Weather factors associated with occurrence (A) and no occurrence (B) periods of stag head phase of white rust (Albugo candida) and downy mildew (Hyaloperonospora parasitica) on mustard (Brassica juncea) in crop seasons Y1 (1976–1977), Y2 (1977–1978), Y3 (1978–1979), Y4 (1979–1980), Y5 (1980–1981), Y6 (1981–1982), and Y7 (1982–1983). Symbol Y represents the number of crop seasons covering the period from 1977–1978 to 1982–1983 under no occurrence periods of stag heads (B) (Kolte et al. 1986) radishes, tubers, conidiophores, and conidia appear to be relatively large, which is probably due to the availability of ample nutrient supply in the tubers (Hammarlund 1931). The incidence and severity of downy mildew on lower leaves of winter oilseed rape was higher in N-treated plots, compared to N-untreated plots under 194 9 Epidemiology and Forecasting 0.71 1.71 0.23 1.12 1.85 15.36 6.52 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Increase 27.50 26.79 25.08 24.85 23.73 21.88 6.52 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Alternaria blight PDI Accumulated 0.07 1.84 1.55 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Increase 2.46 2.39 1.55 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Downy mildew PDI mildew Downy Accumulated 4.24 0.28 0.67 12.80 3.05 3.20 1.14 4.75 15.77 0.00 0.00 0.00 0.00 0.00 Increase 45.88 41.64 41.36 40.71 27.91 24.86 21.66 20.52 15.77 0.00 0.00 0.00 0.00 0.00 Accumulated White rust PDI 0.00 0.00 0.00 1.11 0.00 0.00 0.00 0.00 0.00 0.00 0.00 1.38 0.71 0.00 Rainfall Rainfall (mm) 36.00 43.14 38.42 43.57 42.71 27.71 35.57 36.71 43.71 49.57 51.14 57.57 64.28 60.71 II 80.28 69.14 79.85 83.57 84.42 81.57 86.85 83.28 92.00 90.00 88.57 94.00 87.28 85.00 I Relative humidity (%) Relative C 0 15.18 15.54 14.97 16.97 13.20 11.77 15.37 14.40 13.12 14.40 13.90 16.08 16.01 15.91 Min. 31.41 29.25 29.42 28.51 28.60 27.80 26.54 26.67 26.30 26.48 26.50 23.11 26.02 27.20 Max. Temperature Effect of weather factors on development and severity of white rust, downy mildew, and Alternaria blight (Sangeetha Siddaramaiah 2007 ) mildew, of white rust, downy and severity on development of weather factors Effect 14 13 12 11 10 9 8 7 6 5 4 3 2 1 Observations at Observations 7-day intervals Max = maximum; Min = minimum; I = morning; II = evening Table 9.5 Table 9.4 Disease Development in Relation to Host Nutrition 195

Table 9.6 Correlation value between disease index of white rust, downy mildew, and Alternaria blight of Indian mustard with environmental factors (Sangeetha and Siddaramaiah 2007) Correlation coefficient ‘R’ value Weather factors White rust Downy mildew Alternaria blight Maximum temperature +0.8245** +0.7211** +0.8263** Minimum temperature −0.41139 −0.1853 −0.0340 Relative humidity I −0.3252 −0.2009 0.0983 Relative humidity II −0.4645 −0.3156 −0.6147 Rainfall −0.1723 −0.2548 −0.1052 **Indicate positive and significant correlation between maximum temperature and diseases index

Table 9.7 Effect of planting time on the severity of downy mildew and white rust complex of mustard (Saharan 1984) Percent disease intensity Planting time Hisar Kanpur Pantnagar 06.10.1978 10.0 – – 21.10.1978 8.6 – – 28.10.1978 18.6 – – 06.11.1978 55.4 – – 18.11.1978 68.5 – – 02.12.1978 72.8 – – 01.10.1979 – 24.16 – 10.10.1979 4.6 28.30 – 20.10.1979 10.0 34.34 – 30.10.1979 22.5 36.18 – 09.11.1979 46.8 40.91 – 20.11.1979 57.5 46.15 – 03.10.1981 – – 15.04 23.10.1981 – – 19.85 13.10.1981 – – 32.85

Lithuania conditions (Petraitiene and Brazauskiene 2005). The increase in the N fertilization rate resulted in the increase in infection of downy mildew of oilseed rape under Germany conditions (Sochting and Verret 2004). Jiang and Caldwell (2015) found positive correlation of downy mildew infection in Camelina with applied N rates. 196 9 Epidemiology and Forecasting

Table 9.8 Influence of planting dates on stag head incidence and severity of white rust and downy mildew of rapeseed and mustard in three rabi crop seasons starting from 1977–1978 to 1979–1980 (Kolte et al. 1986) Date of 1977–1978 1978–1979 1979–1980 planting Mustard Yellow Toria Mustard Yellow Toria Mustard Yellow Toria sarson sarson sarson Oct. 1–6 2.7* 0.0 0.0 20.6 24.3 10.2 8.9 4.8 0.7 (11.0)** (0.0) (0.0) (15.6) (23.1) (20.3) (20.3) (15.7) (4.4) Oct. 11–14 6.7 1.2 0.0 14.6 23.7 7.7 14.2 8.3 1.8 (15.6) (8.4) (0.0) (20.4) (24.0) (21.8) (17.7) (32.4) (6.4) Oct. 20–22 10.2 4.1 0.0 10.6 17.2 18.2 7.8 3.6 3.6 (23.4) (24.8) (0.0) (11.3) (14.9) (16.8) (14.6) (17.7) (16.9) Oct. 10.2 8.2 10.2 7.8 (9.7) 11.6 24.9 5.3 11.2 9.2 31–Nov. 1 (20.9) (35.2) (23.9) (17.2) (13.5) (11.5) (43.4) (22.4) Nov. 1–11 9.3 9.7 10.6 9.6 11.2 9.4 3.1 (5.2) 6.7 18.2 (22.4) (25.8) (25.3) (11.1) (16.5) (14.4) (16.3) (34.7) Nov. 14.2 38.2 22.2 9.1 13.7 8.9 4.3 (8.5) 1.4 15.9 20–22 (22.8) (25.7) (24.0) (10.9) (19.0) (15.3) (8.8) (23.2) * = Incidence (%plant affected); ** = Figures in parentheses are disease severity (% racemes affected/plant) CD at 5% 1977–1978 1978–1979 1979–1980 for planting dates Incidence 5.6 2.8 3.6 Severity 5.5 1.7 9.8 Incidence NS 1.8 2.6 Severity 3.4 1.3 4.2

9.5 Disease Interaction with Insecticidal Sprays

The incidence of downy mildew in plots of broccoli sprayed with emulsifiable insecticide formulations containing a solvent and a wetting agent is significantly greater than in plots sprayed with an insecticide formulation containing no solvent or wetting agent or in unsprayed plots (Natti et al. 1956). It is possible that emulsifiable insecticide formulations remove the bloom from the leaves and dissolve the wax from the cuticle of the leaves creating conditions favourable for the germination of H. parasitica spores (Natti et al. 1956).

9.6 Disease Prediction Models

Kolte et al. (1986) developed prediction equations for the stag head severity in relation to planting dates and associated weather factors as under: References 197

Stag head incidenceY()%.=+16 925 0..019XX12−−0 132 0..086X3 + 0 158X4 +−0..030XX1 469 −−R2 06. 8 56

Stag headsSeverity()%.YX=−86 169 1..241 12−−0 129XX0.503 3 +0.054XXX+−0..472 2 125XR−−2 06. 2 456 where X1 = mean max. Temp, X2 = mean min. Temp, X3 = mean RH, X4 = total rain fall (mm), X5 = total rainy days; and X6 = mean bright sunshine period (h/day). Mehta and Saharan (1998) developed the prediction models for the progression of downy mildew of rapeseed-mustard as under:

2 A. Leaf infection: Y = −32.7 + 0.09 X1 + 0.31 X2 + 1.31 X3 + 0.12 X4 + 0.22 (R = 0.36) X5 − 0.03 X6 B. Stag head: 2 i. Incidence: Y = −18.6-2.8 X1 + 2.5 X2 + 4.5 X3 + 0.6 X4 − 0.2 (R = 0.23) X5 + 1.0 X6 2 ii. Length: Y = −17.4-1.5 X1 + 1.5 X2 + 2.6 X3 + 0.4 X4 − 0.1 X5 − 1.0 (R = 0.26) X6

where X1 = Tmax., X2 = Tmin., X3 = Sunshine, X4 = RHmor, X5 = RHeve, and X6 = RF

References

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Jiang Y, Caldwell CD (2015) Effect of nitrogen fertilization on Camelina seed yield, yield components, and downy mildew infection. Can J Plant Sci 96:17–26 Jonsson R (1966) Peronospora on oil yielding Brassicas. Methods for testing resistance in winter rape and their results. Sver Utsadestor Tidskr 76:54–62 Kolte SJ, Awasthi RP, Vishwanath (1986) Effect of planting dates and associated weather factors on staghead phase of white rust and downy mildew of rapeseed and mustard. Indian J Mycol Plant Pathol 16:94–102 Mehta N (1993) Epidemiology of white rust and downy mildew disease complex in mustard and residual toxicity of fungitoxicant. Ph.D. Thesis, CCS. Haryana Agricultural University, Hisar, Haryana, India: 154p Mehta N, Saharan GS (1998) Effect of planting dates on infection and development of white rust and downy mildew disease complex in mustard. J Mycol Plant Pathol 28:259–265 Mehta N, Saharan GS, Sharma OP (1995) Influence of temperature and free moisture on the infection and development of downy mildew on mustard. Plant Dis Res 10:114–121 Nakov B (1972) The effect of ecological factors on the dissemination of Peronospora parasitica (Pers.) Fr. on cabbage. Nauch Trudov Vissh Selskostop Inst V Kolarov; Plovdiv 21:109–116 Natti JJ, Harvey GER, Sayre CB (1956) Factors contributing to the increase of downy mildew of broccoli in New York State and its control with fungicides and agrimycin. Plant Dis Report 40:118–124 Petraitiene E, Brazauskiene I (2005) Incidence and severity of Alternaria blight (Alternaria spp.) and downy mildew (Peronospora parasitica) as affected by winter oilseed rape sowing time and nitrogen fertilizer rate. Agronomijas Vestis (Latvian J Agro) 8:158–162 Quanjer HM (1928) The influence of potash deficiency on the susceptibility of cauliflower to Peronospora parasitica. Tijdschr, Over Plantenziekten 34:254–256 Saharan GS (1984) A review of research on rapeseed mustard pathology in India. Annual Workshop AICORPO ICAR, Jaipur, 6–10 August 1984 Saharan GS, Verma PR, Nashaat NI (1997) Monograph of downy mildew of crucifers. Saskatoon Res Cent Tech Bull, Agriculture and Agri-Food Canada, 1997–01, 197 pp Sangeetha CG, Siddaramaiah AL (2007) Epidemiological studies of white rust, downy mildew and Alternaria blight of Indian mustard [Brassica juncea (Linn.) Czern & Coss.] African J Agric Res 2(7):305–308 Sinobas Alonso J, Diaz Alonso M (1995) The mildew of the crucifers in the term of villa del Prado (Madrid): epidemiological notes. Boletín de Sanidad Vegetal Plagas 21:497–506 Sochting HP, Verret JA (2004) Effects of cultivation systems: soil management, nitrogen fertilization on the epidemics of fungal diseases in oil seed rape (Brassica napus L. Var. napus). J Plant Dis Protect 111:1–29 Townsend GR (1935) Everglades Experimental Station. Report Florida Expt Stn 1933-34, pp. 86–112 Vladimirskaya ME, Ilyina MH, Klinkovskaya IK (1975) Forecasting disease incidence on cabbage crops as influenced with their cultivation practices. Mikol Fitopatol 9:130–132 Williams PH, Leung H (1981) Methods of breeding for multiple disease resistant Chinese cabbage. In: Talekar NS, Griggs TD (eds) Chinese cabbage. Proceedings of the 1st International Symposium. The Asian Vegetable Research and Development Center, Shanhua, Taiwan, pp 393–403 Chapter 10 Association or Mixed Infection of Downy Mildew and White Rust Disease Complex

10.1 Introduction

The association of downy mildew (DM and white rust (WR) infection on oilseed Brassica, vegetable Brassica, wallflowers, and stocks has been observed (Butler 1918; Wiese 1927). On horseradish leaves and petioles, A. candida (white rust) and H. parasitica (downy mildew) are frequently associated with each other causing brown rot commencing at the head of the rootstock and extending downwards (Boning 1936). The association or mixed infections, or DM-WR disease complex, or simultaneous occurrence of A. candida and Hyaloperonospora parasitica (mustard mosaic and downy mildew on leaves) on leaves, stems, inflorescence, and siliquae of crucifers in nature is very common (Saharan and Verma 1992; Saharan et al. 1997, 2005, 2014; Saharan 2010. White rust pathogen, A. candida, is frequently associated with the DM pathogen H. parasitica. Numerous instances are known of considerable damage from combined infection (Magnus 1894; Butler 1918; Ocfemia 1925; Savulescu and Rayss 1930; Butler and Jones 1949, 1961; Vanterpool 1958; Sansome and Sansome 1974). In India, mixed infections of DM and WR are common on B. juncea (Bains and Jhooty 1985; Saharan 1984). A. candida appears first on leaves, which probably predisposes the host to infection by H. parasitica (Bains and Jhooty 1985). The intensity of mixed infections varies from 0.5 to 35%, depending on local weather conditions (Bains and Jhooty 1979; Saharan 1984). Incidence and severity of mixed infections by A. candida and H. parasitica on B. juncea inflorescence are higher on detopped than on normal plants (Bains 1989). Severity of mixed infections on leaves is not related to infections on inflorescence. It seems that greater susceptibility of new inflorescence and their availability over extended periods of time are associated with this phenomenon (Bains and Sokhi 1986). Hyaloperonospora parasitica also causes severe infections and high levels of sporulation on plants of mustard systematically infected with mustard mosaic virus (Bains and Jhooty 1978). The hypertrophied and malformed inflorescences ofB. juncea infected with

© Springer Nature Singapore Pte Ltd. 2017 199 G. S. Saharan et al., Downy Mildew Disease of Crucifers: Biology, Ecology and Disease Management, https://doi.org/10.1007/978-981-10-7500-1_10 200 10 Association or Mixed Infection of Downy Mildew and White Rust Disease Complex

A. candida are usually covered heavily with the white powdery growth of the DM spores consisting of conidia and conidiophores (Chaurasia et al. 1982; Saharan 1984, 1992a, b). Albugo candida alone, on artificial inoculation of flower buds, induces typical hypertrophy of the inflorescence (Verma and Petrie 1980). However, whether H. parasitica alone induces hypertrophy of the inflorescence in B. juncea or other hosts remains to be confirmed. The role(s) played by A. candida and H. parasitica in induction of mixed infections of inflorescences in the field is also yet to be determined. Under field conditions, A. candida is reported to elevate both incidence and severity of infection by H. parasitica in crucifers (Constantinescu and Fatehi 2002). A similar situation has been described for H. arabidopsidis in A. thaliana (Holub et al. 1991) and B. juncea (Cooper et al. 2008) after pre-inoculation with A. candida. After simultaneous co-inoculation of B. juncea plants, Singh et al. (2002) showed that while infection with a virulent isolate of H. parasitica inhibited or adversely affected the development of a virulent isolate of A. candida, an avirulent isolate of A. candida induced host resistance to H. parasitica. Over 20 species of fungi, including several pathogens of crucifers, have been reported in association with A. candida which produced hypertrophies of the inflorescence (stag heads), stem, and pod blisters on turnip rape (B. rapa), wild mustard (B. kaber), and false flax (Camelina microcarpa). The most prevalent fungal associates of Albugo are H. parasitica, Alternaria alternata, Fusarium roseum, F. acuminatum, F. equiseti, Alternaria raphani, A. brassicae, and Cladosporium sp. (Petrie 1986). Alternaria, Cladosporium, and Fusarium spp. reported seem to be common saprophytes under humid conditions. Kaur et al. (2011a, b) reported that the presence of asymptomatic systemic colonization of H. parasitica in a resistant host increases susceptibility to WR disease.

10.2 Symptoms

Symptoms of mixed infection of DM and WR diseases of Brassicas can be seen at both stages, i.e. leaf phase and stag head phase of plant growth. On leaves, WR pustules are surrounded by white or creamy white sporulation of DM pathogen (Plate 10.1). On the malformed floral parts,A. candida produces pustules, and H. parasitica produces a coating of fine white mass of sporangia or conidia. Conidia initially appear at the tip of the stag head, later on covered the entire inflorescence (Plates 10.2 and 10.3). At crop maturity, and under high humidity, malformed inflorescence may be covered with saprophytes turning into dark brown to black colour (Saharan 1992a, b; Saharan and Mehta 2002). 10.3 Yield Losses 201

Plate 10.1 White rust pustules are surrounded by conidial mass of Hyaloperonospora

Plate 10.2 White rust-infected stag head with initial growth of downy mildew

10.3 Yield Losses

Mixed infection of DM and WR causes loss of 17–32% in B. juncea under Punjab conditions (Saharan et al. 1997; Bains and Jhooty 1979). In B. rapa var. toria, yield loss of 34% has been estimated if the average length of hypertrophied raceme is 10 cm (Kolte et al. 1981). Lakra and Saharan (1989) observed that combined infection of WR and DM may cause yield loss up to 89.8% during epidemic years. According to Meena et al. (2014), in B. juncea, DM infected plants reduced yield by 42.6, 46.8, and 66.7% in cvs. Rohini, Varuna, and NRCDR-2, respectively. DM infection significantly reduced average plant height, 1000 seed weight, and oil content of seeds. The number of primary branches per plant, however, was significantly higher in infected than in healthy plants. Generally, the number of silique on main shoot, as well as on primary branches, was also significantly lower in 202 10 Association or Mixed Infection of Downy Mildew and White Rust Disease Complex

Plate 10.3 Conidial growth of Hyaloperonospora covered the white rust pustules in stag head

infected than in healthy plants. Quantification of economic impact showed that the yearly expected loss in India due to DM was about 683.1 million Indian rupees, depending upon the disease severity (Tables 10.1 and 10.2).

10.4 Pathogenesis

The nature of association of mixed infection, or simultaneously occurrence, or DM-WR disease complex seems to be synergistic, since the magnitude of effect on host plant is increased manifold. The sequence of events during the pathogenesis of Hyaloperonospora and Albugo in Indian mustard cv. RH-30 has been explained by Mehta et al. (1995) (Table 10.3). In the treatment where Hyaloperonospora was inoculated 24 h prior to Albugo (HP-AC), the infection by Hyaloperonospora was recorded after 7 days and by Albugo after 5–6 days. Similarly, in reverse treatment, i.e. AC-HP, the incubation period of Hyaloperonospora and Albugo was remaining the same. Likewise, the incubation period was the same when both pathogens were inoculated separately. But in the treatment where both the pathogens were inoculated together (HP + AC), there was a delay in incubation period of Hyaloperonospora. However, Albugo did not show variation in its incubation period. The latent period of each pathogen was recorded 12 days in all the treatments irrespective of their 10.4 Pathogenesis 203 NS NS NS NS NS 3.4 5.3 69.2*** − 1.8 − 6.1 137.9*** − 10.6*** − 66.7*** − 11.9* − 12.8 − 30.7** − 32.5** − 52.9*** – Difference (%) Difference 1.7 4.0 8.8 37.2 66.7 31.4 53.6 42.0 23.2 37.0 27.0 15.4 11.4 139.8 Infected 5.1 3.8 5.2 41.6 13.2 54.6 40.6 26.6 53.4 40.0 16.4 24.2 158.6 – Cultivar NRCDR-2 Cultivar Healthy NS NS NS NS NS NS NS NS NS NS 4.2 53.1* 60.0 − 8.8 − 8.2*** − 5.7 − 5.9 − 1.8 − 4.8 − 2.5 − 3.6 − 42.6*** – Difference (%) Difference − 13.6 3.1 9.8 3.9 57.8 40.1 42.6 44.6 77.4 15.8 24.8 64.0 39.6 24.2 191.0 Infected 5.4 6.4 4.0 63.4 43.7 47.4 78.8 16.6 15.5 66.4 38.0 28.0 202.6 Cultivar Rohini Cultivar Healthy – NS NS NS NS NS – 22.0** 10.3 − 9.9 − 2.6 − 2.1* − 16.1 − 34.4*** − 16.8*** − 80.0** − 32.2*** − 19.8 − 18.1** − 16.9*** Difference (%) Difference 3.8 3.3 46.8 50.4 34.7 41.8 38.0 10.0 30.6 19.0 68.2 27.8 11.8 187.0 Infected 3.9 6.2 8.2 60.1 38.5 63.7 38.8 45.1 23.7 83.3 25.2 14.2 203.3 – Cultivar Varuna Cultivar Healthy ) (Meena juncea of Indian mustard ( Brassica cultivars infection on yield, yield components, and oil contents of different mildew of downy Effect % yield reduction Yield components Yield Length of primary branch (cm) Length of secondary branch (cm) Length of siliquae (cm) No. of siliquae on main shoot 1000 seed weight (g) % oil content Plant height (cm) No. of primary branches/plant No. of siliquae on primary branch No. of siliquae on secondary branch Length of main shoot(cm) No. of secondary branches/plant No. of seeds/siliquae NS = not significant. The asterisk indicates that the difference in main values is significant at 0.10(*), 0.05 (**), and 0.01 (***) percent level of significance. * level values is significant at 0.10(*), 0.05 (**), and 0.01 (***) percent in main The asterisk indicates that the difference significant. NS = not and infected plants in 1000 seed weight between healthy percent yield loss based on difference Table 10.1 Table et al. 2014 ) 204 10 Association or Mixed Infection of Downy Mildew and White Rust Disease Complex

Table 10.2 Estimation of Parameters Value economic loss due to downy Average actual yield loss (kg/ha) 143.2 mildew in Brassica juncea Area affected by downy mildew 11.0 during 2010–2011 crop a season in India (Meena et al. infection (%) 2014) Probability of disease occurrencea 0.15 Total area under mustard (m/ha)b 5.2 Price of output (INR/kg)c 18.5 Expected yield loss (mt) 0.75 Expected value of damage due to 683.1 disease (million INR) Loss of edible oil at 42% oil 0.31 recovery rate (mt) aCalculated from primary survey data bCalculated as 80% of the total area under rapeseed-mustard in India during 2010–2011 (GOI 2011) cThe price of output was taken as the minimum support price announced for the produce by government

Table 10.3 Interaction between Albugo candida and Hyaloperonospora parasitica during pathogenesis of B. juncea (Mehta et al. 1995) Incubation period (days) Latent period (days) Treatment HP AC HP AC HP-AC(24 h) 7 5–6 12 12 HP alone 7 − 12 − AC-HP (24 h) 7 5–6 12 12 AC alone − 5–6 − 12 HP + AC 9–10 5–6 12 12 Hp, Hyaloperonospora parasitica; Ac, Albugo candida time of inoculations (Table 10.3). The results showed that when both pathogens were inoculated together, there was a delay in incubation period of Hyaloperonospora for 2–3 days only.

10.5 Histopathology

In histopathological studies, the transverse sections of inoculated leaves collected at different intervals revealed that the epidermal layer was broken at many places. Generally, there was an overall deformation in the size and shape of mesophyll cells in the inoculated leaves compared to the sections of the healthy leaves. The ana- tomical examinations of leaves collected 24 h after inoculation with either of the pathogen or combination of both did not show any penetration; germinated conidia 10.5 Histopathology 205 or sporangia were seen on the leaf surface (Mehta et al. 1995). The samples collected after 2 days of inoculation indicated that the initiation of infection confined mainly to epidermis. The pathogen penetrated up to 1/3 of mesophyll cells in samples obtained 3 days after inoculation (Plate 10.4 (1); 6 days after inoculation, pathogen invaded deeper into the tissue. In the treatment where H. parasitica was inoculated prior to or after Albugo, mycelium could be seen in the intercellular spaces with globose to knoblike haustoria in the mesophyll cells (Plate 10.4 (2). Similarly, in the treatments where Albugo was inoculated alone or in combination with H. parasitica, the pathogen penetrated up to lower epidermis, and there was development of pustules on the lower epidermis (Plate 10.4 (3). Similar interaction was observed with H. parasitica showing necrosis in the mesophyll cells. However, when both the pathogens were inoculated together (HP + AC), the infection was confined to the upper layer of mesophylls, and there was a limited colonization of the cells, with a limited number of haustoria or mycelium in the intercellular spaces. Leaves collected after 6 days of inoculation with either of the pathogen inoculated alone or in combination showed characteristic disease symptoms (Mehta et al. 1995). The Albugo pustules showed hyaline sporangiophores bearing globose to oval-shaped sporangia in chain (Plate 10.4 (4). The H. parasitica mycelium was intercellular with lobe-shaped haustoria in the distorted tissues of leaves (Plate 10.4 (5). When both the pathogens were inoculated together, infection was extended to mesophyll cells, and there was a development of pustules below the epidermis (Plate 10.4 (6). The leaves samples collected 12 days after inoculation in all the treatments showed complete colonization of the pathogen as evident from the development of necrotic zone by H. parasitica and bursting of pustules in case of Albugo releasing sporangia. The sections of diseased inflorescence showed deep colonization of H. parasitica. The mycelium passed through the epidermis, hypodermis, cortex, and finally reached the pith region. The fungal mycelium was in the cortex and produced conidiophores bearing conidia above the epidermis layer. Similarly, where Albugo was inoculated, numerous sporangiophores bearing sporangia were observed below the epidermis layer in the form of pustules and knob. In the samples where both the pathogens were inoculated together, deep colonization of the tissues was observed, but the individual pathogen could not be distinguished in the host tissues. These studies revealed that the pathogens could invade deeper into the tissues intercellularly and Albugo had faster growth in terms of incubation period than the H. parasitica. In colonized tissues, both the pathogens could not be distinguished on the basis of their somatic morphology (Mehta et al. 1995). The incubation period of H. parasitica was 7 days in all the treatments, except in case where both the pathogens were inoculated together. The incubation period of A. candida was 5–6 days in all the treatments. These results indicate that Albugo colonizes the host tissue earlier than Hyaloperonospora. The DM colonies appear on or around the WR pustules on leaves and malformed inflorescence under field conditions (Plates10.1 , 10.2 and 10.3). The Albugo mycelium while developing in the intercellular spaces, and by disturbing the metabolism of the cell, may create congenial situation for Hyaloperonospora to colonize at later stages. This could be a possible reason why incubation period was delayed in case Plate 10.4 (a) Transverse section of leaf 3 days post-inoculation AC-HP or HP-AC showing mycelium in the intercellular spaces. UEP upper epidermis layer, LEP lower epidermis layer, F fungus GMS × 66 × 8 approx. (Mehta et al. 1995). (b) Transverse section of mustard leaf 6 days post-inoculation (AC-HP) showing mycelium in the intercellular spaces. UEP upper epidermis layer, LEP lower epidermis layer, F fungus GMS × 66 × 8 approx. (c) Transverse section of mustard leaf 6 days post-inoculation (AC alone) showing mycelium in the intercellular spaces and developing white rust pustules. UEP, upper epidermis layer; F fungus, M mesophyll cells, WRP white rust pustules GMS × 66 × 8 approx. (d) Transverse section of mustard leaf 9 days post- inoculation (AC alone) showing fungal mycelium and white rust pustules with sporangia/sporangiophores on abaxial surface. UEP upper epidermis layer, F fungal mycelium, M mesophyll cells, SP sporangia/sporangiophores GMS × 66 × 8 approx. (e) Transverse section of DM-infected mustard inflorescence depicting fungal mycelium and haustoria in the cortical cells and pith. EP epidermis layer, F fungus, CR cortex cells, X xylary vessels, Ph phloem, P pith, HA haustoria; GMS × 66 × 8 approx. (f) Transverse section of white rust-infected mustard inflorescence showing fungal mycelium, haustoria, and oospores in the cortical, xylary vessels. OS oospores, P pith, HA haustoria, F fungus; GMS × 66 × 8 10.5 Histopathology 207 of H. parasitica where mixed inoculations were done. Another possibility could be the competition between the two pathogens for the same site of infection, and H. parasitica may get limited site for infection and development. But once the fungus penetrated, and developed, there was no delay in the production of asexual spores, since the same latent period was recorded in all the treatments (Mehta et al. 1995). Liu and Rimmer (1990) reported 5–6 days of incubation period for Albugo, but contrarily, H. parasitica has been reported to colonize the cotyledonary leaves within 4 days of inoculation (Kluczewski and Lucas 1982). The longer incubation period in case of H. parasitica may be attributed to the fact that inoculation were made on true leaves rather than the cotyledonary leaves. In addition, the weather parameters prevailing at the time of infection may affect the time required for infection. The latent period of 12 days was recorded in all the treatments when inoculated with both the pathogens with different combinations. Kolte (1985) observed a period of 10 days for release of sporangia in case of Albugo under Pantnagar conditions. The slight variation in latent period in the investigations by Mehta et al. (1995) may be due to host influence and effect of micro- and macroclimate prevailing around the plant (Bains and Jhooty 1985). The sections of the leaves collected at different intervals after inoculation with either of the pathogen or in combination of both revealed that penetration of the host started 24 h after inoculation, and complete infection and colonization took place by the third day. Greenhalgh and Dickinson (1975) showed that sporangia penetrated the cotyledonary leaves of crucifers within 24 h of inoculation. Longer incubation period reported by Mehta et al. (1995) may be attributed to the fact that inoculations were made on the true leaves. However, Kluczewski and Lucas (1982) reported colonization of H. parasitica on oilseed rape (B. napus) within 4 days of inoculation on cotyledonary leaves, whereas mycelium development continued up to 5th day accompanied by abundant sporulation. Mehta et al. (1995) reported that sections of leaves obtained after 6 days of inoculations also show production of conidiophores and conidia. The development of necrosis and complete infection of the inoculated leaves after 9 and 12 days of inoculation are in accordance with the findings of Kluczewski and Lucas (1982). In the case of Albugo, the sori (pustules) development was started 3 days after inoculation. According to Verma et al. (1975), Albugo produced the haustorium in B. juncea cotyledonary leaves after 16–18 h of inoculation, and the mycelium occupied the intercellular spaces within 3 days of inoculation. After 4 days, almost all the intercellular spaces of the inoculated tissues were occupied by the mycelia, and WR pustules were visible 5–6 days after inoculation (Liu et al. 1989). Similar observations have also been made by Mehta et al. (1995). However, according to Kaur et al. (2011b, c), pre-inoculation with Hyaloperonospora reduces incubation period and increases severity of WR disease in B. juncea variety resistant to DM. White rust symptoms appear 4 days earlier and are more severe when DM resistant but highly WR-susceptible B. juncea variety is first inoculated withAlbugo followed 10 days later with Hyaloperonospora. DNA extraction of tissues indicated that Hyaloperonospora has colonized the asymptomatic plants infected systemically. Studies of Singh et al. (2002) showed that the infection of B. juncea with a 208 10 Association or Mixed Infection of Downy Mildew and White Rust Disease Complex virulent isolate of H. parasitica inhibited or adversely affected the development of a virulent isolate of A. candida. Under field conditions, A. candida can elevate both incidence and severity of infection by H. parasitica in crucifers (Constantinescu and Fatehi 2002). Cooper et al. (2008) observed broad-spectrum suppression of host defence by A. candida in A. thaliana and B. juncea. It seems that genes governing the virulence of A. candida in Brassica system elude the plant defence mechanism, through reduction of phytoalexin biosynthetic pathway and by producing metabolites preferred as food by the pathogen Hyaloperonospora for colonization of host tissues. The compatibility genes of both the pathogens may be the same or situated on the same locus as alleles or tightly linked or epistatic. However, detailed study is required on these aspects (Saharan 2010).

10.6 Epidemiology

Development of WR and DM disease complex in Indian mustard in relation to planting time and environmental interaction was investigated by Mehta and Saharan (1998). There was no stag head formation in the crops sown during September (Tables 10.4 and 10.5), but significantly higher stag head incidence was observed in crops planted late in October and November. The crops sown in mid-November although had fewer incidences of stag head but their intensity was significantly higher. Stag head length reported comparatively shorter in crops planted in

Table 10.4 Effect of planting dates on the development of stag head due to downy mildew and white rust disease complex in Indian mustard cv. RH-30 during 1991–1992 crop season (Mehta and Saharan 1998) Observations Date of sowing (week) Sept. Sept. Oct. Oct. Dec. Mean 8.a 24.b 19.c 29.d 4.e I L I L I L I L I L I L 1 – – – – 0 0 5 3 3 1 2.5 1.5 2 – – – – 6 2 17 11 3 2 8.8 5.1 3 – – – – 7 2 17 11 5 5 10.0 6.4 4 – – – – 7 2 17 11 6 6 10. 6.6 Mean – – – – 5.2 1.6 14.2 9.5 4.2 3.5 – – I = incidence (%); L = length (cm); − = disease did not appear LSD (0.05) I L Stag head started on Date of sowing (D) 3.81 1.22 a = No. stag head Intervals (I) 4.39 1.41 b = No. stag head D × I 7.62 2.45 c = 14.01.92 d = 14.01.92 e = 04.02.92 10.7 Disease Forecasting 209

Table 10.5 Effect of planting dates on the development of stag head due to downy mildew and white rust disease complex in Indian mustard cv. RH-30 during 1992–1993 crop season (Mehta and Saharan 1998) Observations Date of sowing (week) Sept. Sept. Oct. 15c Nov.5d Nov.18e Dec. 4f Mean 15a 30b I L I L I L I L I L I L I L 1 – – – – 18 7 29 11 16 8 14 14 13.0 6.7 2 – – – – 18 8 29 11 16 8 14 15 13.0 7.1 3 – – – – 18 8 29 11 16 9 14 15 13.0 7.2 4 – – – – 18 9 29 11 16 9 14 15 13.0 7.2 Mean – – – – 18.0 7.8 29.0 11.0 16.0 8.5 14.0 15.0 – – I = incidence (%); L = length (cm); − = disease did not appear LSD (0.05) I L Stag head started on Date of 4.74 2.16 a = No. stag sowing (D) head Intervals (I) NS NS b = No. stag head D × I NS NS c = 31.12.92 d = 31.12.92 e = 14.01.93 f = 04.02.93

December, but it had higher downy mildew incidence (Table 10.4). The maximum stag head incidence was recorded from the first week of January to mid-February for successive 2 years. During the time of inflorescence formation in the September- planted crop, the weather was not favourable for stag head development. However, fewer incidences of stag heads in the December sown crop may be attributed to improper growth of the crop (Table 10.5). Kolte et al. (1986) reported that early- sown crop either escaped or showed least stag head incidence. The late-sown crop, however, had higher incidence and severity of stag heads. High RH and low temperature favoured stag head formation (Lakra et al. 1989; Lakra and Saharan 1990). Bains and Jhooty (1979) found that temperature of 14.30 C and 152 mm rainfall resulted in high infection.

10.7 Disease Forecasting

Correlation between weather parameters and disease progression has revealed that the combination of six independent variables accounted for more than 66% variation in white rust during 1991–1992 and only 16% during 1992–1993, although disease intensity was more (Table 10.6). Similarly, these variables accounted for 21 and 36% variation in DM intensity during 1991–1992 and 1992–1993, respectively. All the test variables influenced floral malformation incidence and intensity to the 210 10 Association or Mixed Infection of Downy Mildew and White Rust Disease Complex

Table 10.6 Prediction equation for progression of white rust and downy mildew complex in relation to environmental factors during 1991–1992 and 1992–1993 crop seasons (Mehta and Saharan 1998)

2 Diseases Year R Y = a1 +b1x1 +b2x2 +b3x3 +b4x4 +b5x5 +b6x6 White rust 1991–92 0.66 56.119 +1.017 −4.086 +8.702 −2.015 +2.723 +0.890 1992–93 0.16 25.041 −4.790 +3.834 +3.429 +1.237 −1.146 +4.270 Downy 1991–92 0.21 −0.511 −0.82 +0.157 −0.075 +0.040 −0.038 −0.005 mildew 1992–93 0.36 −32.744 +0.095 +0.316 +1.316 +0.127 +0.221 −0.038 Malformation 1991–92 0.26 −0.261 +0.048 −0.323 +1.460 −0.202 +0.342 −0.123 (incidence) 1992–93 0.23 −18.687 −2.858 +2.539 +4.540 +0.621 −0.268 +1.056 Malformation 1991–92 0.28 −0.784 +0.079 −0.271 +1.078 −0.172 +0.257 −0.094 (length) 1992–93 0.26 −17.485 −1.598 +1.571 +2.644 +0.413 −0.170 −1.022 ° ° X1 = temp. (maximum) C; X2 = temp (minimum) C; X3 = sunshine (h); X4 = relative humidity (morning) %; X5 = relative humidity (evening) % X6 = rainfall (mm); X = 0.05 level of significance; Y = expected disease incidence

Table 10.7 Correlation coefficient between white rust-downy mildew disease complex and meteorological parameters (Mehta and Saharan 1998) White rust Downy mildew Weather parameters 1991–1992 1992–1993 1991–1992 1992–1993 Temperature (maximum) −0.52 −0.25 0.30 0.45 Temperature (minimum) −0.35 −0.18 0.30 0.42 Sunshine −0.24 −0.08 0.16 0.21 RH (morning) 0.26 −0.08 −0.13 0.01 RH (evening) 0.54 −0.10 −0.24 0.16 Rain fall 0.44 0.22 −0.15 −012 extent of 26 and 28%, respectively, in 1991–1992, and to the extent of 23 and 26%, respectively, in 1992–1993 crop seasons (Mehta and Saharan 1998). The analysis of six test-independent variables that fit the analysis best revealed that each variable played an important role in disease development in addition to other unknown factors which were not included in the study. The prediction equations which showed maximum R2 value are presented in Table 10.6. Results revealed that variable evening RH played a significant role in WR development. Similarly, in the case of DM, maximum and minimum temperatures played a significant role in disease development. The correlation coefficients between diseases and meteorological parameters showed that morning and evening RH and rainfall had positive correlation for WR, whereas temperature had positive correlation for DM development during the 2 years of study (Table 10.7). The infection and progression of WR and DM on rapeseed-mustard was affected by variations in the weather factors. References 211

10.8 Altered Phenotypic Expression of Downy Mildew

Altered phenotypic response of Hyaloperonospora parasitica in Brassica juncea seedlings following prior inoculation with an avirulent isolate of A. candida was studied by Singh et al. (2002). Prior inoculation with the incompatible isolate of A. candida induced resistance to subsequently inoculated H. parasitica. The degree of resistance was proportional to the zoosporangia concentration of the incompatible isolate, and induced resistance was more marked in the cotyledon receiving the inducing inoculum compared to the opposite cotyledon and subsequently emerging true leaves that had not been pre-inoculated. Induction of resistance was also observed if the incompatible isolate of A. candida and H. parasitica was co- inoculated simultaneously. However, the effect was greater with the longer interval between inoculations, up to a period of 4 days. When the incompatible isolate of A. candida was inoculated 4 h after H. parasitica, there was no marked effect on resistance to the latter. In contrast, prior inoculation with the compatible isolate of A. candida increased susceptibility to H. parasitica inoculated subsequently. However, pre- or co-inoculation with H. parasitica suppressed the development of the compatible isolate of A. candida. A spectrum of responses was observed when one cotyledon was inoculated simultaneously with both the incompatible and compatible isolates of A. candida and followed subsequently with H. parasitica after different time intervals. In such combinations, a transition was observed in the host response to H. parasitica from induced resistance/reduced susceptibility, which increased up to 24 h following a simultaneous inoculation with incompatible + compatible isolates of A. candida to an almost neutral reaction after 72 h to induced susceptibility after 96 h. This range of altered responses appeared to reflect the outcome of the differing kinetics and counter-effects of resistance and susceptibility induction.

References

Bains SS (1989) Relationship between detoppings and mixed infections of Albugo candida and Peronospora parasitica on inflorescences of Brassica species. Plant Dis Res 4:148–150 Bains SS, Jhooty JS (1978) Increased susceptibility of virus-infected Brassica juncea to Peronospora parasitica. Plant Dis Report 62:1043–1046 Bains SS, Jhooty JS (1979) Mixed infections by Albugo candida and Peronospora parasitica on Brassica juncea inflorescence and their control. Indian Phytopathol 32:268–271 Bains SS, Jhooty JS (1985) Association of Peronospora parasitica with Albugo candida on Brassica juncea leaves. J Phytopathol 112:28–31 Bains SS, Sokhi SS (1986) Increased levels of mixed infections by Albugo candida and Peronospora parasitica on Brassica juncea inflorescence of detopped plants. Plant Dis Res 1:55–58 Boning K (1936) Investigations on horseradish disease and their control. Angew Bot (J Appl Bot) 18:482–494 Butler EJ (1918) Fungi and diseases in plant. Thaker Spink & Co., Calcutta, pp 297–300 Butler EJ, Jones SQ (1949) Plant pathology. McMillan & Co.Ltd, London Butler EJ, Jones SG (1961) Plant Pathology, p. 68. Macmillan & Co. Ltd, London 212 10 Association or Mixed Infection of Downy Mildew and White Rust Disease Complex

Chaurasia SNP, Singh UP, Singh HB (1982) Preferential parasitism of Peronospora parasitica on galls of Brassica campestris caused by Albugo cruciferarum. Trans Br Mycol Soc 78:379–381 Constantinescu O, Fatehi J (2002) Peronospora-like fungi (Chromista, Peronosporales) parasitic on Brassicaceae and related hosts. Nova Hedwigia 74:291–338 Cooper AJ, Latunde-Dada AO, Woods-Tor A, Lynn J, Lucas JA, Crute IR, Holub EB (2008) Basic compatibility of Albugo candida in Arabidopsis thaliana and Brassica juncea causes broad- spectrum suppression of innate immunity. Mol Plant-Microbe Interact 21:745–756 Greenhalgh JR, Dickinson CH (1975) Differential reactions of three crucifers to infection by Peronospora parasitica (Pers. Ex Fr.) Fr. Phytopathol Z 84:131–141 Holub EB, Williams PH, Crute IR (1991) Natural infection of A. thaliana by A. candida and Peronospora parasitica. Phytopathology 81:1226 Kaur P, Sivasithamparam K, Barbetti MJ (2011a) Host range and phylogenetic relationships of Albugo candida from cruciferous hosts in Western Australia, with special reference to Brassica juncea. Plant Dis 95:712–718 Kaur P, Sivasithamparam K, Barbetti MJ (2011b) Racial/pathotype status and phylogenetic relationships of Albugo candida occurring on cruciferous hosts in Western Australia with special reference to Brassica juncea. (online at http://apsjournals.apsnet.org/doi/pdf/10.1094/ PDIS-10-10-0765) Kaur P, Sivasithamparam K, Li H, Barbetti MJ (2011c) Pre-inoculation with Hyaloperonospora parasitica reduces incubation period and increases severity of disease caused by Albugo candida in a Brassica juncea variety resistant to downy mildew. J Gen Plant Pathol 77:101–106 Kluczewski SM, Lucas JA (1982) Development and physiology of infection by the downy mildew fungus Peronospora parasitica (Pers. ex Fr.) Plant Pathol 31:373–379 Kolte SJ (1985) Diseases of annual edible oilseed crops, Rapeseed-mustard and sesame diseases, vol II. CRC Press, Boca Raton, p 135 Kolte SJ, Sharma KD, Awasthi RP (1981) Yield losses and control of downy mildew and white rust of rapeseed and mustard. Third International Symposium on Plant Pathology, Dec. 14-18, IARI, New Delhi, India, Session 12: 70–71 Kolte SJ, Awasthi RP, Vishwanath (1986) Effect of planting dates and associated weather factors on staghead phase of white rust and downy mildew of rapeseed and mustard. Indian J Mycol Plant Pathol 16:94–102 Lakra BS, Saharan GS (1989) Correlation of leaf and staghead infection intensities of white rust with yield and yield components of mustard. Indian J Mycol Plant Pathol 19:279–281 Lakra BS, Saharan GS (1990) Role of host resistance and planting time in progression of white rust and yield of mustard. Indian Phytopathol 43:201–206 Lakra BS, Saharan GS, Verma PR (1989) Effect of temperature, relative humidity and light on germination of Albugo candida sporangia from mustard. Indian J Mycol Plant Pathol 19:264–267 Liu Q, Rimmer SR (1990) Effect of host genotype, inoculum concentration, and incubation temperature on white rust development in oilseed rape. Can J Plant Pathol 12:389–392 Liu O, Rimmer SR, Scarth R (1989) Histopathology of compatibility and incompatibility between oilseed rape and Albugo candida. Plant Pathol 38:176–182 Magnus P (1894) Das Auftreten der Peronospora parasitica, beeinflusst von der Beschaffenheit und dem Entwickelungszustande der Wirthspflanze. Plant Biol 12:39–44 Meena PD, Thomas L, Singh D (2014) Assessment of yield losses in Brassica juncea due to downy mildew (Hyaloperonospora brassicae). J Oilseed Brassica 5:73–77 Mehta N, Saharan GS (1998) Effect of planting dates on infection and development of white rust and downy mildew disease complex in mustard. J Mycol Plant Pathol 28:259–265 Mehta N, Saharan GS, Babber S (1995) Sequence of events in the pathogenesis of Peronospora and Albugo on mustard. J Indian Bot Soc 74:299–303 Ocfemia GO (1925) The occurrence of the white rust of crucifers and its associated downy mildew in the Philippines. Philipp Agric 14:289–296 Petrie GA (1986) Blackleg and other diseases of canola in Saskatchewan in 1984 and 1985. Can Plant Dis Survey 66:51–53 References 213

Saharan GS (1984) A review of research on rapeseed mustard pathology in India. Annual Workshop AICORPO ICAR, Jaipur, 6–10 August 1984 Saharan GS (1992a) Disease resistance. In: Labana KS, Banga SS, Banga SK (eds) Breeding oilseed Brassicas. Narosa Publication House, New Delhi, India. Ch. 12, pp 181–205 Saharan GS (1992b) Management of rapeseed and mustard diseases. In: Kumar D, Rai M (eds) Advances in oilseeds research. Scientific Publications, Jodhpur, pp 152–188 Saharan GS (2010) Analysis of genetic diversity in Albugo-Crucifer system. J Mycol Plant Pathol 40:1–13 Saharan GS, Mehta N (2002) Fungal diseases of rapeseed-mustard. In: Gupta VK, Paul YS (eds) Diseases of field crops. Indus Publication Company, New Delhi, pp 193–228 Saharan GS, Verma PR (1992) White rust- A review of economically important species. International Development Research Centre, Ottawa, Ontario, Canada, IDRC-MR315e, IV+65p Saharan GS, Verma PR, Nashaat NI (1997) Monograph on Downey mildew of crucifers. Agric. and Agri-Food Canada, Saskatoon Research Centre Technical Bull. 1997–01, Cat. No. A5A- 13/1997-01E, 197pp Saharan GS, Mehta N, Sangwan MS (2005) Development of disease resistance in rapeseed- mustard pp 561-617. In: Saharan GS, Mehta N, Sangwan MS (eds) Diseases of oilseed crops. Indus Publication Co., New Delhi. 643p Saharan GS, Verma PR, Meena PD, Kumar A (2014) Sources of resistance. In: Saharan GS, Verma PR, Meena PD, Kumar A (eds) White rust of crucifers: biology, ecology and management. Springer, New Delhi, pp 181–187 Sansome E, Sansome FW (1974) Cytology and life history of Peronospora parasitica on Capsella bursa-pastoris and of Albugo candida on C. bursapastoris and on Lunaria annua. Trans Br Mycol Soc 62:323–332 Savulescu T, Rayss T (1930) Contribution to the knowledge of the Peronosporaceae of Romania. Ann Mycol 18:297–320 Singh US, Nashaat NI, Doughty KJ, Awasthi RP (2002) Altered phenotypic response to Peronospora parasitica in Brassica juncea seedlings following prior inoculation with an avirulent or virulent isolate of Albugo candida. Eur J Plant Pathol 108:555–564 Vanterpool TC (1958) Rape diseases in Saskatchewan in 1958. Can Plant Dis Survey 38:37–38 Verma PR, Petrie GA (1980) Effect of seed infestation and flower bud inoculation on systemic infection of turnip rape by Albugo candida. Can J Plant Sci 60:267–271 Verma PR, Harding H, Petrie GA, Williams PH (1975) Infection and temporal development of mycelium of Albugo candida in cotyledons of four Brassica spp. Can J Bot 53:1016–1020 Wiese E (1927) A fungus disease of wallflowers and stocks. Gartenwelt 31:486 Chapter 11 Biochemistry of the Host Pathogen Interaction

11.1 Introduction

Biochemical studies of the growth and survival of a pathogen and the changes it induces in its host can ultimately lead to a better understanding of the disease development, its epidemiology, and control. Ideal prerequisites for meaningful studies of the biochemistry of the host-parasite interaction are: (a) A clear understanding of the genetic control of virulence and avirulence in the parasite and of susceptibility and resistance in the host (b) Precise histological and cytological descriptions of spore germination, infection, and the establishment and development of the host-pathogen interaction (c) The availability of methods for maintaining the pathogen alone, and in combination with its host, under controlled conditions. Unfortunately, these criteria have not yet been satisfactorily met for downy mildew of crucifers.

11.2 Metabolic Changes

Many marked shifts were observed in the metabolic processes of plant tissues following infection by biotrophic parasites. These included changes in respiration, photosynthesis, nucleic acid, and protein synthesis and phenol metabolism. There could also be changes in the translocation and accumulation of nutrients and in the levels of endogenous growth substances. The respiration rate was raised sharply soon after H. parasitica infection of cabbage cotyledons reached a maximum, almost twice that of uninfected controls, at the time of the initiation of sporulation (Fig. 11.1). The chlorophyll content of infected and non-infected cotyledons did not differ significantly at any time (Fig. 11.2). The increased respiration rate of the infected tissues did not reflect any significant changes to the pentose phosphate pathway in this infection since no

© Springer Nature Singapore Pte Ltd. 2017 215 G. S. Saharan et al., Downy Mildew Disease of Crucifers: Biology, Ecology and Disease Management, https://doi.org/10.1007/978-981-10-7500-1_11 216 11 Biochemistry of the Host Pathogen Interaction

Fig. 11.2 Chlorophylla plus chlorophyllb, content of infected, and uninfected cotyledons at various times after inoculation: (●-●), infected; (○-○), uninfected (Thornton and Cooke 1974)

acyclic polyhydric alcohols were detected in soluble extracts of either infected leaves or fungal conidia (Thornton and Cooke 1974) (Figs. 11.3 and 11.4). Pathogenesis in Brassica-Hyaloperonospora combinations was observed to be accompanied by large increases in electrolyte leakage (Fig. 11.5, 11.6, 11.7, and 11.2 Metabolic Changes 217

glucose 2.0

1.0

fruclose 2.0

1.0

1.0 sucrose 0.5 Carbohyrate content (mg per g fresh wt)

1.0 inositol 0.5 TT T TT 12345678 Days ofter inoculation

Fig. 11.3 Carbohydrate content of the alcohol-soluble fraction of infected and uninfected cotyledons at various times after inoculation with Hyaloperonospora parasitica: ■ = infected; □ = uninfected; T trace (indicating that the peak height of the TMS derivative was indeterminable at an attenuation of 20 × 103 (Thornton and Cooke 1974)

11.8) and increased activity of glucosidase (Fig. 11.6), ribonuclease (Fig. 11.7), and peroxidase (Fig. 11.8) (Kluczewski and Lucas 1982). The large increase in glucosidase was of pathogen origin, while enhanced ribonuclease activity was due to a new post-inflectional form of the enzyme. In vivo infected leaves of B. juncea produced cellulase, endo-PMG, and endo-PG (Singh et al. 1980). 218 11 Biochemistry of the Host Pathogen Interaction

0.04

0.03

0.02

carbohydrate content 0.01 (mg per g cotyledon fresh wt)

glucose fructose trehalose sucrose inositol

Fig. 11.4 Principal carbohydrates of the alcohol-soluble fraction of sporangia from infected cotyledons and control washings, 7 days after inoculation with Hyaloperonospora parasitica. ■ = infected; □ = uninfected (Thornton and Cooke 1974)

Fig. 11.5 Conductivity changes of de-ionized glass-distilled water containing samples of uninfected cotyledons (...) and cotyledons infected (−) by Hyaloperonospora parasitica isolate from cauliflower (○) and oilseed rape (–) (Kluczewski and Lucas 1982). Each point represents the mean of four replicates 11.2 Metabolic Changes 219

Fig. 11.6 β-glucosidase activity in extracts of control cotyledons (...) and cotyledons infected (−) by either cauliflower ○( ) or oilseed rape (−) isolate of Hyaloperonospora parasitica (Kluczewski and Lucas 1982)

Fig. 11.7 Acid ribonuclease activity in extracts of control cotyledons (...) and cotyledons infected (−) by Hyaloperonospora parasitica isolate from cauliflower ○( ) and oilseed rape (−) (Kluczewski and Lucas 1982) 220 11 Biochemistry of the Host Pathogen Interaction

Fig. 11.8 Peroxidase activity in extracts of control cotyledons (...) and cotyledons infected by either cauliflower ○( ) or oilseed rape (−) isolate of Hyaloperonospora parasitica (−) (Kluczewski and Lucas 1982)

11.3 Role of Natural Biochemical Compounds

There are a number of natural biochemical compounds present in host tissues which may influence the defence mechanism of crucifers against downy mildew infection. The role of phenolic compounds, glucosinolates, phytoalexins, and flavour volatile compounds in providing resistance to crucifers against downy mildew infection has been explained in Sect. 12.6, ‘Biochemical Basis of Resistance’, of Chap. 12, ‘Host Resistance’.

11.4 Biochemistry of Disease Resistance

Plants can employ chemical defence either as preformed antimicrobial substances (phytoanticipins) or as induced antimicrobial substances which accumulate after contact with the pathogen (phytoalexins) (Van Etten et al. 1994). In Arabidopsis, as in other Brassica, glucosinolates come into question as potential phytoanticipins. However, phytoanticipins are generally correlated with non-host resistance (Mansfield 2000), and there is no evidence that glucosinolates play a role in the interaction of Arabidopsis with H. parasitica. The accumulation of defence gene transcripts such as PAL and CHS and the increase in peroxidase activity in cell suspension cultures treated with elicitor were reported by Davis and Ausubel (1989) 11.4 Biochemistry of Disease Resistance 221 and indicated that stimulation of phenylpropanoid metabolism in Arabidopsis could be expected to be associated with resistance, as observed in several other plant species (Dixon and Paiva 1995). In the Arabidopsis-H. parasitica interaction, an early oxidative burst of H2O2 production is observed on penetration of the epidermis by an avirulent race of the pathogen. Following the oxidative burst is the genetically programmed hypersensitive cell death response (HR). In an HR against an avirulent Pseudomonad, a shift from housekeeping to defence metabolism which affected an estimated 10% of the transcriptome was demonstrated (Scheideler et al. 2002), and this is presumably similar in the HR against H. parasitica. Since H. parasitica is an obligate biotroph, the rapid hypersensitive response (HR) which occurs in the epidermal cells adjacent to the penetration hyphae, and sometimes an additional few cells in the mesophyll (Koch and Slusarenko 1990), would be expected to be sufficient to effectively condition resistance by preventing the establishment of the highly co-evolved nutritional relationship between host cell and pathogen, which depends on haustoria. Nevertheless, the HR to H. parasitica is associated with the accumulation of at least one antimicrobial phytoalexin, camalexin (Slusarenko and Mauch-Mani 1991; Tsuji et al. 1992), and it was reported that the phytoalexin- deficient mutants (pad1-1, pad2-1, and pad3-1 and the double mutants pad1- 1/pad2-1, pad1-1/pad3-1, and pad2-1/pad3-1) showed increased susceptibility in incompatible combinations with five races of H. parasitica, namely, EMOY, EMWA, CALA, HIKS, and HIND (Glazebrook et al. 1997). Camalexin, the only Arabidopsis phytoalexin so far described, is an indole thia- zole derivative, as are other phytoalexins so far reported from the Brassicaceae. Thus, camalexin is synthesized from tryptophan, and not from phenylalanine, which is a precursor for phenolics in plants. This metabolic distinction was made use to assess the relative contributions of phenolics and phenolic polymers to the resistance of Arabidopsis to H. parasitica, separately from phytoalexins (Mauch-Mani and Slusarenko 1996). Thus, in Arabidopsis, it is possible to inhibit phenolic metabolism without directly interfering with phytoalexin synthesis (Fig. 11.9). Using specific inhibitors of PAL (2-aminoindan-2-phosphonic acid, AIP) and cinnamyl alcohol dehydrogenase (2-hydroxyphenyl-aminosulphinyl acetic acid 1, 1-dimethyl ester, OH-PAS) to inhibit phenolic metabolism in general or lignifications in particular, it was shown that loss of lignification caused a mild shift towards susceptibility but that a more general inhibition of the phenolic metabolism resulted in complete susceptibility (Mauch-Mani and Slusarenko 1996). Feeding salicylic acid (SA) back into the system restored resistance in the presence of AIP and was interpreted as showing the dependence of resistance expression on SA. These results conform with the observations of Delaney et al. (1994) for transgenic Arabidopsis carrying the bacterial nahG gene and are therefore unable to accumulate SA. However, despite the overwhelming effect of SA, the contribution of lignification to resistance against H. parasitica was shown, and this highlights the multifac- torial nature of the resistant response. It has been suggested that the SA that is required for PR1 gene expression and SAR might be predominantly synthesized via isochorismate, whereas the SA which modulates cell death in the HR might arise predominantly from phenylalanine (Fig. 11.9) (Wildermuth et al. 2001). 222 11 Biochemistry of the Host Pathogen Interaction

Fig. 11.9 Simplified scheme of the biosynthesis of the defence-related compounds camalexin, salicylic acid, and lignin in Arabidopsis. Chorismate is the first branch point, since camalexin arises via tryptophan, while salicylic acid is synthesized via isochorismate and phenylalanine, and lignins arise via phenylalanine. Chorismate is converted into isochorismate by isochorismate synthase (ICS). Phenylalanine ammonia-lyase (PAL) converts phenylalanine into cinnamic acid and is specifically inhibited by aminoindan phosphonic acid AIP( ). From cinnamic acid, the pathway branch to produce salicylic acid or, via cinnamaldehydes and monolignols, lignin. The conversion from cinnamaldehydes to monolignols by cinnamyl alcohol dehydrogenase (CAD) is inhibited by hydroxyphenyl-aminosulphinyl acetic acid dimethyl ester (OH-PAS) (Slusarenko and Schlaich 2003)

The idea that plants might lignify not only their own walls to strengthen them as barriers to pathogen spread (Hijwegen 1963), but that the pathogens themselves might be inactivated by polymerization of monilignols to lignin in their walls as part of the peroxidase catalysed intercellular free radical condensation reaction, was proposed by Hammerschmidt and Kuc (1982) and Ride (1983). However, Slusarenko and Schlaich (2003) have found evidence for the active lignification of hyphae in intercellular spaces (Mauch-Mani and Slusarenko 1996). Arabidopsis host- pathosystem is not an easy subject for biochemical study, and comparative studies are complicated by leaf age- and ecotype-specific differences in basal enzyme activity levels (Mauch-Mani et al. 1993). After the inoculation of plants with virulent or avirulent isolates of H. parasitica, Slusarenko and Schlaich (2003) did not observe a significant pattern of change in the activities of superoxide dismutase, catalase, ascorbate peroxidase, lipolytic acyl-hydrolase, lipoxygenase, or linolenic acid 13-hydroperoxide decomposing activity. All these enzymes have been reported to be important in one or more pathosystems. It is possible that in Arabidopsis, these enzymes have no role in resistance against H. parasitica or, because only relatively few cells show a HR, any changes are diluted out in comparison with the bulk of References 223 non-stimulated cells in the leaf. In contrast, transcripts for lipoxygenase were reported to be induced by treatment with Pseudomonas (Melan et al. 1993), which leads to a quantitatively greater leaf response than in the H. parasitica pathosystem where Slusarenko and Schlaich (2003) found no increased transcript levels using the same probes (Mauch-Mani et al. 1993; Slusarenko 1996). Thus, SA seems to be important for defence against most H. parasitica races, presumably because of its role as a signal amplifier, and it seems that lignification and camalexin might be needed for full resistance, since some resistance is lost when lignifications is blocked or in pad4 or pad1 + 2, 1 + 3, or 2 + 3 mutants.

References

Davis KR, Ausubel F (1989) Characterization of elicitor-induced defense responses in suspension- cultured cells of Arabidopsis. Mol Plant-Microbe Interact 2:363–368 Delaney TP, Uknes S, Vernooij B, Friedrich L, Weymann K, Negrotto D, Gaffney T, Gut-Rella M, Kessmann H, Ward E, Ryals J (1994) A central role for salicylic acid in plant disease resistance. Science 266:1247–1250 Dixon RA, Paiva NL (1995) Stress induced phenyl-propanoid metabolism. Plant Cell 7:1085–1097 Glazebrook J, Zook M, Mert F, Kagan I, Rogers ER, Crute IR, Holub EB, Hammerschmidt R, Ausubel FM (1997) Phytoalexin deficient mutants of Arabidopsis reveal that PAD4 encodes a regulatory factor and that four PAD genes contribute to downy mildew resistance. Genetics 146:381–392 Hammerschmidt R, Kuc J (1982) Lignification as a mechanism for systemic acquired resistance in cucumber. Physiol Plant Pathol 20:61–71 Hijwegen T (1963) Lignification, a possible mechanism of active response against pathogens. Neth J Plant Pathol 69:314–317 Kluczewski SM, Lucas JA (1982) Development and physiology of infection by the downy mildew fungus Peronospora parasitica (Pers. ex Fr.) Fr. in susceptible and resistant Brassica species. Plant Pathol 31:373–389 Koch E, Slusarenko A (1990) Arabidopsis is susceptible to infection by a Downy mildew fungus. Plant Cell 2:437–445 Mansfield JW (2000) Antimicrobial compounds and resistance. The role of phytoalexins and phytoanticipins. In: Slusarenko AJ, Fraser RSS, van Loon LC (eds) Mechanisms of resistance to plant diseases. Kluwer Academic Publishers, Dordrecht, pp 325–370 Mauch-Mani B, Slusarenko AJ (1996) Production of salicylic acid precursors is a major function of phenylalanine ammonia lyase in the resistance of Arabidopsis to Peronospora parasitica. Plant Cell 8:203–212 Mauch-Mani B, Croft KPC, Slusarenko AJ (1993) The genetic basis of resistance of Arabidopsis thaliana L. Heyhn. to Peronospora parasitica. In: Davis KR, Hammerschmidt R (eds) Arabidopsis thaliana as a model for plant–pathogen interactions. APS Press, St Paul, pp 5–20 Melan MA, Dong X, Endara ME, Davis KR, Ausubel FM, Peterman TK (1993) An Arabidopsis thaliana lipoxygenase gene can be induced by pathogens, abscisic acid, and methyl jasmonate. Plant Physiol 101:441–450 Ride JP (1983) Cell walls and other structural barriers in defence. In: Callow JA (ed) Biochemical plant pathology. Wiley, Chichester, pp 215–236 Scheideler M, Schlaich NL, Fellenberg K, Beissbarth T, Hauser NC, Vingron M, Slusarenko AJ, Hoheisel JD (2002) Monitoring the switch from housekeeping to pathogen defense metabolism in Arabidopsis thaliana using cDNA arrays. J Biol Chem 277:10555–10561 224 11 Biochemistry of the Host Pathogen Interaction

Singh SB, Singh DV, Bais BS (1980) In vivo cellulase and pectinase production by Albugo candida and Peronospora parasitica. Indian Phytopathol 33:370–371 Slusarenko AJ (1996) The role of lipoxygenase in plant resistance to infection. In: Piazza GJ (ed) Lipoxygenase and lipoxygenase pathway enzymes. American Oil Chemists Society Press, Champaign, pp 176–197 Slusarenko AJ, Mauch-Mani B (1991) Downy mildew of Arabidopsis thaliana caused by Peronospora parasitica: a model system for the investigation of the molecular biology of host-pathogen interactions. In: Hennecke H (ed) Advances in the molecular genetics of plant– microbe interactions, vol 1. Kluwer Academic Press, Dordrecht, pp 280–283 Slusarenko AJ, Schlaich NL (2003) Downy mildew of Arabidopsis thaliana caused by Hyaloperonospora parasitica (formerly Peronospora parasitica). Mol Plant Pathol 4:159–170 Thornton JD, Cooke RC (1974) Changes in respiration, chlorophyll content and soluble carbohydrates of detached cabbage cotyledons following infection with Peronospora parasitica (Pers. ex. Fr.) Fr. Physiol Plant Pathol 4:117–125 Tsuji J, Jackson E, Gage R, Hammerschmidt R, Somerville SC (1992) Phytoalexin accumulation in Arabidopsis thaliana during the hypersensitive reaction to Pseudomonas syringae pv. syringae. Plant Physiol 98:1304–1309 Van Etten HD, Mansfield JW, Bailey JA, Farmer EE (1994) Letter to the editor: two classes of plant antibiotics: phytoalexins versus ‘phytoanticipins. Plant Cell 6:1191–1192 Wildermuth MC, Dewdney J, Wu G, Ausubel FM (2001) Isochorismate synthase is required to synthesize salicylic acid for plant defence. Nature 414:562–571 Chapter 12 Host Resistance

12.1 Introduction

Genetic resistance is the most important attribute of the host defence against Hyaloperonospora parasitica. Host resistance provides an economical, environmentally benign, and widely accepted method of managing downy mildew of crucifers. Present day’s emphasis on organic agriculture relies mainly on large-scale deployment of disease-resistant crop cultivars in addition to the use of techniques such as cultural practices (sowing time, soil solarization, crop rotation, use of organic manures, plant population, clean cultivation, soil, and plant moisture regulation) and biological disease control. Brassica host-pathogen interaction can be classified into two types, compatible and incompatible interaction, leading to the phenotypes of susceptibility and resistance to certain pathogens. Obviously, the incompatible interaction has been extensively exploited by the Brassica breeders to develop resistant cultivars for increasing crop production. Large-scale deployment of elite cultivars carrying an R gene (monoculture) imposes higher selection pressure on the pathogen carrying the cognate Avr gene to survive, resulting in the modification or deletion of the recognized Avr gene or generation of novel effector genes that can escape the recognition of the old ‘R’ gene. This R-Avr interactive coevolution explains why many resistant cultivars in the field lose their resistance in a relatively short period of time. Hence, knowledge on the molecular mechanisms of Brassica host-pathogen interaction and the use of horizontal, durable, and broad- spectrum resistance has become very important in disease resistance breeding programmes. Both Brassica host and pathogen are constantly evolving for survival, and such a co-evolutionary struggle results in the generation of advanced weapons for attack and counter-attack by the invading pathogen and the Brassica host plant for defence. Pathogenic variability in Brassica host-pathosystem has been reported in Albugo candi (white rust), Hyaloperonospora parasitica (downy mildew), and Sclerotinia sclerotiorum (stem rot) largely due to sexual reproduction and in Alternaria brassicae (Alternaria blight) due to mutation, uniform host selection,

© Springer Nature Singapore Pte Ltd. 2017 225 G. S. Saharan et al., Downy Mildew Disease of Crucifers: Biology, Ecology and Disease Management, https://doi.org/10.1007/978-981-10-7500-1_12 226 12 Host Resistance extensive dispersal, or the existence of a cryptic sexual stage. The interplay of host resistance and pathogen virulence in latter system is likely to be quite different to major gene-based system. In nature, Brassica hosts employ a wide range of mechanisms to restrict the growth of the pathogen. The first mechanism operates before the pathogen contact, while the other mechanisms operate after the pathogen has made contact with the host. Selection for resistance can be done by measuring the growth and development of the pathogen; the higher the reduction, the more resistant the host plant. The nature and mechanism of resistance in the Brassica host-pathosystem are multilay- ered and multicomponents. Very useful information has been generated by the co- operation and co-ordination of plant pathologists and breeders through conventional and modern techniques to ameliorate disease resistance in Brassica crops to be the most effective, and stable, and economical. However, recently breeding for disease resistance has been transformed into molecular genetics, which has been providing knowledge of the genes that underlie natural variation in valued traits of Brassica species Arabidopsis thaliana. Majority of the R genes have been characterized as encoding NBS-LRR receptors, some encodes receptor-like kinase, and some being atypical in terms of protein structures. The broad-spectrum resistance mediated by PAMP (pathogen-associated molecular pattern)-recognizing receptors might be exploited using conventional or transgenic approaches. To comprehend Brassica host-pathogen interactions, we have to first focus on the identification of the pathogen factors, PAMs, and effectors, which are conserved across isolates, essential for pathogens survival, and unable to withstand modification. The plant ‘R’ genes that recognize these conserved pathogen factors have a greater possibility of remaining effective overtime because of their evolutionary constraints on the pathogen factors. The second approach should be the identification and study of QTLs for partial and often durable resistance. Most of these QTLs encode genes directly involved in resistance which activates general defence mechanisms of the host with added bonus of conferring resistance to multiple pathogen species.

12.2 Mechanism of Host Resistance

The first defence barrier in crucifers is the cuticle which is often covered with a waxy layer, a hairy surface, and a few stomata with narrow apertures. The mechanisms of resistance to H. parasitica in Chinese rape, cabbage, and radish were studied by Wang (1949) through observations on pathogen entry, mycelial and haustorial development, and sporulation. All plants, regardless to whether they are susceptible or resistant, were initially penetrated directly through the epidermal cells or by entering the stomata. After penetration, the mycelia grew through the intercellular spaces of the leaf mesophyll, and haustoria penetrate the cells of susceptible hosts. In the resistant and immune hosts, development of mycelia and formation of haustoria were curtailed with the death of the surrounding host cells (Plate 12.1). The 12.2 Mechanism of Host Resistance 227

Plate 12.1 Hyaloperonospora parasitica brassicae race 2. Entry of germ tube of the conidium, through (a) an epidermal cell and (b) a stoma. Mycelium in tissue of (c) the susceptible Chinese rape host and (d) the immune radish host. Legend: Sp, conidium; Ap, appressorium; IH, infection hypha; My, mycelium; Ha, haustorium; Sh, sheath; Ep, epidermis; St, stoma; Sp, spongy mesophyll tissue; and DC, dead host cells (Wang 1949) pathogen sporulated abundantly on susceptible hosts, but necrotic reaction was associated with the infection of the resistant hosts. On the immune hosts, few minute necrotic spots/or occasionally no visible symptoms were observed. Weak light and high moisture conditions may alter the resistant or immune reaction of the host. The growth of two isolates of H. parasitica obtained from cauliflower and oilseed rape (B. napus) was assessed in their respective hosts of origin and also in the alternative combination by Kluczewski and Lucas (1982). Both isolates were capable of infecting either host, but there were marked contrasts in the time course and extent of mycelial development, the amounts of associated host cell necrosis, and eventual intensity of sporulation (Figs. 12.1 and 12.2; Plate 12.2). Oilseed rape 228 12 Host Resistance

Fig. 12.1 Relationship between mycelial development and host-cell necrosis estimated as granulation and browning of cells in (a) cauliflower and b( ) oilseed rape inoculated with Hyaloperonospora parasitica isolates from cauliflower (CI) and oilseed rape (Rl).□ -CI mycelial growth index; ■–CI necrotic cell index; □-Rl necrotic cell index. Bars indicate + standard deviation (Kluczewski and Lucas 1982) which is partially resistant to the isolate from cauliflower exhibits extensive necrosis of mesophyll cells in conjunction with reduced mycelial development and delayed and reduced sporulation by the pathogen (Figs. 12.1 and 12.2; Plate 12.2). The isolate from oilseed rape is virulent on both host species. Pathogenesis in the susceptible 12.2 Mechanism of Host Resistance 229

Fig. 12.2 Time course of sporulation of Hyaloperonospora parasitica isolate from cauliflower (o) and oilseed rape (■) on cauliflower (- -) and oilseed rape (…). Bars indicate + standard deviation (Kluczewski and Lucas 1982) combination is accompanied by a large increase in electrolyte leakage and increased activity of the enzymes β-glucosidase, ribonuclease, and peroxidase. The growth of hyphae in the susceptible cultivar of Japanese radish was reported to be faster than that in the resistant cultivar (Ohguchi and Asada 1991). Five days after inoculation, haustoria were formed in the cells of the 63rd cell layer from the inoculated layer in the susceptible cultivar and in the cells of 12th cell layer in the resistant cultivar. The haustoria formed in both cultivars were similar in size and shape. On the surface of haustoria, spherical or semi-spherical granules, 1.7–3.7 μm in diameter, were often observed in the susceptible cultivars while rarely observed in the resistant cultivar. For resistance of Arabidopsis accessions harbouring the resistance genes to EMoy2 isolate carrying corresponding Avr genes was characterized by the induction of a wide range of defence mechanisms including the encashment of haustoria with callose-containing material and the HR, in invaded and neighbouring cells. 230 12 Host Resistance

Plate 12.2 Cotyledon tissue 4 days after inoculation with Hyaloperonospora parasitica cauliflower isolate stained with trypan blue and cleared in chloral hydrate. (a) Intercellular hyphae in oilseed rape cultivar Primor showing left, developing haustoria (arrows) in host mesophyll cells close behind the hypha apex, and right, necrosis of penetrated host cells in older regions of a hypha x 400; (b) intercellular hyphae in cauliflower cultivar VSAG forming abundant intracellular haustoria. Note absence of host-cell necrosis, x 400 (Kluczewski and Lucas 1982) 12.4 Systemic Acquired Resistance 231

Table 12.1 Expected Differentiation in response to plant sequence of events leading to signal hypersensitive reaction Production of cultivar-specific expression in crucifers to H. elicitors parasitica infection (Lebeda and Schwinn 1994) Primary recognition Activated responses De novo protein synthesis in penetrated cell Irreversible membrane damage and release of phenolics Release of endogenous elicitors Accumulation of wall-bound phenolics Release of secondary signals Secondary recognition Transcriptions of mRNAs controlling biosynthesis of lignin Precursors in surrounding Deposition of lignins in and around the infection site

Limited pathogen development was clearly associated with the HR in FN (minute necrotic flecks and no sporulation) and PN (necrotic pits and no sporulation) phenotypes (Soylu et al. 2004).

12.3 Host-Pathogen Recognition System

The Arabidopsis-H. parasitica system was recently adopted as a model system for studying the recognition process for gene-for-gene interactions (Davis and Hammerschmidt 1993). The determination of specificity and mechanism of recognition is a highly complex phenomenon. The comprehension of this phenomenon depends on better knowledge of molecular biology and genetics of the host-pathogen interaction. Cytological and biochemical studies are being attempted to identify the stages at which the incompatibility recognition events occur. Proposed steps between recognition signal transduction and activated defence during the hypersensitive response in crucifers to H. parasitica are outlined in Table 12.1 (Lebeda and Schwinn 1994).

12.4 Systemic Acquired Resistance

Systemic acquired resistance (SAR) has been demonstrated in Arabidopsis plants treated with chemical inducers such as 2,6-dichloroisonicotinic acid or INA (Uknes et al. 1992) and after biological treatments, such as necrotic infection with 232 12 Host Resistance

Pseudomonas or Fusarium (Cameron et al. 1994; Mauch-Mani and Slusarenko 1994). In addition, the chemical inducers benzothiadiazole (BTH) and β-aminobutyric acid (BABA) have been shown to be effective in inducing resistance against downy mildew (Lawton et al. 1996; Zimmerli et al. 2000). Resistance is expressed within a few days of exposure to the inducer and inhibits subsequent infection by both bacterial and fungal pathogens, including H. parasitica. The degree of protection varies depending upon the concentration of inducer chemical used, but at higher dose rate, sporulation of H. parasitica is completely inhibited. Microscopic examination of induced plants inoculated with the fungus reveals that hyphal growth is restricted to the initial penetration site, associated with a necrotic reaction in host cells. In plants treated with lower doses of the chemical, some hyphal development occurs, but haustoria are reduced in size, and many are encased in material of host origin. Host cells penetrated by haustoria often become necrotic. Similar cytological events occur in uninduced hosts inoculated with incompatible isolates of H. parasitica (Kluczewski and Lucas 1982). Induction of SAR appears to enhance the efficiency of host defence responses and thereby disrupts the development of a biotrophic relationship in a normally compatible host. The biochemical mechanism of SAR is not yet understood, but models envisage a translocated single molecule that induces changes in tissues removed from the initial inoculation site. Development of SAR is associated with induction of PR proteins (Uknes et al. 1992).

12.4.1 Expression of Systemic Acquired Resistance

A mutant (Cpr 5) of Arabidopsis thaliana which constitutively expresses pathogenesis- related (PR) proteins was identified from a screen for expression of system acquired resistance (SAR). This single recessive mutation also leads to spontaneous expression of chlorotic lesions and reduced trichome development. Cpr 5 mutants were characterized by constitutive resistance to two virulent pathogens Pseudomonas syringae pv. maculicola ES 4326 and Hyaloperonospora parasitica NOCO2, endogenous expression of the PR gene 1 (PR-1), and exhibited elevated levels of salicylic acid (SA). Cpr 5 NPR 1 plants retained heightened resistance to H. parasitica

NOCO2 and elevated expression of the defensive gene PDF 1.2, implying that NPR 1-independent resistance signalling also occurs. It was concluded that Cpr 5 leads to constitutive expression of both an NPR 1-dependent and NPR 1-independent SAR pathways. Characterization of this mutation indicated that these pathways were connected in early signal transduction steps and have overlapping functions in disease resistance (Bowling et al. 1997). According to Clarke et al. (1998), the dominant CRP6-1 mutant expressed the SA/NPR 1 regulated PR genes (PR-1, BGL-2, and PR-5) and displayed enhanced resistance to H. parasitica NOCO2 in the absence of SAR induction. Cao et al. (1998a) found that NPR 1 confers resistance to H. parasitica in a dosage-dependant fashion. Overexpression of NPR 1 led to enhanced resistance with no obvious detrimental effect on the plants. However, Thomma et al. (1998) did not find any role 12.5 Genetics of Host-Pathogen Relationship 233 of ethylene-/jasmonate-responsive antimicrobial gene in the host defence against H. parasitica. Mauch-Mani and Slusarenko (1996) observed that production of salicylic acid precursors was a major function of phenylalanine ammonia-lyase in the resistance of Arabidopsis to H. parasitica. According to Lawton et al. (1996), benzothiadiazole (BTH) is a novel chemical activator of disease resistance and works by activating SAR in A. thaliana to induce resistance against H. parasitica. Chemical treatment induced accumulation of mRNAs from the SAR-associated genes, PR-1, PR-2, and PR-5. BTH treatment induced both PR-1 mRNA accumulation and resistance against H. parasitica in the ethylene response mutants etr 1 and ein 2 and in the methyl jasmonate-insensitive mutant, Jar 1, suggesting that BTH action was independent of these plant hormones. Components of systemically induced resistance responses (Bowling et al. 1994; Cao et al. 1998a, b; Delaney et al. 1995) and other potential pathways of defence (Dietrich et al. 1994; Glazebrook and Ausubel 1994; Greenberg et al. 1994; Ausubel et al. 1995) have also been characterized in Arabidopsis.

12.5 Genetics of Host-Pathogen Relationship

Resistance derived from the Broccoli Introduction No. PI 189028 to H. parasitica race 1 is found to be governed by one dominant gene. The distribution of resistant plants in populations segregating both downy mildew resistance and wax less foliage indicates that resistance is independent of foliage wax. Resistance to race 1 and race 2 obtained from the cabbage introduction PI 245015 is found to be inherited independently. Resistance is governed by one dominant gene for each race (Natti et al. 1967). However, in a later study, Hoser-Krause et al. (Hoser-Krauze et al. 1991) found that in broccoli (B. oleracea var. botrytis), resistance to a Polish isolate of H. parasitica at the 4–5-leaf stage is determined by a single recessive gene different from genes determining resistance at the cotyledon stage. Subsequently, they (Hoser-Krauze et al. 1995) found that in broccoli resistance to downy mildew is governed by three or four dominant complimentary genes. Both seedling and mature plant resistance have been reported in B. oleracea with the latter being quantitative (Dickson and Petzoldt 1996). Carvalho and Monterio (Caravalho and Monteiro 1996) crossed ‘R’ B. oleracea cv. Algarvia with ‘S’ cv. Penca de chaves and observed that the resistance to downy mildew is dominant and controlled by two independent loci with Algarvia being heterozygous at both loci (R1 r1, R2 r2) and Penca de chaves being homozygous recessive (r1 r1 r2 r2). Resistance to downy mildew in broccoli and cauliflower is controlled by a single dominant gene. No susceptibility was observed among F2 seedlings derived from intercrossing the R lines indicating that they all are the same or closely linked broad-spectrum R genes (Vicente et al. 2012). In Chinese cabbage resistance to downy mildew at the cotyledon stage expressed as a reduction in the sporulation capacity of H. parasitica was found to be under 234 12 Host Resistance dominant monogenic control (Niu et al. 1983). However, Yuen (1991) while analysing Chinese cabbage lines with a reduced rate of mildew development found additive effects with involvement of several resistant genes. The resistance to H. parasitica in radish cultivars Tokinaski (All Season) and Okura was found to be controlled by two dominant and independent genes (Bonnet and Blancard 1987). Cytoplasmic male sterile (B. campestris) breeding lines with resistance to downy mildew have been identified by Leung and Williams (1983). Downy mildew resistance was expressed in cotyledons as a reduction in the sporulation capacity of H. parasitica. In high and partially resistant hosts, spore production ranged from 7 to 30 spores/g host tissue as compared to 260 spores/g host tissue in the fully susceptible hosts. In the chinensis lines, 80% of the plants showed high to partial resistance to H. parasitica, whereas in the other lines, resistance ranged between 10 and 50%. Differential host resistance to homologous isolates of H. parasitica has been identified in B. rapa (B. campestris), B. napus, B. juncea, and B. oleracea. In B. napus, resistance in the oilseed rape cultivar Cresor is controlled by a single dominant allele (Lucas et al. 1988). In B. oleracea, differential resistance has been located in a land race cauliflower ‘Palermo Green’. A model based on two or possible three major genes has been proposed by Moss et al. (1988) to account for the reaction patterns of individual plants to select fungal isolates within a host population (Table 12.2). In B. campestris, both rapid cycling and commercial genotypes have been identified for differential resistance (Table 5.6). Four homologous isolates have identified a range of differential responses. According to Nashaat et al. (1995a; b, 1996), resistance of the RES-01-1-4 and RES-26 lines of B. napus to isolate P003 of H. parasitica is conditioned by a single dominant resistant gene, whereas resistance of RES-02 is conditioned by two independent dominant resistance genes. Later, Nashaat and Awasthi (1995) selected differential putative homozygous resistant lines from seedling populations of accessions that exhibited a heterogeneous reaction to the isolates from B. juncea (Tables 12.3 and 12.4). In inoculation tests on Arabidopsis thaliana with seven pathogen isolates and eleven host accessions, a range of interaction phenotypes were observed including localized necrosis (flecking), more extensive cell collapse (pitting), delayed sporu-

Table 12.2 Inheritance of resistance in cauliflower to H. parasitica using Palermo Green model (Moss et al. 1988) Resistance Virulence phenotype – R1 – R1 P. parasitica isolates A1 A2 – – R2 R2 Approx. resistant seedlings observed P005 1 2 + – – – 95 P015 1 – + – + – 80 P018 – 2 + + - – 80 P006 – – + + + + 0 % phenotype in seed 5 15 15 65 stock A avirulence gene, R resistance gene, + + susceptibility, – resistance 12.5 Genetics of Host-Pathogen Relationship 235

Table 12.3 Response of groups A, B, C, D, and E of Brassica juncea accessions and of one accession of B. napus at the cotyledon stage to infection with four isolates of Hyaloperonospora parasitica (Nashaat and Awasthi 1995) Disease index Isolates of P. parasitica Brassica species Host categorya IP01/IP02 P003 R1 B. juncea A (4) 1 1 1 B (2) 2–3 1 1 C (19) 6–8 1 1 D (5) 6–8 2–3 1 E (1) 6–7 5–6 1 B. napus Ariana 1 7–8 7–8 a= number of accessions of each group in parentheses

Table 12.4 Examples of a successful selection for putative homozygous resistance response to Hyaloperonospora parasitica from a heterogeneous starting population of Brassica juncea at the cotyledon stage (Nashaat and Awasthi 1995) Isolates of P. parasitica Disease index (standard deviation, n = 24) Brassica junceaa IP01 IP02 P003 R1 Kranti 3.0 (3.0) 4.1 (3.6) 1.0(0.0) 1.0(0.0) RES-BJ01 1.0 (0.0) 1.0 (0.0) 1.0(0.0) 1.0(0.0) Krishna 5.4 (2.5) 5.7 (2.5) 1.0(0.0) 1.0(0.0) RES-BJ02 1.0 (0.0) 1.0 (0.0) 1.0(0.0) 1.0(0.0) Varuna 6.1 (1.8) 7.1 (2.2) 1.0(0.0) 1.0(0.0) RES-BJ03 1.0 (0.0) 1.0 (0.0) 1.0(0.0) 1.0(0.0) a= RES-BJ01, RES-BJ02, and RES-BJ03 lines selected from seedling populations of Kranti, Krishna, and Varuna, respectively lation, or complete susceptibility (Holub et al. 1994). Segregation for these phenotypes among F2 progeny from a half-diallel cross between nine A. thaliana accessions and ten host loci (termed RPP, recognition of H. parasitica loci) has been observed. Four of these loci (RPP1, RPP2, RPP4, and RPP7) were mapped (Tor et al. 1994), along with a further locus RPP5 (Parker et al. 1993). Three loci were apparently clustered together on chromosome 4. There was also evidence for the existence of different alleles at a single locus, although corresponding crosses between pathogen isolates differing in specificity are required to confirm whether different alleles at closely linked loci might explain such results. A genetic model postulating the existence of complementary recognition loci in the fungus designated ATR-A. thaliana recognition has been proposed (Holub et al. 1994). These results provide support for the gene-for-gene model of specificity in crucifer’s downy mildew. The interaction phenotypes between different host lines and pathogen isolates reveal a degree of complexity in the system, with partial dominance, epistasis, and gene dosage effects. 236 12 Host Resistance

Table 12.5 Reaction to Hyaloperonospora parasitica isolate P003 of F1, F2, and back cross F1 (BC1F1) progeny from crosses involving spring Brassica napus accessions RES-26 and Callypso (Nashaat et al. 1997) Number of plants with a disease reaction rating of Generation and cross 1–5a 6–7 Ratio X2 P

F1 RES-26/Callypso and reciprocal 240 (–23) 0

F2 RES-26/Callypso and reciprocal 1581 520 1:1 0.07 0.79

BC1F1RES-26 Callypso and reciprocal/Callypso 920 876 1:1 1.08 0.30

F1 RES-02/Callypso and reciprocal 240 (2–3) 0

F2 RES-02/Callypso and reciprocal 1989 139 15:1 0.29 0.59

BC1F1RES-02/Callypso and reciprocal/Callypso 1532 482 3:1 1.22 0.27

F1RES-26/RES-02 and reciprocal 48 (1.0) 0

F2 RES-26/RES-02 and reciprocal 5000 (1.0) 0 a= except where indicated in parenthesis

The resistance of RES-26 to isolate P003 seemed to be conditioned by a single, partially dominant gene and the resistance of RES-02 which belong to group ‘A’ (resistant to R1 and P003) by two independent partially dominant genes. The genes for resistance to P003 in RES-26 were either closely linked, allelic, or identical to one of the two genes for resistance in RES 02. Resistance of RES-02 to R1 was conditioned by a single incompletely dominant gene. The genes for resistance to isolates R1 and P003 in RES-02 were either closely linked allelic or identical (Table 12.5). The cotyledonary leaves of each seedling responded independently when inoculated simultaneously each with a different isolate of the pathogen (Nashaat et al. 1997). Differential response and genes for resistance to Hyaloperonospora parasitica in Brassica juncea have been identified by Nashaat et al. (2004). The response of a wide range of Brassica juncea accessions to 14 isolates of Hyaloperonospora parasitica, 12 from India (IP00A, IP02, IP03, IP04, IP04A, IP05, IP05B, IP33, and IP33A were derived from B. juncea; IP09, IP14, and IP13A from B. rapa), and 2 from B. napus in the UK (R1 and P003) was screened. Sixteen differential host response groups to these isolates (classified as groups A–P) were identified. Groups ‘A’ and ’B’ expressed the widest resistance profiles to these isolates. Group ‘A’ was susceptible to isolates IP05 and IP05B, moderately resistant to isolate IP33, and resistant to all other isolates. Group ‘B’ was susceptible to isolates IP03, IP04, and IP04A and resistant to the other isolates. Putative homozygous lines resistant to all

14 isolates were selected from the F4 progeny of crosses involving lines RESBJ-200 from group ‘A’ (selection from cv. Kranti) and RESBJ-190 from group ‘B’ (selection from cv. Krishna). Both selections were selfed and tested for uniformity of reactions to all isolates for three generations. The resistance of RESBJ-200 to isolates IP00A, IP04A, and IP33A seems to be conditioned by single dominant genes. The resistance of RESBJ-190 to isolates IP00A, IP05B, and IP33A was also conditioned by single dominant genes. The gene for resistance to IP00A and IP33A in RESBJ-200 seems to be independent of the genes for resistance to the same isolates in RES BJ-190 (Table 12.6). The new genes for differential resistance to H. para- 12.5 Genetics of Host-Pathogen Relationship 237 P 1.0 2 X 0.0 Ratio 15:1 S 0 60 Brassica juncea accessions Brassica R 120 No. of plants 903 IsolateIP33A P 0.81 2 X 0.06 Ratio 15:1 S 0 65 ) progeny from crosses involving from crosses involving ) progeny R 120 No. of plants 1006 Isolate IP00A 1 F 1 P 0.92 0.33 (BC 1 2 X 0.01 0.95 Ratio 3:1 1:1 , and back cross F 2 S 604 0 0 111 , F 1 No. of plants R 1803 184 Isolate IP05B 349 126 P 0.62 0.18 2 0.25 X 1.82 3:1 Ratio 1:1 S 0 520 0 173 R 184 1521 IsolateIP04A No. of plants 355 199 isolates of F isolates parasitica Reaction to Hyaloperonospora RESBJ-200/ RESBJ-190/ 1 1 F F 1 1 RESBJ-200/ RESBJ-200/ 1 1 F F RESBJ-190 and reciprocal RESBJ-190 and reciprocal Generation and cross RESBJ-190//RESBJ-200 RESBJ-200//RESBJ-190 BC BC RESBJ-190 and RESBJ-200 (Nashaat et al. 2004 ) Table 12.6 Table 238 12 Host Resistance sitica will be of value in future studies of the genetics of the host-pathogen interaction and for breeding for disease resistance. In cabbage, resistance to downy mildew was dominantly controlled by two independent loci with Algarvia being heterozygous at both loci (R1r1 R2r2) and Penca de Chaves being homozygous recessive (r1r1 r2r2) (Caravalho and Monteiro 1996). Downy mildew resistance in Indian cauliflower crosses CC x HRS-4, and 3-5-1-1 × 244 was governed by single dominant gene PPA3, but in cross CC x 244, recessive epistasis was observed (Mahajan et al. 1995). According to Hoser-Krauze et al. (1995), in a cross P1246077 × P123210 to H. parasitica, Polish isolate proved to be dominant character. At the cotyledon stage, it was determined by four (in the broccoli-cauliflower line) or three (in the P/S) dominant complementary genes while at the stage of 4–5 leaves by one (in P123210), two (in P1246077), or three (in the broccoli-cauliflower line) additive dominant genes. In A. thaliana, alleles controlling the genotype-specific variation were identified at 19RPP (resistance to H. parasitica) loci, and most were mapped with varying degrees of resolution. RPP loci occur in several clusters, and at least two clusters also contain loci identified by involvement in recognition of other pathogens (Crute et al. 1994). Genetic crosses between A. thaliana genotypes RLD R and Wei-0(s) showed that resistance to H. parasitica was inherited through a monogenic dominant trait. The RPP 11 resistance gene was mapped by following the co-segregation of the resistance phenotype with RFLP markers in a mapping population of 254 F3 families derived from RLD x Wei-0 F2 individuals. Linkage analysis placed the RPP 11 resistance locus on chromosome 3 between marker m 249 (two recombinants) and marker g 2534 (six recombinants) (Joos et al. 1996). It is clear from Table 12.7 that resistance in different cruciferous crops against H. parasitica is governed by one to four dominant genes, closely linked allelic, additive genes, partial or partial dominant genes with epistasis, and gene dosage effects. Twelve host loci associated with different interaction phenotypes were postulated in part by studying segregations among F2 progeny from crosses of half-diallel among nine A. thaliana accessions. Advanced generations including recombinant inbred lines were used to confirm the identity of loci. It was suggested that a single cross between two host accessions could be used to characterize and map several loci associated with isolate-specific recognition of H. parasitica and distinct interaction phenotypes. Partial dominance and genetic epistasis are postulated as being common features of these parasitic symbols (Holub et al. 1994). As a prelude to cloning, the positions in A. thaliana genome were investigated for four loci (RPP1, RPP2, RPP4, and RPP7) of the ten that have been identified as associated with the genotype-specific recognition of the downy mildew pathogen H. parasitica (Tor et al. 1994). Segregating populations of A. thaliana generated from crosses between the susceptible accessions Col-0 and the resistant accessions Ws-0, Pr-0, OY-0, P0-1, Gc-1, Di-1, Ji-1, and Te-0 were screened with H. parasitica isolate N0C0. The genetic data were consistent with the presence of single resistance (RPP) loci in all of these accessions except Oy-0, in which resistance was inherited as a digenic trait. As a first step to molecular cloning, the map positions of four resistant loci were 12.5 Genetics of Host-Pathogen Relationship 239

Table 12.7 Inheritance of resistance in crucifers to H. parasitica Crucifer genotype Resistance gene References Broccoli PI 189028 One dominant gene Natti et al. (1967) and Vicente et al. (2012) Cabbage PI245015 One dominant gene Natti et al. (1967) and Vicente et al. (2012) Broccoli One recessive gene Hoser-Krauze et al. (1991) Broccoli PI 246077 x PI Three or four dominant Hoser-Krauze et al. (1995) 23210 complementary genes B. oleracea Quantitative genes Dickson and Petzoldt (1996) Chinese cabbage One dominant gene Niu et al. (1983) Chinese cabbage Additive genes Yuen (1991) Radish cvs. Tokinaski and Two dominant genes Bonnet and Blancard (1987) Okura B. rapa Partial resistance Leung and Williams (1983) B. napus cv. Cresor One dominant gene Lucas et al. (1988) B. oleracea Two or three dominant genes Moss et al. (1988) B. napus lines One dominant gene Nashaat et al. (1995a, b, 1996) RES 01-1-4 and RES-26 One partial dominant gene Nashaat et al. (1997) B. napus RES-02 B. napus Two partial dominant genes, closely Nashaat et al. (1997) linked, allelic Arabidopsis thaliana Partial dominant epistasis and gene Holub et al. (1994) dosage effects B. oleracea Two dominant genes Caravalho and Monteiro (1996) One dominant gene Vicente et al. (2012) B. juncea One dominant gene Nashaat et al. (2004) RESBJ-200, RESBJ-190 One dominant gene Nashaat et al. (2004) (B. juncea) Indian cauliflower One dominant gene Mahajan et al. (1996) A. thaliana One dominant gene, two dominant Reignault et al. (1996) and genes Joos et al. (1996) Broccoli Partial resistance Jensen et al. (1999a) A. thaliana Four tightly linked genes Botella et al. (1998) determined. These have been designated RPP 14.1 from Ws-0, RPP 14.2 from Pr-0 and RPP 14.3, and RPP 5.2 from Oy-0. RPP 14.1 was mapped to a 3.2 cm interval on chromosome 3 that is linked to a region between the markers Gl-1 and M 249 known to contain other H. parasitica specifications. RPP 14.2 from Pr-0 and RPP 14.3 from Oy-0 were also positioned in this interval. RPP 14.1 and RPP 14.2 showed linkage of <0.05 cm suggesting possible allelism. The second RPP locus from Oy-0, RPP 5.2 was located on chromosome 4 and exhibited strong linkage (<2 cm) to RPP 5.1, a locus previously identified in theArabidopsis accession Landsberg erecta (Reignault et al. 1996). 240 12 Host Resistance

According to Botella et al. (1998) in the Arabidopsis accessions Wassilewskija, the RPP 1 region on chromosome 3 contains four genetically linked recognition specificities conditioning resistance to different isolates of H. parasitica. Three of the four tightly linked genes in this region, designated RPP-1- WSA, RPA-1-WsB, and RPP-1-WsC, encode functional products of the NBS-LRR (nucleotide-binding site-leucine-rich repeat) R protein class. They possess a TIR (Toll, interleukin-1, resistance) domain that is characteristic of certain other NBS-LRR-type R protein, but, in addition, they have unique hydrophilic or hydrophobic N termini. Together, the three RPP-1 genes account for the spectrum of resistance previously assigned to the RPP-1 region and, thus, comprise a complex R locus. The distinct but partially overlapping resistance capabilities conferred by these genes are best explained by the hypothesis that each recognizes a different pathogen avirulence determinant.

12.5.1 Seedling and Adult Plant Resistance to Downy Mildew

Resistance to H. parasitica has been described in Brassica oleracea at the seedling stage (Natti et al. 1967; Monteirio and Williams 1989; Thomas and Jourdain 1992) and in the first true leaves (Thomas and Jourdain 1990). Seedling resistance to H. parasitica has been ascribed to a dominantly inherited single gene in Chinese cabbage (Niu et al. 1983), broccoli, and cabbage (Natti et al. 1967). Resistance in cauliflower seedlings has been characterized as a recessive trait based on a single gene (Hoser-Krauze et al. 1984), but within B. oleracea the action of several genes may also control resistance (Hoser-Krauze et al. 1995). Adult plant resistance to downy mildew has been reported in field trials in cauliflower (Sharma et al. 1991; Mahajan et al. 1995) and has been ascribed to a single gene with dominant effect (Mahajan et al. 1995). From a breeding point of view, assessment of the correspondence between seedling and adult plant resistance may indicate whether resistance at the seedling stage can be used as a reliable indicator of adult plant resistance. Parity between resistance of cotyledons and the first true leaves has been reported previously (Natti 1958; Monteirio and Williams 1989). Seedlings of six cauliflower cultivars (Brassica oleracea convar. botrytis var. botrytis) were assessed for resistance to a Danish isolate of Hyaloperonospora parasitica, under controlled conditions. Resistance, characterized by restricted sporulation and necrotic dark flecks at the inoculation site on the cotyledons, was expressed in the hybrids 9306 F1, 9311 F1, and the open pollinated cultivar Perfection. Testing of the parent lines and F2 generations of the two resistant hybrids suggested that resistance was a dominantly inherited trait controlled by a single gene. Inoculation of the cultivars with seven isolates, from different geographic origins, showed that the resistance was isolate-specific. The two hybrid cultivars expressing cotyledon resistance and two hybrids expressing susceptibility were assessed for adult plant resistance under field conditions. The AUDPC (area under disease progress curve), based on disease incidence and severity, revealed significant differences between the cultivars. At harvest, the cultivars exhibited significantly different levels of defo- 12.5 Genetics of Host-Pathogen Relationship 241

liation and curd attack. The cultivars 9306 F1 and 9311 F1 showed high levels of resistance in all assessments, whereas the two cultivars exhibiting susceptibility at the seedling stage, i.e. 9304 F1, and 9305 F1, also exhibited susceptibility through the adult plant stage. Thus, the resistance exhibited under field conditions resembled that identified at the seedling stage under controlled conditions. The results suggest that cotyledon resistance similar to that described could provide resistance through ought the adult plant stage, including curds (Jensen et al. 1999a).

12.5.2 Inheritance of Partial Resistance to Downy Mildew

Latent period and sporulation capacity are two major components of partial resistance and governed by the same mechanism in the interaction between broccoli (B. oleracea) and H. parasitica. Twenty doubled haploid broccoli lines from breeding material were evaluated for resistance to H. parasitica at the seedling stage. All lines supported sporulation of the pathogen but to varying extents and intensities. Partial resistance of the more resistant lines ‘br8’ and ‘br9’ reduced conidia production on cotyledons by 50–70% compared with the most susceptible lines. Inoculation of the two most resistant lines with 13 isolates of different geographic origins revealed that ‘br9’ showed a rather uniform level of resistance to all isolates, while ‘br8’ showed some isolate specificity. Partial resistance was evaluated in six of the broccoli lines in a half-diallel set of crosses. Disease assessment of seedlings showed that additive genetic effects explained 45.8 and 31.8% of the total variation of sporulation score and conidia production, respectively. This suggests that recurrent selection for partial resistance to H. parasitica in early generation inbreds or in populations of broccoli will be efficient to obtain cotyledon resistance (Jensen et al. 1999b). Seedling resistance conditioned by a single major gene is isolate-specific and may lose the effect through selection of virulent types of the pathogen, if resistant cultivars are produced on large areas (Lucas et al. 1988). Partial quantitative disease resistance by definition provides reduced host invasion without complete protection (Parlevliet 1992) and is generally anticipated to be based on several genes. It may be a source of durable resistance with less isolate specificity than resistance based on single genes (Parlevliet 1992). In Brassica oleracea, partial resistance to H. parasitica at the seedling stage has been reported in Portuguese coles (Monteirio and Williams 1989; Dias et al. 1993) and in gene bank accessions of cabbage (Thomas and Jourdain 1992) and broccoli (Thomas and Jourdain 1990). Jensen et al. (1999b) characterize partial resistance to H. parasitica at the cotyledon stage within 20 doubled haploid broccoli lines originating from a wide range of breeding material. Inheritance of the resistance was analysed in progeny from a half-diallel set of crosses (Jensen et al. 1999b). 242 12 Host Resistance

12.5.3 Molecular Basis of Downy Mildew Resistance

Arabidopsis a cruciferous weed has emerged as an important model for investigat- ing the molecular basis of disease resistance in plants. Numerous isolates of H. parasitica along with isolates of the bacterium Pseudomonas syringae have been particularly useful for characterizing numerous naturally variable genes that determine the isolate specificity of resistance (so-called R genes) as well as associated ‘downstream’ defence genes revealed by analyses of artificially derived mutants that exhibit either enhanced susceptibility or enhanced resistance (Holub and Beynon 1997; Shapiro 2000). More than 20 wild-type specificities of resistance to H. parasitica (RPP) genes have been identified which are distributed among the five chromosomes of Arabidopsis (Holub 1997). Genes from four RPP loci have so far been published, and they all encode receptor-like proteins containing a conserved nucleotide-binding motif and a highly variable leucine-rich repeat domain (so- called NB-LRR genes) (Bittner-Eddy et al. 2000; Botella et al. 1998; McDowell et al. 1998; Parker et al. 1997). They include examples of functional alleles for different specificities of downy mildew resistance at either a single gene locus in the case of RPP13 or at a complex locus (multiple genes in a region of <1 cm) in the case of RPP1. The simple locus RPP8/HRT1 is particularly fascinating as an example in which different functional alleles have been described which confer resistance to widely divergent pathogens (Cooley et al. 2000; McDowell et al. 1998). White rust resistance loci on three Arabidopsis chromosomes are closely linked to downy mildew (H. parasitica) resistance loci. Two accessions of Arabidopsis thaliana (Ksk-1 and Ksk-2) were used to identify and map three loci (RAC1, RAC2, and RAC3) of genes that confer resistance to Albugo candida (white rust). The phenotypes associated with these genes were classified as either FN (necrotic flecks on upper surface of cotyledons and no blisters) for RAC2 and RAC3 or FYN (flecks surrounded by yellowing and no blisters) for RAC1. Both phenotypes exhibited rapid death of host cells penetrated by the parasite (hypersensitive response), with callose deposition commonly encasing the haustorium. F6 recombinant inbred lines were produced specifically for the purpose of mapping each RAC locus relative to molecular markers. Dominant resistance at the locus RAC1 in Ksk-1 was previously mapped to chromosome 1 between RFLP markers, m253, and m254, co-segregating with downy mildew resistance specificity RPP9 in the accession Wei-0. Borhan et al. (2001) supported a fine-scale map interval and co-segregating markers for this locus, which in turn enabled mapping of a previously unnoticed source of resistance in Ksk-1 designated RAC3 that exhibits an FN phenotype hyperstatic to the FYN phenotype of RAC1. RAC3 is closely linked to the RPP8/HRT on chromosome 5, a locus which contains specificities for resistance to downy mildew and turnip crinkle virus. Recombinant inbreds also enabled mapping of recessive resistance at RAC2 in Ksk-2 to the bottom arm of chromosome 3, in the 6 cm interval between two downy mildew resistance loci (RPP1 and RPP13). Phenotypic characterization, and molecular mapping of the Arabidopsis thaliana locus RPP5, determining disease resistance to Hyaloperonospora parasitica was 12.5 Genetics of Host-Pathogen Relationship 243 observed by Parker et al. (1993). An isolate of H. parasitica (denoted NOCO2) was identified that infected Arabidopsis plants of the land race Columbia (Col-0) but not plants of land race Landsberg erecta (La-er). Segregation analysis of F2 plants derived from a La-er x Col-0 cross established that resistance was inherited as a single locus, denoted RPP-5. Macroscopic and microscopic examination of inoculated La-er and Col-0 cotyledons showed that restriction of fungal growth in La-er was accompanied by massive callose accumulation and death of plant cells in direct contact with points of attempted fungal penetration. La-er x Col-0 F1 plants exhibited an intermediate resistance response in all aspects of fungal development, indicating that RPP-5 is semi-dominant in its action. F3 recombinant inbred lines generated between La-er and Col-0 were used to map RPP-5 to a narrow interval (<1.1 cm) on chromosome 4, utilizing existing restriction fragment length polymorphic (RFLP) markers and newly generated random amplified polymorphic DNA (RAPD) markers. The data provide a basis for the isolation of the RPP-5 locus by positional cloning as a first step towards understanding recognition specificity in plant-pathogen interactions at a molecular level (Fig. 12.3).

12.5.4 Mutation Approach to Identify Resistance Genes

A mutational approach has been adopted to identify genes that are necessary for resistance mediated by RPP-5 and RPP-14 and to attempt separation of RPP-14 from the other closely linked RPP gene specificities. Parker et al. (1996) screened mutagenized populations of Ler-0 and Ws-0 for mutations that cause a change from NOCO2 resistance to susceptibility. They have described a recessive mutation of Ws-0 called eds1 (for enhanced disease susceptibility), which abolishes the resistance mediated by RPP-14 as well as by other linked and unlinked RPP genes present in the Ws-0 background. This mutation also partially suppresses resistance of Ws-0 to five Brassica oleracea-infecting isolates of H. parasitica to which all Arabidopsis ecotypes so far tested exhibit resistance, implicating a possible common functional role for the EDS1 protein in downy mildew resistance in Arabidopsis and Brassica plants. The interaction between Arabidopsis and H. parasitica provides an attractive model pathosystem to identify molecular components of the host that are required for genotype-specific recognition of the parasite. These components are the so- called RPP genes (resistance to H. parasitica). Mutational analysis of ecotype Wassilewskija (Ws-0) revealed an RPP- nonspecific locus called EDS1 (for enhanced disease susceptibility) that is required for the function of RPP genes on chromosomes 3 (RPP1/RPP-14 and RPP-10) and 4 (RPP-12). Genetic analysis demonstrated that the eds1 mutation is recessive and is not a defective allele of any known RPP gene, mapping to the bottom arm of chromosome 3 (13 centimorgans below RPP1/RPP14). Phenotypically, the Ws-eds1 mutant seedlings supported heavy sporulation by H. parasitica isolates that are each diagnostic for one of the RPP genes in wild-type Ws-0; none of the isolates is 244 12 Host Resistance

g3843

m226

g2616

pCIT-d23 0.9cM

(SD = 0.018)

g 10086 m326 m226 RPP5, OPC18540 g4539 ag RPP5, OPC18540

0.34cM

(SD = 0.028)

g3845, g4539

10cm g8300

g3713

Fig. 12.3 Linkage map of Arabidopsis chromosome 4 showing location of RPP-5, relative to cosmid (g), λ (m) RFLP markers, and RAPD (OP) markers, based on the segregation analysis of La-er x Col-0 Rls. (a) Mapping data derived from segregation analysis of RFLP markers on 100Rls and RAPD markers on 50Rls; (b) map position of RPP-5 relative to closely linked markers, from the analysis of 289 Rls (Parker et al. 1993) 12.5 Genetics of Host-Pathogen Relationship 245 capable of sporulating on wild-type Ws-0. Ws-eds1 seedlings exhibited enhanced susceptibility to some H. parasitica isolates when compared with a compatible wild-type ecotype, Columbia, and the eds1 parental ecotype, Ws-0. This was observed as earlier initiation of sporulation and elevated production of conidia. Surprisingly, cotyledons of Ws-eds1 also supported low sporulation by five isolates of H. parasitica from Brassica oleracea. These isolates were unable to sporulate on >100 ecotype of Arabidopsis, including wild-type Ws-0. An isolate of Albugo candida from B. oleracea also sporulated on Ws-eds1, but the mutant exhibited no alteration in phenotype when inoculated with several oomycete isolates from other host species. The bacterial resistance gene RPM1, conferring specific recognition of the avirulence gene avrB from Pseudomonas syringae pv. glycinea, was not compromised in Ws-eds1 plants. The mutant also retained full responsiveness to the chemical inducer of systemic acquired resistance, 2,6-dichloroisonicotinic acid; Ws-eds1 seedlings treated with 2,6-dichloroisonicotinic acid became resistant to the Ws-0 compatible and Ws-0 incompatible H. parasitica isolates, Emwa1 and NOCO2, respectively. The EDS1 gene appears to be a necessary component of the resistance response specified by several RPP genes and is likely to function upstream from the convergence of disease resistance pathways in Arabidopsis (Parker et al. 1996). While studying the Arabidopsis downy mildew resistance gene RPP5, Parker et al. (1997) found that it encodes a protein that possesses a putative nucleotide- binding site and leucine-rich repeats, and its product exhibits striking structural similarly to the plant resistance gene products N and L6. Like N and L6, the RPP5 N-terminal domain resembles the cytoplasmic domains of the Drosophila Toll and mammalian interleukin-1 transmembrane receptors. In contrast to N and L6, which produce predicted truncated products by alternative splicing, RPP5 appears to express only a single transcript corresponding to full-length protein. However, a truncated form structurally similar to those of N and L6 is encoded by one or more other members of the RPP5 gene family that are tightly clustered on chromosome 4. The organization of repeated units within the leucine-rich repeats encoded by the wild-type RPP5 gene and an RPP5 mutant allele provides molecular evidence for the heightened capacity of this domain to evolve novel configuration and potentially new disease resistance specificities. Nucleotide sequence data have been submitted to the GenBank database under the accession number U97106.

12.5.5 Genetics of Multiple Disease Resistance

Multiple disease resistance (MDR) in B. rapa to three pathogens showed heritable genetic variation or resistance to each pathogen and a positive genetic correlation between resistance to H. parasitica and Leptosphaeria maculans (Mitchell-Oldts et al. 1995). However, in a later study, Mitchell-Olds and Bradley (1996) found that Hyaloperonospora-resistant genotypes grew 6% slower than Hyaloperonospora- susceptible genotypes in pathogen-free environments, indicating a significant genetic fitness cost to Hyaloperonospora resistance. 246 12 Host Resistance

Genetic costs of resistance to pathogens may be an important factor maintaining heritable variation for resistance in natural populations. Pleiotropic fitness trade- offs occur when genetic resistance causes reduction in other components of fitness. Although costs of resistance have an important influence on plant-pathogen interactions, few previous studies have detected pleiotropic costs of resistance in the absence of confounding effects of linkage disequilibrium. To avoid this potential problem, Mitchell-Olds and Bradley (1996) performed artificial selection experiments on resistance to two fungal pathogens, Leptosphaeria maculans and Hyaloperonospora parasitica, and compared growth rates of resistant and susceptible genotypes of Brassica rapa in the absence of pathogens. Leptosphaeria resistance had no effect on growth rate, indicating cost-free defence. In contrast, Hyaloperonospora-resistant genotypes grow 6% slower than Peronospora- susceptible genotypes in pathogen-free environments, indicating a significant genetic fitness cost to Hyaloperonospora resistance. Such genetic trade-offs could maintain genetic variation in the wild. Another factor that might explain heritable variation for resistance is ecological trade-offs, in which genetic resistance to one species causes susceptibility to another. Such ecological trade-offs do not exist for the pathogens studied in this system. Mitchell-Olds and Bradley (1996) observed that genes conferring resistance can have pleiotropic effects on other aspects of plant performance (Simms 1992). If resistant plants have reduced growth or fecundity in the absence of pathogen attack, then selection may favour intermediate levels of disease resistance. When allocation to constitutive resistance mechanisms varies among genotypes, then resistant genotypes may experience a fitness benefit in the presence of a pathogen, relative to susceptible plants. However, resistant plants would have reduced relative fitness when pathogen attack is reduced or absent, because resources would be expended on unneeded defensive mechanisms. For constitutive defence pathways, a genetic cost to resistance is indicated by a significant negative genetic correlation between growth or fitness in a pathogen-free environment and resistance measured in the presence of pathogens. Few published studies provide clear evidence for pleiotropy causing genetic fitness costs. Berenbaum et al. (1986) showed a negative genetic correlation between insect resistance and fecundity in Pastinaca sativa. Han and Lincoln (1994) found a significant negative genetic correlation between growth rate and levels of secondary metabolites in Diplacus aurantiacus. In Escherichia coli, mutations causing resistance to virus T4 carry pleiotropic costs of resistance, which can be ameliorated by epistatic modifiers (Lenski 1988a; b). Several studies from habitually inbreeding species or near-isogenic lines are consistent with the existence of pleiotropic fitness costs (Bergelson 1994; Brinkmamn and Frey 1977; Burdon and Muller 1987; Chaplin 1970; Frey and Browning 1971; Simons 1979), but linkage disequilibrium and pleiotropy are confounded in these experiments. Additional evidence from ecological and physiological studies is also suggestive of genetic costs of resistance (Simms 1992). 12.5 Genetics of Host-Pathogen Relationship 247

12.5.6 Disease Resistance Increases Competitive Ability of Host Plants

Disease resistance is believed to play a role in the dynamics of plant communities and plant populations. The competitive ability of two Arabidopsis thaliana genotypes, one exhibiting resistance, and the other exhibiting susceptibility to an isolate of H. parasitica was determined by Damgaard and Jensen (2002). The A. thaliana genotypes were grown in competition experiments under controlled conditions at three densities in pure and mixed stands both in the presence and absence of H. parasitica. After seed set, the dry weight of the plants was determined, and the amount of seeds produced was calculated from an established linear relationship. The two genotypes were found to be ecologically different; the susceptible genotype was the strongest competitor (significantly so in the absence of the disease), whereas the resistant genotype produced most seeds. The competitive ability of the disease-resistant A. thaliana genotype increased significantly in the presence of aH . parasitica isolate when competing with the susceptible A. thaliana genotype, whereas the disease did not affect the competitive ability of the susceptible genotype significantly. Assuming that the population only consisted of the two genotypes and mainly was affected by the disease, it was possible to predict the probabilities of four long-term ecological scenarios. Without the disease, the most likely long- term ecological scenario was that the two A. thaliana genotypes would coexist, whereas in the presence of the disease, the resistant genotype most likely would outcompete the susceptible genotype (Fig. 12.4).

12.5.7 Expression of Age-Related Resistance (ARR) to Downy Mildew

Disease symptoms in the field start on the lower leaves and progress upwards (Natti et al. 1956; Coelho et al. 1998; Jensen et al. 1999a). This disease pattern could be attributed to environmental differences, variation in inoculum availability, and changes in leaf resistance. Downy mildew resistance is affected by plant development in Brassica oleracea. The resistance of mature broccoli plants having eight or more leaves is independent of the resistance of young seedlings (Dickson and Petzoldt 1993), and the plants can be susceptible at the cotyledon stage and resistant at the adult stage (Coelho and Monteiro 2003). In ‘Couve Algarvia’, resistance at cotyledon and adult plant stages is under the control of two different genetic systems (Monteirio et al. 2005). Agnola et al. (2003) used leaf discs collected from 17 leaf positions in 10–18-week-old Brassica plants to show that incompatible reactions occur regardless of leaf position, but with compatible reactions, sporulation intensity varies with the position of the leaf on the stem. The increasing resistance to pathogens with plant or leaf ageing is a form of resistance referred to as age- related resistance (ARR) (Kim et al. 1989). ARR in Arabidopsis is a 248 12 Host Resistance

Fig. 12.4 The Bayesian posterior distribution of the competition coefficient of twoA . thaliana genotypes, Nd-1 (susceptible) and C24 (resistant), competing against each other. Solid line: without pathogens. Dashed line: with pathogens. Percentiles (2.5%, 50%, and 97.5%) in the posterior distributions: cC24, without pathogen (−0.23, 0.01, 0.31), with pathogen (0.40, 0.92, 1.60); cNd-1, without pathogen (0.99, 1.52, 2.19), with pathogen (1.28, 1.76, 2.34). The Bayesian posterior distribution of a parameter provides information on our degree of beliefs in the different possible values of the parameter. The mode of the posterior distribution corresponds to the maximum likelihood value, and increasing variance of the distribution corresponds to an increasing degree of uncertainty about the true value of the parameter (increasing experimental variation) (Damgaard and Jensen 2002) developmentally regulated and environmentally sensitive defence response to Pseudomonas syringae (Kus et al. 2002). ARR is frequently reported in mature plants in different plant-pathogen interactions: potato-Phytophthora infestans (Warren et al. 1971), lettuce-Bremia lactucae (Dickinson and Crute 1974), tobacco- Peronospora tabacina (Reuveni et al. 1986; Cohen et al. 1987; Wyatt et al. 1991; Hugot et al. 2004), and pepper-Phytophthora capsici (Kim et al. 1989). Coelho et al. (2009) conducted experiments under controlled environment conditions, using inoculated leaf discs, to determine the influence of leaf position, plant age, and leaf age on the expression of resistance to downy mildew in various Brassica oleracea genotypes. The upper leaves were more resistant than the lower leaves when 7–19-week-old plants of broccoli and tronchuda cabbage were tested. Broccoli lines ‘PCB21.32’ and ‘OL87123-2’ were fully susceptible at the cotyledon stage and showed a clear resistance increase from lower to upper leaves at 6 weeks, and ‘PCB21.32’ was fully resistant 16 weeks after sowing. Immature leaves were more resistant than adjacent fully expanded mature leaves. Susceptibility increased with leaf age when the same leaf was tested at 2–4-week intervals. Leaf age and upper-leaf position on the stem had opposite effects on dis- 12.5 Genetics of Host-Pathogen Relationship 249 ease score, since younger leaves collected from lower positions and older leaves collected from upper positions tended to score similarly in compatible interactions. The progression of downy mildew from the base of the plant upwards on B. oleracea in the field could be due to differences in leaf resistance in addition to environmental variation. To maximize the expression of a compatible reaction in adult plants, lower leaves of Brassica plants that are at least 12 weeks old should be used. In order to clarify these questions, the use of leaf discs is adequate to assess disease resistance variation within adult plants because it is possible to compare, under the same environmental conditions, the resistance of leaves of different ages or leaves collected from different positions. The use of leaf disc evaluation methods is more informative than field or greenhouse tests, and that can also be used to predict – host response to natural epidemics in the field (Coelho et al. 2009).

12.5.8 Different Requirements for Disease Resistance Genes

Mutational analyses in Arabidopsis have identified other wild-type genes that are required for R gene-mediated resistance (Glazebrook et al. 1997). A mutation in NDR1 (non-race-specific disease resistance) abolished resistance conferred by RPS2 and RPS5 to Pseudomonas syringae expressing avrRpt2 and avrPph3, respectively, as well as a dual specificity resistance encoded by RPM1 to the bacterial genes avrB and avrRpm1 (Century et al. 1995). ndr1 plants also were compromised in RPP gene-mediated resistance to several incompatible H. parasitica isolates, suggesting that the wild-type NDR1 protein may function at a common point downstream from the perception of these prokaryotic and eukaryotic pathogens (Century et al. 1995). Another resistance signalling component is encoded by EDS1 (enhanced disease susceptibility) as observed by Parker et al. (1996). A mutation in EDS1 abolished resistance conferred by several RPP loci but had no effect on RPM1- specific resistance, suggesting a function before the convergence of downstream pathways (Parker et al. 1996). The Arabidopsis genes EDS1 and NDR1 were shown previously by mutational analysis to encode essential components of race-specific disease resistance. Aarts et al. (1998) have examined the relative requirements for EDS1 and NDR1 by a broad spectrum of resistance (R) genes present in three Arabidopsis accessions (Columbia, Landsberg erecta, and Wassilewskija). There is a strong requirement for EDS1 by a subset of R loci (RPP2, RPP4, RPP5, RPP21, and RPS4), conferring resistance to the Hyaloperonospora parasitica and to Pseudomonas bacteria expressing the avirulence gene avrRps4. The requirement for NDR1 by these EDS1- dependent R loci is either weak or not measurable. Conversely, three NDR1- dependent R loci, RPS2, RPM1, and RPS5, operate independently of EDS1. Another RPP locus, RPP8, exhibits no strong exclusive requirement for EDS1 or NDR1 in isolate-specific resistance to H. parasitica, although resistance is compromised weakly by eds1. Similarly, resistance conditioned by two EDS1-dependent RPP genes, RPP4 and RPP5, is impaired partially by ndr1, implicating a degree of path- 250 12 Host Resistance way crosstalk. These results provide compelling evidence for the preferential utilization of either signalling component by particular R genes and thus define at least two disease resistance pathways. The data also suggest that strong dependence on EDS1 or NDR1 is governed by R protein structural type rather than pathogen class.

12.5.9 Differential Expression of Downy Mildew Resistance Genes

So far in all, Xiao et al. (2016) identified 129 transcript-derived fragments (TDFs), of which 121 TDFs were upregulated and eight were downregulated using cDNA- AFLP (Turck et al. 2004). By BLAST searching in the Brassica database, these TDFs were classified according to their different functions. The functional catego- rization showed a complex linkage between proteins encoded by the TDFs. Information obtained from this study may provide a foundation for better understanding defence mechanisms of the non-heading Chinese cabbage with H. parasitica incompatible interaction. Several transcripts encoding the group of PR proteins were differentially expressed in the interaction. TDF3 (β-1,3-glucanase) was induced within 24 h.p.i., and its expression peaked at 48 h.p.i. The expression levels of TDF8 (Hapless 8), TDF16 (a thaumatin-like protein), and TDF28 (PR 1-like protein) were induced within 24–72 h.p.i. Chen et al. (2008) cloned the full length of β-1,3-glucanase, hapless 8, and PR 1 genes and analysed their expression patterns in response to H. parasitica infection in ‘Suzhou Qing’ cultivar of non- heading Chinese cabbage. The accumulations of these two transcripts were upregulated during the infection period, suggesting that these proteins may participate in the defence reaction for non-heading Chinese cabbage against H. parasitica. They also found that expression of TDF39 (ATMT-1) peaked at 48 h.p.i. In rice and barley, MT2A genes were induced by stresses such as drought, cold treatment, and wounding or in response to pathogen attacks (Ozturk et al. 2002; Degenhardt et al. 2005; Jin et al. 2006). The products of homologous MT scavenged the reactive oxygen species (ROS), such as OH to H2O (Akashi et al. 2004). Evidences suggest that the generation of ROS occurs at early stage in the plant-pathogen interaction. Rapid accumulation of ROS causes oxidative burst that results in hypersensitive cell death and cell wall cross-link (Das et al. 2008). Several signal transduction-related proteins are involved in the plant-fungus interactions. Xiao et al. (2016) identified many TDFs related to the signal transduction, such as TDF42 (catalase 3), TDF43 (2-Cys PrxB), TDF45 (ATP binding), TDF50 (ATRER1A), TDF58 (nitrilase), TDF48 (BcCAM3_A04, calcium ion binding), and TDF68 (WRKY DNA-binding protein). Calcium binding-like proteins may have a role in signalling pathways against pathogens and wounding. A number of downstream targets of calmodulin (CaM), including nitric oxide synthase, barley MLO protein, maize Ca2+-CaM, and transcriptional regulators, are involved in plant responses to pathogens. Given that calcium ion-binding proteins are important modulators of defence response in pathways for 12.5 Genetics of Host-Pathogen Relationship 251 pathogen sensing in plants, the CAM 3 gene could have a special role as Ca2+ sensors during the plant immune response to the fungus H. parasitica (Xiao et al. 2016). WRKY proteins are signal transcriptional factors recognizing the TTGAC (C/T) W-box elements in the promoters of a large number of plant defence-related genes (Turck et al. 2004). Many of the WRKY genes are upregulated particularly in pathogen-infected, wounded, or abiotic-treated plants (Pandey and Somssich 2009). In this study, expression of WRKY DNA-binding protein peaked at 24 h.p.i., suggesting that the possible role of WRKYs is in the regulation of the genes associated with plant defence responses. However, Xiao et al. (2016) found that expression pattern of TDF60 (WRKY gene) determined by qRT-PCR was inconsistent with that of cDNA-AFLP. The inconsistence may be caused by different paralogues in the genome. An ethylene response factor (BcERF1_A01, TDF93), a regulator of ethylene responses after pathogen attack in Arabidopsis (Berrocal-Lobo et al. 2002), may have a key role in the non-heading Chinese cabbage-H. parasitica interaction. Previous studies have been demonstrated that ERFs are involved in regulating the expression of the defence-related genes during the disease resistance responses (Singh et al. 2002; Cao et al. 2005). Xiao et al. (2016) found that TDF91 (BcABCG36_A07, ATP-binding cassette g36) was inhibited after inoculation by cDNA-AFLP analysis. However, its expression was induced weakly at 24 h.p.i., peaked at 48 h.p.i., and decreased weakly at 72 h.p.i. afterwards by qRT-PCR analysis. TDF100 (BcEDM1_A09) coding for an enhanced downy mildew 1 homologue was found to be induced during the infection period. Its relationship with the fungi, bacteria, and viruses has been identified to be regulators of R gene-mediated resistance in other crop species. Recent studies have revealed that EDM1 homologue gene SGT1 is required for pathogen-induced disease-associated cell death during both compatible and incompatible interactions in tobacco (Wang et al. 2010). Energy metabolism has an important role in plant-pathogen interaction. The photosynthetic carbon cycle (PCC) is part of the dark reactions of photosynthesis and can be roughly divided into three steps: carboxylation, reduction reaction, and regeneration of RuBP. Xiao et al. (2016) found that some TDFs relating to energy metabolism were downregulated, such as TDF76 (chlorophyll a/b binding protein), TDF78 (SIT4 phosphatase-associated family protein), and TDF82 (PP2C-related protein), whereas some were upregulated, such as TDF79 (receptor-like protein 51), TDF80 (NADH-ubiquinone oxidoreductase), TDF83 (respiratory burst oxidase protein), and TDF85 (rubisco small subunit 1b). Previous reports have identified that they are involved in PCC cycle, for example, TDF86 (reduction of transketo- lase) inhibited ribulose-1, 5-bisphosphate regeneration, and photosynthesis. The expression of energy metabolism-related genes is induced and/or suppressed in photosynthesis during abiotic and biotic stresses. The PCC cycle could provide protection function in energy metabolism during non-heading Chinese cabbage against H. parasitica (Xiao et al. 2016). A number of genes related to protein-protein interaction were induced after inoculation, such as TDF110 (BcTPR12_A07, tetratricopeptide repeat protein) and 252 12 Host Resistance

TDF111 (BcZF_A01, zinc-finger family protein). Of which, the gene expression of PAT is induced in the presence of ozone in Arabidopsis (Conklin and Last 1995). The tryptophan biosynthetic enzymes, including anthranilate synthase (ASA) and PAT, are co-ordinately upregulated at both the messenger RNA and protein level during biotic and abiotic stresses (Zhao and Last 1996). Xiao et al. (2016) found that one of BcZF orthologous to A. thaliana was induced after inoculation. Rizhsky et al. speculated that a zinc-finger protein is required for the expression of ascorbate peroxidase, which provides some measure of resistance for plant during oxidative stress. The pathogen-induced accumulation of these protein-protein interaction- related genes suggested that these genes may be involved in some defence mechanisms against H. parasitica indirectly (Xiao et al. 2016). Using the cDNA-AFLP method, Xiao et al. (2016) also detected several unknown functional genes. Their biological role is still unclear. Xiao et al. (2016) examined gene expression patterns in an incompatible interaction between non-heading Chinese cabbage ‘Suzhou Qing’ and the downy mildew pathogen. They obtained 129 TDFs with different expression patterns and classified functional categories using cDNA-AFLP. Fifteen TDFs were randomly selected for validation of cDNA- AFLP expression patterns using qRT-PCR. Results showed that reliability of cDNA- AFLP is suitable for detecting differentially expressed genes. Among the 15 TDFs, four TDFs are related with fungal resistance, namely, TDF14 (BcLIK1_A01), TDF42 (BcCAT3_A07), TDF75 (BcAAE3_A06), and TDF88 (BcAMT2_A05). They further compared expression patterns in ‘Suzhou Qing’ and ‘Aijiao Huang’ using qRT-PCR. Results showed that the four genes displayed similar expression trend in the two lines. Importantly, the expression of genes in the resistant line is higher than that in susceptible line. These gene expression patterns and their putative functions may provide insight in understanding the non-heading Chinese cabbage-downy mildew incompatible interaction. It may also provide a foundation for better understanding molecular mechanisms and can be beneficial in selecting candidate resistance genes for the incompatible interaction between non-heading Chinese cabbage and H. parasitica. Further research is needed to study the comparison between compatible and incompatible interactions to identify novel and common genes that regulate non-heading Chinese cabbage-downy mildew pathosystem.

12.5.10 Cloning of Major Resistance Genes

With the emergence and growth of modern molecular biology and cloning techniques, what had long been fantasy became reality. Thus, map-based or positional cloning strategies to isolate ‘R’ genes could be begun in the early 1990s (Parker et al. 1993). As more ‘R’ genes were mapped, it became clear that these were not randomly distributed on the chromosomes, but that most fell into larger or smaller clusters, often interspersed with major resistance genes which were active against other pathogens. This led to the coining of the term MRC loci (major recognition gene complexes) with MRC-A, MRC-B, MRC-F, MRC-H, and MRC-J on 12.5 Genetics of Host-Pathogen Relationship 253

RPP1* MRC-F RPP2 MRC-H m583 RPP3 Oy-0/CALA RPP4* MRC-H RPP5* MRC-H RPP6 MRC-B m241 RPP7 MRC-B MRC-A RPP8* MRC-J RPP19 RPP9 chromosome 1 RPP20 RPP10 MRC-F RPP9 RPP11 MRC-F m253 g11 RPP12 MRC-H RPP13*MRC-F tt3 RPP14 MRC-F RPP28 m249 MRC-F MRC-J RPP15 not assigned m280 m226 m435 RPP16 MRC-F RPP17 MRC-F MRC-H RPP18 MRC-H MRC-B m600 RPP19 chromosome 2 RPP20 chromosome 2 RPP21 MRC-J RPP22 MRC-J RPP23 MRC-J RPP24 MRC-J RPP25 MRC-B RPP26 MRC-F RPP27 MRC-B RPP28 chromosome 2

Fig. 12.5 Distribution of the mapped RPP genes along the five chromosomes of Arabidopsis thaliana. To the left: a numerical list of the known 27 RPP genes with their chromosomal or MRC location given, where known. RPP3 is not mapped yet; thus, the Arabidopsis ecotype and the H. parasitica isolate are given. Underlined RPP genes have been renamed once. RPP genes with an asterisk have been cloned. To the right: graphical representation of the five Arabidopsis chromosomes with chromosome 1 to the left and chromosome five to the right. The centromeres are shown as black boxes. Mapping markers are given to the left of the chromosome. The region of a MRC is indicated with a black bracket, and the locus of a specific RPP gene is shown with a black arrow head to the right of the chromosome (Slusarenko and Schlaich 2003) chromosomes 1, 3, 4, and 5, respectively (Holub 1997). Candidate ‘R’ genes in Arabidopsis were named RPP genes for recognition of Hyaloperonospora parasitica (Crute et al. 1993). Some 19 RPP genes are grouped in the three MRCs, and four others are scattered at separate loci on chromosomes 1 and 2 (Fig. 12.5; Table 12.8). RPP3, which was identified in Oy-0, has not yet been mapped. Of the several RPP loci postulated on the basis of interaction patterns with isolates of H. parasitica, only a minority have been cloned (Fig. 12.5; Table 12.8). In Zurich, Slusarenko and Schlaich (2003) mapped RPP11, which conditions resistance to WELA in RLD, to chromosome 3 at a position 0.4 cm below the marker m249 (Joos et al. 1996). Meanwhile, Jonathan Jones and Jane Parker at Norwich, Jim Beynon and Eric Holub at East Malling, and Jeff Dangl across the Atlantic at Chapple Hill were chasing RPP5/RPP1, RPP13, and RPP8, respectively. Despite all the advantages for molecular genetics which Arabidopsis offers, the honour of being the first plant from which an ‘R’ gene (sensu Flor) was cloned went to tomato, with the isolation of PTO against Pseudomonas syringae pv. tomato (Martin et al. 1993). The first RPP gene whose cloning was reported was RPP5 (Parker et al. 1997), followed 254 12 Host Resistance

Table 12.8 Recognition specificities of RPP genes (for each RPP gene the MRC or chromosomal location, the formerly assigned number (where applicable), the ecotype, and the isolate(s) recognized are given (Slusarenko and Schlaich 2003) Major recognition Arabidopsis Hyaloperonospora parasitica complexes RPP gene ecotype isolates(s) MRC-A No RPP genes Col-0 MRC-B RPP6 Col-0 WELA RPP7 Nd-0 HIKS RPP25 Ler-0 AHCO RPP27 Ws-0 HIKS MRC-F RPP1-WsA formerly Ws-0 NOCO, EMOY, MAKS, CALA RPP-10 RPP-WsB formerly Ws-0 NOCO, EMOY, MAKS RPP-14 RPP-WsC formerly Nd-1 NOCO RPP-1 RPP1-Nd Nd-1 EMOY, HIKS P-1-Nd formerly Nd-1 WACO RPP26 RPP13-Nd formerly Nd-1 ASWA RPP16 RPP13-Nd formerly Nd-1 EMCO RPP17 RPP13-Nd Nd-1 GOCO,EDCO,MAKS RPP13-RLD RLD WELA formerly RPP11 MRC-H RPP2 Col-0 CALA RPP4 Col-0 EMOY, EMWA RPP5 Ler-0 NOCO RPP12 Ws-0 WELA RPP18 Col-0 HIND MRC-J RPP8 Ler-0 EMCO RPP8 formerly Ler-0 GOWA RPP23 RPP21 Ler-0 MADI, MAKS RPP22 Ler-0 ASWA RPP24 Ler-0 EDCO Not in MRCs RPP3 (not mapped) Oy-0 CALA RPP9 Wei-0 HIKS RPP19 Col-0 HIND4 RPP20 Col-0 WAND RPP28 Col-0 HIND2 12.5 Genetics of Host-Pathogen Relationship 255 closely by RPP1 and RPP8 (Botella et al. 1998; McDowell et al. 1998). The Arabidopsis RPP5 gene specifying resistance to the downy mildew pathogen Hyaloperonospora parasitica was positionally cloned by Parker et al. (1997). It encodes a protein that possesses a putative nucleotide-binding site and leucine-rich repeats, and its product exhibits striking structural similarity to the plant resistance gene products N and L6. Like N and L6, the RPP5 N-terminal domain resembles the cytoplasmic domains of the Drosophila Toll and mammalian interleukin-1 transmembrane receptors. In contrast to N and L6, which produce predicted truncated products by alternative splicing, RPP5 appears to express only a single transcript corresponding to the full-length protein. However, a truncated form structurally similar to those of N and L6 is encoded by one or more other members of the RPP5 gene family that are tightly clustered on chromosome 4. The organization of repeated units within the leucine-rich repeats encoded by the wild-type RPP5 gene and an RPP5 mutant allele provides molecular evidence for the heightened capacity of this domain to evolve nove1 configurations and potentially new disease resistance specificities. Interestingly, while RPP8 conditions resistance against EMCO in Ler-0, alternative alleles condition resistance to turnip crinkle virus (allele HRT) and cucumber mosaic virus (allele RCY1) (Cooley et al. 2000; Takahashi et al. 2002). The RPP8 locus is a good example of how recombination slippage and domain shuffling lead to new recognition specificities (Cooley et al. 2000; McDowell et al. 1998). The mapping of RPP13 was published in 1999 (Bittner-Eddy et al. 1999) and was shown to map at the same locus as RPP11 on chromosome 3 (Joos et al. 1996). Following cloning and characterization, it turned out that RPP13 and RPP11 were indeed allelic and recognized different pathogen avirulence determinants (Bittner-Eddy et al. 2000). Thus, RPP13, which was originally cloned from the ecotype Niederzenz, was designated RPP13-Nd, while RPP11, which Slusarenko and Schlaich (2003) had been chasing in the ecotype RLD, was renamed RPP13- RLD (Bittner-Eddy et al. 2000). Cloning of the RPP1 locus from the Ws-0 ecotype revealed that three functional, genetically linked recognition specificities which had been previously designated RPP1, RPP10, and RPP14 were present together. These were redesignated RPP1-WsC, RPP1-WsA, and RPP1-WsB, respectively (Botella et al. 1998). Until recently there has been little progress in characterizing the products of, or cloning avirulence genes from, H. parasitica. DNA fingerprinting of the pathogen has been attempted, and John Lucas has established a mating system useful for gene mapping with H. parasitica, but this is largely with genetic markers in heterothallic isolates, and the situation is com- pounded by the homothallism observed for the isolates pathogenic on Arabidopsis (Moss et al. 1994; Tham et al. 1994). Nevertheless, the outcrossing of two homothallic H. parasitica isolates revealed subsequent progeny segregation of avirulence loci matching six resistance loci in A. thaliana (Gunn et al. 2002). Additionally, the potential for map-based cloning of genes from H. parasitica was demonstrated in the report by Rehmany et al. (2003), which described the isolation of a BAC contig of four clones spanning less than 250kb of DNA over the region containing the ATR1Nd avirulent gene (Slusarenko and Schlaich 2003). 256 12 Host Resistance

12.5.11 Mapping of Downy Mildew Resistance Genes

During the last few years, there were some advances in the genetic study of the inheritance of downy mildew resistance and in the isolation and cloning of resistance genes in Brassica species. One locus conferring downy mildew resistance at the cotyledon stage in broccoli (Brassica oleracea convar. italica) was genetically mapped by Giovannelli et al. (2002) and located in close linkage to the glucosinolate pathway gene BoGsl-elong on a dense map of B. oleracea (Gao et al. 2007). A second downy mildew resistance gene at seedling stage was recently mapped in Chinese cabbage (Brassica rapa ssp. pekinensis) (Carlier et al. 2011). A dominant and monogenically inherited resistance locus expressed at the adult plant stage was identified in broccoli by Coelho et al. (1998) and named Pp523 (after a pathogen strain). This locus was later located on a new genetic map of RAPD and AFLP markers (Farinhó et al. 2004) within a linkage group assigned to the B. oleracea chromosome C8 (Carlier et al. 2011). Five DNA markers that defined a genomic region of 8.5 cm encompassing this resistance locus were then cloned, sequenced, and remapped as SCAR and CAPS markers. BLAST queries (www.ncbi.nihl.gov/ blast) identified a genomic region syntenic to thisB . oleracea genome segment at the extremity of the top arm of Arabidopsis thaliana L. chromosome 1 (Farinhó et al. 2007). Map-based, or positional, cloning is a common strategy for isolation of genes responsible for phenotypic differences. This strategy was used for the isolation of most of the >100 reference ‘R’ genes so far included in the Plant Resistance Genes database (http://prgdb.cbm.fvg.it; Sanseverino et al. 2010). Map-based cloning, with specific variations, was also the central procedure used for the isolation of the A. thaliana genes RPP5 (Parker et al. 1997), RPP8 (McDowell et al. 1998), RPP1 (Botella et al. 1998), RPP4 (Van der Biezen et al. 2002), and RPP2A/RPP2B (Sinapidou et al. 2004), the single downy mildew resistance genes so far isolated in the Brassicaceae family. One of the major steps in map-based cloning is the physical identification of the genomic region where the gene is located. For genomes, still not fully sequenced, this implies the physical mapping of the gene of interest via construction of a contig of large insert DNA clones, usually BACs. Carlier et al. (2011) analysed the construction of a physical map of a genomic region of 2.9 cm that encompasses the downy mildew resistance locus Pp523 in B. oleracea, carried out by exploiting the conserved synteny between B. oleracea and A. thaliana (Farinhó et al. 2007). One major obstacle to overcome was the triplicated nature of B. oleracea genome (O’Neill and Bancroft 2000; Lysak et al. 2005; Town et al. 2006). The selection of BAC clones for construction of the physical map was carried out by screening gridded BAC libraries with DNA overgo probes derived from both genetically mapped DNA markers flanking the locus of interest and BAC-end sequences that align to Arabidopsis thaliana sequences within the previously identified syntenic region. The selected BAC clones consistently mapped to three different genomic regions of B. oleracea. Although 83 BAC clones were accurately mapped within a 4.6 cm region surrounding the downy mildew resistance locus 12.5 Genetics of Host-Pathogen Relationship 257

Pp523, a subset of 33 BAC clones mapped to another region on chromosome C8 that was 60 cm away from the resistance gene, and a subset of 63 BAC clones mapped to chromosome C5. These results reflect the triplication of the Brassica genomes since their divergence from a common ancestor is shared with A. thaliana, and they are consonant with recent analyses of the C genome of Brassica napus. The assembly of a minimal tiling path constituted by 13 (BoT01) BAC clones that span the Pp523 locus sets the stage for map-based cloning of this resistance gene (Carlier et al. 2011).

12.5.12 Resistance Gene-Mediated Signal Transduction

Signal transduction pathways used by the different RPP genes to mediate resistance are still poorly understood and are complex and diverse. At one extreme, there is RPP13-Nd, for which so far no downstream signalling components are known. It functions, independently of EDS1, NDR1, SA, or NPR1, which in various combinations are required for the function of several other ‘R’ genes (Bittner-Eddy and Beynon 2001). At another extreme is RPP4, which seems to depend upon almost all defence signalling components so far known. Thus, RPP4-conditioned resistance against EMOY or EMWA is compromised in Col-0 plants carrying a mutation in EDS1, PAD4; NDR1, PBS3; SGT1b, RAR1; and SID1, SID2 or DTH9, respectively (Table 12.9). Additionally, it is dependent on NPR1 and the accumulation of SA and even requires a functional RPS5 gene, which is the ‘R’ gene required for the recognition of Pseudomonas syringae pv. tomato isolates carrying the avrPphB gene. Essentially, all RPP genes seem to have individual downstream requirements, being qualitatively or quantitatively dependent on different combinations of, for example, SGT1b, RAR1, EDS1, PAD4, or NDR1 (Table 12.9). In mutants, a complete loss of resistance is often not observed but rather a reduction of resistance which can be additive if two mutations are combined. Thus, resistance in the ecotype Ler against the EMCO isolate (mediated by RPP8) was initially reported to be independent of EDS1 and NDR1 (Aarts et al. 1998) but was later found to be significantly compromised in an eds1/ndr1 double mutant (McDowell et al. 2000). Rather than being a straight forward, a linear path from recognition to resistance expression, the picture which is emerging is of a complicated ‘lattice’ or ‘grid-like’ network of interconnecting signalling circuits (Parker 2000). What is known in terms of a rather ‘linear’ summary of signalling components required for the establishment of full resistance as mediated by the different ‘R’ genes is given in Table 12.9. Thus so far, signalling through RPP gene appears to be independent of COI1, JAR1, or EIN2, components required for jasmonate and ethylene signalling, respectively. These genes are required for resistance to necrotrophic pathogens such as Alternaria or Botrytis (Thomma et al. 1998). A simplified schematic representation of RPP- conditioned signal transduction (Fig. 12.6) has been given by Schlaich and Slusarenko (2009). Table 12.9 Contribution of the various signalling components to the resistance reaction mediated by the various RPP resistance proteins (to be read from top to bottom for each RPP column) (Slusarenko and Schlaich 2003) RPP13 RPP gene RPP1- WsA RPP-WsB RPP2 RPP4 RPP5 RPP6 RPP7 RPP8 RPP12 RLD RPP13 Nd RPP19 RPP20 RPP21 Protein TIR-NB-LRR11 TIR-NB- TIR-NB-LRR22,24 TIR-NB-LRR16 TIR- CC-NB-LRR1,24 CC-NB- CC-NB- CC-NB-LRR12 structure LRR11 NB- LRR10 LRR12 LRR8 Ecotype Ws-0 Ws-0 Col-0 Col-0 Ler Col-0 Col-0 Ler Ws-0 RLD Nd Col-0 Col-0 Ler H. CALA, EMOY, CALA EMOY, EMWA NOCO WELA HIKS EMCO WELA WELA ASWA, HIND4 WAND MADI, parasitica EMOY,HIKS, MAKS, EDCO,EMCO,GOCO, MAKS isolate MAKS, NOCO NOCO MAKS RPS5 +9 ++9,14 +9 NR9 NR14 +9 SGT1b NR22 +++1,22 +++1,22 ++22 ++1 +++1 NR22 +++1 ++1 −/+22 RAR1 NR22 NR5,22 ++4,5,14,16,22 ++22 +5 NR4,5, NR22 NR14 +5 +++5 −/+22 RAR1+NDR14 EDS1 +++3,6,20,21 +++3,6,20,21 +++3 +++3,16 +++3,21 NR2,21 −/+2/ +++6 ++3 NR14 +++3,21 EDS1+NDR12,7 NR3,21 NDR1 −/+3,5,7 −/+3,20/++5,7,9,14,16 NR1 −/+5,9 −/+2,5,7,9 NR3 NR14 −/+5,9 −/+5 Oxidative burst 1,20,23 PBS3 NR5 ++5,16 NR5 NR5 +5 PAD1 NR13 NR13 NR13 NR13 PAD2 NR13 NR13 NR13 +13 PAD3 NR13 NR13 NR13 +13 PAD4 +20,21 +20,21 +++13 +++13,14,16 NR13,21 NR21 NR14 +++13 +21 PAD1+PAD213 PAD2+PAD313 PAD2+PAD313 PAD1+PAD313 SA -/+2 +++1,2,14,15 +21 ++2,15 NR2 ++2 +++15 NR14 -/+2 ++2 NPR1 ++16,18 +16,18 -/+2 +++1,2,14,16 -/+2 NR2 -/+2 +16,18 NR14 ++2 -/+2 PHX6+LSD519 DTH916,17 PHX11+LSD519 SID116 SID216 Quantitative contributions are as follows: NR, not required; –/+, contributes very little to resistance; + or ++, increasing requirement to resistance; +++, required for resistance. The relative position of the signalling components does not necessarily reflect the correct sequential action of the respective gene product along the signalling pathway. Occurrence of the oxidative burst has been shown only for a few RPPs; however, it is assumed that it is a generally important factor in disease resistance signalling, and hence it is shown as if it would occur downstream of all RPPs. 260 12 Host Resistance

Fig. 12.6 Schematic representation of RPP-conditioned signal transduction. H. arabidopsidis isolates are shown above the corresponding RPPs that recognize them below the isolates (hyphenated with the Arabidopsis accession from which they were cloned). RPP1 from Ws (RPP1-Ws) is dependent on PAD4 and EDS1. RPP 2A+B from Col-0 is quantitatively influenced by the resistance protein RPS5 that recognizes the bacterial pathogen Pseudomonas and requires SGT1b, PAD4, and EDS1. RPP4 from Col-0 is particularly sensitive to changes in resistance signalling. Thus, mutations in many defence-related genes lower the defence responses to EMOY and EMWA. RPP5 from Ler requires RAR1 and SGT1b as well as PAD4 and EDS1. Furthermore, RPP5 was shown to be dependent on LURP1. RPP8 from L-er is an exception, because it encodes a CC-NB-LRR-type resistance protein, yet is dependent not only on NDR1 and RIN4 (like other CC-NB-LRRs) but also on PAD4 and EDS1. No downstream signalling components of RPP13-Nd have so far been reported. Please note: Signalling by the TIR-NB-LRR-type RPP genes RPP1-Ws, RPP2A+B-Col-0, RPP4-Col-0, and RPP5-Ler was dependent on PAD4 and EDS1, which have a positive interlocked feedback regulation with salicylic acid (SA) [Slusarenko and Schlaich 2003; Yoshioka et al. (2006); Zhang et al. (2005); Zhang and Li (2005); Knoth and Eulgem (2008); Knoth et al. (2007)] (Schlaich and Slusarenko 2009)

12.6 Biochemical Basis of Resistance

The presence and absence of natural biochemical compounds like glucosinolates and other phenolic compounds play a significant role in providing resistance to the host plant. There is a correlation between high levels of flavour volatiles (e.g. allylisothiocyanate) released by tissue damage and the limitation of fungal growth in both wild and cultivated Brassica lines. In cultivated Brassicas, breeding has resulted in reduced levels of flavour volatiles with a consequent reduction in their general resistance to H. parasitica. The resistance to H. parasitica in the cabbage cultivar ‘January King’ may be attributed to the high concentration of 12.6 Biochemical Basis of Resistance 261 allylisothiocyanate (Greenhalgh and Mitchell 1976). In France, cauliflower cvs. C300 and Maudez resistant to downy mildew contained the highest sinigrin content which can act as biochemical marker for identification of downy mildew resistance (Menard et al. 1999). The incidence and severity of downy mildew are positively correlated with glucosinolate concentration in seeds of oilseed rape (Rawlinson et al. 1989). In oilseed rape, downy mildew severity is lower on cultivars with high concentration of glucosinolate (>100/μmol g−1) and greater on those with lower concentrations (<15 μmol g−1). In all cultivars grown in the UK, incidence is lower in mid-February when glucosinolate products in the leaves reach a maximum level (Anonymous 1985). A large number of B. napus species with different glucosinolate and erucic acid contents have been screened for resistance to four isolates of H. parasitica at the cotyledon stage (Nashaat and Rawlinson 1994). Two groups of accessions with different resistance factors were identified. The first group was different from that of the cultivar ‘Cresar’ which has an isolate-specific gene for resistance to H. parasitica, and the second group was identical to that of ‘Cresar’. There was moderate to full susceptibility at the cotyledon stage, but no clear differential response to any of the isolates. Those with high glucosinolate and high erucic acid content were significantly less susceptible than those with high glucosinolate and low erucic acid or low glucosinolate and low erucic acid content. In preliminary experiments on the effect of treatment with abiotic elicitors on disease reaction in oilseed rape seedlings, salicylic acid and methyl jasmonate reduced the severity of infection when cotyledons were subsequently inoculated with H. parasitica (Doughty et al. 1995).

12.6.1 Role of Phytoalexins in Resistance to Downy Mildew

Glazebrook et al. (1997) studied a phytoalexin-deficient (pad) mutant in Arabidopsis thaliana Columbia (Col-0) to investigate the role of phytoalexin camalexin (3-thiazol-2yl-indole) in disease resistance. Mutations in PAD1, PAD2, and PAD4 caused enhanced susceptibility to Pseudomonas syringae pv. maculicola strain ES4326 (Psm ES4326), while mutations in PAD3 or PAD 5 did not. Camalexin was not detected in the double mutants pad 1-1, pad 2-1; pad 1-1, pad 3-1; or pad 2-1, pad 3-1. Growth of Psm ES4326 in pad 1-1, pad 2-1 was greater than that in pad 1-1 or pad 2-1 plants, while bacterial growth in pad 1-1, pad 3-1 and pad 2-1, pad 3-1 plants was similar to that in pad1-1 and pad 2-1 plants, respectively. The pad 4-1 mutation caused reduced camalexin synthesis in response to Psm4326 infection but not in response to infection by Cochliobolus carbonum, indicating that PAD 4 has a regulatory function. PAD 1, PAD 2, PAD 3, and PAD 4 are all required for resistance to downy mildew (H. parasitica). The pad 4-1 mutation caused the most dra- matic change, exhibiting full susceptibility to four out of six Col-incompatible H. parasitica isolates. Each combination of double mutants between pad1-1, pad2-1, and pad3-1 exhibited additive shifts to moderate or full susceptibility to most of the isolates. 262 12 Host Resistance

12.6.2 Lignification of Host Cells

Lignin formation was observed in cell walls of parenchyma of Japanese radish root infected with downy mildew fungus. The observed lignin was mainly composed of guaincylpropane units and apparently differed from syringyl lignin which was present in vessels of the healthy tissues. Different pathways for lignin biosynthesis in the healthy and the diseased tissues were proposed (Tables 12.10 and 12.11; Figs. 12.7, 12.8, and 12.9) by Asada and Matsumoto (1972). Specific isoperoxidases synthesized de novo in diseased tissues were presumed to play an important role in the formation of the guaincyl lignin (Ohguchi et al. 1974; Ohguchi and Asada 1975). Lignin was also formed in cell walls of parenchyma of Japanese radish root which was infiltrated with 700 × g supernatant of homogenate of downy mildew infected tissues. It began to form about 12 h after infiltration of the homogenate (Matsumoto et al. 1978). The effective component in the homogenate for the induction of lignification is dialyzable, which is highly water soluble, and seems to resemble monilicolin A (Asada et al. 1975). The lignification-inducing factor (LIF) plays a significant role in the induction of systemic resistance. Resistance to H. parasitica was induced in the roots of susceptible radish cultivars when they were preliminarily inoculated with the pathogens or wounded. An increase in L-phenylalanine ammonia-lyase (PAL) activity and lignification of cell walls

Table 12.10 Amounts of degradation products by alkaline nitrobenzene oxidation of the isolated lignin (Asada and Matsumoto 1972) Lignin Product Amount (%) Ratio Healthy p-Hydroxybenzaldehyde (H) 0.39 V/H 6.05 Vanillin (V) 2.36 S/H 3.90 Syringaldehyde (S) 1.52 S/V 0.64 Total 4.27 Diseased p-Hydroxybenzaldehyde (H) 0.99 V/H 2.75 Vanillin (V) 2.72 S/H 0.0 Syringaldehyde (S) 0.00 S/V 0.0 Total 3.71

Table 12.11 Elemental compositions and empirical formulae of the isolated lignins and the related compounds (Asada and Matsumoto 1972)

Lignin C (%) H (%) OCH3 (%) Formula

Healthy root 64.38 7.27 18.93 C9H10.3 O2.20(OCH3)1.16

Diseased root 63.47 5.74 12.58 C9 H8.33 O2.80(OCH3)0.75 a Birch 58.82 6.49 21.51 C9 H9.03 O2.77(OCH3)1.58 a Spruce 63.48 6.35 14.84 C9 H8.83 O2.37(OCH3)0.96 b DHP 64.00 6.00 16.90 C9 H8.21 O2.50(OCH3)1.02

Coniferyl alc. 66.67 6.67 17.22 C9 H9 O2 (OCH3)1 aFrom Bjorkman and Person (1957) bDehydrogenation polymerization product, from Nozu (1967) 12.6 Biochemical Basis of Resistance 263

Fig. 12.7 The UV absorption spectra of the diseased parenchyma cell wall (a), the vessel wall (b), and the healthy parenchyma cell wall (c) of the Japanese radish root (Asada and Matsumoto 1972)

Fig. 12.8 UV absorption spectra of the authentic compounds (a) and the degradation products (b) obtained from the extraction of paper chromatograms. I, P-hydroxybenzaldehyde; II, vanillin; III, syringaldehyde (Asada and Matsumoto 1972) occurred in these tissues after challenge inoculation with downy mildew. Histochemical observations indicated that the cell walls were lignified in the tissues beyond the site of fungal attack (Matsumoto and Asada 1984). High peroxidase activity was located around the lignified cell walls (Asada and Matsumoto 1969; Ohguchi et al. 1974). The higher amount of lignin accumulation was present in the middle portion of the cell wall (Asada and Matsumoto 1971). The most resistant cvs of Japanese radish has lignified cell walls prior to attack by H. parasitica, and the 264 12 Host Resistance

Fig. 12.9 Suggested pathway of lignin biosynthesis in healthy (full lines) and diseased (broken lines) plants. Px, Py, Pz: peroxidase isoenzymes x,y,z (Asada and Matsumoto 1972) hyphae could not penetrate cells (Matsumoto 1994). Following infection of radish by H. parasitica, the deposition of lignin in host cell walls may have a role to play in nonspecific limitations of the growth of biotrophic fungi. Ohguchi and Asada (1975) have defined the pathways and enzymes involved in lignin biosynthesis in radish following infection by H. parasitica. The formulae were obtained from the following equations:

108.%09HO()−×31.%035 CH3 () 108.%9×OCH () X =   Z = 3 1.008D 31.035D

108.%09OO()−×31.%035 CH3 () 12.01 =   DC= ()% − 16D 31.035× OCH % 3 ()

12.7 Sources of Resistance

Differential host resistance to isolates of H. parasitica has been identified in B. campestris, B. juncea, B. napus, B. oleracea, and R. sativus (Bonnet and Blancard 1987; Lucas et al. 1988; Nashaat and Rawlinson 1994; Nashaat et al. 1995a, b, 1997, 2004; Silue et al. 1996). Sources of resistance to Hyaloperonospora parasitica in different host species of crucifers identified from different countries of the world are given in Table 12.12 (Saharan et al. 1997). 12.7 Sources of Resistance 265

Table 12.12 Sources of resistance to Hyaloperonospora parasitica (Saharan et al. 1997 updated) Host species/genotypes References Brassica alba (white mustard) Saharan (1992a, b) All Indian accessions B. carinata (Ethiopian mustard) All Indian accessions Saharan (1992a, b) HC1 Saharan (1996), Dang et al. (2000) B. campestris Saharan (1992a, b) Candle B. campestris var. toria (toria) Kolte and Tewari (1980) 1B-586 B. campestris var. yellow sarson (yellow sarson) Kolte and Tewari (1980) YST-6 B. campestris var. brown sarson (brown sarson) Kolte and Tewari (1980) BS-16 B. juncea (Indian mustard) IC 296685, IC 326253, IC 417020, Bisht et al. (2015) DIR 1507, DIR 1522 Dang et al. (2000) PI340207, PI 340218, PI 347618 Ebrahimi et al. (1976) Domo, RC-781, EC 126743, Zem, YRT3, 45,72 Saharan (1992a, b) PR 8805, RN 248, EC 129126-1, PC-3 Saharan (1996) RESBJ-01, RESBJ-02, RESBJ-03 Nashaat and Awasthi (1995) EC 399296, EC 399299, EC 399301, EC 399313, EC Kolte et al. (2008) 414308 (NRCR-837), EC 414319 (NRCR-836) B. oleracea var. botrytis (cauliflower) C300, Maudez Menard et al. (1999) Igloo, Snowball y, Dok Elgon, RS-355 Kontaxis et al. (1979) PI 181860, PI 188562, PI 189028 (MR), PI 204765, PI Thomas and Jourdain (1990) 204768 PI 204772, PI 204773, PI 204779, PI 241612 Sharma et al. (1991) PI 264656, PI 291567, PI 373906, PI 462225 (MR) Hoser- Krauze et al. (1991) KPS-1 Rooster et al. (1999) PI 231210, PI 189028 Pandey et al. (1995) Aviso, Exale, B1704, Stanley Adam’s white head, IIHR-142, IIHR-217 B. oleracea var. capitata (cabbage) January King Greenhalgh and Mitchell (1976) Balkan Elenkov (1979) Spitz Kool Verma and Thakur (1989) PI 246063, PI 246077, PI 245013 Hoser- Krauze et al. (1991) Tromchuda cabbage ‘Algarvia’ (ISA 207) Caravalho and Monteiro (1996) PI 245015, Geneva 145-1 Sherf and Macnab (1986) B. oleracea var. accephala gr. ornamentalis Vitanova (1996) (Decorative cabbage) (continued) 266 12 Host Resistance

Table 12.12 (continued) Host species/genotypes References B. oleracea (broccoli) Ching-Long 45 Yang et al. (1998) Calabrese, Grand Central Natti et al. (1956) PI 231210, Italian Green Sprouting, Natti (1958) Hyb 1230 (Moran), Green surf (Moran), Laemmlen and Mayberry (1984) 2804 (Qualisal), GSV 82-4310 (Goldsmith), XPH 1117(Asgrow), Hyb. 288 (Moran) Hoser-Krauze et al. (1991) AVX 7631 (Sun Seeds) Baggett and Kean (1985) PI 263056, PI 263057, PI 3573, PI 3574 Sherf and Macnab (1986) PI 418984, PI 418985, PI 418986, PI 418987, PI 418988 OSU CR 2 to OSU CR 8 Citation, Excalibur, Nancy B. napus (rape) Hg Vestal Jonsson (1966) Eurora, Janetzki, Kubla, Lesira Dixon (1975) Mogul, Primar, Rapot, Rapara, Sinus Cultivar 78-22 Chang (1981) Cresor Kluczewski and Lucas (1983) PI 199949, PI 263056 Thomas and Jourdain (1992) Gulivar, Midas, Tower, GS 7027 Saharan (1992a, b), Dang et al. (2000) RES 01-14, RES-02, RES-26 Nashaat and Awasthi (1995), Nashaat et al. (1996, 1997) HNS3, HNS4, GSL-1, GSL 1501 Saharan (1996) EC338986-2, EC338996-1 Kolte et al. (2008) B. chinensis (Chinese cabbage) Bau chin 26, PHW 64707, PHW 64710, PHW 64722, PHW Niu et al. (1983) 64620 Hyb. 77M (3)-27, Hyb.77M(3)-35, Hyb. 82-46, Hyb. Anonymous (1987a, b) 82-46R, Hyb.82-156, Hyb. 82-157 Dwarf Resistant 5, Dwarf Resistant 6 Cao et al. (1998a, b) B. nigra (black mustard) PI 19948 Thomas and Jourdain (1992) B. rapa PI 418984, PI 418988, PI 418987, PI 418988 Thomas and Jourdain (1992) B. rapa subsp. Rapifera Long Blanc de croissy, Stanis, Jaune Boule d’or Silue et al. (1996) Raphanus sativus (radish) Okura Shiraishi et al. (1974) Tokinoshi (all season) Bonnet and Blancard (1987) Bamba, Noir Lon d’orloge, Rave a Forcer Silue et al. (1996) Cheiranthus cheiri (wallflower) Covent Garden Blood Red Greenhalgh and Dickinson (1975) 12.8 Breeding for Disease Resistance 267

12.8 Breeding for Disease Resistance

Plant breeding offers one method for controlling diseases and has obvious advantages if successful. As with other traits, the breeder’s task is firstly to find sources of disease resistance and effective ways to screen for resistant genotypes. The trait has then to be transferred into a useful cultivar or hybrid (Buzza 1995). Transfer of resistance among crucifers and from other species is possible by using conventional and biotechnological techniques: (a) Germplasm evaluation for sources of resistance at national and international levels (b) Selection for disease resistance through (i) pure line selection, (ii) mass selection, (iii) modified recurrent mass selection, and iv) recurrent selection (c) Breeding for disease resistance by increasing the level of resistance through (i) multiple crosses, (ii) recurrent selection, (iii) diallel crossing, and (d) selective mating system (d) Transfer of resistance by (i) intraspecific pedigree, backcross, and modified recurrent mass selection methods and (ii) interspecific genome substitutions, chromosome substitutions, and gene introgression (e) Transfer of resistance through mutation breeding (f) Use of biotechnological and genetic engineering techniques such as (i) genome manipulation, (ii) manipulation of cytoplasmic genomes, (iii) use of transformation and foreign gene expression techniques, and (iv) embryo rescue techniques for wide hybridization (g) Use of genomics and molecular techniques for identification, mapping, cloning, and utilization of ‘R’ genes with other useful traits

12.8.1 Strategies to Breed Downy Mildew Resistance Cultivars of Crucifers

The details of techniques based on current understanding on the molecular mechanisms of crucifer-H. parasitica interaction have been given in Chap. 1 at Sect. 1.11.

12.8.2 Designation and Nomenclature of Resistance Genes

The system and methods adopted to designate the R genes in crucifers have been described in detail in Chap. 1 at Sect. 1.11.6. 268 12 Host Resistance

12.9 Mechanisms and Application of Gene Silencing Techniques to Downy Mildew of Crucifers

Gene silencing is triggered by dsRNA that, under normal cellular conditions, may develop from transposon transcripts, gross overexpression of transcripts (as for viruses), aberrant messenger RNAs, or from non-protein-coding premicro RNA (miRNA) transcripts (Ruiz et al. 1998; Jensen et al. 1999a, b, c; Gazzani et al. 2004; Luo and Chen 2007; Ghildiyal et al. 2008). Alternatively, dsRNA molecules can be introduced into cells either as in vitro synthesized molecules (Timmons and Fire 1998; Clemens et al. 2000) or from antisense or inverted repeat expression vectors (Guo et al. 2003). The basic mechanism of gene silencing follows that dsRNA is bound and cut by the dicer enzyme into shorter dsRNA molecules, typically of 21–24 bp and termed short interfering RNAs (siRNAs). Dicer is a multidomain protein that comprises at least two RNase III domains and additional domains such as dsRNA binding, PAZ, and DEAD-box helicase. The PAZ domain may act in concert with the RNase III domains as a molecular measure to define the number of nucleotides of RNA between the two RNase III cut sites on the dsRNA (Zhang et al. 2004). The presence of a helicase domain may assist in unwinding complex secondary structures, processing of long dsRNAs, or unwinding of siRNAs for the next stage of the silencing pathway. Following dicer digestion, the siRNAs strands are separated, and the antisense strand is incorporated into the RNA-induced silencing complex (RISC), the core of which is the multidomain argonaute protein. Argonaute proteins contain two major domains, PAZ and PIWI. The antisense strand of the siRNA duplex is bound into the RISC via the PAZ domain and acts as the guide strand, which targets the homologous sequence in the native messenger RNA (mRNA) and degrades it through the slicer activity of the PIWI domain (Collins and Cheng 2005). A related class of small RNA species, called miRNA, also develops through dicer or other RNase III enzyme (such as Drosha) digestion of non-protein-coding premiRNA transcripts containing a stem-loop secondary structure. Similar to siR- NAs, the mature miRNA may also associate with argonaute proteins to degrade endogenous mRNAs, thus achieving a level of post-transcriptional regulation. Alternatively, the mature miRNA may bind to the homologous messenger RNA (mRNA) to inhibit translation (Brodersen et al. 2008). SiRNAs and fragments released from RISC degradation of mRNAs can act to prime synthesis of more dsRNA from the native mRNA through the action of the RdRP enzyme. RdRP can act both to spread the silencing target signal along the mRNA and to amplify the signal through production of more dsRNA (Alder et al. 2003). The synthesis of dsRNA by RdRP has been shown to occur in both a primer- dependent and primer-independent manner (Makeyev and Bamford 2002). Secondary siRNAs in Caenorhabditis elegans have been shown to exhibit sequence direction bias suggestive of unidirectional priming (Sijen et al. 2001), whereas aberrant mRNAs can be substrates for primer-independent RdRP synthesis of dsRNA (Baulcombe 2004; Luo and Chen 2007). 12.9 Mechanisms and Application of Gene Silencing Techniques to Downy Mildew… 269

The major cytoplasmic steps of gene silencing are frequently described as post- transcriptional gene silencing (PTGS) or RNAi. That is, the native gene is transcribed, but the mRNA is degraded before translation can occur. The components of PTGS also provide a link to silencing at the transcriptional level (Verdel et al. 2004; Buhler et al. 2007). In many organisms where PTGS operates, it can also lead to epigenetic changes at the DNA level, principally through de novo methylation of cytosines or through deacetylation and methylation of histone proteins. The outcome of these modifications is silencing at the transcriptional level. Plant proteins involved in these processes are RNA polymerase IV, DNA methyltransferase, histone methyltransferase, histone deacetylase, and chromodomain protein DRD1 (Huettel et al. 2007); RdRP, dicer, and argonaute are also involved in initiating and maintaining TGS. Similar processes also operate in fungi and are best characterized in fission yeast Schizosaccharomyces pombe. In S. pombe, silencing operates at the transcriptional level through formation of heterochromatin. Similar to plants, this involves dicer, RdRP, and argonaute proteins. S. pombe argonaute forms part of the RNA-induced transcriptional silencing (RITS) complex, together with proteins CHP1 and TAS3 (Verdel and Moazed 2005). TAS3 has no discernable domains, but CHP1 is a chromodomain protein. The RITS complex, through CHP1, targets DNA sequences homologous to the siRNA bound in the RITS complex to modulate the deacetylation and methylation of histones and thus the formation of heterochromatin (Verdel and Moazed 2005). The methylation/acetylation status of histones is thought to mediate access to the DNA for RNA polymerase II, with any mRNA formed, and targeted by RDRP to form dsRNA and then siRNAs; this process main- tains the silenced state (Grewal and Jia 2007). Similar to S. pombe, TGS that develops from RNAi in the nematode C. elegans can persist through several generations, with its basis being the activity of histone deacetylase, histone acetyltransferase, a chromatin remodelling ATPase, and a chromodomain protein (Vastenhouw et al. 2006). PTGS in C. elegans and other organisms such as Neurospora crassa has been observed as a transient phenomenon that decreases in intensity with time after exposure to the initial silencing stimulus. A component of this ‘release’ from PTGS has been identified fromC. elegans as a dsRNA-specific RNase termed ERI1, which degrades siRNAs (Kennedy et al. 2004). ERI1 mutants exhibit enhanced and more persistent RNAi and TGS (Kennedy et al. 2004; Iida et al. 2006). The recent sequencing of the genome of Arabidopsis downy mildew pathogen and Hyaloperonospora arabidopsidis (Hyaloperonospora parasitica) (Tyler et al. 2006; http://www.broad. mit.edu/annotation/genome/Phytophthora infestans; http://phytophthora.vbi.vt.edu/) has and will continue to assist the discovery of numerous genes and pathways associated with downy mildew biology. Identifying the components of downy mildew gene silencing has also been assisted by the availability of genome sequences. Using the P. infestans genome sequence as an example, genes encoding the key components of PTGS are all present: dicer-like, argonaute, and RdRP (Ah-Fong et al. 2008). Genes encoding other potential PTGS accessory proteins that contain domain characteristics of proteins that are involved in PTGS in other organisms, such as dsRNA-binding proteins and DEAD-box RNA helicases, are also present (Walker et al. 2008). Using both plants, S. pombe, and C. 270 12 Host Resistance elegans as reference systems where TGS has been dissected, genes encoding the major components, such as chromodomain proteins and histone-modifying proteins, are all present within oomycete genome sequences in multiple copies as in other organisms. It remains to be demonstrated which of these protein family members contribute to TGS in oomycetes. In support of the observations from laboratory- based studies, genes encoding proteins for de novo methylation of DNA were not identified (Judelson and Tani 2007). Similarly, subunits of RNA polymerase IV, a plant-specific protein apparently involved in de novo DNA methylation, were not identified in the P. infestans genome. Model gene silencing systems C. elegans, S. pombe, and Arabidopsis thaliana all contain a protein called ERI1, which is an exoribonuclease that digests siRNAs, ultimately leading to a release of PTGS (Kennedy et al. 2004; Iida et al. 2006). The P. infestans genome does not seem to contain the gene encoding this protein. The lack of an ERI homologue may in part explain the persistence of gene silencing in the absence of any homologous transgene (Van West et al. 1999, 2008; Whisson et al. 2005; Gaulin et al. 2007) and the observation that partial silencing in some P. infestans transformants can convert to complete silencing after some time (Ah-Fong et al. 2008). The strategies for application of gene silencing in downy mildew of crucifers can be (1) stable gene silencing and (2) transient gene silencing procedures.

12.9.1 Stable Versus Transient Gene Silencing

Each of the two strategies for achieving gene silencing in downy mildew of crucifers has inherent advantages and disadvantages. Stable, transcriptional silencing has the obvious advantages in maintaining undetectable levels of gene expression, such that effectively null genotypes and their associated phenotypes can be studied. This allows very subtle differences in phenotypes to be identified and studied in detail, such as the 50% reduction in cyst germination observed for silencing of NIF genes (Judelson and Tani 2007). However, transcriptional gene silencing is effectively a gene knockout in terms of gene expression, which signifies that genes for which silencing leads to a lethal phenotype are difficult to study by this approach. Transient gene silencing, in which a range of gene expression knockdown levels are observed, may be better suited to the study of these ‘core biology’ genes. The obvious disad- vantage of transient silencing is that the genotype and phenotype apparently do not persist for more than 20 days, and thus experiments require repetition on each occa- sion that the gene under study is to be characterized phenotypically. However, the recent reporting of stable but partial gene silencing may alleviate this difficulty (Ah-Fong et al. 2008). Stable gene silencing, whether partial or complete, relies on transformation of downy mildew. Although transformation is routine for model downy mildew species, this is not the situation for many other downy mildews. References 271

Table 12.13 Responses of phenylamide sensitive and insensitive isolates of H. parasitica to phenylamide fungicides (Moss et al. 1988) Fungicides μg/ml Compound Isolates 0.05 0.05 5.0 50.00 Factor of insensitivity Metalaxyl P005 32a 0 0 0 0 P006 94 89 100 83 X1000 Cyprofuran P005 84 79 0 0 P006 100 94 78 3 X10 Figures are reciprocals of mean latent periods (time form inoculation to sporulation expressed as a percentage of the untreated control)

12.10 Development of Resistance to Fungicides

The existence of metalaxyl resistant strains of H. parasitica has been reported (Crute 1984; Crute and Gordon 1986; Brophy and Laing 1992). Metalaxyl-resistant isolates were also shown to be cross-resistant to furalaxyl and ofurace, two related phenylamide fungicides. A differential degree of insensitivity to two related phenylamide fungicides has been demonstrated (Table 12.13) (Moss et al. 1988). Metalaxyl was more active against a sensitive isolate (P005) than cyprofuran, but the converse was true with an insensitive isolate (P006). The inheritance of fungicide insensitivity to H. parasitica may reveal the true picture of genes controlling this phenomenon.

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13.1 Introduction

To manage downy mildew of crucifers, no single method or approach is considered feasible, effective, environmentally safe, and economical. It is always essential to integrate the available methods for disease control/management.

13.2 Cultural Practices

Cultural control of crucifers’ downy mildew disease is largely a matter of sanitation, and of manipulating the environment to the advantage of the host, and to the detri- ment of the pathogen. Since the pathogen survives in the form of oospores in the host tissues, removal, destruction, and burning of the infected plant debris along with weeds has been suggested to restrict the source of primary inoculum (Butler 1918; Vasudeva 1958). In addition, clean, well-drained soils with 2 years of crop rotation using non-cruciferous crops were also recommended. Measures to reduce the relative humidity around the plants by adequate aeration, and avoidance of dense sowing, and controlling the growth of weeds also helped to reduce the disease (Butler 1918; Conroy 1960; Schmidt 1960; Sherf and Macnab 1986). Avoidance of continuous cropping of rape on the same field or adjacent to a field sown to rape in the previous year was also advised to reduce infection by H. parasitica (Downey and Bolton 1961). The widespread cultivation of one or only a few cultivars of the same species may favour the disease. In India, the late sown crops of rapeseed- mustard were reported to have a higher incidence of downy mildew than the early (before October) or timely (by middle of October) sown crops (Kolte 1985; Saharan 1984, 1992a) (Table 13.1). Early sowing (first October) of B. juncea resulted in disease escape from downy mildew under Jammu and Kashmir, Indian conditions (Kaul and Singh 1999).

© Springer Nature Singapore Pte Ltd. 2017 285 G. S. Saharan et al., Downy Mildew Disease of Crucifers: Biology, Ecology and Disease Management, https://doi.org/10.1007/978-981-10-7500-1_13 286 13 Disease Management

Table 13.1 Effect of planting dates on the severity of white rust and downy mildew of Indian mustard cv. Varuna (Saharan 1992b) Percent WR and DM disease severity Sowing date Hisar Kanpur Pantnagar 6 October 1978 10.0 – – 21 October 1978 8.6 – – 28 October 1978 18.6 – – 6 November 1978 55.4 – – 18 November 1978 68.5 – – 2 December 1978 72.8 – – 1 October 1979 – 24.16 (29.45) – 10 October 1979 04.6 28.30 (32.14) – 20 October 1979 10.0 34.34 (35.86) – 30 October 1979 22.5 36.18 (36.97) – 9 November 1979 46.8 40.91 (39.77) – 19 November 1979 57.5 46.15 (42.76) – 3 October 1980 – – 15.04 23 October 1980 – – 19.85 13 November 1980 – – 32.85 – = no stag head recorded

In the Lujskaya area, Leningrad regions of the USSR, the level of downy mildew infection was reduced on cabbage plants transplanted between 26 and 30 June. Fertilizer containing 50% humus, 45% peat, and 5% mullein with 3.9 g ammonium nitrate, 4.3 g Kcl, and 8.1 g superphosphate/100 g added to 110 g organic matter applied to the soil reduced the percentage of diseased plants better than the organic manure alone (Kupryanova 1957). Removal of 50% lower leaves at 60 days of crop age reduced disease severity (Kaul and Singh 1999). In Poland cultivation of winter rape after red clover increased infection of H. parasitica (Czajka 1994). Esiyok et al. (1996) found that covering cauliflower plants with polyethylene sheets gave better results than the classical seed production method in the open field.

13.3 Seed Treatment

Fungicidal seed treatment followed by a foliar spray is a common practice to control downy mildew of crucifers. Metalaxyl seed treatment at the rate of 0.3–0.6 g a.i. Kg−1 reduced downy mildew infection on broccoli (Paulus and Nelson 1977) and rapeseed-mustard (Kolte 1985; Saharan 1992a). Jensen et al. (1998) found significant effect of seed treatment with CGA 245704 on the sporulation of H. parasitica, providing a novel strategy for management of seedling infection. Seed treatment with metalaxyl (as Apron 35 DS) at 2 g/Kg seed gave 96.5% control of downy mildew in cauliflower (Verma et al. 1994). A significant yield increase was observed when plants raised from such treated seed were sprayed once or twice with the same 13.4 Soil Treatment 287

Table 13.2 Efficacy, economics, and spray schedule of fungicides against downy mildew of mustard (Mehta et al. 1996) Percent Percent Percent Cost- Conc. Spray disease disease increase in benefit Fungicides (%) no. intensity control yield ratio Dithane M-45 0.2 4 28.4 42.4 28.5 1:2.20 Kavach 0.2 4 28.2 35.0 23.3 1:1.57 Ridomil MZ-72 0.25 3 9.1 81.3 49.3 1:1.21 *Apron 0.25 2 15.8 68.5 34.2 1:1.11 SD-35 + Ridomil MZ-72 Apron SD-35+ 0.2 3 22.9 47.4 22.1 1:2.11 Dithane M-45 Apron SD-35+ 0.2 3 21.9 49.6 20.4 1:1.29 Kavach Apron SD-35 – – 33.6 41.0 15.9 1:20.62 Control – – 49.7 – LSD 0.05 – – 4.1 * Apron SD-35 as seed treatment at 2 g a.i. kg seed compound. Seed treatment with Apron SD 70 (35% metalaxyl and 35% Captan) controlled downy mildew of cauliflower for more than 2 weeks after sowing (Crute 1984). According to White et al. (1984), seed treatment with Apron SD 70 (1 g metalaxyl kg−1) gave complete control of downy mildew on cauliflower inoculated 10 days after sowing. Following seed treatment, metalaxyl was detectable in the cotyledons, true leaves, and roots of cabbage seedlings up to 4 weeks after sowing. An effective and economical schedule for control of downy mildew of mustard through fungicidal seed treatment and/or spray application has been worked out under Indian conditions. Seed treatment with Apron SD-35 (2 g metalaxyl a.i. kg- 1seed) along with two foliar applications of Ridomil MZ-72 at 30 days intervals gave the best control of downy mildew on mustard along with an increase in yield (Mehta et al. 1996; Table 13.2). The maximum cost-benefit ratio was obtained when mustard seeds were treated with Apron SD-35 followed by three spays with mancozeb.

13.4 Soil Treatment

The use of systemic fungicides such as prothiocarb (Dynone) at 5 g m−2 before sowing and fosetyl-aluminium (Aliette) at 10 g m−2 as soil drench gave excellent disease control on cauliflower (Ryan 1977) (Table 13.3). Both these fungicides were as effective as eight sprays of dichlofluanid (Ryan 1977). Prothiocarb also reduced infection of radish leaves and bulbs when applied at 0.1% as a drench (4 litres m−2) at 50% seedling emergence and was much more effective than sprays of dichlofluanid, zineb, captafol, and maneb (Anonymous 1974). 288 13 Disease Management

Table 13.3 Efficacy of fungicidal treatments on the severity of downy mildew of cauliflower (Ryan 1977) Disease severity index Method of Rate of Walk in tunnels Low tunnels Fungicides application application March 28 April 13 April 13 April 25 Dichlofluanid Foliar spray 8 at 1 g/10 m2 1.4 2.8 1.6 2.3 Aliette Foliar spray 3 at 1 g/10 m2 1.7 1.9 1.7 1.9 Prothiocarp Foliar spray 3 at 1 g/10 m2 1.6 2.1 1.5 1.7 Aliette Soil treatment 5 g/m2 1.2 2.6 1.0 2.0 Aliette Soil treatment 10 g/m2 0.7 1.8 0.3 1.2 Aliette Soil treatment 20 g/m2 0.0 0.8 0.0 0.6 Prothiocarp Soil treatment 5 g/m2 0.6 1.4 0.2 1.0 Prothiocarp Soil treatment 10 g/m2 0.2 1.1 0.0 0.9 Prothiocarp Soil treatment 20 g/m2 0.0 0.5 0.0 0.7 Control – – 2.4 3.7 2.0 3.0 LSD 5% 0.72 0.76 0.78 0.56 0 = no disease; 5 = very severe disease

Granular applications of metalaxyl prior to sowing were shown to be an effective control of downy mildew on broccoli (0.56 and 1.2 kg a.i. ha−1) and on cauliflower (0.28 kg a.i. ha−1). In cauliflower, pre-sowing incorporation or a single post-sowing drench (1.5 kg a.i. ha−1) or three high-volume sprays (0.8 g a.i. litre−1) of metalaxyl gave much better disease control than nine sprays of dichlofluanid applied during a 6–8 week period (Chiu 1959).

13.5 Compost Treatment

In the UK, metalaxyl, milfuran + manganese zinc dithiocarbamate, or propamocarb incorporated in the compost provides good control of downy mildew on moduleraised cauliflowers in early summer plantings. In summer cauliflowers, good control was achieved by drenching the compost with propamocarb and fosetyl-aluminium foliar sprays and by applying a dichlofluanid foliar spray programme (Davies and Wafford 1987).

13.6 Foliar Spray of Fungicides

During the period from the mid-1940s to the mid-1960s, control of downy mildew of crucifers rested on frequent applications of sprays or dusts of fungicides such as chloranil (Spergon), copper-based materials, and zineb (Channon 1981). These materials were subsequently superseded by other nonsystemic fungicides like captafol, daconil, dichlofluanid, propineb, bordeaux mixture, copper oxychloride, 13.6 Foliar Spray of Fungicides 289 mancozeb, ziram, chlorothalonil, and fentin hydroxide (Butler 1918; Butler and Jones 1949; Kolte 1985; Saharan and Chand 1988; Sherf and Macnab 1986; Vasudeva 1958). The list of fungicides found effective against downy mildew of crucifers at different locations is given in Table 13.4. The fungicides captafol, mancozeb, difolatan, copper oxychloride, dichlofluanid, propineb, and metalaxyl have been found to be superior to other fungicides on a large number of crucifers at several locations. The time of application of fungicides, and numbers, and interval of sprays depend on the duration and type of crop species grown (Channon et al. 1970; Kolte 1985; Saharan and Chand 1988; Saharan 1992a; Verma et al. 1994; Whitewell and Griffin1967 ).

13.6.1 Brassica Vegetables

In the UK, dichlofluanid gave excellent control of the disease on the cotyledons of cabbage and cauliflower. Dichlofluanid and propineb reduced the level of early mildew infection and increased the size and dry weight of cauliflower plants (Channon et al. 1970; Whitewell and Griffin 1967). RPA 407213 combined with fosetyl-Al was highly active against H. parasitica infecting Brassicaceae (Mercer et al. 1998). In the Irish Republic, downy mildew of Brassica crops, especially cauliflower, has been controlled by fosetyl-aluminium, metalaxyl + mancozeb, cyprofuram, and propamocarb (Ryan et al. 1984). In South Africa, during the initial years of contain- erized seedling production of cabbage, mancozeb (Dithane M-45), chlorothalonil (Bravo), metalaxyl (Ridomil), and metalaxyl plus mancozeb (Ridomil MZ-72) provided adequate control of downy mildew disease. Later on, cymoxanil plus mancozeb consistently provided the most effective control against downy mildew. Oxadixyl plus mancozeb, cupric hydroxide, and chlorothalonil gave significantly better protection than mancozeb (Brophy and Laing 1992). In Australia, neutralized phosphonic acid sprays applied on to cauliflower in the field within 3 weeks of harvest reduced downy mildew under storage conditions. Two applications of 2.4 kg a.i./h, 21, and 7 days before harvest reduced the curd infection development at the post- harvest stage in storage. There was no effect of phosphonic acid on crop appearance and maturity. The maximum phosphonate residue in curds at harvest was 12 μg/g, which was considered a safe limit (McKay et al. 1992). In Thailand, the best control of Chinese cabbage downy mildew was obtained with Ridomil 25 WP at 2 kg/h (Yang et al. 1983). Three sprays at weekly intervals beginning from 28 days after transplanting gave 65% more marketable yield. In India, four sprays with Difolatan (0.3%), Daconil (0.1%), Dithane M-45 (0.2%), Ridomil (0.2%), or Aliette (0.1%) at intervals of 8–10 days were most effective for controlling downy mildew of radish (Sharma and Sohi 1982; Sharma 1983) (Table 13.5). Root yield was significantly higher in sprayed plots. There was a significant reduction in the apparent infection (r) and the basic infection rate (R) of downy mildew in treated plots. According to Gupta and Shyam (1994), even a single application of metalaxyl + mancozeb or cymoxanil + 290 13 Disease Management

Table 13.4 Fungicides found effective against downy mildew of crucifers (Saharan et al. 1997 updated) Fungicide Rate of application References Cabbage Spergon spray (48% a.i.) 4 lbs./100 gallons Borders (1953) Spergon dust (4.8% a.i.) 30 lbs./acre Kolophygon dust (30% sulphur and 1% 30 lbs./acre phygon) Parzate dust (6.5% a.i.) 30 lbs./acre Dithane Z-78 spray (65%a.i.) 2 lbs./100 gallons Dithane Z-78 dust (6.5% a.i.) 30 lbs./acre Yellow cuprocide 1 lb./100 gallons Foster (1947) Dithane B-11 1 lb./100 gallons Spurgeon (Wettable) 4 lbs./100 gallons Dow seed treatment 2 lbs./100 gallons Fermate 2 lbs./100 gallons Phygon ¼ lb./100gallons Phenanthraquinone 1 lb./100 gallons Bordeaux mixture 1:1:10 Anonymous (1938) and Wiese (1927) Spergon (chloranil) (5 & 10% a.i.) − Epps (1955) Dithane Z-78 (zineb) (3.9% a.i.) − Phygon XL (dichlone) (10% a.i.) − Copper no. 30 (4.0% a.i.) − Thiram (5.0% a.i.) − Manzate (maneb) (4.2% a.i.) − Vancide F995 W (6.0% a.i.) − Vancide 51ZW (6.0% a.i.) − Ethyle B-622 (4.0% a.i.) − Metalaxyl 1.12 kg a.i./ha Jaworoski et al. (1982) Captafol (0.25% a.i.) − Channon and Hampson (1968) Daconil 2787 − Dichlofluanid − Propineb − Zineb − Maneb − Mancozeb − Quintozene − Quinomethionate − Copper oxychloride − Dichlone − Chloranil 4 lbs., 48%/100 gallons Anonymous (1953) Nabam-zinc sulphate 1 lb./100 gallons (continued) 13.6 Foliar Spray of Fungicides 291

Table 13.4 (continued) Fungicide Rate of application References Phygon XL-N 1 lb./100 gallons Difolatan 4F (captafol) 0.2% Apandi (1980) Polyram (metiram) 0.2% Dithane Z-78 0.2% Ciferri (1953) Dichlofluanid 0.05–0.2% a.i. Channon et al. (1970) Tribasic copper sulphate, sulphur − Lawson et al. (1998) RPA 407213 + fosetyl-al − Mercer et al. (1998) Aspor 0.3% Nakov (1968) Maneb 0.2% Perotsin 0.3% Nickel sulphate spray 0.05–0.2% Keyworth (1967) Polycaracin spray 0.4% Vasileva (1976) Cymoxanil + mancozeb (6 + 70% a.i.) 200 g/100 litres Brophy and Laing (1992) Cymoxanil + chlorothalonil (6 + 50% 200 g/100 litres a.i.) Oxadixyl (8%a.i.) 80 ml/100 litres Oxadixyl + mancozeb (8% + 56% a.i.) 330 g/100 litres Propamocarb + HCL (72% a.i.) 120 ml/100 litres Propamocarb + mancozeb (72 + 80% 60 ml/100 a.i.) litres + 75 g/100 litres Metalaxyl (Ridomil WP0 25% a.i.) 50 g/100 litres Metalaxyl + mancozeb (25 + 80% a.i.) 50 g/100 litres Fosetyl-AL-mancozeb (44 + 26% a.i.) 350 g/100 litres Chlorothalonil (50% a.i.) 100 ml/100 litres Mancozeb (80% a.i.) 200 g/100 litres Copper oxychloride (80% a.i.) 400 g/100 litres Cupric hydroxide (72% a.i.) 200 g/100 litres CGA-48988 soil application 23 mg a.i./M Gabrielson and Getzin (1979) Broccoli Agrimycin 0.1 lbs./acre Natti et al. (1956) Spergon SL 2 lbs./acre Streptomycin 50 ppm Altman (1958) Agri-strep 3 lbs./acre Natti (1957) Agri-strep + glycerol 3 lbs./acre Copper-zinc 6 lbs./acre Copper-manganese 6 lbs./acre Spergon SL 3 lbs./acre Manzate 4 lbs./acre Thioneb 6 lbs./acre Captan 50 W 6 lbs./acre Vancide M 4 lbs./acre (continued) 292 13 Disease Management

Table 13.4 (continued) Fungicide Rate of application References Kemate 50% 6 lbs./acre Manzate + agri-strep 4 + 0.4 lbs./acre Natti (1957), (1959) Copper-zinc + agri-strep 4 + 0.4 lbs./acre Natti (1957) Agrimycin 500 4.6 lbs./acre CGA-1-82 50 WP 2 lbs./5 ft.band soil Johnston and Springer application (1977) CGA-38140 50 WP 2 oz./acre 14 days after seeding CGA-48988 (Metaxadine) 1 or 2 oz. /100lbs seed Paulus et al. (1978) Cauliflower Dichlofluanid 50 WP 1½ lbs./100 gallons Whitewell and Griffin (1967) Captafol 85WP 3 lbs./100 gallons Zineb 70WP 3 lbs./100 gallons Propineb 70WP 3 lbs./100 gallons Daconil 2787 75 WP 3 lbs./100 gallons Dichlofluanid 0.05–0.2% a.i. Channon et al. (1970) Fosetyl-aluminium Ryan et al. (1984) Metalaxyl + mancozeb Cyprofuram Propamocarb Phosphonic acid 2.4 kg a.i./h McKay et al. (1992) CGA 245704 − Jensen et al. (1998) Apron 35 WS 2 g/kg seed Verma et al. (1994) Metalaxyl − Esiyok et al. (1996) Propineb − Esiyok et al. (1996) Captan − Verma et al. (1994) Rapeseed-mustard Cymoxanil − Gupta and Shyam (1994) Polyram M 2 lbs./100 gallons Perwaiz et al. (1969) Melprex 1.5 lbs./100 gallons Bordeaux mixture 4:4:50 (0.8%) Cuprovit 2 lbs./100 gallons Dithane M-45 2 lbs./100 gallons Dithane M-45 0.3% Bains and Jhooty (1979) Dithane Z-78 0.3% Blitox-50 0.3% Difolatan 80 0.2% Chauhan and Muheet (1976) Ziram 0.2% Dithane M-45 0.2% Thiovit 0.2% (continued) 13.6 Foliar Spray of Fungicides 293

Table 13.4 (continued) Fungicide Rate of application References Difolatan 0.2% Saharan, (1984, 1992a) Dithane M-45 0.2% Dithane Z-78 0.2% Blitox −50 0.3% Ridomil 0.2% Brestan 0.1% Apron SD-35 0.2% seed treatment Apron 35-SD 06 g/kg seed Singh and Singh (2005) Metalaxyl 0.2% Kavach 0.2% Mehta et al. (1996) Ridomil MZ-72 0.25% Apron SD-35 seed treatment + Dithane 2 g a.i./kg seed +0.2% M-45 spray Apron SD-35 seed treatment + Ridomil 2 g a.i./kg seed +0.2% MZ-72 spray Apron SD-35 seed treatment + 2 g a.i./kg seed +0.2% Difolatan spray Apron SD-35 seed treatment + Kavach 2 g a.i./kg seed +0.2% spray Mancozeb 75% WP 0.2% Singh and Singh (2005) Thiophanate methyl − Kumar and Kumar (1996) Iprodione − Kumar and Kumar (1996) Quintal 0.2% Shiwangi et al. (2017) Radish Difolatan seed treatment or spray 0.3% Sharma and Sohi (1982) Daconil seed treatment or spray 0.1% Dithane M-45 seed treatment or spray 0.2% Ridomil seed treatment or spray 0.1% Aliette seed treatment or spray 0.1% Blitox seed treatment or spray 0.2% Captan seed treatment or spray − Copper oxinate seed treatment or spray − Delan seed treatment or spray Dithane Z-78 seed treatment or spray Macuprax seed treatment or spray Stock Zineb 8 lbs./100 gallons Jafar (1963) Strepto spray +glycerol 500 ppm + 1% Trioneb 8 lbs./100 gallons Bordeaux mixture 5:5:50 Fongarid (CGA 38140) 0.05% Trimboli and Hampshire (1978) (continued) 294 13 Disease Management

Table 13.4 (continued) Fungicide Rate of application References Zineb 0.13% Camelina sativa Brestan Zarzycka and Kloczowska (1964) Polyram –M Sadoplon Copper oxychloride 0.4% Zarzycka and Kloczowska (1967)

Table 13.5 Efficacy of fungicidal sprays on downy mildew of radish (Sharma and Sohi 1982) Apparent Basic infection Yield of roots Disease index (%) infection rate (r) rate (R) (Kg/plot) Fungicides A B A B A B A B Aliette NT 6.37 NT 0.078 NT 0.59 NT 20.45 Blitox 17.07 12.27 0.108 0.088 2.36 0.93 18.45 22.02 Captan 17.18 11.03 0.115 0.083 3.08 0.79 19.85 19.65 Copper oxinate 24.46 8.25 0.128 0.083 4.60 0.81 18.85 18.47 Deconil 3.70 3.40 0.055 0.051 0.28 0.21 23.15 24.45 Delan 19.75 8.22 0.123 0.083 4.04 0.81 19.67 19.17 Difolatan 5.88 4.10 0.079 0.053 0.81 0.23 25.67 24.06 Dithane M-45 7.35 7.55 0.084 0.071 0.99 0.48 22.37 24.12 Dithane Z-78 13.67 10.30 0.109 0.092 2.56 1.10 21.52 21.61 Macuprax 26.82 13.20 0.130 0.101 4.84 1.53 17.92 18.43 Control 55.79 19.51 0.171 0.109 13.06 1.93 17.30 17.73 SEM± 1.74 0.942 – – – – 1.156 1.020 CD at 5% 5.049 2.722 – – – – 3.356 2.947 A = December 1979–February 1980; B = June–September 1980; NT = not tested mancozeb gave good control of downy mildew of cabbage. Verma et al. (1994) found best control of cauliflower downy mildew with metalaxyl seed treatment followed by captan.

13.6.2 Brassica Oilseeds

For the control of downy mildew of mustard, difolatan, mancozeb, and metalaxyl have been found to be very effective at different locations (Table 13.6) in India. An effective and economical schedule has been worked out under Indian conditions for the control of downy mildew of mustard through seed treatment and/or spray with fungicides. Three sprays of Ridomil MZ-72 (metalaxyl and mancozeb at 0.25%) at an interval of 20 days starting from 40 days after sowing gave maximum disease 13.6 Foliar Spray of Fungicides 295

Table 13.6 Efficacy of fungicidal treatments on the downy mildew of mustard in India (Saharan 1984, 1992a) Percent disease intensity Fungicide Conc. (%) Durgapura Hisar Pantnagar Difolatan 0.2 10.65 9.80 15.09 Dithane M-45 0.2 8.31 12.00 10.79 Dithane Z-78 0.2 – 14.80 14.50 Blitox 0.2 15.00 14.80 16.77 Ridomil 0.2 16.25 8.00 – Control – 23.00 24.80 16.80 CD at 5% – 5.90 5.69 control (82%) along with >49% increase in yield; seed treatment with Apron SD-35 (metalaxyl at 2 g a.i. kg−1 seed) along with two foliar applications of Ridomil MZ-72 at 30 days interval was relatively less effective (Tables 13.7 and 13.8). These treatments were quite effective in reducing stag head formation in mustard (Table 13.9). When mancozeb (Dithane M-45) and chlorothalonil (Kavach) were sprayed three times following seed treatment with Apron SD-35, disease control of around 47% and 49%, respectively, was achieved. The maximum cost-benefit ratio was either with four sprays of mancozeb or seed treatment with Apron SD-35, followed by three spays of mancozeb (Mehta et al. 1996). Absorption of metalaxyl increased, up to 30 days, when applied as seed treatment; thereafter, it gradually declined and was not detectable after 60 days of sowing (Table 13.10). The maximum residue (average 9.03 ppm) of metalaxyl was found to be 1 day after spraying (Table 13.11). The metalaxyl on mustard plants was almost undetectable 15–30 days after spraying (Table 13.12). The safe waiting period for metalaxyl was calculated to be 62 and 8 days for seed treatment and for foliar application, respectively (Table 13.13). No metalaxyl was detected in mustard seedlings raised from seeds obtained from these treatments (Mehta 1993) (Table 13.14). Quintal at 0.2% was found effective in reducing the downy mildew of mustard severity followed by mancozeb at 0.25% (Shiwangi et al. 2017). Seed treatment with metalaxyl at 6 g/kg seed and seed treatment with metalaxyl combined with 2 sprays of mancozeb + metalaxyl (Ridomil MZ-72) at 0.2% were most effective for the control of downy mildew of rapeseed (Puzari and Saikia 1997). Maximum yield was obtained in mustard with iprodione, but mancozeb + thiophanate methyl gave the best cost-benefit ratio (Kumar 1996). Singh and Singh (2005) observed the effect of fungicidal treatment on percent disease intensity, seed yield, and avoidable yield losses of mustard (Table 13.15). 296 13 Disease Management Average 42.4** 35.0 48.2 81.8** 68.5** 47.4 49.6 51.8 32.7** – (90) 1992–1993 45.2 – – 81.6 66.6 – – – 28.6 – (60) ‡ Percent disease control 1991–1992 39.7 35.0 48.2 82.0 70.3 47.4 49.6 51.8 36.9 – 9.1 28.4 28.2 22.4 15.8 22.9 21.9 20.9 33.6 49.7 (90) 7.0 3.5 2.7 4.0 6.1 0.4 1.2 1.8 8.7 14.2 (60) Average (90) 30.7 – – 10.3 18.7 – – – 40.0 56.0 4.7 11.3 12.3 15.3 23.7 (60) 1992–1993 – – – – – 7.8 26.2 28.2 22.4 12.9 22.9 21.9 20.9 27.3 43.4 (90) (60) 1991–1992 ‡ Percent disease index (DAS) ‡ Percent disease index 2.7 3.5 2.7 0.2 0.0 0.4 1.2 1.8 2.2 4.8 No. of sprays 4 4 4 3 2 3 3 3 – – Conc. (%) 0.2 0.2 0.2 0.25 – 0.25 0.2 0.2 0.2 – – Efficacy and spray schedule of fungicides against downy mildew of mustard during 1991–1992 and 1992–1993 crop seasons (Mehta et al. 1996 ) mildew downy and spray schedule of fungicides against Efficacy Treatments Dithane M-45 Kavach Difolatan Ridomil MZ-72 *Apron SD-35 + Ridomil MZ-72 *Apron SD-35 + Dithane M-45 *Apron SD-35 + Kavach *Apron SD-35 + Difolatan *Apron SD-35 Unsprayed (control) Table 13.7 Table -1 seed, ** two years mean * seed treatment at 2 g a.i.kg -1 seed, ** two days after sowing, of four replications, () DAS ‡ average 13.6 Foliar Spray of Fungicides 297

Table 13.8 Comparative yield increase and cost-benefit ratio of fungicides used against downy mildew of mustard (Mehta et al. 1996) †Average yield/ % Increase in yield Cost-benefit ratio** Treatments. plot* (Kg) over control Rs:Rs:Ps Dithane M-45 1.164 28.5 1:2.20 Kavach 1.116 23.3 1:1.57 Ridomil MZ-72 1.352 49.3 1:1.21 ***Apron SD-35 + Ridomil 1.216 34.2 1:1.11 MZ-72 ***Apron SD-35 + Dithane 1.115 22.1 1:2.11 M-45 ***Apron SD-35 + Kavach 1.090 20.4 1:1.29 ***Apron SD-35 1.050 15.9 1:20.62 Unsprayed (control) 0.950 – – † average of four replications; * plot size, 2.0 × 2.1 m2; **based on prevalent market price in 1992 Raya, Rs. 800/− Q; Dithane M-45, Rs. 152/− Kg; Ridomil MZ-72, Rs. 950/Kg; Kavach, Rs. 333/ Kg Apron SD-35, Rs.2782/Kg; labour, 5 labour/spray/hectare at Rs.40/− per labour; *** seed treatment at 2 g a.i./Kg−1 seed

Table 13.9 Efficacy of fungicides against stag head of mustard due to combined infection of white rust and downy mildew (Mehta et al. 1996) No. of † Stag head ** † Stag head ** † Stag head Treatments sprays incidence (%) length (cm) ** score Dithane M-45 4 8.8 (16.2) 7.9 2.1 Kavach 4 2.4 (8.9) 8.9 1.4 Ridomil MZ-72 3 1.6 (6.4) 5.4 1.5 Apron SD-35 2 5.2 (12.0) 8.1 2.0 *** + Ridomil MZ-72 Apron SD-35 3 2.6 (9.2) 1.9 1.0 *** + Dithane M-45 Apron SD-35 3 2.9 (9.9) 3.7 1.7 *** + Kavach Apron SD-35*** – 13.9 (19.0) 10.4 2.3 Unsprayed (control) – 25.8 (30.4) 14.5 3.4 LSD (0.05) (2.7) () angular transformed values, † average of four replications, *** seed treatment at 2 g.a.i. Kg−1 seed, ** two years mean (1991–1992, 1992–1993), *** seed treatment with Apron SD-35 @ 2 g a.i. per Kg seed 298 13 Disease Management

Table 13.10 Persistence of metalaxyl in mustard foliage after seed treatment (Mehta 1993) Days of Average residue Range * Dissipation Treatments sampling level (ppm) (ppm) (%) SD± Apron SD-35 at 2 g.a.i. 7 1.81 1.58–2.02 0.00 0.23 Kg−1 seed’ 15 3.46 3.03–3.80 +191.46 0.37 30 9.08 8.63–9.84 +242.42 0.54 40 5.82 5.57–6.01 35.90 0.18 60 0.00 – 100.0 – * = average of three replications

Table 13.11 Persistence of metalaxyl in foliage of mustard after foliar application (Mehta 1993) Average residue level* (ppm) Treatments Days 1 5 10 15 30 ** Foliar spray-I 9.03 0.68 0.28 0.0 0.0 Range 7.94–10.08 0.49–0.80 0.25–0.32 – – Dissipation (%) 0.00 92.46 96.89 100.0 100.0 Foliar spray-II 10.37 0.54 0.29 0.0 0.0 Range 9.57–11.10 0.51–0.57 0.29–0.30 – – Dissipation (%) 0.00 94.79 97.20 100.0 100.0 SD± I-spray 0.830 0.169 0.040 – – II-spray 0.812 0.034 0.005 – – * average of three replications, ** foliar spray 40 and 70 days after sowing at 0.25%

Table 13.12 Persistence of metalaxyl in mustard foliage after seed treatment and foliar sprays (Mehta 1993) Average residue level* (ppm) Treatments Days 1 5 10 15 30 ** Foliar spray -I 8.21 0.49 0.26 0.0 0.0 Range 7.89–8.76 0.47–0.52 0.23–0.29 – – Dissipation (%) 0.0 94.03 96.83 100.0 100.0 Foliar spray-II 9.45 0.69 0.27 0.0 0.0 Range 8.64–10.22 0.59–0.80 0.25–0.30 – – Dissipation (%) 0.00 92.69 97.14 100.0 100.0 SD± I-spray 0.382 0.028 0.034 – – II-spray 0.673 0.121 0.029 – – * average of three replications, ** foliar spray 60 and 80 days after sowing at 0.25% 13.7 Biological Control 299

Table 13.13 Safe period and Treatments RL-50 (days) SWP (days) residue half-life values of Seed treatment 17.57 62.33 metalaxyl in mustard (Mehta 1993) Foliar spray-I 2.08 8.69 Foliar spray-II 1.79 7.85 RL-50, residue half-life; SWP safe waiting period

Table 13.14 Translocation Treatments Average residue (ppm) of metalaxyl residues into Seed treatment ND mustard seed following * (apron SD-35 different treatments at harvest (T1) 60DAS (Mehta 1993) Foliar sprays ND ** (Ridomil MZ-72 (T2) 40,70 DAS Seed treatments ND + foliar sprays (T3) 60,80 DAS ND not detectable; DAS days after sowing; *, at 2 g.a.i. Kg−1 seed; **, at 0.25%

13.7 Biological Control

13.7.1 Plant Extracts as Fungitoxicant

Garlic juice or aqueous extracts of garlic was reported to be toxic to H. parasitica which causes downy mildew of radish (Ark and Thompson 1959).

13.7.2 Antagonists for Biocontrol

Bacteria were observed on the mycelium, conidiophores, and conidia of H. parasitica on Lepidium graminifolium (Nicolas and Aggery 1940). This was associated with a reduction in conidial germination. Bedlan (1987) obtained good control of H. parasitica in radish with an isolate of Trichoderma harzianum under greenhouse conditions. Paraffin oil-based Trichoderma formulations reduced downy mildew, white rust, and Alternaria blight of mustard under organic farming (Saxena and Tewari 2017). 300 13 Disease Management 1:8.8 Cost- benefit ratio 1:5.3 1:1.0 1:4.9 1.09 – 705.7 Additional income (Rs.) 10131.7 7845.7 8907.7 6900.7 – 5.6 Avoidable Avoidable loss (%) 46.2 39.9 43.0 36.9 – 58.8 Additional yield (Kg/ha) 844.3 653.8 742.3 575.1 – 1042.7 Yield Yield (Kg/ha) 1828.2 1637.8 1726.3 24.38 1559.0 983.9 9.91 62.5 Mean 26.0 29.8 30.5 36.0 67.1 Downy Downy mildew 42.5 (40.67) 20.4 (26.85) 10.7 (19.07) 24.0 (29.33) 19.5 (26.17) 1.37 46.4 (42.93) 0.56 White rust 55.3 (45.05) 27.0 (30.96) 15.4 (23.05) 32.6 (31.83) 20.6 (26.98) 1.40 61.3 (51.51) 0.57 Pod 78.4 (62.31) 32.6 (34.79) 51.5 (45.84) 37.8 (37.92) 57.8 (49.48) 1.87 82.6 (65.37) 0.76 Alternaria blight Leaf 73.7 (59.18) Percent disease intensity 24.00 (29.35) 41.7 (40.24) 27.6 (31.69) 46.3 (42.86) 1.55 78.0 (62.04) 0.63 Effect of fungicidal treatments on percent disease intensity, seed yield, and avoidable yield loss for 1996–1997 to 1998–1999 (Pooled data) seed yield, and avoidable of fungicidal treatments on percent disease intensity, Effect Seed treatment with apron SD-35 at 6 g/kg seed (ST) Treatments ST-Indofil M-45 (0.20%) ST-Indofil ST-Ridomil MZ-72 ST-Ridomil (0.025%) Indofil M-45 (0.020%) Ridomil MZ-72 (0.25%) Cd (P < 0.05) Untreated check SEM Table 13.15 Table (Singh and Singh 2005 ) 13.10 Integrated Disease Management 301

13.8 Host Resistance

Many sources of resistance to downy mildew of crucifers have been identified in major host species from various parts of the world. All sources of R identified so far are race specific and governed by major genes. Information is also known on the genetics of the host-parasite interaction. Efforts are being made to breed downy mildew-resistant cultivars in various crucifers through conventional and biotechnological techniques.

13.9 Fungicide Resistance

French isolate, P006, and FP 13 of H. parasitica were insensitive to metalaxyl at 0.5, 5.0, and 50 μg metalaxyl/ml (Vishunavat et al. 1998). However, according to Molina et al. (1998), application of systemic acquired resistance activator benzothiadiazole (BTH) in combination with metalaxyl, fosetyl, and Cu(OH)2 fungicides resulted in synergistic effect on H. parasitica in wild-type plants of Arabidopsis and an additive effect in NahG and BTH-unresponsive niml plant. Interestingly, BTH treatment normally induces long-lasting pathogen protection, but in NahG plants, the protection is transient. In contrast, the effectiveness of these fungicides is not altered in Arabidopsis mutants defective in the ethylene or jasmonic acid signal transduction pathways.

13.10 Integrated Disease Management

In the quadrangle of integrated control (chemical-cultural-biological-host resistance) of downy mildew of crucifers, biological control has not been exploited at the field scale. Breeding for resistance has only succeeded in some crucifers. Chemical control of the disease may not always be reliable as resistance has been developed in H. parasitica to metalaxyl, which at one stage proved outstandingly effective in the control of downy mildew (Brophy and Laing 1992; Crute et al. 1985). Thus, there is clearly a need to breed sources of host resistance that would counter pathogenic variation. It is also possible that differential sources of host resistance could be useful in programmes of integrated control if they were deployed together with fungicides; this would potentially prolong the effectiveness of both control procedures (Silue et al. 1996). Other methods involve sanitation, field practices like sowing time, plant density, and the judicious use of nutrition and irrigation so that inoculum levels will not build up too rapidly. Butler (1918) reported long time ago that the disease can be controlled in young crucifer plants by a mulch of sawdust saturated with copper sulphate placed around the base of the plants. 302 13 Disease Management

In the Shanghai region of China, a combination of seed treatments, direct seeding, and application of fertilizer and 2–3 fungicide sprays at the first peak infection period decreases the incidence of H. parasitica in Chinese cabbage and increases yield by 10–18% (Shao et al. 1991). The multiple disease control strategy is mainly dependent on the balanced fertilizer application (N100; P40: K40) and use of available level of tolerance in host varieties along with timely sowing of the crop (first fortnight of October) and seed treatment with Apron 35-SD followed by use of fungicidal spray (metalaxyl + mancozeb = Ridomil MZ-72 at 0.25%) for Alternaria blight white rust + downy mildew disease complex control or carbendazim spray at 0.05% for Sclerotinia rot and powdery mildew disease control (Kolte 2005) (Table 13.16). Godika et al. (2001) and Kolte (2005) reported that the combination of boron (0.53%) with boric acid or zinc (0.22%) spray through zinc oxide showed synergistic effect in the effectiveness of mancozeb and gave 16–20% improvement in disease control in comparison with such treatments when used separately. It indicates the importance of integrating plant nutrients such as boron, potassium, and zinc with foliar sprays of fungicides for management of the downy mildew, white rust, and Alternaria diseases in oilseed rape (Table 13.17). No single method or approach can be viable, stable, effective, environmentally safe, and economically feasible with any biological system, particularly disease management system. Therefore, all means of control measures like cultural, biological, chemical, and host resistance including genetic engineering should be used to manage cruciferous diseases in an integrated way (Mukerji et al. 1999; Saharan and Mehta 2002; Saharan et al. 1997, 2005). The following general practices should be followed to manage the diseases of cruciferous crops under field conditions. Cultural control of diseases of crucifer’s crops is largely a matter of sanitation and of manipulating the environment to the advantage of the host and to the deter- ment of the pathogen. Various cultural practices useful in reducing the diseases are as under: (i) Pathogen H. parasitica survives in the form of thick-walled oospores; diseased debris containing these resting structures should be burnt or destroyed after the harvesting of crop. (ii) Deep ploughing should be done during hot summer months to expose and probably destroy (May–June) the resting propagules of the pathogen. (iii) Crop rotation for a period of at least 3 years should be followed with non- cruciferous crops after epidemics. (iv) Proper plant to plant and row to row spacing should be maintained to make microenvironment less conducive to disease. (v) High relative humidity around the plant canopy should be avoided to have proper aeration. (vi) The cruciferous and other weed host plants should be removed. (vii) Avoidance of continuous cropping of rapeseed-mustard, particularly the same variety, helps in reducing disease severity and build-up of oosporic inoculum (primary source of infection). 13.10 Integrated Disease Management 303 Yield Yield increase over check % 72.03 37.08 84.06 112.79 Yield Yield potential (kg/ha) 1901.6 1105.1 902.7 658.0 1918.1 1042.7 904.8 425.2 900.4 1000 grain weight (g) 3.74 3.36 3.35 3.04 3.76 3.45 3.33 2.99 0.51 DM + WR stag head (severe) 4.64 4.71 15.75 21.50 3.71 5.06 16.79 20.24 13.77 DM + WR stag head (incidence) 1.92 2.13 19.01 16.11 1.76 2.64 20.19 16.85 19.45 , Kranti 2 (Kolte 2005 ) (Kolte a AB pod 2 WBM (%) 46.92 52.74 39.88 47.17 46.65 51.75 38.95 49.16 8.23 , Varuna; V , Varuna; 1 AB leaf 100 DAS (%) 45.45 59.41 52.77 58.97 46.14 59.65 52.77 59.70 10.59 WR leaf (%) index 100DAS 29.34 40.42 35.47 43.31 28.44 40.64 35.24 – 10.47 , sowing date 20 November; V date 20 November; , sowing 2 DM leaf 60 DAS (%) 33.58 36.84 22.06 24.06 34.13 37.57 19.10 21.50 13.24 Integrated disease management module (seed treatment, spray schedule, and fertilizer doses for the control of DM, WR, AB) and its significance WR, disease management module (seed treatment, spray schedule, and fertilizer doses for the control of DM, Integrated DM cotyledons (%) index 17.71 62.28 14.99 44.90 17.71 63.06 14.99 42.03 16.06 0 1 0 1 0 1 0 1 P P P P P P P P 0 1 0 1 0 1 0 1 P P P P P P P P 1 1 1 2 2 2 2 I , recommended plant protection practices, i.e., NPK = 100:40;40 kg ha-1 , no plant protection chemical treatment 1 0 V V V V V V V V I 1 2 2 1 1 2 2 , sowing date 20 October; D , sowing P P 1 0 1 Treatments D D D D CD at 5% D D D D downy mildew; WR white rust; AB Alternaria blight mildew; DM downy D P Apron 35 SD (6 g/kg) Seed treatment with by mancozeb spray at 0.2% 70 and 90 DAS followed Ridomil MZ at 0.25spray 50 DAS of 3 years crop seasons 1999–2000 to 2001–2002 aData indicated in the table are averages Table 13.16 Table higher yield of mustard during 1999–2000 to 2001–2002 in achieving P 304 13 Disease Management

Table 13.17 Some micronutrients as possible inducer for multiple disease resistance in rapeseed- mustard (Kolte 2005) Phytoalexin Disease index Treatments inhibition Under field Yield/20 (%) Conc. zone (mm) Under artificial conditions conditions plants (gm) DM WR AB WR AB Fe EDTA 0.2 95.0** 66.9** 77.0* 39.8 36.7** 40.2** 85.6

MnSO4 0.2 95.9** 43.9** 56.3** 64.5 28.6** 50.2 88.9

CaCl2 1.0 90.9** 88.8 56.9** 24.4** 25.6** 46.2* 82.4

ZnSO4 0.5 119.9** 4.1** 37.0** 39.2 48.3 49.7 109.0*

CuSO4 0.1 119.0** 9.5** 4.0** 54.6 21.9** 47.7* 111.3*

Co(NO3)2 0.5 69.2** 59.3** 53.8** 28.3** 29.1** 46.6* 118.5*

Na2BO7 0.5 53.5** 31.8** 18.4** 23.0** 29.3** 50.2 11.4* Distilled 8.5 92.6 83.6 36.9 47.7 53.5 77.9 – water **significant at P < 0.01, *significant at P < 0.05 DM downy mildew; WR white rust; AB Alternaria blight

(viii) Optimum doses of fertilizers should be used. (ix) Time of sowing of the crop should be adjusted depending upon weather conditions conducive to disease development in particular area. (x) Early sowing (up to 20 October) of the crop reduces incidence of major diseases including DM. (xi) Planting of cultivars with high degree of tolerance. (xii) Plant healthy and treated seeds. (xiii) Use foliar fungicidal spray, if necessary.

References

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White JG, Crute JR, Wynn EC (1984) A seed treatment for the control of Pythium damping-off diseases and Peronospora parasitica on Brassicas. Ann Appl Biol 104:241–247 Whitewell JD, Griffin GW (1967) Chemical control of downy mildew in seedling cauliflower. Proceedings of Fourth British Insecticide Fungicide Conference, Brighton, pp 239–242 Wiese E (1927) A fungus disease of wallflowers and stocks. Gartenwelt 31:486 Yang CY, Hu RLF, Pupipat U (1983) Control of Chinese cabbage downy mildew (Peronospora parasitica) by metalaxyl (Ridomil) and other fungicides. International Symposium on Pesticides in Developing Countries – Present and Future, Ibaraki, Japan. Tropical Agricultural Research Center, pp 125–130 Zarzycka H, Kloczowska T (1964) Results of investigations on the control of downy mildew of false flax (Camelina sativa). Biul Inst Ochr Rosl Poznan 26:231–248 Zarzycka H, Kloczowska T (1967) Investigations on the control of Peronospora parasitica of false flax. Biul Inst Hodowl Aklim Rosl 6:146–159 Chapter 14 Techniques

14.1 Introduction

Laboratory, and field techniques, and bioassays are very important methods, procedures, and tools for the researchers and graduate students to use for their research achievements. In this chapter, a number of standardized, reproducible techniques have been included which will be very helpful for Brassicalogists to validate, refine, and update their research pursuits on all aspect using crucifer-Hyaloperonospora pathosystem.

14.2 Culturing of Hyaloperonospora parasitica

The biotrophic nature of this pathogen implies a sophistication of nutritional requirement which cannot be met upon the death of the host plant. H. parasitica was cultured on disinfected slices of swede root where aerial growth of mycelium was observed with conidiophores, conidia, antheridia, oogonia, and oospores (Guttenberg and Schmoller 1958). Ingram (1969) successfully established and maintained cultures of the fungus on callus tissues derived from (a) a mature leaf of cabbage, (b) a mature root/hypocotyl of rape, and (c) a seedling hypocotyl of swede. The infected calluses were incubated either at 22 °C in the dark or at 15 °C with 12 h fluorescent illumination photoperiod. To maintain the dual culture of the callus and the pathogen, subculturing through transfer of explants of infected callus to fresh uninfected callus was necessary every 14–21 days. Calluses derived from the root/hypocotyl of rape grew faster than those from cabbage leaf and were transferred directly when subculturing. About 4 weeks after inoculation of the rape callus, small nodules of new tissue were developed on the infected callus. These nodules or the whole callus were successfully maintained when transferred to fresh culture medium. Conidia of H. parasitica were produced on infected callus tissue maintained at 15 °C and 12 h/

© Springer Nature Singapore Pte Ltd. 2017 309 G. S. Saharan et al., Downy Mildew Disease of Crucifers: Biology, Ecology and Disease Management, https://doi.org/10.1007/978-981-10-7500-1_14 310 14 Techniques photoperiod, but production was much lower at 22 °C in the dark. Such conidia were used to infect detached cotyledons or leaf callus of cabbage or rape. Attempts were made with limited success by Guttenberg and Schmoller (1958) to culture the fungus in the absence of living plant tissue. They obtained visible mycelial growth in filter, sterilized swede juice, but it ceased after 3 days when a yellow discoloration appeared suggesting that the medium was chemically unstable. Very limited mycelium was developed on swede seed-glucose agar and maize decoctions-glucose agar, but more success was achieved with an agar medium containing 2% beer wort +0.1% phosphate in which hyphae and conidiophores were developed within and outside the agar substrate. Similar but less growth was achieved in oatmeal agar and rice starch agar. Asada and Ohguchi (1981) studied the behaviour of downy mildew fungus of Japanese radish on modified Knop’s medium. An isolate of H. parasitica from naturally infected leaves was cultured on modified Knop’s medium and 0.1% streptomycin, using infected slices of radish root tissue as inoculum. Vigorous hyphal growth was observed spreading into the medium, and numerous haustoria-like bodies were formed. The production of which was favoured by low agar concentration, low pH (4), and high sucrose concentration (50 g/l). Growth was ceased 2 weeks after placing the tissue slices on the medium. The response of H. parasitica to a liver medium was studied by McMeekin (1981). Washed, autoclaved 2 mm cubes of liver were placed in a 9 cm Petri dish and covered with 20 ml of a mixture of 0.01% tryptone (Difco, pancreatic digest of casein), 0.04% K2HPO4, and 2% agar: a 1/10 dilution of the K2HPO4 and tryptone. Ten μg/ml streptomycin (Calbiochem) were added to this medium to control bacterial growth. When the agar was solidified, drops of a suspension of conidia in water were placed on the piece of fibre. The plates were incubated at 18 °C. After 4 days the germ tubes in the control (without liver) disintegrate, but in a 5 mm diam. Zone around the liver pieces, 90% of the germ tubes grow towards the liver. They grow from the surface towards the bottom of the medium and form large swellings or lobes within the agar. The lobed germ tubes reached their maximum size after 4 days. When the plate was flooded with sterile distilled water, either 4 or 7 days after the beginning of conidial germination, the lobes on the germ tubes maintained the same size.

14.3 Maintenance of H. parasitica Isolates and Production of Inoculum

Isolates of H. parasitica were maintained separately on cotyledons obtained from 6-day-old seedlings, raised in soilless compost in a modified plant propagator (35.5 cm × 21.6 cm × 18 cm), sited in the glasshouse (Nashaat and Rawlinson 1994). The propagator was supplied with continuous filtered (spore-free) moist air at 18 ± 2 °C through a central flue conducting air from beneath the propagator to 14.4 Germplasm Screening and Evaluation 311 exhaust at two adjustable ventilators on the cover and the junction between the cover and the base (Jenkyn et al. 1973). Supplementary light was given to maintain a 16 h photoperiod. Cotyledons and a short length of hypocotyl were detached and transferred to folded filter paper (Whatman 12.5 cm, 113 v) supports in glass jars (8 cm diameter, 7 cm depth) containing 20 ml sterile distilled water. Cotyledons were then inoculated in a sterile air flow with 5 μl of conidial suspension on each half cotyledon with the aid of a micropipette. Conidial suspensions were prepared by tapping infected cotyledons to dislodge conidia into sterile distilled water; this minimized bacterial contamination. After inoculating the cotyledons, the glass jars were covered with clear plastic lids, sealed with Parafilm, and incubated in a growth cabinet at 16 °C under 70 μE/m2 s1 irradiance with a 16 h photoperiod for 7 days after which peak sporulation occurred. Once a culture is actively growing, it must be maintained continuously on living host tissue to provide inoculum for experiment. Prepare inoculum as follows: Collect cotyledons and leaves with profuse sporulation from 15 to 20 seedlings placing the tissues in a glass vial containing 3 ml double-distilled water, shake the tube to wash conidiosporangia off, remove the mass of plant tissue with forceps, and transfer the suspension to two 1.5 ml Eppendorf centrifuge tubes. Wash the inoculum as described in Sect. 14.25, combining the pellets after the first centrifugation and replacing the final supernatant with 0.5 ml double-distilled water. Quantify the suspension using a haemocytometer, and adjust the concentration of inoculum to 5 × 106 conidiophores per ml. Inoculate cotyledons of 10–14-day-old seedlings by placing 1–2 μl of inoculum on each cotyledon, thereby delivering about 100–200 conidiosporangia per plant. Incubate in a clear plastic box to maintain high humidity conditions at 16–18 °C, 10 h photoperiod, and 50–100 μE m2 s−1. Spray plants daily with a fine mist of water. H. parasitica will begin to sporulate 4–5 days after inoculation. But peak sporulation will occur after 7–10 days and may continue for another week. Subculturing at weekly intervals is advisable (Dangl et al. 1992).

14.4 Germplasm Screening and Evaluation

Genotypes are grown in propagators (as described under isolate maintenance in Sect. 14.3), except that two adjacent 5 cm ‘Jiffy pots’ for each line or cultivar are used as pots. The pots are placed on capillary matting to ensure a uniform water supply. Each propagator contains up to 13 accessions arranged as two randomized blocks (propagators) with each accession occurring only once in each propagator. Initially, 9–15 seedlings per accession are grown in each propagator, and these are thinned 6 days after sowing from 6 to 10 to decrease variability in growth. Sowing dates are staggered to produce seedlings at the required growth stage for inoculation at the same time. The average times required under these conditions to reach fully expanded cotyledons, first, and second true leaves are 7, 16, and 22 days, respectively (Nashaat and Rawlinson 1994). 312 14 Techniques

Seedlings are inoculated by spraying them to run off with a suspension of conidia (2.5 × 106 conidia/ml). The propagators are sealed after inoculation to allow the relative humidity to rise to 100% and then incubated in growth cabinets under the conditions described for isolate maintenance. Infection phenotypes are recorded 7–9 days after inoculation (on cotyledons and leaves, respectively) using a 0–9 scale (Nashaat and Rawlinson 1994).

Percentage disease Host-pathogen interaction 0.0 No infection 0.1 Traces of infection, generally confined to lower leaves, one plant in ten or fewer with lesions 1 Some plants infected, but one or two lesions per plant 5 Most plants infected with about 5% of the lower leaf area affected 10 Most plants with about 10% of the lower leaf area are affected. Up to 5% infection on the upper stem leaves and bracts 25 About 25% of the lower leaf area is affected. Leaf area may appear to be half affected and half unaffected. Infection frequent on upper stem leaves and bracts (up to 10% area affected) 50 About 50% of the leaf area is affected. Affected area appears to be greater than unaffected 75 About 75% of the leaf area is affected. Very little unaffected tissue observable 100 100% of leaf area affected

According to Williams and Leung (1981), single seeds are grown in 12-pack pots, and when the cotyledons have expanded, after 5–7 days, single 0.01–0.02 ml drops of a freshly collected conidial suspension containing approximately 1 × 105 conidia/ml are placed on each half of the two cotyledons using a finely tipped glass pipette. As plants of each 12 pack are inoculated, they are placed in glass or plastic boxes containing a 1–2 cm depth of warm water in the bottom. A tight-fitting cover is placed on the box after the box is filled with plants. The box is then placed in a darkened incubator at 20 °C for 8–16 h. The atmosphere in the box will maintain the droplets on the cotyledons during which time germination, and penetration will occur. After incubation, plants are placed on a lighted greenhouse bench at 20–25 °C for 5 days and then returned to humidity boxes at 20 °C for 16–24 h. Upon removal from the humid atmosphere, susceptible plants will have a profuse growth of H. parasitica conidiophores on the abaxial sides of the cotyledons, whereas resistant plants will exhibit varying degrees of sporulations and tissue necrosis which can be evaluated on a 0–9 scale (Fig. 14.1). A rating of 1 is given to rapidly occurring (24–48 h) hypersensitive necrotic flecking, without visible sporulation, found immediately under the droplets. Interaction phenotypes representing host-pathogen compatibility are expressed as increasing degree of sporulation on the abaxial side of the cotyledons and decreasing degree of necrosis associated with tissue colonization. It is important to use freshly produced inoculum collected by washing off spores from leaves with distilled water. Older plants may be inoculated by atomizing a suspension of conidia on the foliage and holding them at 100% RH for 8–16 h. 14.5 Preservation of Hyaloperonospora parasitica 313

Fig. 14.1 Rating scale for downy mildew (Hyaloperonospora parasitica) interaction phenotypes on Chinese cabbage (Williams and Leung 1981)

Knight and Furber (1980) used descriptive key for the assessment of downy mildew disease prevalence and severity on winter oilseed rape varieties. A group of plants (10 plants/sample) chosen at random from a number of observation site assessed were as follows. However, different disease assessment scales have been used/suggested to classify downy mildew interaction phenotypes in cruciferous crops (for details refer to Chap. 2 Sect. 2.6).

14.5 Preservation of Hyaloperonospora parasitica

Brassica leaves infected with the pathogen were collected, and conidia from such infected material were inoculated on cotyledons of a susceptible Brassica variety (Paul and Klodt-Bussmann 1993). Cotyledons are put into plastic boxes on moist filter paper and incubated at 15 °C and 70–80% RH. Under these environmental conditions, conidia and conidiophores of the pathogens are ready to be harvested 6 days after inoculation. For each isolate, five cotyledons colonized with fresh conidia were collected in a glass vial in 10% (v/v) glycerine which serves as a cryo- protectant in the suspension medium. For each isolate, six glass vials are filled with the conidial suspension and immediately stored in a freezer at −21 °C. After a storage interval, samples are taken out from the freezer and thawed at room temperature (20 °C). After thawing for 10 min, 2 ml of the conidial suspension are added to each Petri dish which contains 15% (w/v) water agar, and the percentage of conidial germination is assessed after 24 h at 15 °C. After a storage period of 8 days, it was found that the highest germination rate of 73% occurred using 10% (v/v) glycerine. To culture and preserve Japanese radish downy mildew fungus, slices (1 cm) of Japanese radish root cv. Miyashige were inoculated with conidia of the fungus collected from naturally infected leaves and incubated at 20 °C for 6–8 days (Ohguchi and Asada 1981). Conidia produced on the infected slices were then collected to make a suspension and were used to inoculate other healthy slices. Numerous oospores are observed in these sliced tissues 6 days after inoculation. 314 14 Techniques

The conidial viability of H. parasitica derived from cabbage was longest, up to 130 days, when the spores were stored in air-dried soil at a constant temperature of 5 °C. Conidia kept at −25 °C and relatively dry on leaf discs (air-dried at 20 °C) maintained a relatively high rate of germination after 1 year or longer (Krober 1970, 1981). The pathogen is usually preserved by storing few sporulating cotyledons in small vials with tight lids inside a deep freezer (−25 to −30 °C) for up to 6 months. Temperature fluctuations should be avoided during this period. For reviving the pathogen, the vials containing the sporulating cotyledons are taken out of the freezer and, with their lids kept tightly on, transferred immediately to a container containing icy water. The temperature of the container and its content is then allowed to rise gradually, within 1–2 h, to 15 °C. Thereafter, the vials are taken out of the container, and conidial suspension for inoculating fresh cotyledons is prepared in the normal way (Nashaat and Rawlinson 1994). A long-term preservation of H. parasitica conidia in freezer at −21 °C for a period of 1 year was possible using glycerine polyethylene glycol (PEG) 400 or 1000 as cryoprotective agents. Of these, 20% (V/V) glycerine/water solution was the most suitable, resulting in germination rates of the stored conidia of 21% after 1 year (Paul et al. 1998).

14.6 Artificial Inoculation of Excised Cotyledons

Cotyledons of radish, 9 days after sowing, were placed face downwards on a damp filter paper in a transparent plastic box (Bonnet and Blancard 1987). Then kept the box in a growth chamber at 20 °C at daytime and 18 °C at nighttime, with relative humidity of 90% and illumination of 2000 lux for 12 h. With a micropipette, 50 μl of H. parasitica conidial suspension (16,000 sp./ml) was placed on the abaxial surface of the cotyledons. After 5 days, the conidia were collected by rubbing the cotyledons with a brush into 2 ml of water; the concentration of the suspension was then measured and adjusted using a haemocytometer. When 15-day-old plants were inoculated, an excellent correlation was observed between the number of conidia and symptoms on leaves.

14.7 Propagation of Hyaloperonospora parasitica on Cotyledons or True Leaves of Japanese Radish Seedlings

Cotyledons and the true leaves of radish seedlings were subjected to hot water (50 °C) treatment, or the roots were cut off to weaken resistance to H. parasitica (Ohguchi et al. 1989). Each of the 7–11-day-old cotyledons of cvs. Awa-ichigo, Sarakamuri, and Daimaru-shogoin, which had been treated with hot water, was put 14.8 Laboratory Tests of Fungicides 315 in a test tube (2.8 × 19 cm) containing 15 ml of distilled water. The upper surfaces of these cotyledons were inoculated with drops of conidial suspension of the fungus. The percentage of conidiophore formation on the cotyledons grown for 1 week at 20 °C, 1000 lux after inoculation, was highest on the 11-day-old cotyledons from cv. Diamaru-shogoin treated with hot water for 60 s. On the true leaf of 3-week-old seedlings, the percentage was highest in cv. Sarakamuri treated with hot water for 30 s. Since the cotyledons of cv. Shirokubi-miyashige, Heian-tokinashi, and Daimaru-shogoin, which had been grown for 4–6 days in a growth chamber (25 °C; 5000 lux), were very susceptible to hot water treatment, their roots were cut off. Cut surfaces of hypocotyls were wrapped with cotton wetted with sterilized distilled water or a modified Knop solution in order to keep the cotyledons from withering. The lower surfaces of cotyledons were inoculated with the suspension of conidia. The percentage of the conidiophore formation was highest on the 6-days-old cotyledon of cv. Shirokubi-miyashige. In the case of cv. Daimaru-shogoin, the 4-day-old cotyledon was best suited. A dark treatment of 18 h of the infected cotyledons on the 6th day after inoculation stimulated conidiophore formation following synchronized formation of the conidia. Also, many conidiophores were formed on the infected cotyledon when moved into an incubator at 20 °C after being stored in a refrigerator at 5 °C for 2 weeks after the 3rd day after inoculation (Ohguchi et al. 1989).

14.8 Laboratory Tests of Fungicides

A susceptible cultivar of Brassica species must be used for the maintenance of H. parasitica (Channon and Hampson 1968). Sow seeds of susceptible cultivar in boxes. Detach cotyledons bearing 4–5 mm petiole from the seedlings, and lay them in a single layer on sterilized moist, crinkled filter paper in transparent plastic boxes. Add sufficient sterilized tap water to maintain the filter papers adequately moist. Ten to fourteen -day-old cotyledons are suitable for maintaining the cultures. Obtain conidia from an actively sporulating fungus on the leaf or cotyledon. Seedlings can be inoculated with the aid of a small paint brush or spraying or by dipping the cotyledons in the spore suspension. After inoculation, incubate the boxes of cotyledons in growth cabinet at 15 °C with illumination. Supplementary light (400 w mercury fluorescent lamps, 3/4 ft. above the boxes, and each illuminating an area of just under 11 sq. ft) for 12 h per day is essential for the survival of both infected, and uninfected cotyledons. To test the protectant action of fungicides, 10–14-day-old seedlings are cut off at soil level and placed in Weldmesh Racks (14 seedlings per rack) with cut ends of the stems immersed in water in an enamel dish which support the racks (Channon and Hampson 1968). Atomize 2 ml of the test chemical on the upper surfaces of leaves of seedlings in the racks. On the following day, cut off the leaves with petioles, and place on moist filter paper in three to four plastic boxes (3 1/3″ × 1 7/8″ × 7/8″). Inoculate these leaves with a drop (0.01 ml) of spore suspension containing approximately 1000 conidia, and incubate at 15 °C in an illuminated incubator. Record the number of leaves showing sporulation. 316 14 Techniques

14.9 Fungicide Resistance Assay

An assay for resistance and sensitivity of H. parasitica to metalaxyl can be made using cauliflower seedlings of cv. Lawyna (Crute et al. 1985). Nutrient solution (25 ml)-amended metalaxyl (Ridomil 25 WP) at a range of concentration of up to 100 μg ml−1 was contained in 7 cm diameter glass crystallizing dishes and absorbed in an equal volume of vermiculite. Seed was sown into the dishes (30–40 per dish) and placed in a temperature controlled growth room (15 °C, 12 h photoperiod, 100 μEm−2S−1). To avoid problems with vapour activity, each dish was contained within a plastic ‘treacle pot’. Seedlings at the cotyledon stage, 7–10 days after sowing, were inoculated with the conidial suspension of the fungus and incubated under the same conditions. Observations were recorded for the presence or absence of sporulation 5–10 days after inoculation. A standard metalaxyl sensitivity isolate which was completely inhibited at 0.01 μg ml−1 was included in all tests. A modification of the method using cauliflower seed treated with metalaxyl (Ridomil 25 WP) at a rate of 1 g a.i. per Kg clearly discriminated between resistant and sensitive isolates. The bioassay of the plant material revealed 15–20 μg ‘metalaxyl equiva- lents’ per g fresh weight in seedlings during the course of the test. Resistant isolates sporulated profusely on seedlings grown from treated and untreated seed while sensitive isolates only sporulated on the latter (Crute et al. 1985).

14.10 Measuring Systemic Infection of the Downy Mildew Pathogen

According to McMeekin (1971), seeds of Brassicas were first surface sterilized in sodium hypochlorite (10% Commercial Clorox) for 5 min and then placed about 1.5 cm apart on either 1–2% agar or sterilized glass wool moistened with distilled water in the bottom of a moist chamber (McMeekin 1971). They were germinated in the dark at 20 °C. Seven days later most seedlings were about 2 cm tall. The cotyledons and roots were excised from these seedlings. The hypocotyl, relatively free of starch granules and chloroplast, was placed with one end in 10 ml of test solution at the bottom of a Petri dish (5 cm diameter). Ten or more hypocotyls were kept upright in the dish by pushing them through a double layer of cheesecloth stretched over the dish and held in place by a rubber band. Inoculum was applied either on the upper tip or on the side of the hypocotyl. A 1 mm square piece of Brassicas cotyledon covered with conidiophores was used as inoculum. The Petri dish bottoms, containing the inoculated hypocotyls, were placed in a moist chamber lined with wet paper towel. The moist chamber was placed in a 15 °C incubator with a light intensity of 5 ft.-c for about a week. The whole hypocotyl was removed from the solution and fixed on a slide by 0.1% cotton or aniline blue in lactophenol (20% carbolic acid, 20% lactic acid, 40% glycerine, 20% distilled water). The hypocotyl was pressed evenly with another slide until it 14.11 Methods of Breeding for Multiple Disease Resistance 317 was flattened, and then a cover slip was applied. After a few hours, the mycelium was stained and could be seen within the host. The length of time between inoculation and fixation determined the intensity of the stain in the mycelium within the host tissue. The cotton blue stained the protoplasm of the fungus, but not the cell wall. Only the youngest fungal growth at the time of fixation was deeply stained in the final preparation. If the test solution favoured or did not interfere with host or fungal growth, the fungus could grow from the point of inoculation to the lower tip of the hypocotyl that was immersed in the test solution for 4 to 5 days at 150 C. At this time the mycelium in the lower tip would stain dark blue, but the older mycelium at the point of inoculation took little stain. If the test solution was unfavourable, a ‘zone of inhibition’ lacking fungal growth could be measured from the base of the hypocotyl to the point where fungal growth has stopped. Most of the mycelium was parallel to the stele. Solutions containing either antibiotic or sugars were tested with this method, and for a given concentration, the zone of inhibition was very consistent. It was possible to use this method without completely aseptic procedures and not have a serious problem with rotting. However, streptomycin sulphate (0.25 μg/ml) reduced the rotting without appearing to affect host or the pathogen (McMeekin 1971).

14.11 Methods of Breeding for Multiple Disease Resistance

To identify resistance to various Chinese cabbage pathogens, Williams and Leung (1981) developed methods for screening large populations of seedling plants. Screening of seedlings was preferred in the early stages of breeding programmes because it takes less time and space. Plants which exhibited seedling resistance were later evaluated for mature plant resistance. Such procedures may involve simultaneous inoculation and incubation of 1-week-old seedlings with Plasmodiophora brassicae, Hyaloperonospora parasitica, Albugo candida, Phoma lingam, and Alternaria brassicicola or A. brassicae. This can be followed by a sequential inoculation with turnip mosaic virus (TuMV) or/and Erwinia carotovora and/or Xanthomonas campestris. The interactions of phenotypes of more than one pathogen on a single host can be relied on for evaluation (Fig. 14.2) by paying special attention to the following: (a) careful preparation, quantification, and delivery of precise amounts of virulent inoculum, (b) careful cultivation of host ‘target tissues’ of known physiological age, and (c) optional incubation conditions for disease development. Single seeds were sown in 12-pack pots, and when the cotyledons have expanded, after 5–7 days, single 0.01–0.02 ml drops of a freshly collected conidial suspension containing approximately 1 × 105 conidia/ml were placed on each of the two cotyledons with the aid of a finely tipped glass pipette (Williams and Leung 1981). As plants of each 12 pack were inoculated, they were placed in glass or plastic boxes containing a 1–2 cm depth of warm water. A tight-fitting cover was placed on the box when it was filled with plants. The box was then placed in a darkened incubator 318 14 Techniques

Fig. 14.2 Location of inoculum placement of eight pathogens in multiple disease screening of seedling Chinese cabbage. Pb = Plasmodiophora brassicae, Ec = Erwinia carotovora, Ac = Albugo candida, Pl = Phoma lingam, Ab = Alternaria brassicae, Hp = Hyaloperonospora parasitica, Xc = Xanthomonas campestris, and TuMV = Turnip mosaic virus (Williams and Leung 1981)

set at 20 °C for 8–16 h. The atmosphere in the box maintained the droplets in the cotyledons during which time germination and penetration occurred. Plants were then transferred to the greenhouse bench at 20–25 °C for 5 days and then returned to the humidity boxes at 20 °C for 16–24 h. By then, the susceptible plants had a profuse growth of conidiophores on the lower sides of the cotyledons, whereas resistant plants exhibited varying degrees of sporulation and tissue necrosis which was evaluated on a 0–9 scale as illustrated in Fig. 14.1. Williams and Leung (1981) also noted that it is important to use freshly produced inoculum collected by washing off spores from leaves with distilled water and that older plants may be inoculated by atomizing the foliage with the suspension of conidia and keeping them at 100% RH for 8–16 h. Chinese cabbage was grown under the above conditions, and sequentially inoculated with four pathogens (Williams and Leung 1981). Five days after sowing, seedlings were dipped in a spore suspension of Plasmodiophora brassicae spores and transplanted and then 1–2 days later inoculated with Hyaloperonospora parasitica conidia. Twelve days after sowing, the plants were evaluated for downy mildew resistance and the susceptible plants removed. The remaining plants were then inoculated with turnip mosaic virus (TuMV) at 14 days, evaluated, reinoculated, and rouged over the following 14 days. Surviving plants were then inoculated at 21 days after sowing for soft rot resistance. Two weeks later, 35 days after sowing, resistant plants could be removed from the pots and examined for club root. Plants with- 14.11 Methods of Breeding for Multiple Disease Resistance 319

DAYS 0714 21 28 35

SEED TRANSPLANT 25°C

PLASMODIOPHORA BRASSICAE

PERONOSPORA PARASITICA 100% 100% RH 20°C RH 20°C

TuMV 15–20°C

ERWINIA CAROTOVORA

INOCULATION READ INTERACTION PHENOTYPE

Fig. 14.3 Sequence for individual and multiple disease resistance screening in Chinese cabbage (Williams and Leung 1981) standing all four diseases could then be potted, and vernalized or treated with Benlate fungicide, and transplanted to the field. Further inoculations with TuMV, Erwinia, and Hyaloperonospora could be made in the field, and plants not treated with benomyl fungicide could be planted in H. parasitica-infested field plots. The procedures for the sequential inoculations were reported to be essentially the same as those for individual inoculations except when TuMV inoculation was to be followed by E. carotovora or H. parasitica (Williams and Leung 1981). The plants were maintained at 25 °C instead of returning them to cooler temperatures for enhancement of virus symptoms. Inoculation with a combination of any of the four pathogens was possible by following the appropriate portions of the total sequence in Fig. 14.2. Plants can also be inoculated with other pathogens such as Albugo candida, Alternaria spp., Phoma lingam, and Xanthomonas campestris. Inoculations with these pathogens were kept apart from those areas of the cotyledons and leaves which were occupied by Hyaloperonospora (Fig. 14.3). It allowed large F2 and backcross progenies to be efficiently screened. It was possible to screen approximately 600 plants per m2 for the four diseases every 35 days. If resistance to each of the above pathogens were controlled by independent single recessive genes, a theoretical minimum population of 256 plants would be needed to recover the four recombinants. It is likely that far larger populations would be screened to accommodate the differing heritabilities for each form of resistance. An important consideration in selecting dominant forms of resistance in the production of F1 hybrid Chinese cabbages was that resistance to different pathogens can be introduced into the hybrid from different inbred parents. 320 14 Techniques

14.12 Heterothallism and Homothallism

For such studies, isolates of H. parasitica were collected from different host species and also from various geographical locations (Sherriff and Lucas 1989a, b). The isolates were maintained on seedlings of susceptible cultivars of respective hosts. Cotyledons with spores were excised, placed in 50 ml sterile distilled water (SDW), and shaken gently to dislodge the conidia. The conidial suspension was then filtered through three layers of cotton gauze and centrifuged at 1500 g. The conidial pellet was resuspended in SDW, centrifuged, and finally resuspended in 1–2 ml SDW. Cotyledons of 7-day-old susceptible seedlings, rose in a 9 cm pot, were drop inoculated with the conidial suspension with the aid of a Pasteur pipette. Inoculated seedlings were then sealed in a 13 × 21 cm propagator and transferred to a growth room (19 + 1 °C; 16 h d, Osram cool white fluorescent tubes, photon flux density 70μml−2 S−1). Conidia were harvested 5–7 days after inoculation. For microscopic examination, cotyledons and leaf pieces were cleared by boiling for 2 min in a lactophenol-ethanol solution containing 10 g phenol, 20 ml glycerol, 10 ml lactic acid, and 20 ml 96% (V/V) ethanol (Sherriff and Lucas 1989b). Cleared cotyledons were rinsed in water, stored in 70% (V/V) glycerol, and examined under a low-powered microscope. Mature oospores were easily visible due to their brown pigmentation; young oospores tended to take up and retain green pigments from host tissues during cleaning.

14.13 Seed-Borne Nature of H. parasitica

Jang and Safeeulla (1990a, b) studied the seed-borne nature of H. parasitica in Raphanus sativus. Four hundred seeds of test host cultivars were sown in field plots which were observed periodically for the occurrence of downy mildew disease. At the seed-setting stage, seeds from infected plants were subjected to a maceration technique (Shetty et al. 1978). Seeds were placed in 250 ml of 10% NaOH for 24, 36, and 48 h, at 22 °C along with 0.5 g of trypan blue stain. After the alkali treatment, the seeds were agitated in warm water (60–70 °C) for 5 min. Hard seeds were softened by boiling in 5% NaOH for an additional 5–10 min. Seeds were then sieved and excess water drained off, and lactophenol was added to a beaker containing the treated seeds. The lactophenol completed the detachment of the embryo from the seed coat. The beaker with the embryos and the seed coats were placed in a water bath and heated with a low flame until the embryos were cleared. The embryos and seed coats were examined under a stereomicroscope. To determine the viability of the internally borne mycelium, a seedling symptom test was carried out. Four hundred seeds from the above samples were sown under controlled conditions in a glass house which was free from airborne inoculum. Before sowing, the seeds were surface sterilized. Such seeds were sown in pots containing steam-sterilized soil. Daily observations were made following seedling emergence, and the percentage of 14.14 Conidial Germination 321 infected seedlings within each cultivar was recorded. The seeds from the first harvest were subjected to the alkali maceration technique to determine the rate of transmission of the pathogen in the seeds (Jang and Sefeeulla 1990b). To study pathogen infection through the stigma, unfertilized stigma of healthy plants were taken from test cultivars of the host (Jang and Safeeulla 1990b). Unfertilized carpels were removed from healthy plants. The ovaries along with style and stigma were placed on the sporulating surface of infected leaves at 160 C for 3 days. At 12 h intervals, such ovaries were fixed in acetic acid/alcohol (1:3) and subjected to the alkali maceration technique (Shetty et al. 1978). Another method was to spray unpollinated carpels with a conidial suspension or dipping inflorescences of healthy plants in a container with a concentrated conidial suspension. Such treated carpels were covered with moist polyethylene bags to maintain humidity for 2–3 days. The carpels are then fixed in acetic acid/alcohol. They were dehydrated by boiling in alcohol/lactophenol (50:50) for 30–35 min. Followed by maceration in 5% KOH solution for 24 h. The macerated carpels were washed in distilled water and treated with saturated chloral hydrate solution with 0.5% cotton blue for 24 h. The clear ovaries were mounted in lactophenol on slides after squash- ing and then observed microscopically (Jang and Safeeulla 1990b). Achar (1995) developed a tissue culture technique to determine the viability of seed-borne mycelium of H. parasitica in cabbage (Brassica oleracea). The percentage of seeds showing viable mycelium varied with cultivar, and a direct correlation between embryo infection and seed transmission in callus was observed. A similar correlation was established using a seedling system test. The tissue culture technique was quicker and more reliable than the seedling symptom test for determining the viability of mycelium. The possibility of adopting this technique for routine seed health testing was discussed.

14.14 Conidial Germination

To test the effect of temperature and relative humidity on conidial germination and germ tube growth, a conidial suspension was made by washing off the conidia from the donor host leaves into Petri dishes (Lin 1981). A fine stream of cold, sterilized, distilled water delivered by an atomizer was used for this purpose. The donor leaves were usually collected at 4 am, when sporulation was abundant. The conidial suspension was adjusted to the desired concentrations (5 × 103cells/ml) by dilution with distilled water and then sprayed on to 1.25% water agar in Petri dishes. After incubating the inoculated Petri dishes and kept separately at 4 °C, 8 °C, 12 °C, 16 °C, 20 °C, 24 °C, 28 °C, 32 °C, and 36 °C for 24 h, germination and germ tube growth of conidia were determined by microscopic observation of 400 spores per plate. Three replications were used for each treatment. A conidium was considered germinated if the length of the germ tube exceeded the width of the conidium. To determine the effect of humidity on germination of conidia, three drops of conidial suspension were pipetted onto a clean glass slide placed in a Petri dish containing 322 14 Techniques

saturated salt solution to obtain theoretical relative humidity of 0% (CaCl2), 32% (CaCl26H2 0), 55% (Ca (N03)2 4H2 0), 81% (NH4)2 SO4), and 95% (NAHPO4 12H2O). The Petri dishes were sealed and incubated at 16 °C for 24 h. Spore germination was then counted (Lin 1981).

14.15 Sporulation

In the evening, diseased leaves were excised from 40–50-day-old plants grown in the field (Lin 1981). Excised leaves showing fresh symptoms were cut into several 0.5 × 0.5 cm2 pieces. The pieces were first gently dipped into sterilized water to wash off the conidia borne on conidiophores. Six pieces were then placed on a slide with the abaxial surface upward. The slide was put in a Petri dish containing two filter papers previously moistened with distilled water. After incubating the Petri dishes at 4 °C, 8 °C, 12 °C, 16 °C, 20 °C, 24 °C, 28 °C, 32 °C, and 36 °C for 18 h, sporulation was determined by shaking the six pieces in 1 ml distilled water and counting the number of conidia with a haemocytometer under a microscope (Lin 1981).

14.16 Discharge of Conidia

Lin (1981) also measured the discharge of H. parasitica conidia from diseased leaves of the host. A leaf showing typical symptoms of downy mildew was selected and fixed on the hole of a spore collector so that the abaxial surface of the diseased leaf faced directly over 1 of the 24 slides attached to this collector. The surface of the slides was smeared with a layer of Vaseline to intercept the falling conidia. Each slide automatically moved forward one position per hour; thus a 24 h spore collection was obtained. The collection was continued for 3 days beginning at 9 pm. each day, and the periodic conidial discharge was determined by counting the conidia on slides under a microscope. Temperature and relative humidity for each hour during conidia collection were also recorded to establish the relationship between the discharge of conidia and environmental parameters (Lin 1981).

14.17 DNA Fingerprinting of Hyaloperonospora parasitica

RAPD analyses were carried out using genomic DNA from 16 isolates of H. parasitica (Pers. ex. Fr). These included seven field isolates from theB. napus pathotype and four field isolates from the B. oleracea pathotype (Sherriff and Lucas 1990) collected in different years from various sites in the UK. Within each pathotype, isolates were chosen on the basis of differential virulence to specific host cultivars, 14.17 DNA Fingerprinting of Hyaloperonospora parasitica 323 the oospore-derived progeny P003, and P033 segregate for virulence to the oilseed rape (B. napus var. oleifera) cv. Cresor (Lucas et al. 1988), while P005 and P006 differ in virulence to the cauliflower (B. oleracea var. cauliflora) cv. Palermo Green (Moss et al. 1991). Other variable characters were mating type and differences in sensitivity to the acylalanine fungicide metalaxyl.

14.17.1 DNA Isolation

1. Fungal isolates: Genomic DNA was extracted from conidia of 4–5-day-old H. parasitica cultivars maintained on host seedlings (B. napus cv. Mikado; B. napus cv. Cresor or B. oleracea cv. Offenham Compacta) in a growth room at a temperature of 15 ± 20 C under a 14 h photoperiod at a light intensity of 77 μEm−2S−1. Conidia were dislodged from sporulating cotyledons by washing with sterile distilled water. The conidial suspension was centrifuged and the conidial pallet washed at least three times in sterile distilled water. Clean conidia were then vortexed with a mixture of 1 and 6 mm diameter in a lysis buffer containing 100 mM TRIS HCL (pH 7.2), 100 mM EDTA, 10% (w/v) SDS, and 2% (v/v) 2-mercaptoethanol. DNA was recovered from the suspension of broken conidia using the protocol described by Lee and Taylor (1990). The measurement of DNA concentration was done using a TKO 100 dedicated mini-fluorometer. 2. From host plants: Genomic DNA was extracted from cotyledons of uninfected 9–10-day-old seedlings of maintenance hosts using the protocol described by Edwards et al. (1991). 3. Polymerase chain reaction (PCR) materials: The 10 mers used as random primers in the PCR were purchased from Operon Technologies (Alameda, California). Taq DNA polymerase together with 10 × concentration buffer was supplied by Boehringer Mannheim (FRG). Amplification was carried out in a Model 60 Tempcycler (Coy Lab Products, Ann Arbor, Michigan) and in a PHC-3 Dri-Block Thermal Cycler (Techne, Cambridge). Agarose (UltraPure) was supplied by Gibco, BRL. 4. PCR amplification conditions: The amplification conditions were rigorously tested. A standard procedure was determined based on the protocol of Williams et al. (1990). The PCR volume was 25 μl and contained 0.2 μM of primer; 100 μM each of dATP, dCTP, dGTP, and dTTP 25 ng template DNA; and 0.5 U of Taq polymerase in 1 × PCR buffer (10 mM TRIS-HC1 pH 8.3, 1.5 mM MgC12, 50 mM KC1, 0.1 mg/ml gelatine). During manipulations, the tubes were kept on ice. The reaction mixtures were overlaid with 25 μl mineral oil. Standard amplifications were performed in a Coy Model 60 Tempcycler programmed for 45 cycles of 1 min at 94 °C, 1 min at 36 °C, and 2 min at 72 °C or in a Techne PHC-3 Dri-Block Thermal Cycler programmed for 45 cycles of 30s at 93 °C with ramp time of 30 °C/min, 40 s at 37 °C with a ramp time of 30 °C/min, and 1 min 20 s at 72 °C with a ramp time of 20 °C/min. When programmed as above, both machines produced similar temperature profiles and amplification products. 324 14 Techniques

After the last cycle, the samples were kept at 72 °C for an additional 10 min and then cooled to 4 °C. Samples of 15 μl were analysed by electrophoresis in a 1.5% (w/v) agarose gel containing 0.2 μg/ml ethidium bromide with 0.5 × TBE as buffer. 5. Hybridization conditions: Selected amplification products obtained with template genomic DNA from isolate P003 were recovered from the gel and purified using GENECLEAN II. Random-primed DNA labelling was carried out using dCTP32. These fragments were used as probes in hybridization experiments with genomic DNA digested with Eco RI and Hind III. Genomic DNA (0.1 μg) was digested with 2 μl of restriction enzyme, separated by electrophoresis, and transferred to Gene Screen Plus membrane, according to the manufacturer’s instructions. The filter was pre-hybridized for 4 h at 42 °C in a buffer containing 50% (v/v) formamide, 5 × Denhardt’s solution (Denhardt 1966), 3 × SSPE, and 0.5% (w/v) SDS. Hybridization was carried out overnight under the same conditions. The filter was washed once in 3 × SSC with 0.1% (w/v) SDS, followed by another wash in 2 × SSC containing 0.1% (w/v) SDS, each for 20 min at 65 °C. A final rinse in 2 × SSC was carried out at room temperature, and the filter was exposed to X-ray film (Tham et al. 1994).

14.18 Molecular Marker for Identification of H. parasitica

PCR amplification of ribosomal RNA (rRNA) gene block spacers (ITS1 and ITS2) performed in 44 H. parasitica isolates from different Brassica oleracea cultivars and distinct geographic origins revealed no length polymorphisms. ITS restriction analysis with three endonucleases, confirmed by sequencing, showed no fragment length polymorphisms among isolates. Furthermore, ITS amplification with DNA isolated from infected host tissues also allowed the detection of the fungus in incompatible interactions. The combination of the universal ITS4 and ITS5 primers, for amplification of full ITS, with a new specific forward internal primer for ITS2 (Pp ITS 2F), originates a H. parasitica-specific amplicon, suitable for diagnosis. As ITS2 regions of H. parasitica, B. oleracea, other B. oleracea fungal pathogens, and other Hyaloperonospora species are clearly distinct, a fast, and reliable molecular identification method based on multiplex PCR amplification of full ITS and H. parasitica ITS2 is useful for the diagnosis of crucifer downy mildew. The method can be applied to diagnose the disease in the absence of fungal reproductive structures, thus being useful to detect non-sporulating interactions, early stages of infection on seedlings, and infected young leaves packed in sealed plastic bags. Screening of seed stocks in sanitary control is also a major application of this diagnostic method (Casimiro et al. 2004). In fungal genomes, the highly conserved rRNA genes are separated by two less conserved internal transcribed regions, the internal transcribed spacers 1 and 2 (ITS1 and ITS2), which are therefore suitable for polymorphism studies among species or even at infraspecific level (Duncan et al. 1998; Mills et al. 1998). Amplified 14.18 Molecular Marker for Identification of H. parasitica 325 ribosomal DNA restriction analysis (ARDRA), using PCR primers based on conserved regions of the rRNA genes (White et al. 1990), followed by restriction with frequently cutting endonucleases, allows the easy assessment of sequence differences in ITS regions without length polymorphisms (Buscot et al. 1996; Lanfranco et al. 1998). Although fungal ITS1 has been shown to be more polymorphic at sequence level than ITS2 (Duncan et al. 1998), analysis of H. parasitica ITS1 sequences showed no differences among isolates collected from the same host species (B. oleracea and A. thaliana) and only 85% similarity between isolates from different hosts (Rehmany et al. 2000). In fact, host range must be associated with genetic differences of the isolates, classified as belonging to the same species, and these data point to the potential of ITS regions as molecular markers, both at species and forma specialis levels. There are, however, no available data on ITS2 variability and taxonomic relevance. Casimiro et al. (2004) characterize the ITS regions of H. parasitica isolates, from different B. oleracea crops and distinct geographic origins, in order to evaluate their potential as molecular markers for identification purposes. 1. Hyaloperonospora parasitica isolates and conidia isolation: Forty-four H. parasitica isolates, collected from B. oleracea plants, were grown on seedlings of B. oleracea hosts ‘1’ or ‘2’. For each isolate, ca. 50 1-week-old seedlings were inoculated with two droplets of a conidia suspension (5 × 104 conidia ml−1) per cotyledon and maintained in the dark, at 16 °C for 24 h. Then the inoculated seedlings were transferred to a growth room and maintained at 20 ± 1 °C, under a 20 h photoperiod. After 6 days of incubation, the seedlings were transferred to a dark room for 24 h, to induce sporulation. Cotyledons with sporulation were harvested and shaken in 50 ml of sterile distilled water to dislodge conidia. The conidial suspension was gauze filtered and centrifuged at 2600 g for 3 min. The pellet was resuspended in 7.5 ml of sterile distilled water, aliquot in 1.5 ml fractions, and stored at −200 C until use. 2. B. oleracea fungal pathogens selection and growth: Ten isolates of B. oleracea fungal pathogenic species or related species of the same genera were selected, namely, Fusarium culmorum, Trichoderma sp., Alternaria sp., Phoma sp., Phytophthora cinnamomi, and Sordaria sp., from our collection, and from CECT (Collection Espanola de Cultivos Tipo), F. oxysporum (CECT 2154), Sclerotinia sclerotiorum (CECT 2882), Mycosphaerella tassiana (CECT 2665), and Diaporthe phaseolorum (CECT 2022). All fungi were grown on potato dextrose agar medium, with exception of Phytophthora cinnamomi which was grown on corn meal agar, at 280 C for 7 days. 3. DNA isolation: DNA was isolated using an adaptation of the Ferreira and Grattapaglia (1995) method. An aliquot of each H. parasitica conidial suspension was centrifuged at 6400 g for 3 min. The pellet or 100 mg of each fungal mycelium (obtained by colony scraping) was macerated with 200 μl of glass beads (425–600 microns), and 500 μl of extraction buffer (CTAB 2%, 1.4 mol l−1 NaCl, 0.02 mol l–l EDTA, 0.01 mol l−1, Tris–HCl pH 8.0, 1% PVP, 0.2% β-mercaptoethanol, 0.1% Proteinase K), at 65 °C, was added. The suspension 326 14 Techniques

was incubated at 65 °C for 45 min, with mixing by inversion each 15 min. After cooling to room temperature, 500 μl of chloroform/isoamyl alcohol (24:1) was added; the tube was mixed by inversion and centrifuged at 16700 g for 10 min. The upper aqueous phase was collected, and the DNA was precipitated with 600 μl of isopropanol (−20 °C) for 1 h at −70 °C. After a 10 min centrifugation at 16,700 g, the pellet was washed with 500 μl of washing buffer (ethanol 70%, 0.15 mol l−1 NaCl) and centrifuged at 16,700 g for 5 min. The pellet was resuspended in 25 μl of TE (0.01 mol l−1 Tris–HCl, pH 8.0, 0.001 mol l−1 EDTA) and stored at 4 °C until utilization. After maceration with liquid nitrogen and using the method above, DNA was extracted from 100 mg of short cycle B. oleracea CrGC3.1 and cabbage ‘Coracao de boi’ seedling tissue, which were not infected with H. parasitica, and from 100 mg of Tronchuda cabbage ‘Algarvia’ seedling tissue, either infected with the isolate P501 or uninfected. 4. ITS amplification: To amplify ITS1, ITS2, and full ITS regions, either from the fungus or the hosts, the following primers were used (White et al. 1990): ITS2 and ITS5 to amplify ITS1 region, ITS3 and ITS4 to amplify ITS2 region, and ITS4 and ITS5 to amplify full ITS. Each reaction mixture contained 2 μl DNA, −1 −1 PCR buffer 1X, 0.0025 mol l MgCl2, 0.05% W1, 0.0002 mol l of each dNTP, 0.001 mol l−1 of each primer, and 2 μl of Taq DNA polymerase, in a final volume of 50 μl. To each PCR tube, ca. 50 μl of mineral oil was added, and amplification occurred in a RoboCycler 96, according to the following amplification programme: 4 min at 95 °C; 35 cycles of 1 min at 95 °C, 1 min at 56 °C, and 2 min at 72 °C; 4 min at 72 °C. Each reaction sample was run on a 1.5% agarose gel, in 0.5 X TBE (0.05 mol l−1 Tris, 0.045 mol l−1 boric acid, 0.001 mol l−1 EDTA) at 90 V for 2 h 30 min, using 1 kb plus standard as molecular size marker. After ethidium bromide staining, the gels were analysed with KODAK 1D 2.0 software. For each isolate or host ITS regions, amplification was performed two to three times in order to assess the reproducibility of the method. The molecular sizes of H. parasitica, ITS regions were estimated using the arithmetic average, and standard error of the 44 isolates. Molecular sizes of host ITS regions were calculated as the average value of two replicates. 5. ITS restriction assay: To perform restriction digestion of amplified ITS regions, 10 μl samples of each PCR product, not purified, were digested with 5 μl of each one of three restriction endonucleases, RsaI, Hae III, and Sau 3AI, in a final volume of 15 μl, according to manufacturer instructions. After a 3 h incubation period at 37 °C, 1.5 μl of bromophenol blue solution (0.25% bromophenol blue, 0.25% xylene cyanol, 15% Ficoll in water) was added to each sample to stop the reaction. Each reaction sample was run on a 1.5% agarose gel, in 0.5 X TBE at 90 V for 3 h, using a 100 bp standard as a molecular size marker. After ethidium bromide staining, the gels were analysed with KODAK 1D 2.0 software. 14.18 Molecular Marker for Identification of H. parasitica 327

Reproducibility of the method was assessed with duplicate reactions. Molecular sizes of individual restriction fragments produced from H. parasitica, ITS regions were estimated using the arithmetic average and standard error of the 44 isolates. For host ITS regions, molecular size estimations were based on two replications.

6. ITS2 sequencing: In order to sequence the ITS2 region, 15 μl of the PCR reaction of isolate P524 was run in 1% agarose gel, in 0.5 X TBE at 90 V for 1 h 30 min. After ethidium bromide staining, the ITS2 band was extracted with a sterile scalpel and purified with the Concert Rapid Gel Extraction Systems. The purified product was cloned using the pGEM-T Easy Vector Systems, with the following adaptations: 3 μl of the purified PCR product in the ligation reaction; JM109 competent cells, after inoculation in TSS medium (1 X LB (1% tryptone, 0.5% yeast extract, 0.5% NaCl, pH 7.0), 10% PEG 6000, 5% DMSO, 0.05 mol l−1 MgSO4, pH 6.5); SOC medium replaced by LB medium in JM 109 transformation. The recombinant cells were plated in LB medium with 0.15 g l−1 ampicillin, 0.04 g l−1 IPTG, and 0.04 g l−1 XGAL. Screening of recombinant white colonies was performed after an overnight incubation of each colony in 2 ml of LB medium with 0.15 g l−1 ampicillin. Each cell suspension was centrifuged at 18,000 g for 1 min, and the pellet was resuspended in 150 μl TEG (0.05 mol l−1 glucose, 0.025 mol l−1 Tris-HCl, 0.01 mol l−1 EDTA, pH 8.0), followed by the addition of 200 μl 0.2 mol l−1 NaOH, 1% SDS. The suspension was mixed by inversion, and chilled on ice, 200 μl 3 mol l−1 potassium acetate (pH 4.8) were added, and the suspension was centrifuged at 18000 g for 10 min. To the collected upper phase, 500 μl of isopropanol (−20 °C) was added. After a 30 min centrifugation at 18,000 g, the pellet was washed with 500 μl of 70% ethanol and centrifuged at 18,000 g for 5 min. The final pellet was resuspended in 50 μl TE with RNase (0.05 g l−1). Restriction analysis of putative recombinants with the endonuclease Pvu II (Biolabs) occurred for 2 h at 37 °C, according to the manufacturer instructions, in a final volume of 30 μl. Restriction products were resolved by electrophoresis in a 1% agarose gel, in 0.5 X TBE at 90 V for 1 h 30 min. The gel was stained with ethidium bromide, and fragment molecular size was estimated with KODAK 1D 2.0 software. Recombinant colonies containing the insert were reinoculated in LB medium with 0.15 g l−1 ampicillin and incubated overnight at 37 °C. Recombinant plasmid DNA was extracted with the Concert High Purity Plasmid Miniprep System. Sequencing was performed using the CEQ 2000 Dye Terminator Cycle Sequencing Kit and a capillary electrophoresis CEQ 2000-XL sequencer, both directly from purified PCR product and from the cloned fragment. Both DNA strands were sequenced, with the primers T7 and SP6. BLASTN (Altschul et al. 1997) of the two sequences was performed in the GenBank database. 7. Internal primer design and multiplex PCR: ITS sequences of Hyaloperonospora (26 from H. parasitica and 15 from other Peronospora spp.), Albugo (5), Botrytis (2), Alternaria (3), Leptosphaeria (2), Plasmodiophora (4), Fusarium (3), Cladosporium (2), Trichoderma (1), Phoma (1), Diaporthe (1), Phytophthora 328 14 Techniques

(1), Sclerotinia (1), Mycosphaerella (2), and B. oleracea (4) available in the GenBank database (http://www.ncbi.nlm.nih.gov) were aligned with hierarchical clustering (Corpet 1988) at the INRA website (http://prodes.toulouse.fr/multialign/multialign.html). Based on this alignment, internal primers were designed for specific amplification of full ITS and ITS2 regions ofH. parasitica: PpITS1F (5′-CAAYTWTAATTGGGGG TCGTGATCTT-3′), PpITS2F (5′-AAGCGTGACG ATACTAATTTG-3′), and Pp ITS2R (5′-TGAAGTG CGGCCGAAGCTT-3′. Three multiplex PCR amplifications were performed using the following combinations of primers: ITS3 and ITS4 (to amplify any ITS2 region) plus PpITS2F and PpITS2R (to specifically amplify H. parasitica ITS2 region); ITS5 and ITS4 (to amplify any full ITS region) and PpITS1F and PpITS2R (to specifically amplify H. parasitica full ITS region); and ITS5 and ITS4 plus PpITS2F to amplify any full ITS region and H. parasitica-specific ITS2 region. Selectivity of internal primers was tested with samples corresponding to uninfected ‘Algarvia’ cabbage, the same host infected with H. parasitica, the infected host DNA combined with Alternaria sp., and Phytophthora cinnamomi DNA; H. parasitica DNA free from host DNA contamination, and each one of the 10 B. oleracea fungal pathogens. Each reaction mixture contained 2 μl DNA, PCR buffer 1X, 0.0025 mol l−1 −1 −1 MgCl2, 0.05% W1, 0.0004 mol l of each dNTP, 0.001 mol l of each primer, and 2 U of Taq DNA polymerase, in a final volume of 50 μl. To each PCR tube, ca. 50 μl of mineral oil was added, and amplification occurred in a RoboCycler 96 (Stratagene), according to the following amplification programme: 4 min at 95 °C; 35 cycles of 1 min at 95 °C, 1 min at 54 °C, and 2 min at 72 °C; 4 min at 72 °C. Each reaction sample was run on a 1.5% agarose gel, in 0.5 X TBE (0.05 mol l−1 Tris, 0.045 mol l−1 boric acid, 0.001 mol l−1 EDTA) at 90 V for 2 h 30 min, using 1 kb plus standard as molecular size marker. After ethidium bromide staining, the gels were analysed with KODAK 1D 2.0 software. For each sample, amplification was performed two to three times in order to assess the reproducibility of the method. Molecular sizes of ITS regions were calculated as the average value of two replications.

14.19 Leaf Disc Test to Assess Resistance

To study the correlation between the interaction phenotypes obtained with the leaf disc test and in the classical cotyledon test, leaf discs are removed from sufficiently developed leaves (at least 100 cm2). Different conditions are tested; they are either disinfected in a 3% calcium hypochlorite solution or simply rinsed in sterile distilled water and then placed in agar plates (14 cm diameter) with the upper side on the agar surface. The agar medium is completed with nutrients (MS, Murashige and 14.20 Use of Rooted Leaves for Screening Brassica Germplasm 329

Skoog 1962) or no nutrient but contained either 50 or 100 ppm of benzimidazole. Five droplets of 20 μl of a spore suspension (5 × 104spores ml−1) are deposited on each leaf disc. Working with five droplets is a way to spread the isolate on the disc (a spraying process, as used for infecting whole plants, is not possible on leaf disc placed in Petri dishes). After inoculation, Petri dishes are sealed and kept in a growth chamber in the dark for at least 8 h, then for 7 days under a 12 h photoperiod (16 °C at nighttime and 20 °C at daytime, 30 μE light intensity). At least six leaf discs per couple of accession/isolate are tested at least twice for each treatment. In another experiment on the effect of the leaf position on the disc resistance/susceptibility response, each leaf from the bottom (number 1) to the top of the plant is tested (the highest number is 17). These 17 leaf positions are obtained on 10–18-week-old plants, in order to cut leaf disc from sufficiently developed leaves (at least 100 cm2). Ten discs per leaf position are tested for each genotype/isolate combination (Agnola et al. 2003).

14.20 Use of Rooted Leaves for Screening Brassica Germplasm

Fully developed leaves from the middle part of 20–30 days were cut off tightly to the stem, and these whole petioles were immediately immersed into a solution of 10 mg/l of indole butyric acid (IBA) and 5 mg/l of nicotinic acid (NOA) for 20 h. Then the leaves were thoroughly rinsed under tap water and transplanted into moist perlite in 250 ml plastic beakers closed for the following week by another transparent plate cover. Perlite was moistened with tap water, but the leaf blades were weekly sprayed with diluted foliar fertilizer 0.3 g/l. Beakers were incubated in growth chamber (22 °C/18 °C at daytime/nighttime, 16 h day length) under batteries of fluorescent tubes, generating 140 μmol/m2 of irradiance at the plant level. When rooting young cotyledons, the same procedure was used except the cotyledons with petioles were harvested from 10–11-day-old seedlings. Hyaloperonospora isolate OLI from kohlrabi (Brassica oleracea var. gongylodes L.) was propagated on young detached cotyledons of cabbage cv. Kodanske on moist filter paper in plastic Petri dishes under fluorescent tubes at 16 °C. After 7–10 days, the cotyledons covered with sporulating mycelia were frozen and maintained at −80 °C. Water suspensions of conidia from this stock were used for the preparation of inocula diluted to 5 × 10−4 to 5 x 10−5 spores per ml. Surfaces of leaf discs’ rooted leaves or cotyledons were drop-inoculated (7 μl droplets). Closed plastic containers with plant organs growing in perlite were incubated for the next 2 days in the dark at 10 °C and thereafter under fluorescence tubes at 16 °C/10 °C (day/night, day length 16 h) (Havranek et al. 2005). 330 14 Techniques

14.21 Artificial Inoculation Technique Under Growth Chamber (Williams 1985)

Inoculation: Place a 10 μl drop of conidial suspension on each ½ cotyledon at 24–48 h after expansion using micropipette. Prevent droplets from evaporating. Gently agitate inoculum to keep conidia from settling (Fig. 14.4). Incubation: Infectional environment has to be maintained on inocula droplets on cotyledon 12–18 h, 16 °C, 100% RH. Place plants in dew chamber in the dark (Fig. 14.5). The incubation environment needs to be maintained as in case of propagation of the plants. Remove plants from dew chamber, and grow them for 5 days. Again keep the plants to dew chamber for 6 days post-inoculation, and maintain same condition as infectional environment, 16 °C, 100% RH for 24 h. Evaluation: Observe interaction phenotype on 7 days of inoculation. Interaction phenotype (IP) rates for both symptoms range from a hypersensitive necrotic flecking at the site of inocula droplet to irregular narrow streaks of internal necrosis associated with more or less chlorosis. Signs of the pathogen range from no sporulation to very sparse, through increasingly heavy sporulation on both the top and bottom of the cotyledons. Most highly compatible IPs have little or no necrosis, none to light sporulation on the upper, adaxial, surface of cotyledon with heavy sporulation on the lower surface (Fig. 14.1).

Fig. 14.4 Inoculation of leaves with conidial suspension

Fig. 14.5 Inoculated leaves in the dew chamber 14.21 Artificial Inoculation Technique Under Growth Chamber (Williams… 331

Scale: Plants scored 0–9 at 7 days PI after having been 160 C, 100% RH, for 24 h. 0= No infection. 1= Very minute to larger scattered necrotic flecks under inoculum drop, none to small amount of necrosis on lower cotyledon surface, no sporulation. 3= Very sparse sporulation, one to few conidiophores on upper or lower surface. Necrotic flecking often present. Tissue necrosis present. 5= Sparse scattered sporulation on either or both cotyledon surface, tissue necrosis. 7= Abundant to heavy sporulation mainly on lower surface, light to scattered sporulation on upper surface, tissue necrosis, and chlorosis may be present. 9= Heavy sporulation on lower surface, none to light sporulation on upper surface slight to no tissue necrosis. Chlorosis may be present. Disease index

i=0 DI =−∑()ij 9 n n = total plants, i = IP class, j = number of plants/class. Selection: Susceptible controls, IP = 7–9 by 7 days, PI = partial resistance, IP = 3–5, resistant plants, IP = 0–1. Host recovery: Selected plants may be grown on in the multipots until requiring transplanting or vernalized and flowered. Cotyledons of susceptible and partially resistant plants should be removed to prevent the spread of inoculum and movement of the fungus, systemically into the plants. Plants may be rid of H. parasitica by spraying or dropping Ridomil systemic fungicide, 0.1 g/lt on infection (Fig. 14.6). Resistance: A wide range of host-specific variation existing in H. parasitica appears to be adaptable to numerous hosts other than on which it occurs naturally (Dickinson and Greenhalgh 1977). 1. B. oleracea cc: Little effective R exists; monogenic dominant race-specific R is in cc. I P.I. 189,029, and cc. c P.I. 245,015 (Natti et al. 1967), cc. c cv. ‘January king’, and wild cc contain resistance (Greenhalgh and Dickinson 1975). 2. Resistance in cotyledons does not ensure resistance in other parts. Sources in cc. i. R. l. Gabrielson, CrGC. 3. aa: Oligogenic R exists in aa. p (Leung 1981) source Cr GC.

Fig. 14.6 Plants inoculation technique with fungicides 332 14 Techniques

Fig. 14.7 Multiple inoculation methods

4. Race-specific resistance can be found in a number of open pollinated crucifers. 5. There is a great need to identify race nonspecific resistance to H. parasitica. Multiple pathogen inoculation: Plants may be inoculated on one half of a cotyledon with one race of H. parasitica and on other halves with other races. Mark initial inoculation with needle puncture on margin of cotyledon, then proceed clockwise with other races or additional pathogens. Plants may be inoculated first on roots with FOC, PB, or AR, then on cotyledons with HP, AC, or LM. Multiple resistant survivors may later be inoculated with TUM, XCC, EC, and EP. AC and HP inoculum may be mixed, then applied. Interactions among host responses may occur depending on host and pathogen genotypes. For dual HP and AC inoculations, infectional incubation temperature should be 16 °C (Figs. 14.7 and 14.8).

14.22 Microscopic Studies

The light microscopic and electron microscopic techniques have been described by Koch and Slusarenko (1990) using Arabidopsis and downy mildew pathosystem. 1. Cultivation of plants: Arabidopsis was grown in 12-cm-diameter plastic pots in potting compost covered with a layer of fine vermiculite. After watering the soil, seeds were sown densely enough onto the vermiculite to give a lawn of plants. Pots were then covered with Saran wrap and kept up to 4 weeks at 4% until required. Plants were grown in a glasshouse at 23 ± 3 °C. Additional lighting (16 h) was supplied by a high-pressure sodium lamp. In this environment, germination of seeds occurred within 3–4 days, after which the Saran Wrap was removed from the pots. 2. lnoculation of plants: Plants were inoculated 2–3 weeks after germination when four to five true leaves were present. For the initial inoculation, tissue that had been stored at −20 °C for 5 months was thawed at room temperature, and a few millilitres of water was added. After vortexing, the resulting suspension was passed through two layers of cheesecloth and sprayed with a chromatographic sprayer onto the plants. Plants were incubated overnight at 20 °C in a moist chamber and returned to the glasshouse for disease development. Five days after 14.22 Microscopic Studies 333

DAYS

0714 21 28 35 SEED TRANSPLANT 240C

XANTHOMONAS CAMPESTRIS

FUSARIUM OXYSPORUM

APHANOMYCES RAPHANI

ALBUGO CANDIDA 20°C

LEPTOSPHAERIA MACULANS

ALTERNARIA BRASSICICOLA 24°C 24°C

RHIZOCTONIA SOLANI

ERWINIA CAROTOVORA

PLASMODIOPHORA BRASSICAE

TuMV

PERONOSPORA PARASITICA 16°C 16°C

INOCULATION READ GREENHOUSE 100% RH INTERACTION OR GROWTH INCUBATOR PHENOTYPE ROOM

Fig. 14.8 Time course and environmental regime for multiple disease resistance screening of crucifer seedlings 334 14 Techniques

inoculation, plants were again incubated overnight in the moist chamber to promote sporulation of the fungus. After the fungal culture was established, an easier method of inoculation was employed. After moist incubation, plants bearing conidiophores were simply rubbed against uninoculated plants, thus depositing spores onto the latter. The freshly inoculated plants were again incubated in the moist chamber. This method was convenient for routine maintenance of pathogen stocks and resulted in high infection densities that were particularly useful for the histological studies of the infection process. 3. Light microscopy: Infection and development of the fungus were studied in whole leaf mounts stained with lactophenol-trypan blue (1.0 ml of lactic acid, 10 ml of glycerol, 10 g of phenol, 10 mg of trypan blue, dissolved in 10 ml of distilled water) (Keogh et al. 1980). Whole leaves were boiled for approximately 1 min in the stain solution and then decolorized in chloral hydrate (2.5 g of chloral hydrate dissolved in 1 ml of distilled water) for at least 30 min. They were mounted in chloral hydrate and viewed under a compound microscope equipped with interference or phase-contrast optics. 4. Scanning electron microscopy (SEM): lnfected leaves were fixed in the vapour of a 4% (w/v) aqueous solution of osmium tetroxide for 3 h at room temperature, dehydrated in a graded series of acetone, and dried at critical point. After mount- ing on specimen stubs with conductive silver print paint and sputter-coating with an 80%:20% alloy of gold and palladium, the samples were examined in a Cambridge S4 Stereoscan Electron Microscope.

14.23 Light and Transmission Electron Microscopy (TEM)

1. Plant and pathogen: Arabidopsis accessions of wild-type Oy-0 and mutant Ws- eds1, susceptible to H. parasitica isolate Emoy-2, were obtained. Ws-eds1, which is a mutant accession of Ws-0, has been characterized to enhance disease susceptibility of H. parasitica isolates (Parker et al. 1996). Isolate Emoy-2 was derived from single oospore obtained from infected wild populations of Arabidopsis and maintained as distinct genotype by using the asexual conidia to reinfect 4–6-week-old susceptible Arabidopsis accession (Holub et al. 1994). Inoculated plants were placed in a clear plastic box to retain sufficient relative humidity during the experiment. Plants were then transferred to growth rooms at 18–20 °C with 16 h photoperiod (400 μm photons/m2/s). Seeds were sown on the surface of a soil mix consisting of four parts commercial peat compost contain-

ing macro nutrients (Levington F2 mix), one part vermiculite, and one part fine sand. Seven to 10-day-old seedlings with fully expanded cotyledons were inoculated by placing a drop of inoculum (2–4 μl) of H. parasitica (1 × 104 spores/ml) on each cotyledon using a Pasteur pipette. The plants were kept under a micro- propagator, to retain humidity, and the temperature was maintained between 15 and 17 °C. 14.24 C DNA-AFLP Analysis to Reveal Gene Expression 335

2. Light microscopy: Light microscopic examination of infected cotyledon was carried out on a differential interference contrast (DIC) microscope equipped with epifluorescence optics. Infection and development of the pathogen were studied in cotyledon leaves which were cleared in methanol/chloral hydrate and mounted in 50% glycerol as described previously (Shipton and Brown 1962). Aqueous basic aniline blue staining was applied to clear infected cotyledon to reveal sites of callose deposition (O’Brien and McCully 1981) and viewed with epifluorescence microscopy. Callose was detected by bright-pale-blue fluorescence under UV radiation (2A, exciter filter EX 330–380, dichroic mirror DM 400, and barrier filter BA 420). 3. Transmission electron microscopy (TEM): Samples for TEM were excised from inoculated cotyledon leaves of both accessions 1–3, 5, or 7 days after inoculation (dai) and fixed immediately in fixative solution of 3% glutaraldehyde in phosphate buffer (50 mm, pH 7) overnight at 4° C. The samples were then washed in the same buffer, twice each for 10 min, and post-fixed in 1% osmium

tetroxide (OsO4) in 50 mm phosphate buffer at pH 7 for 2 h at room temperature. Following the action of osmium tetroxide fixation and then two 10 min washes in phosphate buffer, samples are to be dehydrated with 10 min stages in a graded series of increasing acetone concentrations (50, 70, 80, and 90, v/v) followed by three changes of 100% dry acetone, each for 30 min. Dehydrated samples were subsequently embedded in Epon araldite mixture. All sectioning is carried out on a Reichert Ultracut E microtome using glass or diamond knives. The resin blocks are trimmed with a razor, and sections 2 μm thick were taken from the tissue blocks with a glass knife. Sections were mounted on glass microscope slides and stained with 1% toluidine blue in 1% borax. This allowed the selection of interesting areas suitable for the preparation of ultra-thin sections. After location of infection sites, ultra-thin sections (80–90 nm) were cut using a diamond knife. The sections were then routinely mounted for staining on uncoated 300 mesh copper grids or Formvar-coated 200-mesh copper grids. Grid-mounted sections with silver-gold interference colour were stained with uranyl acetate and lead citrate, the method being a minor modification of the procedure as described by Roland and Vian (1991). All sections are viewed using Hitachi H7000 1TEM at an accelerating voltage of 75 kV (Soylu and Soylu 2003).

14.24 C DNA-AFLP Analysis to Reveal Gene Expression

1. Plant material, inoculums, and pathogen infection: Two non-heading Chinese cabbage inbred lines, ‘Suzhou Qing’ (resistant to H. parasitica) and ‘Aijiao Huang’ (susceptible to H. parasitica), were used in this study. The transcript- derived fragments (TDFs) were obtained from interaction between ‘Suzhou Qing’ and downy mildew (H. parasitica). ‘Aijiao Huang’ was used for the comparison of expression patterns of the four genes related with fungal resistance between resistant and susceptible line. Plants were grown in plastic nurseries 336 14 Techniques

(inner size, 45 × 45 mm; height, 57 mm) and transferred to a growth chamber under 25 °C day/20 °C night temperature with 85 ± 5% relative humidity and a 12 h light/12 h dark after germination for 36 h under dark. H. parasitica was isolated from leaves of susceptible line ‘Aijiao Huang’. Conidial suspensions were adjusted to 1 × 105 spores per ml, and Tween-20 was added as a surfactant to a final concentration of 0.1%. One hundred of 3-week-old seedlings (with four true leaves) were sprayed with 50 ml pathogen suspension and distilled water (as control), respectively. After inoculation, the seedlings were covered with plastic film separately and transferred to a growth chamber under 20 °C, 100% relative humidity in the dark for the first 24 h to promote sporulation and then moved back to the initiatory conditions. Both control and treated third leaf of five plants were harvested and pooled at 0, 24, 48, and 72 h post-inoculation (h.p.i.), immediately frozen in liquid nitrogen and stored at −70 °C until use. 2. RNA isolation and cDNA-AFLP analysis: Total RNAs were extracted using the RNeasy Plant Mini Kit (Qiagen; https://www.qiagen.com/cn/shop/sample- technologies/rna/rna-preparation/rneasy-mini-kit # ordering information), and synthesis of the first strand of cDNA is made using the M-MLV reverse transcrip- tase according to the manufacturer’s protocol. To synthesize the second strand, the following components were added to the first-strand solution: a volume of 30 μl, 5 × 2nd strand synthesis buffer, 3 μl dNTP mixture, 89 μl RNase-free H2O, 2 μl Escherichia coli DNA polymerase I, 2 μl E. coli RNase H/E. coli DNA ligase mixture, and 4 μl T4 DNA polymerase in a final volume of 150 μl. The components were gently mixed and incubated at 16 °C for 2 h. Double-stranded cDNA was purified using the DNA Fragment Purification Kit Ver.2.0. Second-strand cDNA was digested by two restriction enzymes Tag I (restriction site TCGA) and Ase I (restriction site ATTAAT) and then ligated to Tag I and Ase I double-strand adaptors. The AFLP adaptor primers 5′-GACGATGAGTCCTGAC-3′ and 5′-CGGTCAGGACTCAT-3′ (Taq I-adaptor primers) and 5′-GCGTAGACTGCGTACC-3′ and 5′-TAGGTACGCAGTC-3′ (Ase I-adaptor primers) were ligated onto the restriction fragments: Taq I preamplification primer, 5′-GACGATGAGTCCTGACCGA-3′; Ase I preamplification primer, 5-CTCGTAGACTGCGTA-CCTAAT-3′; Taq I selective amplification primer, 5′-GATGAGTCCAGACCGA + NN-3′; Ase I selective amplification primer, 5′-GACTGCGTACCTAAT + NN-3′ (indicated by N, representing an A, C, G or T). The initial small-scale screen using 96 AFLP primer combinations was done using six Taq I forward selective amplification primers (extension CG, CA, CT, CC, GA, or GT) in combination with 16 Ase I reverse selective amplification primers (extension NN), respectively. Preamplification PCR was carried out with one tenth volume of the restriction/ligation mix; the preamplification PCR was carried out as follows: 94 °C, 3 min; 94 °C, 30 s, 55 °C, 30 s, 72 °C, 60 s, 25 cycles; and 72 °C, 5 min. The products of preamplification was diluted ten- fold, and the selective amplification PCR was carried out as follows: 94 °C, 30 s; 94 °C, 30 s, 65 °C, 30 s (−0.7 °C per cycle), 72 °C, 60 s, 12 cycles; 94 °C, 30 s, 56 °C, 30 s, 72 °C, 60 s, 24 cycles; and 72 °C, 5 min (Xiao et al. 2016). 14.25 Thawing and Revival of Inoculum 337

Selective amplification products were separated on 6% polyacrylamide gels run- ning at 60 W for 2 h and visualized by silver staining. Differential bands were excised from the polyacrylamide gel electrophoresis gels based on the alignment between films and markers on the gels and incubated in 30 μL of water and then at 95 °C for 30 min. The TDFs were then re-amplified by PCR using same primers under the similar conditions. The amplified fragments were retrieved from a 1% agarose gel with the Sephaglas Band Prep Kit, cloned into pGEM-T Easy Vector according to the manufacturer’s protocol, and sequenced by Invitrogen Company (http://en.cellfood.com.cn/culture.aspx), and sequence information was BLASTed in the Brassica database (http://brassicadb.org/brad/). 3. Quantitative real-time PCR: The single-strand cDNA of resistant line ‘Suzhou Qing’ and susceptible line ‘Aijiao Huang’ were diluted to 30 ng μl− 1 and were used for quantitative reverse transcription PCR (qRT-PCR) analyses. Primers were designed by Primer 3 (http://frodo.wi.mit.edu/primer3/) based on the inter- ested cDNA sequence. The qRT-PCR reaction mixtures contained 12.5 μl, 2 × SYBR Green PCR Master Mix (Applied Biosystems; http://www.bio-rad. com/), 10 pM of each primer, 2 μl template, and sterile distilled water to total volume of 25 μL, as well as also performed on CFX96 real-time system. Thermal conditions were 2 min of denaturation at 95 °C, followed by 45 cycles of 95 °C for 10 s, annealing at 55 °C for 20 s and extension at 72 °C for 20 s and 72 °C for 5 min. Three technical replicates were analysed for each biological replicate. All the cycle threshold (Ct) values from one gene were determined at the same threshold fluorescence value of 0.2 using theΔΔ Ct method (Xiao et al. 2016). The primers of gene-specific and housekeeping were listed.

14.25 Thawing and Revival of Inoculum

Remove a tube of inoculum from cold storage, and thaw for 5 min at room temperature. Add 1 ml double-distilled water to the tube, invert, and shake gently by hand to wash conidiosporangia from the host tissue. Remove the mass of plant tissue with forceps ensuring that water from the tissue drains back into the tube. Wash the inoculum as follows: collect inoculum into a pallet by centrifugation (1500 g for 5 min); replace supernatant with 1 ml double-distilled water, leaving the pellet in the tube; resuspend pallet and centrifuge again; repeat washing and centrifugation once again; and then replace final supernatant with 0.1 ml double-distilled water. Examine 1 μl of inoculum microscopically on a haemocytometer slide, and if possible, compare cold stores spores with fresh ones. Visible spores are spherical and non- granular; nonviable ones will appear granular and misshapen. Use the remaining spore’s suspension to inoculate fully expanded cotyledons of susceptible host seedlings. It is worthwhile doing this even if the viability of inoculum is uncertain. Place a 1–2 μl drop on each cotyledon. High humidity and cool temperature are critical, so place the pot in a clear polythene box, and incubate at 16–18 °C, 10 h 338 14 Techniques photoperiod, 50–100 μE m2 s1. Spray plants daily with a fine mist of water, and inspect for sporulation from 5 days after inoculation and every day until 5–7 days later. A low frequency (1–5%) of inoculated plants will usually become infected, so it is necessary to build up inoculum by transforming fresh spores to a pot of healthy plants. Spray plants to be inoculated with water, pick infected seedlings up with forceps, and brush conidiosporangia from infected plants onto the moistened cotyledons of healthy plants. Repeat inoculations to achieve high incidence of infection (Dangl et al. 1992).

14.26 Obtaining New Isolates from Dried Leaf Tissue Containing Oospores

It is possible to obtain new isolates of H. parasitica from stocks of field-collected and lab-produced oospores. Air-dry on a windowsill a large mass of infected plants, coarsely grind the dried tissues, and store in a glass vial at room temperature. To recover asexual cultures of the fungus, grind dried tissues to a fine powder with a mortar and pestle, plant seed of susceptible host, and sprinkle the powder onto the soil surface. Moisten soil, place pot in a polystyrene box, and vernalize for 5 days. Incubate seeds at 16–18 °C, 10 h photoperiod, and inspect seedlings daily (beginning 5–7 days after planting) for infected individuals. Transfer conidiosporangia from infected plants onto healthy plants to increase the incidence of infection. Several generations will be necessary. Maintain separate cultures from each of the original individual seedlings bearing sporulation because each represents infection from an oospore of potentially different genotype (Dangl et al. 1992).

References

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15.1 Introduction

At the international level in the recent past, agriculture research has been revolu- tionized with significant increase in the movement of crucifer’s germplasm and in the cropping patterns of this important group of crop commodity all over the world. The absence of strict measures on the restriction of movement of germplasm and the intensive and/or extensive cultivation of these crops have resulted in large-scale perpetuation, build-up, and dissemination of Hyaloperonospora parasitica virulence on cruciferous species all over the world. The information gathered in this book indicates that some gaps still exist in the complete comprehension of this disease, and this is highlighted below: even after well-developed techniques, procedures, and protocols available using Arabidopsis-Hyaloperonospora pathosystem, progress to use this information has not attained satisfactory level.

15.2 Disease Epidemiology

Epidemiological factors governing disease initiation, development, and consequent progression of the disease are not fully understood. There is need to generate more information on the role of initial inoculum in disease development, in the area of changing host susceptibility over time, and in oosporic multiplication. In Arabidopsis oospore, infection of roots has been observed, but in other crucifers, its role has not been documented. There is a need to reveal the role of oospores in root infection and systemic disease development. Time should always be included as one of the variables in the study of the relationship between pathogen development and host or environmental conditions. Multilocational trials with staggered dates of planting can be helpful in analysing disease development in relation to environmental

© Springer Nature Singapore Pte Ltd. 2017 343 G. S. Saharan et al., Downy Mildew Disease of Crucifers: Biology, Ecology and Disease Management, https://doi.org/10.1007/978-981-10-7500-1_15 344 15 Future Strategies and Priorities of Downy Mildew Disease Management conditions and to develop disease prediction models. With the change in climate pattern, more precise data are required in different ecological zones to develop disease forecasting system at different levels. There is a need to compare historical data with the recent observations.

15.3 Physiological Specialization

Host specificity has been observed since the first report of downy mildew of crucifers, but little efforts have been made on the analysis of this specificity at species level. To analyse the virulence pattern of H. parasitica, identification and standardization of host differentials are necessary. The relationship between pathogenicity on wild hosts and crop plants needs further study since wild hosts may act as a theatre for increased genetic variation in the pathogen. The use of modern techniques like RAPD finger printing may distinguish between pathotypes or even separate clonal population. PCR amplification of ITS (internal transcribed spacer) regions may be useful both for identification of isolates and in estimating their similarity. Ultimately this type of sequence analysis may reveal the evolutionary relationship between different species, genera, or higher taxa. The studies on variation between isolates obtained from different geographic regions might explain the variability with respect to virulence and to other characters such as fungicide sensitivity. A standard set of host differentials at international level in the form of isogenic lines and standardization of pathotypes nomenclature are very essential steps to validate the occurrence of pathotypes. It will also facilitate the process of breeding multigene downy mildew resistance cvs. Along with favourable gene combinations.

15.4 Genetics of Resistance

The search for new sources of resistance is always of high priority. The understanding of population biology and genetics (genetic diversity, relative fitness in geo- graphically separated population, good knowledge of host pathogen variation, availability of reliable markers like virulence/avirulence mating type, allozymes, mitochondria, and nuclear DNA RFLPS) should lead to a more effective management strategy for disease control. On the cellular level in relationship to molecular studies, the development of an axenic culture system would help in studies of heredity. Different resistance mechanisms should be characterized in more detail at the histological and cellular level. Morphological and histological attributes of downy mildew resistance should be exploited. Use of partial R genes can provide better cover towards durable resistance to downy mildew. A single gene-based resistance may not provide durable disease resistance, but the combination of isolate-specific genes that confer broad-spectrum resistance, together with an appropriate strategy for deployment of cultivators, can contribute to the management of crucifer downy mildew disease. 15.5 Molecular Plant Pathology/Genomics/Genetic Engineering 345

15.5 Molecular Plant Pathology/Genomics/Genetic Engineering

On the molecular level, a major emphasis should be placed on the development and improvement of methods for isolation of RNA and DNA; isozyme analysis; use of RFLP analysis for assessment of genetic variation; development of genetic maps; research of transposable elements and plasmids; development of fungal transformation system, i.e. availability of vectors with suitable markers; and methods for intro- ducing DNA. The cloning of avirulence and resistance genes will contribute to the elucidation of the molecular basis of host-pathogen specificity and, from the practical point of view, be helpful in the design of integrated control strategies. Using Arabidopsis-Hyaloperonospora pathosystem, a number of R genes has already been cloned and mapped which should be used and exploited to breed resistance cvs. of crucifers though modern and conventional breeding techniques. The concentrated effort to identify loci in Arabidopsis thaliana associated with specific resistance to H. parasitica should enable cloning of host resistance genes in better way. The eventual isolation and functional analysis of avirulence genes from crucifers downy mildew will be of particular interest, given the apparently separate phylogeny of this group from other plant pathogenic fungi. Based on current information generated and understanding on the molecular mechanisms of crucifer-H. parasitica interaction system, the following strategies for developing downy mildew resistance cvs. of crucifers may be adopted: (i) Identification and utilization of receptor like kinase involved in host immunity. (ii) Identification and utilization of R gene involved in effecter-triggered immunity (ETI). (iii) The utilization of quantitative trait loci (QTLs) to breed for non-race-specific and durable resistance. (iv) Screening and utilization of recessive gene-mediated broad-spectrum resistance. (v) Engineering broad-spectrum resistance through molecular genetics. (vi) Molecular bar coding system for improved and reliable species identification may be developed. (vii) More ATRs have to be cloned. (viii) To learn more details about the transfer (location and mechanism) of ATRs from H. arabidopsidis into host cell. (ix) The phenomena of organ specificity and age-related resistance (ARR) regulated at the molecular level needs in depth study. (x) A similar system and method of designation and nomenclature of R genes and AVR genes at international level should evolve and be adopted. (xi) Since genome of both host (Arabidopsis) and pathogen (H. arabidopsidis) has been sequenced, the application of gene-silencing techniques should be exploited. 346 15 Future Strategies and Priorities of Downy Mildew Disease Management

(xii) Lack of protocols for culture, genetic transformation, and established techniques like P. infestans for H. arabidopsidis. (xiii) Direct or Agrobacterium-mediated transformation for genetic manipulation and analysis of pathogen. (xiv) Establishment of temporal hierarchy of effectors during penetration, colonization, and sporulation. (xv) Molecular basis of biotrophy. (xvi) Role of genetic recombination and epigenetic on the emergence of new effectors. (xvii) To develop tools to know how plant originated molecules regulated pathogen response. (xviii) The relevance of interspecies transfer of small RNAs. (xix) New insights into molecular mechanism to compare and functional analysis of biotrophism.

15.6 Biochemical Aspects of Resistance

The elucidation of the biochemical background of biotrophy, the establishment of an intracellular interface with host cells, and the role of different infection structures should be topics of future research. Also we understand very little of the biochemical mechanisms involved in the hypersensitive reaction and in various types of resistance. There is a need to gather information concerning the effects of the downy mildew fungus on respiration, photosynthesis, and the translocation, accumulation, and transfer of carbohydrates in infected host tissues. The role of hormonal distur- bances in pathogenesis and the basis of systemic versus local lesion infection need more study. Genetical and histo-cytological descriptions of interactions, and the availability of methods for growing parasites alone and in combination with their hosts, are largely lacking.

15.7 Disease Management

There is good information on the efficacy of fungicides against downy mildew pathogen. Efforts should continue to search for low-cost effective chemicals which can provide economically significant disease control. The possibility of biocontrol agents needs to be explored. Study of integrated disease control strategies may be very useful. However, integration of all the means of control needs to be done for each crop and for each geographical region. 15.8 Phylogenetic Relationship and Co-evolution of Cruciferous Hosts and Downy… 347

15.8 Phylogenetic Relationship and Co-evolution of Cruciferous Hosts and Downy Mildew Pathogen

Evolutionary ecology has progressed from the early revolutions led by Karl Linnaeus, the founder of taxonomy, and Charles Darwin, the founder of evolutionary theory, to a molecular age to reassess classifications of species through the inference of familial history or phylogenetics of species with comparative analysis of informative DNA, RNA, and protein sequences. 1. Detailed phylogenetic relationship and evolutionary scenario within downy mildews remain still unresolved whether downy mildews are monophyletic and how the major groups of downy mildews are related to each other. Identification of phylogenetic relationship with additional molecular markers, new methods of phylogenetic information, as well as improved taxon sampling is required. Sound phylogenetic hypotheses are a precondition for detailed insights into the processes of character evolution, adaptive radiation, and speciation/specification of downy mildew on different crucifer hosts. 2. It is still little investigated whether polyploidy in speciation/specification is involved. Detailed studies on nuclear genome size may provide information on genome evolution. 3. Whole-genome analysis in an evolutionary context is better for understanding the evolutionary processes and phylogenetic relationships. The integration of whole-genome data is also promising. 4. Applicable species definitions and need for taxonomic and nomenclatural revisions where some of the species are apparently nonspecific and others are heterogeneous will resolve problem arising in uniform identity of the pathogenic species. Lecto-typification is necessary which for heterogeneous assemblages has great impact on species nomenclature. 5. Molecular bar coding system should be developed for easy, improved, and reliable species identification. Morphology is often not the best basis for identification due to lack of morphological distinctness of numerous genetically well distinct lineages. Molecular bar coding requires strict quality standards for the laboratory routine, the sequence data, as well as identification, documentation, specimen deposition, and nomenclature. Crucial for resolution of a molecular bar coding system is the selection of the sequence region used. One possible candidate is the ITS rDNA, which is a multicopy region easy to amplify in downy mildew of crucifers. It can detect pathogen with high sensitivity and specific as well as universal primers are available. Another very promising candidate for bar coding is COX2, which offers high resolution in closely related species groups. Index

A Biochemical compounds Abiotic elicitors, 261 carbohydrates, 116, 217, 218, 346 Abiotic stress, 3–5, 26, 251 enzymes, 158, 217, 222, 229, 264, 268, Accessions, 15, 17, 21, 23, 59, 70, 73, 76, 108, 324, 336 109, 127, 130, 133, 134, 140, lipids, 107–109, 116, 123, 167 166–168, 170–173, 229, 234–238, nucleic acid, 215 240–242, 245, 260, 261, 311, 329, phenolic compounds, 220, 221, 260 334, 335 protein, 10, 18, 19, 24, 116, 123, 215, 226, Aggressive, 26 232, 240, 242, 243, 249–251, 255, Albugo candida, 22, 40, 157, 176, 200, 204, 258, 259, 268, 269, 347 225, 242, 245, 317–319 Biological control, 12, 70, 225, 299, 301, 302 Allylisothiocyanate effect, 261 Biotic stress, 2, 4, 5, 26, 251 Altered phynotype of DM, 211 Botrytis parasitica, 70 Antibiotics Brassica agrimycin, 291, 292 B. alba, 129, 131, 265 agri-strep, 291, 292 B. alboglabra, 37, 47 streptomycin, 291, 310, 317 B. campestris, 37, 48, 128, 130–132, 134, Appressoria, 88, 93, 114, 157, 159–162, 183 141, 142, 154, 234, 264, 265 Arabidopsis species, 6, 37, 67, 107, 127, 159, B. carinata, 1, 2, 129–131, 133, 265 175, 220, 226, 301, 332, 343 B. caulorapa, 130, 131 Arabidopsis thaliana, 6, 9–12, 14, 17, 20, 21, B. chinensis (Chinese cabbage), 2, 11, 23, 39, 55, 70, 82, 133, 162–165, 36–38, 43, 47, 128–131, 147, 150, 232, 234, 239, 242, 247, 253, 256, 151, 153, 158, 177, 185, 233, 239, 261, 270, 345 240, 250, 251, 256, 266, 289, 302, Area Under Disease Progress Curve 313, 317–319, 335 (AUDPC), 186, 189, 240 B. fructiculosa, 47, 128 Asexual phase, 87, 88 B. juncea, 128 Association, 8, 41, 93, 199–211 B. juncea var. megarrhiza (Chinese- Avirulence (AVR), 18, 20, 23, 24, 86, 172, mustard), 129, 130 215, 225, 229, 240, 245, 249, 255, B. napobrassica, 128 344, 345 B. napus, 1, 2, 11, 21, 37, 39, 47, 128–131, Axenic culture, 149, 344 133, 134, 139, 141, 142, 151, 207, 227, 234–236, 239, 261, 264, 266, 322, 323 B B. napus subsp .oleifera, 2, 130, 140 Biochemical basis of resistance, 9, 220 B. napus subsp. rapifera, 2

Brassica (cont.) Brassica vegetables, 42, 52, 54, 289–294 B. nigra, 2, 39, 44, 47, 128–133, 266 Breeding for disease resistance, 134, 226, 238, B. oleracea var. acephala (Kale), 2, 36, 38, 267, 301, 317–319 43, 47, 48 Broad spectrum resistance, 19, 20, 225, 226, B. oleracea var. alboglobra (Chinese 344, 345 Kale), 2, 37, 47 Brussels sprouts, 2 B. oleracea var. botrytis (broccoli), 233 B. oleracea var. botrytis (cauliflower), 2, 47, 130, 131, 141, 154, 218–220, C 240, 261, 265, 286–289, 292, 294, Callus culture, 309, 321 316, 323 Camalexin, 221–223 B. oleracea var. capitata (cabbage), 2, 47, Camelina sativa, 37, 39, 44, 48, 294 130, 265 Camelina species, 78, 195, 200 B. oleracea var. fruticasa (Branching Bush Capsella bursa-pastoris, 6, 9, 12, 17, 36, 37, Kale), 2 44, 48, 68, 70, 71, 88, 128, 129, B. oleracea var. gemmifera (Brussel’s 133, 152 sprouts), 2, 36, 38, 43, 48 Cardamine impatiens, 37, 48, 71, 75, 78, 81, B. oleracea var. gongylodes (kohlrabi), 2, 84 36, 43 Caronopus didymus, 39 B. oleracea var. italic (broccoli), 2, 48, 256 Causal organism, 12 B. oleracea var. sabauda (savoy cabbage), Chemical dust, 288, 290 2 Chemical spray, 41, 196, 286–299, 302–304, B. pekinensis (Chinese cabbage), 37, 39, 311, 321, 338 44, 48, 128–131, 151 Classification, 12–14, 69, 74, 83, 347 B. rapa, 2 Climate change effect, 24–27 B. rapa subsp. chinensis (Bok choy), 2 Cole crops, 1, 184 B. rapa subsp. nipposinica, 2 Compositae, 6 B. rapa subsp. oleifera (turnip rape), 2, 43, Compost treatment, 288 47, 129, 200 Conidia cross wall, 88, 112, 113 B. rapa subsp. parachinensis, 2 Conidial discharge, 149, 322 B. rapa subsp. pekinensis (Chinese Conidial dispersal, 150, 176, 226 cabbage), 2 Conidial germination B. rapa subsp. rapifera (turnip), 2, 48, 129, relative humidity, 55, 147, 151, 152, 177, 266 180, 183, 187, 285, 302, 312, 314, B. rapa subsp. var. brown sarson (brown 321, 322, 334 sarson), 1, 2, 131, 132 temperature, 147, 150–154, 313, 321 B. rapa subsp. var. toria (toria), 1, 2, 41, Conidial longevity, 148, 177, 207, 314 48, 130–132, 149, 154, 201, 265 Conidial measurement, 3, 7, 70–72, 176 B. rapa subsp. var. yellow sarson (yellow Conidiophore and conidia, 6, 7, 46, 52, 55, 73, sarson), 1, 2, 131, 132, 154, 265 88, 93, 118, 120, 121, 158, 193, B. tournefortii, 48, 128, 130–132 200, 207, 299, 309, 313 B.juncea, 1, 2, 21, 39, 41, 44, 46, 47, 56, Conidiophore development 128, 130, 140, 141, 151, 154, 158, primordium, 111 188, 199, 201, 204, 207, 208, 217, Control, 12, 68, 135, 139, 215, 218–220, 225, 234, 236, 239, 265, 285 234, 240, 247, 285–289, 294–297, B.juncea var. multiceps (Chinese- 299, 301, 302, 310, 336, 344, 346 mustard), 129, 130 Cost-benefit ratio, 287, 295, 297 Brassica oilseeds, 7, 36–52, 129, 185, Crucifers, 1, 35, 67, 148, 157, 175, 183, 199, 294–299 215, 225, 267, 285, 309, 343 Brassica species, 2, 3, 6, 35, 70, 127, Crucifers vegetables, 127 130–132, 134, 135, 139, 143, 226, Cultivar, 2, 18, 19, 45, 46, 127, 130, 148, 153, 235, 256, 315 180, 191, 203, 225, 229–231, 234, Brassica species (rapid cycling), 133–135, 240, 241, 261, 267, 285, 301, 304, 141, 234 311, 315, 320–322, 324 Index 351

Cultural practices ITS, 140, 324–326, 328 crop rotation, 180, 225, 285, 302 RAPD scan, 138, 344 date of sowing, 208, 209 RFLP, 138 sanitation, 285, 301, 302 Downy mildew (DM), 1, 3, 35, 51, 53, 67, spacing, 302 128, 157, 175, 183, 199, 201–204, weeding, 285, 302 206, 208–210, 215, 225, 285, 310, Culture medium, 149, 309 343 Cytology and genetics Downy mildew synergism, 202, 301, 302 chromosomes, 116 mitochondria, 116, 122, 123 nucleus, 122, 123 E RNA, 116, 122 Economic importance brassica oilseeds, 36–41 brassica vegetables, 42 D Electrolyte leakage, 158, 216, 229 Defence mechanism Electron micrograph, 95, 96, 99–106, camalexin, 221, 222 118–121, 159 defence compounds, 220 Electron microscopy, 7, 12, 165 lignin, 222 Environmental factors effect salicylic acid, 20, 221, 222 dew, 180, 330 Detached leaf (cotyledon) culture, 310, 311, leaf wetness, 180 313 rainfall, 180, 183 Differential resistance, 130, 133, 134, 140, relative humidity, 180, 183, 210 234, 236 temperature, 180, 183, 210 Differential virulence, 134, 135, 322, 323, 344 wind velocity, 180 Disease, 2, 35–59, 69, 93, 127, 150, 158, Enzymes, 158, 222, 229, 252, 264, 336 175–179, 183, 199, 201–204, 206, Epidemiology, 8, 12, 142, 183, 208, 209, 215, 208–210, 220–223, 225, 285, 312, 343 343 Eruca sativa (taramira), 2, 11, 36, 37, 41, 49, Disease assessment, 7, 12, 35, 50, 56–59, 241, 55, 69, 82, 128, 130, 131 313 Erucic acid, 261 Disease cycle on Arabidopsis, 175, 177 on Crucifers, 175 F Disease development, 8, 176, 183–190, 192, Fertilizer effect, 192, 302 210, 215, 304, 317, 332, 343 Fungicide residue Disease forecasting, 12, 183, 209, 210, 344 plant, 289, 295 Disease incidence/intensity/severity, 26, 41, seed, 295, 298, 299 57, 59, 184, 189, 190, 202, 210, soil, 286–297 240, 286, 288, 302 Fungicide resistance, 133, 271, 301, 302, 316, Disease index (DI), 56, 149, 187, 235, 294, 319, 331, 344 304 Fungicide spray Disease intensity-yield loss equation, 158, Brassica oilseeds, 294–299 184, 195, 209, 287, 295, 300 Brassica vegetables, 289–294 Disease management, 9, 12, 285, 343 Fungicide tested Disease occurrence, 26, 43, 202, 204, 320 Aliette, 288, 294 Disease perpetuation, 12, 175, 343 Apron 35-WS, 292 Disease prediction, 183, 196, 344 Apron SD -35, 300 Disease rating, 41 Aspor, 291 Disease scoring scale, 56, 59 Blitox 50, 292 Disease symptoms, 35, 42–55, 121, 165, bordeaux mixture, 288, 290, 293 205, 247 Bravo, 289 DNA finger printing Brestan, 293, 294 AFLP, 140 captafol, 287–292 352 Index

Fungicide tested (cont.) prothiocarb, 287 Captan 50W, 291 quinomethionate, 290 CGA 1, 292 quintal, 293, 295 CGA 245704, 286, 292 quintozene, 290 chloranil, 288, 290 Ridomil, 287, 289, 291, 293–297, 299, chlorothalonil, 289, 291, 295 300, 302, 316 copper oxinate, 293, 294 RPA 407213, 289, 291 copper oxychloride, 288–291, 294 Sadoplon, 294 cupric hydroxide, 289, 291 Spergon, 288, 290, 291 cuprocide, 290 sulphur, 290, 291 cymoxanil, 289, 291, 292 thioneb, 291 cyprofuram, 289 thiophanate methyl, 293, 295 daconil, 288–290, 292, 293 thiovit, 292 Delan, 293, 294 thiram, 290 dichlofluanid, 287–292 tribasic copper sulphate, 291 dichlone, 290 trioneb, 293 difolatan, 289, 291–294, 296 Vancide, 290, 291 Dithane B-11, 290 zineb, 287, 288, 290, 292, 293 Dithane M-45, 287, 289, 292–297 ziram, 289, 292 Dithane Z-78, 290–295 Fungicide treatment Dow seed treatment, 290 compost, 288 Dynone, 287 foliage, 318 Ethyle B-622, 290 seed, 41, 286, 287, 316 fentin hydroxide, 289 soil, 287–288 fermate, 290 Future strategies, 343–347 fongarid, 293 fosetyl aluminium, 287, 288, 292 iprodione, 293, 295 G Kavach, 287, 293, 295–297 Garlic juice, 299 Kemate, 292 Gene expression, 26, 221, 267, 335–337 Kolophygon, 290 Genes Macuprax, 293, 294 cloning, 12, 23, 238, 252, 256, 267, 345 mancozeb, 287, 289–291, 294, 295, 302, expression, 10, 20, 26, 221, 232, 250, 251, 303 257, 267, 268, 270, 335–337 mancozeb 75 WP, 41, 293 inheritance, 12, 234, 239, 241, 256, 271 maneb, 287, 290, 291 mapping, 12, 238, 243, 244, 255, 256, 267 manzate, 290–292 mutation, 10, 18, 19, 225, 232, 243, 246, melprex, 292 249, 257, 261, 267 metalaxyl, 288–295, 298, 299, 301, 316 silencing, 268, 270, 345 metaxadine, 292 Genetic engineering, 20, 267, 302, 345, 346 milfuran, 288 Genetics of host-pathogen relationship, 233 Nabam-zinc sulphate, 290 Genomics, 10, 24, 26, 138, 139, 256, 267, 322, nickel sulphate, 291 324, 345, 346 oxadixyl, 289, 291 Geographic distribution parzate, 290 Argentina, 36 perotsin, 291 Australia, 36 phenanthraquinone, 290 Austria, 36 phenylamide, 133, 143, 271 Bavaria, 36 phosphonic acid, 289, 292 Bermuda, 36 phygon, 290, 291 Borneo, 36 polyram M (metiram), 291, 292, 294 Brazil, 36 propamocarb, 288, 289, 291, 292 Brunei, 36 propineb, 288–290, 292 Bulgaria, 36 Index 353

Canada, 36 Philippines, 39 Chile, 36 Poland, 39 China, 36 Portugal, 39 Costa Rica, 36 Puerto Rico, 39 Cuba, 36 Queensland, 39 Cypress, 36 Romania, 39 Czechoslovakia, 36 Russia, 39 Denmark, 36 Sabah, 39 Dominica, 36 Samoa, 39 Ethiopia, 37 Sarawak, 39 Fiji, 37 Saxony, 39 Finland, 37 South Africa, 39 France, 37 Spain, 39 Germany, 37 Sri Lanka, 39 Greece, 37 Sweden, 39 Guatemala, 37 Switzerland, 39 Haiti, 37 Taiwan, 39 Holland, 37 Tanzania, 39 Hong Kong, 37 Thailand, 39 Hungary, 37 Trinidad and Tobago, 39 Iberia (Spain), 37 Turkey, 39 India, 37 U.S.S.R., 40 Iran, 37 Uganda, 40 Iraq, 37 United Kingdom, 40 Ireland, 37 United States, 40 Israel, 37 Uruguay, 40 Italy, 37 Venezuela, 40 Jamaica, 37 Victoria, 40 Japan, 38 Vietnam, 40 Kampuchea, 38 Yugoslavia, 40 Kenya, 38 Zimbabwe, 40 Korea, 38 Germplasm screening, 19, 267, 311–313, 329 Libya, 38 Glucosinolate effect, 261 Ludlow, 38 Growth substances, 99, 215 Malawi, 38 Malaysia, 38 Malta, 38 H Mauritius, 38 Haustoria, 11, 52, 84, 87, 93, 97, 99, 108–110, Mexico, 38 115, 134, 158–163, 165–169, Montpellier, 38 171–173, 176, 179, 184, 205, 206, Moravia, 38 221, 226, 229, 230, 232, 310 Morocco, 38 Haustorium development, 7, 93, 97, 98, 101 Mozambique, 38 Heterothallism, 7, 142, 320 Nepal, 38 Histopathology, 204, 205, 207, 208 New South Wales, 38 Homologous, 10, 20, 68, 133, 143, 234, 268, New Zealand, 38 270 North Borneo, 36 Homothallism, 7, 142, 255, 320 Norway, 38 Horse radish, 11, 36, 40, 43, 47, 199 Pakistan, 38 Host differentials, 129, 134, 135, 140, 344 Palestine, 38 Host-pathogen interaction, 8, 19, 21, 23, 134, Panama, 38 215–222, 225, 226, 231, 238, 312 Papua New Guinea, 39 Host-pathogen interface, 7, 93, 98 Peru, 39 Host-pathogen recognition system, 8, 231 354 Index

Host penetration, 7, 93, 97, 158, 183 I Host range, 12, 15–17, 26, 35, 43, 68, 73, 86, Immune, 18, 23, 24, 58, 128, 226, 227 127, 129, 325 Immunity, 18, 19, 345 Host resistance, 6 Infection process, 12, 159–163, 334 adult plant, 240, 247, 249, 256 Inheritance of resistance, 19, 20, 22, 233, biochemical, 8, 12, 69, 93, 215, 220, 222, 238–242, 319, 345 231, 232, 260, 346 additive genes, 239, 241 genetical, 10, 12, 15, 17, 68, 75, 80, 82, 87, allelic, 239 93, 140, 221, 346, 347 dominant genes, 239 histological, 12, 215, 344 epistatic, 246 molecular, 6, 10, 12, 13, 15–17, 19, 21, 24, partial 68, 70, 74, 79, 81–84, 86, 127, 138, partial dominant, 238, 239 225, 226, 231, 238, 242, 243, 252, recessive genes, 19, 20, 233, 239, 319, 267, 324–327, 344, 345, 347 345 morphological, 3, 6, 12, 13, 15, 16, 43, single gene, 240–242 67–69, 73, 74, 81, 83, 84, 99, 129, tightly linked genes, 22, 239, 240 344, 347 Inoculum concentration, 158 multiple, 2, 15, 20, 107, 135, 161, 226, Inoculum dose relationship, 302 242, 245, 302, 304, 317–319, 332 Insecticide spray, 196 seedlings, 6, 43, 57, 59, 127, 129, 184, nutrition, 192 211, 233, 240, 241, 243, 247, 311, planting time, 192 316, 320, 321, 323–325, 333, 338 Integrated disease management, 9, 301–304 Host response and reaction, 135, 236 Interaction phenotype (IP), 21, 50, 57, 59, 134, Host responses, 93, 115, 133, 165, 211, 249, 136, 137, 234, 235, 238, 312, 313, 332 328, 330 Hyaloperonospora, 4, 35, 67, 99, 127, Isolate, 6, 43, 68, 166, 177, 200, 218, 226, 148–150, 157, 192, 199, 216, 225, 299, 310, 334, 344 309, 343 Hyaloperonospora arabidopsidis, 11, 16, 23, 24, 86, 87, 107, 269 K Hyaloperonospora brassicae, 84–86, 135–137 Knop's medium, growth on, 310 Hyaloperonospora parasitica, 9, 10, 12, 14, Kohlrabi, 37–39, 45, 48, 147, 149, 158, 177, 17, 36–40, 43, 67, 108, 109, 329 118–124, 129–132, 134, 138, 141, 142, 148–150, 152–154, 159–166, 176, 179, 189–193, 199, 204, 211, L 217–220, 225, 227–230, 232, Leaf wetness, 8, 180, 186, 187, 190, 191 235–237, 240, 242, 246, 249, 253, Lepidium, 39, 40, 44, 49, 71, 75, 78, 84, 150, 254, 264–266, 269, 309, 310, 299 313–315, 317, 318, 325, 343 Light, 4, 7, 12, 46, 50, 52, 54–57, 59, 76, 86, Hyaloperonospora parasitica-Albugo candida, 93, 99, 137, 151, 154, 159–165, mixed infection, 40, 46, 192, 193, 169, 185, 188, 227, 311, 315, 316, 199, 200, 204, 225, 317–319 323, 329–332, 334, 335 Hyaloperonospora species, 17, 43, 74, 75, 80, Lignin/lignification, 9, 134, 221–223, 262, 264 82, 87, 324 Local infection, 6, 157, 346 Hybridization, 139, 143, 267, 324 Hypersensitive reaction, 162, 163, 231, 250, 346 M Hypertrophy (malformation), 5, 46, 149, 157, Maledmia africana, 37 200, 209, 210 Matthiola species (stocks), 44, 49, 54, 55, 88 Hyphae, 52, 88, 93, 95, 97, 103, 105, Mechanism of resistance, 8, 12, 226 108–110, 149, 159–162, 165–168, Medium, 154 170–173, 179, 185, 221, 222, 229, Metabolic changes, 8, 215, 217 230, 264, 310 Micronutrients, 180, 304 Index 355

Mildews, 3, 110 biological forms, 127, 141 Milestones in DM research, 6, 10–17, 86 biological strains, 128, 141 Mixed infection identification, 18, 19, 24, 128, 140 DM+ mustard mosaic, 5, 180, 199 isolates, 127, 129, 130, 134, 135, 140 DM+WR, 8, 46, 192, 199, 201–204, 206, Pathotype 208–210 designation, 20, 21, 23, 140 Molecular bar coding, 345, 347 nomenclature, 20, 21, 23, 140, 344 Molecular plant pathology, 23, 24, 345, 346 races, 129, 140, 141 Morphology, 6, 12, 14, 15, 26, 69, 70, 72–76, races, 16, 20, 21, 23 78, 80–87, 205, 347 strains, 138 Mustard, 2, 11, 27, 38, 40–44, 47–51, 130, Perofascia, 6, 11, 13, 14, 68, 73, 75, 76, 140, 141, 148, 149, 177, 186, 188, 84, 85 189, 191–193, 195, 199, 200, Peronospora, 3, 67, 246, 327 202–204, 206, 208, 209, 265, 286, Peronospora brassicae, 67 287, 294, 295, 297, 298, 303, 304 Peronospora parasitica, 6, 10, 14, 84, 94 Mustard mosaic virus, 5, 180, 199 Peronospora parasitica-Albugo candida, Mycelium, 3, 8, 35, 54, 87, 88, 123, 147, 149, mixed infection, 41, 199 161, 175, 177, 179, 205–207, 227, Peronospora ramicis, 70 299, 309, 310, 317, 320, 321, 325 Peronospora species, 15, 70, 110 Peroxidase, 158, 217, 220, 222, 229, 252, 263, 264 N Perpetuation, 8, 12, 147–154, 175, 343 Nutrition, 8, 180, 192, 221, 301, 309 Phenolic compounds, 220, 260 Phylogeny, 12, 69, 70, 73, 345 Physiologic specialization, 127–145 O Phytoalexin, 208, 220, 221, 261, 304 Oilseed rape, 41, 43, 47, 57, 130, 138–140, Planting time, 8, 192, 195, 208 142, 192, 193, 207, 218–220, Primary infection, 176 227–230, 234, 261, 302, 313, 323 Oospore germination age, 154 Q light, 154 Quantitative trait loci (QTLs), 19, 226, 345 pH, 154 relative humidity, 154 temperature, 154 R Oospore infection, 148, 149, 177, 343 Race nomenclature, 20, 21, 23, 81, 82, 140, Oospore perpetuation 344, 345, 347 over summering, 147, 175 Radish, 2, 11, 36–40, 42, 43, 50, 54, 87, over wintering, 147, 175 127–130, 141, 147–149, 154, 158, Oospores, 8, 11, 50, 54, 70, 89, 142, 143, 177, 226, 234, 262, 287, 293, 294, 147–149, 164, 165, 175–177, 179, 299, 314 206, 285, 302, 309, 313, 320, 338, Rainfall (water), 8, 25, 180, 183, 185, 187, 343 194, 195, 209, 210 Rape, 36–38, 41, 47, 128, 134, 141, 158, 285, 286, 309 P Rapeseed-mustard, 1, 25, 27, 43, 58, 59, 148, Parasite, 10, 12, 13, 17, 67, 69, 70, 75, 78, 177, 187, 192, 197, 285, 286, 292, 81–83, 86, 93, 106, 127, 161, 175, 302 215, 242, 243, 301, 346 Raphanus raphanistrum, 2, 44, 128 Pathogen, 40, 67–89, 127, 147, 159, 175, 183, Raphanus sativus, 2, 11, 50, 71, 128, 130–132, 199, 225, 285, 309, 343 134, 148, 266, 320 Pathogenesis, 8, 12, 18–20, 157–173, 175, Raphanus sativus L., 71 202, 204, 216, 228, 346 Rating scales, 57 Pathogenic variability Receptor like kinases, 18, 19 356 Index

Relative humidity, 55, 147, 151, 152, 177, Stock, 36–39, 43, 44, 49, 54, 55, 199, 293, 180, 183, 187, 192, 194, 195, 210, 329, 334, 338 285, 302, 312, 314, 321, 322, 334, Survival 336 conidia, 8, 147, 148 Reproduction and reproductive structures, 7, mycelium, 8, 147 87–89 oospores, 8, 148, 177 Resistance, 9, 18–22, 127, 172, 220, 221, 233, Susceptible, 1–3, 18, 20, 23, 43, 46, 54, 56, 234, 236, 240, 243 59, 108–110, 128, 133–135, 158, Resistance genes (R-genes), 10, 20, 21, 23, 165, 172, 226, 228, 236, 238, 247, 250, 267, 345 248, 261, 313, 315, 334, 335 cloning, 12, 23, 252, 256, 345 Swede, 2, 36, 38, 43, 47, 309, 310 designation, 20–23, 267, 345 Symptoms, 200 different requirements, 249 Arabidopsis, 55, 165 differential expression, 10, 250–252 oilseed brassica, 42–52 identification, 12, 19, 24 rapeseed-mustard, 43 mapping, 12 rocket, 44, 45, 49, 50, 55 mutation, 10, 18, 19 stock, 43, 44, 50, 54, 55 nomenclature, 20, 21, 23 vegetable brassica, 200 Resistant, 18, 20, 56, 58, 128, 130, 134, 135, wall flower, 36, 39, 43, 44, 49, 54 139, 142, 165, 168–170, 200, 207, Systemic acquired resistance (SAR), 9, 10, 20, 221, 226, 228, 236, 238, 240, 245, 180, 231, 232, 245, 301 248, 316, 335 Systemic infection, 6, 54, 56, 176, 316, 317 Rutabaga, 2, 43, 127

T S Taxonomy, 12–14, 69, 70, 72–76, 78–87, 347 Secondary infection, 176 Techniques, 9, 12, 24, 69, 225, 226, 252, 267, Seed-borne, 320, 321 301, 309–338, 343–346 Seed transmission, 148, 177, 321 Temperature, 8, 24–26, 88, 147, 150–152, 154, Seed treatment, 41, 286, 287, 293–295, 175, 177, 183–185, 189, 194, 209, 298–300, 302, 303 314, 316, 322, 334 Sexual phase, 88, 89 Temperature effect (conidia, conidiophores), Signal transduction, 231–233, 250, 257, 260, 147, 152, 175 301 Temperature effect (disease), 24–27, 158, 184, Sinapis alba, 37, 44, 129 186, 188, 189 Sinapis species (white mustard), 78, 86 Temperature effect (oospores), 154, 175, 177, Sisymbrium species, 39, 84, 129 338 Soil borne, 176 Toria, 1, 2, 41, 48, 132, 149, 201 Soil treatment, 287, 288 Transmission electron microscopy Sources of inoculum infected tissues, 54, 109, 148, 149, 177, 262 primary inoculum, 285 Turnip, 2, 36–40, 43, 44, 48, 52, 87, 127, 129, secondary inoculum, 180 130, 141 Sources of resistance, 9, 264–266, 301, 344 Turnip rape, 2, 43, 47, 200 Sporangia, 14, 55, 154, 200, 205–207, 218 Sporangiophore, 11, 13, 14, 50, 55, 59, 205, 206 U Sporulation, 12, 24, 26, 44–46, 55–57, 89, U’s triangle, 3 111, 137, 152, 154, 161, 162, 165, Ultra structures 166, 183, 184, 199, 215, 216, 226, haustorium, 109, 115 227, 233, 241, 271, 286, 311, 315, host cell, 97, 107, 109, 159 316, 321, 322, 325, 334, 338, 346 hypae, 109 Index 357

W Y Wallflower, 36, 39, 43, 44, 48, 54 Yield increase, 286, 297 Watercress, 11, 36, 40, 43, 44, 49 Yield loss assessment, 7, 11, 12, 16, 40–42, White rust (WR), 5, 8, 21, 40–42, 46, 50, 52, 59, 201–204, 295, 300 187, 192–195, 199, 201–204, 206, 208–210, 286, 297, 299, 302–304