Handbook of Florists’ Crops Diseases 9783319396705, 9783319396682, 9783319396699

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Handbook of Plant Disease Management Series Editors: Robert J. McGovern · Wade H. Elmer

Robert J. McGovern Wade H. Elmer  Editors

Handbook of Florists’ Crops Diseases

Handbook of Plant Disease Management Series Editors Robert J. McGovern NBD Research Co. Ltd. Lampang, Thailand Department of Entomology and Plant Pathology Chiang Mai University Chiang Mai, Thailand Wade H. Elmer Department of Plant Pathology and Ecology The Connecticut Agricultural Experiment Station New Haven, CT, USA

Our objective for the Handbook of Plant Disease Management series is to provide up-to-date, field-tested information on integrated disease management that will be useful to a diverse technical audience interested in a wide variety of crops. The series will have an international focus and will address these questions: “What is the disease?” and “How can it be managed?” The Handbook of Plant Disease Management series will include: Handbook of Florists’ Crops Diseases, Handbook of Vegetable Diseases, Handbook of Tropical Fruit Diseases, Handbook of Temperate Fruit Diseases. More information about this series at http://www.springer.com/series/13570

Robert J. McGovern • Wade H. Elmer Editors

Handbook of Florists' Crops Diseases With 522 Figures and 49 Tables

Editors Robert J. McGovern NBD Research Co. Ltd. Lampang, Thailand Department of Entomology and Plant Pathology Chiang Mai University Chiang Mai, Thailand

Wade H. Elmer Department of Plant Pathology and Ecology The Connecticut Agricultural Experiment Station New Haven, CT, USA

ISBN 978-3-319-39668-2 ISBN 978-3-319-39670-5 (eBook) ISBN 978-3-319-39669-9 (print and electronic bundle) https://doi.org/10.1007/978-3-319-39670-5 Library of Congress Control Number: 2017948983 # Springer International Publishing AG 2018 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, express or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. Printed on acid-free paper This Springer imprint is published by Springer Nature The registered company is Springer International Publishing AG The registered company address is: Gewerbestrasse 11, 6330 Cham, Switzerland

Preface

Florists’ crops production has evolved considerably through new technological advances in irrigation, media, and environmental control, along with the appearance of new centers of large-scale production of plant material. In addition, many florists’ crops have changed their production cycles by shifting the initial production stage to offshore facilities. While this offers new benefits in product quality, quantity, and cost, it removes some opportunities to introduce management strategies to suppress future outbreaks. These changes have necessitated the development of innovative ways of suppressing pathogenic fungi, bacteria, viruses, and nematodes. The objective of the Handbook of Florists’ Crops Diseases is to provide research-based information on the diagnosis and management of diseases of cut-flowers and potted flowering plants. We believe that following such an integrated approach will help to minimize nontarget effects of flower production and increase its sustainability. The introductory chapters, “Components of Integrated Disease Management,” present disease management strategies that are applicable to all florists’ crops. The chapters on the individual crops that follow in the “Florists’ Crops Diseases” section are grouped alphabetically within three subsections: “Cut-flowers,” “Potted Flowers,” and “Flowering Geophytes”. The grouping of crops is somewhat arbitrary; some like orchid could fit into more than one subsection: cut-flowers and potted flowers. These chapters present information on the geographic occurrence and impact, symptoms/ signs, biology and epidemiology, and integrated management of major diseases of these plants. While it is not possible to cover the diagnosis and management of all diseases on every florists’ crop, we believe that the extensive cross-referencing and indexing used in the handbook will provide assistance where specific information is lacking. We hope that the global perspective of the handbook and the detailed information provided will make it useful to florists’ crops producers, researchers, extension personnel, and students around the world. This volume is one of a series of Handbooks of Plant Disease Management being published by Springer.

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Acknowledgments

We thank the many participating authors (over 60 from 11 countries) who freely gave of their time to share their considerable expertise on the management of florists’ crops diseases. We appreciate not only the efforts of these individuals on their own chapters but also their generosity in providing diagnostic images to authors of other chapters. We also thank the many other individuals acknowledged throughout the handbook who provided images of and information on florists’ crops diseases. We are indebted to the Springer editorial team of Sylvia Blago, Simone Giesler, Zuzana Bernhart, and Priya Ponnusamy for their guidance in initiating and shaping the Handbook of Florists’ Crops Diseases.

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Contents

Volume 1 1

Florists’ Crops: Global Trends and Disease Impact . . . . . . . . . . . . Robert J. McGovern and Wade H. Elmer

Part I

1

Components of Integrated Disease Management . . . . . . . . .

11

2

Fundamentals and Advances in Plant Problem Diagnostics . . . . . Tim Schubert, Ayyamperumal Jeyaprakash, and Carrie Harmon

13

3

Nutritional Disorders of Florists’ Crops . . . . . . . . . . . . . . . . . . . . . Rosa E. Raudales

41

4

Insect Management for Disease Control in Florists’ Crops . . . . . . Raymond A. Cloyd

69

5

Breeding for Disease Resistance in Florists’ Crops Zhanao Deng

............

87

6

Environment Modification for Disease Management . . . . . . . . . . . Maria Lodovica Gullino and Angelo Garibaldi

119

7

Fungicides and Biocontrols for Management of Florists’ Crops Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cristi L. Palmer and Ely Vea

8

Soil/Media Disinfestation for Management of Florists’ Crops Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Erin N. Rosskopf, Nancy Kokalis-Burelle, Steven A. Fennimore, and Cheryl A. Wilen

137

167

9

Sanitation for Management of Florists’ Crops Diseases . . . . . . . . . Warren E. Copes

201

10

Mineral Nutrition and Florists’ Crops Diseases . . . . . . . . . . . . . . . Lawrence E. Datnoff and Wade H. Elmer

237

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11

Contents

Postharvest Disease Management . . . . . . . . . . . . . . . . . . . . . . . . . . Anastasios I. Darras

Part II

Florists’ Crops Diseases: Cut Flowers . . . . . . . . . . . . . . . . . .

253

281

12

Diseases of Anthurium Anne M. Alvarez

..................................

283

13

Diseases of Carnation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Silvia M. Wolcan, Ismael Malbrán, Cecilia A. Mourelos, Marina N. Sisterna, Mirian del P. González, Adriana M. Alippi, Andrés Nico, and Gladys A. Lori

317

14

Diseases of Celosia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ann B. Gould

379

15

Diseases of China Aster . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Harukuni Horita and Robert J. McGovern

419

16

Diseases of Chrysanthemum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jane C. Trolinger, Robert J. McGovern, Wade H. Elmer, Nancy A. Rechcigl, and Christine M. Shoemaker

439

17

Diseases of Delphinium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Stephen N. Wegulo

503

18

Diseases of Gerbera . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Elizabeth I. Brisco-McCann and Mary K. Hausbeck

533

19

Diseases of Gypsophila . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Silvia M. Wolcan, Cecilia A. Mourelos, Marina N. Sisterna, Mirian del P. González, Adriana M. Alippi, Andrés Nico, and Gladys A. Lori

561

20

Diseases of Lisianthus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Robert J. McGovern

583

21

Diseases of Orchid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Prasartporn Smitamana and Robert J. McGovern

633

Volume 2 22

Diseases of Peonies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Andrea R. Garfinkel and Gary A. Chastagner

663

23

Diseases of Proteaceae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Brett A. Summerell

693

24

Diseases of Rose . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jay W. Pscheidt and Tatiana Gomez Rodriguez

713

Contents

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25

Diseases of Snapdragon . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Stephen N. Wegulo and A. R. Chase

743

26

Diseases of Stock . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Steven T. Koike

767

27

Diseases of Sunflower . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Thomas J. Gulya, Febina Mathew, Robert Harveson, Samuel Markell, and Charles Block

787

28

Diseases of Zinnia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Dorota Szopińska

839

Part III

Florists’ Crops Diseases: Potted Flowers . . . . . . . . . . . . . . .

871

29

Diseases of Azalea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Robert G. Linderman

873

30

Diseases of Begonia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cristina Rosa and Gary W. Moorman

891

31

Diseases of Coleus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Blair R. Harlan and Mary K. Hausbeck

911

32

Diseases of Gardenia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. J. Palmateer and A. R. Chase

927

33

Diseases of Geranium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cristina Rosa and Gary W. Moorman

941

34

Diseases of Holiday Cacti: Schlumbergera and Hatiora . . . . . . . . . Robert L. Wick

975

35

Diseases of Hydrangea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yonghao Li, Margaret T. Mmbaga, Boru Zhou, Jacqueline Joshua, Emily Rotich, and Lipi Parikh

987

36

Diseases of Kalanchoe . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1007 Robert L. Wick

37

Diseases of Poinsettia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1021 Margery L. Daughtrey and A. R. Chase

Part IV

Florists’ Crops Diseases: Flowering Geophytes . . . . . . . . .

1071

38

Diseases of Caladium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1073 H. M. Bowman, J. E. Polston, and Robert J. McGovern

39

Diseases of Cyclamen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1099 Wade H. Elmer and Margery L. Daughtrey

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Contents

40

Diseases of Daffodil (Narcissus) . . . . . . . . . . . . . . . . . . . . . . . . . . . 1129 Gordon R. Hanks and Gary A. Chastagner

41

Diseases of Lily . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1229 Gary A. Chastagner, Jaap M. van Tuyl, Martin Verbeek, William B. Miller, and Becky B. Westerdahl

42

Diseases of Gladiolus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1289 Wade H. Elmer and Kathryn K. Kamo

43

Diseases of Tulip . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1313 Robert J. McGovern and Wade H. Elmer Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1339

About the Editors

Prof. Dr. Robert J. McGovern received his M.S. (1983) and Ph.D. (1986) in Plant Pathology from Cornell University. He joined Flower Time/Frank’s Nursery & Crafts, Inc., the largest garden center chain in the USA, in 1985 and served first as its plant pathologist/academic liaison, and then as director of the Horticulture and Diagnostics Department. He joined the University of Florida, Institute of Food and Agricultural Sciences (UF-IFAS), Department of Plant Pathology in 1990. Dr. McGovern served as an assistant professor with extension, research, and teaching responsibilities for integrated disease management in vegetables and citrus (Southwest Florida Research and Education Center, 1990–1995) and as an associate professor focused on management of ornamental and vegetable diseases (Gulf Coast Research and Education Center, 1995–2002). He was promoted to full professor in 2002 and moved to the UF-IFAS main campus to become director of the Plant Medicine Program, which he led from 2002 to 2011. During this period, Dr. McGovern also helped to establish a number of other plant medicine/plant health programs in Asia and the USA, and served as co-director/director of the Southern Plant Diagnostic Network, a component of the USDA-coordinated National Plant Diagnostic Network. He retired from UF-IFAS in 2011 and is a professor emeritus of its Department of Plant Pathology. Currently, Dr. McGovern is a senior consultant with the NBD Research Co., Ltd., Lampang, Thailand, and an adjunct professor in the Chiang Mai University, Department of Entomology and Plant Pathology, Chiang Mai, Thailand. His research and teaching interests include integrated plant disease management, scientific writing and editing, and multidisciplinary training. Dr. McGovern is the author of over 150 scientific publications.

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About the Editors

Dr. Wade H. Elmer is the chief scientist of the Department of Plant Pathology and Ecology at the Connecticut Agricultural Experiment Station in New Haven, CT, USA. He received his Ph.D. from Michigan State University in 1985 and began his career at the Connecticut Agricultural Experiment Station in 1987. He specializes in soil-borne diseases of ornamentals and vegetables. Dr. Elmer has devoted much attention to the epidemiology and suppression of diseases caused by fungal pathogens in the genus Fusarium. Understanding the mechanisms of disease suppression through mineral nutrition has been his longterm goal. Dr. Elmer is currently investigating the influence of nanoparticles of metal oxide composites on plant health and root disease. In 2008, Dr. Elmer coedited a book entitled Mineral Nutrition and Plant Disease (APS Press) which was awarded the CHOICE Award for Best Outstanding Academic Title and has been a best seller for 8 years. Throughout his career, Dr. Elmer has traveled widely and presented invited lectures and presentations in Australia, Brazil, Canada, Chile, China, Colombia, France, Germany, Iceland, India, Italy, Thailand, The Netherlands, and Peru. Dr. Elmer has authored or coauthored over 95 peer-reviewed research papers, two books, 16 book chapters, and over 100 articles for trade magazines, symposia, fact sheets, and bulletins.

Contributors

Adriana M. Alippi Centro de Investigaciones de Fitopatología (CIDEFI–UNLP– CICBA), Facultad de Ciencias Agrarias y Forestales, Universidad Nacional de La Plata, La Plata, Buenos Aires, Argentina Comisión de Investigaciones Científicas de la Provincia de Buenos Aires (CICBA), La Plata, Buenos Aires, Argentina Anne M. Alvarez Department of Environmental Protection Sciences, University of Hawaii, Honolulu, HI, USA Charles Block Seed Science Center, Iowa State University, Ames, IA, USA H. M. Bowman Biofire Defense LLC, Salt Lake City, UT, USA Elizabeth I. Brisco-McCann Department of Plant, Soil and Microbial Sciences, Michigan State University, East Lansing, MI, USA A. R. Chase Chase Agricultural Consulting LLC, Cottonwood, AZ, USA Gary A. Chastagner Department of Plant Pathology, Puyallup Research and Extension Center, Washington State University, Puyallup, WA, USA Raymond A. Cloyd Kansas State University, Manhattan, KS, USA Warren E. Copes USDA ARS Thad Cochran Southern Horticultural Laboratory, Poplarville, MS, USA Anastasios I. Darras Department of Agricultural Technology, Technological Educational Institute (TEI) of Peloponnese, Kalamata, Messinia, Greece Lawrence E. Datnoff Department of Plant Pathology and Crop Physiology, Louisiana State University Agricultural Center, Baton Rouge, LA, USA Margery L. Daughtrey Section of Plant Pathology and Plant-Microbe Biology, Long Island Horticultural Research and Extension Center, Cornell University, Riverhead, NY, USA Mirian del P. González Cátedra de Fitopatología, Facultad de Ciencias Agrarias, Universidad Nacional de Rosario, Zavalla, Santa Fe, Argentina xv

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Contributors

Zhanao Deng Department of Environmental Horticulture, Institute of Food and Agricultural Sciences, Gulf Coast Research and Education Center, University of Florida, Wimauma, FL, USA Wade H. Elmer Department of Plant Pathology and Ecology, The Connecticut Agricultural Experiment Station, New Haven, CT, USA Steven A. Fennimore Department of Plant Sciences, UC Davis, Davis, CA, USA Andrea R. Garfinkel Department of Plant Pathology, Puyallup Research and Extension Center, Washington State University, Puyallup, WA, USA Angelo Garibaldi Centre of Competence Agroinnova, University of Torino, Grugliasco, Italy Tatiana Gomez Rodriguez GR Chia SAS, Centro Empresarial Centro Chia Oficina 304 Chia, Cundinamarca, Bogotá, Colombia Ann B. Gould Department of Plant Biology, School of Environmental and Biological Sciences, Rutgers, The State University of New Jersey, New Brunswick, NJ, USA Maria Lodovica Gullino Centre of Competence Agroinnova, University of Torino, Grugliasco, Italy DISAFA, University of Torino, Grugliasco, Italy Thomas J. Gulya USDA-Agricultural Research Service, Sunflower and Plant Biology Research Unit, Fargo, ND, USA Gordon R. Hanks Warwick Crop Centre, Wellesbourne Campus, University of Warwick, Wellesbourne, Warwickshire, UK Blair R. Harlan Department of Plant, Soil and Microbial Sciences, Michigan State University, East Lansing, MI, USA Carrie Harmon Department of Plant Pathology, University of Florida, Gainesville, FL, USA Robert Harveson Department of Plant Pathology, Panhandle Research and Extension Center, University of Nebraska, Scottsbluff, NE, USA Mary K. Hausbeck Department of Plant, Soil and Microbial Sciences, Michigan State University, East Lansing, MI, USA Harukuni Horita Department of Plant Development, Hokkaido Central Agricultural Experiment Station, Hokkaido Research Organization, Naganuma, Hokkaido, Japan Ayyamperumal Jeyaprakash Division of Plant Industry, Florida Department of Agriculture and Consumer Service, Gainesville, FL, USA

Contributors

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Jacqueline Joshua Department of Agricultural and Environmental Sciences, Tennessee State University, Nashville, TN, USA Kathryn K. Kamo USDA-ARS, Beltsville, MD, USA Steven T. Koike University of California Cooperative Extension, Monterey County, Salinas, CA, USA Nancy Kokalis-Burelle USDA Horticultural Research Lab, Ft. Pierce, FL, USA Yonghao Li Department of Plant Pathology and Ecology, The Connecticut Agricultural Experiment Station, New Haven, CT, USA Robert G. Linderman Plant Health, LLC, Corvallis, OR, USA Gladys A. Lori Centro de Investigaciones de Fitopatología (CIDEFI–UNLP– CICBA), Facultad de Ciencias Agrarias y Forestales, Universidad Nacional de La Plata, La Plata, Buenos Aires, Argentina Comisión de Investigaciones Científicas de la Provincia de Buenos Aires (CICBA), La Plata, Buenos Aires, Argentina Ismael Malbrán Centro de Investigaciones de Fitopatología (CIDEFI–UNLP– CICBA), Facultad de Ciencias Agrarias y Forestales, Universidad Nacional de La Plata, La Plata, Buenos Aires, Argentina Consejo Nacional de Investigaciones Científicas y Tecnológicas (CONICET), La Plata, Buenos Aires, Argentina Samuel Markell Department of Plant Pathology, North Dakota State University, Fargo, ND, USA Febina Mathew Department of Agronomy, Horticulture, and Plant Science, South Dakota State University, Brookings, SD, USA Robert J. McGovern NBD Research Co. Ltd., Lampang, Thailand Department of Entomology and Plant Pathology, Chiang Mai University, Chiang Mai, Thailand William B. Miller School of Integrative Plant Sciences, Horticulture Section, Cornell University, Ithaca, NY, USA Margaret T. Mmbaga Department of Agricultural and Environmental Sciences, Tennessee State University, Nashville, TN, USA Gary W. Moorman Department of Plant Pathology and Environmental Microbiology, The Pennsylvania State University, University Park, PA, USA Cecilia A. Mourelos Centro de Investigaciones de Fitopatología (CIDEFI–UNLP– CICBA), Facultad de Ciencias Agrarias y Forestales, Universidad Nacional de La Plata, La Plata, Buenos Aires, Argentina Consejo Nacional de Investigaciones Científicas y Tecnológicas (CONICET), La Plata, Buenos Aires, Argentina

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Contributors

Andrés Nico Cátedra de Horticulture and Floriculture, Facultad de Ciencias Agrarias y Forestales, Universidad Nacional de La Plata, La Plata, Buenos Aires, Argentina A. J. Palmateer Department of Plant Pathology, Tropical Research and Education Center, University of Florida, Homestead, FL, USA Cristi L. Palmer IR-4 Project, Rutgers University, Princeton, NJ, USA Lipi Parikh Department of Agricultural and Environmental Sciences, Tennessee State University, Nashville, TN, USA J. E. Polston Department of Plant Pathology, University of Florida, Gainesville, FL, USA Jay W. Pscheidt Department of Botany and Plant Pathology, Oregon State University, Corvallis, OR, USA Rosa E. Raudales Department of Plant Science and Landscape Architecture, University of Connecticut, Storrs, CT, USA Nancy A. Rechcigl Syngenta Crop Protection, LLC, Greensboro, NC, USA Cristina Rosa Department of Plant Pathology and Environmental Microbiology, The Pennsylvania State University, University Park, PA, USA Erin N. Rosskopf USDA Horticultural Research Lab, Ft. Pierce, FL, USA Emily Rotich Department of Agricultural and Environmental Sciences, Tennessee State University, Nashville, TN, USA Tim Schubert Division of Plant Industry, Florida Department of Agriculture and Consumer Service, Gainesville, FL, USA Christine M. Shoemaker Syngenta Flowers, LLC, Gilroy, CA, USA Marina N. Sisterna Centro de Investigaciones de Fitopatología (CIDEFI–UNLP– CICBA), Facultad de Ciencias Agrarias y Forestales, Universidad Nacional de La Plata, La Plata, Buenos Aires, Argentina Comisión de Investigaciones Científicas de la Provincia de Buenos Aires (CICBA), La Plata, Buenos Aires, Argentina Prasartporn Smitamana Agricultural and Industrial Clinic Co. Ltd., Chiang Mai, Thailand Department of Plant Pathology, Faculty of Agriculture, Chiang Mai University, Chiang Mai, Thailand Brett A. Summerell Royal Botanic Gardens and Domain Trust, Sydney, NSW, Australia Dorota Szopińska Department of Phytopathology, Seed Science and Technology, Poznań University of Life Sciences, Poznań, Poland

Contributors

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Jane C. Trolinger Syngenta Flowers, LLC, Gilroy, CA, USA Jaap M. van Tuyl Wageningen University and Research, Wageningen, The Netherlands Ely Vea IR-4 Project, Rutgers University, Princeton, NJ, USA Martin Verbeek Wageningen University and Research, Wageningen, The Netherlands Stephen N. Wegulo Department of Plant Pathology, University of Nebraska-Lincoln, Lincoln, NE, USA Becky B. Westerdahl Department of Entomology and Nematology, University of California, Davis, CA, USA Robert L. Wick Stockbridge School of Agriculture, University of Massachusetts, Amherst, MA, USA Cheryl A. Wilen UC IPM, San Diego, CA, USA Silvia M. Wolcan Centro de Investigaciones de Fitopatología (CIDEFI–UNLP– CICBA), Facultad de Ciencias Agrarias y Forestales, Universidad Nacional de La Plata, La Plata, Buenos Aires, Argentina Comisión de Investigaciones Científicas de la Provincia de Buenos Aires (CICBA), La Plata, Buenos Aires, Argentina Boru Zhou Department of Forest Protection, Northeast Forestry University, Harbin, China

1

Florists’ Crops: Global Trends and Disease Impact Robert J. McGovern and Wade H. Elmer

Abstract

The production of cut-flowers, potted flowering plants, and related propagative material is a dynamic, fast-growing, and highly lucrative global industry. In the past, flower production sites were primarily located in or near the major markets – developed countries in North America and Europe. However, because of lower production costs, including labor inputs, more favorable climatic conditions, and the development of efficient transportation methods, there has been a shift in production to exporting countries closer to the equator such as those in Latin America, sub-Saharan Africa, and Asia. The direct effects of pathogens on florists’ crops are the extensive losses that can occur in their production and postharvest handling. Furthermore, the dissemination of pathogens and their vectors on cut-flowers and potted flowering plants and propagative material may also damage unrelated crops. Management of pathogens of florists’ crops is very challenging and costly because these crops have an essentially zero pest damage threshold, and because of the increasing demand by the public and government agencies that all of agriculture, including floriculture, should adopt a sustainable approach which minimizes/eliminates its nontarget effects. Keywords

Cut-flowers • Potted flowers • Production • International trade • Plant disease • Crop losses • Pathogen spread • Pest dissemination R.J. McGovern (*) NBD Research Co. Ltd., Lampang, Thailand Department of Entomology and Plant Pathology, Chiang Mai University, Chiang Mai, Thailand e-mail: [email protected] W.H. Elmer Department of Plant Pathology and Ecology, The Connecticut Agricultural Experiment Station, New Haven, CT, USA e-mail: [email protected] # Springer International Publishing AG 2018 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florists’ Crops Diseases, Handbook of Plant Disease Management, https://doi.org/10.1007/978-3-319-39670-5_8

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R.J. McGovern and W.H. Elmer

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Global Production Trends . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Direct Disease Impact . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Indirect Disease Impact . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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2 2 5 5 7

Introduction

Flowers affect our emotions in a number of positive ways (Haviland-Jones et al. 2005). Flowers have apparently benefited humankind for millennia; the ceremonial use of flowers in burials has been documented as long as 60,000 years ago, during the Neanderthal era (Solecki 1975). Humans have actively sought positive psychological and environmental modification through flower cultivation for thousands of years (Ackerson 1967; Haviland-Jones et al. 2005). Today the production of cutflowers, potted flowering plants, and related propagative material (seeds, cuttings, bulbs, etc.) is a dynamic, fast-growing, and highly lucrative global industry. Global trade volume of florists’ crops has been estimated to be more than $100 billion annually (African Business Magazine 2012). In an industry where extremely high value criteria prevail, even a slight loss in quality can render the crop unmarketable. In Table 1, recent estimated losses caused by pathogens in representative florists’ crops range up to 100%. If we assume a conservative estimate that pathogens reduce the value of florists’ crops by 10%, it would suggest a staggering $10 billion in annual losses from floral diseases. This appears to be a very conservative estimate in that postharvest diseases alone have been reported to cause losses of 10–30% of the total florists’ crops yield worldwide (Dole and Wilkins 2004). Pathogens directly reduce the production and marketability of cut- and pottedflowers and propagative material. An indirect impact results from the dissemination of pathogens and their vectors as a result of the global movement of flowering ornamentals from offshore production facilities into new areas, which, in turn, may damage unrelated crops.

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Global Production Trends

In the past, flower production sites were primarily located in or near the major markets, primarily the developed countries in Europe and North America. As of 2012, countries in the EU led by the Netherlands still accounted for a hefty 44% of world production of flowers and potted plants (European Commission Advisory Group 2012). However, because of lower production costs, including lower labor inputs, more favorable climatic conditions, and the development of efficient transportation methods, exporting countries located closer to the equator have steadily

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Table 1 Estimated losses caused by pathogens in representative florists’ crops Crop Anthurium

Caladium

Carnation

Celosia

China aster

Chrysanthemum

Geranium

Gerbera

Gladiolus

Gypsophila Lily

Disease/Pathogen Bacterial blight/ Xanthomonas axonopodis pv. dieffenbachiae Fusarium tuber rot/Fusarium solani Sclerotinia stem rot/Sclerotinia sclerotiorum Root knot/ Meloidogyne incognita Yellows/’ Candidatus Phytoplasma spp.’ Fusarium wilt/ F. oxysporum f. sp. callistephi Tobacco rattle virus Tomato spotted wilt virus White rust/ Puccinia horiana Bacterial wilt/ Dickeya chrysanthemi Tomato spotted wilt virus, Impatiens necrotic spot virus Bacterial blight/ Xanthomonas hortorum pv. pelargonii Tomato spotted wilt virus

Country USA (Hawaii)

Loss (%)1 50–100

Reference Alvarez et al. 2006

USA (Florida)

80–100

McGovern et al. 2003

India

38.5

Vinod Kumar et al. 2015

India (Bangalore area) Israel

40–60

Parvatha Reddy 2014

up to 100

Tanne et al. 2000

USA (Connecticut, Florida) Japan

up to 70

Greece

37.7

Turkey (Izmir province) Hungary (Budapest area) Colombia

80

Elmer and McGovern 2013 Komuro et al. 1970 Chatzivassiliou et al. 2000 Göre 2008

Fusarium corm rot/ F. oxysporum f. sp. gladioli Powdery mildew/ Erysiphe buhrii Fire/Botrytis spp.

up to 63

~100

Végh et al. 2014

up to 80

Vasquez and Angarita 1999

USA

up to 100

Rosa and Moorman 2017

Serbia, Venezuela

up to 30

India

60–80

Stankovic et al. 2011; Marys et al. 2014 Chandel and Deepika 2010

Korea (Iksan City) Most lilygrowing areas

~100

Choi et al. 2014

up to 100

Chastagner et al. 2017 (continued)

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Table 1 (continued) Crop Lisianthus

Orchid

Rose

Snapdragon

Multiple crops (Gerbera, Lisianthus, Rose)

Disease/Pathogen Fusarium crown and stem rot/F. avenaceum Bacterial brown spot/Burkholderia gladioli Foliar nematodes/ Aphelenchoides besseyi and Aphelenchoides fragariae Crown rot/Phytophthora nicotianae Crown gall/ Agrobacterium tumefasciens Gray mold/Botrytis cinerea Downy mildew/ Peronospora antirrhini Phytophthora stem rot and wilt/ Phytophthora spp. Gray mold/Botrytis cinerea

Country USA (Florida)

Loss (%)1 up to 70

Reference McGovern et al. 1997

China

10–25

You et al. 2016

USA (Hawaii)

up to 90–95

Kawate and Sewake 2014

Argentina

10–25

Wolcan et al. 2007

Kenya

5–60

Real IPM/CSL 2007

Ethiopia (East Shoa Zone) USA

up to 31 (postharvest) up to 100

Aman 2014

USA

50

Gill 1960

The Netherlands Flower Exchange

15–30 (postharvest)

Vrind 2005

Yarwood 1947

1

Loss estimates refer to those observed in production unless stated otherwise.

increased their market share in cut-flowers, starting in the 1970s with Colombia and Ecuador, and more recently in sub-Saharan African countries including Kenya and Ethiopia, and Asian countries such as Malaysia (van Rijswick 2015). Although the export value of cut-flowers from the Netherlands rose by 2% to an industry-leading $7.2 billion in 2011, the number of Dutch exporters fell sharply to 724 compared to 857 in 2008 (ITC 2012). In 2013, Colombia exported $1.34 billion worth of flowers (PMA 2015), and Ecuador exported $786 million (OEC 2013). Between 2000 and 2007, cut-flower exports from sub-Saharan Africa expanded by an average annual rate of 20% (WTO 2008). Africa’s share of world exports doubled to 8% during this period. Kenya, Africa’s leading exporter of cut-flowers, is the fourth-largest world supplier (behind the Netherlands, Colombia, and Ecuador) and the largest outside supplier to EU countries. The export value of Kenyan ornamentals (live trees, plants, bulbs, roots,

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cut-flowers, etc.) was more than $720 million in 2013 (ITC 2013). From 2003 to 2007, exports of cut-flowers from Ethiopia increased by an average annual rate of 142%, while Zambia’s exports increased at an annual rate of 44% between 2000 and 2006 (WTO 2008). With a compounded annual growth rate of 30%, the value of the Indian floriculture sector was projected to exceed $140 million by 2015 (Bahri 2012). Another Asian giant, China, is rapidly becoming one of the largest flower growing and consuming countries in the world with an annual production of 2.7 billion fresh flowers and sales worth about $260 million (Canada-China Agriculture and Food Development Exchange Centre 2015). Interestingly, floriculture is also an important source of food security in developing countries because of the income it brings to thousands of workers – most of whom are women (FAO 2002). Although not a major exporter, the wholesale value of USA-produced floricultural crops remains very substantial, being estimated at $2.13 billion in 2012, up 1% from 2011 (PMA 2012).

3

Direct Disease Impact

Because of the inherent esthetics involved, florists’ crops have an essentially zero pest damage threshold including even slight flower blemishes caused by pathogens. Noticeable residue from fungicides and other materials is also unacceptable. Disease management is additionally challenging because of the increasing demand by the public and government agencies that all of agriculture, including floriculture, should adopt a sustainable approach which minimizes/eliminates its nontarget effects. Therefore, along with other flower production practices, disease management must be equally precise, intensive, and sustainable. The direct effects of pathogens on florists’ crops are the losses that can occur in their production and postharvest handling. There have been many reports of extensive losses caused by pathogens in florists’ crops. Table 1 presents recent disease loss estimate percentages for a number of these plants. However, estimates assigning a monetary value to such losses in these high value crops have been rare. The University of Georgia annually reports the financial impact of disease on the state’s crops. In 2013, total losses due to diseases of floricultural crops in the state of Georgia, USA, were estimated to be about $42 million, or approximately 17% of the 2010 farm gate value for these crops (Martinez-Espinosa 2013).

4

Indirect Disease Impact

Increasing globalization of trade can rapidly introduce new ornamental pathogens and their biotypes. In addition, the dissemination of pathogens and their vectors on cut-flowers, potted flowering plants, and propagative material may also damage unrelated crops. Ralstonia solanacearum is a major bacterial pathogen of many horticultural crops. Race 3 biovar 2 (R3bv2) of the bacterium is considered to be

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particularly damaging because of its ability to cause rot and wilt in eggplant, geranium, potato, and tomato, as well as persist in areas with temperate climates. The bacterium is considered a serious quarantine pest by national regulatory agencies like the USDA Animal and Plant Health Inspection Service (APHIS), and multinational regulatory organizations such as the North American Plant Protection Organization (NAPPO), and the European Plant Protection Organization (EPPO). In 2003 and 2004, R3bv2 was detected in geraniums that were imported as cuttings from Kenya and Guatemala and grown in greenhouses in Canada and the USA (USDA-CSREES 2004). The costly eradication measures mandated for the pathogen included quarantine of the affected greenhouses and total crop destruction. In addition, movement of pesticide-resistant pathogens in global agricultural trade is an important issue. The pathogenic fungus Botrytis cinerea has acquired fungicide resistance due to the use of single-site mode of action fungicides to suppress Botrytis blight without rotation. The dissemination of fungicide resistant strains of Botrytis and other fungal pathogens in imported flowers and related materials to geographically distinct regions may compromise local production of vegetables, small fruit, and other crops. The movement of viruses and other graft-transmissible agents in ornamentals has been a long-standing regulatory concern (Lawson and Hsu 2006). Preventing the dissemination of virus vectors is also of critical importance. The Bemisia tabaci whitefly complex is viewed as one of the most damaging agricultural pest groups because of the many viruses that it vectors and its large host range which includes many florists’ crops (Lapidot and Polston 2007; Oliveira et al. 2001). It has been hypothesized that the B biotype of B. tabaci (aka silverleaf whitefly), which originated in the Middle East, was moved to the Western Hemisphere on ornamental plants (CABI 2015). Poinsettia infested with the B biotype may have further spread the pest within the USA (Parella et al. 1992). The Q biotype of B. tabaci, which is reported to be resistant to commonly used neonicotinoid insecticides, was recently spread from the Mediterranean region to China on ornamentals (Chu et al. 2006). Regulatory organizations like the EPPO, NAPPO, USDA/APHIS, etc. maintain lists of quarantinable pathogens and other plant pests and their hosts. The USDA/ APHIS also operates a remote program, the Offshore Pest Information System (OPIS), to gather information about new pest outbreaks of potential concern to US agriculture. Furthermore, the USDA/APHIS conducts preclearance of a number of crops in cooperation with export countries to help reduce the spread of quarantinable pathogens and other pests. In addition, there are joint industry-government certification programs to insure that plant propagative material is free of pathogens and other pests. In the Netherlands such a program, the Naktuinbouw Elite certification system (http://www. naktuinbouw.eu/node/192), combines pathogen-free propagation methods with compulsory pathogen testing.

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References Ackerson C (1967) Development of the chrysanthemum in China. Natl Chrysanth Soc Bull 23 (4):146–155 African Business Magazine (2012) The global flower trade. African Business Magazine. 19 Feb 2012. http://africanbusinessmagazine.com/special-reports/sector-reports/floriculture/the-globalflower-trade. Accessed 19 Apr 2016 Alvarez AM, Toves PJ, Vowell TS (2006) Bacterial blight of Anthuriums: Hawaii’s experience with a global disease. Online. APSnet features. 10.1094/APSnetFeature-2006-0206 Aman M (2014) Postharvest loss estimation of cut rose (Rosa hybrida) flower farms: economic analysis in East Shoa zone, Ethiopia. Int J Sustain Econ 6(1):82–95. http://www. inderscienceonline.com/doi/abs/10.1504/IJSE.2014.058518. Accessed 19 Apr 2016 Bahri C (2012) Flower power. IndiaNow Oct–Nov 2012, p 36 http://www.ibef.org/IndiaNow Magazine_e-Versions/Vol-3-Issue-4/files/assets/downloads/page0038.pdf. Accessed 19 Apr 2016 CABI (2015) Bemisia tabaci (B biotype) (silverleaf whitefly) Invasive Species Compendium, Data Sheet 8925. http://www.cabi.org/isc/datasheet/8925. Accessed 19 Apr 2016 Canada-China Agriculture and Food Development Exchange Centre (2015) China’s flower industry in full bloom. http://www.ccagr.com/content/view/31/113/. Accessed 19 Apr 2016 Chandel S, Deepika R (2010) Recent advances in management and control of fusarium yellows in gladiolus species. J Fruit and Ornamental Plant Res 18(2):361–380. http://www.insad.pl/files/ journal_pdf/journal_2010_2/full34%202010(2).pdf. Accessed 19 Apr 2016 Chastagner GA, van Tuyl JM, Verbeek M, Miller B, Westerdahl BB (2017) Diseases of lily. In: RJ MG, Elmer WH (eds) Handbook of florists’ crops diseases. Springer, Dordrecht Chatzivassiliou EK, Livieratos I, Jenser G, Katis NI (2000) Ornamental plants and thrips populations associated with Tomato spotted wilt virus in Greece. Phytoparasitica 28:257–264 Choi IY, Kim BS, Cho SE, Park JH, Shin HD (2014) First report of powdery mildew caused by Erysiphe buhrii on Gypsophila paniculata in Korea. Plant Dis 98(7):1013 Chu D, Zhang Y-J, Brown JK, Cong B, Xu B-Y, Wu Q-J, Zhu GR (2006) The introduction of the exotic Q biotype of Bemisia tabaci from the Mediterranean region into China on ornamental crops. Fla Entomol 89(2):168–174. doi:10.1653/0015-4040(2006)89[168:TIOTEQ]2.0.CO;2 Dole JM, Wilkins HF (2004) Floriculture: principles and species. Pearson Prentice Hall, New Jersey Elmer WH, McGovern RJ (2013) Epidemiology and management of Fusarium wilt of China asters. Plant Dis 97:530–536. http://apsjournals.apsnet.org/doi/pdf/10.1094/PDIS-05-12-0445-RE. Accessed 19 Apr 2016 European Commission Advisory Group (2012) Flowers and ornamental plants (working document) AGRI-C2. http://ec.europa.eu/agriculture/fruit-and-vegetables/product-reports/flowers/statis tics-2012_en.pdf. Accessed 24 May 2016 FAO (2002) A thorn on every rose for Kenya’s flower industry. http://www.fao.org/english/news room/news/2002/3789-en.html. Accessed 19 Apr 2016 Gill DL (1960) A stem and branch rot of snapdragon. Plant Dis Rep 44(12):946–947 Göre ME (2008) White rust outbreaks on chrysanthemum caused by Puccinia horiana in Turkey. Plant Pathol 57:786. doi:10.1111/j.1365-3059.2007.01796.x. http://onlinelibrary.wiley.com/ doi/10.1111/j.1365-3059.2007.01796.x/pdf. Accessed 19 Apr 2016 Haviland-Jones J, Hale Rosario H, Wilson P, TR MG (2005) An environmental approach to positive emotion: flowers. Evol Psychol 3:104–132. http://evp.sagepub.com/content/3/1/ 147470490500300109.full.pdf. Accessed 19 Apr 2016 International Trade Centre (ITC) (2012) Cut flowers and ornamental plants. Marketing News Service Bulletin No. MO2 Feb 3, 2012. http://www.intracen.org/uploadedFiles/intracenorg/ Content/Exporters/Market_Data_and_Information/Market_Insider/Cut_Flowers_and_Orna mental_Plants/Floriculture_Monthly_M02_12.pdf. Accessed 19 Apr 2016

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International Trade Centre (ITC) (2013) Exporters, floriculture, Kenya. http://2013.intracen.org/ country/kenya/. Accessed 19 Apr 2016 Kawate M, Sewake KT (2014) In: Tarutani K (ed) Pest management strategic plan for potted orchid production in Hawaii. University of Hawaii, Manoa, 143 pp. http://www.ipmcenters.org/pmsp/ pdf/HI_orchid_PMSP.pdf. Accessed 9 May 2016 Komuro Y, Yoshino M, Ichinohe M (1970) Tobacco rattle virus isolated from aster showing ringspot syndrome and its transmission by Trichodorus minor. Colbran Ann Phytopath Soc Japan 36(1):17–26 (in Japanese) Lapidot M, Polston JE (2007) Biology and epidemiology of bemisia-vectored viruses. In: Czosnek H (ed) Tomato yellow leaf curl: virus disease management, molecular biology, breeding for resistance. Springer, Dordrecht, pp 227–231 Lawson RH, Hsu H-T (2006) Quarantine viruses, viroids and phytoplasmas that affect movement of ornamental plants. Acta Hortic 722:17–30. doi:10.17660/ActaHortic.2006.722.1 Martinez-Espinosa A (2013) Plant disease loss estimates – 2013. UGA Extension AP 102–6, 22 pp. http://extension.uga.edu/publications/detail.cfm?number=AP102-6. Accessed 19 Apr 2016 Marys E, Mejías A, Rodríguez-Román E, Avilán D, Hurtado T, Fernández A, Zambrano K, Garrido M, Brito M (2014) The first report of tomato spotted wilt virus on Gerbera and Chrysanthemun in Venezuela. Plant Dis 98:1161–1161 McGovern RJ, Harbaugh BK, Polston JE (1997) Severe outbreaks of Fusarium crown and stem rot of lisianthus in Florida. Phytopathology 87:S64. (Abstract) McGovern RJ, Elmer WH, Harbaugh BK, Geiser DC (2003) Biology, epidemiology and integrated management of Fusarium tuber rot of caladium. American Floral Endowment Special Research Report #110: Disease Management. http://endowment.org/wp-content/uploads/2014/03/110dm. pdf. Accessed 19 Apr 2016 Observatory of Economic Complexity (OEC) (2013) Ecuador. http://atlas.media.mit.edu/en/profile/ country/ecu/. Accessed 19 Apr 2016 Oliveira MRV, Henneberry TJ, Anderson P (2001) History, current status, and collaborative research projects for Bemisia tabaci. Publications from USDA-ARS / UNL Faculty. Paper 352. http://digitalcommons.unl.edu/usdaarsfacpub/352/. Accessed 19 Apr 2016 Parella MP, Bellows TS, Gill RJ, Brown JK, Heinz KM (1992) Sweetpotato whitefly: prospects for biological control. Calif Agric 46:25–26 Parvatha Reddy P (2014) Ornamental crops. In: Plant growth promoting rhizobacteria for horticultural crop protection. Springer, Dordrecht, pp 267–274 Produce Marketing Association (PMA) (2012) New trends are transforming the European Marketplace. Fresh Magazine Apr 2014, pp 12–13. http://www.nxtbook.com/nxtbooks/pma/fresh_ 201404/index.php?startid=12#/14. Accessed 19 Apr 2016 Produce Marketing Association (PMA) (2015) Colombia floral industry overview. Mar 2015. http:// www.pma.com/~/media/pma-files/research-and-development/colombia-floral-market-final.pdf? la=en. Accessed 19 Apr 2016 Real IPM/CSL. (2007) Crown gall workshop proceedings. 3&5th Oct 2007. Cited in: Maina G, Mutitu EW, Ngaruiya PN (2011) The Impact of Agrobacterium tumefaciens and other soil borne diseases on productivity of roses in East African region. http://erepository.uonbi.ac.ke/ bitstream/handle/11295/34815/2.ESA_.D02_KFC_Seminar_Issue_-_Agrobacterium.pdf? sequence=1. Accessed 19 Apr 2016 Rosa C, Moorman GW (2017) Diseases of geranium (Pelargonium). In: RJ MG, Elmer WH (eds) Handbook of florists’ crops diseases. Springer, Dordrecht Solecki RS (1975) Shanidar IV: a neanderthal flower burial in northern Iraq. Science 190 (4217):880–881 Stankovic I, Bulajic A, Vucurovic A, Ristic D, Jovic J, Krstic B (2011) First report of tomato spotted wilt virus on Gerbera Hybrida in Serbia. Plant Dis 95(2):226–226 Tanne E, Kuznetsova L, Cohen J, Alexandrova S, Gera A (2000) Phytoplasmas as causal agents of celosia disease in Israel. Hortscience 35:1103–1106

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USDA (2004) National pest alert: Ralstonia solnacearum race 3 biovar 2. USDA–CSRESS Integrated Pest Management Centers in cooperation with NPDN, APHIS and ARS. https://www. aphis.usda.gov/plant_health/plant_pest_info/ralstonia/downloads/ralstoniapestalert.pdf. Accessed 19 Apr 2016 van Rijswick C (2015) World floriculture map 2015: gearing up for stronger competition. Rabobank Industry Note #475. https://www.rabobank.com/en/images/World_Floriculture_Map_2015_ vanRijswick_Jan2015.pdf. Accessed 19 Apr 2016 Vasquez S, Angarita A (1999) Production of a polyclonal antiserum against two tospoviruses of chrysanthemum (Dendranthema grandiflora): tomato spotted wilt virus (TSWV) and impatiens necrotic spot virus (INSV). Acta Hortic 482:203–208 Végh A, Zs N, Salamon P, Mándoki Z, Palkovics L (2014) First report of bacterial wilt on chrysanthemum caused by Dickeya chrysanthemi (syn. Erwinia chrysanthemi) in Hungary. Plant Dis 98(7):998. doi:10.1094/PDIS-09-13-0948-PDN Vinod Kumar S, Rajeshkumar P, Senthilraja C, Nakkeeran S, Fernando WGD (2015) First report of Sclerotinia sclerotiorum causing stem rot of carnation (Dianthus caryophyllus) in India. Plant Dis 99(9):1280. doi:10.1094/PDIS-02-15-0240-PDN Vrind TA (2005) The Botrytis problem in figures. Acta Hortic 669:99–102. http://www.actahort. org/books/669/669_11.htm. Accessed 19 Apr 2016 Wolcan SM, Ronco L, Lori GA (2007) Basal rots of Gypsophila paniculata (Caryophylaceae) in Argentina: causal agents and their potential pathogenicity on Dianthus caryophyllus (Caryophylaceae) (in Spanish). Bol Soc Argent Bot 42:159–167 World Trade Organization (WTO) (2008) II. Merchandise trade by product. https://www.wto.org/ english/res_e/statis_e/its2008_e/its08_merch_trade_product_e.pdf. Accessed 19 Apr 2016 You Y, Lü FB, Zhong RH, Chen HM, Li HP, Liu JM, Zhang JH (2016) First report of bacterial brown spot in Phalaenopsis spp: caused by Burkholderia gladioli in China. Plant Dis. doi:10.1094/PDIS-09-15-0963-PDN Yarwood CE (1947) Snapdragon downy mildew. Hilgardia 17:241–250

Part I Components of Integrated Disease Management

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Fundamentals and Advances in Plant Problem Diagnostics Tim Schubert, Ayyamperumal Jeyaprakash, and Carrie Harmon

Abstract

Plant disease/problem diagnostics and pathogen detection are fundamental components of successful agriculture. Any corrective action for poor plant performance should arise out of thorough assessment and diagnosis of the problem(s), to be followed by appropriate management suggestions. Among the factors to consider in the choice of diagnostic techniques are speed, cost, accuracy, availability of technology, training, value of the crop, and likelihood of generating a successful management strategy. Many exciting new technologies are entering the field of diagnostics, and plant disease/problem diagnostics and pathogen detection are not exceptions. Among the favorites are molecular-based tests that rely on the specificity of gene sequences, serological tests in various formats, direct and indirect biosensor technologies, and spectral imaging. We have entered an era in which the entire biome both inside and outside a plant can be identified, diseased plants can be analyzed non-destructively, and the overall health of an entire crop is quickly discernible. Now we must learn how to interpret all that data and put it to practical use. In spite of the enormous potential of all these new developments, successful disease diagnosis will still depend fundamentally on basic skills that have been honed and perfected in practice for more than a century. Without the foundational information from thorough and focused observation on-site and access to all horticultural information and inputs for the crop in question, no amount of advanced technology can reliably correct for deficiencies in the primary steps in diagnosis.

T. Schubert (*) • A. Jeyaprakash (*) Division of Plant Industry, Florida Department of Agriculture and Consumer Service, Gainesville, FL, USA e-mail: [email protected]; Timothy.Schubert@freshfromflorida.com; ayyamperumal. jeyaprakash@freshfromflorida.com C. Harmon (*) Department of Plant Pathology, University of Florida, Gainesville, FL, USA e-mail: clharmon@ufl.edu # Springer International Publishing AG 2018 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florists’ Crops Diseases, Handbook of Plant Disease Management, https://doi.org/10.1007/978-3-319-39670-5_1

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Keywords

Plant disease diagnosis • Plant problem diagnosis • Triage • Microscopy • ELISA • PCR

Contents 1 2 3 4

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . General Laboratory Diagnostic Techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Special Features Related to Ornamental Disease Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . An Argument for Continued Use of Fundamental Diagnostic Techniques . . . . . . . . . . . . . . . . 4.1 Attention to Details . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Discerning Indications of Abiotic Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Many Advanced Techniques Yield Only Binary Answers . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4 Interpretation Challenges with Genetic Sequencing Data . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5 The Complications of Host Predisposition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6 Balancing Diagnostic Costs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7 Diagnosticians Are the Major Ambassadors of Phytopathology . . . . . . . . . . . . . . . . . . . . 5 The Challenges of Abiotic Disease Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 A Universal Diagnostic Process . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1 Step One: Assemble a Comprehensive Syndrome Description . . . . . . . . . . . . . . . . . . . . . . 6.2 Step Two: Check for Spatial Patterns . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.3 Step Three: Establish the Disease Chronology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.4 Step Four: Assemble a Comprehensive History of the Horticultural Treatment of the Crop . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.5 Step Five: Consult Literature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.6 Step Six: Begin Laboratory Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.7 Step Seven: Consolidate Information . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 A Special Diagnostic Category: Indexing Plant Propagative Material for Cryptic Pathogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 Diagnosis of Biotic Diseases: Direct Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.1 Microscopic Methods for Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2 Microbiological Methods for Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.3 Serological Methods for Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.4 Metabolic Tests for Pathogen Identification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.5 Molecular Methods for Pathogen Detection and Identification . . . . . . . . . . . . . . . . . . . . . . 9 Indirect Methods of Plant Pathogen or Disease Detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.1 Detection of Volatile Organic Compounds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.2 Spectral Imaging of Host Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 Accurate and Timely Reporting of the Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Introduction

When a crop fails to perform up to expectations, it is logical to turn to a diagnostic process to determine what is preventing the attainment of full potential. This chapter will explain the full spectrum of options for determining what might be limiting the growth of a particular plant while considering various factors that influence which tools are most suitable for the particular diagnosis in question. In an era of rapidly advancing modern technologies promising extraordinary accuracy and precision

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heretofore practically unimaginable, the wise deployment of the entire spectrum of available tools maximizes the accuracy and usefulness of the diagnosis for the client and the grower community with a minimum of expense in a reasonable amount of time. There are three caveats at the beginning: Firstly, for this discussion, we use two different terms or phrases to describe the process of figuring out what is wrong with a plant – plant problem diagnosis and plant disease diagnosis. It is not necessary for the reader to accept any strict definitions here, as the two can have nearly the same meaning, and that is how we intend to use them. We draw attention to both terms to make certain that the reader understands that we are including some techniques for the diagnosis of plant problems that have no biotic (living) causal agent, i.e., they are due to some abiotic (nonliving) cause. Some consider plant disorders caused by abiotic influences something other than disease, with the term “disease” restricted to plant disorders caused by living causal agents (giving some latitude for the viruses and viroids and perhaps prions which are incapable of life functions outside a host cell). Others classify these disorders as noninfectious diseases or physiological diseases. In this abiotic category are factors such as excesses or deficiencies of nutrients, water, or light; poor-quality water, soil/media, or air in the growing environment; temperatures outside optimal ranges for plant growth; mechanical damage; etc. Though some may prefer to make a distinction, we will be using plant problem diagnosis and plant disease diagnosis as essentially the same thing. Secondly, a further clarification of terms is helpful in our quest for plant health. Plant disease or plant problem diagnosis represents the final step or culmination of a process that is preceded first by plant disease surveillance, followed by plant disease or plant pathogen detection. The semantics here are important. A grower must engage in continual surveillance to monitor overall plant health. This answers the question, “Is the crop normal? Is it on schedule? Is there a problem?” Plant disease or plant pathogen detection is the process of following up on the findings of surveillance. It addresses the question, “Is there a problem?” by defining the nature of the problem found, and discovers what organisms or causal agents are possibly associated with the syndrome detected. Finally, plant disease or problem diagnosis is the term reserved for the process of assigning the blame for the problem detected in the surveillance. The techniques and technologies discussed in this chapter can help in one or all of these three steps (Stack and Fletcher 2007; Miller et al. 2009). Thirdly, many clients of plant disease diagnostic services automatically assume that the diagnosis takes place entirely or mostly in the lab using esoteric methods, specialized knowledge, and complicated machinery. Actually, overlooking the field aspects of the diagnosis can derail the entire process and yield faulty conclusions. Though we will start our discussion on lab techniques, at several points through the chapter, we will remind the reader of what must be accomplished in the field with the goal of reinforcing the idea that good diagnostics requires more than modern equipment and skilled operators. The indispensable detective work in the field at the time the sample is taken sets the stage for an accurate and holistic diagnosis that leads to optimal management decisions. Furthermore, very often the sample-taker

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does not participate in the lab work and vice versa, so good communication networks between all the parties (the client, the sample-taker, and the lab scientist) are vital.

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General Laboratory Diagnostic Techniques

In general, the techniques normally restricted to the laboratory and used for plant disease or plant problem diagnosis fall into these four major categories: • Microscopy – examining the afflicted plant’s tissues at various magnifications to visualize symptoms and evidence of any pathogen(s). This would include a hand lens, dissecting (reflected light) and compound (transmitted light) microscopy, and scanning and transmission electron microscopies. • Microbiology – using various culture methods to separate the pathogen from diseased host tissue to study it in isolation for identifying features. If a pathogen can be isolated from its host and grown in culture or, in the case of viruses and viroids, purified by various methods, proof of pathogenicity by reinoculation is possible using Koch’s postulates. Failure to recover or witness any organisms associated with the syndrome would push the diagnosis in the direction of an abiotic causal agent. • Serology – using antisera developed against specific antigens from suspect pathogens to identify the causal agent in whatever matrix it may be found. • Molecular biology – using nucleic acid visualization and sequencing to find identifying sequences in DNA or RNA extracts from isolated suspect pathogens or taken directly from samples of diseased tissues suspected of containing the pathogen. This technology has the sensitivity to detect latent disease and inform management decisions proactively (Michailides et al. 2005). These categories are what many clients in need of diagnostic services believe to be the major or even sole methods employed. Each is in a typical sequence of steps and in relative order of cost to perform, and each requires special equipment and training to be successful in their use and interpretation. As technology advances, some of the work formerly confined to the lab is being “field hardened” to permit quick deployment on-site (Boonham 2014). Examples are more sophisticated field microscopes, dipstick and lateral flow devices for serological detection of microbes, and field PCR units. These methods can be used separately or in combination to reach the desired level of specificity in the diagnosis. However, it is extremely important to remember that for the best diagnosis, much of the successful process takes place in the field, prior to sample processing in the lab. While still in the field at the time of sample taking, a diligent effort must be made to acquire background information about the problem. Further questions may arise during lab processing that will make it necessary to revisit the site, resample, discuss issues again with growers, etc. These frequently underappreciated, overlooked, or poorly performed primary aspects of the diagnostic process are covered in greater detail in Sect. 6.

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Special Features Related to Ornamental Disease Diagnosis

At the outset, it is important to note that plant problems in ornamental crops as opposed to food, fuel, or fiber crops present some unique features that need special consideration in the diagnostic context. First, the value of ornamental plants is aesthetic – based upon appearance. Perfection (or near perfection) in appearance can make a huge difference in profitability of the enterprise, so much so that addressing any factor(s) that prevents attainment of that aesthetic standard could require the utmost in diagnostic expertise and technology. Plus, specifically ornamental floral crops are by their very nature more fragile than even other ornamental plants such as foliage or woody ornamentals. Conversely, cosmetic imperfections that blemish the non-harvested portions of food, fuel, or fiber crops (or even the harvested portions to a degree) can be tolerated with little or no impact on yield or product value. (In making note of this fact, we must at the same time admit much room for improvement in consumer tolerance of minor blemishes in food products in order to feed the growing masses and still provide a fair economic return to the grower.) Calculating the amount of pest damage to tolerate on any crop with no loss in value is fundamental to good stewardship and maximizing profits. So, in the final analysis, that allowance for imperfections while remaining profitable tends to be much smaller for the ornamental crop, especially floral products. Furthermore, commerce involving the propagative material of ornamental plants often comes with a much greater plant pest regulatory risk than for food, fiber, and fuel crops because the ornamental commodity is frequently destined not for some devitalizing use such as consumption as food, manufacture into an inanimate object, or burned as fuel. Instead, it is planted in a new location for a continued existence, complete with all the pests and pathogens that accompanied it on its original journey. The default “apparently free of pests and diseases” standard for most routine interstate and international phytosanitary certification leaves considerable latitude for cryptic organisms to accompany and disperse from propagative material. This added risk potential makes high-level diagnostics especially appropriate in the ornamental plant arena. And lastly, high-value ornamental crops as a rule receive much more attention in the way of horticultural inputs, each one designed to maximize quality production at minimum cost in the shortest amount of time. Crops “pushed hard” in this manner are more apt to suffer abiotic disorders from such things as excesses in fertilizer or growth regulators, phytotoxic pesticide applications and/or mechanical injury from repotting and handling for merchandizing.

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An Argument for Continued Use of Fundamental Diagnostic Techniques

Before addressing what might be considered by many to be the main topic in this chapter (that of advancements in biotic disease diagnostic techniques), we consider it entirely fitting to present a strong case for continued employment of more fundamental standard diagnostic methods. For the most part, few diagnostic tests are

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necessarily outmoded or automatically unsuitable with the passage of time. Especially neglected in this category are macroscopic and microscopic diagnostic approaches. Several reasons support this position, and these are addressed singly and in order.

4.1

Attention to Details

First, one’s initial approach to a diagnostic puzzle should commence with fundamental and historically proven tactics which depend upon keen attention to all aspects of the disease syndrome. Unfortunately, in every aspect of modern life, forces work against a person’s ability to simply give attention, to focus intently on a subject, and to notice pertinent details (Crawford 2015). It is at this initial stage where disciplined and polished observational skills and inquiries pay great dividends and start the diagnostic quest in the right direction. Furthermore, in most cases, the sample collection process is performed by someone other than the diagnostician, so good collaboration and communication with the sample collector is critical. It is no exaggeration to assert that the sample collector is an absolutely essential participant in the diagnostic process. Every available means of modern communication (both real time and recorded) should be made available and used to help the lab personnel grasp the disease presentation in the field and guide the sample collection process to full advantage. Done properly, much information will be gathered in these early stages. Much of it will prove unhelpful and tangential to the final diagnosis, but there is no substitute for a thorough job at this stage. The essential clues will be distilled from that voluminous assemblage of seemingly random information. A sloppy job here will frequently alter the course of the entire diagnosis, and advanced techniques have little or no inherent value to correct the situation.

4.2

Discerning Indications of Abiotic Disease

Second, it is important to remember that abiotic causes are responsible for about half the samples submitted to a typical plant disease diagnostic clinic. The fact that ornamental plants are subjected to a high degree of deliberate manipulation by caretakers translates into a quite reasonable expectation that abiotic diseases might be even more common here. Assignment of an abiotic cause is, of course, based on our current fragmentary understanding of what can cause various symptoms in plants, but it would be unwise to assume that our understanding does not remain fragmentary even with advanced diagnostic tools at our disposal. Abiotic diseases seldom leave signs of the causal agent in the classic sense (a recognizable part of the causal agent left behind at the scene), as is more often the case with a pathogen/biotic disease. Only symptoms remain, and symptoms are notoriously general with multiple paths of causation. Genomic tools are not as valuable under such abiotic disease circumstances, though one day, transcriptome (RNA molecules present) or

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metabolomics (chemical fingerprints) characterization technologies might prove helpful in understanding how a syndrome developed, and that could yield clues as to what abiotic condition may be responsible (Adams 2013).

4.3

Many Advanced Techniques Yield Only Binary Answers

Third, many (not all) of the more advanced diagnostic techniques are limited to providing only yes/no answers inasmuch as they can only detect a single organism. These are more accurately deemed detection tools that contribute to a diagnosis and do not constitute a diagnosis in and of themselves. This is especially true of PCR techniques and also serological tests to a lesser degree. They are more suited and useful for diagnostic confirmation than for diagnostic exploration.

4.4

Interpretation Challenges with Genetic Sequencing Data

Fourth, more advanced nucleic acid sequencing techniques and metagenomics have enormous potential to reveal genetic clues about all the organisms present in a sample of diseased plant tissue. One must be careful in the interpretation of the data procured with sequencing methods however. Our understanding of apparently innocuous or beneficial endophytic and epiphytic organisms associated with plants is too rudimentary to jump to conclusions about causality for a plant disease syndrome. Furthermore, associating gene sequences of an organism and its ability to cause disease is still a loose and tenuous relationship for many pathogen (e.g., Fusarium oxysporum). Many surprises await us, and understanding the significance of the biome and microbiome associated with a diseased plant will probably be a long learning curve. The more traditional diagnostic skills of isolating a pathogen from diseased tissues offer a cleaner substrate to take to the lab for nucleic acid sequencing, but limitations in what can be successfully isolated from diseased tissues have been known at some level for a long time. The automatic bias against fastidious microbes as pathogens is something the diagnostician must consciously avoid. The more we learn about the plant biome/microbiome, the more we understand about the limitations of culturing or otherwise separating microbes from diseased plant tissue. Even though these limitations are greater than we may have imagined, it is still a good idea to attempt to isolate the suspected causal agent from the host tissues because the clarity of genomic analyses of the clean isolate is far greater than an extract taken directly from diseased plant tissues.

4.5

The Complications of Host Predisposition

Fifth, as diagnostic skills advance, it is becoming more apparent that, under certain (perhaps many) circumstances, predisposing abiotic events precede and may even be

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essential for biological invasions leading to plant disease development. Since plant tissues can behave something like ecosystems in miniature, it is not unreasonable to assume that abiotic predisposing events and colonization events could very well take place in a somewhat orderly succession. Although it is helpful to know the biotic agent responsible for the eventual symptoms, the most effective plant disease management techniques would recommend the avoidance or minimization of the predisposing conditions to lessen or prevent the biotic phase(s) of the disease altogether. Starting with metagenomics in such a situation would render a very complicated and incomplete diagnosis and probably generate an ineffective management scheme. And if preliminary genomic findings have any early message for growers and diagnosticians, we should set aside notions of genetic stability of pathogens and, as a consequence, discount any parallel belief that a sequencebased diagnostic method will reach perfect performance and permanence. Clearly, pathogens are much more dynamic and adaptable than once thought. Fortunately for both client and diagnostician, a 100% accurate diagnosis is not always necessary to yield a helpful (if incomplete) determination(s) and provide recommendations that improve plant health.

4.6

Balancing Diagnostic Costs

Sixth, it remains imperative that any clinical methods be commensurate with the goal of the diagnosis, triage in one sense of the word. Every sample does not call for advanced techniques. With no intent to diminish the importance of any diagnosis, clinics need to maximize the resources at their disposal to provide as much service to society as possible without limiting their capacity to respond to extraordinary circumstances with every tool possible when the need arises. Most plant problem clinics operate with some level of public support, so there is an obligation to serve any tax-paying client to the fullest extent possible and warranted. Charging a fee for diagnostic services to supplement operating budgets and recoup costs may make larger staff and more costly procedures possible, but those same fees can also interrupt vital links to the very community that can serve as front-line detectors and “citizen scientists” for early disease detection and effective intervention. These fragile links need nurturing, and diagnostic fees as such tend to be counterproductive. The balance between affordability, applicability, accuracy, and sensitivity must be decided jointly by the clinician and client while keeping all these competing factors in view. One trend that helps in this regard is that of illustrated electronic communications to initiate a diagnosis. Assuming good photographic skills on the part of the client or disease surveyor/scout, a good diagnostician should be able to recognize the routine problems and dispatch them with minimum expense and time to save resources for the major events that will require “all hands on deck” and full spectrum diagnostics. In short, when it comes to advanced techniques, discretion in deployment may be as important as capabilities.

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Diagnosticians Are the Major Ambassadors of Phytopathology

Seventh, related to the theme of how the costs of diagnostic services are met, we must further acknowledge that disease diagnostic services and the essential cooperative nature of this enterprise with the client are almost certainly the sole link between the phytopathology profession and most of the general public. That being the case, the diagnostic clinic is well advised to put its best foot forward for the reputation of the profession. Additionally, even though all life on planet Earth is based directly on plant health, the scarcity of providers of diagnostic plant pathology services contributes to the general obscurity of phytopathology in the public service arena. For comparison purposes, according to the US Bureau of Labor Statistics, the medical profession has in the neighborhood of 700,000 diagnostic physicians and surgeons and the veterinary profession about 70,000. Diagnostic phytopathologists, on the other hand, are very generously estimated to number only about 1000 nationwide based on the number of members in the American Phytopathological Society that participate in the Diagnostics Committee plus the estimated number of employees at state- and federally sponsored diagnostic labs and clinics around the country. Diagnosticians undoubtedly serve as the foremost ambassadors for the entire profession. Deciding on the method(s) to be used in a diagnostic project must take into account much more than cost and applicability but must also consider the representation of the profession in the public forum and the potential contributions to be made to the common good.

5

The Challenges of Abiotic Disease Diagnosis

In many respects, the diagnostic approach for any disease, biotic or abiotic, will follow the same pathways. Exceptions are granted when the diagnostician knows a crop extremely well and is fully aware of all the horticultural inputs and their effects. Familiarity with common biotic diseases for the crop also helps narrow the diagnostic focus. Usually, however, a thorough abiotic disease diagnosis is a meticulous process of eliminating possible biotic causal agents, then resorting to attempts to match abiotic causal agents to the symptoms observed.

6

A Universal Diagnostic Process

Stepwise, here is a rational process for any disease diagnosis. We emphasize that much of a successful and accurate diagnostic process occurs before moving into the lab. It gathers enough information to evaluate both categories, biotic and abiotic:

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Step One: Assemble a Comprehensive Syndrome Description

Begin with a careful examination of the syndrome to record all symptoms plus any signs of a pathogen with the goal of fully describing the problem. Is the presentation actually abnormal? How should the crop appear at this stage in the production cycle? At this early juncture, it is essential that detailed record keeping be initiated and practiced throughout the diagnostic process. These documents are vital for legal disputes and for trace back – trace forward investigation.

6.2

Step Two: Check for Spatial Patterns

Look for any patterns in the symptom expression on both the individual plants and within the plant population as a whole. Do not confine the exam to just the single crop, but also evaluate neighboring flora, even weeds if they are present. Examine whole plants as much as possible, internally and externally both above and below ground, and be prepared to sacrifice representatives to supply samples for further lab work.

6.3

Step Three: Establish the Disease Chronology

Determine the chronology of symptom appearance and any changes in the syndrome over time. When did the syndrome start? Was the onset sudden? Do symptoms abate, stay the same, or get worse?

6.4

Step Four: Assemble a Comprehensive History of the Horticultural Treatment of the Crop

Find out as much as possible about how the crop has been grown. Is the crop new to the grower, a new variety, growing in a different structure, or has something changed in the horticultural regimen (physical and chemical qualities of the growing media, water quality and amount, fertility, light levels and quality, temperature, growth regulator use)? Pay particular attention to the mode of delivery of any horticultural treatments. Phytotoxic agents can reach a crop in any number of ways, including by direct and deliberate application, drift, contamination of pesticide or fertilizer product or application equipment, rate miscalculations, poor application techniques, contamination of soil or surroundings, mulch or soil amendments, in irrigation water or runoff, etc.

6.5

Step Five: Consult Literature

Check references on the plant in question to learn of any unusual sensitivities, preferred growing conditions, idiosyncrasies in growth cycles and plant appearance,

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and reported diseases and disorders. APS compendia and reference manuals such as this one are good sources of information (Chase et al. 1995; Daughtrey et al. 1995; Gill et al. 2006; Gleason et al. 2009; Horst and Cloyd 2007; Horst and Nelson 1997; Horst 2013).

6.6

Step Six: Begin Laboratory Analysis

Based on the information gathered, determine what laboratory testing might be warranted to attempt confirmation. (It is here that the more technical laboratory work introduced earlier in Sect. 1 and addressed in more detail in Sect. 8 usually begins). Determine whether any biotic agents discovered using laboratory techniques and coupled with symptoms and disease chronologies are primary or secondary. Soil and tissue analysis are advised for confirmation of symptoms associated with known syndromes of nutrient deficiencies/excesses or soil pH problems. A bioassay using indicator plant species can confirm problems with suspected harmful soil residues. Testing of pesticide products for foreign components or testing of plant tissues for residues of pesticides is much more involved and expensive, usually requiring some idea of the nature of the residue sought.

6.7

Step Seven: Consolidate Information

Review all the gathered information to synthesize a likely diagnosis. The process for an abiotic disease diagnosis can eventually assemble large amounts of information because it is fundamentally a process of elimination, and much of the facts gathered may end up having little or no bearing on the final conclusions. Nevertheless, the quality of the diagnosis depends on thoroughness of the overall investigation. As stated earlier, using the most advanced diagnostic tools available cannot make up for deficiencies in steps 6.1 through 6.5.

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A Special Diagnostic Category: Indexing Plant Propagative Material for Cryptic Pathogens

Indexing for plant pathogens in propagative material is a different sort of diagnostic challenge altogether. The most sensitive detection methods available deserve top consideration, especially if the propagative material is to be moved to a new location within a country or to another country and must meet the attendant phytosanitary requirements. The potential gains in overall plant health (and thus profitability) from indexing of asexually propagated perennial plants in particular can be substantial. The rewards for indexing annual plants are different but similarly valuable because the crop customarily starts from seed each year anyway. With annuals, indexed seed is the goal. Indexed seed and stock plants are extremely valuable, though they can be quite expensive to acquire and maintain. Furthermore, these costs come up front,

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before any actualization of profits. Rewards may or may not prove to be worth the investment. Costs of the alternative (imperfect regulatory protection in place of starting with clean stock) are seldom fully considered in any cost-benefit ratio calculations; indeed such values are difficult to commodify without advanced economic skills which are outside the circle of most growers’ acquaintances. Assigning costs to any of the parties involved, especially at the outset, is also controversial, but eventually the potential for a higher profit margin and premium price for the superior product could supply ample financial returns to cover the setup and day-to-day operating expenses. Acquiring the equipment for the most advanced indexing techniques is another matter. Prices for advanced sequencing equipment and subsequent data processing are becoming more affordable with the passage of time. Grants and sharing resources collaboratively with other professions needing similar services (e.g., medical/veterinary health and ecosystem services) may be a possible solution. Still, in a free market, there will always be a niche for the cheapest items of any commodity, so we should not fool ourselves into thinking that every grower will eagerly adopt propagation from high-quality indexed stock. Regulatory oversight and public funding to protect consumers and natural resources from the consequences of substandard plant material are still necessary and advisable.

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Diagnosis of Biotic Diseases: Direct Methods

Historically, plant disease diagnosis has targeted direct detection of the organism (s) associated with a syndrome to arrive at a diagnosis. In the case of an abiotic disease, there may be no physical evidence of a causal agent to detect and obvious shortcoming. Still, direct methods can be used in a process of elimination toward an abiotic diagnosis, and some abiotic agents do indeed leave physical evidence amenable to direct methods. Lately, progress has been made in indirect methods of disease diagnosis. These are covered in Sect. 9.

8.1

Microscopic Methods for Diagnosis

The careful observation of diseased plants and tissues using various means of magnification is the oldest, potentially the quickest, lowest cost, and most accessible diagnostic method available. Unfortunately, in many respects, the more advanced microbiological, serological, and molecular methods have distracted diagnosticians from the inherent value of simple and thorough light microscopic techniques. Skill in microscopic methods definitely requires good equipment and the time and material for plenty of practice. An apprenticeship with an adept microscopist is a great advantage. The more one practices microscopy and the discriminating observation and attention skills required to spot signs of a pathogen, the more exciting discoveries appear. Some good advice when examining diseased plant tissues for clues about the pathogen: have patience and expect to discover useful information using the microscope, do not give up easily, and do not restrict your examination to just a

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few pieces of symptomatic tissue. Carefully examine all stages of the symptoms insofar as you can reconstruct a time frame for the syndrome. If you see nothing noteworthy upon initial examination, try a moist chamber incubation for a few days to encourage fungal sporulation, and then reexamine. Inducing fungal sporulation to permit microscopic examination might require many weeks and manipulation of incubation conditions in some cases. In the emerging era of identification at the molecular level, there remains a significant place for morphometric identifications based on reproductive structures (Hyde et al. 2010) using fungal identification literature (Barnett and Hunter 1998; Leslie et al. 2006; Siefert et al. 2011). Highresolution digital photographic equipment for both macroscopic and microscopic images allows the sharing and cataloguing of images with and by experts over the Internet, thereby enhancing the technique even further. Good light microscopic equipment for both reflected and transmitted light microscopy equipped with digital cameras can cost from $3,000 to $4,000 at the low end to as much as $15,000 at the high end. Electron microscopy provides even more imaging capacity which can prove especially useful for diagnosis of viral and viroid diseases. Virus and viroid particles can be fairly quickly detected in expressed sap dried down and negatively stained on a coated grid. Electron microscopes small enough for a lab bench or tabletop capable of both scanning and transmission electron microscopy can be obtained for around $175,000 to $200,000. However, the specialized training and consumables make this equipment a likely purchase only when in support of a larger program or one with an emphasis in virus diagnosis or morphologically supported taxonomy and systematics. Microscopic detection methods suffer from the sensitivity inherent in the process. It is impossible to examine large volumes of host tissues or the environment using a light microscope to find suspected targets. The sampling problem is compounded when using the electron microscope. Extraordinary observation skills, training, and experience can lessen the inefficiency to a degree.

8.2

Microbiological Methods for Diagnosis

Several excellent treatises and lab manuals for the microbiological techniques traditionally used for plant disease diagnosis are available (Burns 2009; Dhingra and Sinclair 1995; Dugan 2006; Schaad et al. 2001; Shurtleff and Averre 1997; Streets 1979;Tuite 1969; Waller et al. 2002) plus there are many methods described online at http://wiki.bugwood.org/Diagnosticians_cookbook. Bacterial differentiation to the genus level is possible with the use of a few semiselective culture media coupled with tests for oxidase, Gram reaction, anaerobic growth, and hypersensitive response. These tests are not particularly difficult, but selection of isolates to carry forward from streaked plant tissue is based on experience. Careful attention to the chemistry and recipes of the culture media will encourage success. One is reminded to use fresh solutions and young cultures for best results. A few useful additions to the diagnostician’s bench include

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semiselective media for fungi that can help to separate out oomycetes from true fungi, can encourage sporulation, and can be amended to identify fungicide resistance.

8.3

Serological Methods for Diagnosis

Serological methods rely on the specificity inherent between an antigen and antibody developed as they are naturally formed by the immune system after injecting the antigen into the bloodstream of an animal, usually a mammal. This specific recognition phenomenon can be exploited to great advantage to seek out pathogen-specific antigens in any matrix (expressed plant sap, ground tissue extracts, etc.) and coupled with stains for light and/or electron microscopy, spectrophotometry, etc. Antisera produced in this traditional fashion are polyclonal and a mixture of antibodies generated to recognize different parts of a surface protein molecule or antigen. Most antibodies are now produced using hybridoma technology in tissue cultured cells that can be kept indefinitely to produce more antisera as needed. Antisera produced via hybridoma technology are monoclonal and recognize as a specific part of the antigen. Immunodiagnostics are quite reliable depending of the selectivity and sensitivity of the antiserum. Several enzyme-linked immunosorbent assays (ELISAs) have been developed to detect pathogens, especially bacteria and viruses. However, this does require that a species-specific antibody is already available to perform ELISA. The collective commercially available catalog of ELISA assays that have been developed so far to diagnose selected viral and bacterial pathogens is limited, but the availability of commercially available positive and negative controls encourages standardization of testing and verification of results. Interpretation of results must be done with care. Sutula et al. (1986) discuss the potential pitfalls and how to avoid erroneous interpretation. Once set up, an ELISA assay can be performed rapidly, but the costs of superior antigen identification and antibody development plus signal attachment can be a constraint. Once all reagents are developed and validated, a 96-well plate ELISA assay format can be utilized to reduce cost when the test volume is high. A microplate reader will be required to perform ELISA. Some serological assays have been incorporated into a very handy single sample dipstick or a lateral flow format that can be employed even in the field when and where the samples are collected. Both types of antisera, polyclonal and monoclonal, are often used in double-antibody sandwich ELISA (DAS-ELISA) and the aforementioned dipstick or lateral flow device. Monoclonal antibodies are more expensive to produce but since they are highly specific to the pathogen antigen, can detect single strains of a virus or a bacterial species or potentially subspecies. Results between tests are highly reproducible. Polyclonal antibodies are produced as a mixture, which has the advantage of being less expensive to produce as well as detecting multiple strains of a virus or multiple species of a bacterial genus. However, they are more likely to cross-react with other proteins. The lower specificity of polyclonal antisera may have an additional advantage in that they may provide clues to help identify unknown or as-yet uncharacterized viruses.

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DAS-ELISA kits including all reagents and controls are available from many vendors globally, costing between $1–12 per test well in kit cost. This plate-based assay is scalable, making diagnosis of large numbers of samples relatively easy and decreasing cost per sample. Quick tests such as dipsticks and lateral flow devices are readily available but for a smaller number of pathogens than plate-based ELISA and cost $1–5 per test. The limits of detection using ELISA are 105 to 106 CFU/ml. Serological specificity has been adapted a step further by attaching the antibodies to a fluorescent marker that emits a visible signal when viewed under a specially equipped microscope upon binding with its antigen while illuminated with a particular wavelength of light, usually in the UV range. This immunofluorescent microscopy depends on the availability and specificity of the antiserum. Flow cytometry (or FCM), used sparingly in plant pathology, can also take advantage of immunofluorescence technologies to detect pathogen antigens in large volumes of individualized plant cells. Flow cytometry offers enormous potential for indexing plant tissues for cryptic pathogens or a genetic sequence of interest, targeting any antigen or genetic sequence for which a specific antibody can be devised and a fluorescent probe attached (D’Hondt et al. 2011). It is even possible to sort the scanned cells based on the signal detected, thus permitting further work specifically with the selected tissues. Pathogen detection and viability in environmental samples is another application of flow cytometry (Chitarra and van den Bulk 2003). The need to develop antibodies and affix a fluorochrome is unnecessary to make use of flow cytometry if the target cells have an innate signal fluorescence when illuminated by a particular wavelength of light from the laser light source in the instrument. A basic FCM instrument without sorting capabilities costs from approximately $40,000 to $100,000. The sensitivity of FCM is in the range of 104 CFU/ml.

8.4

Metabolic Tests for Pathogen Identification

Microplate-based metabolic assays such as Biolog ® have successfully guided bacterial diagnosis, and also limited fungal diagnosis of plant pathogens. This system records the ability of a given active culture to metabolize specific carbon sources; the resulting activity matrix is compared to a library of known responses for over 2000 bacteria and yeasts. A separate setup can identify a smaller number of filamentous fungi. Originally developed for assessment of bacterial mammalian pathogens, the current GenIII library includes many plant pathogens as well, and also tests for Gram reaction. However, the use of this technology to identify specific organisms may be limited to genus, with a suggestion as to the species. Additionally, this system is limited for obvious reasons to culturable organisms. But it requires little specialized expertise and can be piggy-backed onto a plate reader that can also be used for ELISA, creating some efficiency in equipment purchase and upkeep for both systems. Biolog ® can cost about $20 per sample, assuming the use of the lesscostly nonautomated equipment. Identification of bacteria and fungi via fatty acid methyl ester (FAME) analysis is exemplified by the SherlockTM Microbial ID system (MIDI). It is based on gas

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chromatography and, in disease diagnosis, has been most heavily used for identification of culturable aerobic plant pathogens. This well-plate-based assay can be fully automated to resolve the identification of a microbe and compare its profile to a large library of known organisms. As with Biolog, this system generally resolves bacteria to genus reliably and suggests species and subspecies. The MIDI system develops a dendrogram of relatedness with references in the library, as chosen by the user, and generates a similarity index to the closest match. The MIDI tests generally cost $30–50 per sample, with savings possible for increasing numbers of samples run together. Diagnosticians tend to couple data from Biolog ® or FAME tests with other sequence-based, cultural, or plant-based tests to confirm the identified organism is capable of causing the disease symptoms of the affected plant. One thing these two tests can provide that molecular detection such as PCR and sequencing cannot is the verification that the detected organism is viable. If any or all these traditional approaches fail to provide a complete and/or clear diagnosis, molecular diagnosis at DNA or RNA level (Sect. 8.5) is an appropriate next step.

8.5

Molecular Methods for Pathogen Detection and Identification

Plant pathogen identification has always been a challenging task requiring increasingly more sophisticated tools and techniques as our understanding of host-parasite relationships and pathogenicity advances. As stressed earlier, even with molecular methods, initial assessment still requires techniques of visual inspection to recognize the symptoms caused by the infection such as chlorosis, necrosis, stunting, distortion, dieback, etc. Properly targeted sampling and careful shipment of the selected plant parts or nucleic acid extracts to a diagnostic laboratory is required for a methodical analysis to identify the pathogens at genus, species, or strain level. Molecular analysis requires various pieces of equipment (water bath, refrigerator, freezer, microcentrifuge, tissue homogenizer, nucleic acid quantification equipment, PCR and real-time PCR machines, gel-documentation system, Sanger and next-generation sequencing platforms, sequence analysis software and computers), funding for the lab supplies (extraction kits, reagents, pipettes tips and tubes, etc.), and a skilled technician to do the work. Some labs charge a price for every sample submitted for analysis to cover the operational cost. Other labs cover costs using grants from grower associations and local and federal governments, thus providing diagnostic help without any additional charge to the sample submitter. Cost estimates are given under the individual subject headings.

8.5.1 DNA and RNA Extractions The starting material for DNA and RNA extraction varies depending on the pathogen. Harvested bacterial/fungal cultures grown on plates or finely ground up infected

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plant leaves, stem, or root tissues (100–200 mg) are extracted using spin-column kits or chemical purification such as CTAB methods kit to obtain genomic DNA. Elution using ultrapure, PCR-grade water (100–200 μl) is always preferred because the elution buffers provided in many extraction kits contain enzyme inhibitors which could interfere with downstream applications. The majority of the viral diseases are caused by RNA viruses, and their detection will require extracting total RNA from infected plant tissues (100–200 mg) using a kit such as a QIAGEN RNeasy Plant Mini Kit. A microcentrifuge is required to perform this task. The nucleic acids can be stored at
20 C until ready for analysis. The cost for the QIAGEN column is reasonable ($2–3). The quality of the DNA and RNA extracted must be quite good for successful downstream applications.

8.5.2 Molecular Identification The precise strategy used for identification of plant pathogens can vary depending on the organism and the information available about the pathogen. The NIH GenBank ® database (www.ncbi.nlm.nih.gov/genbank) has nucleic acid sequences for numerous well-known species, and the database is growing constantly as scientists from all over the world deposit sequences. Several million sequence deposits already exist in the GenBank, and that includes sequences from a large number of important plant pathogens. There is a good chance that the sequence information about a suspected plant pathogen could be already present in the GenBank. Sequence information for rare plant pathogens may not exist in the GenBank, so this would require using universal primers (ribosomal RNA or mitochondrial COX1) or degenerate primers (β-tubulin, actin, histone, etc.) to amplify a marker sequence by PCR and perform phylogenetic analysis to match to a closely related species for identification. Therefore, a strategy for molecular identification has to be selected carefully. 8.5.3

Fungal Species Identification

Species-Specific PCR Positive identification of plant-associated fungi is tremendously aided by modern genetic analysis methods. One must remember, however, that simply identifying a plant-associated fungus based on a genetic sequence is not strictly the equivalent of a disease diagnosis, though it can be a step in the process. Pathogenicity testing will be required to confirm the virulence of any isolates from the diseased tissues. Proving pathogenicity can be a simple or very complicated process which may need to take into account groups of organisms which work in concert in a chronological set or physiological sequence. Furthermore, environmental predisposition of the host to achieve a susceptible state can be complicated to determine and recreate. If a fungal plant pathogen is suspected to be a particular species based on morphology and culture characteristics, but a confirmation is required, then the sequences can be obtained from the GenBank for that species, and species-specific primers can be designed using CLUSTAL W sequence alignment generated by comparing to all closely related species and then selecting a unique sequence sites to design a species-specific primer. A straightforward PCR assay can be performed,

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and the size of the DNA band amplified should match the sequence information already existing in the GenBank, thus providing confirmation. Further confirmation can be derived by digesting the PCR products with a restriction enzyme and developing a banding pattern to detect unique restriction fragment length polymorphisms (PCR-RFLP), which should match the expected sizes. This approach has an added advantage, as it can be used for plant pathogens that are difficult to culture because the DNA extracted from infected plant could be used for PCR. The primers and a PCR test per sample could cost around $20. A PCR machine and an electrophoresis gel system would be required. A basic gradient block PCR thermocycler can be purchased for as little as $5000; gel electrophoresis equipment can be $12,000 or more, depending on the camera and size of the setup. Sequencing Several nuclear or mitochondrial marker sequences can be used to identify plant pathogenic and other plant-associated fungi. This strategy requires using universal primers or degenerate primers to amplify a specific nuclear or a mitochondrial marker, sequencing, and then comparing the sequences to those already existing in the GenBank using a BLAST search tool. The markers generally used for fungal taxonomy are as follows. Ribosomal RNA The ribosomal RNA region (18S-ITS1-5.8S-ITS2-28S) has been well investigated from numerous fungal pathogens. 18S rRNA, 28S rRNA, and ITS1-ITS2 spacers are the markers that are extensively investigated from fungal pathogens. The ITS1 and ITS2 spacers are known to accumulate mutations, and it is often possible to find unique species-specific sequences in this region to generate phylogenetic trees for identification or to design species-specific primers. The ITS region is multicopy, but surprisingly all the copies that are present in a fungal species turned out to be identical. This was exploited by fungal taxonomists to build a rich collection of sequence information in the GenBank database. Universal primers are available that amplify well from both known and unknown fungal cultures grown on media (Table 1) (White et al. 1990). Culturing on a plate is necessary before performing this procedure to eliminate all the other background fungi present in the diseased tissue. Other Markers Mitochondrial cytochrome oxidase 1 (COX1), the nuclear elongation factor-1 α (EF1α), β-tubulin (Tub), histone (His), etc. could be used as well, but the sequence information library is not as rich for fungal species as it is for the ribosomal sequences. Multilocus sequence analysis (MLSA) involving several different markers greatly improves fungal species identification. The PCR products can be shipped to a commercial sequencing company or university core laboratory to obtain the sequence information. The cost for sequencing PCR products or cloned inserts is steadily decreasing, around $10 per sequencing run. Multiple sequence runs as well as sequencing both forward and reverse strands would greatly improve sequence

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Table 1 Generic primers for amplifying fungal and bacterial pathogen DNA Organism Fungi

Marker 18S ITS

28S

Bacteria

16S

Primer NS1 NS8 ITS1 ITS4 ITS5 LROR LR5 LR7 27f 1495r

Sequence 50 –30 GTA GTC ATA TGC TTG TCT C TCC GCA GGT CAC CTA CGG A TCC GTA GGT GAA CCT GCG G TCC TCC GCT TAT TGA TAT GC GGA AGT AAA AGT CGT AAC AAG G ACC CGC TGA ACT TAA GC TCC TGA GGG AAA CTT CG TAC TAC CAC CAA GAT CT GAG AGT TTG ATC CTG GCT CAG CTA CGG CTA CCT TGT TAC GA

accuracy and provide a clear identification. Occasionally, diagnostic labs do have their own Sanger sequencing machine (Applied Biosystems 3130XL Genetic Analyzer), with equipment costs starting around $150,000. Real-Time PCR The conventional standard PCR procedure can be quite labor-intensive, is prone to aerosol contamination, and is good for screening only a few samples. If the sample size is large, then a real-time PCR (also called quantitative PCR or qPCR) assay, such as one based on TaqMan technology, is ideal. This does require two primers (forward and reverse) and a species-specific fluorescent probe (6FAM or TET). This technology exploits the Taq DNA polymerase exonuclease activity, which shreds the probe bound to the species-specific site releasing the fluorescent marker and producing a positive result (Ct value 16–32). The probe fails to bind other non-target species DNA due to mismatches, producing a negative result (Ct value 32–40). The Ct values 0.5%) and has been associated with improved plant

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Table 1 Concentration of essential plant elements in foliar tissuea Element Macronutrients Carbon Oxygen Hydrogen Nitrogen Potassium Calcium Sulfur Phosphorus Magnesium Micronutrients Nickel Molybdenum Copper Zinc Boron Manganese Iron Chlorine a

Chemical symbol

Concentration in dry matter (ppm or percentage)

C O H N K Ca S P Mg

45% 45% 6% 0.5–6% 0.8–8% 0.1–6% 0.1–1.5% 0.15–0.5% 0.05–1%

Ni Mo Cu Zn B Mn Fe Cl

0.05 ppm 0.1 ppm 6 ppm 20 ppm 20 ppm 50 ppm 100 ppm 100 ppm

Based on Epstein and Bloom 2005

health (Epstein 1994). Essential plant micronutrients include molybdenum (Mo), copper (Cu), zinc (Zn), boron (B), manganese (Mn), iron (Fe), nickel (Ni), and chlorine (Cl). Sodium (Na) is an essential micronutrient only for C4 and CAM plants (Brownell and Crossland 1972). Carbon (C), oxygen (O), and hydrogen (H) are also essential elements for plant growth. However, discussion of these elements is beyond the scope of this section. Further discussion will focus on the remainder of the essential elements for plant health, traditionally known as mineral elements.

3.1

Macro- and Micronutrients

In general, plants have very similar nutrient requirements. However, the small variances in nutrient needs are typically what result in nutrient disorders. Plants have specific needs on the quantity and type of nutrients required at a given time. Critical concentration is the nutrient concentration in plant tissue just below (typically 10%) the optimum level for plant growth, quality, and profit (Bates 1971). Deficiency symptoms develop when the concentration of an essential element is below the critical concentration. The optimum nutrient concentration is specific to plant genus, species, varieties, and cultivars. The typical concentration of nutrient elements is provided in Table 1. Nonetheless, it is recommended that specific

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guidelines for each crop are used as a reference when matching tissue analysis results to optimum levels. Micronutrient toxicity and deficiency are the most common nutrient disorders observed in floriculture. Micronutrients are required at such low concentrations that there is a fine line between toxic, optimum, and deficient levels. Fe, Mn, Cu, B, and Zn are more available when the pH is below 6.0 and less available at pH higher than 6.0 (Marschner 1995). Plants’ sensitivity to micronutrient level is very specific and narrow. For example, petunias are inefficient at taking up micronutrients, especially iron, from the root zone and are prone to iron deficiencies when then pH is above 6.0 (Argo and Fisher 2002). Other crops, like marigolds, are very efficient at taking up iron and manganese from the root zone and are prone to Fe and Mn toxicity. When conducting visual diagnoses, it is important to know whether the plant genus or species is sensitive to minor changes of pH or nutrients in the root zone. A general initial practice when diagnosing micronutrient disorders is to measure the pH of growing media. Nutrient deficiency is the prevalent nutrient disorder for macronutrients. Macronutrient deficiencies occur when the essential elements are below the critical concentration in the plant tissue caused by low nutrient levels in the root zone or incapacity of the plant to uptake nutrients. Toxicity caused by macronutrients is typically associated with high concentration cations (i.e., calcium, potassium). Extremely high concentrations of macronutrients result in imbalance nutrition and therefore suboptimal growth. In some instances high concentration of an element reduces uptake or availability of another element resulting in an indirect nutrient deficiency (specific examples are provided under each element in the section below).

3.2

Nutrient Function and Movement

Classification of the essential elements by function and movement provides a good understanding of plant physiology (Marschner 1995; Taiz and Zeiger 2006; Welch and Shuman 1995) and is also a strong tool to diagnose nutrient disorders (Table 2). By understanding the functions and translocation of nutrients in plants, we can discriminate between elements causing the problems by making inferences based on symptoms and the organs where the symptoms are observed. In practice, this is especially important in circumstances when there is no background information on a specific species or cultivar. For example, calcium is involved with cell structure and membrane integrity; therefore deficiencies lead to distortion of organs. Iron is involved in chlorophyll synthesis; therefore deficiencies lead to chlorosis. The functions of each element are listed in the section below and in Table 2. Nutrient translocation is the movement of nutrients in plants from source to sink via the phloem; in other words it is the movement of nutrients from one part of the plant to another (Marschner 1995; Taiz and Zeiger 2006). Translocation of nutrient elements depends on whether the element moves exclusively via xylem or also via

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Table 2 Function and translocation of essential plant elements Important functions Constituent of cell wall, involved in seed and flower development, nutrient absorption and cell division Constituent of cell wall, cell membrane structure and permeability, signaling and regulating enzyme activity Activates enzymes, osmosis, and ionic balance

Translocationa Immobile

Copper

Enzyme component and activator

Immobile

Iron

Involved in chlorophyll synthesis, nitrate reduction, and enzymatic reactions

Immobile

Magnesium

Constituent of chlorophyll and enzyme activation Enzyme activator

Moderately mobile Immobile

Nitrogen fixation and nitrate reduction Component of urease and required in N-fixing plants Constituent of organic compounds, proteins, amino acids, enzymes, coenzymes, and nucleic acids Energy acquisition, storage, and utilization and component of genetic material Enzyme activity, regulate cell turgor, sugar translocation, and stomatal opening Components of amino acids, proteins, coenzymes, and vitamins and contribute to aroma of plants

Mobile

Element Boron

Calcium

Chlorine

Manganese Molybdenum Nickel Nitrogen

Phosphorus

Potassium

Sulfur

Immobile

Mobile

Mobile Mobile

Common deficiency symptoms Distortion of tip and margins of new leaves, hard and dull growth, distorted flowers Distorted and necrotic leaf tip, leaf scorching, strap-like growth, and blossom end rot in fruits and flowers Wilting of the leaf tips, followed by chlorosis and necrosis. Plants wilt during the day and recover at night or on cloudy days. Reduction of overall plant growth and distortion and necrosis of new leaves. Interveinal chlorosis in younger leaves, advanced symptoms result in completely chlorotic leaves and necrotic margins. Interveinal chlorosis in older leaves Necrotic leaf spots and mottling and leaf scorch Interveinal chlorosis and necrosis in older leaves Not known to be a problem in floriculture crops. General chlorosis and reduction of overall plant size and development

Mobile

Slow growth, stunted plants, and dark-green leaves turn reddish

Mobile

Slow growth, marginal chlorosis, and leaf scorching

Mostly immobile

General chlorosis and growth reduction

(continued)

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Table 2 (continued) Element Zinc

Important functions Enzyme activator and involved in the formation of chlorophyll

Translocationa Mobile

Common deficiency symptoms Short internodes, small and distorted leaves, rosette growth habit. Chlorosis of older leaves and flower abscission

Based on Epstein and Bloom (2005) Evans and Sorger (1966), Gibson et al. (2007), Marschner (1995), Mengel and Kirkby (2001), and Taiz and Zeiger (2006) a Mobile elements translocate from older tissue to younger tissue; therefore, early deficiency symptoms appear on older leaves. Immobile elements do not translocate after absorption; therefore, deficiency symptoms first appear on younger leaves.

phloem. All inorganic elements move from the rhizosphere to the upper parts of the plant through the xylem via mass flow (unidirectional water movement driven by transpiration) (Taiz and Zeiger 2006). Not all elements move multidirectionally or translocate. Mobile elements are nutrients that translocate from older parts of the plant to younger tissue. If an essential element is mobile, deficiency symptoms are noticeable first on the older leaves. Immobile elements are elements that do not translocate in plants or do it slowly. If an essential element is immobile, deficiency symptoms tend to appear first on younger leaves. For example, Mg is a structural component of chlorophyll and Fe is involved in chlorophyll synthesis; consequently both Mg and Fe cause interveinal chlorosis in leaves. This pattern is developed because the chlorophyll in the vascular systems withstands deficiencies for prolonged periods (Taiz and Zeiger 2006). However, Mg is mobile and Fe is immobile; therefore early symptoms of interveinal chlorosis in older leaves are most likely caused by Mg deficiency and in younger leaves are most likely caused by Fe deficiency. The translocation ability of each essential element is listed in Table 2 and is further discussed for each element in the section below.

4

Characteristics of Nutrient Disorders by Element

In this section, we provide characterization of nutrient disorder symptoms generally observed across species. However, specific symptoms may vary by species and cultivar.

4.1

Nitrogen (N)

Function. Nitrogen is a constituent of organic compounds and all proteins, amino acids, and nucleic acids. Translocation. Nitrogen is a very mobile element in the plant. Nitrogen moves from older tissue to younger tissue. Plants uptake inorganic nitrogen in the form of nitrate (NO3 ) and ammonium (NH4+) (Marschner 1995). Ammonium is readily

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assimilated by plants to form organic compounds. Consequently, excessive ammonium can quickly accumulate in plant tissue and result in phytotoxicity, especially in cold months when growth is slow. In contrast, nitrate moves through the xylem and can be stored in the vacuoles of roots, shoots, and storage organs. Nitrate has to be reduced to ammonia to be incorporated as part of organic, which makes nitrate absorption a slower process compared with ammonium.

Visible Symptoms Deficiency. Nitrogen deficiency symptoms begin with loss of color (light green or chlorosis) in the older leaves and size reduction of younger leaves (Jones 1965). Overall the plant is chlorotic and plant size and stem girth are smaller than normal (Barnes 2010; Barnes et al. 2012; Bennett 1993; Gibson et al. 2007; Winsor and Adams 1987). With prolonged deficiency, younger leaves also become chlorotic, while senescence and dehiscence of older leaves is accelerated (Hewitt 1984b). In some species (i.e., chrysanthemum, begonia, celosia, marigold, and pansy), the older leaves turn red, orange, or purple as a result of reduced chlorophyll content (Gibson et al. 2007; Hewitt 1984b; Laurie and Wagner 1940; Winsor and Adams 1987). Flowering is in general delayed and reduced in number and size (Barnes 2010; Laurie and Wagner 1940) (Fig. 2). The stem size is strongly reduced, affecting both cut and container-grown flowers (Laurie and Wagner 1940). An overall biomass reduction is observed across species (Barnes 2010). Toxicity. Toxicity symptoms of N alone are not well defined. However, excessive vegetative growth and reduction of yields are observed when N levels are high. Ammonium, which is readily assimilated by plants, can result in toxicity. Ammonium toxicity symptoms include chlorosis or interveinal chlorosis of older leaves, and the margins of the leaves become scorched and rolled upward or inward (Gibson et al. 2007; Winsor and Adams 1987) (Fig. 3). Foliar and root biomass are significantly reduced with ammonium toxicity (Cox and Seeley 1984; Gaffney et al. 1982; Gibson et al. 2007; Winsor and Adams 1987). Roots are typically short, unbranched, and discolored. Management Nitrogen deficiency is easy to correct by applying nitrogen in the preferred form. Develop a nutrition program that matches the fertilizer rate to the optimum level of a specific crop and growth stage. Avoid using fertilizers with high ammonium concentration (>40%) in winter months. Ammonium toxicity is corrected by maintaining the growing media pH at > 5.5 (to avoid conversion of nitrate to ammonia) and by avoiding fertilizer with high concentration of ammonium and urea.

4.2

Potassium (K)

Function. Potassium builds up turgor and maintains osmotic potential in plants. Potassium also serves as a carrier across cell membranes and is involved in cell elongation, protein synthesis, enzyme activation, photosynthesis, and water uptake.

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Fig. 2 Nitrogen deficiency symptoms in bacopa (Bacopa monnieri) (top) and New Guinea impatiens (Impatiens hawkeri) (bottom). Healthy plants (on the left of each photo) and nitrogen-deficient plant (right of each photo). Nitrogen deficiency symptoms: loss of green color in foliage (more accentuated in older leaves), reduction of leaf size, flowers, and stem girth. With prolonged deficiency, younger leaves also become chlorotic. Overall reduction of plant size and delayed flowering (P. Fisher # 2017. All Rights Reserved.)

Translocation. Potassium is not structurally bound in any plant component. It is mobile, and therefore it can be redistributed within plants. Potassium moves directly from the rhizosphere to the growing points and also translocates from older tissue to new tissue.

Visual Symptoms Deficiency. Potassium deficiency symptoms begin with mild marginal or interveinal chlorosis, followed by leaf scorch (marginal necrosis or necrotic spots) of recently matured leaves (Barnes et al. 2012). In some cases, the leaf spots can be pronounced red or violet caused by accumulation of anthocyanin (Laurie and Wagner 1940). Consistently across species, as symptoms progress, older leaves become completely necrotic and symptoms progress bottom up. Dieback (terminal and lateral buds die) is observed in woody plants. Young plants are shorter and darker (dark green or blue green) than normal. In Phalaenopsis the tip or margin of the oldest leaves turns

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Fig. 3 Ammonium phytotoxicity symptoms on vinca (Vinca minor). Ammonium toxicity symptoms include chlorosis or interveinal chlorosis of older leaves (top), and the margins of the leaves become scorched and rolled upward or inward (bottom). Foliar and root biomass are significantly reduced. Roots are typically short, unbranched, and discolored (R. Raudales # 2017. All Rights Reserved.)

yellow and then chlorosis progresses from the bottom up in the flowering stem (Wang 2007). As symptoms develop, the older leaves turn to bronze color and then the leaves die. K deficiency does not affect the number of leaves in Phalaenopsis (Fig. 4). Toxicity. Potassium toxicity is rare in practice. Excess potassium in nutrient solutions or soils results in competition with other cations (e.g., Mg+2). Cations are ions that have less electrons than protons, therefore give a positive charge (e.g., Ca+2, Mg+2, Mn+2). However, the deficiency of competing cations is what results in abnormal plant growth and not a direct effect of potassium toxicity (Ulrich and Ohki 1965). In many cases, high levels of potassium can cause salt damage symptoms.

Management Potassium is an element which is associated with cell processes and is a mobile element. Consequently, potassium deficiency symptoms are visible when the deficiency has progressed, and in many cases corrective actions to manage potassium deficiency are not effective (Ulrich and Ohki 1965; Wang 2007). When developing a

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Fig. 4 Potassium deficiency symptoms on phalaenopsis (Phalaenopsis sp.). The tip or margin of the oldest leaves turns yellow and then chlorosis progresses from the bottom up in the flowering stem. As symptoms develop, the older leaves turn to a bronze color (top) and then the leaves die (bottom). The number of leaves is not affected by K deficiency (Y.T. Wang # 2017. All Rights Reserved.)

management strategy for K deficiency, it is important to identify whether the deficiency is caused by insufficient levels of K in the growing media or by imbalances of K with other nutrients. For example, a high ratio of N:K (2:1) or high concentration of sodium (>40 mg/L) inhibits K uptake. In general, potassium levels should be at similar levels to nitrogen. Supplemental fertilization of potassium at the right ratio may correct potassium deficiency. Deficiency caused by low levels of K in the growing media can be amended by supplementing with fertilizers.

4.3

Phosphorus (P)

Function. Phosphorus is an integral component of energy storage molecules, nucleic acids, and membranes. Translocation. Phosphorus is a mobile element in the plant, moving from older tissue to younger tissue in response to low P levels.

Visual Symptoms Deficiency. Early symptoms of P deficiency include slow emergence and growth, and the petioles are purple and foliage turns dark green. As the deficiency progresses, plants exhibit severe spindling and stunting and the foliage changes from dull green or bronze to purple or dark red (Fig. 5). Older leaves become deep green or red and

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Fig. 5 Phosphorus deficiency symptoms in seed geraniums (Pelargonium hortorum, top photo) and impatiens (Impatiens walleriana, bottom photo). Healthy (left in both photos) and P deficient (right in both photos). Early symptoms of P deficiency include slow emergence and growth and petioles are purple. As the deficiency progresses, plants exhibit severe spindling and stunting and the foliage changes from dull green or bronze to purple. Older leaves become deep red and dull. In some cases, P deficiency symptoms are not distinctive and result in reduced growth and flowering (P. Fisher and R. Dickson # 2017. All Rights Reserved.)

become dull (Barnes et al. 2012; Epstein and Bloom 2005; Gibson et al. 2007; Hewitt 1984a). The leaves curl downward (Fig. 5). Flowering is significantly reduced. Toxicity. Excessive concentrations of P do not cause direct damage to plants; however, high P concentrations can result in Cu or Zn deficiencies (Bingham 1965).

Management P deficiencies result from insufficient P fertilization, slow growth (e.g., during cold weather), low B concentration (low B inhibits P uptake), high Fe, Zn or nitrate, or excessive leaching (Gibson et al. 2007). Corrective procedures include adjusting fertilization to match levels to the plant’s needs. Excess P in the root zone can be managed by increasing leaching and then using fertilizers low in P (e.g., 15-0-15).

4.4

Calcium (Ca)

Function. Calcium is a component of cell walls, maintains cell wall integrity, and regulates membrane permeability. Ca also enhances pollen development and germination and is involved in enzyme activation, cell mitosis, division, and elongation.

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Translocation. Calcium is not mobile and is not redistributed after the initial absorption. Plants uptake Ca during transpiration; therefore Ca deficiencies generally appear when growth is slow (e.g., during cold weather, drought, high humidity, etc.).

Visual Symptoms Deficiency. Calcium deficiency symptoms appear first on apical meristems of shoots and roots. The symptoms are known as “death of growing points” (Epstein and Bloom 2005; Hewitt 1984a). Young leaves become distorted, small, and chlorotic. Leaf margins and distorted areas might become necrotic as symptoms progress. The plant overall presents dieback symptoms. Dark spots are developed in some species. Flowers and fruit present “blossom end rot.” Roots are short and thick and develop few root hairs (Barnes et al. 2011) and sometimes necrotic. Flowers and petals drop, and vase life of flowers is shortened. Flower size is smaller than usual. Toxicity. Direct negative effects of excess of calcium are rare. However, excessive calcium concentration can reduce boron availability and result in B deficiency, especially at alkaline pHs when boron is less available. Calcium uptake can compete with other cations (e.g., Mg+, K+). Management Calcium deficiency is caused by insufficient Ca in the root zone (e.g., low Ca fertilizer or excessive leaching), slow growth (caused by low temperatures, high humidity, drought, etc.), or high concentrations of competing cations (e.g., Na+, Mg+) (Gibson et al. 2007) or total salts (Gislerod 1999). Ca reacts with phosphates or sulfates and forms insoluble compounds that plants cannot uptake resulting in Ca deficiency. This problem occurs when the elements are at high concentrations such as stock fertilizer solutions or high concentrations in the soil. Consequently, nutrient analysis of the growing media will provide information about the availability of Ca in the substrate. Management options include reducing leaching, balancing the cations, and supplementing Ca in the fertilization program. In addition, having a separate stock tank for calcium prevents formation of precipitates. Excess calcium potentially comes in the water source, excessive lime application, or over fertilization. Corrective procedures include blending or changing the water source and revising the fertilization program to lower calcium levels or increase the cations that are deficient.

4.5

Magnesium (Mg)

Function. Magnesium is a component of the chlorophyll molecule and activates enzymes involved in photosynthesis, respiration, and DNA synthesis. Translocation. Magnesium is a mobile element that is translocated from mature to young leaves.

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Visual Symptoms Deficiency. Early symptoms appear first in older leaves as interveinal chlorosis. Magnesium and iron are both important for chlorophyll formation; however Fe is immobile and Mg is mobile. Therefore, the symptoms can be distinguished by observing the location of the chlorosis during its early stages. Mg deficiency symptoms are not always as consistent across species. Overall, plants with Mg deficiency are smaller than healthy plants, and leaves turn chlorotic or present interveinal chlorosis. Older leaves turn reddish brown or reddish purple and veins remain green (Gibson et al. 2007; Hewitt 1984a; Laurie and Wagner 1940). Toxicity. Magnesium phytotoxicity has not been reported. However, Mg can inhibit uptake of other cations (e.g., K+). Management Mg deficiency can result from low Mg from the fertilizer program or high content of other cations (e.g., Ca+2, Na+2), dysfunctional roots, and slow growth. Magnesium sulfate (Epsom salts) or fertilizer blends with Mg are commonly used to correct Mg deficiency. High levels of Mg in tissue can come from high Mg in the water source, excessive fertilizer, or low levels of other cations. Corrective procedures include adjusting the fertilizer program (lower Mg or increase other cations) or changing the water source.

4.6

Sulfur (S)

Function. Sulfur is a constituent of amino acids, coenzymes, and vitamins. Sulfur is also a structural component of aromatic compounds. Translocation. Not easily mobilized, translocation varies by species.

Visual Symptoms Deficiency. Sulfur deficiency symptoms are very similar to nitrogen (both are constituents of amino acids and proteins). Deficiency symptoms include general chlorosis, stunting, and reddish foliage (anthocyanin accumulation) (Barnes 2010; Hewitt 1984a). In some cases, the foliage is chlorotic and petioles are red (Dale et al. 1990). Root biomass is reduced, but the color is not altered. Sulfur is generally immobile; consequently, early symptoms appear in younger leaves. However, sulfur translocation varies by species and deficiency might appear in all leaves simultaneously (Taiz and Zeiger 2006). Toxicity. Sulfur toxicity in floriculture crops is rarely observed in practice. High concentrations of sulfur inhibit nitrogen uptake and might result in N deficiency. As a consequence, S toxicity and deficiency could be easily confused. Management Sulfur deficiencies result from inadequate supply of sulfur in the nutrition program. High sulfur comes from the fertilizers, acidifying agents (i.e., sulfuric acid, iron sulfate, elemental sulfur), or irrigation water; to correct the problem, discontinue application of sulfur. If water is the source of the problem, then blend with or change to water from another source.

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Iron (Fe)

Function. Iron plays an important role in chlorophyll synthesis, nitrate reduction, and other enzymatic reactions. Translocation. Iron is an immobile element.

Visual Symptoms Deficiency. The visual symptoms of Fe deficiency are consistent across plants (Barnes et al. 2012; Gibson et al. 2007; Hewitt 1984a; Laurie and Wagner 1940). Deficiency symptoms start with interveinal chlorosis of young leaves, where the veins remain green and the tissue between the veins is chlorotic (Fig. 6). With prolonged deficiency or deficiency at the seedling stage, the whole leaf will turn chlorotic and dehiscence will occur (Fig. 6). Fe is immobile; hence deficiency symptoms appear in younger leaves. Mg deficiency symptoms are similar to Fe, but Mg symptoms first appear in older leaves. Toxicity. Iron toxicity begins with a mild chlorosis followed by bronzing of leaves. Symptoms advance as necrotic spots or necrosis of whole leaves and death (Fig. 7). Management Fe is more soluble at lower pH (acidic) and less soluble at higher pH (basic). Therefore, plants which are “inefficient” at absorbing iron (e.g., petunias, pansies, snapdragons, etc.) tend to show iron deficiency symptoms when the pH is  6.2. In contrast, other species (e.g., seed geraniums, marigolds, lisianthus, etc.) are efficient at taking up Fe and Mn and tend to present Fe and Mn phytotoxicity when the pH is 150 plant genera and to survive as mycelium in plant debris (Booth and Waterston 1964). McGovern et al. (2003) indicated that F. avenaceum could survive on Styrofoam transplant trays and as an endophyte in the root systems of a number of ornamentals including wax begonia (Begonia x semperflorens-cultorum), carnation (Dianthus caryophyllus), exacum (Exacum affine), Gebera daisy (Gerbera jamesonii), Madagascar periwinkle (Catharanthus roseus), French marigold (Tagetes patula), pansy (Viola tricolor subsp. hortensis), petunia (Petunia x hybrida), rudbeckia (Rudbeckia sp.), salvia (Salvia sp.), and verbena (Verbena sp.). In addition, isolates of the fungus from several other hosts were pathogenic to lisianthus, indicating that F. avenaceum may be pathogenic on lisianthus regardless of its phylogenetic origin (Nalim et al. 2009). Seed infection by F. avenaceum has been reported in a number of hosts (Booth and Waterston 1964), but the fungus was not isolated from lisianthus seed (McGovern et al. 2003; Pecchia et al. 2000). Short-distance aerial movement of macroconidia of F. avenaceum in a lisianthus transplant production house in California and two lisianthus cut-flower facilities in Florida was documented (McGovern et al. 2003; Seijo et al. 2000). The fungus is readily spread over long distances via infected transplants, and serious outbreaks of Fusarium crown and stem rot in cut-flower production facilities in the USA were attributed to infected transplants resulting from propagation in reused transplant

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trays without prior disinfestation (McGovern et al. 2003). The fungus can also be spread by fungus gnats (Bradysia spp.), shore flies (Scatella spp.), and moth flies (Psychoda spp.) (McGovern and Harbaugh 1997; El-Hamalawi and Stanghellini 2005). Common feeding sites for fungus gnats in lisianthus are roots at the plant base which are covered by decomposing leaves closely appressed to the growing medium; these sites provide moist infection courts for F. avenaceum. The fungus may also be spread from plant to plant on cutting tools (McGovern et al. 2003). Management • Cultural practices – Use pathogen-free transplants. Avoid overwatering to prevent the buildup of fungus gnat and shore fly populations and control these vectors by biological or chemical means. • Sanitation – Avoid reusing transplant trays without thorough chemical or steam disinfestation; an alternative is the use of disposable plastic inserts that fit within Styrofoam trays. Disinfest cutting tools between plants. Locate cull piles away and downwind from production facilities. Eliminate weeds. Disinfest soil between crops using chemical (fumigation) or physical measures (steam or soil solarization). • Fungicides – McGovern et al. (2002a) reported that preventive application of azoxystrobin reduced disease incidence and plant mortality to ~20% and 0%, respectively, in a growth chamber trial; fluazinam, fludioxonil, myclobutanil, and thiophanate methyl were generally less effective. In a greenhouse experiment in the Netherlands, van der Wurff and Hamelink (2007) found that fludioxonil + cyprodinil and trifloxystrobin were effective in reducing Fusarium crown and stem rot. • Resistance – Harbaugh and McGovern (2000) evaluated the susceptibility of 46 lisianthus cvs. under high disease pressure in growth chamber experiments and found that the lowest disease frequencies 55 days after inoculation with F. avenaceum occurred in “Ventura Deep Blue” and “Hallelujah Purple” (25%), “Bridal Pink” (23%), and “Heidi Pure White” (53%), representing the blue/ purple, pink, and white flower color groups, respectively.

2.7

Fusarium Wilt (Fusarium oxysporum (Schlechtend):Fr. f. sp. eustomae)

Geographic occurrence and impact. Fusarium wilt of lisianthus has occurred in Ecuador, Israel, Italy, Japan, Korea, Poland, the Netherlands, the UK, and the USA (de Werd 2003; Elad et al. 2014; Hahm 1998; McGovern unpublished data; O’Neil and Green 2010; Orlikowski 2001; Raabe 1991; Rapetti et al. 2002; Tomita et al. 2004). Losses of 50–70% observed at a lisianthus cut-flower facility in the Netherlands were attributed to F. oxysporum f. sp. eustomae possibly in combination with Pythium irregulare (de Werd 2003). Losses of 10–50% and 100% were observed at cut-flower sites in Ecuador and the USA (California), respectively (McGovern, unpublished data). Fusarium wilt was reported to be widespread throughout the

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Fig. 7 Fusarium wilt symptoms/signs: root and basal stem rot and white masses of conidia in sporodochia (arrow) on the stem of a wilted plant (R.J. McGovern # 2017. All Rights Reserved.)

ROK especially in alpine production locations where it occurred at incidences of 5–30% (Hahm et al. 1998). Up to 40% crop loss occurred in Poland (Orlikowski 2001). Symptoms/signs. The F. oxysporum f. sp. eustomae infects and causes a dark discoloration of roots and the stem base. The pathogen moves upward through the xylem causing yellowing of leaves and wilting; vascular discoloration is visible when infected stems are cut lengthwise. As the disease progresses white masses of spores (sporodochia) are produced on stems, the leaves become tan, and plants wilt entirely and die (Fig. 7). The pathogen may also cause pre- and postemergence damping-off. Biology and epidemiology. Fusarium oxysporum f. sp. eustomae produces three types of asexual spores: microconidia and macroconidia which enable aerial dissemination and thick-walled, stress-resistant chlamydospores which enable longterm survival. Survival by the fungus in association with plant debris similar to other formae speciales of F. oxysporum is very likely. The fungus has also been shown to survive as a symptomless endophyte in the roots of Aubrieta sp., rapeseed (Brassica

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napus), and stock (Matthiola incana) (O’Neil 2006a). The sexual stage of F. oxysporum has not been reported. The disease is favored by warm temperatures (20–28  C/68–82  F) (Bertoldo et al. 2015; McGovern et al. 2002b). Genetic and virulence variability of the pathogen was reported in Italy (Bertoldo et al. 2015), and a PCR assay was developed which enabled its detection in the roots and stems of infected but symptomless plants (Li et al. 2010). Management • Biological control – A commercial formulation which combines Bacillus sp., Pseudomonas sp., Streptomyces sp., and Trichoderma sp. Consistently reduced plant mortality from F. oxysporum f. sp. eustomae in growth chamber experiments (McGovern et al. 2002c; McGovern unpublished data). The biocontrol Trichoderma lignorum significantly reduced the incidence of damping-off caused by Pythium sp. and F. oxysporum and increased grower profitability in research in commercial facilities in Ecuador (Sacoto Bravo 2009). • Fungicides – In controlled environment experiments, fludioxonil and triflumizole were very effective in reducing both the incidence of and mortality due to F. oxysporum f. sp. eustomae; azoxystrobin, myclobutanil, thiophanate methyl, and trifloxystrobin were less effective, and thiophanate + chlorothalonil was ineffective (McGovern et al. 2002b, c). Chase (2011) reported that the severity of the disease was significantly reduced by the fungicide metconazole and the inducer of systemic acquired resistance, acibenzolar-S-methyl. • Soil disinfestation – Preplant soil disinfestation is an important tool in the management of Fusarium wilt. Failure to fumigate the soil between successive lisianthus crops and inadequate fumigant distribution due to high soil compaction led to serious outbreaks of the disease in the USA (California) and Colombia, respectively (McGovern, unpublished data). O’Neil and Green (2010) reported that sheet steaming (>80  C/176  F for 10 h), a steam plough (>80  C/176  F for 1 h), chloropicrin applied via drip line irrigation, dazomet incorporated into the soil, and formaldehyde and metam sodium drenched onto the soil significantly reduced but did not eliminate the incidence of Fusarium sp. in buried, naturally infected roots and stems of lisianthus in commercial greenhouse experiments in the UK; calcium cyanamide incorporation was ineffective. Preplant steaming and application of metam sodium and dazomet provided acceptable control of Fusarium sp. for cut-flower production in Argentina (Salles et al. 2001). • Resistance – Greenhouse trials conducted in Italy indicated that the lisianthus cvs. Mariachi Green and Echo Dream Yellow were partially resistant to the disease (Gilardi et al. 2006). • Integrative strategies – O’Neil and Green (2007) found that the efficacy of sheet steaming in reducing the viability of F. oxysporum was not improved by combination with a number of biocontrols (Gliocladium catenulatum, nonpathogenic F. oxysporum, Trichoderma harzianum) or fungicides (calcium cyanamide, carbendazim).

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Diseases of Lisianthus

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Phomopsis Stem and Leaf Blight (Phomopsis sp.)

Geographic occurrence and impact. The disease occurred in the USA (Florida) where losses ranging from 4% to 80% were observed in two potted lisianthus production facilities (McGovern et al. 2000a). Symptoms/signs. Stem necrosis is rapidly followed by leaf blight and the production of numerous dark pycnidia in diseased tissue. As the stem blight progresses, infected plants collapse and die (Fig. 8). Biology and epidemiology. The fungus is spread by spores and infected plants and survives as pycnidia and mycelium in infected tissue. Management. Refer to the introductory chapters on integrated disease management.

2.9

Powdery Mildew [Leveillula taurica (Lév.) G. Arnaud (Anamorph: Oidiopsis taurica (Lev.) E. S. Salmon)]; Oidium sp.

Geographic occurrence and impact. Powdery mildew of lisianthus has been reported to be caused by L. taurica (Oidiopsis spp.) in Brazil, Israel, Poland, Spain, Venezuela, and the USA and by Oidium sp. in Argentina and Japan (Cabrera et al. 2009; Cedeño et al. 2009; Elad et al. 2007; Koike et al. 1995; Melgares de Aquilar Cormenzana 1996; Okamoto et al. 2002; Orlikowski 2001; Reis et al. 2007). Fig. 8 Phomopsis stem and leaf blight symptoms/signs. Note the production of numerous black pycnidia (arrow) in infected tissue (R.J. McGovern # 2017. All Rights Reserved.)

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Fig. 9 Powdery mildew symptoms/signs: yellow blotches on leaves covered with profuse, white fungal growth (mycelium, conidiophores, and conidia) (R.J. McGovern # 2017. All Rights Reserved.)

Genetic analysis should elucidate the taxonomic relationships among the various powdery mildew pathogens of lisianthus. Symptoms/signs. Symptoms and signs include yellow blotches on both the upper and lower leaf surfaces and stems which become covered with profuse white fungal growth (mycelium, conidiophores, and conidia). Floral infection and leaf distortion and premature abscission may also occur (Fig. 9). Biology and epidemiology. Only the anamorphic stage of powdery mildew fungi has been observed on lisianthus; the role of the teleomorphic stage is currently unknown. The fungus infects the epidermis but, unlike most powdery mildew pathogens, it also colonizes mesophyllic tissue by means of stomata through which conidiophores subsequently emerge. Elad et al. (2007) found that conidial germination of O. taurica was optimal at 20  C/68  F and a relative humidity of 75–85% and that severe infection occurred at 15–20  C/59–68  F in pepper research in Israel; prolonged temperatures above 25  C/77  F decreased disease severity. Conidia of the fungus are spread by air currents. It has been suggested that L. taurica is a composite species consisting of a number of host-specific races (Braun 1987, 1995). However, Correll et al. (1987) found that L. taurica isolates from a number of different genera and families could cause powdery mildew in tomato (Solanum lycopericum). Okamoto et al. (2002) demonstrated that an isolate of Oidium sp. from 4 o’clock flower (Mirabilis jalapa) could cause powdery mildew symptoms on both lisianthus and broad bean (Vicia faba). Reis et al. (2007) reported that in addition to lisianthus, nasturtium (Tropaeolum majus), calla lily (Zantedeschia aethiopica), impatiens (Impatiens balsamina), and balloon plant (Asclepias physocarpa) were hosts of Oidiopsis. Management. The endoparasitic nature of Leveillula taurica makes it more difficult to control by contact fungicides than ectoparasitic powdery mildew pathogens which infect host tissue more superficially. • Cultural practices – In greenhouse research on pepper (Capsicum annuum) in Israel, Elad et al. (2007) found that powdery mildew caused by Leveillula taurica

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was decreased by increasing nighttime temperatures by heating and daytime temperatures by manipulation of the greenhouse vents. • Fungicides – Numerous fungicides and a number of biocontrols are available for management of powdery mildew (refer to ▶ Chap. 7, “Fungicides and Biocontrols for Management of Florists’ Crops Diseases”). Fungicides should be rotated by mode of action and applied preventively.

2.10

Pythium Root Rot (Pythium spp.)

Geographic occurrence and impact. Pythium root rot in lisianthus may be caused by a number of Pythium species including P. irregulare, P. myriotylum, P. spinosum, as well as unidentified species of the pathogen and has been reported in Ecuador, Israel, Japan, Norway, Spain, and the USA (Aegerter et al. 2002; Elad et al. 2014; McGovern unpublished data; Melgares de Aquilar Cormenzana 1996; Sacoto Bravo 2009; Tomita et al. 2004; Toppe 2005). Losses exceeding 75% occurred in greenhouse in the USA (California) (Aegerter et al. 2002). Symptoms/signs. The root systems become darkly discolored and rotted, and infected plants rapidly wilt. The outer rotted and discolored layer of Pythiuminfected roots is often easily pulled off leaving behind the central, lighter-colored stringy root fiber (stele). The pathogen may also cause pre- and postemergence damping-off (Fig. 10). Biology and epidemiology. The genus Pythium is closely related to Phytophthora and also is fungus-like but not a true fungus. The pathogen is disseminated as

Fig. 10 Pythium root rot symptoms: discolored roots (left), rapid wilting (right) (R.J. McGovern # 2017. All Rights Reserved.)

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zoospores through irrigation, surface water, and rain, and long-term survival is enabled by chlamydospores and sexually produced oospores. Pythium root rot is common in waterlogged soils with poor aeration; cycles of under- and overwatering which lead to root damage are very conducive to outbreaks of Pythium. Fungus gnats (Bradysia spp.) create infection sites through their root-feeding and may disseminate the pathogen. Pythium, depending on the species, may have a very broad host range. Management • Biological control – The biocontrol Trichoderma lignorum significantly reduced the incidence of damping-off caused by Pythium sp. and F. oxysporum and increased grower profitability in research in commercial facilities in Ecuador (Sacoto Bravo 2009). • Fungicides – A number of isolates of P. irregulare recovered from lisianthus in the USA (California) were found to be resistant to the commonly used fungicide mefenoxam (Aegerter et al. 2002).

2.11

Rhizoctonia Crown and Stem Rot Blight [Rhizoctonia solani Kühn (Teleomorph: Thanatephorus cucumeris (A.B. Frank) Donk)]

Geographic occurrence and impact. Rhizoctonia root, crown, and stem rot is a major plant disease and has been documented in lisianthus in Israel, Japan, Spain, and the USA (Alfieri et al. 1994; Meir et al. 2010; Melgares de Aquilar Cormenzana 1996; Yoshimatsu 1993). Tomita et al. (2004) observed 2.8–31.6% losses due to damping-off in the field from R. solani in Japan. Symptoms/signs. Rhizoctonia solani can infect the roots, stems, and foliage of lisianthus. The fungus most typically produces a dark discoloration and rot in stems at the soil line, leading to wilting and rapid collapse of the entire plant. Under humid conditions, mycelial growth of R. solani may rapidly envelop and blight leaves and shoots; this form of the disease is known as Rhizoctonia aerial blight. Discrete leaf lesions may have a concentric appearance. The pathogen may also cause pre- and postemergence damping-off (Fig. 11). Biology and epidemiology. Rhizoctonia solani has a very large host range which encompasses most economically important cultivated plants including ornamentals and weeds; but some host specialization has been demonstrated (Mordue 1974). The fungus is very active at warm temperatures and effectively colonizes and survives as hyphae and sclerotia in plant debris, soil, and other growing media. Although R. solani does not produce spores, it can be spread by infected propagative material including seeds, water splash, and airborne particulate matter. The relationship of the sexual stage of R. solani to plant disease has not been well studied.

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Fig. 11 Rhizoctonia crown and stem rot and foliar blight symptoms (R.J. McGovern # 2017. All Rights Reserved.)

Management • Biological control – Meir et al. (2010) observed a consistent though nonsignificant increase in lisianthus survival in soil infested by both F. solani and R. solani by prior inoculation with the arbuscular mycorrhizal fungus Rhizophagus intraradices (formerly Glomus intraradices). • Soil disinfestation – Chloropicrin applied by chemigation (through drip irrigation tubing) resulted in significant but minor reduction of R. solani soil densities in greenhouse research in the UK (O’Neil 2006b).

2.12

Thielaviopsis Black Root Rot [Thielaviopsis basicola (Berk. and Broome) Ferraris (1912)]

Geographic occurrence and impact. The disease has occurred in Canada, Switzerland, and the USA (Florida) (Joshi 2000; McGovern unpublished data, Michel 2015). A loss of ca. 70% occurred with one planting of a single cultivar in Florida. Symptoms/signs. A black discoloration occurs in the small, feeder roots, or these roots are entirely absent. Infected plants are stunted, turn yellow, wilt, and die (Fig. 12). Biology and epidemiology. Thielaviopsis basicola produces black, thick-walled, resistive chlamydospores (also referred to as macroconidia), and microconidia, which are involved in its survival and dissemination. The fungus is worldwide in distribution especially in cool, wet climates and infects a broad range of cultivated plants in many unrelated families. Black root rot is most severe in cool, wet soils (17–23  C/62–73  F) and at a soil pH above 5.5 (Shew and Lucas 1991).

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Fig. 12 Thielaviopsis black root rot symptoms: note the black discoloration of small, feeder roots (R.J. McGovern # 2017. All Rights Reserved.)

Management • Cultural practices – The disease can be reduced by lowering the soil pH to below 5.5 if feasible, and crop rotation. Additional Fungal Diseases. The following fungal diseases of lisianthus have also been reported: – Alternaria blight (Alternaria sp.) (Israel, Auscher 1997) – Brown root rot [Subplenodomus drobnjacensis (Bubák) Gruyter, Aveskamp, and Verkley 2012] (Japan, Kondo et al. 2014) – Curvularia leaf blotch (Curvularia sp.) (USA, Jones and Harbaugh 1995) – Myrothecium leaf spot and blight (Myrothecium roridum Tode) (The Netherlands, Ludeking and Arkesteijn 2011) – Penicillium root rot (Penicillium sp.) (Japan, Tomita et al. 2004) – Phytophthora root rot and stem blight (Phytophthora spp.) (Japan, Uematsu et al. 1996) – Phyllosticta leaf spot (Phyllosticta sp.) (USA, Daughtrey 2000) – Sclerophoma stem blight (Sclerophoma eustomis Taubenhaus and Ezekiel) (USA, Taubenhaus and Ezekiel 1935) – Sclerotium stem blight [Sclerotium rolfsii Sac. (teleomorph: Athelia rolfsii (Curzi) C.C. Tu and Kimbr.) (USA, Japan, McGovern et al. 2000b; Tomita et al. 2004) – Sclerotinia stem rot [Sclerotinia sclerotiorum (Lib.) de Bary] (Wolcan et al. 1996) – Stemphylium blight (Stemphylium lycopersici Wollr.) (Venezuela, Cedeño et al. 2011) – White rust [Albugo swertiae (Bed. et Komm.) Wilson (Dingley 1969)] (New Zealand, McKenzie 1987) – White blister rust (Pustula centaurii (Hansf.) Thines, C. Rost et Y. J. Choi, comb. nov, MB519564) (Australia, Tasmania, Ploch et al. 2011)

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For an up-to-date list of fungi associated with lisianthus, access http://nt.ars-grin. gov/fungaldatabases/fungushost/FungusHost.cfm and enter “Eustoma” in the host name box.

3

Bacterial and Phytoplasma Diseases

3.1

Bacterial Crown Rot [Burkholderia gladioli (Severini 1913) Yabuuchi et al. 1993, comb. nov.]

Geographic occurrence and impact. The disease has been reported in Poland and the USA (Schollenberger and Zamorski 2008; Seijo et al. 2002). Symptoms/signs. The pathogen can infect plants of various ages ranging from small seedlings to flowering plants and causes tan veinal necrosis in the leaves, rot in the tap root, crown, and stem, and wilting and death (Fig. 13). Biology and epidemiology. Burkholderia gladioli (formerly Pseudomonas gladioli) can infect a number of plant hosts including gladiolus (Gladiolus x hortulanus), onion (Allium cepa), and mushroom (Agaricus bitorquis), and based on its host range was divided into three pathovars: pv. gladioli, pv. alliicola, and pv. agaricicola, respectively (Young et al. 1996). However, Burkholderia gladioli from lisianthus has not yet been characterized to the level of pathovar. The bacterium has also been isolated from the pulmonary tissue of cystic fibrosis patients (Beringer and Appleman 2000). The disease is favored by warm temperatures (28 °C/82 °F), and the bacterium can be seed-borne (Seijo et al 2002). Whitby et al. (2000) developed a species-specific PCR assay that distinguishes between B. gladioli and

Fig. 13 Bacterial crown rot symptoms: crown and stem rot and veinal necrosis (left) wilting and stem collapse (right) (R.J. McGovern # 2017. All Rights Reserved.)

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a number of other closely related species such as B. caryophylli which causes similar symptoms in lisianthus. Management • Cultural practices – Use pathogen-free seed, and avoid unnecessarily wounding plants. Rapidly remove infected plants.

3.2

Burkholderia Wilt [Burkholderia caryophylli (Burkholder 1942) Yabuuchi et al. 1993, comb. nov.]

Geographic occurrence and impact. This disease has occurred in Japan (Furuya et al. 2000). Symptoms/signs. The symptoms are similar to those caused by Ralstonia solanacearum (see next disease). The pathogen causes root rot and yellowing, wilting, and browning of the foliage through destruction of the water-conducting tissue; a cross section of infected stems reveals a typical tan to yellow-brown discoloration of the cortical tissue. The lower leaves are affected first. Biology and epidemiology. Burkholderia caryophylli (formerly Pseudomonas caryophylli) also causes rot in carnation, onion, and statice (Limonium sinuatum) (Ballard et al. 1970; Jones and Engelhard 1984; Palleroni 1984). Shao et al. (2011) developed a real-time fluorescent PCR assay for specific detection of B. caryophylli. Management • Cultural practices – Rapidly remove infected plants.

3.3

Ralstonia Wilt (Ralstonia solanacearum (Smith 1896) Yabuuchi et al. 1996, comb. nov.)

Geographic occurrence and impact. Bacterial wilt caused by Ralstonia solanacearum (formerly Pseudomonas solanacearum) is one of the most important bacterial diseases of cultivated plants and has been reported to occur in hundreds of plant species representing more than 40 families (Buddenhagen and Kelman 1964). This disease of lisianthus has occurred in Japan and Taiwan (Chao et al. 1995; Tomita et al. 2004). Symptoms/signs. Disease symptoms include wilting of lower leaves and then the entire plant, followed by browning of the foliage and plant death. In cross section, infected stems exhibit a characteristic brown discoloration in cortical tissue; infected stems also feel hollow when pressed. Biology and epidemiology. Five groups (races) of Ralstonia solanacearum have been described based on host range (Buddenhagen 1986; Buddenhagen et al. 1962).

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Thus far, all strains of the bacterium in Taiwan have been classified as race 1, including the strain that infects lisianthus (Lee et al 2001). Race 1 infects many solanaceous crops, ornamentals, and weeds and has a very high temperature optimum (35–37  C/95–98  F). The bacterium can survive in the soil in association with plant debris (Sugawara and Ishii 2009). In carnation the pathogen has been reported to move from one cutting to another in irrigation water during propagation, but plant to plant transmission appeared to be slow and limited to neighboring plants (Bradbury 1973). Lee et al. (2001) have developed a method to detect race 1 strains of R. solanacearum by PCR. Management • Cultural practices – Sugawara and Ishii (2009) recommended the avoidance of successive plantings of lisianthus, rapid removal of infected plants, disinfestation of soil and equipment, and improvement of drainage.

3.4

Phytoplasma Diseases

Geographic occurrence and impact. Diseases of lisianthus caused by phytoplasmas have been reported to occur in Brazil and Israel (Rivas et al. 2000; Weintraub et al. 2007). However, it is likely that additional reports of phytoplasma infection of lisianthus will be forthcoming given the global distribution of both host and pathogen. Symptoms/signs. Symptoms caused by phytoplasmas may include yellowing of foliage, stunting and proliferation of shoots (witch’s brooms), and the abnormal development of floral parts into leafy structures (phyllody) and of green coloration in flowers (virescence). Biology and epidemiology. Phytoplasmas are prokaryotes that lack cell walls and are obligate plant parasites vectored by phloem-feeding insects in the Cicadellidae family and Fulgoroidea superfamily such as leafhoppers and planthoppers, respectively. Based on 16S rDNA analysis, the phytoplasma detected in lisianthus and those detected in other plants in Israel including apricot (Prunus sp.), cyclamen (Cyclamen sp.), grape (Vitis vinifera), papaya (Carica papaya), and pepper (Capsicum annuum) have been placed in the Stolbur phytoplasma group (Weintraub et al. 2007). Six confirmed leafhopper vectors and one suspected planthopper vector of phytoplasmas occur in Israel (Weintraub et al. 2007). Management. Management of phytoplasma diseases, like those caused by viruses, is based on monitoring and reduction of insect vector populations through chemical or biological means, vector exclusion by physical barriers such as fine mesh screening, vector disorientation and repellence in field-grown crops through the use of reflective mulches, and elimination of, and avoidance of growing near, alternate hosts especially established, infected crops.

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4

Viral Diseases

4.1

Bromoviridae

Geographic occurrence and impact. Viruses in the Bromoviridae commonly cause major worldwide crop losses. Two members of this group have been reported in lisianthus: Cucumber mosaic virus (CMV, genus Cucumovirus) and Tobacco streak virus (TSV, genus Ilarvirus) (de Freitas et al. 1996a; Fujinaga et al. 2006; Gera and Cohen 2007; Hahm et al. 1998; Providenti 1985; Rivas et al. 2000). The geographic occurrence of these viruses is indicated in Table 1. CMV was detected at an incidence of 8.6% in Japan (Nagano Prefecture) (Fuginaga et al. 2006) and 10% in alpine production areas in the ROK in combination with a Fabavirus, Broad bean wilt virus (Hahm et al. 1998). Symptoms/signs. Symptoms caused by plant viruses, including the Bromoviridae, can be variable and depend on the host, environment, virus strain, and coinfection with other viruses; different Bromoviridae genera may produce similar symptoms. Symptoms caused by the Bromoviridae in lisianthus are indicated in Table 1. Biology and epidemiology. Viruses in the Bromoviridae are acquired and spread by aphids (Hemiptera: Aphididae) or thrips (Thysanoptera: Thripidae) in a noncirculative (nonpersistent) manner; these viruses are mouthpart-borne and do not enter the insect’s hemolymph. Spread of TSV-infected pollen by thrips has been demonstrated (Klose et al. 1996). Seed transmission of CMV has been reported in many plant species at incidences of less than 1% up to 50% (Garcia-Arenal and Palukaitis 2008). Seed transmission of TSV in various hosts has been observed at incidences of 0.7–90.6% (Kaiser et al. 1982). Management. Virus management, including species in the Bromoviridae, is based on monitoring and reduction of vector populations through chemical and/or biological measures, vector exclusion by physical barriers such as fine mesh screens, vector disorientation in field-grown crops through the use of reflective mulches, and elimination of, and avoidance of growing near, alternate hosts especially established, infected crops. Cross protection, prior infection with an attenuated strain of CMV, was shown to reduce subsequent infection by a severe strain of the virus in cucumber, lisianthus, petunia, and tomato (Sayama 1996). Resistance to CMV has been introduced into a wide variety of vegetables through genetic modification. Additional information on integrated disease management of viruses may be found in the introductory (▶ Chap. 4, “Insect Management for Disease Control in Florists’ Crops”).

4.2

Bunyaviridae

Geographic occurrence and impact. Plant viruses in the Bunyaviridae and genus Tospovirus cause extensive losses in nearly all cultivated plants. Thus far seven

Tobacco streak virus (TSV)

Viruses Cucumber mosaic virus (CMV)

Occurrence in lisianthus Brazil, Israel, Japan, the ROK, the USA Brazil

Frankliniella occidentalis, F. schultzei Microcephalothrips abdominalis, Thrips parvispinus, T. tabaci, and by seed

Transmission >80 aphid species including Aphis gossypii and Myzus persicae and by seed

Table 1 Bromoviridae infecting lisianthus

A wide range of vegetables, ornamentals, and weed species

Other natural hosts A very large number of vegetable, ornamental, and weed species Irregularly shaped, necrotic streaks or ring spots on leaves; flower size reduction and premature senescence

Symptoms in lisianthus Leaf mosaic and distortion, stunting, flower color-breaking, and malformation

References Garcia-Arenal and Palukaitis (2008), Gera and Cohen (2007), Hahm et al. (1998), Providenti (1985), and Rivas et al. (2000) de Freitas et al. (1996a), Kaiser et al. (1982), and Klose et al. (1996)

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viruses in this genus have been detected in lisianthus including: Chrysanthemum stem necrosis virus (CSNV), Groundnut ringspot virus (GRSV), Impatiens necrotic spot virus (INSV), Iris yellow spot virus (IYSV), Lisianthus necrotic ringspot virus (LNRV), Tomato chlorotic spot virus (TCSV), and Tomato spotted wilt virus (TSWV) (Alexandre et al. 1999; de Freitas et al. 1996b; Fujinaga et al. 2006; Kritzman et al. 2000; McGovern unpublished data; McGovern et al. 1997b; Momonoi et al. 2011; Mumford et al. 2008; Shimomoto et al. 2014; Wolcan et al. 1996). Alexandre et al. (1999) found that three lisianthus plants in Brazil were co-infected with either CSNV + TCSV + GRSV or those three viruses + TSWV. The geographic occurrence of these viruses is indicated in Table 2. CSNV was observed at an incidence of 10% in a commercial greenhouse in Brazil (Duarte et al. 2014). A very low incidence (900 species in >90 monocotyledonous and dicotyledonous plant families

Not determined

Necrotic spots and ring spots in leaves (Fig. 18)

Ring spots, necrosis, leaf deformation, stunting

Necrotic spots and ring spots in leaves (Fig. 17)

Alexandre et al. (1999), de Freitas et al. (1996b), McGovern (unpublished data), Melgares de Aquilar Cormenzana (1996), Pappu et al. (2009), Veerakone et al. (2015), and Wolcan et al. (1996)

Alexandre et al. (1999)and Polston et al. (2013)

Shimomoto et al. (2014) and Zen et al. (2008)

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Fig. 14 Symptoms of CSNV: stem and foliar necrosis (Momonoi et al. 2011)

Fig. 15 Symptoms of INSV: distortion and yellow spots in newly developing leaves (left) and yellow-tan and ring spots in mature leaf (right) (R.J. McGovern # 2017. All Rights Reserved.)

Fig. 16 Symptoms of IYSV: stem necrosis (left) and systemic necrosis (right) (Srinivasan et al. 2011)

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Fig. 17 Symptoms of LNRV (Shimomoto et al. 2014)

Fig. 18 Symptoms of TSWV: yellow-tan ring spots (Florida Department of Agriculture and Consumer Services # 2017. All Rights Reserved.)

resistant to TSWV. Peng et al. (2014) produced transgenic tobacco plants (Nicotiana benthamiana Domin.) that were resistant to multiple tospoviruses including GNRV, INSV, IYSV, and TSWV. A method for transformation of lisianthus was developed and the TSWV nucleoprotein was introduced, suggesting that genetic engineering of this crop for tospovirus resistance is feasible (Semeria et al. 1996). • Integrative strategies – Zen et al. (2010) reported that the combined use of acephate with reflective screening was very effective in reducing IYSV in lisianthus in greenhouses in Japan. The combination of UV-reflective mulch, the systemic acquired resistance activator acibenzolar-S-methyl, and insecticides (methamidophos, spinosad) was very effective in reducing TSWV incidence in field-grown tomato in the USA (Florida) (Momol et al. 2004). Additional information on integrated disease management of viruses may be found in the introductory (▶ Chap. 4, “Insect Management for Disease Control in Florists’ Crops”).

4.3

Geminiviridae Tomato Yellow Leaf Curl Virus (TYLCV, Genus Begomovirus)

Geographic occurrence and impact. The virus has become a limiting factor for lisianthus production in Israel; TYLCV incidences of near 100% have occurred in

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Fig. 19 Symptoms of TYLCV: stunting, yellowing and upward curl of leaves, and flower bleaching in infected plant (left); healthy plant (right) (R.J. McGovern # 2017. All Rights Reserved.)

commercial facilities (Cohen et al. 1995). An incidence of 5% was reported in a greenhouse in the ROK (Kil et al. 2014). Two other B. tabaci-vectored begomoviruses, Ageratum yellow vein virus and Papaya leaf curl guangdong virus were reported to infect lisianthus in Taiwan (Cheng et al. 2005; Chen et al. 2016). Symptoms/signs. TYLCV symptoms in lisianthus include stunting, upward curling, veinal swelling (on the lower leaf surface), and yellowing of leaves, flower bleaching, and failure to flower (Fig. 19). Biology and epidemiology. TYLCV comprises a virus complex that has become a worldwide limiting factor for production of tomato (Solanum lycopersicum). The virus is transmitted circulatively and non-propagatively by the whitefly Bemisia tabaci (Gennadius) (Hemiptera: Aleyrodidae). Once TYLCV is acquired, the whitefly remains viruliferous for life. Besides lisianthus and tomato, other natural hosts of TYLCV include a number of solanaceous plants [black nightshade (Solanum nigrum), chili pepper (Capsicum chinense), Jimsonweed (Datura stramonium), petunia (Petunia x hybrida), sweet pepper (C. annuum), and tobacco (Nicotiana tabacum)] and common bean (Phaseolus vulgaris) (Díaz-Pendón et al. 2010). Management. Whitefly management through physical, chemical, and biological means and avoidance of locating lisianthus transplant and production facilities near established TYLCV-infected crops are essential for management of the virus.

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Additional information on integrated disease management of viruses may be found in the introductory (▶ Chap. 4, “Insect Management for Disease Control in Florists’ Crops”).

4.4

Potyviridae

Geographic occurrence and impact. Viruses in the genus Potyvirus infect most cultivated plants and weeds. A number of potyviruses cause disease in lisianthus including: Bean yellow mosaic virus (BYMV), Pepper veinal mottle virus (PVMV), Turnip mosaic virus (TuMV), and Watermelon mosaic virus (WMV). The geographic occurrence of these viruses is indicated in Table 3. Symptoms/signs. The range of symptoms caused by potyviruses in lisianthus is presented in Table 3. Biology and epidemiology. Potyviruses are spread by aphids (Hemiptera: Aphididae) in a noncirculative (nonpersistent) manner. It was suggested that other ornamentals, such as freesia (Freesia alba) or gladiolus, known to be hosts of BYMV, may have provided a reservoir for outbreaks of the virus in lisianthus (Lisa and Dellavalle 1987). Management. Uga et al. (2004) demonstrated that dwarf lisianthus plants that had been infected with an attenuated strain of BYMV (B-33) were protected from subsequent infection by a virulent isolate of the virus. Refer to the Bromoviridae Management section of this chapter and introductory (▶ Chap. 4, “Insect Management for Disease Control in Florists’ Crops”).

4.5

Secoviridae Broad Bean Wilt Virus (BBWV, Genus Fabavirus)

Geographic occurrence and impact. BBWV was detected in lisianthus in Japan (Nagano Prefecture) at an incidence of 5.7% (Fujinaga et al. 2006) and at 10% in combination with CMV in alpine production areas of the ROK (Hahm et al. 1998). Symptoms/signs. Mosaic Biology and epidemiology. BBWV can be found worldwide and infects many economically important vegetable and ornamental crops. It is transmitted in a noncirculative manner by a number of aphids including A. gossypii and M. persicae (Belliure et al. 2009). Management. Refer to the Bromoviridae Management section of this chapter and introductory (▶ Chap. 4, “Insect Management for Disease Control in Florists’ Crops”).

Watermelon mosaic virus (WMV)

Pepper veinal mottle virus (PVMV) Turnip mosaic virus (TuMV)

Viruses Bean yellow mosaic virus (BYMV)

Japan

Brazil, Taiwan

Taiwan

Occurrence in lisianthus Brazil, Israel, Italy

>35 aphid species

Aphis gossypii, A. spiraecola, Myzus persicae, Toxoptera citricida 40–50 aphid species especially M. persicae and Brevicoryne brassicae

Transmission At least 19 aphid species with M. persicae being the most effective

Table 3 Potyviruses infecting lisianthus Other natural hosts 14 plant families including vegetables (legumes) and ornamentals (freesia, gladiolus, etc.) Mainly infects cultivated and weedy solanaceous plants (black nightshade, pepper, petunia, tomato, etc.) Chinese mustard (Brassica campestris ssp. chinensis), Chinese white cabbage (B. campestris ssp. chinese var communis), radish (Raphanus sativus), rape (B. campestrisi) and mustard (B. juncea), etc. >170 species in 26 mono- and dicotyledonous families including cultivated plants and weeds Chlorotic or whitish ring spots on leaves

Stunting, systemic yellow spotting

Symptoms in lisianthus Stunting, mosaic and leaf curl, flower colorbreaking Stunting, yellow blotches, and ring spots on leaves

Inoue and Kasuyama (2001)and Lecoq and Desbiez (2008)

Alexandre et al. (2005), Chao et al. (2000), and ICTVdB Management (2006b)

References Alexandre et al. (2005), Bos (2010), Gera and Cohen (2007), Lisa and Dellavalle (1987), and Swenson (1957) Cheng et al. (2009)and ICTVdB Management (2006a)

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4.6

Diseases of Lisianthus

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Tombusviridae

Geographic occurrence and impact. Three members of the Tombusviridae infect lisianthus including Carnation mottle virus (CarMV, genus Carmovirus), Lisianthus necrosis virus [LNV (a strain of Eggplant mottled crinkle virus), genus Necrovirus], and Moroccan pepper virus (MPV, genus Tombusvirus). The geographic occurrence of these viruses is indicated in Table 4. CarMV was detected at an incidence of 20% in field-grown lisianthus in Taiwan (Chen et al. 2011). About 85% of the lisianthus surveyed in greenhouses in the Pakdasht region of Iran exhibited symptoms of MPV infection (Beikzadeh et al. 2011). Symptoms/signs. The range of symptoms caused by the Tombusviridae in lisianthus is presented in Table 4. Biology and epidemiology. The Tombusviridae contains virus genera that persist in, and are spread by, soil, plant debris, water, beetles, chytrid fungi such as Olpidium brassicae, and mechanically. It has been suggested that plants may become infected when their roots come in contact with sloughed-off virus-infected root tissue. Tombusviridae generally have a limited host range compared to other virus groups infecting lisianthus. The mode of infection of lisianthus by mechanically transmissible viruses with no known vectors in the outbreaks of CarMV and MPV mentioned above remains unexplained. It is possible that these viruses were spread by cutting tools contaminated by use in another infected cut-flower crop or through the action of an unknown vector. Management. Refer to the introductory (▶ Chap. 4, “Insect Management for Disease Control in Florists’ Crops”).

4.7

Virgaviridae

Geographic occurrence and impact. Three members of this virus group infect lisianthus: Tobacco mosaic virus (TMV, genus Tobamovirus), Tomato mosaic virus (ToMV, genus Tobamovirus), and Pepper ringspot virus (PepRSV, genus Tobravirus). The geographic occurrence of these viruses is indicated in Table 5. Symptoms/signs. The symptoms of these viruses are presented in Table 5. Biology and epidemiology. The Virgaviridae is a relatively new family of six rod-shaped plant virus genera including Tobamovirus and Tobravirus (Adams and Antoniw 2009). These viruses are transmitted mechanically and in some cases by seed, are resistant to degradation, and may persist in the soil.

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Table 4 Tombusviridae infecting lisianthus Viruses Carnation mottle virus (CarMV)

Occurrence in lisianthus Taiwan

Transmission Mechanical, especially through asexual plant propagation

Lisianthus necrosis virus (LNV)

Japan, Taiwan

Mechanical and by the chytrid fungus Olpidium sp.

Moroccan pepper virus (MPV)

Iran, Japan

Mechanical

Other natural hosts 30 species in 15 plant families, including ornamentals such as calla lily (Zantedeschia spp.), carnation, geranium (Pelargonium hortulanum), hibiscus (Hibiscus sp.), narcissus (Narcissus sp.) Calla lily, carnation, and a number artificially inoculated ornamentals such as petunia and zinnia (Zinnia elegans)

Jimsonweed, lettuce (Lactuca sativa), pelargonium (Pelargonium zonale), pepper, tomato

Symptoms in lisianthus Systemic necrotic spots (Fig. 20)

Initially yellow spots appear on upper leaves followed by systemic necrotic leaf lesions, ring spots and distortion, tip necrosis, and flower colorbreaking (Fig. 21) Stunting, necrotic leaf spots, distortion, and necrosis of leaves and stems (Fig. 22)

References Chen et al. (2011)and Qu and Morris (2008)

Chang et al. (2007), Chen and Hsu (2002), Chen et al. (2000), and Iwaki et al. (1987)

Beikzadeh et al. (2011), Fischer and Lockhart (1977), Okhi et al. (2014), Vetten and Koenig (1983), and Wintermantel and Hladky (2013)

Management. Use virus-free seed. Promptly rogue infected plants and have workers disinfest their hands before resuming regular duties. Disinfest cutting tools between plants. Areas where a high virus incidence has been observed should be scheduled last for cultural activities.

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Fig. 20 Symptoms of CarMV: systemic necrotic spots (Y.-K Chen # 2017. All Rights Reserved.)

Fig. 21 Symptoms of LNV: systemic yellow turning necrotic leaf lesions and ring spots, leaf distortion, and flower color-breaking (Y.-K Chen # 2017. All Rights Reserved.)

5

Nematode Diseases

5.1

Root Knot (Meloidogyne spp.)

Geographic occurrence and impact. Root knot in lisianthus has been reported in Italy, Israel, and the USA (California and Florida) (Elad et al. 2014; Russo and di Vito 2005; Schochow et al. 2004; Wang and McSorley 2004). Symptoms/signs. Yellowing of foliage, stunting, flower delay and number reduction, and root galls (Fig. 23).

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Fig. 22 Symptoms of MPV: necrotic leaf spots (left); distortion and necrosis of leaves and stems (right) (Ohki et al. 2014)

Table 5 Virgaviridae infecting lisianthus Viruses Tobacco mosaic virus (TMV) Tomato mosaic virus (ToMV) Pepper ringspot virus (PepRSV)

Occurrence in lisianthus Israel

Transmission Mechanical, seed

Other natural hosts Very broad host range

Symptoms in lisianthus Stunting, leaf mosaic

Italy, Taiwan

Mechanical, seed

Very broad host range

Stunting, mosaic, and necrosis of leaves

Brazil

Mechanical

Some vegetable species and ornamentals such as Gerbera jamesonii, Gloxinia sylvatica

Stunting, mosaic, and necrotic line patterns in leaves, flower color-breaking

References Gera and Cohen (2007) Jan et al. (2003) and Lisa and Gera (1995) Rivas et al. (2000)

Biology and epidemiology. Root knot in lisianthus can be caused by three species of Meloidogyne (Tylenchida: Heteroderidae): M. hapla, M. incognita, and M. javanica. Root-knot nematodes begin as eggs in root tissue or the soil and pass through four juvenile stages (J1–J4) before reaching maturity. The J2 stage is the only one capable

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Fig. 23 Symptoms of root-knot nematode (R.J. McGovern # 2017. All Rights Reserved.)

of infecting roots, where they produce giant cells in the host in which they feed, enlarge, and undergo their final developmental stages. The mature females become globose and produce eggs, while the adult males migrate from the host and become free-living. Root-knot nematodes are primarily spread by water, equipment contaminated with infested soil, and in infected plant material in the case of vegetatively propagated crops. Based on reproduction on cotton and/or tobacco, M. incognita was divided into four races (Taylor and Sasser 1978). Wang and McSorley (2004) reported that lisianthus “Avila Rose Rim” and “Echo Pink” were relatively poor hosts of M. incognita races 1 and 2. Schochow et al. (2004) found that lisianthus was a better host for M. javanica than for M. incognita and a poor host for M. hapla; but that all three species reduced flower number per plant. Management. Prevention of nematode dissemination through disinfestation of equipment is critical. Disinfestation of soil in florists’ crops production sites has utilized fumigation, chemical drenches, and heat treatment by steam or soil solarization. Refer to ▶ Chap. 8, “Soil/Media Disinfestation for Management of Florists’ Crops Diseases.” Additional Nematode Pathogens. The following nematode pathogen of lisianthus has also been reported: Needle nematode (Longidorus spp.) (Israel, Elad et al. 2014)

References Adams MJ, Antoniw JF (2009) Virgaviridae: a new family of rod-shaped plant viruses. Arch Virol 154:1967–1972. doi:10.1007/s00705-009-0506-6 Aegerter BJ, Greathead AS, Pierce LE, Davis RM (2002) Mefenoxam-resistant isolates of Pythium irregulare in an ornamental greenhouse in California. Plant Dis 86(6):692. doi:10.1094/ PDIS.2002.86.6.692B. Accessed 22 Oct 2016 Alexandre MAV, Duarte LML, Rivas EB, Chagas CM (1999) Mixed infections by Tospovirus species in ornamental crops in São Paulo State, Brazil. Summa Phytopathol 25:353–356

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Alexandre MAV, Vagueiro Seabra P, Borges Rivas E, Lembo Duarte LM, Galleti SR (2005) Vírus, viróides, fitoplasmas e espiroplasmas detectados em plantas ornamentais no período de 1992 a 2003. Rev Bras Hortic Ornam Campinas 11(1):49–57 (in Portugese) Alfieri SA, Langdon KR, Kimbrough JW, El-Gholl NE, Whelburg C (1994) Diseases and disorders of plants in Florida. Florida Department of Agriculture and Consumer Services, Division of Plant Industry Bulletin No. 14. Gainesville, FL Allen WR, Matteoni JA (1991) Petunia as an indicator plant for use by growers to monitor for thrips carrying the tomato spotted wilt virus in greenhouses. Plant Dis 75:78–82 Aloj B, Scalcione M, Nanni B, Marziano F (1990) Studies of a new phytopathogen in Italy: Peronospora of Eustoma (Lisianthus) russelianum. Annali della Facoltà di Scienze Agrarie della Università degli Studi di Napoli, Portici. 24:45–52 (in Italian, English abstract) Arthurs S, Heinz KM (2003) Thrips parasitic nematode Thripinema nicklewoodi (Tylenchida: Allantonematidae) reduces feeding, reproductive fitness, and Tospovirus transmission by its host, Frankliniella occidentalis (Thysanoptera: Thripidae). Environ Entomol 32(4):853–858 Auscher R (1997) Implementation of integrated pest management in Israel. Phytoparasitica 25 (2):119–141 Bag S, Schwartz HF, Cramer CS, Havey MJ, Pappu HR (2014) Iris yellow spot virus (Tospovirus: Bunyaviridae): from obscurity to research priority. Mol Plant Pathol. doi:10.1111/mpp.12177 Ballard RW, Palleroni NJ, Doudoroff M, Stanier RY (1970) Taxonomy of the aerobic pseudomonads: Pseudomonas cepacia, P. marginata, P. alliicola and P. caryophylli. J Gen Microbiol 60:199–21 Beikzadeh N, Peters D, Hassani-Mehraban A (2011) First report of Moroccan pepper virus on Lisianthus in Iran and worldwide. Plant Dis 95(11):1485. doi:10.1094/PDIS-04-11-0342. Accessed 22 Oct 2016 Belliure B, Gómez-Zambrano M, Ferriol I, La Spina M, Alcácer L, Debreczeni DE, Rubio L (2009) Comparative transmission efficiency of two Broad bean wilt virus 1 isolates by Myzus persicae and Aphis gossypii. J Plant Pathol 91(2):475–478 Bennison J, Maulden K, Barker I, Morris J, Boonham N, Smith P, Spence N (2001) Reducing spread of TSWV on ornamentals by biological control of western flower thrips. In: Marullo R, Mound L (eds) Thrips and tospoviruses proceedings of the 7th International Symposium on Thysanoptera pp 215–219 http://www.ento.csiro.au/thysanoptera/Symposium/Section7/33Bennison-et-al.pdf. Accessed 22 Oct 2016 Bennison J, Maulden K, Tomiczek M, Morris J, Barker I, Boonham N, Spence N (2007) Use of entomopathogenic nematodes for the management of western flower thrips and tospoviruses. J Insect Sci 7(28):4. doi:10.1673/031.007.2807 Beringer PM, Appleman MD (2000) Unusual respiratory bacterial flora in cystic fibrosis: microbiologic and clinical features. Curr Opin Pulm Med 6:545–550 Bertoldo C, Gilardi G, Spadaro D, Gullino ML, Garibaldi A (2015) Genetic diversity and virulence of Italian strains of Fusarium oxysporum isolated from Eustoma grandiflorum. Eur J Plant Pathol 141(1):83–97 Booth C, Waterston, JM (1964) Fusarium avenaceum. CMI descriptions of pathogenic fungi and bacteria no. 25. CAB International Bos L (2010) Legume viruses. In: Mahy BWJ, van Regenmortel MHV (eds) Desk encyclopedia of plant and fungal virology. Elsevier/Academic, Amsterdam, pp 418–425 Bradbury JF (1973) Pseudomonas caryophylli. CMI descriptions of pathogenic fungi and bacteria no. 373. CAB International Braun U (1987) A monograph of the Erysiphales (powdery mildews). Beih Nova Hedwigia 89:1–700 Braun U (1995) The powdery mildews (Erysiphales) of Europe. Nord J Bot 16(2):121–232 Buddenhagen IW (1986) Bacterial wilt revisited. In: Persley GJ (ed) Bacterial wilt disease in Asia and the South Pacific, PRO13. Proceedings of an International Workshop, PCARRD, Los Banos, Philippines. Australia Centre for International Agricultural Research, Canberra, Australia pp 126–143

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Schollenberger M, Zamorski C (2008) New bacterial disease of lisianthus. Prog Plant Prot 2:520–523 Seijo TE, McGovern RJ, Morrison RH (2000) Use of a Rotorod spore sampler to examine potential airborne dispersal of Fusarium avenaceum causing crown and stem rot of lisianthus. Phytopathology 90(6 Supplement):S128, http://www.apsnet.org/members/divisions/south/meetings/ Pages/2000MeetingAbstracts.aspx. Accessed 22 Oct 2016 Seijo TE, McGovern RJ, Dickstein ER, Harbaugh BK (2002) Bacterial crown rot of lisianthus caused by Burkholderia gladioli. Online. Plant Health Prog. doi:10.1094/PHP-2002-0520-01-HN Semeria L, Ruffoni B, Rabaglio M, Genga A, Vaira AM, Accotto GP, Allavena A (1996) Genetic transformation of Eustoma grandiflorum by Agrobacterium tumefaciens. Plant Cell Tissue Org Cult 47:67–72 Shao X-L, Gan Q-H, Li Y, Zhao W-J, Wu X-H, Su Z-P (2011) Detection of Burkholderia caryophylli by TaqMan real-time fluorescent PCR. Acta Phytopathol Sin 41(1):24–30 Sherman JM, Moyer JW, Daub ME (1998) Tomato spotted wilt virus resistance in chrysanthemum expressing the viral nucleocapsid gene. Plant Dis 82:407–414 Shew HD, Lucas GB (eds) (1991) Compendium of tobacco diseases. APS Press, St. Paul Shimomoto Y, Kobayashi K, Okuda M (2014) Identification and characterization of Lisianthus necrotic ringspot virus, a novel distinct tospovirus species causing necrotic disease of lisianthus (Eustoma grandiflorum). J Gen Plant Pathol 80:169–175. doi:10.1007/s10327-014-0503-9 Shinners L (1957) Synopsis of the genus Eustoma (Gentianaceae). Southwest Natl 2:38–43 Shpialter L, Rav David D, Dori I, Yermiahu U, Pivonia S, Levite R, Elad Y (2009) Cultural methods and environmental conditions affecting gray mold and its management in lisianthus. Phytopathology 99:557–570 Spadotti DMA, Leão EU, Rocha KCG, Pavan MA, Krause-Sakate R (2014) First report of Groundnut ringspot virus in cucumber fruits in Brazil. New Dis Rep 29:25. doi:10.5197/ j.2044-0588.2014.029.025. Accessed 22 Oct 2016 Srinivasan R, Sundaraj S, Pappu HR, Diffie S, Riley DR, Gitaitis RD (2012) Transmission of Iris yellow spot virus by Frankliniella fusca and Thrips tabaci (Thysanoptera: Thripidae). J Econ Entomol 105(1):40–47. doi:10.1603/EC11094. Accessed 22 Oct 2016 Stegmark R (1994) Downy mildew on peas (Peronospora viciae f. sp. pisi). Agronomie EDP Sci 14 (10):641–647 Stovold G (1998) Fungicide resistance in isolates of Botrytis from ornamentals and development of control strategies. Horticultural Research & Development Corporation, New South Wales, Australia, Report NY 306. http://www.ngia.com.au/Attachment?Action=Download&Attach ment_id=1237. Accessed 22 Oct 2016 Strandberg J (2003) Colletotrichum leaf and flower blight of lisianthus. University of Florida-IFAS, Mid-Florida Research and Education Center Sugawara T, Ishii T (2009) Bacterial wilt of Eustoma grandiflorum. National Agriculture and Food Research Organization, Japan, http://www.naro.affrc.go.jp/flower/kakibyo/plant_search/ta/ lisianthus/post_290.html (in Japanese). Accessed 22 Oct 2016 Swenson KG (1957) Transmission of Bean yellow mosaic virus by aphids. J Econ Entom 50 (60):727–731 Taubenhaus JJ, Ezekiel WN (1935) Fusarium crown and root rot, and sclerophoma stem blight, of the Texas Bluebell. Bull Torrey Bot Club 62(9):503–510 Taylor AL, Sasser JN (1978) Biology, identification, and control of root-knot nematodes (Meloidogyne species). North Carolina State University Graphics, Raleigh Tomita Y, Chiba T, Ogawara T, Nagatsuka H (2004) Occurrence of several diseases during Russell Prairie gentian cultivation in Ibaraki prefecture. Bull Hortic Instit Ibaraki Agric Center 12:28–38 (in Japanese) Toppe B (2005) Fungal diseases in Eustoma grandiflorum – occurrence and control. Grønn Kunnskap 9(2):70–75, http://www.bioforsk.no/ikbViewer/Content/18614/toppe2.pdf (In Norwegian). Accessed 22 Oct 2016

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Truter M, Wehner FC (2004) Crown and root infection of lisianthus caused by Fusarium solani in South Africa. Plant Dis 88(5):573. doi:10.1094/PDIS.2004.88.5.573A Uematsu S, Okubo H, Suzui T, Shiota A, Chiba T (1996) First report of phytophthora rot of eustoma grandiflorum caused by phytophthora spp. Ann Phytopathol Soc Jpn 62(3):266, http://ci.nii.ac. jp/naid/110002733867/ (in Japanese). Accessed 22 Oct 2016 Uga H, Kobayashi YO, Hagiwara K, Honda Y, Omura T (2004) Selection of an attenuated isolate of Bean yellow mosaic virus for protection of dwarf gentian plants from viral infection in the field. J Gen Plant Pathol 70:54–56. doi:10.1007/s10327-003-0091-6 USDA-NASS (2012) Floriculture: U.S. Summary. 8 pp http://www.nass.usda.gov/Statistics_by_ State/Ohio/Publications/Reports_by_Title/flor2012.pdf . Accessed 22 Oct 2016 van de Wetering F, Posthuma K, Goldbach R, Peters D (1999) Assessing the susceptibility of chrysanthemum cultivars to tomato spotted wilt virus. Plant Pathol 48:693–699 van der Burg AMM, De Kreij C (2003) Voorkomen uitval bij Lisianthus: Onderzoek op 3 praktijkbedrijven. Wageningen, Praktijkonderzoek Plant & Omgeving B.V. PPO rapport nr.420002. http://www.tuinbouw.nl/sites/default/files/documenten/00021689.pdf (in Dutch). Accessed 22 Oct 2016 van der Wurff A, Hamelink R (2007) Middelentoets uitval Lisianthus: Vergelijkende effectiviteitsproef van middelen tegen Fusarium avenaceum en Myrothecium roridum in Lisianthus (Eustoma sp.) cv. Picolo White Wageningen UR Glastuinbouw (in Dutch). http://edepot.wur.nl/ 272656. Accessed 22 Oct 2016 Veeracone et al (2015) A review of the plant virus, viroid, liberibacter and phytoplasma records for New Zealand. Aust Plant Pathol 44:463–514. doi:10.1007/s13313-015-0366-3 Vetten HJ, Koenig R (1983) Natural infection of tomato and pelargonium in Germany by a tombusvirus originally described from pepper in Morocco. Phytopathol Z 108:215–220 Vrind TA (2005) The Botrytis problem in figures. Acta Hortic 669:99–102, http://www.actahort. org/books/669/669_11.htm. Accessed 22 Oct 2016 Wang K-H, McSorley R (2004) Host status of several cut flower crops to the root-knot nematode, Meloidogyne incognita. Nematropica 35(1):37–44 Webster CG, Frantz G, Reitz SR, Funderburk JE, Mellinger HC, McAvoy E, Turechek WW, Marshall SH, Tantiwanich Y, McGrath MT, Daughtrey ML, Adkins S (2015) Emergence of groundnut ringspot virus and tomato chlorotic spot virus in vegetables in Florida and the Southeastern United States. Phytopathology 105(3):388–398. doi:10.1094/PHYTO-06-14-0172-R Wegulo SN, Vilchez M (2004) Evaluation of fungicides for control of Botrytis blight of lisianthus. Fungicide Nematicide Tests 61:OT030 Wegulo SN, Vilchez M (2007) Evaluation of lisianthus cultivars for resistance to Botrytis Cinerea. Plant Dis 91:997–1001 Weintraub PG, Zeidan M, Spiegel S, Gera A (2007) Diversity of the known phytoplasmas in Israel. Bull Insectol 60(2):143–144 Whitby PW, Pope LC, Carter KB, LiPuma JJ, Stull TL (2000) Species-specific PCR as a tool for the identification of Burkholderia gladioli. J Clin Microbiol 38:282–285 Wintermantel WM, Hladky LL (2013) Complete genome sequence and biological characterization of Moroccan pepper virus (MPV) and reclassification of Lettuce necrotic stunt virus as MPV. Phytopathology 103:501–508 Wolcan SM (2005) Occurrence of Pseudocercospora eustomatis on Eustoma grandiflorum in Argentina. Australas Plant Pathol 34(4):617–618 Wolcan S, Ronco L, Dal Bo E, Lori G, Alippi H (1996) First report of diseases on Lisianthus in Argentina. Plant Dis 80:223. doi:10.1094/PD-80-0223A Wolcan S, Lori G, Ronco L (2001a) First report of Fusarium solani causing stunt on lisianthus. Plant Dis 85(4):443, http://dx.doi.org/10.1094/PDIS.2001.85.4.443C. Accessed 22 Oct 2016 Wolcan SM, Lori GA, Ronco L, Mitidieri AF, Fernandez R (2001b) Enanismo y podredumbre basal de Eustoma grandiflorum y su relación con la densidad de Fusarium solani en el suelo. Fitopatol Bras 26:710–714, http://www.scielo.br/pdf/fb/v26n4/8177.pdf (in Spanish). Accessed 22 Oct 2016

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Yang HC, Hsieh TF (1998) Salisb Plant Protect Bull (Taichung) 40(1):37–48 Yoshimatsu H (1993) Root and foliage rot of Eustoma grandiflorum caused by Rhizoctonia solani Kuhn. Ann Phytopathol Soc Jpn 59(3):284 (in Japanese) Young JM, Saddler GS, Takikawa Y, De Boer SH, Vauterin L, Gardan L, Gvozdyak RI, Stead DE (1996) Names of plant pathogenic bacteria 1864–1995. Rev Plant Pathol 75:721–763 Yunis H, Elad Y (1989) Survival of Botrytis cinerea in plant debris during summer in Israel. Phytoparasitica 17:13–21 Zen S, Okuda M, Fuji S, Iwanami T (2008) The seasonal occurrence of viruliferous Thrips tabaci and the incidence of Iris yellow spot virus disease on lisianthus. J Plant Pathol 90(3):511–515 Zen S, Nakashima S, Tashiro N, Okuda M, Fuji S (2010) Control of lisianthus necrotic ringpot disease caused by Iris yellow spot virus using reflective net and insecticide application adjusted to the immigrating periods of viruliferous onion thrips, Thrips tabaci Lindeman. Jpn J Phytopathol 76:17–20, https://www.jstage.jst.go.jp/article/jjphytopath/76/1/76_1_17/_pdf. Accessed 22 Oct 2016 Zulfiqar M, Brlansky RH, Timmer LW (1996) Infection of flower and vegetative tissues of citrus by Colletotrichum acutatum and C. gloeosporioides. Mycologia 88:121–128

Diseases of Orchid

21

Prasartporn Smitamana and Robert J. McGovern

Abstract

Orchids are monocotyledonous plants that belong to the Orchidaceae family which has a diverse range of habitats from tropical to temperate zones. Orchids grow at different elevations from sea level to the very high mountainous level of the Himalayas. Due to great variation, orchids can be classified in many genera and can be infected with many of the same fungi, bacteria, viruses, and nematodes as other plants. In this chapter, economically important diseases of orchids and their management are described. Keywords

Botrytis • Phytophthora • Pythium • Fusarium • Rhizoctonia • Sclerotium • Acidovorax • Dickeya • Burholderia • Viruses • Foliar Nematodes

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Black Rot (Phytophthora and Pythium spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Shoot Rot or Top Rot [Phytophthora nicotianae (syn. Phytophthora parasitica); Phytophthora cactorum] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Botrytis Spot (Botrytis cinerea) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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P. Smitamana (*) Agricultural and Industrial Clinic Co. Ltd., Chiang Mai, Thailand Department of Plant Pathology, Faculty of Agriculture, Chiang Mai University, Chiang Mai, Thailand e-mail: [email protected] R.J. McGovern NBD Research Co. Ltd., Lampang, Thailand Department of Entomology and Plant Pathology, Chiang Mai University, Chiang Mai, Thailand e-mail: [email protected] # Springer International Publishing AG 2018 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florists’ Crops Diseases, Handbook of Plant Disease Management, https://doi.org/10.1007/978-3-319-39670-5_21

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2.4 Petal Blight (Curvularia eragrostidis, Alternaria alternata) . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Sclerotium Stem Rot (Sclerotium rolfsii) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 Black Leg/Dry Rot and Slow Decline (Fusarium spp. Rhizoctonia solani) . . . . . . . . . 2.7 Leaf Spots (Fusarium, Phyllosticta, Guignardia, Septoria, Cercospora spp.) . . . . . . 2.8 Anthracnose [Colletotrichum gloeosporioides (syn. Glomerella cingulata)] . . . . . . . . 2.9 Rust (Sphenospora kevorkianii) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Bacterial and Phytoplasma Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Leaf Spot (Acidovorax avenae subsp. cattleyae) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Soft Rot [Dickeya chrysanthemi (syn. Pectobacterium chrysanthemi); D. dieffenbachiae] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Black Spot [Burkholderia gladioli (syn. Pseudomonas gladioli)] . . . . . . . . . . . . . . . . . . . 3.4 Phyllody (“Candidatus Phytoplasma” sp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Viral Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Nematode Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 Foliar Nematodes (Aphelenchoides spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1

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Introduction

Orchid belongs to the Orchidaceae which is a highly diverse family and can be found worldwide, but most species are found in the tropics, especially in the tropical mountains (Dressler 1993). Orchidaceae is possibly the largest family of the flowering plants and consists of 899 genera that include 71,391 species, but only 27,135 are accepted species names (The Plant List 2013). Among the orchids, 73% are epiphytes (Atwood 1986) and show stereotypical flower structure and remarkably diverse plant structure (Benzing 1986a, b). Orchids are economically important ornamental crops as potted plants and cut flowers but also provide raw materials for the fragrance and pharmaceutical industries. In some countries, orchids serve as the major source of income for growers. The commercial production of orchid began in natural habitats but has been adapted for large-scale industrial production. This change has made the environment conducive for the outbreak of many diseases. Due to vegetative propagation, the planting materials are the major source of inoculum, and pathogens are easily distributed, especially viruses. Major diseases of orchids and their control measures are listed below.

2

Fungal and Fungus-Like Diseases

2.1

Black Rot (Phytophthora and Pythium spp.)

Geographic occurrence and impact. Black rot is a common disease found in all orchid genera, especially after heavy rainfall or when in highly humid environments for extended periods, such as occur in poorly ventilated, crowded collections. Black rot is caused by one or both pathogens belonging to Phytophthora and Pythium spp. Fusarium can frequently occur as a secondary pathogen. Pythium ultimum, Phytophthora cactorum, and Phytophthora palmivora are commonly reported as the

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major pathogens (Hine 1962; Burnett 1974; Uchida 1994; Uchida and Aragaki 1991a; Orlikowski and Szkuta 2006; Cating et al. 2013). Phytophthora multivesiculata was found in Cymbidium in New Zealand (Ilieva et al. 1998; Hill 2004). Black rot is a common disease in all nurseries worldwide, especially in the tropics and subtropics, including Australia, Bulgaria, China, New Zealand, Poland, Taiwan, and the USA (Hawaii) (Burnett 1974; Chen and Hsieh 1978; Hall 1989; Duff 1993; Uchida 1994; Ilieva et al. 1998; Hill 2004; Orlikowski and Szkuta 2006; Tao et al. 2011). Symptoms/signs. In seedlings and young plantlets, water soaked lesions and discoloration occurs at the base of the plant and roots during periods of high humidity. This condition can lead to damping off. In monopodial type orchids, e.g., Vanda, Phalaenopsis, and Rhynchostylis, the disease usually starts on new leaves as brown to black lesions (Figs. 1a, b and 2b).

Fig. 1 Black rot lesions surrounded by a yellowish region on Vanda (a) and Rhyncostylis (b) orchids; infected root velamen has dried out (b) (P. Smitamana # 2017. All Rights Reserved.)

Fig. 2 Black rot of Catteleya pseudobulb (a) and Vanda leaves (b) on which the fungal mycelia are visible under high moisture conditions (a, P. Smitamana # 2017. All Rights Reserved; b, Kamjaipai 1983)

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The pathogens rapidly invade downward into the stem and roots. The infected shoot can easily be pulled out and, if unattended, the whole plant will die. In the sympodial type orchids, e.g., Cattleya, Dendrobium, and Oncidium, a light yellow-brown discoloration on the pseudobulbs is observed. After the disease progresses, softening and rotting of the pseudobulbs which turn brown to black occurs (Fig. 2a). The infected roots become darkly discolored as the velamen rots. Leaf symptoms first appear on the underside as small, water-soaked, irregular spots that are yellowish brown in color, then turn brown or black with a yellowish margin. Under high temperature and humidity, the lesions rapidly enlarge and become soft with ooze if pressed. If the humidity decreases, the lesions dry and turn black into which other pathogens can easily invade. Pathogens can also infect the leaves via wounds. In general, root symptoms initially appear as brown to black rot of root tips; the stem and/or pseudobulbs may rapidly rot. Infected roots can also be stunted, lose the velamen, and dry out. In Cymbidium, a dry rot of leaves can occur that has alternating horizontal light- and dark-brown zebra-like stripes. The pseudobulb tissue usually becomes wet and brown-black (Ilieva et al. 1998). Biology and epidemiology. Phytophthora and Pythium are fungus-like, but not true fungi, and are favored by high humidity. Both species spread rapidly if free water is available giving rise to zoospores that swim freely to, penetrate, and infect plant tissue (Agrios 2005). Following infection, mycelia spread rapidly through the infected tissues. The lesions rapidly expand and exhibit brown-black necrotic regions. Motile zoospores produced in the infected lesions are spread easily in irrigation water and are splash-dispersed on uninfected tissue during watering (Uchida 1994). Management • Cultural practices – Carefully check newly acquired plants to be sure that they are free from disease. Separate new stock from other plants and monitor them for disease symptoms for least 6 weeks before moving them into the nursery. Elevating the plants 60–90 cm above the ground to avoid splashing from the soil below or keeping them on a solid surface can also help prevent infections. Growers should maintain good ventilation through adequate plant spacing to reduce duration of leaf wetness. • Sanitation – If early symptoms of black rot are found, immediately excise infected tissue with a clean knife and discard it in order to prevent the spread of the disease on that plant or to other plants. Application of a fungicide (fosetyl-al, mefenoxam, etc.) may be effective when used immediately after. Use only clean water for watering orchids. Deep well water is safer than surface water unless surface water is disinfested. Discard or burn severely infected plants to prevent the spread of the disease. • Fungicides and biocontrols – Use suitable fungicides (fosetyl-al, mefenoxam, promocarb hydrochloride, trifloxystrobin, etc.) as a preventative measure, particularly during hot humid periods. The highest rate of the harpin protein, which induces systemic acquired resistance, significantly reduced black rot in Vanda

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caused by P. palmivora compared to the control; Bacillus subtilis + Trichoderma harzianum and etridiazole + quintozene were ineffective (Maketon et al. 2015). Growers should follow all label instructions.

2.2

Shoot Rot or Top Rot [Phytophthora nicotianae (syn. Phytophthora parasitica); Phytophthora cactorum]

Geographic occurrence and impact. This is a common disease found mostly on Vanda (upper stem) and Cattleya (leaf shoot) and is favored by high humidity and temperature, and poor ventilation due to high plant densities. This disease is caused by Phytophthora nicotianae, which can cause the death of the whole plant (Vanda); or in the case of Cattleya caused by Phytophthora cactorum, which causes rot of the shoots, leaves, pseudobulbs, rhizomes, and flower buds. Like black rot, shoot rot is commonly found in humid and high temperature environments especially in the tropical and subtropical zones: Thailand, USA (Florida, Hawaii), Poland, and Australia (Daly et al. 2013; Kamjaipai 1983; McMillan et al. 2009; Orlikowski and Szkuta 2006; Uchida 1994; Uchida and Aragaki 1991a, b). Symptoms/signs. A symptom of top rot includes the death of new leaves, which turn dark brown. This discoloration advances down the stem similar to the symptom caused by black rot. This disease may also start at the base of the stem and spread upward resulting in the same dark brown stem discoloration. Shoot rot was detected on Cattleya orchids in Darwin, Australia, and in USA (Florida) causing the rapid death of new side shoots, turning them almost black. The pathogen will spread back along the rhizomes to the next shoot causing the same symptoms. This disease has been reported on only Cattleya and Vanda but may affect others. Biology and epidemiology. As stated before, Phytophthora is fungus-like and its infection and spread are favored by wet conditions. Phytophthora cactorum is a homothallic oomycete that produces a sexual spore, an oospore, for survival under unfavorable environments. In addition, this fungus also asexually produces survival spores called chlamydospores as well as sporangia. The pathogen spreads by means of the pear or lemon shaped motile zoospores produced by either oospores or sporangia under wet conditions. After encysting and germination, zoospores can penetrate the plant directly or enter through wounds to cause infection. Management • Sanitation and fungicides – Immediately cut the infected top with a clean knife, and discard severely infected plants to prevent the spread of the pathogen, and spray the plants with a fungicide (fosetyl-al, mefenoxam, etc.). Use suitable fungicides (see Sect. 2.1 above) as a preventative measure, particularly during hot humid periods, following label instructions. Improve ventilation to reduce leaf wetness.

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• Biocontrol – Certain biocontrol products containing Streptomyces lydicus, Bacillus subtilis, Trichoderma species, and Gliocladium virens applied prior to the disease occurring in the growing area have been reported to help prevent some soilborne and aerial diseases (Bottom 2016). Spraying infected plants with a spore suspension of Trichoderma harzianum or T. viride, or in the case of Vanda directly injecting the solution into the infected shoot, may be helpful (Smitamana, unpublished data).

2.3

Botrytis Spot (Botrytis cinerea)

Geographic occurrence and impact. Botrytis spot is a fungal disease affecting many types of orchids, mostly during cool, damp weather and in nurseries with poor air circulation. Phalaenopsis and Cattleya are particularly susceptible to Botrytis cinerea infection, but the disease may be found in a wide range of orchid genera. This disease has been reported in the tropics and the temperate regions including the USA (Florida, Hawaii) and Thailand (Jones 2003; Kamjaipai 1983; Uchida and Aragaki 1991b). Symptoms/signs. Older flowers are highly susceptible to infection. Brown necrotic or pale spots on orchid flowers are typical symptoms. The spots may enlarge and increase in number as the infection progresses and may be surrounded by a yellowish or pale pink margin depending on the background color of the flowers. Under cool and high moisture conditions, fungal mycelial webbing may be visible. Biology and epidemiology. Botrytis cinerea is an airborne plant pathogen commonly found in nurseries. Hyaline conidia of the fungus are produced on grey, branching conidiophores. Sclerotia, highly resistant survival structures, can be produced on dying infected tissue. In the temperate zone, B. cinerea overwinters in dead and dying plant material and begins producing and dispersing spores during cool, damp weather in the spring and autumn. Spores are distributed by wind, rain, irrigation water, or any mechanical action. Temperatures in the range of 18–25  C/ 64–77  F and wet plant surfaces or at least 92% RH are conducive factors for infection, and the pathogen can quickly proliferate, infecting the surrounding healthy plant tissue in less than 14 h (Agrios 2005; Sumbali 2005).

Management • Cultural practices – Growers should avoid wetting the flowers as much as possible. In tropical nurseries, the growers always spray or water the whole nonflowering plants early in the morning to make sure that the plants will be dry in the late afternoon or by nightfall. Water that remains on petals or leaves after a rain or watering promotes fungal growth. Avoid damp environments and poor air circulation that promote pathogen growth. Facilitate good air circulation through proper plant spacing and decrease the humidity during cool, damp

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weather by heating and venting greenhouses in temperate climates, especially when plants are in the blooming stage. • Sanitation – Remove any plant debris, fallen flowers, leaves, and infected tissue from the growing area in closed containers to reduce the infection potential and the possibility of spreading the fungus. Burning or burying all affected plant tissue is recommended. Eliminate unwanted residual ornamental crops and weeds in the growing area. Make sure that newly introduced plants are free of any disease; normally keep them in an isolated area for at least 2 weeks before bringing them into the nursery. • Fungicides – Spray preventively with fungicides such as chlorothalonil, thiophanate methyl, iprodione, or vinclozolin. Resistance to thiophanate methyl and vinclozolin has been widely reported in B. cinerea; therefore, rotation of these fungicides with those of different modes of action is essential.

2.4

Petal Blight (Curvularia eragrostidis, Alternaria alternata)

Geographic occurrence and impact. This disease is found mostly in the Dendrobium hybrids whose flowers are most susceptible to the pathogens. Petal blight has been reported in Australia, Thailand, and the USA (Burnett 1957, 1965; Daly et al. 2013; Kamjaipai 1983). Symptoms/signs. Small brown spots develop on the flower petals, then the spots enlarge and merge to form necrotic lesions; under high moisture conditions or after rainfall these spots will rot (Fig. 3a, b). Biology and epidemiology. Alternaria and Curvularia belong to the fungal family Dermatiaceae which produces both dark hyphae and conidia. No organized fruiting bodies are formed, and the majority of these groups are saprophytic but some are plant pathogens (Sumbali 2005). Conidia of Alternaria are typically club or pear shaped and multicellular with both transverse and longitudinal cross walls. Conidia are easily detached and carried by air currents (Agrios 2005).

Fig. 3 Petal blight of Dendrobium flowers: water-soaked lesions turning brown or rust colored on white flower petals (a); and whitish yellow lesions on “Madame Pompadour” flowers (b). (Kamjaipai 1983)

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Curvularia conidia are curved, clavate, broadly fusiform, obvoid, or pyriform with three or more transverse septa, pale or dark brown, with the end cells paler than the others (Ellis 1977). Both Alternaria and Curvularia can survive in plant debris, infected, or dry tissue, and the conidia are easily spread by air currents or water drops splashed off the orchid plants during overhead irrigation. Management • Cultural practices – Keep orchids in bloom under solid rooves in order to protect them from rain. Avoid damp environments caused by incorrect watering practices and poor air circulation that promote pathogen infection. • Fungicides and biocontrols – The highest rate of the harpin protein, which induces systemic acquired resistance, significantly reduced rust spot in Dendrobium flowers caused by Curvularia lunata, compared to the control. Formulations of Bacillus subtilis + Trichoderma harzianum and mancozeb were intermediate in effectiveness (Maketon et al. 2015).

2.5

Sclerotium Stem Rot (Sclerotium rolfsii)

Geographic occurrence and impact. This disease is found mostly in Ascocenda, Brassidium, Cymbidium, Dendrobium, Neofinetia, Phaius, Paphiopedilum, Phalaenopsis, and Vanda orchids. Stem rot is commonly found in the tropics especially in nurseries that do not have roofs to protect the foliage from rainfall. Pots with poor drainage, such as those used to grow Paphiopedilum, are conducive to the disease. Stem rot has been reported in both the tropics and temperate zones including Australia, India, Korea, Thailand, and the USA (Florida) (Bag 2004a, b; Cating et al. 2009a; Cating et al. 2013; Daly et al. 2013; Han et al. 2012; Kamjaipai 1983; Tara et al. 2003). Symptoms/signs. The symptoms of this disease usually start on the lower shoots, pseudobulbs, and leaves which turn yellow, grow poorly, and then rapidly collapse (Fig. 4a). Rotting of the stem (Fig. 4c), roots, and leaf tissue gradually occurs, and in severe infections the entire plant wilts and dies. White fluffy mats of fungal mycelia and small yellow cream to brown spherical, mustard seed-size bodies called sclerotia appear on infected tissue (Fig. 4d). On Dendrobium plants, the pathogen can infect the leaves and basal part of pseudobulbs on which white mycelia are found; or in certain cases, the plants can be infected from the top causing rapid collapse and rotting of shoot apices (Fig. 4b). On Phaius flavus and Paphiopedilum venustum the disease may be found on the base of the pseudobulbs and collar region, respectively, and causes rot and leaf yellowing or collapse, which spreads upward, until the entire plant turns brown to black and dies. Diagnostically characteristic white mycelia and small sclerotia are found on the infected areas.

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Fig. 4 Infected Vanda basal shoot with white mycelia and small sclerotia (a), young Dendrobium plant with severe leaf rot and white mycelia (b), severely infected Vanda showing dry rot of stem (c), and sclerotia on the infected stem and leaves (d) (Kamjaipai 1983)

Biology and epidemiology. Sclerotium rolfsii is characterized by small cream to dark brown sclerotia produced on septate, white fluffy mycelia with hyphal clamp connections (Sumbali 2005). Sclerotia can be spread by water and act as sources of infection. The fungus has a wide host range and is favored by a warm, moist environment. In the temperate zone, the pathogen can overwinter in the sclerotia and starts infecting new plants when the temperatures become warm. Sclerotia can survive in soil for many years.

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Management. Managing Sclerotium stem rot is quite difficult because of the pathogen’s wide host range and long survival in soil via sclerotia. Infected plants should be immediately destroyed, and an appropriate fungicide such as PCNB should be applied when the disease is observed. Using biocontrol agents, such as Trichoderma spp., as an alternative measure for preventing and controlling the pathogen is also recommended.

2.6

Black Leg/Dry Rot and Slow Decline (Fusarium spp. Rhizoctonia solani)

Geographic occurrence and impact. Fusarium solani, F. oxysporum, F. subglutinans, and F. proliferatum have been associated with the disease; in addition, Rhizoctonia solani has also been involved in some areas. Susceptible orchids include Dendrobium, Cattleya, Phalaenopsis, Vanda, Rhyncostylis, and Vanilla. Black leg, dry rot, and slow decline have been reported in the tropics and temperate zones including Australia, Brazil, Indonesia, Malaysia, Taiwan, Thailand, the USA (Florida) (Daly et al. 2013; Dehgahi et al. 2014; Kamjaipai 1983; Pinaria et al. 2010; Su et al. 2010; Pedroso-de-Moraes et al. 2011; Wedge and Elmer 2008). If uncontrolled, Fusarium rot can affect up to 100% of an orchid crop (Kawate and Sewake 2014). Fusarium is considered a quarantine pest. Symptoms/signs. This disease is mostly found in orchids grown in damp environments or waterlogged pots especially after heat stress and heavy fertilization (Wedge and Elmer 2008). Plantlets after transferring from tissue culture vessels to community trays or pots are most susceptible to infection. Infected plantlets show water soaked root tips followed by leaf yellowing in whole plantlets. In mature plants, the symptoms are mostly observed after transplanting when the pathogens can easily infect the plants through wounds. Infected plants show discoloration of the root tips and a gradual dry rot. Pseudobulbs become spongy and discolored, successive new shoots tend to get smaller in height, and stem thickness is also reduced. The leaves will yellow and drop off, one by one, until none are left and the plant dies (Fig. 5). Biology and epidemiology. Fusarium and Rhizoctonia are both soilborne pathogens. Fusarium at first produces colorless mycelia but with age the mycelia color will change to cream, pale yellow, pale pink, or purplish. Fusarium produce three kinds of asexual spores: microconidia which have 1–2 cells; curved macroconidia which have 3–5 cells, and 1–2 cell, thick-walled resistive chlamydospores (Agrios 2005). The microconidia and macroconidia are dispersed easily by air currents and spread with rain and irrigation water. Fusarium can also be spread by fungus gnats. Rhizoctonia produces sterile colorless mycelia which will change to yellowish or light brown with age. Hyphae consist of long cells, with branching at an approximate right angle to the main hypha with slight hyphal constrictions at the junction. Under certain conditions, Rhizoctonia produces sclerotia which function as survival structures (Agrios 2005). The fungus can overwinter as mycelia or sclerotia in or on

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Fig. 5 Primary symptoms of Vanda plantlet rot caused by Fusarium spp. include root tip necrosis in the community tray (a); yellowish leaves and leaf drop, dark brown roots, and plantlet death (b); Dendrobium roots showing brown discoloration with white fungal mycelia, the root system totally dried out and the pseudobulb turning brown (c); Paphiopedilum showing decline symptoms including yellowish leaves and brown and rotten collars caused by Rhizoctonia sp. from contaminated coconut husks used as the planting medium (d) (P. Smitamana # 2017. All Rights Reserved.)

infected plants or by contaminating planting materials and containers, e.g., coconut husks and pots. The fungus spreads by rain or irrigation water. Management • Resistance – Brassocattleya “Orquidacea’s Melody” and Brassocattleya “Orquidacea’s Rare Bird,” were resistant and moderately resistant, respectively, to Fusarium wilt caused by Fusarium oxysporum f. sp. Cattleyae (Pedroso-deMoraes et al. 2011). • Cultural practices – Growers should inspect newly introduced plants and keep them in an area removed from general production for 4–6 weeks to observe the health of the plants and make sure that they are free from any diseases. Transplant uninfected pseudobulbs into pathogen-free media and new pots. Adjust the pH of the irrigation water and medium to 6.0–6.5 to help reduce infection by Fusarium. • Sanitation – For slightly infected Vanda, Vanilla, and Rhyncostylis with good aerial roots, cut above the diseased portion using a sterile cutting tool, pot in a fresh medium, remove and discard or burn infected tissue, and apply an appropriate fungicide or biocontrol (see below).

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• Fungicides and biocontrols – Use biocontrol agents, e.g., Trichoderma harzianum, by incorporation into fresh media in order to prevent contamination by pathogens. If chemical control is needed, apply cyprodinil + fludioxonil, chlorothalonil, azoxystrobin, or thiophanate methyl when early symptoms are observed, being sure to rotate different classes of fungicides. Prochloraz and tebuconazole were very effective in reducing Fusarium bulb and root rot in Cymbidium goeringii (Jee et al. 2003). Quaternary ammonium salts, sodium hypochlorite, and other materials should be used to disinfest benches and containers (see ▶ Chap. 9, “Sanitation for Management of Florists’ Crops Diseases” for more information).

2.7

Leaf Spots (Fusarium, Phyllosticta, Guignardia, Septoria, Cercospora spp.)

Geographic occurrence and impact. Leaf spots of orchids can be caused by many fungi. Symptoms in the diseased orchid are varied based on the causal agent and host. Many orchids are susceptible to these pathogens, e.g., Brassolaeliocattleya, Cattleya, Cymbidium, Dendrobium, Epidendrum, Laelia, Laeliocattleya, Odontoglossum, Oncidium, Phalaenopsis, and Vanda. Leaf spots of orchids are found worldwide: Australia, Brazil, Japan, the Netherlands, Thailand, and the USA (Hawaii) (Daly et al. 2013; Ichikawa and Aoki 2000; Jones 2003; Kamjaipai 1983; Silva and Pereira 2007; Silva et al. 2008; Uchida and Aragaki 1980; Uchida 1994). Leaf spots are common in most nurseries and can lead to serious losses in shipping containers due to the high moisture and low light conditions which favor the growth and infection of the pathogens. Symptoms/signs Fusarium spp. – Two spot types; yellow and black are caused by F. subglutinans and F. proliferatum, respectively, on Cymbidium. The yellow spot symptoms start as water soaked patches on the leaves then the lesions enlarge with sunken centers and turn reddish-brown surrounded with a yellowish swelling without a definitive boarder. In the case of black spot, small black speckles are found at the early stage, then the lesions enlarge to form irregular, angular black spots; some yellow halos are found in certain Cymbidium cultivars. Phyllosticta spp. – Symptoms may start anywhere on the leaf, flower, or pseudobulb. Tiny yellow and slightly sunken spots are initially observed on the leaves. As the lesions enlarge, they become more sunken with round or oval shapes (Fig. 6a). The fungal growth within the tissue turns the lesion tan or brownish as the disease develops. Severely infected leaves may drop prematurely and a black web-like pattern formed by the fungus is observable (Fig. 6b). Spot symptoms on flowers are small, faint, and commonly blue or lavender on purple cultivars. Guignardia spp. – Symptoms start as tiny, dark purple, elongated lesions which can be found on either leaf surface. With age, the lesions enlarge and run parallel to the veins and form elongated purple streaks. Large irregular lesions are formed by

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Fig. 6 Leaf spots caused by Phyllosticta sp. in Vanda: sunken lesions (a); merging lesions with black fungal growth in the leaf tissue (b); yellow green area with brown spots on Dendrobium leaf caused by Cercospora sp. (c) (P. Smitamana # 2017. All Rights Reserved.)

the merging of the spots, and the center of the lesions turns tan. Black fruiting bodies of the fungus develop in the affected area and can be seen as tiny raised black spots. Cercospora spp. – Cercospora attacks the underside of the leaf and induces small yellow spots. After the disease progresses, yellow green areas are observed on the top surface of the leaf (Fig. 6c). Subsequently, the spots enlarge and form irregular patterns with slightly sunken areas which turns brown to black. The necrotic lesions can enlarge to cover the whole leaf area and severely infected leaves may drop prematurely. Septoria spp. – Septoria can infect both sides of the leaf. Primary symptoms start from small sunken yellow lesions which gradually enlarge and form dark brown to black, circular, or irregular lesions. Lesions may merge to form irregular areas on the leaf which may prematurely drop. Biology and epidemiology Phyllosticta belongs to the Sphaeropsidales and produces asexual spores in pycnidia, small, black globose, elongate, or cup-like fruiting bodies. Pycnidia may be superficial or immersed in the tissue. Conidia are hyaline, single-celled, globose, or ovoid (Sumbali 2005). High light levels and dry conditions inhibit the growth and spread of the fungus, whereas low-light and high humidity, or rainy weather, promote Phyllosticta outbreaks (Jones 2003). Guignardia is the perfect stage of Phyllosticta and belongs to the Ascomycetes, and produces eight single celled ascospores in perithecia (Silva and Pereira 2007).

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Cercospora belongs to the Dermatiaceae and produces dark colored hyphae and long slender hyaline or dark, straight to curved, multicellular conidia on short dark conidiophores. Clusters of conidiophores arise from the plant surface through the stoma and form conidia on the growing tips (Agrios 2005). No fruiting bodies are formed (Sumbali 2005). Septoria belongs to the Sphaeropsidales like Phyllosticta and produces long, slender, hyaline, generally curved conidia with one or more septa (Sumbali 2005). Conidia of these fungi are easily detached from the conidiophores, and ascospores discharge from perithecia into the air and are carried great distances. Conidia are splashed from older diseased leaves to other leaves nearby and infect new plants. Management • Cultural practices – Good sanitation with adequate air movement is important to control these diseases. Discard severely infected plants. Collect all fallen leaves, old or dry pseudobulbs, and sheaths from pots, benches, and the ground and discard or burn them. Water the orchids early in the day to allow leaves to dry before dark. • Fungicides – Spray with mancozeb, chlorothalonil, or another effective fungicide to kill the fungal spores and reduce the inoculum level (see ▶ Chap. 7, “Fungicides and Biocontrols for Management of Florists’ Crops Diseases” for more information).

2.8

Anthracnose [Colletotrichum gloeosporioides (syn. Glomerella cingulata)]

Geographic occurrence and impact. This is a disease that commonly affects Dendrobium, Vanda, Cattleya, Phalaenopsis, and Ceologyne orchids. Anthracnose in orchids has been reported in Australia, India, Thailand, and the USA (Florida) (Bottom 2016; Chowdappa et al. 2014; Daly et al. 2013; Jadrane et al. 2012; Kamjaipai 1983; McMillan 2011; Prapagdee et al. 2008; Prapagdee et al. 2012). This disease is considered only a minor pathogen and is usually a result of some type of injury to the leaf, whether it is mechanical, chemical, or insect damage. Nevertheless, anthracnose can result in reduced plant quality and growth; disease development may be sporadic as it is affected by levels of pathogen inoculum and environmental conditions. Symptoms/signs. Colletotrichum infects the aerial portion of the plant of which leaves are most often attacked. Infected leaves show brown discolorations which are irregularly shaped sunken lesions. Leaf tips turn brown and the affected area gradually enlarges toward the base. With age, the lesions become dark brown or light gray patches, and sometimes brownish black concentric rings of fruiting bodies or numerous dark bands develop across the leaf (Fig. 7). Flower infection is characterized by water soaked spots and black or brown pustules which are usually raised

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Fig. 7 Typical symptom of anthracnose on the black orchid (Ceologyne pandurata): the early symptom showing a brownish water-soaked area starting at the leaf tip (a); later stage lesions showing dark concentric rings on the upper leaf surface (b) and lower leaf surface (c) (P. Smitamana # 2017. All Rights Reserved.)

and occur on the underside of old sepals and petals. The spots may merge and cover the entire flower. In white flowered Phalaenopsis, the infected petals show anthracnose-like, sunken lesions which are surrounded by a ring of green tissue. Biology and epidemiology. Colletotrichum gloeosporioides is a member of the Melanconiaceae and is the imperfect stage of the ascomycete Glomerella cingulata. The conidia are produced in acervuli and are single celled, hyaline, ovoid, cylindrical, typically elongated with round ends and slightly narrower in the middle. Large brown or black setae are formed among the conidiogenous cells (Agrios 2005; Sumbali 2005). The conidia are disseminated by air currents, wind, rain, or irrigation water. The pathogen is most active in warm weather when light is low and moisture is high. Colletotrichum can survive or overwinter on plant residue in soil or on infected plants. Anthracnose is common after orchids pass through a stress period, e.g., cold or hot temperature, mechanical injuries, or chemical damage. Management • Cultural practices – Good sanitation and air movement, lower temperatures (if possible), and increased light may help reduce the spread and severity of this disease. • Fungicides and biocontrols – Alternation of protectant fungicides like mancozeb and systemic fungicides such as thiophanate methyl is recommended. The bacterium Bacillus subtilis was shown to reduce anthracnose caused by C. gloeosporiodes in Paphiopedilum concolor in Thailand (Kuenpech and Akarapisan 2014). Application of culture filtrates of the bacterium Streptomyces hygroscopicus prevented anthracnose on orchid (Propagdee et al. 2008).

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Rust (Sphenospora kevorkianii)

Geographic occurrence and impact. Sphenospora kevorkianii has been found infecting orchids in Brazil and in the USA (Linder 1944; Pereira et al. 2002; Pereira and Silva 2009). Known hosts are Cyrtopodium punctatum, Epidendrum difforme, E. secundum, E. xanthinum, Notylia lyrata, Pleurothallis mentigera, Prescottia sclerophylla, Stanhopea graveolens, and Zygostates lunata which are epiphytes found in tropical forests. Symptoms/signs. Rust fungi are very easy to identify from their rust colored spore masses on leaves and stems. The symptoms start out as flecks or spots and grow into bumps, and most commonly appear on the underside of leaves. Biology and epidemiology. Sphenospora kevorkianii produces uredospores in ovoid, ellipsoid, or subspherical uredinia on the lower leaf surface (Pereira et al. 2002). The uredospores are easily detached and dispersed by wind, rain, and irrigation water. Management. Cut off and discard or burn the infected parts. Discard or burn severely infected plants. Reducing nitrogen fertilizer application may help. If needed, spray with appropriate systemic fungicides (see ▶ Chap. 7, “Fungicides and Biocontrols for Management of Florists’ Crops Diseases” for more information).

3

Bacterial and Phytoplasma Diseases

3.1

Leaf Spot (Acidovorax avenae subsp. cattleyae)

Geographic occurrence and impact. This disease has been reported in Australia, China, India, Italy, Korea, Taiwan, Poland, and the USA (Florida) (Borah et al. 2002; Cating and Palmeteer 2011; Ding et al. 2010; Hseu et al. 2012; Kim et al. 2015; Li et al. 2009; Lin et al. 2015; Pulawska et al. 2013; Scortichini et al. 2005; Stovold et al. 2001). Acidovorax avena is a pathogen of quarantine importance and causes serious losses to orchid production especially in Cattleya, Cypripedium, Dendrobium, Oncidium, Phalaenopsis, and Vanda. Symptoms/signs. Symptoms usually start as small brown soft water-soaked spots. The spots turn black forming cavities in the parenchyma which quickly expand over the entire leaf. Eventually, the pathogen invades the growing point of the plant causing death (Fig. 8a, b). Phalaenopsis, due to its succulent leaves, is generally reported in many countries as the most susceptible genus. Biology and epidemiology. Acidovorax avenae subsp. cattleyae is a gram-negative rod shaped, nonfluorescent bacterium, which moves using a single polar flagellum. The bacteria are easily spread by rain, irrigation, and contaminated tools commonly

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Fig. 8 Leaf spot of Phalaenopsis caused by Acidovorax avenae: brownish water soaked area which begins at a mechanical injury (a) and the soft rot of leaf (b) (P. Smitamana # 2017. All Rights Reserved.)

used in the nurseries. High moisture and temperature promote the growth of bacteria and accelerate the progress of disease development. Management. Management of bacterial orchid diseases is challenging and must rely on an integrated approach. • Cultural practices – Avoid overhead watering to minimize the dispersal of the pathogen in nurseries or water early in the morning to avoid long hours of leaf wetness. If the disease is found, reducing the use of nitrogen fertilizer and increasing potassium fertilizer has been associated with disease suppression. • Sanitation – Growers should discard severely infected plants. Trim and discard diseased areas from slightly infected plants and keep them in an area removed from production for monitoring. Early detection of disease outbreaks is essential. Apply a bactericide as soon as the disease is observed (see below). Clean tools after each use by soaking in a 10% bleach solution. • Bactericides – If the infection becomes widespread, growers may dip plants in a labeled quaternary salt material, orthophenylphenol, or natriphene, or spray with a copper compound. Formulations of hydrogen peroxide may help reduce the disease. If antibiotic use is allowed, streptomycin, tetracycline, or a mixture of both may be sprayed. Small-scale testing for phytotoxicity is recommended. However, reliance on chemical control alone for management of bacterial diseases of orchids is most often ineffective.

3.2

Soft Rot [Dickeya chrysanthemi (syn. Pectobacterium chrysanthemi); D. dieffenbachiae]

Geographic occurrence and impact. This disease mainly affects Oncidium, Vanda, Tolumnia, Cattleya, Dendrobium, Phalaenopsis, and Miltoniaorchids and has been reported in China, Taiwan, and the USA (Florida) (Cating et al. 2008;

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Cating et al. 2009b; Li et al. 2009; Cating et al. 2009b; Hseu et al. 2012; Keith and Sewake 2009; Zhou et al. 2012). This is an important quarantine disease that affects the export and import of orchids in many countries. Symptoms/signs. The symptoms are typical of other bacterial soft rots and start with water-soaked tissue at the infection site. On Oncidium, the initial spots are commonly found at the base of the stem, and the lesions expand and become elongated. Gradually, the stem and leaf tissues become soft and watery. Bacterial ooze can be observed by cutting the edge of infected stem and leaf tissues and dipping them in a beaker or Petri dish containing water; bacterial streaming can be observed from the cut tissue by eye or under a stereomicroscope. Infected Phalaenopsis plants initially show water-soaked, pale-to-dark brown pinpoint spots on leaves; some may be surrounded by a yellow halo. Under high humidity and temperature, the spots expand rapidly and extend over the leaf which turns a light tan color with a darker brown border in a few days. A typical odor of rotten tissue can be detected. When the bacteria invade the stem, plant death follows. In Miltonia, Cattleya, and Oncidium, symptoms start with small water-soaked lesions which become a dark brown to black color and gradually expand throughout the leaves which subsequently rot. Some Dendrobium cultivars appear to be resistant to this disease and only small water-soaked spots are produced. In the susceptible types, the spots expand and coalesce to form larger necrotic areas; after that, the infected leaves dry out. If the bacteria invade the pseudobulbs, they turn soft and watery. In Vanda, infected areas show water soaked spots, some are brown color, and the leaves are rapidly rotted. Biology and epidemiology. Dickeya spp. are gram-negative bacteria, facultative, anaerobic rods with peritrichous flagella. Infection by the pathogen normally starts in cutting wounds or other mechanical injuries. Drops of bacterial ooze from the infected leaves or stems are dispersed by rain, irrigation, and wind. The bacteria enter and colonize plants mostly at the basal part of leaves or pseudobulbs. Management. Management of soft rot is the same as for Acidovorax leaf spot listed above.

3.3

Black Spot [Burkholderia gladioli (syn. Pseudomonas gladioli)]

Geographic occurrence and impact. Burkholderia gladioli was first characterized as the causal agent of leaf and corm disease in gladiolus. This pathogen is now known to cause serious losses in many plants especially orchids, e.g., Phalaenopsis, Cattleya, Dendrobium Oncidium, Odontioda, and Miltonia. This pathogen is found in tropical and subtropical zones of such countries as China, the USA (Hawaii), Indonesia, Taiwan, and Thailand (Chuenchitt et al. 1983; Hseu et al. 2012; Joko et al. 2014; Kamjaipai 1983; Keith et al. 2005; Lee et al. 2013; Takahashi et al. 2004; Uchida 1995; You et al. 2016).

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Symptoms/signs. Infected leaves initially show small dark green to brown spots surrounded by a water-soaked area. Sometimes only small brown spots surrounded by yellow halos, usually circular, or in rare cases irregularly shaped occur. Under high humidity or after rain, the wet rot area rapidly expands and causes soft rot of the whole leaf, especially the succulent leaves of Phalaenopsis. The bacteria can invade the stem or pseudobulbs and sometimes cause defoliation. Under low humidity, the infected leaf may dry and crack in the center of the lesion. However, if the humidity increases, bacterial ooze can be seen on the cracks and the rotten tissues. Symptoms on the young plants spread rapidly and cause the whole plant to die. Biology and epidemiology. Burkholderia gladioli is a gram-negative rod that may be straight or slightly curved. It is aerobic, catalase positive, urease positive, and non-sporeforming. Like other phytopathogenic bacteria, it is easily spread by water splash from rain, overhead sprinklers, and wind-blown water. Management. Management of soft rot is the same as for Acidovorax leaf spot listed above.

3.4

Phyllody (“Candidatus Phytoplasma” sp.)

Geographic occurrence and impact. Only a Sarcochilus hybrid in Australia has been reported to be infected by a phytoplasma (Gowanlock et al. 1998). Symptoms/signs. The infected Sarcochilus hybrid exhibited phyllody, the abnormal development of floral parts into leafy structures, and stunting. Biology and epidemiology. Phytoplasmas are bacteria-like microorganisms lacking cell walls, but bounded by a unit membrane. Phytoplasmas are found located in the sieve tubes of phloem. Transmission of phytoplasmas is by phloem-feeding insects in the Cicadellidae family and Fulgoroidea superfamily such as leafhoppers and planthoppers, respectively. Management. Avoid introducing diseased plants into nurseries. Remove and burn symptomatic plants.

4

Viral Diseases

Geographic occurrence and impact. Viruses are among the most damaging pathogens of orchids because of their debilitating effects on flower production and quality and extreme difficulty to control in a vegetatively propagated crop. Orchids can be infected by a very wide range of viruses. The most common and economically important orchid viruses worldwide are Cymbidium mosaic virus (CymMV) and Odontoglossum ringspot virus (ORSV), also known as Tobacco mosaic virus-orchid strain (TMV-O); and these two viruses are often found in mixed infections. Other

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orchid viruses of note include: Orchid fleck virus (OFV), Impatiens necrotic spot virus (INSV), Vanilla mosaic virus (VanMV), and Calanthe mild mosaic virus (CalMMV). The geographic occurrence and orchid host range of these viruses are indicated in Table 1. Orchids have also been reported to host a large number of other viruses in the following genera: Carmovirus – Carnation mottle virus (CarMV) Closterovirus – Dendrobium vein necrosis virus (DVNV) Cucumovirus – Cucumber mosaic virus (CMV) Nepovirus –Tomato ringspot virus (ToRSV) Potyvirus – Bean yellow mosaic virus (BYMV); Ceratobium mosaic virus (CerMV); Clover yellow vein virus (ClYVV); Colombian datura virus (CDV); Dasheen mosaic virus (DsMV); Dendrobium mosaic virus (DeMV); Diurus virus Y (DVY); Habenaria mosaic virus (HaMV); Pecteilis mosaic Virus (PcMV); Phalaenopsis chlorotic spot virus (PhCSV); Spiranthes mosaic virus 2 (SpiMV2); Sarcochilus virus Y (SVY); Spiranthes mosaic virus 3 (SpiMV3); Turnip mosaic virus (TuMV); Vanilla necrosis virus (VanNV) Rhabdovirus – Colmanara mosaic virus (CoIMV); Dendrobium ringspot virus (DRV) Tobravirus – Tobacco rattle virus (TRV) Tombusvirus – Cymbidium ringspot virus (CymRSV); Lisianthus necrosis virus (LNV): Tomato bushy stunt virus (TBSV) Tospovirus – Capsicum chlorosis virus (CaCV); Tomato spotted wilt virus (TSWV) Symptoms/signs. Orchids can show variation in viral symptoms that depend on the virus, virus strain, mixed viral infections, and orchid variety. In addition, different viruses can cause similar symptoms in the same orchid, making diagnosis of virus infection by symptoms alone unreliable. Common symptoms of virus infection of orchids are presented in Table 1. Biology and epidemiology. As obligate pathogens, viruses need a living host for their survival and replication. However, the molecular structure of Tobamoviruses, such as ORSV, makes them resistant to degradation for long periods on tools and in soil when they are released from host tissues. A number of important orchid viruses, such as CymMV, ORSV, VanMV, and CalMMV, are mechanically transmissible in plant sap. This characteristic is very important for virus spread in a vegetatively propagated crop such as orchid. In addition, potyviruses including VanMV and CalMMV are vectored in a nonpropagative (nonpersistent) manner on the stylets of aphids such as Myzus persicae. INSV, a tospovirus, is vectored in a circulative and propagative manner (it enters and replicates within the hemolymph of the insect) by a number of thrips species including the Western flower thrips, Frankliniella occidentalis. Only the first and early second larval stages are able to acquire tospoviruses, and only immature thrips that acquire these viruses or adults derived from such immatures are vectors. Adult thrips remain viruliferous for life; but tospoviruses are not transovarial. INSV also infects a large number of other

Geographic occurrence Argentina, Brazil, China, Colombia, Cook Islands, Costa Rica, French Polynesia, Guam, India, Indonesia, Japan, New Caledonia, Netherlands, Niue, Palau, Puerto Rico, Reunion Island, Samoa, Singapore, Taiwan, Thailand, Tonga, USA, Vanuatu, Venezuela

Argentina, Brazil, China, Colombia, Cook Islands, Costa Rica, Fiji, French Polynesia, India, Indonesia, Japan, Netherlands, New Zealand, Niue, Puerto Rico, Reunion Island, Singapore, Taiwan, Thailand, Tonga, USA, Vanuatu, Venezuela

Virus, genus Cymbidium mosaic virus (CymMV), Potexvirus

Odontoglossum ringspot virus (ORSV), (also known as TMV-orchid strain and TMV-O), Tobamovirus

Table 1 Selected orchid viruses

Mechanical

Transmission Mechanical

A wide range including: Aerides, Bulbophyllum, Calanthe, Cattleya, Cymbidium, Dendrobium, Oncidium, Phalaenopsis, and Vanilla

Orchid hosts A wide range including: Aranthera, Arachnis, Calanthe, Cattleya, Cymbidium, Dendrobium, Gromatophyllum, Phalaenopsis, Phaius, Oncidium, Rynchostylis, Vanda, and Vanilla

Chlorotic or necrotic sunken lesions on leaves and flowers. In highly susceptible varieties, flowers may be smaller, distorted, and exhibit color breaking (Fig. 9c, d). Root tip necrosis is also a common symptom

Symptoms Chlorotic mosaic patterns, seen clearly on the youngest leaves in many orchid varieties, to black necrotic streaks, spots, or rings and sunken patches on Cattleya orchids. Flower size reduction with abnormal coloration and twisted young shoots with chlorotic streaks are common symptoms on young Dendrobium shoots (Fig. 9a, b)

Diseases of Orchid (continued)

References Baker et al. 2007a; Cánovas et al. 2016; Davis and Ruabete 2010; Elliot et al. 1996; Farreyrol et al. 2001; Freitas-Astua et al. 1999; Gara et al. 1996; Hu et al. 1993; Inouye and Gara 1996; Jensen 1952; Kamjaipai 1983; Sherpa et al. 2003; Singh et al. 2007; McMillan et al. 2006; Sutrabutra 1989; Tanaka et al. 1997; Wong et al. 1994; Zheng et al. 2010; Zhou et al. 2004 Cánovas et al. 2016; Davis and Ruabete 2010; Elliot et al. 1996; Farreyrol 2001; FreitasAstua et al. 1999; Hu et al. 1993; Inouye and Gara 1996; Kamjaipai 1983; McMillan et al. 2006; Pearson et al. 2006; Sherpa et al. 2006; Sutrabutra 1989; Tanaka et al. 1997; Thomson and Smirk 1967; Wong et al. 1994; Zhou et al. 2004

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Mechanical; (Myzus persicae) Mechanical; aphid

Japan French Polynesia, Reunion Island

Calanthe, Phalaenopsis, and Tetragonia expansa Vanilla

Phalaenopsis Vanilla

Mechanical Mechanical; green peach aphid (Myzus persicae)

Taiwan Cook Islands, Fiji, French Polynesia

Phalaenopsis chlorotic spot virus Potyvirus Vanilla mosaic virus (VanMV), Potyvirus Calanthe mild mosaic virus (CalMMV), Potyvirus Cucumber mosaic virus Cucumovirus

Orchid hosts Cymbidium, Maxillaria, Renanthera, Phalaenopsis

Cymbidium, Dendrobium, and Phalaenopsis and many other ornamentals Phalaenopsis

China, Taiwan, USA Taiwan

Impatiens necrotic spot virus (INSV), Tospovirus Capsicum chlorosis virus (CaCV), Tospovirus

Transmission False spider mite (Brevipalpus californicus)

A number of thrips species including the Western flower thrips (Frankliniella occidentalis) Mechanical

Geographic occurrence Australia, Brazil, Costa Rica, Denmark, Germany, Japan, Korea, USA

Virus, genus Orchid fleck virus (OFV), Dichorhavirus

Table 1 (continued)

Chlorotic spots Chlorotic and necrotic flecks, mosaic and deformation in leaves; dieback of shoots Mild leaf mosaic leaf and flower breaking Severe stunt, conspicuous stem and leaf deformation

Symptoms Yellow flecks and yellow or necrotic ringspot lesions on leaves (Fig. 9e). In Dendrobium, Miltonia, Odontoglossum, Oncidium, and Paphiopedilum, the chlorotic areas often with necrotic centers or rings Large chlorotic or necrotic ringspots on leaves; leaf yellowing and distortion (Fig. 9f) Chlorotic spots with concentric necrosis

Gara et al. 1998 Farreyrol et al. 2001; Farreyrol et al. 2010

Bakardjieva et al. 1998; Baker et al. 2007b; Koike and Mayhew 2001; Zhang et al. 2010; Zheng et al. 2008a Zheng et al. 2008a Zheng et al. 2008b Davis and Ruabete 2010

References Brunt et al. 1997; Dietzgen et al. 2014; Freitas-Astúa et al. 2002; Kondo et al. 2003; Kitajima et al. 2001; Kubo et al. 2009

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Fig. 9 Symptoms of orchid viruses. CymMV: small, discolored Dendrobium flower (a), twisted shoot with yellow streaks (b); ORSV: distorted Dendrobium flower with color-breaking (c), mosaic and yellow patches in Dendrobium (d); OFV: yellow concentric rings in Phalaenopsis leaf (e); INSV: chlorotic patches and necrotic ringspots and distortion in Oncidium leaves (a-e, P. Smitamana # 2017. All Rights Reserved.; f, S. Koike # 2017. All Rights Reserved.)

ornamentals which can serve as reservoirs of the virus. OFV is spread by the false spider mite, Brevipalpus californicus. Management. Once infected by a virus, unlike with other pathogens, nothing short of complicated and exacting tissue culture procedures can rid plants of viruses, and

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even these extreme measures are not always successful. Therefore, management of orchid viruses must be based on prevention of infection by using virus-free propagative material and good sanitation practices. Isolate suspect plants and new introductions and monitor for symptoms for 6 weeks or until proven by a diagnostic test (ELISA, PCR, etc.) to be virus-free. Promptly discard or destroy by burning all confirmed diseased plants. Use only sterile cutting tools; soak the tools in a 10% bleach solution or saturated trisodium phosphate solution for 10 min; or use disposable razor blades when dividing plants and cutting flowers. Wear latex gloves when handling plants and discard those gloves when finished. Always change to new gloves when handling a new plant. Disinfest containers by first washing with soap to remove residual organic matter, then soak in a 20% bleach solution for 1 h, after that soak them for 1 h in a quaternary ammonium salt solution as per label instructions. Exclude insect-vectored viruses by using fine mesh screening, pesticides, and/or biological controls. Refer to chapter “▶ Chap. 4, “Insect Management for Disease Control in Florists’ Crops” for additional information.

5

Nematode Diseases

5.1

Foliar Nematodes (Aphelenchoides spp.)

Geographic occurrence and impact. A number of Aphelenchoides spp. including A. besseyi, A. fragariae, and A. ritzemabosi have been reported to attack orchids in Singapore, Thailand, and the USA (Florida) (Esser et al. 1988; Kawate and Sewake 2014; Latha et al. 1999). Because warm, wet greenhouse environments are conducive to foliar nematode infection, the occurrence of these nematodes in orchids is undoubtedly more widespread. This disease affects many orchids including Cattleya, Cymbidium, Dendrobium, Oncidium, Paphiopedilum, and Vanda. Losses from uncontrolled foliar nematodes have been estimated at up to 90–95% in potted orchid production in Hawaii (Kawate and Sewake 2014). Symptoms/signs. Foliar nematodes infect immature buds and prevent flower formation and development on Vanda; diseased buds abscise or adhere and become blackened. Symptoms on Dendrobium leaves include yellow-green blotches which become brown as the lesions age. On Oncidium, browning of foliar buds and dark streaks in pseudobulbs can occur (Uchida and Sipes 1998). Biology and epidemiology. A film of water enables foliar nematodes to move up stems and over leaf surfaces to seek new infection sites made by wounds or natural openings such as stomates. Infection and symptom expression are, therefore, enhanced by warm, wet conditions. The Aphelenchoides spp. that attack orchids have a very broad host range that includes many ornamentals and other cultivated crops. Adults and 4th stage juveniles feed on mesophyll and epidermal tissue within leaves (endoparasitism). These tissues collapse and turn brown. Females lay eggs within green leaf tissue. Foliar nematodes also feed externally on stems, buds, and

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flowers (ectoparasitism). Adults and 4th stage juveniles can survive in association with living and desiccated plant tissues for long periods. Foliar nematodes can be disseminated by splashing water, infected plant propagative material, and debris (Kohl 2011). Management. Prevention of nematode introduction and good sanitary practices are the keys to managing foliar nematodes. Isolate suspicious plants and new introductions to monitor for symptom expression. Discard plant debris and infected plants. All surfaces coming in contact with plants (pots, stakes, tools, etc.) should first be cleaned with soap and water and surface-disinfested with a 10% solution of household bleach for 10 min after each use (Uchida and Sipes 1998). Growers should avoid wetting plant surfaces as much as possible. Increasing air circulation through increased plant spacing will help to dry plants. Only propagate from clean material. Hot water treatment of Vanda cuttings for 15 min at 46  C (115  F) or 5–10 min at 49  C (121  F) eliminated nematodes without injuring the plants (Uchida and Sipes 1998).

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Pereira OL, Cavallazzi JRP, Rollemberg CL, Kasuya MCM (2002) Sphenospora kevorkianii, a rust fungus (uredinales: raveneliaceae) on the orchid Pleurothallis mentigera. Braz J Micro 33: (short communication). http://www.scielo.br/scielo.php?script=sci_arttext&pid=S151783822002000200011. Accessed 14 Feb 2016 Pinaria AG, Liew ECY, Burgess LW (2010) Fusarium species associated with vanilla stem rot in Indonesia. Australas Plant Pathol 39:176–183 Prapagdee B, Akrapikulchart U, Mongkolsuk S (2008) Potential of a soil-borne Streptomyces hygroscopicus for biocontrol of anthracnose disease caused by Colletotrichum gloeosporioides in orchid. J Biol Sci 8(7):1187–1192. http://dspace.li.mahidol.ac.th/handle/123456789/6716 Prapagdee B, Tharasaithong L, Nanthaphot R, Paisitwiroj C (2012) Efficacy of crude extract of antifungal compounds produced from Bacillus subtilis on prevention of anthracnose disease in Dendrobium orchid. Environ Asia 5:32–38 Pulawska J, Mikicinski A, Orlikowski LB (2013) Acidovorax cattleyae causal agent ofbacterial brown spot of Phalaenopsis lueddemanniana in Poland. J Plant Pathol 95:407–410 Scortichini M, D’Ascenzo D, Rossi MP (2005) New record of Acidovorax avenae subsp. cattleyae on orchid in Italy. J Plant Pathol 87:244 Sherpa AR, Hallan V, Vij SP, Pathak P, Garg ID, Zaidi AA (2003) The first report on Cymbidium mosaic virus (CymMV) in orchids from India. New Dis Rep 7:10 Sherpa AR, Bag TK, Hallan V, Zaidi AA (2006) Detection of Odontoglossum ringspot virus in orchids from Sikkim, India. Australas Plant Pathol 35:69–71 Silva M, Pereira OL (2007) First report of Guignardia endophyllicola leaf blight on Cymbidium (Orchidaceae) in Brazil. Aust Plant Dis Notes 2:31–32 Silva M, Pereira OL, Braga IF, Lelis SM (2008) Leaf and pseudobulb diseases on Bifrenaria harrisoniae (Orchidaceae) caused by Phyllosticta capitalensis in Brazil. Australas Plant Dis Notes 3:53–56 Singh MK, Sherpa AR, Hallan V, Zaidi AA (2007) A potyvirus in Cymbidium spp. in northern India. Aust Plant Dis Notes 2:11–13 Stovold GE, Bradley J, Fahy PC (2001) Acidovorax avenae subsp. cattleyae (Pseudomonas cattleyae) causing leafspot and death of Phalaenopsis orchids in New South Wales. Australas Plant Pathol 30:73–74 Su JF, Lee YC, Chen CW, Hsieh TF, Huang JH (2010) Sheath and root rot of Phalaenopsis caused by Fusarium solani. Acta Hortic 878:389–394. doi:10.17660/ActaHortic.2010.878.49 Sumbali G (2005) The fungi. Alpha Science International, 298 p Sutrabutra T (1989) Virus and virus-like of important plants in Thailand. Funny Publishing, Bangkok. 310 p (in Thai) Takahashi Y, Takahashi K, Watanabe K, Kawano T (2004) Bacterial black spot caused by Burkholderia andropogonis on Odontoglossum and intergeneric hybrid orchids. J Gen Plant Pathol 70:284–287 Tanaka S, Nishii H, Ito S, Kameya-Iwaki M, Sommartya P (1997) Detection of Cymbidium mosaic potexvirus and Odontoglossum ringspot tobamovirus from Thai orchids by rapid immunofilter paper assay. Plant Dis 81:167–170 Tao YH, Ho HH, Wu YX, HE YQ (2011) Phytophthora nicotianae causing Dendrobium blight in Yunnan Province, China. Int J Plant Pathol 2:177–186 Tara BP, McMillan RT, Graves WR (2003) Sclerotium rolfsii Southern blight of Brassidium hybrid orchid. Proc Fla State Hort Soc 116:195–196 The Plant List (2013) Version 1.1. Published on the Internet; http://www.theplantlist.org/1.1/ browse/A/Orchidaceae/. Accessed 4 Apr 2016 Thomson AD, Smirk BA (1967) An unusual strain of tobacco mosaic virus from orchids. N Z J Bot 5:197–202. https://doi.org/10.1080/0028825X.1967.10428740. Accessed 4 Mar 2016 Uchida JY (1994) Diseases of orchids in Hawaii. Plant Dis 78:220–224 Uchida J (1995) Bacterial diseases of Dendrobium, Research extension series, vol 158. Institute of Tropical Agriculture and Human Resources. University of Hawaii

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Uchida JY, Aragaki M (1980) Nomenclature, pathogenicity, and conidial germination of Phyllostictina pyriformis. Plant Dis 64:786–788 Uchida JY, Aragaki M (1991a) Phytophthora diseases of orchids in Hawaii, Research extension series, vol 129. College of Tropical Agriculture and Human Resources. University of Hawaii Uchida JY, Aragaki M (1991b) Fungal diseases of Dendrobium flowers, Research extension series, vol 133. College of Tropical Agriculture and Human Resources. University of Hawaii Uchida JY, Sipes BS (1998) Foliar namatodes on orchids in Hawaii. University of Hawaii at Manoa, PD-13, 7 pp. http://www.ctahr.hawaii.edu/oc/freepubs/pdf/PD-13.pdf Wedge D, Elmer H (2008) Fusarium wilt of orchids. Int Commer Orchid Growers Organ (ICOGO) Bull 2:9–10 Wong SM, Chng CG, Lee YH, Tan K, Zettler FW (1994) Incidence of cymbidium mosaicand Odontoglossum ringspot viruses and their significance in orchid cultivation in Singapore. Crop Prot 13:235–239 You Y, Lü FB, Zhong RH, Chen HM, Li HP, Liu JM, Zhang JH (2016) First report of bacterial brown spot in Phalaenopsis spp. caused by Burkholderia gladioli in China. Plant Dis PDIS-0915-0963-PDN. http://apsjournals.apsnet.org/doi/abs/10.1094/PDIS-11-15-1373-PDN. Accessed 25 Mar 2016 Zhang Q, Ding YM, Li M (2010) First report of impatiens necrotic spot virus infecting Phalaenopsis and Dendrobium orchids in Yunnan Province, China. Plant Dis 94:915–915 Zheng YX, Chen CC, Yang C, Yeh SD, Jan FJ (2008a) Identification and characterization of a tospovirus causing chlorotic ringspots on Phalaenopsis orchids. Eur J Plant Pathol 120:199–209 Zheng YX, Chen CC, Chen YK, Jan FJ (2008b) Identification and characterization of a potyvirus causing chlorotic spots on Phalaenopsis orchids. Eur J Plant Pathol 121:87–95 Zheng YX, Shen BN, Chen CC, Jan FJ (2010) Odontoglossum ringspot virus causing flower crinkle in Phalaenopsis hybrids. Eur J Plant Pathol 128:1–5 Zhou G, Chen X, Li M, Zhou J, Tang T, Feng S, Guo L, Zhang W (2004) Identification and detection of two major viruses infecting orchids by molecular technique. Virol Sin 19:149–152 Zhou JN, Lin BR, Shen HF, Pu XM, Chen ZN, Feng JJ (2012) First report of a soft rot of Phalaenopsis aphrodita caused by Dickeya dieffenbachiae in China. Plant Dis 96:760–760

Diseases of Peonies

22

Andrea R. Garfinkel and Gary A. Chastagner

Abstract

Herbaceous peonies (Paeonia lactiflora Pall.) are grown throughout temperate regions of the world for their use as high-value cut flowers. Despite their popularity, little information is available on diseases and disease management of this flower compared with other ornamental geophytes, such as tulips or lilies. From the literature, it is clear that the fungal diseases Botrytis gray mold and leaf blotch are among the most economically important diseases of peony; however, many other fungi, viruses, bacteria, and nematodes affect peonies worldwide. Management recommendations used for pathogens of peonies rely mainly on general disease management strategies and information on pathogen biology and epidemiology from other pathosystems.

Keywords

Botrytis • Cut flowers • Fungi • Paeonia • Plant pathogen

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Botrytis Gray Mold (Botrytis cinerea Pers. Fr. and Botrytis paeoniae Oud.) . . . . . . 2.2 Leaf Blotch (Graphiopsis chlorocephala [Fresen.] Trail) . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Alternaria Leaf Spot (Alternaria spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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A.R. Garfinkel (*) • G.A. Chastagner Department of Plant Pathology, Puyallup Research and Extension Center, Washington State University, Puyallup, WA, USA e-mail: andrea.garfi[email protected]; [email protected] # Springer International Publishing AG 2018 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florists’ Crops Diseases, Handbook of Plant Disease Management, https://doi.org/10.1007/978-3-319-39670-5_46

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2.4 2.5

Powdery Mildew (Erysiphe spp. and Podosphaera spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . Phytophthora Blight and Phytophthora Root Rot (Phytophthora cactorum [Lebert & Cohn] J. Schröt) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 Anthracnose (Gloeosporium spp. and Colletotrichum spp.) . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Crown Rot (Sclerotium rolfsii Sacc. and Sclerotium delphinii Welch) . . . . . . . . . . . . . 2.8 White Stem Rot (Sclerotinia sclerotiorum [Lib.] deBary) . . . . . . . . . . . . . . . . . . . . . . . . . . 2.9 Fungal Root Rots (Multiple Species) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.10 Peony Red Spot (Mycocentrospora acerina [Hartig] Deighton) . . . . . . . . . . . . . . . . . . . 2.11 Leaf Rust (Cronartium flaccidum [Alb. & Schw] Wint.) . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Bacterial and Phytoplasma Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Bacterial Diseases (Multiple spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Phytoplasma Diseases (Candidatus Phytoplasma spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Viral Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Tobacco rattle virus (TRV; Family Virgaviridae, Genus Tobravirus) . . . . . . . . . . . . . . . 4.2 Lemoine Disease (Suspected Virus) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Nematode Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 Root-Knot Nematode (Meloidogyne spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2 Foliar Nematode (Aphelenchoides spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Introduction

The vast majority of peonies produced for cut flowers are hybrids crossed with the herbaceous peony species, Paeonia lactiflora Pall. This species, native to China, comprises some of the oldest commercial cut flowers, rising first to popularity in Europe in the late 1700s (Rogers 1995). Hybridization of the herbaceous peony took place shortly thereafter and many of the still-popular cut flower types are relics of these nineteenth century breeders (Rogers 1995). The propagation of these cultivars is done exclusively through vegetative division of the root crown and fleshy storage roots. Peony is undergoing a resurgence in popularity, and cut flowers are available nearly year-round at the Dutch flower auctions as nontraditional flower-producing regions, such as the state of Alaska in the United States, are producing peonies at a rapidly increasing rate to fill gaps in availability (Auer and Greenburg 2009; Holloway and Buchholz 2013). Despite the rise in production and consumption of cut peonies, the information about diseases and disease management is, at best, disjointed. This chapter will seek to consolidate the somewhat limited information available about peony diseases and their management. Where specific epidemiology and disease control methods are not known in peony, an attempt has been made to supplement with general information about the pathogen group or information about other pathosystems. Diseases of other Paeonia species not typically used for cut flowers, including tree peonies (Paeonia suffruticosa), will not be specifically discussed; however, a few mentions of diseases on tree peonies and other Paeonia species have been included when appropriate.

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2

Fungal and Fungus-Like Diseases

2.1

Botrytis Gray Mold (Botrytis cinerea Pers. Fr. and Botrytis paeoniae Oud.)

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Geographic occurrence and impact. Botrytis gray mold (also “Botrytis blight”) affects peonies in nearly every region in which they are cultivated and can be considered, economically speaking, the most important peony pathogen (Whetzel 1939). Botrytis can infect every part of the peony plant including the foliage, stems, flowers, and the tuberous roots and root crown (Wallace et al. 1962; Whetzel 1939). Due to its ability to cause disease on floral tissues under storage conditions (Droby and Lichter 2007), Botrytis is both a pre- and postharvest concern for cut peonies. Symptoms/signs. Early in the season, Botrytis blight can be seen on emerging shoots as a dark brown soft rot that causes toppling of stems (Whetzel 1939). Later in the season, lesions on expanded stems and foliage are dry and tough and exhibit dark and light brown zonation (Whetzel 1939). Lesions originating on the foliage can move down the petiole into the stem. Lesions at the base of the stem can girdle the stem and cause the upper foliage to become red and wilted (Whetzel 1939). Botrytis can colonize undeveloped or “blasted” buds causing them to turn black. Fully developed buds that become infected turn brown, and the petals rot inside causing failure of the flower to open (Whetzel 1939). Botrytis infection of the flower petals develops as brown spots that quickly coalesce, blighting the petals and causes them to drop off (Whetzel 1939). Infections of the flower or flower bud can move down the stem which will sometimes exhibit the zonation described above. Under moist conditions, conidia develop on the surface of lesions giving the tissue a gray fuzzy texture lending inspiration for the name “gray mold” (Fig. 1). Biology and epidemiology. The source of a Botrytis gray mold infection on peony depends on which species of the fungus is the causal agent of disease. Botrytis paeoniae is thought to be a host-specific pathogen (causing disease only on peony); therefore, the inoculum from this pathogen will be from previous or nearby crops of peony (Holz et al. 2007; Whetzel 1939) or potentially infected planting stock. B. cinerea inoculum, on the other hand, could come from any of its over 250 hosts. Primary inoculum for both organisms is considered to be conidia. The first conidia of the season arise from sclerotia which overwinter on the surface of decayed peony foliage. Subsequent crops of conidia (secondary inoculum) are produced on infected plant material throughout the growing season. Conidia enter the host through natural openings, directly, or through wounds (Jarvis 1980). Colonized peony petals and other plant tissues can also serve as an important inoculum source, whereby mycelium directly infects healthy tissue from saprophytically colonized tissue (Daughtrey et al. 2000; Holloway personal communication 2015; Holz et al. 2007). Conidia are mostly dispersed by wind (Holz et al. 2007).

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Fig. 1 Botrytis gray mold on peony foliage showing alternating tan and dark brown zones (left), a peony basal stem lesion showing zonation (center), and an infected flower bud with sporulation (right) (Andrea R. Garfinkel # 2017. All Rights Reserved.)

Disease development is typically considered to be favored by cool, wet weather conditions in which there is free moisture on the plant surface (Jarvis 1980). While B. paeoniae and B. cinerea are the most commonly cited Botrytis species to infect peony, preliminary investigations have identified several other Botrytis species infecting peony including the fenhexamid-resistant B. pseudocinerea (Garfinkel and Chastagner unpublished data; Muñoz et al. 2015). Management • Cultural practices – Reducing leaf wetness is one of the most important mechanisms for limiting Botrytis gray mold development (Daughtrey and Benson 2005). Therefore, wide plant spacing to avoid overcrowding of plants and allow for maximum air movement and sun exposure can help limit disease development (Whetzel 1939). Avoiding overhead irrigation can also help reduce disease incidence. Keeping flowers dry and reducing relative humidity and fluctuations in temperatures in storage areas will help decrease postharvest disease of peony (Whetzel 1939). • Sanitation – Removal of all infected plant parts during the season, especially spent flowers (Pfleger et al. 2007) and other dead plant tissues (Daughtrey and Benson 2005), is an important way to reduce sources of secondary inoculum (Whetzel 1939). All foliage should be cut, removed from the field, and disposed of at the end of the season to eliminate plant material on which the fungus can survive and infect the next crop (Whetzel 1939). Planting of clean propagative material is also an important way to minimize in-field inoculum. Although there are no studies available on the effectiveness of hot-water treatment for controlling Botrytis on peony roots, this practice may be useful as it has proven effective in other geophyte crops.

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• Fungicides – Fungicides are among the most important tools in an integrated Botrytis management program for many ornamentals (Daughtrey and Benson 2005). However, their efficacy is hampered by widespread resistance to fungicides, such as the dicarboximides and benzimidazoles (Leroux 2007). The prevalence of fungicide resistance in Botrytis populations on peonies is unknown. Fungicides typically effective against Botrytis include chlorothalonil, azoxystrobin, iprodione, mancozeb, fenhexamid (except B. pseudocinerea, see above), and copper-based products. Wallace et al. (1962) found that soil drenches containing dicloran were effective at controlling basal stem rot caused by infected roots. • Resistance – Winters (1930) published a list of cultivars and their relative susceptibility to Botrytis paeoniae. The susceptibilities of some peony cultivars are described in Table 1. Itoh hybrids, crosses between the herbaceous P. lactiflora and the tree peony P. suffruticosa, which are gaining popularity in the cut flower market, appear to be more resistant to gray mold than other P. lactiflora types (Beckerman and Lerner 2009).

2.2

Leaf Blotch (Graphiopsis chlorocephala [Fresen.] Trail)

[Shubert et al. (2007) and Braun et al. (2008) conducted investigations into the genetics of herbarium samples of Cladosporium paeoniae Pass., syn. Cladosporium chlorocephalum (McKemy and Morgan-Jones 1991), casual agents of peony leaf blotch, and have suggested that this fungus be renamed Graphiopsis chlorocephala.] Geographic occurrence and impact. Leaf blotch appears to affect many different species of peonies wherever they are grown (McKemey and Morgan-Jones 1991) and ranks perhaps only second in importance to Botrytis gray mold. Leaf blotch is not, as the name would suggest, limited to the leaves of the peony, but can impact all aboveground parts of the plant severely hampering marketability of stems, flowers, and foliage. Symptoms/signs. Leaf blotch goes by many names (also Cladosporium leaf blotch, Cladosporium spot, Cladosporium red spot, red spot, or measles) that describe the various symptoms infected plants may display. Symptoms can range from purple-red, irregularly shaped flecks or streaks with diffuse margins on stems, foliage, flowers buds, and flower petals to large, expanding purple lesions on stems or foliage with dark centers (Fig. 2). The leaf spots as viewed from the underside of the leaves appear pale to chestnut brown. Infected stems and leaves maintain their succulence (Meuli 1937). The development of different symptoms appears to be correlated with leaf phenology; young, succulent tissue limits the infection to flecking, while large, expanding lesions develop on more mature tissues (Weiss 1940). Biology and epidemiology. The most important source of initial inoculum, conidia, of G. chlorocephala arises from plant debris from the previous year in

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Table 1 Susceptibility of peony cultivars to Botrytis paeoniae as determined by Winters (1930) Susceptibility Resistant

Moderately resistant

Susceptible

Very susceptible

Cultivars Akalu, Argus, Arthemise, Attraction, Avalanche, Balliol, Baroness Schroeder, Baron James Rothschild, Black Prince, Cavalleria Rusticana, Chalice, Christine Gowdy, Christine Ritcher, Comte de Nanteuil, Dorothy, Dorothy Echling, Dorothy E. Kibby, Ella Wheeler Wilcox, Eucharis, Eureka, Fragrans, Fulgida, General Bertrand, General Cavaignac, Gloire de Chenonceaux, Glorious, Flory of Somerset, Gretchen, Griff Thomas, Hermes, Hogioku, Itenshikai, King of England, Kumagaye, Lady Bellew, Lady Mayoress, L’étincelante, Lord Salisbury, Luetta Pfeiffer, Madame Lemonnier, Madame Schmidt, Maud L. Richardson, Meissonier, Monsieur Boucharlataine, Mrs. Gwyn-Lewis, Mr. L. van Leeuwen, Old Silvertip, Petite Renée, Plutarch, Princess Ellen, Purpurea Superba, Queen Wilhelmina, Red Bird, Ruigegno, Sarah Bernhardt, Speedwell, Sweet Home, Yeso Admiral Dewey, Agnes Mary Kelway, Alb^atre, Albert Crousse, Alfred de Musset, American Beauty, Archie Brand, Asa Gray, Augustin d’Hour, Aunt Ellen, Béranger, Bullock, Carnea Elegans, Carnot, Charles Binder, Charles Verdier, Charlotte Cushman, Clarisse, Comet, Conqueror, Couronne d’Or, Daubenton, Daybreak, Dorchester, Dorothy Kelway, Duchess of Portland, Duke of Devonshire, Edwin Forrest, Emile Lemoine, Enchantment, Eternal City, Etta, Faust, Favorite, Festiva Maxima, Flambeau, Frances Shaylor, Graziella, Grizzel Muir, Gypsy, Henry Avery, Hon. Mrs. Portman, Innocence, John Fraser, June Day, Jupiter (Calot), Kelway’s Queen, La Coquette, Lady Alexandra Duff, Lady Somerset, La Fraicheur, Lake of Silver, La Perle, La Sublime, La Tulipe, La Vestale, L’étincelante (Dessert), Lord Lytton, Louis van Houtte, Mabel L. Franklin, Madame Coste, Madame de Guerle, Madame de Vatry, Mademoiselle Gaillant, Mafeking, Mary L. Hollis, Masterpiece, Mathilde de Roseneck, Mazie Terry, Monsieur Paillet, Monsieur Pasteur, Muchelny, Norfolk, Octavie Demay, Pallas, Phoebe Cary, Pierre Duchartre, Pink Enchantress, Princess Maud, Queen of Beauty, Rauenthal, Rhoda, Rubicunda, Ruth Brand, Simonne Chevalier, Sir Robert Gresly, Snowflake, Sosthenes, Sully Prudhomme, Torquemada, Triumphata, Trojan, Venus, Victoria, Ville de Nancy, Waterloo, Welcome Guest Adam Bede, Agnes Barr, Amalthea, Armand Rousseau, Bertha, Camille Calot, Canariensis, Carlotta Grisi, Carnea Triumphans, Caul, Chrysanthemiflora, Comte de Cussy, Comte de Paris, Countess of Clancarty, Daniel d’Albert, Delachei, Duc de Cazes, Duc de Wellington, Eastern Beauty, Edmond Lebon, Etienne Mechin, Frances Shaylor, General Grant, Grandiflora, Jules Calot, Lady Beresford, Lutetiana, Madame de Verneville, Madame Emile Galle, Madame Hutin, Magnifica, Marie Lemoine, Marquise d’Ivry, Mathilde Méchin, Meadowvale, Monsieur Chevreul, Mrs. Lowe, Myrtle, Pottsi, Princess Beatrice, Pulcherrima, Queen’s Perfection, Roem de Boskoop, Sappho, Sea Foam, Snowball (Hollis), Souvenir de Gaspard Calot, Strasbourg, Sunrise, Thomas S. Ware, Torch, Triomphe du Nord, Turana, Vicomtesse de Belleval, Virginie, Virgo Maria, Viscountess Folkestone, Whitleyi Antione Porteau, Armandine Méchin, Assmanshausen, Belle of France, Charles Toche, General Bedeau, Grandiflora Lutescens, Irma, Lutea Plenissima, Nivea Plenissima, Paradise, Territorial, Victoire Modeste

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Fig. 2 Flecking on peony leaves and petiole (left), a severe infection on an unopened peony bud (center), and large leaf lesions on peony foliage (right) caused by Graphiopsis chlorocephala (Andrea R. Garfinkel # 2017. All Rights Reserved.)

which the fungus has overwintered as mycelium (McKemy and Morgan-Jones 1991; Meuli 1937; Weiss 1940). Ying and Guihua (1984) found that optimal growth and germination of G. chlorocephala conidia occurred at 20–24  C and Weiss (1940) found that infection of peony could occur from 10  C to 27.5  C, but with significant increase in latent period at lower temperatures. Infection can occur on both wounded and unwounded tissues on tree peonies (Ying and Guihua 1984). Free moisture on the leaf is necessary for conidia germination and infection. Conidia can be produced on infected peony foliage throughout the year; however, its occurrence appears to be rare (Meuli 1937). Management • Sanitation – Removal of infected plant material from the previous year’s foliage is effective at reducing leaf blotch severity in peony (Meuli 1937; Weiss 1940). Burning the plant debris in the field has also been shown to be effective (Meuli 1937). • Fungicides – While testing products for Botrytis management on peony, researchers found that fungicides tebuconazole, fluoxastrobin, chlorothalonil, triticonazole, and copper hydroxide and fungicide mixes boscalid + pyraclostrobin and cyprodinil + fludioxonil reduced the severity of leaf blotch (Palmer and Vea 2014). • Resistance – Itoh hybrids (see “Resistance” in “Botrytis Gray Mold” above) are reported to be more resistant to leaf blotch than P. lactiflora species peonies (Beckerman and Lerner 2009). Meuli (1937) reported that the cultivars Oshkosh White, Felix Crousse, and Livingstone are relatively susceptible to the disease, whereas cultivars Augustin d’Hour, Mathilde de Roseneck, Louis van Houtte, Edulis Superba, Jules Calot, Gigantea, and Humei carnea are more resistant.

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Alternaria Leaf Spot (Alternaria spp.)

Geographic occurrence and impact. Alternaria spp. have been reported on peony in the United States (Anonymous 1960), Canada (Coulson 1920), China (Zhang 2003), Korea (Cho and Shin 2004), and Bulgaria (Bobev 2009). Most reports of Alternaria spp. on peony do not identify the organism to species; however, it appears that at least two species are associated with the herbaceous peony, A. alternata (Zhang 2003, China) and A. infectoria (Anderson et al. 2009, the Netherlands). Three additional species are reported on the tree peony in China, A. suffruticosae, A. suffruticosicola, and A. tenuissima (Zhang et al. 2008). Symptoms/signs. The symptoms associated with this fungus are not well documented. At least two references question if the isolation of Alternaria from peony is due to a secondary infection by the fungus (Anonymous 1960; Coulson 1923) which is characteristic of A. alternata (Rotem 1998). The Texas Plant Disease Handbook (Anonymous n.d.) describes the leaf spots as irregularly shaped ranging in color from purplish-brown to reddish-brown. Later, leaves may yellow and fall off (Texas Plant Disease Handbook). Coulson (1923) suspects Alternaria as the causal agent of yellow leaf spots that were irregularly shaped with a clearly defined, dark border. Alternaria spp. have been isolated from peony leaves with light brown, irregularly shaped lesions in the US Pacific Northwest (Fig. 3, Garfinkel and Chastagner, unpublished). Leaf spots caused by the three Alternaria spp. infecting tree peony are described as circular or irregular, brown to black in color (Zhang et al. 2008). Biology and epidemiology. No refereed information has been located on the biology and epidemiology of Alternaria spp. on peony. Generally, Alternaria spp. persist in plant debris or perennial plant structures as mycelium or spores, which are heavily pigmented (Rotem 1998). Conidia are primarily aerially dispersed, but have also shown to be dispersed with water and require free moisture on the surface of a

Fig. 3 Leaf spots on peony associated with an Alternaria spp. (Andrea R. Garfinkel # 2017. All Rights Reserved.)

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plant to germinate (Rotem 1998). A. alternata is typically regarded as a weak pathogen, capable of infecting only weakened plants or plant tissues, and only causes infection on healthy plants in some pathosystems (Rotem 1998). Management • Cultural – Cultural management to prevent wounding of plant tissues and long periods of leaf wetness including wider plant spacing and avoiding overhead irrigation may help prevent Alternaria leaf spot (Rotem 1998). Removing, destroying, or burying plant debris will help to prevent the fungus from overwintering (Rotem 1998). • Fungicides – Information on the effectiveness of fungicides in controlling Alternaria leaf spot is lacking, although there are many products registered for use for this pathogen in peonies in the United States including iprodione, thiophanate-methyl, propiconazole, trifloxystrobin, chlorothalonil, fluoxastrobin, azoxystrobin, pyraclostrobin, fludioxonil, and mancozeb.

2.4

Powdery Mildew (Erysiphe spp. and Podosphaera spp.)

Geographic occurrence and impact. Powdery mildew has been reported on peonies in Iran (Khodaparast and Abbasi 2009), the former Soviet Union (Amano 1986), Korea (Cho and Shin 2004; Shin 2000), China (Braun and Takamatsu 2000; Tai 1979; Zheng and Yu 1987), Japan (Kobayashi 2007; Takamatsu et al. 2006), Hungary (Braun 1995), and the United States (Anonymous 1960; French 1989). Symptoms/signs. Powdery mildew is characterized by conspicuous white circles or a mat of mycelium on the surface of aboveground parts of the plant (Fig. 4). Later in the season, small, round, black, fruiting structures, called chasmothecia, can be seen on top of or buried within the white mycelium (Fig. 4). Infected peony tissues will sometimes turn purple underneath the mycelium in response to the presence of the pathogen. Biology and epidemiology. Powdery mildews are considered highly specialized, obligate biotrophs which means that they are mostly host-specific and require living host tissue for growth and development. Powdery mildews can survive as mycelium or as chasmothecia, which contain sexually produced ascospores. Mycelium, conidia, or ascospores can serve as the source of initial inoculum in the spring. Asexual conidia are produced on the mycelium throughout the year and serve as a source of secondary infection. Powdery mildew spores, unlike many fungi, do not require free moisture to germinate. Management • Cultural – Removal of infected plant debris during and at the end of the year can help prevent inoculum buildup. Increasing plant spacing to provide as much sun

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Fig. 4 A severe powdery mildew infection on a peony seed pod and foliage (left) and small black (mature) and orange (immature) chasmothecia among the mycelial mat on the lower leaf surface (right) (Andrea R. Garfinkel # 2017. All Rights Reserved.)

exposure to foliage as possible may also help reduce infection (Glawe and Grove 2015). • Fungicides – Fungicides typically effective against powdery mildews include those that contain sulfur or petroleum-based oils, sterol-biosynthesis inhibitors (DMIs), or quinone outside inhibitors (QoIs) (Glawe and Grove 2015).

2.5

Phytophthora Blight and Phytophthora Root Rot (Phytophthora cactorum [Lebert & Cohn] J. Schröt)

Geographic occurrence and impact. Phytophthora blight on peonies has been officially reported in the United States (Anonymous 1960; Thurston and Orton 1921), Canada (Conners and Savile 1948), and Japan (Kobayashi 2007; Villa et al. 2006). Symptoms/signs. Symptoms of Phytophthora blight can be easily confused with Botrytis gray mold as aboveground symptoms involve a dieback of stems and foliage. However, dieback caused by Phytophthora is sunken and black as compared to a Botrytis infection which is typically brown in color. Furthermore, incubation of the tissue in moist conditions does not result in the appearance of gray conidiophores; a sparse white mycelial mat can sometimes be observed. Phytophthora cactorum can infect the enlarged roots of the peony and will form chocolate brown lesions on the roots with defined margins (Fig. 5). Severely infected plants may fail to send up shoots in the spring.

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Fig. 5 Lesion on a fleshy peony root caused by Phytophthora cactorum (Andrea R. Garfinkel # 2017. All Rights Reserved.)

Biology and epidemiology. Phytophthora cactorum is not a true fungus, but a fungal-like oomycete or “water mold.” P. cactorum has over 200 plant host species and grows optimally at 25–30  C (Erwin and Ribeiro 1996; Kim et al. 2003). Hyphae differentiate to produce sporangia that can either germinate directly or from which swimming zoospores emerge. These zoospores have been shown to be infective on the leaves of tree peonies (Kim et al. 2003). Some isolates can produce chlamydospores which can persist in soil and germinate in favorable conditions (Erwin and Ribeiro 1996). The organism is also reported to overwinter as mycelium and oospores (Erwin and Ribeiro 1996). Management • Sanitation – Plants with severe Phytophthora blight of the crown should be removed from the field along with surrounding plants and soil which may have become infected or infested by the pathogen. • Fungicides – Fungicides can be applied to foliage or as a drench. Little information is available on the fungicide control of Phytophthora on peonies. General recommendations for fungicides to control P. cactorum include the use of metalaxyl or fosetyl-Al (Erwin and Ribeiro 1996) if registered for use on peony. Chemistries effective in managing Phytophthora spp. in other pathosystems include aluminum tris (O-ethyl phosphonate), captan, cyazofamid, dimethomorph, famoxadone + cymoxanil, fenamidone, fluopicolide, fluoxastrobin + myclobutanil, mandipropamid, mefenoxam, metalaxyl, pyraclostrobin, and fungicides containing phosphorous acids (Palmer and Vea 2010). Fungicide resistance can be a problem when using fungicides to manage Phytophthora diseases, so it is important to rotate

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materials with different modes of action (see ▶ Chap. 7, “Fungicides and Biocontrols for Management of Florists’ Crops Diseases”). • Cultural – Avoid the presence of standing water in the field which is favorable to pathogen development and movement of zoospores. Overhead irrigation should be avoided as splash can disperse the pathogen within a field.

2.6

Anthracnose (Gloeosporium spp. and Colletotrichum spp.)

Geographic occurrence and impact. Anthracnose has been reported on peonies in the United States (Anonymous 1960; Weiss 1940), Korea (Cho and Shin 2004; Korean Society of Plant Pathology 1998), Japan (Kobayashi 2007), and New Zealand (Pennycook 1989). Symptoms/signs. The anthracnose fungi can cause lesions and cankers on all aboveground parts of peony. Cankers on the stems of peony are elongate and gray in color and often cause severe twisting or curling of the stem (Fig. 6). Pink conidomata can sometimes be observed in the cankers under favorable conditions. Foliar leaf spots and infections of flowers and flower buds are similarly gray in color and can cause irregular growth of plant tissues (Fig. 6) or interfere with flower development. Biology and epidemiology. “Anthracnose” is a general term used to describe a disease that “appears as a black, sunken, leaf, stem, or fruit lesions, caused by fungi that produce their asexual spores in an acervulus” (Agrios 2005). Anthracnose fungi are generally thought to survive in leaf litter; therefore, infested debris can serve as an important source of initial inoculum. Anthracnose fungi on peony appear to

Fig. 6 Cankers (left) (Andrea R. Garfinkel # 2017. All Rights Reserved.) on and twisting of stem below bud (right) (Published with kind permission of J. Mullane # 2017. All Rights Reserved.) on peony caused by Colletotrichum spp.

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sporulate under favorable conditions during the season; thus, secondary inoculum may play a role in the disease cycle. Management. Follow general disease management strategies for fungi including cultural practices to reduce leaf wetness instance and duration (see Management of Botrytis above for specific tactics), and remove and destroy all foliage at the end of the season.

2.7

Crown Rot (Sclerotium rolfsii Sacc. and Sclerotium delphinii Welch)

Geographic occurrence and impact. Crown rot or southern blight has been reported on peonies in North America (Anonymous 1960; Parris 1959) and Japan (Kobayashi 2007). Limited information is available on the impact of this disease on peonies, but Sclerotium rolfsii (teleomorph Athelia rolfsii) and the closely related S. delphinii can cause extensive mortality on a number of ornamental geophytes. Symptoms/signs. Symptoms will appear as small, water-soaked lesions at the base of a stem, quickly expanding to girdle the stem causing wilt and necrosis of the foliage above. As the disease progresses, a white mycelial mat can be observed on the stems and in the soil surface, within which, small orange-tan to brown sclerotia are formed (Fig. 7).

Fig. 7 Immature white to orange colored sclerotia (arrow) of Sclerotium rolfsii developing at the base of a bulbus iris shoot (Andrea R. Garfinkel # 2017. All Rights Reserved.)

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Biology and epidemiology. The crown rot fungi persist in the soil on dead organic matter as mycelia or in the soil as sclerotia that germinate to produce mycelia. The mycelia directly infect crown tissues of the plant host. The production of basidiospores on infected tissues and on mycelium has been observed; however, it is thought that their production is rare (Mullen 2001). Thus, it is not common to practice management strategies based on secondary infection by basidiospores. Instead, spread is thought to occur by mycelial growth to adjacent plants and movement of infected plant material and infested soil. S. rolfsii and S. delphinii are favored by warm, humid conditions, with S. rolfsii specifically limited by low temperatures (Xu et al. 2008). Management • Fungicides – Preventing introduction of the crown rot pathogens by ensuring clean planting stock is an important element of disease management. Chemical treatment of planting stock can be an option to ensure clean planting materials; however, information on the chemical control of this disease on peonies is not available. Soil applications of azoxystrobin, flutolanil, PCNB, and/or preplant dips in PCNB have been used to control crown rot (Chastagner and Riley 2002; Chastagner et al. 1990). Soaking bulbs, such as iris, in hot water plus formalin is also recommended. • Sanitation – Removal of infected plant material and surrounding soil is recommended to limit spread of the pathogen. • Soil disinfestation – Preplant soil fumigation with metam-sodium has been effective in limiting disease development on other geophytes. Soil solarization or sterilization can be effective as preplant treatments if high temperatures and humidity can be maintained for a sufficient duration to kill sclerotia (Mullen 2001).

2.8

White Stem Rot (Sclerotinia sclerotiorum [Lib.] deBary)

Geographic occurrence and impact. White stem rot (also called “white mold” or “Sclerotinia stem rot”) has been reported on peonies in the United States and Canada (Anonymous 1960; Ginns 1986). It has also been reported on tree peonies in Korea (Cho and Shin 2004). Symptoms/signs. Plants infected with S. sclerotiorum initially display aboveground symptoms of wilt and necrosis. The pathogen can later be seen growing at the plant crown as a fluffy white mycelium. Large black sclerotia (white when immature) can be seen forming on the surface or inside the plant tissue. S. sclerotiorum is also able to cause lesions on foliage and flowers that turn brown and can become covered in a white, fluffy mycelial growth under moist conditions. Biology and epidemiology. S. sclerotiorium overwinters as sclerotia formed on or inside plant tissue. Sclerotia germinate either to produce mycelium or apothecia which contain sexual ascospores that become airborne and can infect senescent plant

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tissues. S. sclerotiorum does not produce conidia; thus, there is no repeating cycle throughout the year. However, healthy plants or plant parts can be infected if brought in contact with colonized host tissue or by mycelial spread on the soil surface. Management • Fungicides – Little information is available relating to fungicides that are effective in controlling white mold on peonies. On other crops, fungicides such as thiophanate-methyl, iprodione, boscalid, fludioxonil + cyprodinil, and fluazinam are recommended and can reduce the number of sclerotia in soil. • Cultural – Sanitation to remove infected plant material and senescent plant tissues, especially flower petals, is crucial for preventing pathogen spread. Infected plants should be removed along with surrounding soil to remove any overwintering sclerotia. General fungal disease management strategies to decrease periods of leaf wetness (see Management of Botrytis above for specific tactics) can also be effective at reducing ascospore germination.

2.9

Fungal Root Rots (Multiple Species)

Geographic occurrence and impact. The fungal species reported to cause root rot of peony include Armillaria mellea (United States, Anonymous 1960), Rhizoctonia spp. (United States, Anonymous 1960; Canada, Ginns 1986; Korea, Cho and Shin 2004; Bulgaria, Bobev 2009), Verticillium spp. (United States, Anonymous 1960), Thielaviopsis basicola (United States, Anonymous 1960; French 1989), Fusarium spp. (United States, Anonymous 1960), and Rosellinia necatrix (Japan, Kobayashi 2007; United States, French 1989; Greece, Pantidou 1973). Symptoms/signs. The specific information about these root-rotting pathogens on peony is limited. Aboveground symptoms of root rots typically first display as general wilting, chlorosis, or necrosis as a result of damage to root tissues. Many soilborne fungal pathogens will display characteristic disease patterns in the field with circular foci. Other symptoms are disease-specific. Roots infected with Armillaria often will have thick, black, specialized fungal structures called rhizomorphs, which resemble fine root hairs. Root lesions caused by Rhizoctonia are typically brown to red in color. Verticillium is diagnosed by discoloration or darkening of the vascular tissue in roots and stems. Roots infected by Thielaviopsis often show dark brown to black lesions due to the presence of pigmented chlamydospores that are visible with a microscope in the host tissue. In tree peony, Fusarium root rot causes reddish brown to dark brown lesions on the root (Guo et al. 2012). Infection by Rosellinia is characterized by a very dense, light colored mycelial mat growing up around the base of the stems on the soil surface. Biology and epidemiology. While all are soilborne plant pathogens, the biologies of the root-rotting fungi are unique. Armillaria largely relies on underground spread through mycelial transfer by root-to-root contact and by rhizomorphs. Like

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Armillaria, spores in Rosellinia seem to have little importance in the disease cycle with the majority of spread occurring underground by infected roots coming in contact with healthy roots. Rhizoctonia, Verticillium, and Thielaviopsis form persistent structures called sclerotia, microsclerotia, and chlamydospores, respectively. These structures germinate to produce mycelia which can infect healthy plant tissues. Root-to-root contact between infected and healthy roots can also spread these pathogens. Like most root-infecting pathogens, infected planting stock can play a large role in introducing the pathogen to a new field planting site. Management. Ensuring clean planting material is a best first defense against root-rotting pathogens. For many of these pathogens, rogueing infected plants is recommended, as well as the closest adjacent soil and plant(s), which, although they may not be showing symptoms, could be infected and serve as a source of further spread of the pathogen. Cultural management strategies to alter the pH, temperatures, or saturation of the soils can be successful for some soilborne pathogens. Fungicide drenches or fumigants are available and effective for some, but not all, soilborne pathogens. It is important to always check the pesticide label for proper use.

2.10

Peony Red Spot (Mycocentrospora acerina [Hartig] Deighton)

Geographic occurrence and impact. This disease has only been officially reported in Chile (Gilchrist et al. 2015); however, the geographic distribution is likely much larger as it has also been found on peony in the continental United States and Alaska (Garfinkel and Chastagner unpublished data). As it has only recently been described on peony (Gilchrist et al. 2015), the impact of peony red spot is yet to be determined. The name “red spot,” given to this disease by Gilchrist et al. (2015), is not to be confused with leaf blotch (see earlier in this chapter) which is sometimes referred to as “red spot” in older literature (Weiss 1940). Symptoms/signs. M. acerina can cause disease on all parts of the peony except the flower, although infections of the first few petals of the flower bud have been observed (Gilchrist et al. 2015). On aboveground plant parts, symptoms are apparent as small ( 70%. Management • Cultural practices – Avoid injuring canes. Prune canes with a disinfected, sharp knife or pruner immediately above a node. Avoid leaving long stubs above a node. Frequently clean and disinfect cutting tools (refer to ▶ Chap. 9, “Sanitation for Management of Florists’ Crops Diseases”). Cut and destroy infected canes well below the affected area. Avoid water splashes caused by irrigation, prefer the irrigation drip. Use a winter protecting mulch that does not hold much water such as sand, rock pumice, or coarse bark.

2.12

Petal Spots (Ghost Spot) (Many Fungi Possible Such as Bipolaris, Botrytis, Cercospora, and/or Cladosporium)

Geographic occurrence and impact. Worldwide. Damage makes flowers unmarketable. Symptoms. Numerous small spots develop on petals as buds are opening through flowering. Spots may have colored or darkened margins depending on the cultivar and fungus involved.

Fig. 6 Myrothecium galls can start at pruning wounds (Tatiana Gomez Rodriguez # 2017. All Rights Reserved.)

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Biology and epidemiology. Warm, humid/wet conditions favor disease development especially during rainy weather. Management • Cultural practices – Keep rainfall off blooms. Prune off symptomatic buds as soon as disease is noticed.

2.13

Powdery Mildew (Podosphaera pannosa (Wallr.: Fr.) de Bary 1870)

Geographic occurrence and impact. Commonly found worldwide on roses and other members of the rose family. Symptoms/signs. White, powdery, fungal growth on young leaves, shoots, and buds. When scouting, check on the underside of leaves with good lighting conditions, as this is where it may first start. Distorted growth is common especially if young tissues are infected. If the disease is severe, foliage is stunted (Fig. 7). Biology and epidemiology. The fungus overwinters in infected buds, rose leaves, twigs and branches. Infected buds grow in spring but are stunted and white with fungal conidia. Conidia are released in response to abrupt decreases in relative humidity. Wind blows conidia to healthy foliage where they start new mildew colonies. Germination and growth of conidia happen most readily on nights with high humidity or heavy dew and at temperatures near 21  C (70  F). Too much water, such as flowing water or rain, destroys spores by causing them to burst. The fungus does not grow in a leaf, but rather across the surface. Small anchor cells of the fungus, haustoria, remain in the leaf and take nourishment from the rose plant, but the main filaments and the multitudes of spores it produces are outside the leaf. Newly unfurled leaves are more susceptible to infection than mature leaves. Increasing day lengths to 20–22 h were shown to reduce conidial production on greenhouse roses while maintaining postharvest quality (Suthaparan et al. 2010). Brief exposure to red light during the dark period may also do the same. Small, black fruiting structures (chasmothecia) also allow the fungus to

Fig. 7 Powdery mildew symptoms are not generally seen on the flower petals but early infections can deform flower buds and affect the bloom (Jay W. Pscheidt # 2017. All Rights Reserved.)

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overwinter on plant parts and debris. Ascospores from chasmothecia initiate new infections. The disease can develop under relatively dry conditions so long as the air is humid. Plants in shade tend to have more problems because that environment favors disease development. Multiple disease cycles occur during the growing season. Climbers, ramblers, and hybrid teas are susceptible. The natural genetic variability of the fungus means roses found resistant in one location may be susceptible in another due to the presence of different fungal strains. Also, resistant roses may become susceptible after a few years due to changes in the local fungal population. Management • Cultural practices – Plant cultivars known to be powdery mildew resistant in your area. Note that the cultivar Carefree Spirit was the first landscape shrub to survive for 2 years in AARS tests without any fungicide and voted a winner. Isolate susceptible cultivars. Space plantings for good air circulation between plants. Prune canes when dormant for an open habit also for good air circulation. Remove and destroy diseased canes. Rake and destroy fallen leaves. Use a highpressure water hose to thoroughly wet all leaf and cane surfaces in the early afternoon so plants dry quickly. May need to do this two or three times a week. In Western Oregon, this reduced but did not eliminate powdery mildew. Briefly exposing plants to red light during the dark interval has some utility against powdery mildew in greenhouse production. • Fungicides – Applying a dormant spray of lime sulfur at one part lime sulfur to nine parts water may help a planting that had severe powdery mildew the year before. Foliar applications of potassium salts have been found in Colombia to help reduce fungal sporulation. Apply foliar fungicides during the growing season, starting in early spring when young growth first appears. May need frequent applications, depending on the fungicide, to control the disease. Almost every fungicide on the market is registered for this disease on this crop, but only one or two materials are needed at any one time. Alternate or tank-mix products with different modes of action to prevent buildup of resistant fungal populations. Rosarians may not like the plant growth regulation effect (slight stunting) that may result from the use of FRAC group 3 or group 8 fungicides. Use of sulfur, which is a very effective organic material, can also stunt shoots, and frequent handling of sprayed foliage can irritate skin. Chlorothalonil-based products may be a problem on “Knock Out” and “Double Delight” roses resulting in damaged foliage. Always watch for warnings on labels of various products. • Biological control – Various Bacillus-based products are available and may have efficacy beyond the use of water alone but will not be as effective as many synthetic materials.

2.14

Replant Disease (Many)

Geographic occurrence and impact. The problem appears to be important on roses in Europe but has not been recognized in the Pacific Northwest of North America.

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Symptoms. The poor growth produced in replant situations may not be noticed unless you had a comparison. Roses planted into soil that has never grown roses (virgin soil) or on fumigated or sterilized soil would produce much more luxuriant, vigorous growth than when the same rose is replanted in an area that has grown roses for many years. Biology and epidemiology. Replant diseases are complexes of biological and environmental factors that vary by geographic region (Pscheidt and Ocamb 2016). This is a real disease complex in the Pacific Northwest (PNW) of the USA for many crops in the rose family including pome fruit such as apple and pear and stone fruit such as cherry and peach. Similar problems exist for other crops such as strawberry (black root rot). In the PNW, roses planted into grounds that have grown roses before appear to grow fine. The Washington Park International Rose Test Garden in Portland OR reports that they routinely remove a thousand plants every winter and replant a thousand new ones. They have not noted a replant problem. Management • Cultural practices – When replanting roses, follow correct planting practices. Correct any soil problems such as pH or drainage. You can also plant into new areas of your production site or rotate out of rose crops for several years before planting roses again on the same ground.

2.15

Rust (Phragmidium mucronatum (Pers.) Schltdl. 1824 and Many Other Species.)

Geographic occurrence and impact. Worldwide. Repeated defoliation leads to low vigor, inferior blooms, and high susceptibility to winter injury. Phragmidium americanum (Peck) Dietel 1905 – Eastern North America and Japan P. andersonii Shear 1902 – Northern North America; Asia; Europe P. bulbosum (F. Strauss) Schltdl. 1824 – Africa, Asia, Europe P. butleri Syd. & P. Syd. 1907 – Asia P. fusiforme J. Schröt. 1870 – Temperate northern hemisphere P. kamtschatkae (H.W. Anderson) Arthur & Cummins 1933 – Asia, Europe P. montivagum Arthur 1909 – Temperate North America and Asia; USSR P. mucronatum (Pers.) Schltdl. 1824 – Worldwide P. occidentale Arthur 1901 – North America P. rosae-arkansanae Dietel 1905 – Central North America P. rosae-californicae Dietel 1905 – Western North America P. rosae-pimpinellifoliae Dietel 1905 – North America; temperate Europe P. rosicola (Ellis & Everh.) Arthur 1934 – Central North America P. speciosum (Fr.) Burrill 1875 – North America P. tuberculatum J. Müll. 1885 – Asia, Africa, Europe, North and Central America

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Fig. 8 Note the two different rust pustules on this leaf, orange uredia and black telia (Jay W. Pscheidt # 2017. All Rights Reserved.)

Symptoms/signs. Small orange pustules (aecia) appear early in spring on both leaf surfaces. Later, the pustules enlarge and become more numerous on lower leaf surfaces (Fig. 8). Mottled and chlorotic areas may develop on upper leaf surfaces opposite the spots (uredinia) on the lower surfaces. In late summer and fall, the small pustules turn black (telia) and contain the winter spore stages of the rust. Stems occasionally are infected. Biology and epidemiology. Many species of these autoecious fungi (fungi that complete their life cycle on one plant host) are found throughout the world. They overwinter on diseased leaves and stems. Wind blows spores to healthy foliage. They germinate and infect through the stomata when leaves are wet for 2–4 h. Mild, humid weather favors disease development. Management • Cultural practices – Rake up all dead leaves and prune out infected and dead wood during the dormant season. Plant resistant cultivars. Removing infected leaves early in the season may be effective in some small plantings. • Fungicides – Focus applications during wet weather. Wetting agents will help with many of fungicides if allowed by the label. Many fungicides used for black spot will be helpful to control rust.

2.16

Spot Anthracnose (Elsinoe rosarum Jenk. & Bitanc. 1957)

Geographic occurrence and impact. Temperate regions. Repeated defoliation leads to low vigor, inferior blooms, and high susceptibility to winter injury. Symptoms/signs. At first, the appearance of red spots that vary from brown or dark purple on the upper leaf surfaces occurs (Bagsic et al. 2015). Spots up to 0.5 cm in diameter may be scattered or grouped and sometimes overlapping. Chlorosis or

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yellowing of the leaves may also occur. Later, spots become ashen white with a dark red margin. This tissue may fall off of the lower leaf surface, leaving a thin papery membrane or fall out entirely resulting in a shothole symptom. Symptoms occur on stems, hips, and pedicels as well as leaves. Can be easily confused with black spot or Cercospora leaf spot. Biology and epidemiology. The fungus overwinters on infected leaves and stems with conidia continuously being formed in early spring and through the summer. Spore dispersal primarily occurs with the aid of water through rain and irrigation. There were 5 races of the fungus identified in Germany (Bagsic et al. 2015). Management • Cultural practices – Remove and destroy infected plant parts. Space or prune bushes to allow for good airflow. • Fungicides – No specific chemical control has been reported. Fungicides in the FRAC group 3 used to control black spot, powdery mildew, or rust are also being used by growers to control this disease (Bagsic et al. 2015).

2.17

Verticillium Wilt (Verticillium dahliae Kleb. 1913)

Geographic occurrence and impact. Worldwide. Symptoms. At first, leaves near the growing point of young canes wilt, and lower leaves yellow (Hammett 1971). Sometimes mature leaves become necrotic between veins while the veins remain green. Defoliation progresses from the base of canes to the tip. Permanent wilt, defoliation, and death also may occur. If only a few canes are infected, they may grow normally next season or dieback. Symptoms generally are more severe in the greenhouse than in the field. The characteristic vascular discoloration in other plants is not evident in rose. Biology and epidemiology. This fungus survives a long time in soil and can infect a wide range of hosts. Rootstocks such as Rosa odorata and “Ragged Robin” are susceptible; R. multiflora and “Dr. Huey” have more resistance. R. chinensis var. manetti is very resistant. Under favorable growing conditions, plants may be able to tolerate infection. The fungus grows into the xylem where it colonizes the plant through mycelial growth and conidial production. Fluid movement in the xylem passively transports the conidia. Once in the xylem, this fungus partially blocks water movement and produces toxins that result in wilt symptoms. Wilting occurs under periods of water stress such as midsummer heat and drought. After diseased plant parts die, microsclerotia form and live several years in soil. Many weeds are susceptible and can help the fungus survive and disperse. Send soil samples to any of various private and public laboratories to assay for Verticillium propagules. Nurseries may wish to test individual core samples to

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determine the distribution in a particular field. The presence of any microsclerotia in the soil should be interpreted as a potential disease risk. Management • Cultural practices – Use resistant rootstocks. Avoid planting in old vegetable fields. In the greenhouse, use steam-sterilized soil or use a sterile soilless potting mix and/or hydroponics. Remove and destroy symptomatic or dead branches preferably before leaves fall and thus before new inoculum gets incorporated into the ground. Clean pruning equipment after use. Mulch with conifer-based products as conifers are resistant to infection. General yard waste mulch could contain infected plants and thus inoculum of the fungus.

2.18

Miscellaneous Fungal and Oomycete Diseases of Minor Importance or Considered Rare

Alternaria leaf spot – Alternaria spp. Phytophthora root rot – Phytophthora spp. have been identified on rose samples with root rot from the PNW several times by the Oregon State University (OSU) Plant Disease Clinic. Also reported from Asia, Europe, and North America. Septoria leaf spot – Septoria rosae Desm. 1831. Southern blight – Sclerotium rolfsii Sacc. 1911. Rhizoctonia root and stem rot – Rhizoctonia spp. – Japan. Note: Fungal names based on Farr and Rossman (2015).

3

Bacterial and Phytoplasma Diseases

3.1

Bacterial Cane Blight (Pseudomonas syringae – Undetermined Pathovar)

Geographic occurrence and impact. Treasure Valley of Idaho, USA (Mohan and Bijman 2010). Symptoms/signs. The symptoms usually start at the base of a vegetative bud or at leaf scars or wounds, as reddish-brown areas on the bark that later turn dark purple to black and necrotic (Fig. 9). The necrotic areas expand around and along the cane, often involving a major part or even the entire cane. Vegetative buds on the affected parts of the cane turn brown and dried. The surface of the necrotic areas of the bark is glossy, and the tissue beneath the epidermis is brown to dark brown and moist in the early stages. Often confused with winter injury. Biology and epidemiology. This disease is different from the various symptoms on roses previously attributed to P. syringae and/or P. syringae pv. morsprunorum

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Fig. 9 Glossy, dark purple to black, necrotic bark of canes with bacterial cane blight (Krishna Mohan # 2017. All Rights Reserved.)

(see below) in the literature. It has been an aggressive cane blight observed in the Treasure Valley of Idaho, USA, since 1996. Symptoms were common under cool, wet conditions in spring (March to May), and the level of incidence and severity of the disease varied from year to year. Several cultivars of climbing, floribunda, grandiflora, hybrid tea, hybrid perpetual, miniature, and shrub roses can show severe symptoms. Management • Cultural practices – The following is suggested in the absence of specific research to control this problem. Remove and destroy infected stems. Disinfect pruning shears before cutting more stems. Copper-based products are also recommended before fall rains begin and again when half the leaves have fallen.

3.2

Bacterial Leaf Spot and Blast (Pseudomonas syringae pv. morsprunorum)

Geographic occurrence and impact. Pacific Northwest of North America – rare. Symptoms. Dark brown, sunken spots appear on leaves, flower stalks, and calyx parts. Flower buds may die without opening. Black streaks appear on 1-year-old stems. Biology and epidemiology. The disease has been diagnosed only four times in the PNW from 1962 to 1992. Most common in cool, wet spring weather.

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Management • Cultural practices – Remove and destroy infected stems. Disinfect pruning shears before cutting more stems. Copper-based products are also recommended before fall rains begin and again when half the leaves have fallen. Repeat in spring to protect new growth.

3.3

Crown Gall (Agrobacterium radiobacter (Beijerinck and Van Delden 1902) Conn 1942 or Also Known as Rhizobium radiobacter (Beijerinck and Van Delden 1902) Young et al. 2001)

Geographic occurrence and impact. Worldwide and common. Symptoms. Galls are often at or just below the soil surface in the basal or root crown region. They may frequently be on roots but less frequently on aerial plant parts such as stem and leaves. Galls are usually rounded with a rough, irregular surface (Fig. 10). They first appear as small protuberances on the plant surface. Young, actively developing galls are light green or nearly white, and the tissue is soft. As they age, galls become dark and woody. Outer portions can slough off with age. Sometimes galls have a rather smooth surface, which makes it difficult to distinguish between gall and callus growth, especially if the gall is at the plant base or at the graft or bud union. Plants can be stunted and have reduced vigor, poor foliage, and fewer blossoms. A single gall at the plant’s base may be more detrimental than several galls on canes and roots. Symptoms may not develop for over a year if infection occurs when temperatures are below 15  C (59  F). Biology and epidemiology. The bacterium enters plants through wounds, either natural or caused by pruning, grafting, mechanical injury from cultivation, heaving

Fig. 10 Galls can be seen above, below ground or starting at pruning wounds (Jay W. Pscheidt # 2017. All Rights Reserved.)

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of frozen soils, chewing insects, or the emergence of lateral roots. Systemic populations that initiate disease do not seem to be as important in rose as they are in grape and walnut. After the bacterium enters a wound, a small piece of its DNA is transferred into the plant’s DNA. The foreign DNA transforms normal plant cells in the wounded area into tumor cells. Once transformed, tumor cells proliferate automatically. The result is a gall: a disorganized mass of hyperplastic and hypertrophic tissue. Pruning tools that cut through galls can become contaminated with the bacteria and spread them to cut surfaces of subsequently pruned plants. Galls breaking down in soil release bacteria, which can be transported by moving soil or water. In the absence of plant roots, bacterial populations gradually decrease; however, the pathogen may survive in soil for as long as 3 years. Management • Cultural practices – Use only pathogen-free nursery stock. Inspect new plants; do not plant any rose that has galls. Avoid wounding plants, especially at planting. If root pruning at planting, soak in the biocontrol agent listed below. Use plants with resistant rootstocks. Rootstocks differ widely in susceptibility. Prune off any galls on aerial parts of the plant. Disinfect pruning shears frequently. Clean shears and long soak times improve the disinfectant’s efficacy. Remove and destroy badly affected plants. Preplant soil solarization has been effective against this disease for cherry nursery stock grown in Western Oregon and may be useful for roses. Place clear plastic (anti-condensation coating) on rototilled ground, irrigated to near field capacity, from mid-July to mid-September. Solarization is more effective on sandy loam soil. The technique may help after removing diseased plants from a bed in which roses will be planted again. • Biological control – Agrobacterium radiobacter strain 84 has been used successfully with roses in Australia, New Zealand, and Spain but has not been effective in limited trials in the USA. Strain K 84 is preventive only. Agrobacterium radiobacter strain K1026 is a genetically modified strain of K84 that will help reduce the potential for development of resistant crown gall bacteria. Latent infections (symptomless) and existing galls are not controlled. A suspension of strain 84 may be used as a soak or spray. Thoroughly cover grafting wood, roots, and crown. Spray to runoff. To be effective, it must be applied a few hours after wounding.

3.4

Miscellaneous Bacterial Diseases of Minor Importance or Considered Rare

Hairy root – Agrobacterium rhizogenes (Riker et al. 1930) Conn 1942

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4

Viral Diseases

4.1

Rose Leaf Curl ((A virus is suspected))

Geographic occurrence and impact. California. Symptoms/signs. Spring leaves are small, leaflets detach easily, leaf epinasty, necrosis of shoot tips, and yellow vein flecking occur. Shoots are pointed with a broad base. Plants may recover in summer but show symptoms again in fall. Biology and epidemiology. Little is known about this disease other than natural spread appears to be slow. Management • Cultural practices – Purchase stock that is clean and/or certified as tested and free of all known viruses. Remove and destroy infected plants. Heat-treat scion stock plants 4 weeks at 38  C (100  F) before grafting.

4.2

Rose Mosaic [Prunus necrotic ringspot virus (PNRSV), Apple mosaic virus (ApMV), Arabis mosaic virus (ArMV), and Strawberry latent ringspot virus (SLRSV)]

Geographic occurrence and impact. Worldwide. Symptoms/signs. Symptoms may range widely depending on time of year, temperature, and type of virus(es) infecting the plant. Characteristic symptoms include chlorotic line patterns (zigzag pattern), ringspots, and mottles in leaves sometime in the growing season (Fig. 11). There may also be yellow net and yellow mosaic symptoms. Symptoms often are evident in spring and early summer but may not be on leaves produced in summer. Vein banding may be on leaves in long hot periods. Flower distortion, reduction in flower production, flower size, stem caliper at the graft union, winter survival, and early leaf drop and increase susceptibility to cold injury have all been reported. Some infected cultivars may not show any symptoms at all. Biology and epidemiology. Several viruses are associated with the range of symptoms of rose mosaic, including Prunus necrotic ringspot virus (PNRSV), Apple mosaic virus (ApMV), Arabis mosaic virus (ArMV), and Strawberry latent ringspot virus (SLRSV). The disease does not spread naturally and has no known insect vector, but grafting transfers it to healthy plants. Transmission of ArMV and/or SLRSV could occur by Dagger nematodes, but this has not been extensively studied. Viruses can be in the rootstock or scion or both and may not show symptoms.

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Fig. 11 Each of these leaves were from the same plant with rose mosaic (Jay W. Pscheidt # 2017. All Rights Reserved.)

“Madame Butterfly,” “Ophelia,” and “Rapture” are highly susceptible. Some report the disease does not spread; others indicate it may spread very slowly over many years. Root grafting between infected and healthy plants can also spread the disease (Golino et al. 2011). Management • Cultural practices – Purchase stock that is clean and/or certified as tested and free of all known viruses. Remove and destroy infected plants. However, the disease will not spread unless you propagate from or onto an infected bush. Heat-treat scion stock plants 4 weeks at 38  C (100  F) before grafting.

4.3

Rose Rosette (Rose Rosette Virus)

Geographic occurrence and impact. Widespread east of the Rocky Mountains of North America. Plants decline and die after a few years. Symptoms. Rose rosette symptoms are complex and variable as plants of the same cultivar may have different symptoms at the same or different location(s). Infected plants may have foliage that is bright red throughout the summer rather than just in the spring and fall. Leaves may become unusually long and thin or strapped shaped.

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Increased thorniness and flattening of stems (fasciation) is often but not always observed. Symptomatic foliage is often more susceptible to winter kill and/or desiccation. Multiple shoots can emerge from a single node to form a witches’ broom. Witches’ brooms are easier to recognize in the winter months after most of the foliage has fallen. Unusually large masses of distorted flower buds may also occur that, in most cases, do not open. Infection can resemble herbicide (glyphosate) injury. Infected bushes will decline and die in 3–5 years. Cane mortality is usually observed in spring when symptomatic canes fail to push out new foliage. Large commercial plantings or private rose gardens can be decimated by rose rosette if the disease is left unchecked. Biology and epidemiology. All cultivated roses (shrub type, hybrid tea, floribunda, grandiflora, and miniature roses) are thought to be susceptible to the disease. The Knock Out rose cultivars are as susceptible to rose rosette as other types. The virus is vectored by an eriophyid mite, Phyllocoptes fructiphilus. Although these mites do not fly, they may “balloon” in air currents, as do dust particles, and thus can be spread surprisingly long distances. The closer a healthy rose is planted to an infected rose, the more likely it is to become infected. In Tennessee, rose beds located near a source of the virus have a pronounced edge effect where roses nearest the source are more likely to become infected. Distribution of infected plants in a large rose bed will appear random if the plants were infected prior to planting or if there is a great distance between the rose planting and the inoculum source. Management • Cultural practices – Purchase stock that is clean and/or certified as tested and free of all known viruses. Roses should be inspected for symptoms before being purchased. Purchase plants from a nursery where all roses appear to be healthy. Inspected regularly for symptoms and rogue entire plants soon as possible when found. Do not just prune off symptomatic stems. Rogued plants should be bagged on site before removal. At this writing, preliminary studies have not demonstrated that miticides are effective. Also, it is not known if disinfecting pruning shears will also aid in disease management. Roses reported to be resistant include: R. setigera, R. aricularis, R. arkansana, R. blanda, R. palustris, R. carolina, and R. spinosissima.

4.4

Rose Spring Dwarf (Rose Spring Dwarf-Associated Virus)

Geographic occurrence and impact. California and Chile. Symptoms/signs. A rosetting or balling of new growth (Salem et al. 2008). Leaves are curved, very short, and show vein clearing or a netted pattern. Leaves occur on

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arrested shoots, which may not elongate for several months. Canes may grow in a zigzag pattern during the growing season. Biology and epidemiology. Easily transmitted through vegetative propagation techniques. The virus has been found in aphids on affected plants but is not known if they are a vector of the disease or not (Rivera and Engel 2010). Management • Cultural practices – Purchase stock that is clean and/or certified as tested and free of all known viruses. Remove and destroy infected plants. Heat-treat scion stock plants 4 weeks at 38  C (100  F) before grafting.

4.5

Rose Yellow Mosaic (Rose Yellow Mosaic Virus)

Geographic occurrence and impact. North America. Symptoms/signs. Symptoms associated with RoYMV infection include yellow mosaic, ring mosaic, premature leaf senescence, and necrotic dark brown rings on canes. Symptoms first appeared in new growth 4–6 weeks postinoculation. Necrotic cane symptoms are observed only in the cultivar Ballerina. Symptoms persist throughout the season (Lockhart et al. 2008). Biology and epidemiology. Potyviruses such as RoYMV are known to be aphid transmitted. Management • Cultural practices – Purchase stock that is clean and/or certified as tested and free of all known viruses. Remove and destroy infected plants.

4.6

Miscellaneous Viral or Virus-Like Diseases of Minor Importance or Considered Rare

Rose flower break – England, New Zealand, and Australia – unknown causal agent Rose Streak – eastern and mid-North America and Europe – suspect Rose streak virus The following have only been associated with virus symptoms in rose and are not fully characterized; however, the four viruses commonly found in the rose mosaic complex were not present in plants infected with these viruses (Lockhart et al. 2008): Rose yellow leaf virus Rosa rugosa leaf distortion virus Rosa multiflora cryptic virus Rose chlorotic ringspot virus Rose necrotic mosaic virus

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Nematode Diseases (Meloidogyne hapla, Pratylenchus Penetrans, P. vulnus, and Xiphinema diversicaudatum)

Geographic occurrence and impact. Worldwide. More of a problem on sandy soils. Symptoms/signs. Specific diseases have not been linked with specific nematodes; however, specific nematodes are associated with various root symptoms. In general, nematodes disrupt the root system, which will result in general aboveground symptoms that can include reduced vigor, poor growth or flowering, stunting, nutrient deficiencies, chlorosis, and/or wilting. These may be combined and considered a general plant decline. Root swellings or galls occur from the feeding activity of root-knot nematodes (Meloidogyne hapla) or dagger nematodes (Xiphinema spp.). Dagger nematode form galls on the tips of feeder roots. Root-knot nematodes tend to form galls on smaller roots and, with magnification, the swollen females can also be seen. Excessive root branching may also be observed on root systems with root knot. Small root lesions that become necrotic can be observed with an infestation of Root-lesion nematodes (Pratylenchus spp.). The severity of the symptoms will depend on the soil type and ability of the rose to tolerate various sized populations of root-lesion nematodes. Biology and epidemiology. Nematodes that affect plant growth are classified by feeding behavior. Depending on the nematode, this will dictate what type of samples is needed for diagnosis: roots, soil, or both. Root-knot nematodes (Meloidogyne spp.) are sedentary endoparasites, which tunnel into the roots, establishing permanent feeding sites from which they do not move. The feeding site causes the formation of giant cells, hyperplasia of cortical and vascular parenchyma, and retardation of meristematic activity in root tips (Fig. 12). Root-lesion nematodes (Pratylenchus spp.) are migratory endoparasites that tunnel inside roots, feed inside roots, and freely move back into soil and on to new roots (Fig. 13). Dagger nematodes (Xiphinema spp.) are migratory ectoparasites feed from outside roots, moving from cell to cell and piercing them to feed without entering root

Fig. 12 Adult Meloidogyne sp. and egg mass (Tatiana Gomez Rodriguez # 2017. All Rights Reserved.)

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Fig. 13 Adult Pratylenchus sp. and Xiphinema sp. (Tatiana Gomez Rodriguez # 2017. All Rights Reserved.)

tissue. The galls that are produced at this feeding site are caused by a hyperplastic response of cortical cells. The cells increase two to three times in size and meristematic activity is retarded. Although barely studied in rose, these nematodes can be vectors for some of the viruses in the rose mosaic complex such as Arabis mosaic virus (ArMV) (Fig. 13). Soil types that are more sandy generally result in more nematode injury at lower population levels than less porous soils. Host ranges are wide for many of these nematodes making crop rotation impractical. Management • Cultural practices – Avoidance in floral production is accomplished by eliminating soil from the production system with soilless media and/or hydroponics. If soil must be used then steam treatment is necessary to eradicated nematodes from the soil. Although difficult, soil in planting beds can also be steam treated. Soil solarization is useful to reduce populations in the top foot of soil in order to get plants establish in infested soil. Thermal therapy after infestation of plants is also difficult but possible. Pretreat plants at 38 C for 24 h followed by immersing plants into hot water at 48 C for 35 min. Damage to rose roots with this treatment is possible. Use of resistant rootstocks is complicated by diverse reactions to different nematodes from or in different locations. • Chemical practices – Preplant fumigation can be effective in the field, but available (registered) nematicides are becoming limited each year.

References Aegerter BJ, Nunez JJ, Davis RM (2002) Detection and management of downy mildew in rose rootstock. Plant Dis 86:1363–1368 Bagsic I, Linde M, Debener T (2015) Genetic diversity and pathogenicity of Sphaceloma rosarum (teleomorph Elsinoë rosarum) causing spot anthracnose on roses. Plant Pathol. doi:10.1111/ ppa.12478

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Bowen KL, Roark RS (2001) Management of black spot of rose with winter fungicide treatment. Plant Dis 85:393–398 Chase AR (2015) Rose diseases and their control. Chase Agricultural Consulting. http://www.chaseagri culturalconsultingllc.com/resources/pdfs/articlesPdf/48ROSEDISEASESANDTHEIRCONTROL.pdf Farr DF, Rossman AY (2015) Fungal databases, systematic mycology and microbiology laboratory, ARS, USDA. http://nt.ars-grin.gov/fungaldatabases/ Golino DA, Sim ST, Cunningham M, Rowhani A (2011) Evidence of root graft transmission of two rose mosaic viruses, Prunus necrotic ringspot virus and Apple mosaic virus in rose rootstocks. Phytopathology 101:S62 Hagan AK, Akridge JR (2005) Chemical control of cercospora leaf spot on Fuchsia Meidiland ® shrub rose. Alabama Cooperative Extension PP-587 Hammett KRW (1971) Symptom differences between rose wilt virus and Verticillium wilt of roses. Plant Dis Rep 55:916–920 Hausbeck MK, Moorman GW (1996) Managing Botrytis in greenhouse-grown flower crops. Plant Dis 80:1212–1219 Horst RK, Cloyd RA (2007) Compendium of rose diseases and pests, 2nd edn. APS Press, St. Paul Intrama S (1968) Coniothyrium rose canker in Oregon. PhD thesis, Oregon State University. p 95 Lockhart B, Zlesak D, Fetzer J (2008) Identification and partial characterization of six new viruses of cultivated roses in the USA. In: XII international symposium on virus diseases of ornamental plants. Acta Hortic: Ag Exp Station Technical Bulletin #8. Oregon State College, Corvallis, Oregon 901:139–147 Milbrath JA (1946) Control of black mold fungus Chalaropsis thielavioides Peyr. on Manetti rose, vol 8, Oregon State University technical bulletin Mohan SK, Bijman VP (2010) Bacterial cane blight of rose caused by Pseudomonas syringae. Acta Hortic 870:109–113 Philley G, Hagen AK, Chase AR (2001) Chapter 76. Rose diseases. In: Jones RK, Benson DM (eds) Diseases of woody ornamentals and trees in nurseries. American Phytopathological Society Press, St. Paul Pie K, De Leeuw GTN (1991) Histopathology of the initial stages of the interaction between rose flowers and Botrytis cinerea. Eur J Plant Pathol 97:335–344 Pscheidt JW, Ocamb CM (eds) (2016) Pacific Northwest plant disease management handbook. Oregon State University, Corvallis Rivera PA, Engel EA (2010) Presence of rose spring dwarf-associated virus in Chile: partial genome sequence and detection in roses and their colonizing aphids. Virus Genes 41(2):295–297 Salem N, Golino DA, Falk BW, Rowhani A (2008) Identification and partial characterization of a new Luteovirus associated with rose spring dwarf disease. Plant Dis 92:508–512 Schnabel G, Agudelo P, Henderson GW, Rollins PA (2012) Aboveground root collar excavation of peach trees for Armillaria root rot management. Plant Dis 96:681–686 Suthaparan A, Stensvand A, Torre S, Herrero ML, Pettersen RI, Gadoury DM, Gislerød HR (2010) Continuous lighting reduces conidial production and germinability in the rose powdery mildew pathosystem. Plant Dis 94:339–344

Diseases of Snapdragon

25

Stephen N. Wegulo and A. R. Chase

Abstract

Snapdragons (Antirrhinum majus) are grown as bedding and container plants and as cut flowers. Diseases can be a limiting factor in the production of snapdragons. Fungal diseases commonly encountered in snapdragons include downy mildew, powdery mildew, Botrytis blight, and various leaf spots and root, crown, and stem rots. The main bacterial disease is a seedling blight caused by Pseudomonas spp. Impatiens necrotic spot virus (INSV) and tomato spotted wilt virus (TSWV) are the two major virus diseases affecting snapdragons. Root-knot and lesion nematodes can also attack snapdragons. Yield loss can be reduced and profitability increased by concerted efforts to manage these diseases. Management tactics include host resistance and cultural, chemical, biological, and physical control. Integration of as many of these tactics as practical will maximize the effectiveness of disease management. This chapter presents information on the occurrence, symptoms, biology, epidemiology, and management of individual diseases of snapdragon. Details are provided for diseases on which research has been done and published. Some diseases have only been observed and research has not been done or published. Such diseases are only briefly mentioned. Keywords

Anthracnose • Blight • Leaf spot • Root and crown rot • Wilt • Mildew • INSV • TSWV

S.N. Wegulo (*) Department of Plant Pathology, University of Nebraska-Lincoln, Lincoln, NE, USA e-mail: [email protected] A.R. Chase (*) Chase Agricultural Consulting LLC, Cottonwood, AZ, USA e-mail: [email protected] # Springer International Publishing AG 2018 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florists’ Crops Diseases, Handbook of Plant Disease Management, https://doi.org/10.1007/978-3-319-39670-5_24

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Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Anthracnose (Colletotrichum antirrhini, C. destructivum) . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Botrytis Blight (Botrytis cinerea) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Cercospora Blight (Cercospora antirrhini) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Downy Mildew (Peronospora antirrhini) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Phyllosticta Blight (Phyllosticta antirrhini) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 Phytophthora Stem Rot and Wilt (Phytophthora cactorum, P. cryptogea, P. parasitica) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Powdery Mildew (Golovinomyces orontii (Formerly Erysiphe cichoracearum); Oidium spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.8 Pythium Root Rot (Pythium spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.9 Rhizoctonia Basal Stem Rot (Rhizoctonia solani) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.10 Rust (Puccinia antirrhini) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.11 Verticillium Wilt (Verticillium albo-atrum, V. dahliae) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.12 Additional Fungal Diseases and Pathogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Bacterial Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Seedling Blight (Pseudomonas antirrhini) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Additional Bacterial Diseases and Pathogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Viral Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Impatiens Necrotic Spot Virus (INSV) and Tomato Spotted Wilt Virus (TSWV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Additional Virus Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Nematode Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 Root-Knot Nematode (Meloidogyne spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2 Additional Nematode Pathogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 Additional Parasites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 Nonparasitic Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1 Tip Blight . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2 Phytotoxicity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.3 Additional Nonparasitic Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Introduction

Snapdragons (Antirrhinum majus) are popular plants that have been grown for centuries. They originated in the Mediterranean region (Maree and Wyk 2010; Rogers 1992), but are now grown in many parts of the world as cut flowers and garden, bedding, and potted plants. Cultivar groups differ principally in plant size. Dwarf types are used in gardens, whereas intermediate and taller types are grown for cut flowers. They display a range of beautiful flower colors from white to various shades of orange, yellow, peach, pink, red, and purple (Creel and Raymond Kessler 2007; Maree and Wyk 2010). In the USA, snapdragons were first included in the Agricultural Census of Horticultural Specialty Crops in 1959. At that time, they ranked seventh as the most valuable cut flower and made up 3.2% ($4.5 million) of the total wholesale value of flowers produced in the USA ($142.5 million), mostly in midwestern and northeastern states (Rogers 1992). In 2006, the wholesale value of snapdragon as a cut

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flower in the USA for operations with sales worth $100,000 or more was $12.2 million or 3% of the total value ($411.3 million) of cut flowers (Jerardo 2007). Diseases can significantly reduce marketability of snapdragons as plants or cut flowers. The most common diseases include Botrytis blight, powdery mildew, downy mildew, leaf spots, and root and crown rots. Viruses and nematodes also can attack snapdragons. The diseases found in a given location or geographical area will vary depending on environmental conditions. Losses due to diseases can be mitigated by integrating available management tactics which include cultural practices, physical control, chemical control, biological control, and host resistance.

2

Fungal and Fungus-Like Diseases

2.1

Anthracnose (Colletotrichum antirrhini, C. destructivum)

Geographic occurrence and impact. Snapdragon anthracnose has been reported in the USA (Stewart 1900; Horst 2013) and in Japan (Tomioka and Nishikawa 2011). It occurs most commonly on snapdragons grown in the field or landscape, but can also occur on greenhouse-grown snapdragons. Symptoms/signs. The disease can occur on snapdragons at any growth stage. Leaf spots are circular and slightly sunken. They are initially yellowish green with indefinite borders, but soon become whitish with definite borders. Spots on stems are circular to elliptical and initially whitish with a narrow brown or reddish-brown border. Spots may enlarge and girdle the stem base causing death of lateral shoots. Spots on older, woody stems are sunken. Under high humidity, numerous acervuli form in diseased tissues on the leaves and stems, causing them to become smoky brown (Forsberg 1975; Nelson and Strider 1985; Pirone 1978). Biology and epidemiology. Conidia are produced in acervuli in a sticky mass. They are dispersed by splashing water and may be carried by wind over long distances. Free water is necessary for conidial germination and infection. Disease development is favored by moisture and high humidity. Therefore, the risk for epidemics is higher in outdoor than in greenhouse snapdragons especially during wet growing seasons (Nelson and Strider 1985). C. antirrhini is a synonym of C. gloeosporioides which has a wide host range (Farr et al. 1989). Management. Adjust heating and ventilation to lower humidity and prevent condensation in the greenhouse. Avoid overhead watering. Apply fungicides labeled for anthracnose control on ornamental crops. Some of the most effective products contain a strobilurin (like pyraclostrobin or trifloxystrobin), a triazole (like triadimefon) or chlorothalonil. Combinations like pyraclostrobin and boscalid are especially effective on anthracnose diseases on ornamentals. Where available and known, plant resistant cultivars.

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Botrytis Blight (Botrytis cinerea)

Geographic occurrence and impact. Botrytis blight on snapdragon has been reported in the USA (Horst 2013). It is most common on greenhouse-grown snapdragons, but occurs in field production of cut flowers and in the landscape. Symptoms/signs. Often the first symptom observed is wilting of flower spikes, a result of girdling of the stem in the region of the lowest flowers. The pathogen also causes a soft decay of flowers, stems, and seedlings which is accompanied by wilting (Fig. 1). On mature plants, most infections start in the flowers and progress downward. A gray mold consisting of mycelia and spores of B. cinerea covers diseased plant parts. Sclerotia (compact masses of mycelia) may form inside or on the surface of infected plants (Agrios 2005; Forsberg 1975; Nelson and Strider 1985; Pirone 1978). Biology and epidemiology. Botrytis blight is favored by high relative humidity and cool temperatures. The optimum temperature range for growth and sporulation of B. cinerea is 18–23  C/64–73  F. Free water on the plant surface is required for infection to occur. During favorable conditions, B. cinerea sporulates profusely, producing a mass of gray mycelium and spores. The fungus survives as mycelium or sclerotia in soil and plant debris. Spores are disseminated by air currents and will germinate and infect healthy plants if they land on wounds or senescent flowers. Insects such as moths, thrips, and fruit flies also can spread spores from infected to healthy plants (Forsberg 1975; Holz et al. 2004; Nelson and Strider 1985; Pirone 1978). Management. Scouting for detection of diseased plants coupled with sanitation and good cultural practices can be effective in reducing damage and losses caused by B. cinerea. Removal and destruction of infected plants and infested plant debris can help to reduce the amount of inoculum. Good aeration keeps the plant surface dry, Fig. 1 Wilting (center) of snapdragon caused by Botrytis cinerea (A.R. Chase # 2017. All Rights Reserved.)

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which prevents spores from germinating. In the greenhouse, humidity can be reduced by ventilation and heating. Use of resistant cultivars where available is an effective and inexpensive means of managing Botrytis blight. Application of chemical and biological fungicides was shown to be effective in controlling Botrytis blight on lisianthus (Wegulo and Vilchez 2006a) and is likely to be effective on snapdragon as well. Rotation is critical since resistance to many previously effective active ingredients has been common for Botrytis in ornamental crops (see Botrytis blight in ▶ Chap. 20, “Diseases of Lisianthus”).

2.3

Cercospora Blight (Cercospora antirrhini)

Geographic occurrence and impact. Cercospora blight has been reported on snapdragon in Guatemala (Muller and Chupp 1950), the USA (Bolick 1959), and Argentina (Nelson and Strider 1985). The disease is capable of causing significant economic losses. Symptoms/signs. Symptoms of Cercospora blight on snapdragon were described by Jackson (1960). On leaves, lesions are subcircular and discrete to confluent with dull-white or gray centers surrounded by light-brown, slightly raised margins. They measure 0.5–7 mm in diameter. Under high humidity, rapid expansion of lesions may occur, resulting in irregular, poorly defined gray or tan necrotic areas measuring up to 15 mm in diameter with occasional faint concentric zones. The fungus sporulates profusely in the centers of lesions. Heavily infected leaves become chlorotic and drop. On stems, lesions occur mostly at the base near the soil line. They are depressed and dull white to gray with brown margins. Initially they are discrete, subcircular, oval, or elliptical. Eventually they become confluent, measuring up to 4 cm long. Cortical tissue on the stem becomes necrotic and small longitudinal cracks develop in it. Sporulation is sparse in stem lesions. Infected plants with leaf and stem lesions are chlorotic and stunted. Biology and epidemiology. Conidia are disseminated by air currents, wind, or splashing water. Free water is necessary for spore germination and infection. Incubation period in greenhouse- and field-grown snapdragons inoculated at 8 weeks of age with a suspension of conidia and mycelial fragments was 21–30 days at an incubation temperature range of 16–32  C/61–90  F. Disease development was favored by warm temperatures (30  C/86  F) and wetness. The pathogen survived in dry leaf and stem tissues for at least 14 months and in leaf debris or in the soil for at least 3 months (Porter and Aycock 1967). Because seed transmission is common in Cercospora spp. plant host systems, it is possible that C. antirrhini can be seed transmitted. Farr et al. (1989) list Antirrhinum as the only host of C. antirrhini. Management. Remove and destroy infected plants or plant parts. In the field or landscape, burying plant debris by plowing will reduce inoculum. Rotate with non-host crops. Use pathogen-free seed. Avoid overhead irrigation. Space plants to

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allow adequate air circulation. In the greenhouse, keep humidity low through ventilation and aeration. Integrate these cultural tactics with spraying fungicides labeled for leaf spot control on ornamental crops. Thiophanate-methyl remains an excellent rotational choice in addition to strobilurins, triazoles, chlorothalonil, and fludioxonil.

2.4

Downy Mildew (Peronospora antirrhini)

Geographic occurrence and impact. Downy mildew on snapdragon has been reported in Ireland (Murphy 1937), England (Green 1937), Australia (Anon 1941), Italy (Garibaldi and Rapetti 1981), and the USA (Harris 1939; Kirby 1945; Yarwood 1947). The disease can be very destructive both in greenhouse- and field-grown snapdragons. Total loss can result from the disease (Yarwood 1947). Symptoms/signs. Downy mildew affects primarily seedlings, but snapdragons can be infected at all growth stages. On seedlings, a downward curling of leaves and a reduction in the size of the plant and leaves occur. Infected leaves appear pale green on the upper surface and have a downy gray, lavender to white fungal growth on the lower surface (Figs. 2 and 3). This fungal growth can also occur on stems and upper leaf surfaces of young succulent plants. Systemically infected plants can be severely stunted, wilt, and eventually die. Seedling death progresses from the top down to the soil surface. On larger plants, symptoms of systemic infection include stunting (Fig. 4), pale green leaves (Fig. 5), and lack of flowering. Systemic infection can result in rosetting of growing points. Commonly, the shoots die and infected plants produce many secondary shoots from the base. Biology and epidemiology. Snapdragon downy mildew is favored by high relative humidity and cool, wet conditions. Optimal temperatures for disease development range from 5  C to 21  C/41 to 70  F. Local infection is characterized by pale round areas on leaves and is rarely destructive. Systemic infection can be very destructive Fig. 2 Pale green appearance on the upper surface of a downy mildew-infected snapdragon leaf (S.N. Wegulo # 2017. All Rights Reserved.)

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Fig. 3 The lower surface of a downy mildew-infected snapdragon leaf showing a gray to white fungal growth (S.N. Wegulo # 2017. All Rights Reserved.)

Fig. 4 Stunting and wilting of a snapdragon plant (frontcenter) due to systemic infection by downy mildew (S.N. Wegulo # 2017. All Rights Reserved.)

especially on seedlings. The incubation period can be as short as 4 days. Environmental conditions during the incubation period are most critical in determining how much infection occurs in a snapdragon planting or seedling tray (Yarwood 1947). P. antirrhini sporulates in cool, humid conditions. During sporulation, examination of the underside of infected leaves reveals a grayish downy growth consisting of sporangiophores and sporangia (spores). Sporangia are spread by wind, air currents, or splashing water. When they land on healthy plants, they germinate by means of a germ tube and cause new infections. In Michigan, Byrne et al. (2005) found that in field-grown snapdragons, dew periods of 6 h or longer were associated with large releases of sporangia, whereas consecutive days with short leaf wetness periods were associated with low sporangial concentrations in the atmosphere. Thick-walled resting spores (oospores) form abundantly in the roots, stems, and petioles of

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Fig. 5 A snapdragon plant infected with downy mildew. Note the pale green color on the upper surfaces of leaves (S.N. Wegulo # 2017. All Rights Reserved.)

systemically infected plants and are the means by which the pathogen survives during unfavorable conditions. The host range of P. antirrhini is limited to Antirrhinum (Farr et al. 1989). Management. Keep relative humidity in the greenhouse below 85% by balancing heat and ventilation. Keeping leaves dry using fans can be effective in preventing infections, but can also increase disease spread because spores can be disseminated by air currents. Avoid overhead irrigation. Thoroughly inspect all seedlings or plug trays for downy mildew before transplanting. Remove and destroy infected plants taking care not to spread the spores. After handling infected plants, thoroughly wash hands with soap before handling healthy plants. Some snapdragon cultivars have been shown in tests to have good resistance to snapdragon downy mildew (Byrne et al. 2004). Fungicides have been shown to be effective in controlling snapdragon downy mildew. Wegulo and Vilchez (2006b) found fenamidone, mancozeb, fosetylAl, and dimethomorph to have very good to excellent control of snapdragon downy mildew. To achieve effective control, apply fungicides preventively. The best management strategy for downy mildew is to integrate as many management tactics as practically and economically possible.

2.5

Phyllosticta Blight (Phyllosticta antirrhini)

Geographic occurrence and impact. Phyllosticta blight, also known as Phyllosticta leaf spot, has been reported in the USA (Guba and Anderson 1919). It is damaging primarily on snapdragons grown outdoors, but can also occur in the greenhouse (Nelson and Strider 1985). Symptoms/signs. The pathogen can attack snapdragons at any growth stage. On leaves, initial symptoms are chlorotic spots that form green to black lesions. They

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Fig. 6 Phyllosticta leaf spot on snapdragon (A.R. Chase # 2017. All Rights Reserved.)

enlarge and form yellow to brown concentric rings. The spots are visible on both sides of the leaf and become cream colored and slightly sunken in the center (Fig. 6). In wet weather, the spots continue to enlarge until the entire leaf collapses. Infection of young leaves causes them to become distorted and curled, and they may shrivel and die but remain clinging to the stem. Lesions on petioles are brown and elongated and may girdle the petiole causing the leaf to droop and die. From the petioles, lesions spread to the stem and cause infections at the leaf axil. Pinpoint size black pycnidia form in diseased tissues. On stems, lesions are dark green to brown with no definite margins. They may elongate up to 3 cm and girdle the stem. They turn dark brown, brittle, and dry and pycnidia eventually form in them. Damping-off occurs as a result of rapid wilting and dying of all aboveground parts of the plant (Forsberg 1975; Guba and Anderson 1919; Nelson and Strider 1985). Biology and epidemiology. The pathogen overwinters as pycnidia on host debris. Moisture is required for conidial germination and infection. Conidia released from pycnidia are dispersed by splashing water. Spore germination was best at 25  C/ 77  F (Guba and Anderson 1919), indicating that moderate to warm temperatures

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favor disease development. Disease development continues until the plant dies under wet conditions but is arrested or considerably slowed under dry conditions. In greenhouse infection experiments, lesions appeared on unwounded and wounded leaves of snapdragon plants 9 and 5 days after spray inoculation with conidia, respectively, indicating that wounding is not necessary for infection. Pycnidia began to appear on the fourth or fifth day after lesion appearance (Guba and Anderson 1919). In addition to Antirrhinum, Penstemon is also a host of P. antirrhini (Farr et al. 1989). Management. Remove and destroy infected plants as soon as they are noticed. Remove and destroy plant debris at the end of the growing season. Avoid excessive and overhead watering. Space plants to allow good air circulation. Apply fungicides labeled for control of leaf spots on ornamental crops.

2.6

Phytophthora Stem Rot and Wilt (Phytophthora cactorum, P. cryptogea, P. parasitica)

Geographic occurrence and impact. Phytophthora stem rot and wilt has been reported in the USA (Gill 1960; Harris 1934). Gill (1960) cited reports of the disease in other countries including Rhodesia (now Zimbabwe), Madagascar, Mauritius, England, South Africa, India, and Australia. Losses of 50% or more in greenhousegrown snapdragons were reported by Gill (1960), indicating the destructive nature of the disease. Symptoms/signs. Symptoms of the disease as observed by Harris (1934) include wilting caused by girdling of the stem at or slightly above the soil line. Lesions initially appear as water-soaked areas on healthy white stem tissue. As the lesions enlarge, their older portions become yellow, brown, and eventually almost black. They enlarge, extending up and down, until the stem is girdled (Fig. 7). The outer portion of the stem may slough off exposing the hard woody xylem tissue. When Fig. 7 Phytophthora stem rot and wilt on snapdragon (A.R. Chase # 2017. All Rights Reserved.)

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initially infected, plants may wilt slightly during the day and recover at night. Severely affected plants wilt permanently within 2–3 days and later become dry and brown. Under humid and wet conditions, lesions may extend up the main stem onto the side branches. Biology and epidemiology. Phytophthora spp. have a wide host range. They survive as oospores, chlamydospores, or mycelium in infected plant tissues or in soil. Long-term survival is by means of oospores and chlamydospores. Under favorable conditions, oospores and chlamydospores germinate by means of zoospores, whereas mycelium grows and forms sporangia which release zoospores. The zoospores swim in water and infect roots or are splash dispersed onto aboveground plant parts where they cause infections. Development of Phytophthora stem rot and wilt of snapdragon is favored by moisture and moderate to warm temperatures (Gill 1960). Management. Harris (1934) observed differences in susceptibility of three snapdragon cultivars to Phytophthora stem rot and wilt, with disease incidence ranging from 20% in Cheviot Maid and 30% in Jenny Schneider to 75% in Roman Gold. Plant snapdragon cultivars are known to have good resistance to Phytophthora stem rot and wilt. Because the disease is favored by wetness, ensure good soil drainage. Avoid overhead watering. Keep plant surfaces dry by maintaining sufficient air circulation. Apply fungicides labeled for Phytophthora control on ornamental crops. Hausbeck and Glaspie (2009a, b, c) found the fungicides mandipropamid, fluopicolide, fenamidone, mefenoxam, and mono- and dipotassium salts of phosphorous acid to be effective in controlling Phytophthora root rot of snapdragons when applied as drenches.

2.7

Powdery Mildew (Golovinomyces orontii (Formerly Erysiphe cichoracearum); Oidium spp.)

Geographic occurrence and impact. Powdery mildew on snapdragon has been reported in England (Moore 1947) and the USA (Guba 1936). Symptoms/signs. A powdery, white fungal growth appears on both surfaces of leaves, but mostly on the upper surface. Plants can be infected at any growth stage. Infections usually begin on the lower leaves, but can become severe on the upper leaves, stems, and flowers. Severely infected leaves may wilt and die. Biology and epidemiology. Powdery mildew fungi are obligate parasites. Fungal growth occurs on the plant surface and is nourished by haustoria (special absorption structures that form in epidermal cells following infection). The fungal growth consists of mycelia and conidia which are dispersed by wind or air currents. When the conidia land on healthy plants, they germinate and cause new infections. Cleistothecia form in older infected plant tissues and produce ascospores. Disease

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development is favored by shade and high relative humidity but not free water. These fungi typically have a very narrow host range and survive as mycelium in infected plants or as cleistothecia in plant debris. Management. Space plants to allow good air circulation. Avoid planting snapdragons in shaded areas. In the greenhouse, reduce relative humidity by heating and venting. Apply fungicides labeled for powdery mildew control on ornamental crops. In trials conducted by Raabe et al. (1970), the systemic fungicide benomyl achieved complete control of powdery mildew on snapdragons when applied as a spray or a drench. Fungicides are most effective when applied preventively and especially include triazoles and strobilurins.

2.8

Pythium Root Rot (Pythium spp.)

Geographic occurrence and impact. Pythium root rot on snapdragons has been reported in the USA (Hanan et al. 1962). Symptoms/signs. Snapdragons can be attacked at any growth stage. Infection of roots in the root hair zone and colonization of cortical tissues causes a brown, watersoaked rot. Basal stem tissues may also rot; however, stem tissues above the soil line are rarely colonized. Pre- or postemergence damping-off of seedlings may occur. Infected seedlings that survive and are transplanted and young plants infected after transplanting are stunted, chlorotic and wilt when exposed to sunlight. Infected plants have a reduced root system. Wilt collapse is a common symptom observed in mature plants (Fig. 8) especially at flowering when water demand cannot be met by the reduced root system. Older plants tend to be more resistant to symptom development than younger plants. In experiments conducted by Mellano et al. (1970), 15-day-old or younger seedlings infected with Pythium ultimum died within 6 days due to rapid and unrestricted colonization of host tissue. In contrast, Fig. 8 Wilting caused by Pythium root rot (A.R. Chase # 2017. All Rights Reserved.)

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25-day-old plants were tolerant to infection and host colonization. Wilting and stunting intensified by high temperature was observed in infected tolerant plants. Biology and epidemiology. Pythium spp. have a wide host range. They live in the soil and produce a white mycelium that grows rapidly and gives rise to sporangia. A sporangium germinates directly by giving rise to a germ tube or by producing a short hypha bearing a balloon-like vesicle at its end. Within the vesicle, 100 or more zoospores are produced. When released from the vesicle, the zoospores move around in a swarm for a few minutes then form cysts which germinate by giving rise to a germ tube. The germ tube infects host tissue by direct penetration. The mycelium can also produce a female oogonium and a male antheridium. The antheridium produces a fertilization tube which enters the oogonium. Nuclei of the antheridium move through the tube and fuse with nuclei of the oogonium to form a zygote which becomes an oospore by producing a thick wall around itself. The oospore is resistant to adverse environmental conditions and is the survival and resting stage of Pythium spp. When environmental conditions become favorable, oospores germinate in a manner similar to that of sporangia by directly forming a germ tube or by first forming zoospores and cysts. Rootlets can be infected at any stage of plant growth. Root rot progresses upward to the crown and stem. Disease development is favored by wet and waterlogged soils (Agrios 2005). Zoospores are disseminated by flowing water or splash dispersal. In the greenhouse, the primary means of introducing Pythium propagules are contaminated plant material and soiled hands, tools, and hose ends (Daughtrey et al. 1995). Management. Cultural methods of control include good water drainage. If the planting site is poorly drained, plant snapdragons in raised beds. Composted bark can be incorporated into the soil in the raised bed to improve aeration. Drainage tiles can be used to facilitate drainage and direct water away from the planting site. Do not reuse pots or trays from a previous crop for propagation. If pots or trays are reused, they should sanitized or sterilized first. Fungicides registered for Pythium control such as sodium, potassium and ammonium phosphites, etridiazole + thiophanate-methyl, mefenoxam, and etridiazole can be used to control root rot in snapdragons (Pscheidt and Ocamb 2016). Del Castillo Múnera and Hausbeck (2015) found that the efficacy of fungicides in controlling Pythium root rot on snapdragons depended on the species Pythium. Mefenoxam and fenamidone reduced root rot caused by P. aphanidermatum and P. ultimum; whereas fluopicolide and etridiazole reduced root rot caused by P. ultimum; and fenamidone, potassium phosphite, and the biological control agent Trichoderma harzianum effectively controlled root rot caused by P. irregulare. In the same study, differences in intensity of root rot caused by Pythium spp. were observed among snapdragon cultivars, although none of the cultivars was highly resistant.

2.9

Rhizoctonia Basal Stem Rot (Rhizoctonia solani)

Geographic occurrence and impact. Rhizoctonia basal stem rot on snapdragon has been reported in the USA (Baker and Sciaroni 1952) and Taiwan (Yang and Leu 1981).

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Symptoms/signs. Snapdragons can be infected at any growth stage. They are most susceptible at the seedling stage and immediately following transplanting (Nelson and Strider 1985). Symptoms include pre- and postemergence damping-off. Infected seedlings that survive have red-brown lesions on the basal stem at the soil line. Wilting, collapse, and death occur if lesions girdle the stem. Affected tissues show a coarse brown mycelium of R. solani in wet and humid environments. Biology and epidemiology. R. solani has a wide host range. It survives as mycelium or sclerotia in infected plants or plant parts, soil, or plant debris. It is spread by water, contaminated tools, or infected plants or propagative plant material. Basal stem rot on snapdragon is favored by high temperatures and moderately moist soils (Baker and Sciaroni 1952). Management. Treat soil with heat (steam or solarization), fumigants, or fungicides labeled for control of Rhizoctonia in ornamental crops. Ensure good soil drainage and avoid excessive watering. Plant snapdragons on raised beds if the area is poorly drained. Use sterilized soil, trays, and tools to raise seedlings before transplanting. Transplant disease-free seedlings. In the greenhouse, keep relative humidity low by venting and heating. Space plants to allow good air circulation. Some of the most effective fungicides include fludioxonil, thiophanate-methyl, and strobilurins (like azoxystrobin and pyraclostrobin).

2.10

Rust (Puccinia antirrhini)

Geographic occurrence and impact. Snapdragon rust was first reported in the USA by Blasdale (1903) who had found it at Berkeley, CA, in 1895 (McClellan 1953). Peltier (1919) noted that the disease caused much loss in Illinois in the 4 years following its discovery in the state in 1913. McClellan (1953) echoed the seriousness of the disease by stating that “Of the numerous diseases of snapdragons, rust probably causes the most concern.” Snapdragon rust occurs throughout the world wherever snapdragons are grown (Gawthrop and Brooks 1979; Nelson and Strider 1985). Symptoms/signs. Seedlings, cuttings, and mature plants can be attacked by rust in the greenhouse as well as in the field. It is most severe on cuttings and on plants just before flowering (Peltier 1919). All above-ground parts of the plant except the florets can be infected. Initial symptoms on leaves are small, chlorotic swellings. These swellings increase in size until the epidermis ruptures, exposing red-brown uredinial pustules. Pustules are limited to the lower leaf surface. On the upper surface of the leaf above each pustule, a circular yellow area is apparent. The fungus continues to grow under favorable environmental conditions and in 4 or 5 days forms one or two concentric rings of smaller secondary pustules around the original pustule (Nelson 1962) (Fig. 9). In the presence of moisture and high relative humidity, the pustules are invaded by secondary microorganisms, mainly Fusarium spp., which cause irregularly shaped necrotic spots around the pustule (Dimock and Baker 1951). On stems, initial

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Fig. 9 Rust pustules on the undersides of snapdragon leaves (A.R. Chase # 2017. All Rights Reserved.)

small pustules elongate, causing swollen cankers which serve as entry points for secondary invaders. These invaders cause lesions that girdle the stem, resulting in wilting and plant death. Severely affected plants have a scorched or brown appearance. Flowers open prematurely and are smaller than normal. Biology and epidemiology. The host range of P. antirrhini is limited to Antirrhinum (Farr et al. 1989). Like other rust fungi, it is an obligate parasite. Urediniospores are spread by wind over long distances and by wind-blown rain or splashing water over short distances (Nelson and Strider 1985). They land on healthy plants and initiate new infections. Optimal conditions for infection are the presence of free water and a temperature range of 10–13  C/50–55  F. Disease development is optimal under a temperature range of 21–24  C/70–75  F and is slowed at higher temperatures (Dimock and Baker 1951). Under optimal conditions for disease development, new uredinia are formed within 8–16 days following inoculation (Peltier 1919), and the cycle of spore production is repeated. The urediniospore is the repeating spore of P. antirrhini. Teliospores form later in the growing season, but do not play an important role in the disease cycle. Under semiarid conditions, injury is primarily from desiccation through the rust pustules. In the presence of moisture and high relative humidity, severe injury is caused by secondary invaders, primarily Fusarium spp. (Dimock and Baker 1951). There are no known alternate hosts of P. antirrhini in the USA. The fungus survives as mycelium or urediniospores on snapdragon plants in the greenhouse or field. Management. Cuttings are extremely susceptible (Peltier 1919). Therefore, only cuttings from rust-free snapdragons should be used. Otherwise start plants from seed. Allow good drainage and good air circulation. Keep temperatures in the greenhouse above 24  C/75  C for several days and not below 16  C/61  F at night (Pscheidt and Ocamb 2016). Avoid overhead watering. Integrate the above cultural management tactics with application of fungicides registered for rust control on

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ornamental crops. Fungicides shown to provide good results on snapdragon rust include strobilurins (especially azoxystrobin) and triazoles which can show eradicant benefits at times. Mancozeb is also excellent but only as a true preventive application.

2.11

Verticillium Wilt (Verticillium albo-atrum, V. dahliae)

Geographic occurrence and impact. Verticillium wilt on snapdragons has been reported in the USA (Baker and Sciaroni 1952). Symptoms/signs. The pathogens infect roots and colonize the xylem vessels, inhibiting water movement. Initially the lower leaves may become yellow, wilt, and drop. Later wilting and collapse of single branches may occur, causing the plant to appear as if wilting on one side (Nelson and Strider 1985; UC IPM 2014). The entire plant finally wilts, collapses, and dies. During periods when cool days or nights alternate with warm days, wilting and plant death can occur rapidly. Wilting often occurs after the onset of blossom development. Vascular tissues are discolored continuously from the base of the stem to the top of the plant. Black microsclerotia may form in diseased tissue. Biology and epidemiology. Verticillium spp. are soilborne and have a wide host range. They survive over long periods of time as microsclerotia in the soil or plant debris. Introduction into non-infested fields or greenhouses is through microslerotia-contaminated soil or tools and mycelium in infected plant material such as cuttings. Disease development is favored by moisture and cool temperatures. Dutta (1981) showed that the severity of Verticillium wilt on snapdragon was higher in alkaline compared to acidic soil. Organic (chitin and green manure) and inorganic (ammonium sulfate, calcium nitrate, and combined NPK) soil amendments reduced disease severity, and this was attributed to boosting antagonistic microorganisms in the soil and direct nutritional effects on snapdragon plants (Dutta and Isaac 1979a, b; Isaac 1956). Excessive nitrogen fertilization increased disease severity (Isaac 1957). Management. Treat soil with heat (steam or solarization) or fumigants before planting or transplanting snapdragons. In the greenhouse, sanitize trays, tools, and benches. Remove and destroy infected plants and infested plant debris. Use diseasefree plants or cuttings. Avoid planting snapdragons in poorly drained soils.

2.12

Additional Fungal Diseases and Pathogens

The following fungal diseases and pathogens have been reported on snapdragon (Horst 2013): southern blight (stem rot, Sclerotium rolfsii), stem and crown canker (Myrothecium roridum), collar rot (Rhizoctonia solani), petal rot (Bipolaris setariae), charcoal rot (Macrophomina phaseoli), root rot (Phymatotrichum

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omnivorum, Thielaviopsis basicola), and stem rot and wilt (Fusarium sp., Sclerotinia sclerotiorum, S. minor). In addition, seed blight (Alternaria alternata) has been reported (Harman et al. 1973).

3

Bacterial Diseases

3.1

Seedling Blight (Pseudomonas antirrhini)

Geographic occurrence and impact. Seedling blight on snapdragon has been reported in Japan (Takimoto 1920), Australia (Valder 1963), and the United Kingdom (Simpson et al. 1971). Chase (2001, 2004) listed Pseudomonas as a causal agent of Pseudomonas leaf spot on certain cut flower cultivars of snapdragon in the USA (Fig. 10), but did not specify the species. Symptoms/signs. The disease is actually a leaf spot, but when severe it is better described as a seedling blight (Simpson et al. 1971). Leaf lesions are initially small, pale green, and less than 0.5 mm in diameter. They enlarge and become brown with a green halo. When fully developed, they are discrete, almost circular, and measure 4–5 mm in diameter. They appear sunken and papery brown with a defined dark brown margin. Water-soaked dark green zones may be seen around the lesions. As disease development progresses, the lesions coalesce, forming irregular blotches followed by leaf collapse. On dark-leaved cultivars, lesions are less discrete and the tissue surrounding the lesions may appear chlorotic (Simpson et al. 1971). Biology and epidemiology. Simpson et al. (1971) observed that seedling blight appeared on snapdragon seedlings usually after the first transplanting and before reaching a height of 15–18 cm. Beyond this growth stage, the disease did not attack newly emerged foliage. Disease development during the susceptible growth stages is Fig. 10 Seedling blight caused by Pseudomonas spp. (A.R. Chase # 2017. All Rights Reserved.)

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favored by moisture such as that from mist irrigation during propagation. The pathogen is spread by splashing water from rain or irrigation. Pathogenicity tests on 6 to 8-cm-tall seedlings showed that the incubation period was 10–12 days. Injury to leaf surfaces by an abrasive agent resulted in severe symptoms (Simpson et al. 1971). Seed transmission studies confirmed that P. antirrhini can be transmitted through seed. The host range of this pathogen is limited to members of the family Scrophulariaceae, of which snapdragon is the most susceptible. Management. Use pathogen-free seed and propagative plant material. Avoid excessive moisture on the foliage of seedlings. Remove and destroy infected seedlings. Plant resistant cultivars where known and available. Preventively apply bactericides labeled for disease control on ornamental crops. Bactericides can only prevent or reduce infections; they cannot eliminate established infections.

3.2

Additional Bacterial Diseases and Pathogens

Crown gall (Agrobacterium tumefaciens) has been reported on snapdragon (Horst 2013).

4

Viral Diseases

4.1

Impatiens Necrotic Spot Virus (INSV) and Tomato Spotted Wilt Virus (TSWV)

Geographic occurrence and impact. Impatiens necrotic spot virus has been reported on snapdragon in the USA (Daughtrey et al. 1997; Horst 2013). Significant losses can result from infection by these viruses because symptomatic plants are not marketable. Symptoms/signs. Symptoms of INSV on snapdragons can be variable and include brown to white spots on leaves, stunting, stem necrosis which causes tissues to turn black (Fig. 11), and plant death. Because symptoms are variable and resemble those caused by other diseases, they are not reliable for diagnosis. Confirmation is done by performing laboratory tests such as enzyme-linked immunosorbent assay (ELISA) or polymerase chain reaction (PCR). Biology and epidemiology. INSV is one of two viruses that infect a wide range of plant species including ornamentals, vegetables, and field crops. The other virus is tomato spotted wilt virus (TSWV). Both belong to the genus Tospovirus in the family Bunyaviridae and are transmitted by the western flower thrip (Daughtrey et al. 1997). Spread of these viruses is through movement of infected plants or viruliferous thrips which feed on and transmit the viruses to healthy plants. Many plant hosts can be infected but show no symptoms. These plant hosts which include weeds serve as reservoirs for the viruses and their vectors.

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Fig. 11 Necrosis on snapdragon caused by Impatiens necrotic spot virus (INSV) (A.R. Chase # 2017. All Rights Reserved.)

Management. Effective control can be achieved by virus exclusion and prevention of virus spread. Virus exclusion involves use of virus-indexed propagation material which can be purchased from certain suppliers. INSV and TSWV can also be excluded by preventing thrips from entering the greenhouse or propagation and storage areas. This can be achieved through the use of fine mesh screens. Weeds and other thrips and virus plant hosts growing in the vicinity of greenhouses should be controlled with herbicides or other means. Workers should take care not to introduce thrips into the greenhouse or propagation areas by changing clothes before entry and by not wearing clothes with colors such as yellow, green, or blue which attract thrips. Prevention of virus spread is achieved by controlling resident thrip populations with insecticides or biocontrol agents and by roguing and disposing of infected plants. Because virus-infected plants are more susceptible to the feeding and oviposition of thrips (Bautista et al. 1995), such plants should be disposed of as quickly as they are noticed.

4.2

Additional Virus Diseases

Cucumber mosaic virus, an unidentified mosaic virus, and an unidentified ring spot virus have been reported on snapdragon (Horst 2013).

5

Nematode Diseases

5.1

Root-Knot Nematode (Meloidogyne spp.)

Geographic occurrence and impact. Root-knot nematode has been reported on snapdragon in the USA (Horst 2013; Pirone 1978). Symptoms/signs. Root galling and stunting.

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Biology and epidemiology. Root-knot nematodes have a wide host range. They feed and reproduce within plant roots, inducing small to large galls known as rootknots. The adult females are sedentary in plant roots and have a round to pear-shaped body. Adult males are vermiform, leave the root, and become free-living in the soil. Adult females lay eggs in gelatinous sacs that are deposited on the surface of galled roots or may remain within the galls. The egg develops into a vermiform first-stage juvenile (J1) which molts into a vermiform second-stage juvenile (J2) that emerges from the egg into the soil. If the J2 (the only infective stage) encounters a susceptible host, it enters the root and becomes sedentary. It feeds on the root cells surrounding its head and grows, becoming sausage shaped. The saliva it secretes causes the root cells to enlarge and form galls. The J2 molts into a third-stage juvenile (J3) which molts into a fourth-stage juvenile (J4). Both the J3 and J4 are sedentary within the root. The J4 undergoes a final molt into a mature male which leaves the root or a mature female which remains sedentary in the root and lays eggs. Root-knot nematodes are spread by contaminated equipment, infected plant material, or water. Nematodes attack ornamental plants in the landscape or field; they do not seem to be present in greenhouse- or container-grown crops (Creswell 2009; Ismail 1980; McSorley 1994; McSorley and Fredeik 1994). Management. Sanitize equipment after use. Contaminated soil can be disinfested by fumigation, drenching with nematicides, or heat treatment with steam or solarization. Refer to ▶ Chap. 8, “Soil/Media Disinfestation for Management of Florists’ Crops Diseases” for specific strategies.

5.2

Additional Nematode Pathogens

The lesion nematodes Pratylenchus spp. have been reported on snapdragon (Baker and Sciaroni 1952; Horst 2013).

6

Additional Parasites

Dodder (Cuscuta sp.) has been reported on snapdragon (Horst 2013).

7

Nonparasitic Disorders

7.1

Tip Blight

A tip blight of unknown cause has been described on snapdragons (Forsberg 1975; Horst 2013). Leaves gradually wilt starting at the plant tip and the wilt progresses to affect petioles and stems. Sunken, water-soaked lesions may form on the stem and girdle it, causing the plant tip to wilt and die. This disorder usually occurs in winter when it is cloudy and disappears in spring when brighter weather returns.

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Fig. 12 Phytotoxicity (second and third leaves from left) caused by an experimental fungicide on snapdragon. The first, fourth, and fifth leaves from left are from plots sprayed with commercial fungicides. The sixth leaf is from a non-sprayed check plot (S.N. Wegulo # 2017. All Rights Reserved.)

7.2

Phytotoxicity

Certain pesticides may cause injury to snapdragons. Figure 12 shows injury to snapdragon leaves following application of an experimental fungicide.

7.3

Additional Nonparasitic Disorders

Fasciation has been reported on snapdragon. It is probably a genetic abnormality (Horst 2013).

References Agrios GN (2005) Plant pathology, 5th edn. Elsevier Academic Press, New York Anonymous (1941) Downy mildew of snapdragons. Agric Gaz N S W 52:538–539 Baker KF, Sciaroni RH (1952) Diseases of major floricultural crops in California. California State Florists’ Association, Los Angeles Bautista RC, Mau RFL, Cho JJ, Custer DM (1995) Potential of tomato spotted wilt tospovirus plant hosts in Hawaii as virus reservoirs for transmission by Frankliniella occidentalis (Thysanoptera: Thripidae). Phytopathology 58:953–958 Blasdale WC (1903) On a rust of the cultivated snapdragon. J Mycol 9:81–82 Bolick JH (1959) Cercospora antirrhini found in Florida. Plant Dis Rep 43:511

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Byrne JM, Sconyers LE, Hausbeck MK (2004) Evaluation of snapdragon cultivars for resistance to downy mildew, 2000 and 2001. B&C Tests 19:O010 Byrne JM, Hausbeck MK, Sconyers LE (2005) Influence of environment on atmospheric concentrations of Peronospora antirrhini sporangia in field-grown snapdragon. Plant Dis 89 (10):1060–1066 Chase AR (2001) Controlling bacterial diseases of ornamentals. Southeast Floriculture, July/ Aug:20–21 Chase AR (2004) Bacterial diseases: are we losing the battle? GPN Mag, April 2004:32–35 Creel R, Raymond Kessler J (2007) Greenhouse production of bedding plant snapdragons. Alabama Cooperative Extension System. Publication No. ANR-1312 Creswell T (2009) Nematode management in bedding plants in the landscape. NCSU. Ornamental Disease Note 31 Daughtrey ML, Wick RL, Peterson JL (1995) Compendium of flowering potted plant diseases. APS Press, St. Paul Daughtrey ML, Jones RK, Moyer JW, Daub ME, Baker JR (1997) Tospoviruses strike the greenhouse industry. Plant Dis 81(11):1220–1230 Del Castillo Múnera J, Hausbeck MK (2015) Integrating host resistance and plant protectants to manage Pythium root rot on geranium and snapdragon. HortScience 50(9):1319–1326 Dimock AW, Baker KF (1951) Effect of climate on disease development, injuriousness, and fungicidal control, as exemplified by snapdragon rust. Phytopathology 41:536–552 Dutta BK (1981) Effect of the chemical and physical condition of the soil on Verticillium wilt of Antirrhinum. Plant and Soil 63(2):217–225 Dutta BK, Isaac I (1979a) Effects of inorganic amendments (N, P and K) to soil on the rhizosphere microflora of Antirrhinum plants infected with Verticillium dahliae Kleb. Plant and Soil 52 (4):561–569 Dutta BK, Isaac I (1979b) Effects of organic amendments to soil on the rhizosphere microflora of Antirrhinum infected with Verticillium dahliae Kleb. Plant and Soil 53(1):99–103 Farr DF, Bills GF, Chamuris GP, Rossman AY (1989) Fungi on plants and plant products in the United States. Univeristy of Illinois: APS Press, St. Paul Forsberg JL (1975) Diseases of ornamental plants. Special publication no. 3 revised, University of Illinois: Urbana-Champaign Garibaldi A, Rapetti S (1981) Grave epidemia di peronospora su antirrino [Antirrhinum majus L.] (Downy mildew attacks on Antirrhinum majus L.). Colture Protette 11:35–38 Gawthrop F, Brooks A (1979) The menace of Antirrhinum rust. Garden J Royal Hortic Soc 104:68–70 Gill DL (1960) A stem and branch rot of snapdragon. Plant Dis Rep 44(12):946–947 Green DE (1937) Downy mildew on Antirrhinum majus: a disease new to Great Britain. Gard Chron 102:27–28 Guba EF (1936) Plant disease notes from Massachusetts. Plant Dis Rep 20:302–303 Guba EF, Anderson PJ (1919) Phyllosticta leaf spot and damping off of snapdragons. Phytopathology 9:315–325 Hanan JJ, Langhans RW, Dimock AW (1962) Soil aeration and the Pythium root rot disease of snapdragon. Bull N Y State Flower Grow 195:1–6 Harman GE, Heit CE, Pfleger FL, Braverman SW (1973) Snapdragon seed blight – a serious problem caused by seedborne fungi. Plant Dis Rep 57(7):592–595 Harris MR (1934) A Phytophthora disease of snapdragons. Phytopathology 24:412–417 Harris MR (1939) Downy mildew on snapdragon in California. Plant Dis Rep 23:16 Hausbeck MK, Glaspie SL (2009a) Residual control of Phytophthora root rot of snapdragons, 2007. Plant Disease Management Reports 3:OT014 Hausbeck MK, Glaspie SL (2009b) Control of Phytophthora root rot of dwarf snapdragons with fungicide drenches, 2008. Plant Disease Management Reports 3:OT015 Hausbeck MK, Glaspie SL (2009c) Control of Phytophthora root rot of cut flower snapdragons with fungicide drenches, 2008. Plant Disease Management Reports 3:OT014

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Holz G, Coertze S, Williamson B (2004) The ecology of Botrytis on plant surfaces. In: Elad Y, Williamson B, Tudzynski P, Nelson N (eds) Botrytis: biology, pathology and control. Kluwer, London, pp 9–27 Horst RK (2013) Field manual of diseases on garden and greenhouse flowers. Springer, New York Isaac I (1956) Some factors affecting Verticillium wilt of Antirrhinum. Ann Appl Biol 44:105–112 Isaac I (1957) The effects of nitrogen supply upon the Verticillium wilt of Antirrhinum. Ann Appl Biol 45:512–515 Ismail W (1980) Susceptibility of some ornamental plants to the attack of Meloidogyne incognita. Indian J Hortic 37:326–328 Jackson CR (1960) Cercospora blight of snapdragon. Phytopathology 50:190–192 Jerardo A (2007) Floriculture and nursery crops yearbook/FLO-2007. Economic Research Service, United States Department of Agriculture Kirby RS (1945) Downy mildew on snapdragon in Pennsylvania. Plant Dis Rep 29:371 Maree J, Wyk B-E (2010) Cut flowers of the world: a complete reference for growers and florists. Timber Press, Portland McClellan WD (1953) Rust and other disorders of snapdragon. In: Yearbook of agriculture 1953. United States Department of Agriculture, Washington, DC, pp 568–572 McSorley R (1994) Susceptibility of common bedding plants to root-knot nematodes. Proc Fla State Hortic Soc 107:430–432 McSorley R, Fredeik JJ (1994) Response of some common annual bedding plants to three species of Meloidogyne. J Nematol 26(4S):773–777 Mellano HM, Munnecke DE, Endo RM (1970) Relationship of seedling age to development of Pythium ultimum on roots of Antirrhinum majus. Phytopathology 60:935–942 Moore WC (1947) British fungi. Trans Br Mycol Soc 31:86–91 Muller AS, Chupp C (1950) Cercospora in Guatemala. Ceiba 1:171–178 Murphy PA (1937) Irish Free State: a new outbreak of Peronospora antirrhini in the country. Int Bull Plant Prot 11:176 Nelson P (1962) Diseases. In: Langhans RW (ed) Snapdragons, a manual of the culture, insects and diseases and economics of snapdragons. New York State Flower Growers Association, Ithaca, pp 70–80 Nelson PE, Strider DL (1985) Snapdragons. In: Strider DL (ed) Diseases of floral crops, vol 2. Praeger Publishers, New York, pp 489–510 Peltier GL (1919) Snapdragon rust. Ill Agric Exp Sta Bull 221:535–548 Pirone PP (1978) Diseases and pests of ornamental plants. Wiley, New York Porter DM, Aycock R (1967) Snapdragon leaf spot caused by Cercospora antirrhini. NC Agric Exp Sta Tech Bull No. 179 Pscheidt JW, Ocamb CM (2016) Pacific northwest plant disease management handbook. Oregon State University. http://pnwhandbooks.org/plantdisease/ Raabe RD, Hurlimann JH, Sciaroni RH (1970) Powdery mildew control with benomyl for greenhouse-grown snapdragons. Calif Agric 24(1):8 Rogers MN (1992) Snapdragons. In: Larson RA (ed) Introduction to floriculture, 2nd edn. Academic, New York, pp 93–112 Simpson CJ, Elis Jones G, Taylor JD (1971) A seedling blight of Antirrhinum caused by Pseudomonas antirrhini. Plant Pathol 20:127–130 Stewart FC (1900) An anthracnose and stem rot of the cultivated snapdragon. NY (Geneva) Agric Exp Sta Bull 179:105–110 Takimoto S (1920) On a bacterial leaf-spot of Antirrhinum majus L. Bot Mag (Tokyo) 34:253–257 Tomioka K, Nishikawa J (2011) Anthracnose of snapdragon caused by Colletotrichum destructivum. J Gen Plant Pathol 77:60–63 UC IPM (2014) Pest management guidelines: floriculture and ornamental nurseries, UC ANR Publication 3392. http://www.ipm.ucdavis.edu/PMG/r280111211.html Valder PG (1963) New plant diseases. Plant Disease Survey, 33rd Report, Biol Brch NSW Dep Agric, p 35

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Wegulo SN, Vilchez M (2006a) Evaluation of fungicides for control of Botrytis blight of lisianthus, 2004. F&N Tests 61:OT030 Wegulo SN, Vilchez M (2006b) Evaluation of fungicides for control of downy mildew of snapdragon, 2003. F&N Tests 61:OT027 Yang HC, Leu LS (1981) The occurrence of snapdragon wilt disease in Taiwan. Plant Prot Bull 23:55–57 Yarwood CE (1947) Snapdragon downy mildew. Hilgardia 17:241–250

Diseases of Stock

26

Steven T. Koike

Abstract

The ornamental plant stock (Matthiola incana R. Br.) is a popular and versatile flowering species that can be grown as a garden or bedding plant as well as a field grown cut flower. Profitability of stock production depends on the management of a number of diseases. As part of the Brassicaceae, stock is subject to a number of pathogens that infect vegetable crucifers as well. Major pathogens include Sclerotinia, downy mildew, two wilt pathogens (Fusarium, Verticillium), Xanthomonas, and Turnip mosaic virus. Keywords

Fusarium • Sclerotinia • Verticillium • Xanthomonas

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Cottony Rot (Sclerotinia sclerotiorum) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Downy Mildew (Peronospora parasitica = P. matthiolae or Hyaloperonospora parasitica) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Fusarium Wilt (Fusarium oxysporum f. sp. mathioli) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Rhizoctonia Foot Rot (Rhizoctonia solani) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Verticillium Wilt (Verticillium dahliae and Verticillium zaregamsianum) . . . . . . . . . . . 2.6 Additional Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Bacterial and Phytoplasma Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Bacterial Blight (Xanthomonas campestris pv. incanae) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Blossom Blight (Pseudomonas syringae pv. maculicola) . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Viral Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Turnip Mosaic Virus (TuMV, Genus: Potyvirus) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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S.T. Koike (*) University of California Cooperative Extension, Monterey County, Salinas, CA, USA e-mail: [email protected] # Springer International Publishing AG 2018 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florists’ Crops Diseases, Handbook of Plant Disease Management, https://doi.org/10.1007/978-3-319-39670-5_26

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Introduction

Stock (Matthiola incana R. Br.) is also known as annual stock, column stock, garden stock, or tensweek stock. A closely related ornamental species is M. longipetala (Vent.) DC which is named night-scented stock or evening stock. Likely originating from the Mediterranean area, stock is a popular floral commodity that is strongly scented and is grown as a garden or bedding plant and also as a cutflower (Armitage and Laushman 2003). Depending on the species and region, stock can be grown as annuals, biennials, or perennials. As a member of the Brassicaceae, stock is susceptible to many of the diseases that affect commercial cole crops (broccoli, cabbage, cauliflower, etc.) vegetables (Koike et al. 2007).

2

Fungal and Fungus-Like Diseases

2.1

Cottony Rot (Sclerotinia sclerotiorum)

Geographic occurrence and impact. Cottony rot (also known as white mold and Sclerotinia crown rot) has been reported on stock in Australia, Greece, Korea, New Zealand, Scotland, South Africa, and the United States (California, Michigan, and Pennsylvania). The disease can occasionally be important on stock. Symptoms/signs. Symptoms vary depending on the type of fungal inoculum that comes in contact with stock. The soilborne survival structure, the sclerotium, can germinate directly and the resulting mycelium can infect stock crowns at the soil line. Crown infections first result in a gray to brown lesion on crown tissues. Lesions expand, crowns become completely rotted, and the plant will suddenly collapse (Fig. 1). If conditions are moist enough, the diseased crown will support the growth of white mycelium and large (5–10 mm/0.2–0.4 in. in length), black, irregularly shaped, hard sclerotia (Fig. 2). A second type of inoculum produced by S. sclerotiorum, the ascospore, is airborne, lands on aboveground stock tissues, and causes a blight on stems, leaves, or flowers. To infect the host, the ascospore must land on tissue that is damaged or senescent. Once germinated, the ascospore produces mycelium that will colonize the compromised tissue and also spreads to adjacent healthy tissue, resulting in a brown, soft decay. White mycelium and black sclerotia also develop from these foliar infections. Biology and epidemiology. Cottony rot is caused by Sclerotinia sclerotiorum. There is one report of another species, S. minor, causing a crown rot on stock in Australia. S. sclerotiorum survives and overwinters as sclerotia in soil. Sclerotia can germinate and form infective mycelium or alternatively will produce small, tan, mushroom-like structures called apothecia (Fig. 3). Apothecia emerge from the soil and release airborne ascospores that are carried by wind and infect susceptible stock leaves, stems, and flowers. When infected stock plants are plowed back into the soil,

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Fig. 1 Stock plants infected with Sclerotinia species can suddenly wilt and collapse (Steven Koike # 2017. All Rights Reserved.)

sclerotia that had formed on infected parts are returned to the soil and enable the pathogen to persist. S. sclerotiorum has an extremely broad host range, so sclerotia and ascospores produced from other crops, such as gazania, gaillardia, cauliflower, lettuce, pepper, and many others, can provide inoculum that can infect stock. Management • Cultural practices – Avoid planting stock in fields having a history of cottony rot. Avoid planting stock adjacent to vegetable or other flower crops that are highly susceptible to Sclerotinia and which exhibit symptoms of the disease. • Fungicides – Apply protectant fungicides to stock foliage prior to infection.

2.2

Downy Mildew (Peronospora parasitica = P. matthiolae or Hyaloperonospora parasitica)

Geographic occurrence and impact. This disease occurs worldwide on stock. Severe outbreaks of downy mildew can cause significant damage to the stock foliage and reduce flower quality and marketability. Symptoms/signs. The first symptoms are irregular, light green to slightly yellow, diffuse patches on the top surfaces of leaves. As disease develops, the yellow color intensifies and large portions of the leaf can be affected (Fig. 4). Irregular dark specks can be seen within the yellow patches. Diseased leaves can be twisted and, in

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Fig. 2 The hard, black sclerotia (arrows) of Sclerotinia sclerotiorum can form on the outside of stock stems (Steven Koike # 2017. All Rights Reserved.)

Fig. 3 Small, tan apothecia of Sclerotinia sclerotiorum will release spores that can infect stock foliage (Steven Koike # 2017. All Rights Reserved.)

advanced stages, the entire leaf can turn brown and die. White tufts of the pathogen are seen on the undersides of the lesions (Fig. 5). With a hand lens the tufts can be seen to consist of the branching, treelike structures that bear the downy mildew spores. If conditions are favorable, the sporulation can also occur on the top side of leaves. For some stock cultivars, extensive sporulation can develop well before any

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Fig. 4 Stock plants infected with downy mildew will first develop yellow patches on leaves (Steven Koike # 2017. All Rights Reserved.)

Fig. 5 The fine, white growth of downy mildew can be seen on the surfaces of stock leaves (Steven Koike # 2017. All Rights Reserved.)

symptoms are seen on the top of the leaves (Koike 2000). Downy mildew has been known to kill stock seedlings if conditions allow seedling cotyledons to be infected early and severely. Biology and epidemiology. Downy mildew is caused by the oomycete Peronospora parasitica. Researchers also refer to the pathogen as Peronospora matthiolae or Hyaloperonospora parasitica (Constantinescu and Fatehi 2002; Jafar 1963). Oomycete organisms are no longer considered true fungi but are more closely related to algae. This obligate pathogen forms mycelia and haustoria inside host tissues, and spore-bearing sporangiophores emerge through the leaf stomata. Some P. parasitica isolates and races form the sexual oospore stage, though this has not been observed on stock (Sherriff and Lucas 1989). There is considerable host specialization within the P. parasitica group of pathogens, and strains infecting stock apparently only infect this host (Sherriff and Lucas 1990). Cool (10 to 15  C/50 to 59  F), moist or high humidity conditions favor downy mildew sporulation. Spores,

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which are short-lived, are dispersed via winds and splashing water. Germination of sporangia and infection usually require the presence of free moisture on the leaf surface. Lesion expansion is most rapid at 20  C/68  F. Management • Cultural practices – Irrigating early in the day, to enhance drying of the foliage, may reduce disease severity. • Fungicides – Applying protectant foliar fungicides prior to infection is the best means of controlling downy mildew. Fungicides such as metalaxyl can be used as seed treatments for controlling early downy mildew on seedlings. • Resistance – Differences in susceptibility may exist among different stock cultivars but resistant cultivars do not appear to be available.

2.3

Fusarium Wilt (Fusarium oxysporum f. sp. mathioli)

Geographic occurrence and impact. Fusarium wilt has been reported on stock in Germany (Gerlach 1975), Japan (Saito et al. 2008), South Africa, the United Kingdom, and the United States (Arizona, California) (Baker 1948). The disease has caused considerable damage in the United Kingdom (O’Neill et al. 2003, 2005; O’Neill and Mason 2014, Fig. 6). Symptoms/signs. In some cases, if young seedlings are infected early, these plants will rapidly wilt and die. On larger plants, the lower leaves are affected first,

Fig. 6 Fusarium wilt can cause considerable damage to stock plantings (T.M. O’Neill # 2017. All Rights Reserved.)

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Fig. 7 In advanced cases Fusarium wilt will cause all the leaves to dry up (T.M. O’Neill # 2017. All Rights Reserved.)

developing a clearing of the veinal tissue, followed by a general yellowing and then wilting. The yellowing and wilting symptoms may occur on only one side of the plant. Plants will be stunted and wilt during the warmer times of the day. In advanced cases, the leaves can dry up and the plant can die (Fig. 7). Examination of the stem and taproot vascular tissues will reveal a brown discoloration, though the root is not decayed (Fig. 8). Infected stock seed plants may exhibit seed pods that appear flattened and tan in color. On stock, Fusarium wilt symptoms closely resemble those of Verticillium wilt. Biology and epidemiology. Like most Fusarium wilt pathogens, the stock pathogen (F. oxysporum f. sp. mathioli) is host specific to its original host and does not infect related species such as cabbage, kale, and radish. The report from Japan names the pathogen as F. oxysporum f. sp. conglutinans (Saito et al. 2008). This stock pathogen is typical of all F. oxysporum wilt isolates and forms four- to six-celled, fusiform, curved macroconidia, one- to two-celled oval- to kidney-shaped microconidia, and resilient chlamydospores. The pathogen can be seedborne and can survive via chlamydospores for long periods in the soil. The disease is most severe if the crop is grown during the warm summer and in regions where temperatures are high. The fungus is favored by temperatures around 25  C/77  F.

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Fig. 8 Fusarium wilt causes a brown discoloration in stem vascular tissue (T.M. O’Neill # 2017. All Rights Reserved.)

Management • Cultural practices – Avoid planting stock in fields known to be infested with the pathogen. Since the pathogen may survive in soil for 2 or more years, practice crop rotation with nonstock plants. Stock grown during the cooler winter or spring seasons and in cool coastal locations may escape the disease or experience less severe impacts from Fusarium wilt. • Soil disinfestation – Preplant soil treatments such as fumigation and steaming can help reduce soil inoculum.

2.4

Rhizoctonia Foot Rot (Rhizoctonia solani)

Geographic occurrence and impact. While this pathogen is broadly distributed throughout the world, Rhizoctonia foot rot of stock has been reported only from Australia, Greece, the United Kingdom, the United States (California, Florida), and Zimbabwe. The disease has also been called Rhizoctonia root rot or wirestem, and the pathogen also is implicated in damping-off disease of emerging stock seedlings. Symptoms/signs. The damping-off phase of the disease pertains only to young stock seedlings. For direct seeded crops, the fungus can attack seed or newly germinated seedlings and kill them prior to emergence (preemergence dampingoff), or can attack roots and lower stem (hypocotyl) tissues shortly after the plant has emerged above ground (postemergence damping-off). For recently emerged plants, the stems in contact with soil become water soaked and later brown in color; these stems are often girdled and the plants fall over and die.

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Fig. 9 Rhizoctonia solani infects the stock stem and causes the “wirestem” symptom (T.M. O’Neill # 2017. All Rights Reserved.)

The wirestem phase pertains to older seedlings and consists of browning and cracking of the stem epidermis that is in contact with soil (Dimock 1941, Fig. 9). These stem infections develop into lesions, with the outer tissue decaying away. When the outer stem layers deteriorate, only the fibrous inner xylem remains intact as a wiry strand, hence the name wirestem. Affected plants develop yellow foliage and later wilt. Seedling stems may break at the soil line, resulting in the plant falling over. Plants that survive will likely remain stunted and behind in development. Signs of the pathogen consist of coarse mycelium that can be observed, with a hand lens, in the lesion. The mycelium sometimes causes soil particles to adhere to and dangle from diseased stems. The pathogen is also known to cause a root rot on stock. Biology and epidemiology. Rhizoctonia foot rot or wirestem is caused by the soil inhabitant Rhizoctonia solani. In culture, R. solani forms coarse, brown, relatively thick hyphae that are characterized by right-angle branching. Hyphae are constricted at branch points, and a cross wall with a dolipore septum is formed right after each branch. Older cultures form small, brown, loosely aggregated clumps of mycelia that function as sclerotia. The fungus is multinucleate and has a sexual phase, Thanatephorus cucumeris (Sneh et al. 1991), though this stage has not been reported on stock. This species is extremely diverse and can be divided into anastomosis groups

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(AG) based on the process of hyphal fusion between compatible isolates. AG-2-1 is associated with Rhizoctonia diseases on Brassicaceae hosts (Keijer et al. 1997). While AG-2-1 isolates are pathogenic on plants in this family, these isolates may infect plants outside of the Brassicaceae. Likewise, non-AG-2-1 isolates may infect Brassicaceae hosts. Therefore, these AG pathogens are not absolutely restricted to certain plant groups. R. solani survives saprophytically in soil as mycelia or sclerotia; these fungal structures are the inocula for infecting stock seedlings and older plants. The pathogen is favored by warm soil conditions (25 to 30  C/77 to 86  F) but is capable of causing problems at much lower temperatures as well. Management • Cultural practices – When placing transplants in the field, avoid planting them too deeply in the soil as the hypocotyl stem tissue is the most susceptible part of the plant. Practice crop rotation so that nonhosts are included in the rotation (Henis et al. 1978). Do not plant stock in fields having undecomposed crop residues, since R. solani may be actively colonizing such residues. • Fungicides – When direct seeding in the field, use seed that has been treated with a fungicide. If available, direct-spray fungicides to the base of young stock plants in the field. • Sanitation – Practice thorough sanitation at nurseries to prevent contamination by R. solani. Clean and sanitize transplant trays and benches. Ensure that rooting media are not contaminated by infested soil or diseased plant residues.

2.5

Verticillium Wilt (Verticillium dahliae and Verticillium zaregamsianum)

Geographic occurrence and impact. Verticillium wilt has been reported on stock in Japan, New Zealand, and the United States (California, New York). Symptoms/signs. Lower leaves are affected first, becoming yellow and then wilted. In some cases, the leaf interveinal tissue turns yellow first and the veins remain green longer. Plants will be stunted and wilt during the warmer times of the day. The yellowing and wilting of leaves advances up the stem until the stock plant becomes completely affected and can die (Raabe and Wilhelm 1958). Examination of the stem and taproot vascular tissues can reveal a light tan to brown discoloration (Fig. 10). On stock, Verticillium wilt symptoms closely resemble those of Fusarium wilt. Biology and epidemiology. Verticillium wilt of stock appears to be caused by two Verticillium species (Inderbitzin et al. 2011). Most cases of stock Verticillium wilt are caused by V. dahliae. However, Verticillium wilt of stock in Japan is caused by V. zaregamsianum. This etiology is supported by inoculation experiments. For example, stock is susceptible to a tomato isolate of Verticillium which is the V. dahliae species; however, even though stock is a member of the Brassicaceae, stock is not susceptible to a V. longisporum isolate that infects Brussels sprouts and

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Fig. 10 Internal stem tissues infected with Verticillium will turn light tan to brown (Steven Koike # 2017. All Rights Reserved.)

other brassica crops. In culture these stock pathogens form the typical Verticillium structures consisting of hyaline, verticillate conidiophores bearing three to four phialides at each node. Conidia formed from phialides are hyaline, oval, and single-celled. Older cultures form darkly pigmented, multicelled microsclerotia. The pathogen can survive as resilient microsclerotia for long periods in the soil. Management • Cultural practices – Avoid planting stock in fields known to be infested with the pathogen. Since the pathogen may survive in soil for many years, practice crop rotation with nonhost plants. • Soil disinfestation – Preplant soil treatments such as fumigation and steaming can help reduce soil inoculum.

2.6

Additional Fungal and Fungus-Like Diseases

Alternaria leaf spot (Alternaria japonica) (Davis et al. 1949; Simmons 2007). Club root (Plasmodiophora brassicae) (Samson and Walker 1982). Damping-off (Pythium species) (Fig. 11). Gray mold (Botrytis cinerea) (Baker et al. 1954). Phytophthora root rot (Phytophthora species) (Fig. 12). Powdery mildew (Erysiphe cruciferarum) (Braun and Cook 2012). White rust (Albugo candida) (Choi et al. 2007; Garcia-Blazquez et al. 2006).

3

Bacterial and Phytoplasma Diseases

3.1

Bacterial Blight (Xanthomonas campestris pv. incanae)

Geographic occurrence and impact. Bacterial blight is a significant disease of stock that can cause significant losses in yield and quality (Fig. 13). The disease is

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Fig. 11 Pythium infects stock seedlings early and results stunted growth and/or plant death (T.M. O’Neill # 2017. All Rights Reserved.)

Fig. 12 Phytophthora infections result in rotted roots and a reduced root system (T.M. O’Neill # 2017. All Rights Reserved.)

known to occur wherever stock is grown because it is seedborne (Minardi et al. 1988; Rahimian and Okhovatian 1989). Symptoms/signs. Young seedlings can develop the disease and exhibit yellowing of the stem and leaves, followed by complete wilting of the foliage and plant death. On older plants, initial symptoms consist of yellowing and wilting of the lower leaves. Where lower leaves are attached, the stem develops water-soaked lesions that later turn dark brown to black and are sunken (Fig. 14). Lesions enlarge and can girdle the stem, resulting in complete wilting of the plant. These lesions also weaken the stem and are the points where the stem cracks and causes the plant to fall over (Fig. 15). Examination of the vascular tissue shows a darkened discoloration.

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Fig. 13 Bacterial blight can cause significant losses in stock plantings (Steven Koike # 2017. All Rights Reserved.)

Biology and epidemiology. The pathogen, Xanthomonas campestris pv. incanae, is apparently host specific to stock and did not infect cabbage, cauliflower, or kale in inoculation experiments. In culture the pathogen is typical of most X. campestris pathovars and is characterized by slow growing, mucoid, yellow colonies. The pathogen is seedborne (Kendrick 1938) and can also persist in the soil in association with diseased stock plant residues. Disease is initiated by planting infested seed or by planting stock into soil containing plant residues of diseased stock. Once disease has developed, the bacteria are splashed to other plants via rain and overhead sprinkler irrigation. Cool, wet weather favors infection and disease development. Management • Cultural practices – Stock seed can be treated with hot water (50 to 55  C/122 to 131  F for 10 min) to disinfest seed; however, such treatments can result in loss of seed germination and need to be implemented carefully. Use drip or furrow irrigation instead of overhead sprinklers. Seed crops should be produced in regions where rain does not occur during seed production. Avoid planting stock in back-to-back plantings; a crop rotation of nonstock crops for 2–3 years likely results in eradication of the pathogen from soil. For stock grown in nurseries and containers, remove and discard symptomatic plants. • Resistance – Resistant M. incana cultivars are not yet available, though resistance has been found in M. aspera, M. longipetala, and M. tricuspidata (Ecker et al. 1995).

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Fig. 14 Stock plants infected with bacterial blight will first develop dark lesions on stems (Steven Koike # 2017. All Rights Reserved.)

Fig. 15 Bacterial blight infections weaken the stems of stock, causing them to crack and break (Steven Koike # 2017. All Rights Reserved.)

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3.2

Diseases of Stock

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Blossom Blight (Pseudomonas syringae pv. maculicola)

Geographic occurrence and impact. Blossom blight has been reported only from Australia (Cother and Noble 2009). However, the blossom blight pathogen is widely distributed worldwide on cole crop and other crucifer vegetables (Cintas et al. 2001; Koike et al. 2007; Peters et al. 2004; Shackleton 1996). Symptoms/signs. Disease first develops on flowers located on the lower portion of the spikes. Infected petals turn brown and later collapse (Fig. 16). Collapsed petals adhere to adjacent flowers. As disease develops, infections progress up the spike and in severe cases affect all the flowers (Cother and Noble 2009). Lesions can also develop where the flower base is attached to the spike. Foliar symptoms consist of small, angular, brown leaf spots that can be seen from both upper and lower leaf surfaces (Cother and Noble 2009, Fig. 17). This symptom has been referred to as Fig. 16 Brown, collapsed petals caused by Pseudomonas syringae pv. maculicola (E.J. Cother # 2017. All Rights Reserved.)

Fig. 17 The blossom blight pathogen can also cause small, brown leaf spots that may be surrounded by yellow borders (E.J. Cother # 2017. All Rights Reserved.)

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Fig. 18 A brown discoloration in stem tissue can be caused by Pseudomonas syringae pv. maculicola (E.J. Cother # 2017. All Rights Reserved.)

Fig. 19 Stock seedpods infected with Pseudomonas syringae pv. maculicola can develop dark brown lesions (E.J. Cother # 2017. All Rights Reserved.)

“pepper leaf spot” on vegetable crucifers (Koike et al. 2007). Leaf spots may be surrounded by yellow borders. A slight discoloration can be observed in stem vascular tissue (Fig. 18). For stock plants producing seed, seed pods may develop lesions (Fig. 19). Biology and epidemiology. The cause of blossom blight is Pseudomonas syringae pv. maculicola (Cother and Noble 2009). The pathogen is an aerobic, Gram-negative bacterium that on standard microbiological media forms colonies that are cream to light yellow in color and smooth (Cintas et al. 2001). When cultured on King’s medium B, strains that infect stock do not produce the diffusible pigment that fluoresces blue under ultraviolet light; this stock pathogen, therefore, is a nonfluorescing type of P. syringae. Consistent with P. syringae pv. maculicola isolates from crucifer vegetable crops, this stock pathogen can infect cauliflower and tomato (Cother and Noble 2009; Wiebe and Campbell 1993). The pathogen is seedborne in

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vegetable hosts and appears also to be seedborne in stock. More details on disease epidemiology are not available. Management. Disease control options are based on standard integrated disease control principles since the exact details are not available on how the disease develops. Resistant M. incana cultivars are not available. • Cultural practices – Stock seed can presumably be infested with the pathogen and can be treated with hot water, though such treatments can result in loss of seed germination and need to be implemented carefully. Use drip or furrow irrigation instead of overhead sprinklers. Seed crops should be produced in regions where rain does not occur during seed production. Avoid planting stock in back-to-back plantings. This pathogen likely does not persist for long periods in soil; however, a crop rotation using nonbrassica and nonstock crops for 2 years is advisable. For stock grown in nurseries and containers, remove and discard symptomatic plants.

4

Viral Diseases

4.1

Turnip Mosaic Virus (TuMV, Genus: Potyvirus)

Geographic occurrence and impact. Stock is reported to be susceptible to a dozen or more viruses wherever the flower is grown. Some of the more familiar virus pathogens are Alfalfa Mosaic Virus, Beet Curly Top Virus, Cauliflower Mosaic Virus, Cucumber Mosaic Virus, Tomato Spotted Wilt Virus, and Turnip Mosaic Virus. It is not rare to have a stock plant infected with more than one virus (Alioto et al. 1994; Yoon et al. 1998). It appears that Turnip Mosaic Virus (TuMV) is perhaps the most commonly reported virus pathogen of stock and occurs worldwide (Bahar et al. 1985; Rosciglione and Cannizzaro 1983). Symptoms/signs. As with most viruses, symptoms caused by TuMV vary depending on the strain of the virus, stock cultivar, age of plant when infected, and environmental conditions. Leaves will exhibit a range of symptoms consisting of mottling, mosaic, formation of dark green patches or “green islands,” and clearing of veins. Leaves may also be curled, twisted, or otherwise distorted. The dark-colored (pink, red, purple) flowers that contain anthocyanin will show breaking, in which the petal coloration is broken up by streaks and sectors of nonpigmented tissue (Severin and Tompkins 1948, 1950; Tompkins 1939, Fig. 20). Cultivars with white and yellow colored flowers do not show flower breaking. Infected plants may be distorted. Biology and epidemiology. The virus survives between stock crops in living plants such as cruciferous weeds and volunteer stock. Aphids feed on these reservoir hosts, move to stock plants, and inject the virus during feeding. The virus is borne on the stylet of aphids and is transmitted in a nonpropagative (nonpersistent) manner.

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Fig. 20 If infected with Turnip mosaic virus, darkcolored stock flowers will “break” and develop white streaks and sectors (Steven Koike # 2017. All Rights Reserved.)

Management • Cultural practices – Old and infected stock plantings should be disked and buried in the soil. Remove cruciferous weeds and volunteer stock from around fields. • Vector management – Control the aphids by using insecticides and other IPM methods. • Resistance – Some stock lines and cultivars are resistant to TuMV (Johnson and Barnhart 1956; San Juan and Pound 1963).

References Alioto D, Stavolone L, Aloj B (1994) Serious alterations induced by Cauliflower mosaic virus (CaMV) and Turnip mosaic virus (TuMV) on Matthiola incana. Inf Fitopat 44:43–46 Armitage AM, Laushman JM (2003) Specialty cut flowers, 2nd edn. Timber Press, Portland, 586 pp Bahar M, Danesh D, Dehghan M (1985) Turnip mosaic virus in stock plant. Iranian J Plant Pathol 21(11–12):33–39 Baker KF (1948) Fusarium wilt of garden stock (Matthiola incana). Phytopathology 38:399–403 Baker KF, Matkin OA, Davis LH (1954) Interaction of salinity injury, leaf age, fungicide application, climate, and Botrytis cinerea in a disease complex of column stock. Phytopathology 44:39–42 Braun U, Cook RTA (2012) Taxonomic manual of the Erysiphales (Powdery mildews). CBS-KNAW Fungal Biodiversity Centre, Utrecht, 707 pp Choi Y-J, Shin HD, Hong S-B, Thines M (2007) Morphological and molecular discrimination among Albugo candida materials infecting Capsella bursa-pastoris world-wide. Fung Divers 27:11–34 Cintas NA, Bull CT, Koike ST, Bouzar H (2001) A new bacterial leaf spot disease of broccolini, caused by Pseudomonas syringae pv. maculicola, in California. Plant Dis 85:1207 Constantinescu O, Fatehi J (2002) Peronospora-like fungi (Chromista, Peronosporales) parasitic on Brassicaceae and related hosts. Nova Hedwigia 74:291–338 Cother EJ, Noble DH (2009) Identification of blossom blight in stock (Matthiola incana) caused by Pseudomonas syringae pv. maculicola. Australas Plant Pathol 38:242–246 Davis LH, Sciaroni RH, Pritchard F (1949) Alternaria leafspot of garden stock in California. Plant Dis Rep 33:432–433

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Dimock AW (1941) The Rhizoctonia foot-rot of annual stocks (Matthiola incana). Phytopathology 31:87–91 Ecker R, Zutra D, Barzilay A, Osherenko E, Rav-David D (1995) Sources of resistance to bacterial blight of stock (Matthiola incana R. Br.). Genet Res Crop Evol 42:371–372 Garcia-Blazquez G, Constantinescu O, Telleria MT, Martin MP (2006) Preliminary checklist of Albuginales and Peronosporales (Chromista) reported from the Iberian Peninsula and Balearic Islands. Mycotaxon 98:185–188 Gerlach W (1975) The first case of Fusarium wilt on garden stock (under glass) in Germany. Nachrichtenblatt des Deutschen Pflanzenschutzdienstes 27:17–20 Henis Y, Ghaffar A, Baker R (1978) Integrated control of Rhizoctonia solani damping-off of radish: effect of successive plantings, PCNB, and Trichoderma harzianum on pathogen and disease. Phytopathology 68:900–907 Inderbitzin P, Bostock RM, Davis RM, Usami T, Platt HW, Subbarao KV (2011) Phylogenetics and taxonomy of the fungal vascular wilt pathogen Verticillium, with the descriptions of five new species. PLoS One 6:1–22 Jafar H (1963) Studies on downy mildew (Peronospora matthiolae (Roume-guere) Gaumann) on stocks (Matthiola incana R.Br.). N Z J Agric Res 6:70–82 Johnson BL, Barnhart D (1956) Transfer of mosaic resistance to commercial varieties of Matthiola incana. Proc Am Soc Hort Sci 67:522–533 Keijer J, Korsman MG, Dullemans AM, Houterman PM, de Bree J, van Silfhout CH (1997) In vitro analysis of host plant specificity in Rhizoctonia solani. Plant Pathol 46:659–669 Kendrick JB (1938) A seed-borne bacterial disease of garden stocks, Matthiola incana. Phytopathology 28:12 Koike ST (2000) Downy mildew of stock, caused by Peronospora parasitica, in California. Plant Dis 84:103 Koike ST, Gladders P, Paulus AO (2007) Vegetable diseases: a color Handbook. Manson Publishing Ltd, London, 448 pp Minardi P, Mazzucchi U, Parrini C (1988) Epidemics of bacterial blight of stock (Matthiola incana R. Br.) caused by Xanthomonas campestris pv. incanae in Tuscany. Inf Fitopat 38:43–46 O’Neill TM, Mason L (2014) Contrasting effects on Fusarium wilt of column stock following use of bark as a soil amendment. Acta Hortic 1044:139–143 O’Neill TM, Shepherd A, Inman AJ, Lane CR (2003) Wilt of stock (Matthiola incana) caused by Fusarium oxysporum in the United Kingdom. New Dis Reports 8:10 O’Neill TM, Green KR, Ratcliffe T (2005) Evaluation of soil steaming and a formaldehyde drench for control of Fusarium wilt in column stock. Acta Hortic 698:129–133 Peters BJ, Asha GJ, Cother EJ, Hailstones DL, Nobleb DH, Urwin NAR (2004) Pseudomonas syringae pv. maculicola in Australia: pathogenic, phenotypic and genetic diversity. Plant Pathol 53:73–79 Raabe RD, Wilhelm S (1958) Verticillium wilt of garden stock (Matthiola incana). Phytopathology 48:610–613 Rahimian H, Okhovatian H (1989) Bacterial blight of stock in Mazandaran. Iranian J Plant Pathol 25(11–12):29–37 Rosciglione B, Cannizzaro G (1983) Matthiola incana R. Br., natural host of Turnip mosaic virus in Sicily. Tecnica Agricola 35:251–257 Saito K, Domon K, Honma T, Kawasaki T, Ogata M, Hori Y (2008) Soil reduction disinfection of stock wilt caused by Fusarium oxysporum f. sp. conglutinans race 3. Annu Rep Soc Plant Protect N Jpn 59:71–73 Sampson PJ, Walker J (1982) An Annotated List of Plant Diseases in Tasmania. Department of Agriculture, Tasmania. 121 pp San Juan MO, Pound GS (1963) Resistance in Matthiola incana to the Turnip mosaic virus. Phytopathology 53:1276–1279

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Severin HHP, Tompkins CM (1948) Aphid transmission of mild mosaic virus of annual stock. Hilgardia 18:539–547 Severin HHP, Tompkins CM (1950) Aphid transmission of severe mosaic virus of annual stock. Hilgardia 20:93–108 Shackleton DA (1996) A bacterial leaf spot of cauliflower in New Zealand caused by Pseudomonas syringae pv. maculicola. N Z J Sci 9:872–877 Sherriff C, Lucas JA (1989) 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 Simmons EG (2007) Alternaria: an identification manual. Centraalbureau voor Schimmelcultures, Utrecht, 775 pp Sneh B, Burpee L, Ogoshi A (1991) Identification of Rhizoctonia Species. APS Press, St. Paul, 133 pp Tompkins CM (1939) Two mosaic diseases of annual stock. J Agric Res 58:63–77 Wiebe WL, Campbell RN (1993) Characterization of Pseudomonas syringae pv. maculicola and comparison with Pseudomonas syringae pv. tomato. Plant Dis 77:414–419 Yoon J-Y, Choi H-S, Ryu H-Y, Harm Y-I, Choi J-K (1998) Colour breaking syndrome of Matthiola incana caused by double infection of Cucumber mosaic virus and Turnip mosaic virus. Korean J Plant Pathol 14:220–222

Diseases of Sunflower

27

Thomas J. Gulya, Febina Mathew, Robert Harveson, Samuel Markell, and Charles Block

Abstract

Sunflower (Helianthus annuus L.) has become a popular cut flower in the United States and globally. Easy to grow, bright and cheerful large flowers, with ray flowers from white to yellow to orange to magenta, it is often a centerpiece in mixed bouquets. Sunflower and the rest of the Helianthus genus are native to North America, and, as such, there is a native population of disease organisms and insect pests that can be a production challenge. Fortunately, ornamental sunflower and oilseed sunflower are the same species, and thus genetic advances made with oilseed germplasm can be readily transferred to ornamentals, if the need arises. The major diseases posing threats to ornamental sunflower are the same as those

Thomas J. Gulya was retired. T.J. Gulya (*) USDA-Agricultural Research Service, Sunflower and Plant Biology Research Unit, Fargo, ND, USA e-mail: [email protected] F. Mathew Department of Agronomy, Horticulture, and Plant Science, South Dakota State University, Brookings, SD, USA R. Harveson Department of Plant Pathology, Panhandle Research and Extension Center, University of Nebraska, Scottsbluff, NE, USA S. Markell Department of Plant Pathology, North Dakota State University, Fargo, ND, USA C. Block Seed Science Center, Iowa State University, Ames, IA, USA # Springer International Publishing AG 2018 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florists’ Crops Diseases, Handbook of Plant Disease Management, https://doi.org/10.1007/978-3-319-39670-5_27

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threatening oilseed sunflower hybrids, namely downy mildew, rust, Phomopsis stem canker, Sclerotinia wilt, Verticillium wilt, and charcoal rot. However, greenhouse production presents a different environment, and thus, there can be diseases affecting cut sunflowers that are seldom seen under field production. General management strategies for diseases of florists’ crops may be found in the introductory chapters on integrated disease management. Keywords

Alternaria • Botrytis • Fusarium • Phomopsis • Plasmopara halstedii • Puccinia helianthi • Phoma • Sclerotinia sclerotiorum • Verticillium • White rust

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Alternaria Leaf Spot and Blight (Alternaria spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Botrytis Blight or Gray Mold (Botrytis cinerea Persoon) . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Charcoal Rot (Macrophomina phaseolina (Tassi) Goidànich) . . . . . . . . . . . . . . . . . . . . . 2.4 Damping-off (Pythium spp., Phytophthora spp., and Rhizoctonia solani Kühn) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Downy Mildew (Plasmopara halstedii (Farlow) Berlese and de Toni) . . . . . . . . . . . . 2.6 Fusarium Stalk Rot and Wilt (Fusarium spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Leaf Smut (Entyloma spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.8 Phoma Black Stem (Phoma macdonaldii Boerema) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.9 Phomopsis Stem Canker (Diaporthe helianthi and Other Diaporthe spp.) . . . . . . . . 2.10 Powdery Mildews (Erysiphe and Other Genera) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.11 Rhizopus Head Rot (Rhizopus spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.12 Rust – Puccinia helianthi Schwein. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.13 Rusts – Other Minor Ones (Puccinia spp., Coleosporium helianthi Arthur) . . . . . . 2.14 Sclerotinia Stalk Rot and Head Rot (Sclerotinia sclerotiorum (Libert) de Bary and Sclerotinia minor Jagger) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.15 Septoria Leaf Spot and Blight (Septoria spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.16 Sclerotium Blight (Sclerotium rolfsii Saccardo) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.17 Verticillium Leaf Mottle/Wilt (Verticillium dahliae Klebahn) . . . . . . . . . . . . . . . . . . . . . . 2.18 White Rust (Albugo tragopogonis (DeCandolle) Gray = Pustula helianthicola Rost and Thines) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Bacterial and Phytoplasma Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Apical Chlorosis and Bacterial Leaf Spot (Pseudomonas syringae pv. tagetis and Pseudomonas syringae pv. helianthi) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Bacterial Stalk Rot/Head Rot (Erwinia = Pectobacterium spp.) . . . . . . . . . . . . . . . . . . . . 3.3 Aster Yellows (Phytoplasma spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Viral Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Sunflower Mosaic Virus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Nematodes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 Root-Knot Nematodes (Meloidogyne Species) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2 Root-Lesion Nematodes (Pratylenchus Species) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3 Other Nematodes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Introduction

Sunflower is a unique crop in that it is grown for ornamental use, as an oilseed crop, and for human consumption. All three uses dictate a different germplasm base, but all are Helianthus annuus. Additionally, wild H. annuus is native to the United States, and it occurs in almost all of the 48 contiguous states. Another unique feature of sunflower is that it is one of the few crops whose progenitors are native to the United States and was domesticated by Native Americans (Heiser 2008). There are 53 wild Helianthus species, of which all but one are found in the United States (Heiser et al. 1969; Schilling 2006). Among the 53 Helianthus species, 14 are annual species and 39 are perennial species. Many of these species are adapted to unique environmental niches and thus potential sources of genes, which can be transferred to cultivated sunflower through conventional breeding methods. The popularity of ornamental sunflowers is relatively recent, and this has spurred university research in many areas of the country, addressing the challenges of many different environments. Oilseed sunflower, in comparison, has been a large and essential global oilseed crop for many decades, and there are numerous sunflower disease guides in many languages besides English (Chattopadhyay et al. 2015; Gulya et al. 1997) including French (CETIOM 2015), Portuguese (Leite 1997), Serbo-Croatian (Maric et al. 1987), Spanish (Alonso et al. 1988), and Russian (Artokhin and Ignatova 2013), whose information is just as relevant to cut flower production as for oilseed sunflower. In addition, the American Phytopathology Society has recently issued the Compendium of Sunflower Diseases (Harveson et al. 2016), which will have similar information to this chapter, but with extensive images which should aid with identification. In the United States, extension agencies of several universities have addressed the need for information on the production of cut and potted sunflowers, but few have detailed information on disease management (Garfinkel and Panter 2015; Gill et al. 2003; Schoellhorn et al. 2003; Whipker et al. 1998). The vast majority of public sunflower research in the United States is centered in Fargo, North Dakota, where both North Dakota State University and the USDA-ARS Sunflower and Plant Biology Research Unit are located. North Dakota State University and the USDA-ARS in Fargo have excellent research information available online, at www.ag.ndsu.ext/crops/sun flower and www.ars.usda.gov/main/site_main.htm?modecode=30-60-05-15, respectively. Lastly, there are two other organizations that foster sunflower research, namely the National Sunflower Association (NSA) and the International Sunflower Association (ISA), whose websites are www.sunflowernsa.com and www.isasunflower.org, respectively. The NSA website is publically accessible and contains archives of magazine articles, papers and posters presented at over 30 past “research forums,” and a photograph gallery, one part of which deals with diseases and deformities.

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2

Fungal and Fungus-like Diseases

2.1

Alternaria Leaf Spot and Blight (Alternaria spp.)

Geographic occurrence and impact. Alternaria leaf blight affects sunflower throughout the world and is a major defoliating pathogen in warm, humid climates. The fungus may also cause linear spots on the stems and water-soaked, sunken lesions on the back of sunflower head. Symptoms/signs. Many Alternaria species can cause leaf spots on sunflower, but symptoms are similar for all, making field identification impractical. The primary symptom is dark brown lesions on leaves, and also on stems, petioles, and bracts. Initially the leaf spots are small, dark, and angular (Fig. 1), but with time, they coalesce into large, necrotic areas resulting in defoliation. Defoliation starts with the lower leaves where the microclimate is most favorable. Stem lesions typically are narrow (1–3 mm) black streaks up to 3 cm long. Biology and epidemiology. Although Alternaria helianthi (Hansford) Tubaki and Nishihara, now reclassified as Alternariaster helianthi (Hansford) Simmons, is the major causal agent, there are eight other species reported on sunflower, including A. zinniae Pape, A. helianthicola Rao and Rajagopalan, A. helianthiinficiens Simmons, Walcz and Roberts, A. leucanthemi Nelen (syn. Teretispora leucanthemi (Nelen)

Fig. 1 (a) Alternaria leaf blight lesions. (b) Alternaria stem lesions (T.J. Gulya. USDA.)

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Simmons), A. protenta Simmons, A. roseogrisea Roberts, A. tenuissima (Nees) Wiltshire, and A. alternata (Fries) Keissler. Most of these Alternaria spp. are specific for sunflower (except the opportunistic saprophyte A. alternata). However, A. zinniae has a broad host range that includes several Asteraceae genera, including weeds (Bidens, Cardus, Eupatorium, and Xanthium) and ornamentals (Aster, Calendula, Chrysanthemum, Dahlia, Tagetes, Tithonia, and Zinnia). Because symptoms caused by the Alternaria pathogens look very similar, field identification based on symptoms is difficult and may be inaccurate. Conidial morphology (Simmons 2008) combined with genetic analysis is the only sure means to delineate species. As A. helianthi is the primary causal agent and most widespread, this section addresses just this species (Allen et al. 1983a, b, c). Alternaria helianthi overwinters on infected plant residue, but wild or volunteer sunflowers may also serve as reservoirs. All species including A. helianthi may also be seedborne. The conidia are windborne and spread by splashing water onto the lower leaves. They germinate best at temperatures >26  C and require a minimum of 4 h of leaf wetness for sporulation. Disease progress is also heavily dependent on the duration of leaf wetness following initial infection, as the generation of new spores can occur within 2 days. Young seedlings are more susceptible than older plants, but senescing lower leaves on mature plants frequently are defoliated by Alternaria spp. Management • Cultural practices – including removal of wild and volunteer sunflowers, removal or incorporation in the soil of previous sunflower residues, and minimizing extended leaf wetness will all reduce disease potential. • Fungicides – Seed treatments (captan, thiram, mancozeb) may offer some control (Jeffrey et al. 1985), but most growers rely upon multiple applications of fungicides containing chlorothalonil, iprodione, procymidone, or vinclozolin. More recent fungicide tests with ornamental sunflowers demonstrated that seed treatments with difenoconazole, prochloraz, pyrifenox or triadimenol were effective (Wu and Wu 2003). Consult government fungicide guides for specific products and rates. • Resistance – Disease resistance has been found in oilseed sunflower and some attempts have been made to incorporate that into hybrids, but there have been no reports of resistance in ornamental cultivars.

2.2

Botrytis Blight or Gray Mold (Botrytis cinerea Persoon)

Geographic occurrence and impact. The fungus causing gray mold, Botrytis cinerea (syn. Botryotinia fuckeliana (de Bary) Whetzel), is found worldwide and impacts many floral, fruit, and vegetable crops. On sunflower, the pathogen causes mostly a head rot. However, it can also cause spots on leaves and petals and is most often observed in areas with cool summers accompanied by frequent rains. It is rarely seen outdoors in most areas of the United States but could be a problem in greenhouses where sunflower leaves remains wet for extended periods of time.

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Fig. 2 Botrytis head rot (T.J. Gulya. USDA.)

Symptoms/signs. Gray mold first appears as sunken brown spots on the back surface of the head. During periods of high humidity or rainfall, these lesions will be covered with conidia, giving the infected tissue a gray color (Fig. 2). The gray color is an easy, macroscopic way to distinguish Botrytis head rot from other head rots. If heads are infected early and wet conditions persist, Botrytis may form small, black sclerotia within the sunflower head. Botrytis has also been observed to cause petiole infections in Egypt, India, and Pakistan, which can progress to the stem, resulting in lesions up to 10 cm long. Biology and epidemiology. Botrytis cinerea is a ubiquitous fungus and a facultative pathogen with a wide host range encompassing most ornamentals, soft fruit, and monocots. B. cinerea overwinters in the soil as sclerotia or as mycelium on infected plant debris and is also seedborne (Coley-Smith et al. 1980; Elad et al. 2007). Sclerotia germinate to produce conidia, which can colonize dead organic matter as well as living host tissue. Conidia are windblown and water splashed and require a wound or senescent tissue to gain entry into a plant. Optimal conditions for infection are 15–25  C and 90% relative humidity. Under optimal conditions, Botrytis may produce a new crop of conidia every 5–7 days. Botrytis produces copious amounts of spores, which develop rapidly in decaying vegetation and on senescing flowers. Because of the wide host range, the inocula for sunflower infection may come from many other floral crops or weeds. Management. For control of Botrytis on greenhouse or field grown ornamental sunflowers, fungicide applications coupled with cultural practices are useful. • Cultural practices – Since spores readily form on infected plant tissue, elimination of infected plants is essential. Avoidance of overhead watering, especially prior to bloom, is important, or at least irrigate early in the day to promote drying. Wider spacing between plants will minimize the RH within the canopy, but this may not be practical under the intensive plant populations as seen in commercial

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field operations. The fungus can develop at low temperatures, so cut flowers with no apparent infection may develop gray mold even under refrigeration. As free water is necessary for infection, minimizing conditions that lead to condensation in cold storage is critical (Gibson et al. 2014). • Resistance – Oilseed germplasm with resistance to Botrytis has been identified, but to date these genes have not been incorporated into either oilseed hybrids or ornamental germplasm (Kanyion and Friedt 1993).

2.3

Charcoal Rot (Macrophomina phaseolina (Tassi) Goidànich)

Geographic occurrence and impact. Charcoal rot is a serious root-infecting disease that is found worldwide but is most severe in hot, dry climates. It is both soil- and seedborne (Raut 1983). The fungus has little effect on plants until they become stressed; the stress may be due to heat, lack of soil moisture, or seed filling. The disease may kill plants outright, but it also results in reduced head size. Symptoms/signs. Damping-off may occur if M. phaseolina-infected seeds germinate in soil at or near 35  C. Usually, symptoms appear on older plants after flowering, with a silvery-gray lesion at the base of the plant and eventually encircling the stem. Upon splitting open the stem longitudinally, one will notice pith disintegration, and often a characteristic compression of the pith into layers. Examination with a 5 to 10X hand lens will reveal tiny microsclerotia, which appear as pepper grains scattered within the pith and on the inside of the rind. Severe infections, especially on stressed plants, will cause premature death (Fig. 3a, b). Biology and epidemiology. Charcoal rot is incited by Macrophomina phaseolina. This fungus has had 17 binomials as synonyms, of which Rhizoctonia bataticola and

Fig. 3 (a) Charcoal rot, showing pith compression and microsclerotia. (b) Charcoal rot on sunflower, showing pith with microsclerotia (T.J. Gulya. USDA.)

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Sclerotium bataticola are most commonly found in older literature. The fungus is characterized by producing black microsclerotia, 60–200 μm in diameter. Occasionally an isolate will be found which produces pycnidia (and pycnidiospores) in culture, but this is very rare in infected sunflower tissue. The fungus has a wide host range, but there is no evidence of isolates having host specificity. Macrophomina phaseolina has an extremely broad host range, encompassing 500 species of both monocots and dicots (Dhingra and Sinclair 1978). Thus, nearly all ornamentals are hosts, as well as field crops, vegetables, and weeds. The fungus does not produce any conidia, but persists by forming small, irregular, black microsclerotia (60–200 μm in diameter) which are barely visible with the unaided eye. Microsclerotia in infected plant residue are released into soil as the host tissues decompose, and they can persist in the soil for at least 3 years. The fungus is unusual in its affinity or tolerance of high temperatures and grows best at soil temperatures >35  C. Other conditions which favor disease development include low soil moisture, light sandy soils, as well as stress of the host plants due to herbicides or nematodes (Singh et al. 2012). Charcoal rot is frequently associated with Fusarium stalk rot. Additionally, the sunflower stem weevil has been shown to vector Macrophomina phaseolina in sunflowers (Yang and Owen 1982). Management • Cultural practices – Minimizing plant stress by lowering plant densities, and providing adequate soil moisture, will reduce the onset of the disease. However, since sunflowers grown for cut flowers are harvested in the late bud stage, charcoal rot should seldom be seen. Crop rotations will have little effect on the fungus due to its wide host range and the longevity of the microsclerotia. • Fungicides – Seed treatments with fungicides will minimize the introduction of M. phaseolina (Raut 1983) and protect against early infection.

2.4

Damping-off (Pythium spp., Phytophthora spp., and Rhizoctonia solani Kühn)

Geographic occurrence and impact. Damping-off of oilseed sunflower is rarely noted in the literature, but is more likely to be observed with ornamental sunflowers due to their intensive management practices. Damping-off, as the name implies, leads to seedling death, whether pre- or postemergence, and thus can be quite devastating. Pathogens inciting damping-off are global in occurrence, although exact species may differ from region to region. Symptoms/signs. Damping-off may be the result of seeds or seedlings rotting prior to emergence, or more frequently, the seedlings collapse as they emerge as the roots and/or stem at the soil line becomes necrotic and the plant collapses. When older seedlings are affected, the plant becomes stunted and dies later. Pythium spp.

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generally attack the root tips, causing a dark brown to black rot, while various Phytophthora spp. may attack the fine roots including the tap root, crown, and all belowground parts (Banihashemi 1975). Rhizoctonia solani commonly causes a brown to black lesion encircling the stem at the soil line and may occur both on seedlings and older plants (Srinivasan and Visalakchi 2010). Biology and epidemiology. All three pathogen groups have broad host ranges. Pythium species reported on sunflower include P. aphanidermatum (Edson) Fitzpatrick, P. irregulare Buisman, P. debaryanum Hesse, P. rostratum Butler, and P. splendens Hans Braun (Hendrix and Campbell 1973). Phytophthora species reported on sunflower include P. cryptogea Pethybridge and Lafferty, and P. drechsleri Tucker. Rhizoctonia solani isolates from sunflower belong to anastomosis groups AG-3 and AG-4. Pythium spp., Phytophthora spp., and R. solani are all soilborne and found worldwide, and Rhizoctonia has been shown to be seedborne in sunflower (Lakshmidevi et al. 2010). They can survive for years in soil or infected crop residues with overwintering structures (sclerotia for Rhizoctonia and oospores for Pythium and Phytophthora). Sclerotia of R. solani germinate to form mycelium which attacks seedling roots. The oospores produced by Pythium spp. and Phytophthora spp. germinate to form motile zoospores that are chemotactically attracted to root exudates. Thus, water-logged soils are necessary for infection by Pythium spp. and Phytophthora spp., with longer periods of saturation leading to more damping-off. Management • Cultural practices – Crop rotation is generally of little use due to the broad host ranges and the longevity of overwintering fungal structures. At this time, disease management is aimed at prevention. Using disease-free planting media and avoiding overwatering are the easiest means of minimizing damping-off. • Sanitation – Discard pots or flats with plants showing damping-off. • Soil disinfestation – For field production in warmer climates, solarization has been reported to provide control. • Fungicides and biocontrols – Fungicidal seed treatments, initially registered for control of downy mildew (metalaxyl and mefenoxam) will generally give control of Pythium and Phytophthora, while broad-spectrum strobilurin fungicides will help manage Rhizoctonia. Various biocontrol formulations of Bacillus, Gliocladium, and Trichoderma spp. are available that are applied as seed inoculations, incorporated into potting soil, or used as soil drenches. Fungicide soil drenches are also used in nonorganic production. Consult government publications for recommended products. • Resistance – Although differences in susceptibility have been noted among oilseed sunflowers to these three pathogens, resistance has not been incorporated into either oilseed or ornamental sunflower.

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2.5

Downy Mildew (Plasmopara halstedii (Farlow) Berlese and de Toni)

Geographical occurrence and impact. The pathogen causing downy mildew is found worldwide, with the exception of Australia. Downy mildew is one of the major diseases affecting oilseed sunflower, and can be devastating on ornamentals as well. The pathogen primarily causes systemic infection, which renders the plant unsalable thus affecting florists’ trade. Symptoms/signs. The most commonly seen symptoms are those caused by root infection of seedlings leading to a systemic infection (Friskop et al. 2009). Affected plants have chlorosis on the upper surface of leaves (either the entire leaf or the interveinal areas), with a white covering of spores on the underside of the leaves. Affected plants are severely stunted, and if they do not die within a few weeks, they will produce a head which is horizontal rather than vertical. The pathogen, by means of airborne spores, can also incite leaf spots typical of other downy mildews, with chlorotic spots on the upper surface, with a white coating of spores on the underside of each lesion. These “local lesions” do not lead to systemic infection and are of less impact as the affected leaves can be removed (Fig. 4a, b). Biology and epidemiology. Downy mildew is caused by the obligate pathogen Plasmopara halstedii (Farlow) Berlese & de Toni (syn. Peronospora halstedii Farlow). The host range of the causal pathogen includes sunflower and all wild Helianthus species, plus a number of related Asteraceae weeds and native plants, including Ambrosia, Bidens, Iva, Eupatorium, Silphium, Solidago, and Xanthium. P. halstedii is also reported to cause downy mildew on some ornamental genera such as Centaurea, Coreopsis, Dimorphotheca, Erigeron, Gerbera, Rudbeckia, Senecio, and Verbena; however, cross-pathogenicity studies to prove isolates from these can infect sunflower, and vice versa, have not been done. Plasmopara halstedii exists as physiological races on sunflower, and there is evidence that there are biotypes infective on specific genera, even though morphologically the pathogen isolates are indistinguishable.

Fig. 4 (a) Downy mildew infected plants showing stunting and leaf chlorosis. (b) Downy mildew infected leaves showing chlorosis on upper surface and white sporulation on bottom surface (T.J. Gulya. USDA.)

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Diseases of Sunflower

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Plasmopara halstedii overwinters as resistant oospores, which germinate in water-logged soils to produce motile zoospores. The spores swim towards sunflower roots and infect young seedlings to produce a systemic infection. Infection of older plants will produce a club root without leading to systemic infection. Once the fungus becomes systemic and produces spores on the undersurface of leaves, these spores can be blown some distance and can initiate foliar infection (i.e., local lesions). As the plants die, the fungus produces oospores which can remain viable for years in the soil. The fungus and disease development is fostered by cool (4 h). Thus, long periods of dew or fog producing noninterrupted periods of free water on the plant is more conducive to infection than short periods of rain, regardless of the amount. Temperatures > 25  C and lack of free water are detrimental to ascospore germination/infection (Masirevic and Gulya 1992). The ascospores can be dispersed short distances (100 m) by wind currents, as well as by splashing water. The fungus quickly produces abundant white mycelia in all infected plant parts, and then sclerotia, which can be found externally on plant roots and the lower stem, inside the lower stem, and on the head. S. minor sclerotia, in contrast, germinate only to form mycelium, which initiates root infection. Thus, S. minor does not produce head rot since it produces no airborne ascospores. Sclerotinia sclerotiorum has an extremely broad host range, encompassing almost 400 plants in 278 genera of herbaceous plants with the Asteraceae, Cruciferae, Leguminosae, and Solanceaceae among the most important families. Important

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crop hosts include lettuce, beans, soybeans, canola, and sunflowers, plus many Asteraceae ornamentals. Sclerotinia minor has a slightly less broad host range of 94 species in 66 genera. Important hosts include lettuce, Brassica crops, peanuts, and sunflower. Management. Diseases caused by either of the Sclerotinia species are very difficult to control, and the best management is aimed at prevention. • Cultural practices – Since infected plants can spread Sclerotina via root-to-root contact, lower plant densities will minimize wilt incidence (Nelson et al. 1989). To minimize head infection, anything which decreases foliage, increases air movement, or decreases head/leaf wetness will help, including lower nitrogen fertilization, lower plant densities, and furrow/drip versus overhead irrigation. Many different biocontrol/cultural practices have been shown to be partially effective at decreasing soil populations of sclerotia, and thus minimizing both root and head infections. Deep plowing, with a moldboard plow to completely invert the soil profile to >15 cm, will bury sclerotia into an aerobic environment where microbes should degrade the sclerotia (Mueller et al. 2002). However, there is conflicting data on this practice, and most experimental trials suggest that shallow burial leads to faster degradation of sclerotia (Ćosić et al. 2012; Subbarao et al. 1996, P. 48). Planting dense cover crops of cereal grains will produce the microenvironment conducive to apothecial formation, and if this crop is tilled under just prior to a susceptible crop, the net effect is to sap the sclerotia of energy/biomass sufficient to prevent production of more apothecia (Mueller et al. 2002). Rotation is of minimal use, since there are so many susceptible crops, but if several years of a nonhost monocot (for example, grasses or cereals) are planted, this also would hasten lowering the sclerotial soil population (Mueller et al. 2002). Once either stem rot or head rot are observed, there are no curative measures to save those plants. The best option at that point is to physically remove affected plants and dispose of them away from the field, thus minimizing soil contamination with more sclerotia. • Biological control – Many commercial biocontrol products are available, based on fungi such as Coniothyrium, Gliocladium, and Trichoderma, and bacteria such as Bacillus, and these applied as soil drenches immediately following a Sclerotinia infestation will hasten sclerotial degradation and shorten the interval between planting another susceptible crop. • Soil disinfestation – Preventative measures include using pasteurized soil, solarization, soil fumigation, or, where practical, flooding of fields for several weeks, all of which are aimed at decreasing the soil population of sclerotia. Biofumigation by planting Brassica cover crops and tilling them under will release isothiocyanates, which are toxic to a range of fungi (Griffiths et al. 2011). • Resistance – In certain sunflower hybrids, some progress has been made in developing partial (incomplete) resistance or tolerance to S. sclerotiorum. There are no hybrids available with complete resistance to either of the two Sclerotinia spp.

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Septoria Leaf Spot and Blight (Septoria spp.)

Geographic occurrence and impact. Septoria leaf spot is noted on sunflower throughout the world on every continent. The disease is most destructive in areas with heavy rainfall but is seldom observed in areas with drier climates. Under optimal disease conditions, premature leaf senesce and defoliation will result. Symptoms/signs. Septoria blight is characterized by angular to circular necrotic leaf spots reaching 10–15 mm in diameter. Lesions are tan to gray with dark brown margins, and may be surrounded by a yellow halo. Symptoms may be observed on cotyledons and leaves of young seedlings, but it is more common to see severe symptoms on older plants. One characteristic feature which distinguishes Alternaria and Septoria leaf spots is the presence of pycnidia in the lesions produced by Septoria spp., which appear as small, black dots, barely visible with the naked eye. In addition, Septoria lesions are generally much larger than Alternaria lesions. Lesions coalesce in time, turning the leaf necrotic and defoliating the plant from the bottom leaves up (Fig. 14a, b). Biology and epidemiology. Two species of Septoria are known to infect sunflower, Septoria helianthi Ellis and Kellerman found worldwide (Holliday and Punithalingam 1970) and Septoria helianthina Petrov and Arsenijevic, initially described in Yugoslavia (Petrov and Arsenijevic 1996). Both Septoria spp. are thought to be restricted to sunflower and wild Helianthus spp. Septoria spp. overwinter as pycnidia on infected plant residue. Spores ooze from the pycnidia when wet and are spread by splashing rain and wind. Initial infections occur on lower leaves, where the microclimate is more conducive, and the upper leaves become affected later. Rapid disease development is fostered by abundant rain and moderately high temperatures. The development of disease is arrested with hot, dry weather but can resume with the return of favorable conditions. Management • Cultural practices – Any cultural practice such as crop rotation and tillage of crop residue will lessen the carryover of inocula.

Fig. 14 (a) Septoria leaf blight (T.J. Gulya. USDA.). (b) Closeup (10X) of lesion showing the fungal pycnidia, visible as tiny black dots (S. Markell # 2017. All Rights Reserved.)

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• Fungicides – Broad spectrum fungicides commonly used to control foliar diseases on other ornamental crops should be effective against these Septoria spp. • Resistance – Genetic resistance has been noted in oilseed germplasm (Block 2005; Carson 1985), but this has not been incorporated into commercial hybrids nor into ornamental varieties.

2.16

Sclerotium Blight (Sclerotium rolfsii Saccardo)

Geographic occurrence and impact. Sclerotium blight or wilt is found in Africa, Asia, Australia, Europe, and both Americas in areas that have tropical or semitropical climates (Punja 1985). In the United States, it is most frequently seen in Florida and the Gulf coast. Thus, ornamental sunflowers grown indoors in hot, humid conditions have the potential for significant losses due to this disease. Symptoms/signs. The main symptom of Sclerotium blight or “Southern blight” is a girdling lesion at the soil surface, generally appearing on plants as they reach the bud stage. This tan to dark brown lesion may have alternating light and dark bands, corresponding to the diurnal growth of the fungus. Under moist conditions, a white mycelial mat may form over the lesion and around the base of the plant. Sclerotia will form on the exterior of the lesion. The sclerotia are uniformly round, 0.5–2 mm, and brown, in contrast to the black sclerotia formed by Sclerotinia spp. The fungus will rot the pith but the rind will be intact, and thus, there seldom is lodging (Fig. 15).

Fig. 15 Sclerotium blight (T.J. Gulya. USDA.)

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Biology and epidemiology. Sclerotium rolfsii is the asexual stage of this pathogen; the sexual stage Athelia rolfsii (Curzi) Tu and Kimbrough is seldom observed in nature but can be produced in culture. The mycelium produced by this fungus is abundant, white, and cottony, and very similar to that produced by Sclerotinia spp. The sclerotia are the best means to distinguish S. rolfsii from the two Sclerotinia spp. Sclerotinia sclerotiorum produces sclerotia which are black, irregularly shaped but generally long and thin, and can reach up to 5 cm long. S. minor produces small round sclerotia (0.5–2 mm) black in color, while S. rolfsii sclerotia are similarly small (0.5-2 mm) but tan to dark brown. Soilborne sclerotia germinate to form a mycelium which infects stems at or slightly below the soil line (Aycock 1966). Sclerotial germination is favored by moist, but not saturated soil at temperatures from 25  C to 35  C. From initially infected plants, mycelia can move plant-to-plant via root contact or even grow on the soil surface under wet conditions. Sclerotia can survive for several years in soil, but do not withstand alternating freeze/thaw cycles, so this fungus is thus not seen in colder climates. Like the Sclerotinia spp., this pathogen has a wide host range of nearly 500 dicot plants, but it also infects some shrubs and trees. Management • Cultural practices – Crop rotation with nonhosts (for example, cereals and grasses) and deep burial of sclerotia (>15 cm) will aid disease control (Hagan 2004). Early season planting may have less disease due to cooler soil temperatures. • Soil disinfestation – Solarization or fumigation is more effective against sclerotia than soil-applied fungicides. • Fungicides – Seed or soil-applied fungicides for control of the seedling phase of the disease can be effective.

2.17

Verticillium Leaf Mottle/Wilt (Verticillium dahliae Klebahn)

Geographic occurrence and impact. The fungus causing Verticillium wilt is found globally, and with its wide host range and longevity in the soil, it can be a serious problem not just for sunflower but for any other floral crops grown in the same field/soil. Symptoms/signs. External symptoms of Verticillium infection are more accurately described as a leaf mottle (Sackston et al. 1957). Leaf symptoms generally appear on plants as they reach bud stage, with symptoms appearing on lower leaves and progressing up the plant. Interveinal areas become chlorotic and then quickly die to produce dark brown-black areas surrounded by a yellow margin. These symptoms are often mistaken for the initial leaf symptoms of Phomopsis stem canker. However, with Phomopsis stem canker, the necrotic areas are centered on the veins, while with Verticillium wilt, they are always between the veins. The entire leaf surface will be affected with this mottle pattern, and eventually the first affected leaves will senesce

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Fig. 16 (a) Verticillium leaf mottle/wilt. (b) Stem sections showing vascular browning caused by Verticillium (T.J. Gulya. USDA.)

and die. Wilting is not always observed, despite the inaccurate common name. Severely affected plants may die prior to maturity, producing small heads. Another way to confirm Verticillium infection is to cut the lower stem, either longitudinally or in cross section. The fungus will cause the vascular elements to be discolored brown. In severely affected plants, when the stem is cut open, the surface of the pith will be uniformly black, due to the presence of tiny microsclerotia (Fig. 16). Biology and epidemiology. Older literature incorrectly identified the causal fungus as V. albo-atrum, “microsclerotial form” but Verticillium dahliae is the correct causal agent (Sackston et al. 1957). Another fungus, Phialophora asteris (Dowson) Burge and Isaac (syn. Cephalosporium asteris Dowson), produces symptoms in sunflower very similar to Verticillium, and its morphology in culture is also similar, making identification difficult (Hawksworth and Gibson 1976). Verticillium dahliae produces conidia, but the microsclerotia are the propagules that overwinter and are responsible for infection. Verticillium dahliae is very slow growing in culture, and isolations made from diseased plant tissue are often mixed with faster growing contaminating fungi, further complicating pathogen identification. Verticillium dahliae has a host range of over 400 species of herbaceous and woody plants, including trees, vegetables, flowers, and weedy species. Susceptible flower crops include aster, chrysanthemum, cineraria, dahlia, geranium, gerbera, marigold, rose, snapdragon, statice, and stock, among others. Among field crops, potatoes and mint are severely affected.

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Verticillium dahliae overwinters in soil or plant residue as microsclerotia, and these remain viable for many years. Microsclerotia are stimulated to germinate by root exudates to form a mycelium which penetrates the roots, enters the vascular system, and eventually becomes systemic throughout the plant. Moist, but not waterlogged soil, and temperatures between 21  C and 27  C are optimal for sclerotial germination and root infection. The fungus produces a toxin that is translocated to the leaves, which induces the foliar symptoms. Thus, isolation from symptomatic leaves may yield no Verticillium as the symptoms are the result of the toxin produced some distance away. Wilting, when it occurs, is the result of fungal mycelia and polysaccharides plugging the vascular system. If the mycelium reaches the head, the seeds will become infected. The fungus produces copious amounts of black microsclerotia internally, which gives the pith a charred, blackened appearance. The microsclerotia are smaller than those produced by Macrophomina phaseolina (charcoal rot) and cannot be seen with the naked eye or even a 10X lens. Management. As with other soilborne fungal pathogens forming sclerotia, management of Verticillium is challenging (Berlander and Powelson 2000). The sclerotia, despite their small size, are difficult to kill with fungicides; they exist in large numbers in the soil, and the fungus has a large host range. Since Verticillium is a serious problem on some high cash value crops like potatoes, there are commercial laboratories that assay soil samples to determine the presence and quantity of Verticillium microsclerotia. • Soil disinfestation – Thus, prevention is the first aim of a management program, and this starts with clean seed and pathogen-free soil, via pasteurization, solarization, or fumigation. Flooding the soil for 20 days significantly reduces the number of microsclerotia, as does soil acidification with aluminum sulfate. As mentioned for Sclerotinia wilt, the practice of biofumigation with Brassica cover crops has been shown to decrease the number of microsclerotia and the severity of the disease. • Cultural practices – While extra irrigation might be considered to help plants infected with a wilt pathogen, research has shown that frequent or excessive irrigation actually increases the severity of Verticillium. • Resistance – Genetic resistance is available (Hoes et al. 1973) and widely deployed in oilseed sunflower germplasm. The fungus exists as several races (Gulya 2007) and vegetative compatibility groups (VCGs) (García-Carnero et al. 2014), and this needs to be known so that breeders can select the appropriate single, dominant resistance gene.

2.18

White Rust (Albugo tragopogonis (DeCandolle) Gray = Pustula helianthicola Rost and Thines)

Geographic occurrence and impact. Albugo or white rust is a disease historically found primarily in the Southern Hemisphere (Argentina, Australia, South Africa)

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(Kajornchaiyakul and Brown 1976; Delhey and Kiehr-Delhey 1985) but more recently has been noted on both ornamental and oilseed sunflowers in Europe (Crepel et al. 2006; Thines et al. 2006), China (Chen et al. 2006), and, very infrequently, in the United States (Gulya et al. 2002b). The disease requires very specific environmental conditions, and the pathogen appears to be very host specific, and thus the disease is unlikely to appear year-in and year-out. While the disease is usually not devastating to the oilseed crop, the white blisters on the foliage detract from the value of ornamental sunflower. Symptoms/signs. The disease is usually manifested by raised, small to fairly large (1–2 mm or 5–10 mm) pustules or blisters on the upper leaf surface, with white sporulation on the underside of the leaf (Kajornchaiyakul and Brown 1976). With age, the Albugo pustules become necrotic and the infected tissue may fall out, leaving a “shot hole” appearance. The causal fungus can also cause similar white blisters on floral bracts, and may also initiate faintly black, water-soaked lesions on petioles and the stem. Stem lesions are usually internodal and may be colonized by secondary fungi, leading to lodging in severe cases (Krüger et al. 1999; Van Wyk et al. 1995) (Fig. 17). Biology and epidemiology. The fungus causing this disease is not a true rust and is actually more closely related to the downy mildews. The causal agent of white rust was classified as Albugo tragopogonis for over a century but was recently renamed Pustula helianthicola (Rost and Thines 2012). The fungus overwinters as oospores on infested plant debris and may also be seedborne (Viljoen et al. 1999). Oospores, the primary inoculum, are spread by rain and splashing water. They germinate to form motile zoospores that enter through leaf stomata. The white asexual sporangia produced on the underside of leaf pustules can be windblown to cause secondary infections on leaves, stems, petioles, and bracts. Oospores typically develop only in stem and petiole lesions, which cause the subtle, dark lesions. The disease is dependent on free water, from rain, dew, or irrigation. Optimum infection takes

Fig. 17 White rust on sunflower – leaves (T.J. Gulya. USDA.)

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place at night between 10  C and 15  C, while optimal disease development is at temperatures between 20  C and 25  C. Thus, the disease is favored in environments with cool nights and warm days (Kajornchaiyakul and Brown 1976). Management • Cultural practices – Overhead irrigation at night will foster conditions conducive to foliar infection, and should be avoided. • Sanitation – Removal of plant debris from previous sunflower crops, either in the same or nearby fields, will diminish the possibility of infection. Leaves stripped prior to shipping cut sunflowers, if infected, should be disposed of outside the field, and preferably composted or removed entirely. • Fungicides – Seed-applied fungicides which control downy mildews (metalaxyl, mefenoxam, and strobilurins such as azoxystrobin) will provide early season control. Thereafter, foliar applications of these fungicides should provide protection against leaf infection (Lava et al. 2015). • Resistance – In oilseed sunflower germplasm, resistance to white rust has been noted and is controlled by multiple genes, each governing resistance in different plant parts. No effort has been made to incorporate any type of white rust resistance into ornamental sunflowers, although it has been noted to occur in one older commercial cultivar (“Abendsonne”). Many annual and most perennial Helianthus species appear to be resistant (Lava et al. 2015).

3

Bacterial and Phytoplasma Diseases

3.1

Apical Chlorosis and Bacterial Leaf Spot (Pseudomonas syringae pv. tagetis and Pseudomonas syringae pv. helianthi)

Geographic occurrence and impact. Apical chlorosis was first identified on marigold in Australia (Trimboli et al. 1978) and later on sunflowers in the United States in the early 1980s (Gulya et al. 1982). The causal bacterium is found worldwide on many Asteraceae flowers and weedy species. Bacterial leaf spot is caused by a very closely related bacterium, and it also is found worldwide (Piening 1976). Less is known about its host range, and it may be restricted solely to Helianthus. Apical chlorosis is more likely to be observed and cause alarm to growers due to its spectacular appearance, but since its symptoms are confined to the foliage, which is stripped off for cut flower production, its impact is lessened. Symptoms/signs. Apical chlorosis first manifests itself as a general chlorosis of the youngest leaf, either a portion or the entire leaf (Gulya et al. 1982). The affected area may be a pale yellow, or in extreme cases the leaf will be almost white. This symptom was initially confused with nitrogen deficiency, but the chlorosis is much more pronounced. The chlorosis will affect subsequent leaves as they develop, but the chlorotic leaves will not turn necrotic. When temperatures warm up, the newly emerging leaves will be normal green in color, but lower, chlorotic leaves will

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Fig. 18 Apical chlorosis (T.J. Gulya. USDA.)

remain so (Fig. 18). Symptoms of bacterial leaf spot vary, depending upon the environment and sunflower cultivar. Lesions begin as small necrotic spots of varying size and shape and may be surrounded by chlorotic haloes. Under severe conditions the individual lesions coalesce to form large necrotic areas, giving the leaf a shothole appearance. Lesions may also form on petioles and stems, often leading to defoliation. Biology and epidemiology. Pseudomonas syringae pv. tagetis (Hellmers) Young, Dye, and Wilkie is the causal bacterium for apical chlorosis, while Pseudomonas syringae pv. helianthi (Kawamura) Young, Dye, and Wilkie causes bacterial leaf spot (Piening 1976). Recent molecular studies suggest that both bacteria may be identical, differing only in a toxin production gene present in P. syringae pv. tagetis. Pseudomonas syringae pv. tagetis has a wide host range within the Asteraceae, affecting ornamental flowers such as Tagetes and Zinnia, as well as many weedy genera such as Ambrosia, Cirsium, and Taraxacum (Rhodehamel 1985). Less research has been done with P. syringae pv. helianthi, but its host range appears to

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be restricted to Helianthus spp. Both bacteria can easily be identified as P. syringae pathovars based on the fluorescence under UV light of colonies grown on King’s B medium. Both bacteria survive in the soil and are seedborne. Apical chlorosis also persists on infected weedy hosts. Both are spread by splashing water and are typically seen early in the growing season during cool weather with wet soils. Apical chlorosis is typically seen in low areas of a field. Individual plants may be affected or there may be several affected plants in a row. Pseudomonas syringae pv. tagetis produces a toxin that interferes with chloroplast formation in developing leaves. In severe cases, the leaf may completely lack chlorophyll, but other pigments are not affected. Chlorotic leaves remain so throughout the life of the plant, but the appearance of new chlorotic leaves will cease as soils dry out and air temperatures warm up. Bacterial leaf spot is quite common, and its appearance is not restricted to cool early season and wet soil (Arsenijevic et al. 1994). Infection generally starts on lower leaves where the microclimate is more favorable for both fungal and bacterial infection. Management. Since there are no curative bactericides, there is no management option once symptoms are observed. • Cultural practices – In field situations, limiting irrigation to prevent water-logged soils and water splashing, especially during cool weather, will likely preclude the appearance of apical chlorosis. • Resistance – Differences in susceptibility to both bacteria have been noted in oilseed germplasm, but neither disease has been severe enough to warrant transferring that resistance into either oilseed hybrids or ornamental cultivars. • Sanitation – As the symptoms of apical chlorosis will cease when temperatures warm up, the damaged leaves can be stripped off ornamental sunflowers, and similarly with bacterial leaf spot.

3.2

Bacterial Stalk Rot/Head Rot (Erwinia = Pectobacterium spp.)

Geographic occurrence and impact. The bacteria causing stem and head rot are ubiquitous in soils and are present in every continent. Stem infections, though uncommon, would render ornamental sunflowers unsaleable. Head rot, although potentially serious on oilseed sunflower, does not occur in the bud stage, so it is unlikely to be observed on cut sunflowers but could be present in potted plants or homegrown sunflowers. Symptoms/signs. Bacterial stem rot and head rot can usually be recognized by smell as well as visual symptoms. The exterior of affected stems may have no discernible lesions or may have a blackened area, often centered around a petiole (Gudmestad et al. 1984). Upon splitting the stem open longitudinally, the pith will

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Fig. 19 Bacterial stalk rot (T.J. Gulya. USDA.)

appear very water soaked, and if the infection has progressed long enough, there will be a strong odor of rotting potatoes. In time, due to gas produced by the bacteria, the stem will split open longitudinally and bacterial slime may ooze out, which often attracts flies and other insects. After this phase the stalks will either lodge or then may dry up and turn completely black, and the odor will be gone (Fig. 19). The bacterial head rot phase is characterized by symptoms that initially are identical with those caused by many fungal incited head rots. Small, depressed, watery, soft rotted areas develop on the back of the heads that enlarge and turn from tan to brown. Again there is a distinct odor of rotting potatoes, and if one grasps such a head, the soft mushy condition will be evident. Head infections may also produce copious amounts of slime and exudate, which may smell either of rotting potatoes, or in some cases, of alcohol, due to the presence of other secondary organisms such as

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yeasts. These exudates, combined with the bacterially produced gas, often look like foam on the head and stem. The smell and sugars in the exudates attract many insects, including flies, bees, and various beetles. Biology and epidemiology. Pectobacterium carotovorum (Jones) Waldee and Pectobaterium atrosepticum (van Hall) Lehmann and Neumann are both responsible, alone or in combination, for soft rots of sunflower stems and heads. These “soft-rotting bacteria” can affect many plants, causing rots of flowers, soft fruit, and vegetables. A species from Mexico, Pectobacterium cacticidum, whose primary hosts are cacti, was recently isolated from sunflower (Valenzuela-Soto et al. 2015). Both Pectobacterium spp. are soft-rotting bacteria that are found in soils and decaying plant debris (Charkowski 2015). They are very weak pathogens and require wounds to infect healthy tissue. These wounds can be mechanical damage, or caused by insects, birds, or hail. Free water and high temperatures speed the development of soft rot once the bacteria have colonized a wound. If insect larvae tunnel into the stems, they will not only vector the bacteria but will hasten its spread within a plant. Once the bacteria become established internally within the stem or head, the influence of air temperature and rainfall is negligible. Management. Bacterial diseases, especially caused by ubiquitous organisms, are hard to control, and prevention is the best approach. Controlling insects and birds to minimize wounds through which the bacteria gain entrance is a sound first step (Chattopadhyay et al. 2015). Ground irrigation rather than overhead will minimize the free water necessary for infection. Once an infected plant is noted, it should be physically removed and disposed of outside the greenhouse or field, which will eliminate the inoculum that feeding insects might spread.

3.3

Aster Yellows (Phytoplasma spp.)

Geographic occurrence and impact. Aster yellows has been previously confused with virus diseases, then classified as caused by a mycoplasma (now known as phytoplasmas) (Bertacinni and Duduk 2009). The causal agent causes a “yellows” disease on over 200 crops and is the most widely seen “yellows” on nursery crops. The complex of phytoplasmas affecting sunflower has been reported from North and South America, Europe, and Northern Africa. Symptoms/signs. Sunflower plants affected by “yellows” exhibit a range of symptoms including leaf chlorosis (of the entire leaf or in sectors), proliferation of secondary shoots, and various abnormalities of the head. There may be wedgeshaped sectors of the head showing hypertrophy, and bracts and ray petals may appear in the center of the head (Fig. 20).

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Fig. 20 Aster yellows (T.J. Gulya. USDA.)

Biology and epidemiology. Aster yellows is caused by one or more closely related phytoplasmas. These organisms are smaller than bacteria, larger than viruses, and a few can, with difficulty, be grown in vitro. The phytoplasma causing aster yellows in sunflower and other crop is referred to as “Candidatus Phytoplasma asteris” (Harrison and Helmick 2008). Another phytoplasma causing similar symptoms has been studied in Argentina and tentatively termed “Candidatus Phytoplasma Sunflower Phyllody” (Guzmán et al. 2014), and a third one was characterized in Iran (Salehi et al. 2015). Phytoplasmas are vectored by several genera of leafhoppers. The phytoplasma survives in perennial Asteraceae weed species and leafhopper vectors. Aster yellows multiplies within the body of the leafhopper and is transmitted through the insects saliva when feeding. The Aster yellows phytoplasma is believed to have a host range of more than 150 species, including many flower crops, vegetables (carrot, lettuce, potato, tomato), and even grains and grasses. Cool, wet weather seems to favor both the leafhopper vector and the disease. Management • Vector management – No chemicals are directly effective against phytoplasmas. Controlling the insect vectors, however, whether done by insecticides, biological control, or cultural practices (i.e., exclusion netting on greenhouse vents) will all but eliminate the disease. Removing weeds and volunteer sunflowers from the field and perimeter will reduce the inocula and vectors. • Cultural practices – Some of the more susceptible floral crops include Petunia, Tagetes, Viola, Chrysanthemum, Rudbeckia, and Salvia, and thus it would be wise not to plant these in the vicinity of sunflowers, and to monitor all susceptible crops closely for symptoms and leafhoppers. • Resistance – Aster yellows was once severe enough in Canada on oilseed sunflower to warrant research, and resistance was found within oilseed germplasm (Putt and Sackston 1970). Thus, it would be possible to transfer that level of resistance into ornamental sunflowers.

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4

Viral Diseases

4.1

Sunflower Mosaic Virus

Geographic occurrence and impact. Sunflower mosaic virus is the only virus documented to occur naturally on sunflower in the United States, and currently is known nowhere else. Its range within the United States at the moment is restricted to southern Texas. As such it is classified as a quarantine pathogen by many importing countries, and while its impact upon ornamental sunflowers may not be great, the threat of restricted international movement of plants or seeds would be an economic hardship. Several other viruses have been confirmed on sunflowers in other countries, but their occurrence has usually been sporadic and limited to the specific countries. Thus, sunflower chlorotic mottle virus (SuCoMoV) has been found in Argentina (Giolitti et al. 2010), Pelargonium zonate spot virus (PZSV) also in Argentina (Giolitti et al. 2014), sunflower necrosis caused by tobacco streak virus (TSV) in India and Australia (Sharman et al. 2009; Prasada Rao et al. 2000), Bidens mottle virus (BiMV) in Florida (Wisler 1984) and Taiwan (Liao et al. 2009), plus several other instances of virus diseases of undetermined origin (Loebenstein and Thottappilly 2003). Symptoms/signs. Sunflower mosaic is characterized by a mild mosaic pattern on leaves of young plants, becoming more pronounced, and then fading as the plant matures (Gulya et al. 2002a). The virus does not cause stunting, or any change in leaf or head morphology, nor does it cause ringspots (Fig. 21). Biology and epidemiology. The disease is caused by Sunflower mosaic virus (SuMV), a Potyvirus. Virus particles are long, flexuous rods. Their long size,

Fig. 21 Sunflower mosaic virus (T.J. Gulya. USDA.)

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along with characteristic pinwheel inclusion bodies, is diagnostic for Potyviruses. SuCoMoV is also a Potyvirus, while PZSV and TSV are Ilarviruses, with small, isometric virus particles. The Rio Grande river valley of southern Texas, where the disease is endemic, is subtropical, so that both wild sunflower and insect vectors can be found year-round. The virus was vectored experimentally by aphids (Myzus persicae and Capitphorus elaegni), both of which are common greenhouse inhabitants. Sunflower plants are optimally susceptible for 1 month, after which they become progressively less receptive to the virus. The virus was found to be seed transmitted. SuMV has a very narrow host range, consisting of Helianthus, Zinnia, and Sanvitalia. Other sunflower viruses, such as SuCoMoV, TSV, and PZSV, have much broader host ranges. Management. While resistance to SuMV and SuCoMoV has been noted, it has not been transferred to commercial oilseed hybrids or to ornamental cultivars (Gulya et al. 2002a; Lenardon et al. 2005). The most effective management for these sunflower viruses include the use of virus-free seed, minimizing insect vectors (aphids and thrips) through insecticide sprays or cultural methods, and removing infected plants. The use of silver reflective mulch in field plantings has been shown to repel aphids, as well as conserving soil moisture.

5

Nematodes

5.1

Root-Knot Nematodes (Meloidogyne Species)

Geographic occurrence and impact. Root-knot nematodes are found worldwide on many hosts but are especially serious on sandy soils and in warm climates. Countries where root-knot nematodes have had the greatest impact on sunflower are Egypt, Mozambique, South Africa, and Zambia in Africa (Bolton et al. 1989; Lawn et al. 1988); Pakistan and India in Asia (Ravindra et al. 2015); and Greece and Italy in Europe (Tzortzkzkis et al. 2014; Zazzerini et al. 1997). In the United States, root-knot nematodes have been documented on sunflower in Florida and Texas (Rich and Dunn 1982; Isakeit 2011). There are currently 30 Meloidogyne species in the United States, of which five are known to parasitize sunflower experimentally. M. hapla is the most widespread, found in every state except Alaska, while M. incognita, M. arenaria, and M. javanica are most common in southern and coastal states, and M. chitwoodi is only found in the western states of WA, OR, CA, CO, UT, and NM, and in Mexico (Walters and Barker 1994). M. incognita appears to be most serious on sunflower. Lack of confirmation of root-knot nematodes on sunflower in a particular state may be due to the lack of nematologists to confirm their presence. Symptoms/signs. Symptoms of root-knot nematode infection may not be apparent aboveground, or the plant may display stunting, chlorosis, and daytime wilting. Affected plants may also have fewer and smaller leaves and flowers. Root galls are

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Fig. 22 Root-knot nematode damage on sunflower hybrid “954”, with smaller galls caused by Meloidogyne hapla (left) and larger galls caused by M. arenaria (right) (M.K. Beute # 2017. All Rights Reserved.)

the belowground symptom (Fig. 22). The extent and size of root galls or “knots” depends upon the nematode population density, the species of Meloidogyne, and the host cultivar. Among ornamentals and field crops, sunflower is rated as highly susceptible (Crow 2014; Ferris et al. 1993). In the absence of galls, only a soil analysis in a nematology lab will confirm Meloidogyne as the causal agent, and be able to pinpoint the exact species. Biology and epidemiology. There are five Meloidogyne species documented to parasitize sunflower: M. arenaria (Neal) Chitwood; M. chitwoodi Golden, O’Bannon, Finley and Santo; M. hapla Chitwood; M. incognita (Kofoid and White) Chitwood; and M. javanica (Treub) Chitwood. Root-knot nematodes are sedentary endoparasitic nematodes, meaning they spend most of their life cycle within plant tissue. They hatch from eggs, stimulated by both temperature and exudates from plant roots. The juvenile nematodes migrate through the soil, penetrate the tips of fine roots, and the nematode’s head becomes embedded in the vascular bundle where it begins feeding and becomes immobile. Root cells around the nematode enlarge and form galls, which inhibit root function, leading to the aboveground symptoms. Female nematodes can produce eggs parthenogenetically, and up to several hundred eggs in a gelatinous mass can be found on the root surface. Generation time can vary from 20 to 45 days, allowing for multiple infections in a growing season.

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Management • Cultural – Depending on the root-knot species present, rotation with nonhosts (arugula, onion, sesame, sorghum-sudangrass) will decrease the nematode population but will not completely eliminate Meloidogyne (Dover et al. 2004; El-Ghonaimy et al. 2015; Kokalis-Burelle et al. 2013). Conversely, planting of “trap crops” of susceptible hosts, such as cowpea, and removing them completely, has also been shown to decrease damage on subsequent sunflower crops. Selecting preceeding crops in a commercial nursery which are least susceptible will minimize damage to sunflower (Creswell 2000). Solarization, in climates where feasible, will reduce nematode populations. • Chemical – While no nematicides are registered for use by home owners, several fumigants are commonly used in high-value annual crops and the nursery trade which are effective against all soilborne nematodes, including chloropicrin, 1,3-dichloropropene, metam sodium, and the nonfumigant ethoprop (Stapleton et al. 2014). • Genetic Resistance – Evaluations of oilseed sunflowers have demonstrated that there is considerable variation in susceptibility to Meloidogyne spp., but to date there have been no public releases of root-knot tolerant or resistant germplasm (Fabiyi and Atolani 2013; Rehman et al. 2006). • Biological – Several nematode-trapping fungi and antagonistic bacteria have been identified and shown to reduce, but not eliminate, populations of Meloidogyne spp. (Karssen et al. 2013; Duncan & Moens 2006). At least two mycopesticides, Myrothecium verrucaria (DITERA), and Paecilomyces lilacinus (MELOCON) are registered for use on ornamentals in the United States (Stapleton et al. 2014; Crow 2014).

5.2

Root-Lesion Nematodes (Pratylenchus Species)

Geographic occurrence and impact. Root-lesion nematodes, of the genus Pratylenchus, have a very wide host range and are found throughout the world, ranging from cool temperate to hot tropical climates (Davis and MacGuidwin 2005). They rank third behind root-knot and cyst nematodes as the nematodes of greatest economic impact on all crops worldwide. Their small size (0.4–0.7 mm long) relative to other plant parasitic nematodes enables them to survive in almost any texture soil. At least eight species have been found on sunflower worldwide, with P. penetrans the most widely distributed and important species. It interacts with wiltinciting fungi like Fusarium and Verticillium, thus augmenting the physical damage it causes (Castillo and Vovlas 2007). Symptoms/signs. Aboveground symptoms due to root-lesion nematodes are nonspecific, and like root-knot damage, are manifested by stunting and chlorosis, and wilting during hot days. Nematode damage in fields often occurs in

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random patches which are associated with higher populations of the nematodes. Root-lesion nematode feeding will cause brown lesions on the exterior of roots, while feeding inside roots will result in cracking and eventual rotting. Biology and epidemiology. Pratylenchus species documented on sunflower include P. alleni Ferris, P. brachyurus (Godfrey) Filipjev and Schuurmans Stekhove, P. crenatus Loof, P. hexincisus Taylor and Jenkins, P. penetrans (deMan) Filipjev, P. scribneri Steiner, P. thornei Sher and Allen, and P. zeae Graham. Pratylenchus spp. are usually migratory endoparasites, and remain vermiform throughout their life (Castillo and Vovlas 2007). Males only occur in some species, and thus reproduction is mostly by parthenogenesis. The life cycle can be completed in 28–56 days, depending upon the species, plant host and environment, so several generations can occur during a growing season. As with all plant-feeding nematodes, there are four juvenile life stages following the egg, and culminating in the adult. All juvenile and adults feed on roots, burrowing into the cortical tissue, and they can enter and leave roots during any life stage. Reports on how far they may travel range from 2 cm up to 1–2 m in the soil to reach other plants. Movement of soil and/or infected plant material provides a much wider dispersion. Pratylenchus overwinter in infected plants, including weedy hosts, in any life stage. Eggs can also dehydrate and become dormant and remain viable for over 2 years. Management. Cultural, biological, and chemical management recommendations for root-lesion nematodes are much the same as for root-knot nematodes, covered previously (Creswell 2000; Stapleton et al. 2014; Thompson et al. 2008). Differences in susceptibility to Pratylenchus in oilseed sunflower have been noted (Bolton and De Waele 1989), but there have been no reports of resistance in any ornamental sunflower germplasm.

5.3

Other Nematodes

Many other nematodes have been identified as causing damage on sunflower. These include dagger (Xiphinema), pin (Paratylenchus), spiral (Helicotylenchus), sting (Belonolaimus), stubby-root (Paratrichodorus), and stunt (Tylenchorhynchus) nematodes, among others (Bridge and Starr 2007). Identification of the causal nematodes always depends upon soil analysis by a nematology lab, as the aboveground symptoms are not nematode specific.

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Rost C, Thines M (2012) A new species of Pustula (Oomycetes, Albuginales) is the causal agent of sunflower white rust. Mycol Prog 11:351–359 Ruegg J (1990) Biology and control of Entyloma eryngii, the cause of leaf spot of the blue thistle (Eryngium alpinum L.). J Phytopathol 130:9–16 Sackston WE, Mcdonald WC, Martens J (1957) Leaf mottle or Verticillium wilt of sunflower. Plant Dis Rep 41:337–343 Salehi M, Esmailzadeh SA, Salehi E (2015) Characterisation of a phytoplasma associated with sunflower phyllody in Fars, Isfahan and Yazd provinces of Iran. BSPP New Dis Rep 31:6 Schilling EE (2006) Helianthus. In: Flora of North America Editorial Committee (ed) 1993. Flora of North America North of Mexico, vol 21. New York/Oxford, pp 141–200 Schoellhorn R, Emino E, Alvarez E (2003) Specialty cut flower production guides for Florida: sunflowers. Univ Florida IFAS Ext Ser Pub ENH885 Schoen JF (1983) Identification of seed-like structures: a taxonomic review of sclerotial-forming fungi. Seed Sci Technol 1:639–650 Sharman M, Persley DM, Thomas JE (2009) Distribution in Australia and seed transmission of Tobacco streak virus in Parthenium hysterophorus. Plant Dis 93:708–712 Shtienberg D (1997) Rhizopus head rot of confectionary sunflower: effects on yield quantity and quality and implications for disease management. Phytopathology 87:1226–1232 Simmons EG (2008) Alternaria: an identification manual. CBS Press, Utrecht, 775 pp Singh S, Tyagi S, Prasad D (2012) Nematode-fungal disease complex involving Rotylenchus reniformis and Macrophomina phaseolina on Helianthus annuus. Ann Plant Prot Sci 20:434–436 Sood PN, Sackston WE (1972) Studies on sunflower rust. XI. Effect of temperature and light on germination and infection of sunflowers by Puccinia helianthi. Can J Bot 50:1879–1886 Srinivasan K, Visalakchi S (2010) First report of Rhizoctonia solani causing disease of sunflower in India. Plant Dis 94:488 Stapleton JJ, McKenry MV, Ploeg AT (2014) U CA IPM pest management guidelines: floriculture and ornamental nurseries. UC ANR Publ 3392 Nematodes Subbarao KV, Koike ST, Hubbard JC (1996) Effect of deep plowing on the distribution and density of Sclerotinia minor sclerotia and lettuce drop incidence. Plant Dis 80:28–33 Thines M, Zipper R, Schauffele D, Spring O (2006) Characteristics of Pustula tragopogonis (syn. Albugo tragopogonis) newly occurring on cultivated sunflower in Germany. J Phytopathol 154:88–92 Thompson JP, Owen KJ, Stirling GR, Bell MJ (2008) Root-lesion (Pratylenchus thornei and P. neglectus): a review of recent progress in managing a significant pest of grain crops in northern Australia. Aust Plant Pathol 37:235–242 Thompson SM, Tan YP, Young AJ, Neate SM, Aitken EAB, Shivas RG (2011) Stem cankers on sunflower (Helianthus annuus) in Australia reveal a complex of pathogenic Diaporthe (Phomopsis) species. Persoonia 27:80–89 Thompson SM, Tan YP, Shivas RG, Neate SM, Morin L, Bissett A, Aitken EAB (2015) Green and brown bridges between weeds and crops reveal novel Diaporthe species in Australia. Persoonia 35:39–49 Trimboli D, Fahy PC, Baker KF (1978) Apical chlorosis and leaf spot of Tagetes spp. caused by Pseudomonas tagetis Hellmers. Aust J Agric Res 29:831–839 Tzortzkzkis EA, Anastasiadis AI, Simoglou KB, Cantalapiedra-Navarrete C, Palomares-Rius JE, Castillo P (2014) First report of the root-knot nematode, Meloidogyne hispanica, infecting sunflower in Greece. Plant Dis 98:703 Valenzuela-Soto JH, Maldonado-Bonilla LD, Hernandez-Guzma G, Rincon-Enriquez G, MartınezGallardo NA, Ramırez-Chavez E, Cisneros-Hernandez I, Hernandez-Flores JL, Delano-Frier JP (2015) Infection by a coronatine-producing strain of Pectobacterium cacticidum isolated from sunflower plants in Mexico is characterized by soft rot and chlorosis. J Gen Plant Pathol 81:368–381

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Van Wyk PS, Jones BL, Viljoen A, Rong IH (1995) Early lodging, a novel manifestation of Albugo tragopogonis infection on sunflower in South Africa. Helia 18:83–90 Viljoen A, van Wyk PS, Jooste WJ (1999) Occurrence of the white rust pathogen, Albugo tragopogonis, in seed of sunflower. Plant Dis 83:77 Walters SA, Barker KR (1994) Current distribution of five Meloidogyne species in the United States. Plant Dis 78:772–774 Whipker J, Dasoju S, McCall I (1998) Guide to successful pot sunflower production. NC State Coop Ext Ser Hort Inf Leaflet 562 Wisler GC (1984) Bidens mottle virus of bedding plants. FL Dept Agric Consumer Serv. PP Circ 262 Wu HC, Wu WS (2003) Sporulation, pathogenicity and chemical control of Alternaria protenta, a new seedborne pathogen of sunflower. Aust Plant Pathol 32:309–312 Yang SM, Owen PT (1982) Symptomatology and detection of Macrophomina phaseolina in sunflower plants parasitized by Cylindrocopturus adspersus larvae. Phytopathology 72:819–821 Zazzerini A, Tosi L, Vicente PM (1997) First report of root-knot nematodes (Meloidogyne spp.) on sunflowers in Mozambique. Plant Dis 81:1333

Diseases of Zinnia

28

Dorota Szopińska

Abstract

Modern garden zinnias are widely cultivated ornamentals produced for cut flowers and flowerbeds. Diseases often reduce the quantity and quality of zinnia flowers, causing their production to be unprofitable. Three major, widespread diseases occurring on zinnias are: Alternaria blight (Alternaria zinniae), powdery mildew (Golovinomyces cichoracearum, formerly Erysiphe cichoracearum), and bacterial leaf spot (Xanthomonas campestris pv. zinniae). Other pathogens, such as Botrytis cinerea, Cercospora zinniae, Fusarium spp., Rhizoctonia solani, aster yellows phytoplasma, and several viruses may also be harmful to zinnias; however, their economic impact on zinnia production is less significant. Various management strategies for zinnia diseases currently studied include chemical, biological, and physical control and integration of these methods. Keywords

Alternaria zinniae • Golovinomyces cichoracearum • Xanthomonas campestris pv. zinniae • Integrated management

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Alternaria Blight (Alternaria zinniae Pape) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Botrytis Blight, Gray Mold, Stem Canker (Botrytis cinerea Pers ex Pers) . . . . . . . . . 2.3 Cercospora Leaf Spot (Cercospora zinniae Ellis & G. Martin) . . . . . . . . . . . . . . . . . . . . 2.4 Fusarium Root Rot (Fusarium spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Pythium Root Rot (Pythium spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 Phytophthora Root and Crown Rot (Phytophthora spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . .

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D. Szopińska (*) Department of Phytopathology, Seed Science and Technology, Poznań University of Life Sciences, Poznań, Poland e-mail: [email protected] # Springer International Publishing AG 2018 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florists’ Crops Diseases, Handbook of Plant Disease Management, https://doi.org/10.1007/978-3-319-39670-5_28

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2.7

Powdery Mildew [Golovinomyces cichoracearum (DC.) V.P. Heluta (Formerly Erysiphe cichoracearum DC. ex Merat); Euoidium sp.] . . . . . . . . . . . . . . . . 2.8 Rhizoctonia Root Rot [Rhizoctonia solani Kühn (Teleomorph: Thanatephorus cucumeris (A.B. Frank) Donk)] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.9 Sclerotinia Stem Rot [Sclerotinia sclerotiorum (Lib.) de Bary] . . . . . . . . . . . . . . . . . . . . 2.10 Sclerotium Stem Blight, Southern Blight [Sclerotium rolfsii Sac. (Teleomorph: Athelia rolfsii (Curzi) C.C. Tu & Kimbr.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Bacterial and Phytoplasma Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Apical Chlorosis [Pseudomonas syringae pv. tagetis (Hellmers) Young, Dye & Wilkie, syn. Pseudomonas tagetis (Hellmers)] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Bacterial Leaf Spot (Xanthomonas campestris pv. zinniae Hopkins and Dawson) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Ralstonia Wilt (Ralstonia solanacearum (Smith 1896) Yabuuchi et al. 1996, comb. nov.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4 Aster Yellows (“Candidatus Phytoplasma Asteris”) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Viral Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Bromoviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Bunyaviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Caulimoviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4 Closteroviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5 Comoviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6 Geminiviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7 Potyviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.8 Other Viruses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Nematode Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 Angular Spots on the Leaves (Aphelenchoides spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2 Root Knot (Meloidogyne spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1

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Introduction

The genus Zinnia (family Asteraceae) consists of 19 species of annual or perennial plants originating in North and South America and particularly in Mexico (Spooner et al. 1991; Stimart and Boyle 2007). Among them common zinnia (Zinnia elegans Jacq., syn. Zinnia violacea Cav.) is the most widely cultivated species. Variation in ray floret morphology and color, plant height, and habit has led to development of many cultivars useful for landscape borders or cut flowers. The other two commonly planted species, Zinnia angustifolia Kunth (syn. Zinnia linearis Benth.) and Zinnia haageana Regel, are less conspicuous, characterized by a large number of small inflorescences and often with single whorl of ray florets (Linderman and Ewart 1990; Spooner et al. 1991; Stevens et al. 1993). The common zinnia is highly susceptible to three pathogens: Alternaria zinniae Pape (Alternaria blight), Golovinomyces cichoracearum (DC.) V.P. Heluta (powdery mildew), and Xanthomonas campestris pv. zinniae Hopkins and Dawson (bacterial leaf and flower spot), which may cause severe epiphytotics in zinnia, resulting in plant losses, decrease of ornamental value, or both. Zinnia angustifolia on the other hand is highly resistant to these pathogens and has been used as a source of

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resistance genes for Zinnia breeding programs (Boyle and Wick 1996; Jones and Strider 1979; Spooner et al. 1991). Management of the most important zinnia diseases is presented in this chapter. General management strategies, especially referring to widespread pathogens causing diseases of numerous crops, may be found in the introductory chapters.

2

Fungal and Fungus-Like Diseases

2.1

Alternaria Blight (Alternaria zinniae Pape)

Geographic occurrence and impact. The pathogen is widely distributed. The disease has been reported in North and South America (Dimock and Osborn 1942; Gombert et al. 2001; Judd 1976; Palacios et al. 1991), Europe (Beaumont et al. 1958; Christova et al. 1964; Dias and Lucas 1978; Gambogi et al. 1976; Imre 1974; Łacicowa et al. 1979, 1991; Mowsiesjan 1976; Szopińska 2013, 2014a, b; Szopińska et al. 2012), Asia (Rao 1971; Wu and Yang 1992), and New Zealand (Dingley and Brien 1956). Symptoms/signs. The most conspicuous symptom of Alternaria blight is spotting of the foliage. Characteristic reddish-brown spots, sometimes with grayish-white centers, may also appear on stems and blossoms. The spots on the leaves are first circular and vary in size from 2 to 10 mm in diameter but soon enlarge and become irregular. The affected leaves become brown and desiccated. On blossoms and stems, the spots are relatively smaller, rarely more than 2 mm across. On the ray flowers, sporulation on the spots is often profuse and secondary infection may be abundant leading to darkening and withering of petals. On stems, internodal spots frequently show elongation and are usually superficial. Nodal lesions do not remain superficial and the distal parts of the affected stems may be killed by girdling at the nodes. Dark brown to black cankers with sunken centers may also appear near the soil line. Infected roots may turn dark gray, rot, and slough off resulting in wilting and death of the plant. Seedlings may also wilt and collapse (damp-off) (Fig. 1a, b). Biology and epidemiology. Alternaria spp. usually overseason as mycelium or spores in plant debris and seeds. Fungal sporulation and infection are enhanced by moderate temperatures. In the case of A. zinniae, the optimum temperatures for conidial germination in vitro and plant infection are 20–24  C and 22–26  C, respectively. Free moist is required for infection, but the pathogen survives better under dry conditions. The fungus can persist in the seeds for up to 6–7 years, although in plant debris usually not longer than 2 years (Rotem 1994). Seeds are a very important source of inoculum for Alternaria blight. Deeply seated, severe seed infections may lead to preemergence death of seedlings, while superficial seed infections usually cause diseases on plants after emergence (Gambogi et al. 1976).

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Fig. 1 Alternaria blight symptoms: first spots (left), irregular patches (right) (D. Szopińska # 2017. All Rights Reserved.)

Management • Cultural practices – The seeds and transplants used for propagation should to be free of A. zinniae. Rotation to nonsusceptible crops for 2 years, control of susceptible weeds, plant spacing ensuring good air circulation, and proper time and type of irrigation are the most important cultural control measures. Overhead irrigation as well as late afternoon and evening irrigation prolong the time during which the plants remain wet, and therefore favor spread of the disease (Gombert 1998). • Sanitation – Crop debris from the planting area should to be removed and destroyed after harvest. • Physical treatment – To eradicate the pathogen, the seeds can be treated with hot water at 51.5  C for 20 min (Beaumont et al. 1958; Lamboy and Call 2001). A high risk of germination percentage reduction, especially if older seeds are treated, is a reason why this approach is not widely use by the ornamental seed industry (Daughtrey and Benson 2005). • Fungicides – The plants should be treated with protective fungicides at regular intervals (7–14 days), especially during warm wet weather or during warm dry weather if overhead irrigation is applied. Fungicides which are known to control Alternaria diseases include mancozeb, iprodione, chlorothalonil, copper, and triflumizole (Hagan 2009). • Organic treatment – Szopińska (2013) observed that treating zinnia seeds with 1.0% and 2.5% lactic acid for 30 min decreased the percentage of seeds infected

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with A. zinniae from 56.0% to 33.0% and 23.0%, respectively. Significant reduction of zinnia seed infestation with this pathogen was also observed if the seeds were treated with 6% hydrogen peroxide (H2O2) for 60 min and 12% H2O2 for 20 min. These treatments decreased seed infestation from 76% to 28.0% and 27.5%, respectively (Szopińska 2014b). • Resistance – All Z. elegans selections are moderately to highly susceptible to Alternaria blight (Gombert et al. 2001), while Z. angustifolia is highly resistant to this disease (Hagan 2009). Therefore, colchicine-induced amphiploids have been developed in breeding zinnia plants resistant to A. zinniae (Terry-Lewandowski and Stimart 1983, 1985; Terry-Lewandowski et al. 1984). A new species of Zinnia marylandica, an artificial hybrid between Z. angustifolia and Z. elegans, was described and illustrated by Spooner et al. in 1991. Plants of this species exhibit high levels of resistance to A. zinniae and G. cichoracearum and moderate to high levels of resistance to X. campestris pv. zinniae (Terry-Lewandowski and Stimart 1983).

2.2

Botrytis Blight, Gray Mold, Stem Canker (Botrytis cinerea Pers ex Pers)

Geographic occurrence and impact. The pathogen is distributed worldwide and affects over 200 plant species in temperate and subtropical regions (Williamson et al. 2007). Botrytis cinerea has been reported on zinnias in Canada, Great Britain, Poland, the USA, and Venezuela (Bolton 1976; Kiecana and Mielniczuk 2010; Moore 1959; Palacios et al. 1991; Ruhl et al. 1987). Symptoms/signs. Large areas of petals, leaves, and stems turn brown. The infected plant parts develop a dusty gray mycelium during humid conditions (Fig. 2a, b).

Fig. 2 Botrytis blight symptoms/signs: leaf blight (left) flower and stem blight with profuse sporulation (right) (D. Szopińska # 2017. All Rights Reserved.)

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Biology and epidemiology. The pathogen can survive as mycelia and conidia or for extended periods as sclerotia in crop debris (Williamson et al. 2007). Botrytis cinerea produces tremendous quantities of airborne spores, which are moved around by wind or splashing water onto blossoms or young leaves. Cool temperatures (7–15  C) and high humidity (93%) favor spore germination; however, the disease may occur within wide range of temperatures. The pathogen invades host plant tissues directly or through natural openings or wounds (Daughtrey et al. 2000). Management • Cultural practices – Crucial for controlling B. cinerea is reducing free water on plant surfaces and lowering humidity (Daughtrey and Benson 2005). • Sanitation – Strict sanitation is required to prevent the disease from spreading. The dead or dying tissue from the plants and from the soil surface as well as old blossoms and leaves should to be removed and destroyed. • Fungicides and biocontrols – There is a high risk of resistance arising in B. cinerea if a chemical product is applied repeatedly. Since 1980s, there have been numerous reports about the resistance of B. cinerea to benzimidazoles (e.g., benomyl, carbendazim, thiophanate-methyl) and dicarboximides (e.g., iprodione, procymidone, vinclozolin) (LaMondia and Douglas 1997; Northover and Matteoni 1986; Vali and Moorman 1992; Yourman and Jeffers 1999). This phenomenon has also been recorded to a lesser extent in the case of the anilinopyrimidines (e.g., cyprodinil, pyrimethanil), the hydroxyanilide fenhexamid, and some multisite fungicides and fluazinam (Leroux 2004; Leroux et al. 2010). Therefore, application of multisite fungicides such as chlorothalonil, copper hydroxide, copper sulfate pentahydrate, and mancozeb as well as mixed spray programs have been advised (Daughtrey et al. 2000; Williamson et al. 2007). Fungicide resistance has also led to the intensive evaluation of microbial antagonists for control of gray mold. The greatest potential for control of B. cinerea has been shown by the fungi Trichoderma harzianum, Gliocladium roseum, and Ulocladium oudemansii; the bacteria Bacillus subtilis, Pseudomonas syringae, and Streptomyces griseoviridis; and the yeasts Candida oleophila and Pichia pastoris (Elad and Stewart 2004).

2.3

Cercospora Leaf Spot (Cercospora zinniae Ellis & G. Martin)

Geographic occurrence and impact. The pathogen has been reported on Z. elegans in India, Jamaica, Mauritius, Thailand, the USA, and Venezuela (Felix 1960; Palacios et al. 1991; Pande 1975; Pereira 2008; Phengsintham et al. 2013; Wehlburg 1969). Symptoms/signs. Cercospora leaf spot closely resembles Alternaria leaf spot, and symptoms of both diseases often occur together on the same leaf. Leaf symptoms include relatively large, round, reddish brown or dark purple spots, with light gray

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Fig. 3 Cercospora leaf spot symptoms (R.J. McGovern # 2017. All Rights Reserved.)

centers, which become ashen gray, papery, and brittle. Severely infected leaves become brown and desiccated. The dead tissues often crack and tear (Fig. 3). Biology and epidemiology. The fungus is favored by warm weather and high humidity. Conidia of C. zinniae germinate within 6 h after inoculation of the zinnia leaf surface. Germ tubes penetrate leaf tissue through the stomatal pores. Spots on the leaves appear 5–7 days after inoculation. Development of the first conidia begins on the spots at 9 days after inoculation. The spores are dispersed in the field by wind, insects, splashing water, and humans (Pereira 2008). Management • Cultural practices – Avoiding overhead irrigation and proper plant spacing are important to prevent spread of this pathogen. Crop rotation with hosts not affected by Cercospora zinniae is recommended. • Fungicides – Under field conditions mancozeb (0.2%) followed by carbendazim (0.1%) and chlorothalonil (0.2%) reduced Cercospora leaf spot percent disease index from 53.62 (water spray) to 5.94, 13.36, and 18.26, respectively (Yadahalli et al. 1994). Similar results were recorded by Raghavendra Rao and Chacko (1986) and Madhumeeta and Shyam (1989).

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Fusarium Root Rot (Fusarium spp.)

Geographic occurrence and impact. Several species of Fusarium, e.g., F. avenaceum, F. equiseti, F. culmorum, F. oxysporum, and F. solani have been isolated from diseased zinnia plants in Poland. Fusarium equiseti showed the highest pathogenicity towards zinnia seedlings (Kiecana and Mielniczuk 2010; Łacicowa et al. 1979). Symptoms/signs. Disease symptoms include reduction of growth, necrotic streaks on the hypocotyls, necrotic spots with yellow halos limited to the edge of leaves, and root rotting of severely infected plants. Fusarium spp. may also cause pre- and postemergence damping-off. Biology and epidemiology. Fusarium root rot is favored by high air and soil temperatures that exceed 23  C. At lower soil temperatures, disease symptoms may not occur, and even though the plant is diseased, it may appear to be healthy. The pathogens survive in the soil associated with plant debris but also directly in the soil as mycelium, spores, and, except in the case of F. avenaceum, as chlamydospores (especially in cooler regions). They spread by means of water and contaminated equipment. Growing mycelia invade the root system of healthy plants through wounds or at the point of formation of lateral roots. Management. The fungi are so persistent and widespread that control measures, such as crop rotation and soil sterilization, are of limited value. Some biological methods were proposed but chemical control is currently the only practical way to manage Fusarium spp. effectively in zinnia.

2.5

Pythium Root Rot (Pythium spp.)

Geographic occurrence and impact. Pythium spp. belongs to the kingdom Chromista, class Oomycetes. These pathogens have been reported on zinnia in the USA (Lewis et al. 1996; Lumsden and Locke 1989). Symptoms/signs. The pathogens attack roots which become dark and rot. Infected plants wilt very quickly. Pythium spp. may also cause pre- and postemergence damping-off. Biology and epidemiology. The pathogens from genus Pythium are distributed as zoospores with water, and long-term survival is enabled by chlamydospores and sexually produced oospores. Wet soil with poor aeration favors Pythium growth and development. Pythium spp. have wide range of hosts worldwide.

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Management • Biocontrol – Gliocladium virens controlled damping-off of zinnia caused by P. ultimum and R. solani in a nonsterile soilless mix. Disease control efficacy lasted for at least 2 m when G. virens was introduced with the pathogen inoculum and the mix was planted at intervals with zinnia seeds (Lumsden and Locke 1989). Alginate prills, formulated with the isolate GI-22 of G. virens and various food bases (wheat bran, corn cobs, peanut hulls, soy fiber, castor pomace, and chitin) also significantly reduced the damping-off of zinnia in a soilless mix caused by these pathogens (Lewis et al. 1996).

2.6

Phytophthora Root and Crown Rot (Phytophthora spp.)

Geographic occurrence and impact. Phytophthora spp. also belong to the kingdom Chromista and class Oomycetes. These pathogens infest a wide range of crops, including zinnia (Moore 1959). Symptoms/signs. Symptoms include rot of roots and lower stems and damping-off of seedlings. Infected plants wilt and die. Biology and epidemiology. Phytophthora spp. are disseminated as zoospores with water through irrigation and rain. Wet soil favor incidence of this disease. Long-term survival is by chlamydospores and sexually produced oospores. Management. Susceptible crops should be planted in pathogen free soil. Crop rotation with resistant or less susceptible crops is advised.

2.7

Powdery Mildew [Golovinomyces cichoracearum (DC.) V.P. Heluta (Formerly Erysiphe cichoracearum DC. ex Merat); Euoidium sp.]

Geographic occurrence and impact. Powdery mildews are common and widespread diseases, usually more abundant in semiarid regions than in areas of high rainfall (Kamp 1985). The disease caused by G. cichoracearum has been reported on zinnia in Egypt, India, Japan, New Zealand, the Republic of South Africa, the USA, and Venezuela (Amano 1986; Baker and Locke 1946; Dingley 1965; Gombert et al. 2001; Hegazi and El-Kot 2010a, b; Husain and Akram 1995; Kamp 1985; Linderman and Ewart 1990; Mir et al. 2012; Palacios et al. 1991; Ruhl et al. 1987; Schmitt 1955; Terry-Lewandowski and Stimart 1983; Tesfagiorgis 2008). A powdery mildew caused by Euoidium sp. has been reported on zinnia in Japan (Hoshi et al. 2013). High incidence and severity of powdery mildew on zinnias in some

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regions of Asia and North America appears to be a major factor for the declining commercial value of these plants (Mir et al. 2012; Spooner et al. 1991). Symptoms/signs. White fungal growth forms on the upper surface of leaves and may form on flower petals. Leaves die from base of plant upwards. Affected plants lose vigor and cease growth. Biology and epidemiology. The fungi causing powdery mildews are obligate parasites. The mycelium of most powdery mildew fungi grows only on the surface and produces usually short conidiophores. When environmental conditions become unfavorable, the fungus may produce sexual structures called cleistothecia, containing one or a few asci. The spores can be released, germinate, and cause infection even if there is no film of water on the plant surface as long as the relative humidity in the air is moderately high (Agrios 2005). Baker and Locke (1946) revealed that seed transmission of the pathogen is possible; however, primary inocula of G. cichoracearum for zinnia infections are likely to come from adjacent wild or cultivated Asteraceae (Boyle and Wick 1996).

Management • Cultural practices – For powdery mildew, unlike Alternaria blight and bacterial leaf and flower spots, drip irrigation should to be avoided, because it may produce greater powdery mildew severity than overhead irrigation (Gombert 1998). • Fungicides – Currently, the only practical method for controlling powdery mildew on zinnias in commercial production is by applying synthetic chemicals. Fungicide applications must be started when the first sign of the white powdery colonies on the lower leaves appears and should to be reapplied according to the specifications on the fungicide label. Such fungicides as tebuconazole, chlorothalonil, triforine, propioconazole may be use to control powdery mildew (Hagan 2009). Kamp (1985) demonstrated that the use of a polymer-based antitranspirant significantly reduced powdery mildew on Z. elegans. The author suggested that the epidermal coating may interrupt pathogen development on the leaf surface by repelling the film of free water from the leaves and disorientation of pathogen germ tubes by changes in surface topology. Locke et al. (2006), on the other hand, observed delay of expression of powdery mildew on zinnia grown hydroponically in Hoagland’s solution fortified with silicon. Tesfagiorgis (2008) found that zinnia plants have the ability to accumulate high levels of silicon in the leaves, which may support an active mechanism of defense within the host plant. • Biological control and organic treatment – Hegazi and El-Kot (2010a) sprayed zinnia plants four times at 1 week intervals with 25 and 50% culture filtrates of Trichoderma harzianum, Epicoccum sp., Streptomyces endus, as well as two plant extracts, miswak (Salvadora persica) and henna (Lawsonia inermis) to

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control powdery mildew. All applied treatments significantly decreased disease incidence and severity; however, Epicoccum sp. and T. harzianum at 50% concentration followed by henna and miswak extracts inhibited spread of disease to the highest extent. The authors also observed a significant decrease of powdery mildew incidence and severity as well as improvement of plant growth parameters (i.e., plant height, number of branches per plant, leaf area, fresh and dry weights of shoots and roots, root length) when zinnia plants were treated with cinnamon, clove, and ginger oils at a concentration of 0.1 ppm (Hegazi and El-Kot 2010b). In organic farming use of preparations based on copper, organic, and mineral oils (e.g., neem oil, paraffinic oil) and Bacillus subtilis strain QST 713 are recommended (Hagan 2009). • Resistance – There are only a few Z. elegans cultivars that are resistant to G. cichoracearum (Gombert et al. 2001). Artificial hybrids between Z. angustifolia and Z. elegans, on the other hand, are highly resistant or immune to powdery mildew (Terry-Lewandowski and Stimart 1983, 1985; TerryLewandowski et al. 1984). This disease has rarely been noted on the Crystal and Star series as well as the hybrid Profusion series zinnias (Hagan 2009).

2.8

Rhizoctonia Root Rot [Rhizoctonia solani Kühn (Teleomorph: Thanatephorus cucumeris (A.B. Frank) Donk)]

Geographic occurrence and impact. Rhizoctonia diseases occur worldwide and cause losses in almost all crops. Rhizoctonia root, crown, and stem rot have been documented in zinnia in Australia, Poland, and the USA (Łacicowa et al. 1979; Lumsden and Locke 1989; Schisler et al. 1994). Symptoms/signs. The first sign of infection in seedlings is damping off. In older plants, the infection begins on the stem as dark-brown lesion below the soil line and then a cottony, white mycelium grows upward in the plant. Root discoloration and rot can also occur. Numerous small black sclerotia may be produced on all infected tissues. Biology and epidemiology. The pathogen overwinters as mycelium or sclerotia in the soil and in some species, including zinnia, may be carried in the seeds. Very young seedlings may be killed before or after emergence. The fungus spreads with rain and irrigation, tools contaminated with soil, and with infected seeds. High temperatures (about 35  C) and moderately wet soil favor infection. Management • Cultural practices – Disease-free seeds should be planted. • Biocontrol – Gliocladium virens controlled damping-off of zinnia caused by Pythium ultimum or R. solani in a nonsterile soilless mix (Lewis et al. 1996; Lumsden and Locke 1989).

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Sclerotinia Stem Rot [Sclerotinia sclerotiorum (Lib.) de Bary]

Geographic occurrence and impact. The pathogen has been reported on zinnias in Argentina, Great Britain, Poland, and the USA (Grabowski and Malvick 2015; Kiecana and Mielniczuk 2010; Kiehr et al. 2010; Łacicowa et al. 1979; Moore 1959). Symptoms/signs. Sclerotinia sclerotiorum causes crown rot, bulb rot, stem rot, wilt, and death of a wide variety of plants. The typical early sign of Sclerotinia disease on zinnia is the appearance on the infected plant of a white fluffy mycelium in which variable sized sclerotia soon develop. The mycelium first develops on the base of stem. When the fungal infection develops and the stem rots, the foliage above the lesion wilts and dies. The sclerotia of S. sclerotiorum may be also formed inside the stem. Moreover, the pathogen is frequently responsible for damping-off of zinnia seedlings (Łacicowa et al. 1979). Biology and epidemiology. The pathogen overwinters as sclerotia in the soil and as sclerotia and mycelium on or within infected tissues of dead plants. The sclerotia germinate in the spring or early summer producing apothecia which discharge a large number of ascospores into the air. The spores reaching available sources of food (e.g., old plant parts) start to germinate and cause infection. Infection may also start from the soil when sclerotia produce mycelial strands which invade young plants (Agrios 2005). The pathogen may cause pre- and postemergence damping-off. Management • Cultural practices – Susceptible cultivars should be planted in well-drained soil. Weeds as a potential hosts should to be removed. Because sclerotia may survive in the soil, at least a 3-year rotation with non-susceptible crops is recommended.

2.10

Sclerotium Stem Blight, Southern Blight [Sclerotium rolfsii Sac. (Teleomorph: Athelia rolfsii (Curzi) C.C. Tu & Kimbr.)

Geographic occurrence and impact. Sclerotium rolfsii has been reported on zinnia in India, Japan, and New Zealand (Boesewinkel 1977; Ishii and Abiko 1997; Ramakrishnan 1930; Thakur 1969). Symptoms/signs. The pathogen attacks the stem bases, leaves turn yellow, the plant initially wilts, and then dies. A large number of white turning tan, mustard seed-sized sclerotia are formed on the infected tissue. Biology and epidemiology. Wet and warm weather favor disease progress. Optimal temperatures for fungal growth and disease development are 25–35 oC. High relative humidity is required for germination of sclerotia (Mordue 1974). Sclerotia in the soil are the main source of this disease.

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Management • Cultural practices – Management strategies include a 3- to 4-year rotation with non-susceptible crops, raising the soil pH, and removal of crop debris (Mordue 1974). Additional fungal pathogens of zinnia. The following fungi have also been reported as pathogenic to zinnia: Alternaria carthami S. Chowdhury – in Taiwan (Wu and Chou 1995) Colletotrichum acutatum J. H. Simmonds – in India (Kulshrestha 1976) Colletotrichum falcatum Went. – in Venezuela (Palacios et al. 1991) Nigrospora sphaerica (Sacc.) E.W. Mason – in the USA (Meepagala et al. 2015) Sclerotinia minor Jagger – in Argentina (Kiehr et al. 2010)

3

Bacterial and Phytoplasma Diseases

3.1

Apical Chlorosis [Pseudomonas syringae pv. tagetis (Hellmers) Young, Dye & Wilkie, syn. Pseudomonas tagetis (Hellmers)]

Geographic occurrence and impact. This disease has been reported on zinnia in Australia and the USA (Gulya et al. 1982; Jutte and Durbin 1979; Trimboli et al. 1978). Symptoms/signs. Leaf spots, sometimes with chlorotic haloes, and apical chlorosis are typical symptoms of this disease (Fig. 4). Biology and epidemiology. Pseudomonas syringae pv. tagetis is a Gram negative aerobic rod. The pathogen overseasons in infected plant debris in the soil. In sunflower, this pathogen is seed-borne, but it is not known if this type of transmission occurs in zinnia. Weeds as well as cultivated crops within the family Asteraceae may provide natural sources of inoculum. Wet weather favors spread of this disease. Fig. 4 Apical chlorosis symptoms caused by Pseudomonas syringae (T.J. Gulya. USDA.)

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Management. A 3-year or longer rotation is recommended to avoid building up large soil populations of the bacterium. Plants should to be watered in a manner that keeps the leaves dry to inhibit spread of this disease.

3.2

Bacterial Leaf Spot (Xanthomonas campestris pv. zinniae Hopkins and Dawson)

Geographic occurrence and impact. The disease has been observed in many countries including Australia, Brazil, Bulgaria, Hungary, India, Italy, Korea, Malawi, Pakistan, Rhodesia, Sierra Leone, and the USA (Akhtar and Khokhar 1988; Bertus and Hayward 1971; Boyle and Wick 1996; Deighton 1957; Dimitrov and Tsoneva 1977; Hopkins and Dowson 1949; Jones and Strider 1979; Myung et al. 2012; Nannizzi 1929; Peregrine and Siddiqi 1972; Rangaswami and Gowda 1963; Robbs 1954; Sahin et al. 2003; Schwarczinger et al. 2008a, b; Sleesman et al. 1973; Strider 1973, 1976, 1979a, b, 1980; Terry-Lewandowski and Stimart 1983). Symptoms/signs. The pathogen can attack all aboveground parts of zinnia plants, causing necrotic lesions of leaves, stems, and flowers. The first symptoms of the disease are small (1–2 mm), dispersed, transparent spots encircled by broad yellowish haloes. Under wet conditions, the lesions slowly enlarge. The spots become angular to irregularly circular and develop a reddish center. The lesions may merge to form irregular dead areas (0.5–1.0 cm long), which often crack as they dry. During very humid weather, small brown spots may appear on the ray flowers. Severely infected flower heads are seriously disfigured and may completely decay (Fig. 5). Biology and epidemiology. The pathogen has been reported to overseason in diseased plant residue in the field and in or on seeds (Strider 1973, 1979a). Infected seeds are known to be the major means of long-distance dispersal and primary inoculum source of the pathogen (Strider 1979b). Xanthomonas campestris pv. zinniae is capable of spreading rapidly through warm, humid, and rainy weather. Fig. 5 Bacterial leaf spot symptoms (R.J. McGovern # 2017. All Rights Reserved.)

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Infected zinnias develop characteristic spots on the leaves within 2 weeks. Jones and Strider (1979) observed that plants at a 29–31/21–23  C (max/min) temperature regime developed the most severe disease symptoms. Management • Cultural practices – The use of pathogen free seeds and transplants has been recommended for control of bacterial leaf spot (Strider 1979a, 1980). Whenever possible, plants should be irrigated in a manner that keeps the leaves dry to inhibit spread of this disease. • Sanitation – Plants with leaf spots should be discarded and diseased plant debris should be removed from the growing area. Washing hands after handling diseased plants or soil is recommended. Handling of wet foliage should be avoided. • Chemical control – Chemical control of bacterial leaf spot on zinnia is difficult if not impossible to obtain. Copper hydroxide may provide some protection from this disease. To be effective, treatments have to be applied on a weekly schedule, beginning when the spots first appear on the leaves (Hagan 2009). Bactericides are only marginally effective in controlling Xanthomonas spp. Strider in 1980 recorded the efficacy of captan as a bactericide for control bacterial leaf spot on zinnia. • Resistance – Most common zinnia cultivars as well as Z. haageana “Persian Carpet”, Z. angustifolia “Pixie Sunshine”, Z. tenuifolia “Red Spider”, and Z. pumilia “Cut and Come Again” are susceptible to X. campestris pv. zinniae (Hagan 2009; Gombert et al. 2001). However, hybrids of Z. elegans and Z. angustifolia exhibit moderate to high levels of resistance to this pathogen (Terry-Lewandowski and Stimart 1983). The Crystal and Star series zinnias are highly resistant or immune to bacterial leaf spots. The Profusion series zinnias are moderately resistant to this disease. Some spotting of the leaves in the lower canopy as well as occasional but unobtrusive leaf death has been seen on Profusion series zinnias in the field (Hagan 2009).

3.3

Ralstonia Wilt (Ralstonia solanacearum (Smith 1896) Yabuuchi et al. 1996, comb. nov.)

Geographic occurrence and impact. Ralstonia solanacearum (formerly Pseudomonas solanacearum) causes a severe and devastating wilt to many economically important crops around the world (Hayward 1991). This disease has been reported on zinnia in Australia, India, and Malaysia (Abdullah 1992; Papdival and Deshpande 1978; Thammakijjawat et al. 2001). Strider et al. (1981) reported that a strain of R. solanacearum isolated from geranium was also pathogenic on zinnia. Ralstonia solanacearum is a pathogen of quarantine importance. Symptoms/signs. Infected young plants die rapidly. On older plants, symptoms appear first on the youngest leaves or one-sided wilting and stunting is observed. Finally plants wilt completely and die. The vascular tissues of stem and roots turn brown (Agrios 2005).

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Biology and epidemiology. According to host range, R. solanacearum strains have been classified into five races (Buddenhagen and Kelman 1964; Hayward 1991). A zinnia strain of R. solanacearum originating in Australia has been classified as race 1 (Thammakijjawat et al. 2001). This race attacks all the solanaceous and many non-solanaceous crops and appears as a sudden wilt. The bacteria overwinter in plant debris and spread in soil with water and then enter plants through wounds made in roots by insects, nematodes, or by handling. Management. There are no chemicals or biological agents to effectively control this bacterium. Infected plants have to be discarded as soon as possible. Pathogenfree propagating plant material is the main way to avoid problems with Ralstonia. Soil and equipment should to be disinfested.

3.4

Aster Yellows (“Candidatus Phytoplasma Asteris”)

Geographic occurrence and impact. This disease has been reported on zinnia in Canada and India (Rao et al. 2012; Singh et al. 2011; Wang and Hiruki 2001). Symptoms/signs. The pathogen causes general yellowing and dwarfing of the plant, abnormal production of shoots (formation of witches’ brooms), and malformation of organs (greening or sterility of flowers, shortening of the internodes). The disease reduces yield and may lead to more or less rapid dieback, decline, and death of the plant. Biology and epidemiology. Taxonomically, phytoplasmas are member of the class Mollicutes and are currently classified within the provisional genus “Candidatus Phytoplasma” based primarily on 16S rDNA sequence analysis (Marcone 2014). The phytoplasmas detected in zinnia have been placed in the subgroup II “Candidatus Phytoplasma asteris” (16SrI-B) (Rao et al. 2012; Wang and Hiruki 2001). Phytoplasmas are prokaryotic organisms that have no cell walls and can rarely be grown on artificial nutrient media. Most plant mollicutes are vectored by leafhoppers (Cicadellidae) but some by planthoppers (Fulgoroidea). Phytoplasmas are generally present in the sap of a small number of sieve tubes. Insects become vectors when feeding on leaves and stems of infected plants. The phytoplasma then multiplies and infects various body organs of insect vectors, a process that can take 1–3 weeks. When infected leafhoppers feed on healthy plants, the phytoplasmas in their salivary glands are injected into the plant phloem and disease symptoms develop in 10–40 day. A leafhopper carrying the aster yellows phytoplasma can continue infecting plants for the rest of its life. Outbreaks of aster yellows may be expected in cool and wet summers, which favor both leafhoppers and phytoplasma development (Agrios 2005). Management. There are currently few effective control strategies for aster yellows. Infected plants should to be promptly removed and destroyed. Insect vectors should to be monitored and controlled systematically, because they are essential for

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dispersal of the pathogen and long-term survival in nature. Management of aster yellows includes chemical and biological control of leafhoppers, repellence and disorientation of insects in the field, and avoidance of plant production near infected crops or alternate hosts.

4

Viral Diseases

4.1

Bromoviridae

Geographic occurrence and impact. This family contains five genera of viruses: Alfamovirus, Bromovirus, Cucumovirus, Ilarvirus, and Oleavirus. Several members of these genera are distributed worldwide. Four viruses from the Bromoviridae have been reported in zinnia: Alfalfa mosaic virus (AMV, genus Alfamovirus) in Israel, Cucumber mosaic virus (CMV, genus: Cucumovirus) in India, Iran, and the USA, Tomato aspermy virus (TAV, genus Cucumovirus), and Asparagus virus 2 (AV2, genus Ilarvirus) in Japan (Fujisawa et al. 1983; Kameya-Iwaki et al. 1996; Nitzany and Cohen 1960; Price 1935; Raj et al. 1997; Shahmohammadi et al. 2015). According to Brunt et al. (1996–), zinnia is also susceptible to the following viruses from this family: Peanut stunt virus from genus Cucumovirus and Elm mottle virus, Humulus japonicus virus, Plum American line pattern virus, Prune dwarf virus, Prunus necrotic ringspot virus, and Tobacco streak virus from genus Ilarvirus. Symptoms/signs. All viruses detected in zinnia produce similar symptoms: leaf mosaic and distortion, stunting, and flowers malformation (Fig. 6). Biology and epidemiology. Aphids (Hemiptera: Aphididae) and thrips (Thysanoptera: Thripidae) are main vectors of Bromoviridae. Aphis gossypii, A. craccivora, A. fabae, and Myzus persicae are responsible for CMV transmission in zinnia (Nitzany and Cohen 1960). All cucumoviruses are transmitted by aphids in

Fig. 6 CMV symptoms (R.J. McGovern # 2017. All Rights Reserved.)

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noncirculative (nonpersistent) manner. They may be also transmitted in some species by seeds or mechanically by handling of the plants in the field or greenhouse (e.g., CMV). Seed transmission by zinnia seeds has been reported only for Asparagus virus 2 (Fujisawa et al. 1983). Management. Controlling insect vectors, avoiding growing near infected crops, and removing weeds that serve as hosts help to control viral diseases. Vector populations may be reduced by chemical and biological methods and various physical barriers. Use of reflective mulches disorients the vectors and protects the plants.

4.2

Bunyaviridae

Geographic occurrence and impact. Tospoviruses make up one genus of the viruses within this family, including several economically important species with an extremely wide host range. The main member of this genus – Tomato spotted wilt virus (TSWV) – has been detected in zinnia in Bulgaria and Mexico (Bakardjieva et al. 1998; Morales-Díaz et al. 2008). Symptoms/signs. Typical symptoms on zinnia are chlorotic rings and spots on the leaves, wilting of the leaves, and plant death. Biology and epidemiology. Tospoviruses are transmitted by thrips (e.g., Frankliniella occidentalis, F. fusca, F. schultzei, Thrips tabaci, T. palmi, etc.). The virus is acquired from infected plants by thrips larvae after at least 30 min of feeding. The thrips transmits the virus to healthy plants for the rest of its life. Management. Refer to the Bromoviridae management section above and introductory (▶ Chap. 4, “Insect Management for Disease Control in Florists’ Crops”).

4.3

Caulimoviridae

Geographic occurrence and impact. Dahlia mosaic virus DMV belonging to the genus Caulimovirus has been reported on zinnia in the USA (Kitajima et al. 1969). Symptoms/signs. The symptoms range from none to occasional chlorotic spots to severe mosaic and stunting. Biology and epidemiology. DMV can be transmitted by 16 different aphid (Hemiptera: Aphididae) species, mechanical inoculation, and through the seed (not recorded for zinnia). Management. Refer to the Bromoviridae management section above and introductory (▶ Chap. 4, “Insect Management for Disease Control in Florists’ Crops”).

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4.4

Diseases of Zinnia

857

Closteroviridae

Geographic occurrence and impact. Two viruses belonging to genus Crinivirus have been reported in zinnia in Taiwan: Tomato chlorosis virus (ToCV) and Tomato infectious chlorosis virus (TICV) (Tsai et al. 2004). According to Brunt et al. (1996), zinnia is also susceptible to two other viruses from this genus: Beet pseudoyellows virus and Lettuce infectious yellows virus. Symptoms/signs. Pronounced yellowing symptoms on the lower leaves of plants, similar to those caused by nitrogen deficiency; the brittleness of the discolored leaves; and occasional upward leaf rolling were observed on infected zinnia plants. Biology and epidemiology. Criniviruses are confined to the phloem and phloem parenchyma cells. TICV is transmitted exclusively by the whitefly Trialeurodes vaporariorum, whereas ToCV is vectored by T. vaporariorum, and another whitefly, Bemisia tabaci. Management. Refer to the Bromoviridae management section above and introductory (▶ Chap. 4, “Insect Management for Disease Control in Florists’ Crops”).

4.5

Comoviridae

Geographic occurrence and impact. Two nepoviruses have been reported on zinnia: Tobacco ringspot virus (TRSV) in Japan and the USA and Tomato black ring virus (TBRV) in Kenya (Iizuka 1973; Kaiser et al. 1978; Moore and McGuire 1968). According to Brunt et al. (1996–), zinnia is also susceptible to the following viruses from this family: Cowpea mosaic virus from genus Comovirus and Arabis mosaic virus, Artichoke vein banding virus*, Caraway latent virus*, Cherry leaf roll virus, Dogwood mosaic virus*, Peach enation virus*, Croton yellow vein mosaic virus, and Soybean crinkle leaf virus from genus Nepovirus. (*Species not recorded currently in the ICTV Master Species List). Symptoms/signs. Common symptoms of plant infection by these viruses are mosaic and ring spot, sometimes accompanied by systemic necrosis. Biology and epidemiology. Tobacco ringspot virus is transmitted by the nematode Xiphinema americanum and also nonspecifically by insects and mites such as: Aphis gossypii, Myzus persicae, Melanopus sp. Epitrix hirtipennis, Thrips tabaci, and Tetranychus sp. Iizuka (1973) reported 5% transmission of TRSV by zinnia seeds. This virus causes systemic infection in susceptible cultivars, moving from infected leaves to the tips of stems and into roots. Movement from roots to leaves is uncommon (Moore and McGuire 1968). Kaiser et al. (1978) detected seed transmission of TBRV in 1 seedling infected out of 4 tested.

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Management. Refer to the Bromoviridae management section above and introductory (▶ Chap. 4, “Insect Management for Disease Control in Florists’ Crops”).

4.6

Geminiviridae

Geographic occurrence and impact. These viruses are responsible for a significant amount of crop damage worldwide. Four species in genus Begomovirus were detected in zinnia: Ageratum enation virus (AEV) in India, Ageratum yellow vein China virus (AYVCNV) and Alternanthera yellow vein virus (AlYVV) in Vietnam, and Tomato yellow leaf curl virus (TYLCV) in China. Two other species Zinnia leaf curl virus (ZLCV) identified in India and Zinnia mosaic virus recorded in India and Spain are not recorded currently in ICTV Master Species List and their taxonomic position is unsure (Ha et al. 2008; Huertos 1953; Jabri et al. 1985; Kumar et al. 2010; Li et al. 2013, 2014); Maritan et al. 2004; Panday and Tiwari 2012; Verma and Singh 1973). Zinnia plants are also susceptible to the following viruses from this family: Beet curly top virus belonging to genus Becurtovirus and two species from genus Begomovirus: Croton yellow vein mosaic virus and Soybean crinkle leaf virus (Brunt et al. 1996–). Symptoms/signs. Symptoms caused by AEV, AYVCNV, TYLCV, and ZLCV have been described as leaf curling, yellow mosaic, and stunting (Figs. 7 and 8). Zinnia mosaic symptoms have been characterized as a strong mosaic, stunting, decrease of the number of flowers, and color breaking. Biology and epidemiology. Begomoviruses are transmitted circulatively and nonpropagatively by the whitefly Bemisia tabaci. Once the virus is acquired, the whitefly remains viruliferous for life. Management. Refer to the Bromoviridae management section above and introductory ▶ Chap. 4, “Insect Management for Disease Control in Florists’ Crops”).

4.7

Potyviridae

Geographic occurrence and impact. Potyviruses include many of the viruses causing some of the most severe and economically important plant diseases. Five species belonging to this genus have been identified in zinnia: Bidens mottle virus (BiMoV) and Sunflower mosaic virus (SuMV) in the USA, Sunflower chlorotic mottle virus (SuCMoV) in Brazil, and Vanilla distortion mosaic virus (VDMV) and Zinnia mild mottle virus* in India (Balaji et al. 2014; Gulya et al. 2002; Logan et al. 1984; Maritan et al. 2004; Padma et al. 1972, 1974).

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Fig. 7 Ageratum enation virus symptoms (Kumar et al. 2010)

Fig. 8 Tomato yellow leaf curl virus symptoms (Li et al. 2014)

According to Brunt et al. (1996–), zinnia is also susceptible to many other viruses from this family: Sweet potato mild mottle virus from genus Ipomovirus and Bean yellow mosaic virus, Beet mosaic virus, Bidens mosaic virus, Carrot thin leaf virus, Celery mosaic virus, Clover yellow vein virus, Lettuce mosaic virus, Nasturtium

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mosaic virus*, Patchouli mosaic virus, Pea seed-borne mosaic virus, Pepper veinal mottle virus, Plum pox virus, Primula mosaic virus, Tobacco etch virus, Tropaeolum 2 virus, Turnip mosaic virus, Watermelon mosaic 2 virus, and Wisteria vein mosaic virus belonging to genus Potyvirus. (*Species not recorded currently in ICTV Master Species List). Symptoms/signs. Diseases caused by potyviruses appear primarily as mosaics, mottling, chlorotic rings or color breaking on foliage, flowers, and stems. Many of these viruses may cause severe stunting of young plants, stem malformations, and necroses of various tissues. Biology and epidemiology. Potyviruses are spread by aphids (Hemiptera: Aphididae) in a noncirculative (nonpersistent) manner and several are transmitted through seeds (not recorded for zinnia). These viruses overseason in perennial cultivated and weed hosts. Management. Refer to the Bromoviridae management section above and introductory ▶ Chap. 4, “Insect Management for Disease Control in Florists’ Crops”).

4.8

Other Viruses

According to the VIDE Database (Brunt et al. 1996–), zinnia is also susceptible to the following viruses: Family Flexiviridae, genus Carlavirus: Artichoke latent S virus*, Butterbur mosaic virus*, Cassia mild mosaic virus*, Pea streak virus*, Poplar mosaic virus; genus Potexvirus: Asparagus 3 virus, Cymbidium mosaic virus, Foxtail mosaic virus, Plantain X virus Family Luteoviridae, genus Luteovirus: Beet mild yellowing virus, Beet western yellows virus, Subterranean clover red leaf virus Family Rhabdoviridae, genus Nucleorhabdovirus: Sonchus yellow net virus Family Secoviridae: Strawberry latent ringspot virus Family Tombusviridae, genus Carmovirus: Cucumber leaf spot virus, Galinsoga mosaic virus; genus Dianthovirus: Carnation ringspot virus; genus Necrovirus: Lisianthus necrosis virus*, Tobacco necrosis virus; genus Tombusvirus: Cymbidium ringspot virus, Pepper Moroccan virus, Tomato bushy stunt virus Family Virgaviridae, genus Tobamovirus: Maracuja mosaic virus, Odontoglossum ringspot virus, Ribgrass mosaic virus Genus Sobemovirus: Lucerne transient streak virus Genus Ourmiavirus: Ourmia melon virus (*Species not recorded currently in ICTV Master Species List).

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Diseases of Zinnia

5

Nematode Diseases

5.1

Angular Spots on the Leaves (Aphelenchoides spp.)

861

Geographic occurrence and impact. Foliar nematodes are widespread pathogens of ornamentals grown in greenhouses, nurseries, and in the field. There are three species of economic importance: A. besseyi, A. fragariae, and A. ritzemabosi. Aphelenchoides besseyi prefers warmer climates, while A. fragariae and A. ritzemabosi are often found in temperate zones (Kohl 2011). All three species can cause angular spots on Z. elegans leaves; however, A. ritzemabosi has been the most prevalent. Aphelenchoides spp. have been reported on zinnias in India, New Zealand, Poland, and the USA (Boesewinkel 1977; Crossman and Christie 1936; Gill and Sharma 1979; Gill and Uppal 1979; Gill and Walia 1980; Kohl 2011; Madej et al. 2000). Symptoms/signs. Infected plants appear stunted and dwarfed. Infected leaves become shrunken, occasionally accompanied by discoloration, with blotches and chlorotic patches turning into brown and white-yellow areas limited by the veins. Biology and epidemiology. Water is necessary for the movement and dispersal of foliar nematodes; however, spread from the direct contact of an infected leaf with uninfected plant tissue is possible. Adult and fourth-stage juvenile Aphelenchoides spp. enter leaf tissue through stomata on the leaf undersurface where feeding and reproduction occurs. The nematodes feed by piercing neighboring cells with their stylets. Eggs are laid within healthy, green sections of the leaf tissues. The endoparasitic feeding results in the disintegration of the spongy parenchyma and palisade cells; however, nematodes also feed ectoparasitically on stems, buds, and flowers. Males are necessary for reproduction. Fertilized females are able to lay eggs even after emergence from months of dormancy in an anhydrobiotic state. Each female lays approximately 32 eggs, which hatch in 4 days. The second stage juveniles reach reproductive maturity in 6–7 days. The life cycle of A. ritzemabosi can be completed in 14 days, with 5 days for embryonic development and another 5 days for maturation. Adults and fourth-stage juveniles are able to overwinter in an anhydrobiotic state within desiccated plant tissue (dried leaves, dormant buds, but not in plant roots) and can survive for several months to up to 3 years. Low air temperature and high relative humidity increase nematodes populations in leaves (Kohl 2011). Management. Foliar sprays of quinalphos and methyl parathion at 0.05% applied five times at 10-day intervals effectively controlled A. ritzamabosi on Z. elegans, reducing both the symptoms of infestation and the final nematode population (Gill and Walia 1980). However, due to environmental concerns and toxicity, these nematicides are no longer available. Many other nematicides were proposed (e.g., chlorfenapyr); however, the most effective chemical controls will be those that are truly systemic within the plants (Kohl 2011). Integration of chemical methods with

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cultural practices such as sanitation, removal of crop debris from the planting area, management of leaf wetness by reducing overhead irrigation, and increasing air flow through a crop are necessary to manage foliar nematodes (LaMondia 1999).

5.2

Root Knot (Meloidogyne spp.)

Geographic occurrence and impact. Root knot nematodes have been reported on zinnias in Pakistan, Taiwan, and the USA (Jabri et al. 1985; Mc Sorley and Frederick 1994; Tsay et al. 2004). Symptoms. Infected plants are stunted and poorly growing with yellowing leaves. Infected root systems show characteristic knots or galls. Biology and epidemiology. Zinnia elegans exhibited some galling in response to Meloidogyne incognita, but incidence of galling increased when the plants were infected with Zinnia mosaic virus (Jabri et al. 1985). Mc Sorley and Frederick (1994) found that Z. elegans cv. Scarlet was nearly free of galling from M. incognita and M. arenaria but was susceptible to M. javanica (3,400 eggs per plant). Tsay et al. (2004) also observed that some cultivars of Z. elegans and Z. hageana were highly and moderately resistant to M. incognita, while Z. angustifolia was more susceptible. The life cycles of Meloidogyne spp. differ slightly (De Guiran and Ritter 1979). The life cycle of M. incognita can be completed in 37 days. Second-stage juveniles penetrate root tips, occasionally invading roots in the zone of root elongation. Invasion of nematodes results in development of giant cells in the meristematic, cortical, and xylem tissues of the root and galling of roots occurs. Third- and fourthstage juveniles and young females occur after about 6–8 and 15 days, respectively. Adult females are observed after 20 days and egg laying begins after 25 days. Meloidogyne spp. are mostly spread by water and through equipments contaminated with infested soil. Management. Control of root knot nematodes is difficult and options are limited, especially in field production. Equipment should to be frequently cleaned to prevent dissemination of the nematodes from field to field. In greenhouses soil, disinfestation may be obtained by steam heat treatment, solarization, fumigation, or chemical drenches. • Biocontrol – The biological control agent Paecilomyces lilacinus strain 251 has been successfully applied in several crops to control M. incognita (Anastasiadis et al. 2008; Hashem and Abo-Elyousr 2011; Kiewnick and Sikora 2006). • Resistance – There have been several reports about the resistance of various zinnia cultivars against root knot nematodes (Mc Sorley and Frederick 1994; Tsay et al. 2004).

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Stimart DP, Boyle TH (2007) Zinnia. Zinnia elegans, Zinnia angustifolia. In: Anderson NO (ed) Flower breeding and genetics: issues, challenges and opportunities for the 21st century. Springer, Dordrecht, pp 337–357 Strider DL (1973) Bacterial leaf and flower spot of zinnia in North Carolina. Plant Dis Rep 57(12):1020. http://babel.hathitrust.org/cgi/pt?id=uc1.31175001263642;view=1up;seq=486 Strider DL (1976) An epiphytotic of bacterial leaf and flower spot of zinnia. Plant Dis Rep 60(4):342–344. http://babel.hathitrust.org/cgi/pt?id=uc1.31175001303299;view=1up;seq=370 Strider DL (1979a) Detection of Xanthomonas nigromaculans f. sp. zinniae in zinnia seed. Plant Dis Rep 63(10):869–873. http://babel.hathitrust.org/cgi/pt?id=uc1.31175005867711;view=1up; seq=377 Strider DL (1979b) Eradication of Xanthomonas nigromaculans f. sp. zinniae in zinnia seed with sodium hypochlorite. Plant Dis Rep 63(10):873–876. http://babel.hathitrust.org/cgi/pt?id=uc1. 31175005867711;view=1up;seq=377 Strider DL (1980) Control of bacterial leaf spot of zinnia with captan. Plant Dis 64(10):920–922. http://www.apsnet.org/publications/plantdisease/backissues/Documents/1980Articles/PlantDis ease64n10_920.pdf Strider DL, Jones RK, Haygood RA (1981) Southern bacterial wilt of geranium caused by Pseudomonas solanacearum. Plant Dis 65(1):52–53. doi:10.1094/PD-65-52 Szopińska D (2013) The effects of organic acids treatment on germination, vigour and health of zinnia (Zinnia elegans Jacq.) seeds. Acta Sci Pol Hortorum Cultus 12(5):17–29. http:// wydawnictwo.up.lublin.pl/acta/hortorum_cultus/2013/streszczenia2013_5/02%20Szopinska% 20Hort%2012_5_%202013.pdf Szopińska D (2014a) Alleviation of Zinnia elegans Jacq. seed deterioration using hydrogen peroxide and organic acids. Ecol Chem Eng S 21(2):309–326. doi:10.2478/eces-2014-0024 Szopińska D (2014b) Effects of hydrogen peroxide treatment on the germination, vigour and health of Zinnia elegans seeds. Folia Hort 26(1):19–29. doi:10.2478/fhort-2014-0002 Szopińska D, Tylkowska K, Deng CJ, Gao Y (2012) Comparison of modified blotter and agar incubation methods for detecting fungi in Zinnia elegans Jacq. seeds. Seed Sci Technol 40:32–42 Terry-Lewandowski VM, Stimart DP (1983) Multiple resistance in induced amphiploids of Zinnia elegans and Z. angustifolia to three major pathogens. Plant Dis 67:1387–1389. http://www. apsnet.org/publications/plantdisease/BackIssues/Documents/1983Articles/PlantDisease67n12_ 1387.pdf Terry-Lewandowski VM, Stimart DP (1985) The inheritance of resistance to powdery mildew in interspecific hybrids and induced amphiploids of Zinnia elegans Jacq. and Z. angustifolia HBK. Euphytica 34(2):483–487. http://link.springer.com/article/10.1007/BF00022945 Terry-Lewandowski VM, Bauchan GR, Stimart DP (1984) Cytology and breeding behavior of interspecific hybrids and induced amphiploids of Zinnia elegans and Zinnia angustifolia. Can J Genet Cytol 26(1):40–45. doi:10.1139/g84-007 Tesfagiorgis HB (2008) Studies on the use of biocontrol agents and soluble silicon against powdery mildew of zucchini and zinnia. PhD thesis. School of Agricultural Sciences and Agribusiness, Faculty of Science and Agriculture, University of KwaZulu-Natal, Pietermaritzburg, Republic of South Africa. pp. 182. http://siliconconference.org.za/fotos/Habtom%20PhD%20Thesis% 20Final.pdf#page=158 Thakur RN (1969) Sclerotium root rot of Zinnia elegans from Jammu and Kashmir. Labdev J Sci Tech 6(B):119–120 Thammakijjawat P, Thaveechai N, Kositratana W, Chunwongse C, Frederick RD, Schaad NW (2001) Genetic analysis of Ralstonia solanacearum strains from different hosts in Thailand using PCRrestriction fragment length polymorphism. Kasetsart J Nat Sci 35:397–408. http:// www.thaiscience.info/Article%20for%20ThaiScience/Article/2/Ts-2%20genetic%20analysis% 20of%20ralstonia%20solanacearum%20strains%20from%20different%20hosts%20in%20thai land%20using%20pcr-restriction%20fragment%20length%20polymorphism.pdf Trimboli D, Fahy PC, Baker KF (1978) Apical chlorosis and leafspot of Tagetes spp. caused by Pseudomonas tagetis Hellmers. Aust J Agri Res 29:831–839. doi:10.1071/AR9780831

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Tsai WS, Shih SL, Green SK, Hanson P, Liu HY (2004) First report of the occurrence of Tomato chlorosis virus and Tomato infectious chlorosis virus in Taiwan. Plant Dis 88(3):311. doi:10.1094/PDIS.2004.88.3.311B Tsay TT, Wu ST, Lin YY (2004) Evaluation of Asteraceae plants for control of Meloidogyne incognita. J Nematol 36(1):36–41. http://www.ncbi.nlm.nih.gov/pmc/articles/PMC2620738/ pdf/36.pdf Vali RJ, Moorman GW (1992) Influence of selected fungicide regimes on frequency of dicarboximide-resistant and dicarboximide-sensitive strains of Botrytis cinerea. Plant Dis 76:919–924. doi:10.1094/PD-76-0919 Verma VS, Singh S (1973) Zinnia leaf curl virus. Die Gartenbauwissenschaft 38(20), No. 2:159–162. http://www.jstor.org/stable/43387245?seq=1#page_scan_tab_contents Wang K, Hiruki C (2001) Use of heteroduplex mobility assay for identification and differentiation of phytoplasmas in the aster yellows group and the clover proliferation group. Phytopathology 91:546–552 Wehlburg C (1969) Two leaf spot diseases of zinnia. Plant Pathology Circular 86, Florida Department of Agriculture, Division of Plant Industry. https://www.freshfromflorida.com/content/ download/11093/142849/pp86.pdf Williamson B, Tudzynski B, Tudzynski P, Van Kan JAL (2007) Botrytis cinerea: the cause of grey mould disease. Mol Plant Path 8(5):561–580. doi:10.1111/J.1364-3703.2007.00417.X Wu WS, Chou JK (1995) Chemical and biological control of Alternaria carthami on zinnia. Seed Sci Technol 23:193–200 Wu WS, Yang YH (1992) Alternaria blight, a seed-transmitted disease in Taiwan. Plant Pathol Bull 1:115–123 Yadahalli KB, Kulkarni S, Anahosur KH (1994) In vitro and in vivo evaluation of fungicides against leaf spot of Zinnia caused by Cercospora zinniae Ell. and Mart. Karnataka J Agric Sci 7 (3):363–365. http://14.139.155.167/test5/index.php/kjas/article/viewFile/5996/6223 Yourman LF, Jeffers SN (1999) Resistance to benzimidazole and dicarboximide fungicides in greenhouse isolates of Botrytis cinerea. Plant Dis 83:569–575. doi:10.1094/ PDIS.1999.83.6.569

Part III Florists’ Crops Diseases: Potted Flowers

Diseases of Azalea

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Robert G. Linderman

Abstract

Florist azaleas (Rhododendron obtusum (Lindl.) Planch) are grown worldwide in greenhouses as a potted flower plant. They belong to the evergreen group but vary in their cold hardiness, so they must be protected against cold winter conditions. Many cultivars have been developed based largely on their horticultural traits, especially flower size and color. Some cultivars are prone to develop witches’ broom, thought to be a genetic disorder. However, most florist azaleas are susceptible to a number of diseases, but little attention has been paid to their resistance/susceptibility through breeding. Of concern to growers are diseases caused by species of Cylindrocladium, Pythium, Phytophthora, Rhizoctonia, and occasionally Exobasidium. Many of the diseases of florist azaleas, such as infections caused by Cylindrocladium, occur during the propagation phase of rooting cuttings, with surviving, infected-but-symptomless plants later succumbing to the disease.

Keywords

Phytophthora spp. • Pythium spp. • Cylindrocladium spp. • Exobasidium spp. • Rhizoctonia spp. • Witches’ Broom

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Phytophthora Root Rot, Dieback, and Wilt (Phytophthora spp.) . . . . . . . . . . . . . . . . . . . . 2.2 Pythium Root Rot (Pythium spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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R.G. Linderman (*) Plant Health, LLC, Corvallis, OR, USA e-mail: [email protected] # Springer International Publishing AG 2018 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florists’ Crops Diseases, Handbook of Plant Disease Management, https://doi.org/10.1007/978-3-319-39670-5_29

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2.3 Cylindrocladium Blight and Wilt (Cylindrocladium scoparium Morgan) . . . . . . . . . . . 2.4 Leaf and Flower Gall (Exobasidium spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Rhizoctonia Root Rot and Web Blight (Rhizoctonia spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Genetic Abnormalities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Witches’ Broom . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Introduction

Evergreen azaleas originated in Japan and are classified in the genus Rhododendron, subgenus Tsutsutsi. These grow indoors and outdoors. Most azaleas bloom in spring, although evergreen florist’s azaleas can be forced into bloom under greenhouse conditions for gift giving at any time. Evergreen azaleas vary in cold hardiness with the particular cultivar, but the group spans US Department of Agriculture hardiness zones 5 through 9. A significant component of the florist industry involves the sale of potted evergreen azaleas (Rhododendron obtusum (Lindl.) Planch). Plants are grown in production greenhouses for protection against winter cold, since they vary in cold hardiness. Accordingly, they can be “forced” into flower any time by manipulating the environmental conditions to set flower buds that will bloom predictably to match sales for special occasions. Many cultivars have been selected, mainly for their predictable and extraordinary flowering, with variation in flower size and color. Cultivar development has paid little attention to disease susceptibility/ resistance. Some diseases, such as Cylindrocladium leaf spot, blight, and/or root rot, have caused tremendous losses in the nursery industry over the years. Other diseases, common to many other greenhouse and nursery crops, include root rots caused by species of Pythium, Phytophthora, and Rhizoctonia. Some foliage and flower diseases, such as leaf and flower gall, caused by species of Exobasidium occur mainly on landscape plants and rarely on florist azaleas, mainly because it is readily controlled by judicious removal of galls and application of chemical sprays. Most of the more serious diseases are caused by soilborne pathogens that infect newly rooted cuttings. Surviving plants are infected but remain symptomless until some later time when poor growth, dieback, and mortality occur. Soilborne root pathogens are much more challenging to manage because of the multiple ways they can invade the greenhouse cultural system and the limitations on effective management. Nonetheless, numerous methods have been identified to block entry, largely by eliminating the pathogens from the sources. For example, sanitation of containers with heat, treatment of irrigation water, using pathogen-free growth media, chemically sanitizing cutting propagation material, and maintaining strict sanitation practices within the greenhouse environment can be effective measures for preventing entry into the greenhouse. Some pathogens like Cylindrocladium spp. are especially troublesome because they can disperse spores (inoculum) via an aerial phase within the greenhouse environment.

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2

Fungal and Fungus-Like Diseases

2.1

Phytophthora Root Rot, Dieback, and Wilt (Phytophthora spp.)

875

Geographic occurrence and impact. Phytophthora spp. are known to infect azaleas and rhododendrons worldwide. These fungus-like organisms (Phylum Oomycota and genetically related to the brown algae) infect many other plants within the Ericaceae family as well (Jones and Benson 2001; Linderman and Benson 2014). Interestingly, some of those plants occur as wild species, but the diseases do not occur in natural settings, even when the pathogen is present, for unknown reasons. They cause root rot, crown canker, and branch dieback, depending on the species. Some species can cause all of those symptoms, while others are confined to the roots and crown. Most of the Phytophthora diseases occur in landscapes or in nursery field plantings, in ground or in containers. When the diseases occur in the nurseries, significant economic losses result. Beyond the nursery, after shipment and sale, further disease development, even mortality, may occur in the hands of the consumer. Those occurrences are frequently a result of earlier undetected infections or contamination that develops after time and/or a change in environmental and soil conditions. Sometimes, infected liner plants appear healthy and are shipped to other wholesale nurseries for growing on, and with further development of the infections, symptoms appear. Symptoms/signs. Root infections may develop at any time in the life of florist azaleas, from cutting propagation to field and landscape plantings. Most frequently involved are P. cinnamomi Rands and P. nicotianae Breda de Haan (syn. P. parasitica Dastur), but in some areas, other species such as P. citricola Sawada are recovered. Infected roots are brown and become necrotic. When enough roots are infected, the root system becomes impaired, and as a result leaves may become chlorotic and then necrotic but rarely wilt. Those leaves generally drop off the plant, causing partial defoliation (Fig. 1). Leaves on infected plants, as well as the plants themselves, are also often smaller than with healthy plants. With some Phytophthora species, root infections progress to the lower stem tissue where lethal crown cankers occur. These cankers are revealed by removal of the bark, exposing the internal wood which appears brown. Biology and epidemiology. Root, crown, stem, and foliage infections caused by a number of species of Phytophthora can occur on florist azaleas when sufficient inoculum is present and environmental conditions are conducive. Those infections occur as a result of inoculum being disseminated by air movement, or by water splash of spores from the soil to aboveground tissues landing on susceptible sites. Free water is required for spore germination and infection. The source of inoculum (zoospores, sporangia, chlamydospores, or oospores, depending on the species) in greenhouse culture of florist azaleas is often unknown. Temperature conditions in the

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Fig. 1 Mortality of azalea (left) caused by Phytophthora citricola (R.G. Linderman # 2017. All Rights Reserved.)

greenhouse systems are generally in the right range for infections by Phytophthora spp., as with Pythium spp. infections. So the reduction of infections by environmental conditions that might occur outdoors does not apply for florist azaleas. Temperature is modified by heating and cooling to enhance plant growth. When flower bud setting is involved, temperatures are usually lowered. This allows plants destined for the florist trade to be programmed for forcing in time for special holiday sales. The source of Phytophthora spp. inoculum varies, depending a lot on the practices of the nursery. For soilborne, root-infecting pathogens, the medium, recycled containers, irrigation water, and plant propagules used for cutting propagation could be sources. Strict sanitation practices within the greenhouse, as well as the propagation area, must be maintained. Soilless growth media are used in growing florist azaleas, so they are least likely to be a source. Use of well or municipal water is also not likely to be a source. Likely sources are reused containers, recycled irrigation water, and source of propagation cuttings. Management. The primary management strategy for Phytophthora-caused diseases of florist azaleas is prevention. Eliminating the pathogen from the major sources of irrigation water, the growth medium, the containers, and even the cuttings can go a long way to preventing these diseases. Thereafter, application of preventative chemical fungicides provides some protection against invasion from a number of sources, including greenhouse workers. The chemicals used for Phytophthora control are the same as for Pythium control, and they generally target only those genera and some other fungus-like pathogens, but not fungal pathogens. There are preventative chemicals such as products containing etridiazole, mefenoxam, cyazofamid, fenamidone, and phosphonates. The insensitivity of some species of Pythium to mefenoxam has also been reported for Phytophthora spp. However, there

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can be variation in effectiveness of specific chemicals to different Phytophthora spp. (Linderman and Davis 2008a). Rotation or combinations with other chemicals should be practiced, as with management of Pythium spp. In addition to chemical agents, there are some biological control agents that target certain species of Phytophthora and Pythium. Species of Trichoderma and Streptomyces have been shown to have that activity but usually have not been tested on a range of pathogen species. Suppressive soils that prevent or limit some root diseases, including those caused by species of Phytophthora, have been reported. Often suppression is created by incorporating mixtures of microbes with great diversity into the growth medium. Within that microbial population are members with specific capacity to block the development of specific diseases. Some have the capacity to block sporangium development, and therefore reduce zoospore release (Linderman et al. 1983). Some are able to lyse the pathogen mycelium, while others produce other antibiotic compounds that inhibit growth of the pathogen in soil. With production of florist azalea, however, such practices generally are not used.

2.2

Pythium Root Rot (Pythium spp.)

Geographic occurrence and impact. Roots of many plant species can be infected by species of Pythium. Most often Pythium spp. are known as seedling damping-off pathogens, but they are root pathogens of many woody plants, including florist azaleas (Ivors 2014). They do not cause immediate death of plants but cause root rot that can debilitate plants to various degrees, depending on the time and extent of infection of the root system. Young rooted azalea cuttings can be killed if enough of the root system is involved. That will depend on when the pathogen begins to infect, relative to the rooting process. If infection occurs after a substantial number of roots have formed, the infections might not be detected, and the plants become infectedbut-symptomless until the percentage of infected roots increases. A number of species of Pythium can be involved in azalea root rot. Some may be more pathogenic than others (Weiland et al. 2013). Pythium spp. might also be secondary invaders to infections by species of Phytophthora. The net result and impact of Pythium root infections, therefore, is difficult to determine. If growing conditions in the greenhouse are favorable, Pythium infections might go unnoticed until some other stress occurs on the plants. Then the combination of Pythium root rot and stress causes the plants to show symptoms on the foliage, such as leaf chlorosis, small leaves, even dieback of young branches. Examination of the roots and detection of root rot suggest the role that Pythium played in the syndrome. Such plants, however, do not make it into the florist trade because they are discarded. Symptoms/signs. A range of symptoms of Pythium root rot can occur, including failure of cuttings to root, rooting followed by root rot, young plant mortality, stunting of plants at later growth stages, chlorosis of foliage, wilting under water deficiency, loss of vigor, necrotic larger roots, and leaf drop (Linderman and Benson

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Fig. 2 Symptoms of Pythium root rot of azalea: Left photo showing greater root browning of plant on left. Right photo showing aboveground symptoms of smaller plant, smaller leaves, and off-colored foliage, compared to plant on the-left (R.G. Linderman # 2017. All Rights Reserved.)

2014) (Fig. 2). Examination of the root ball of affected plants, compared to nonaffected healthy plants, usually shows a greatly reduced size. Often it is difficult to distinguish many of the aboveground symptoms from a nutritional disorder. Examining the root ball is critical. Over-watering or drought episodes may hasten the development of aboveground symptoms, but plants without Pythium root rot generally show no or fewer symptoms. Biology and epidemiology. Identifying the species of Pythium involved in root rot of azalea is extremely difficult, even for the specialist. Ten species have been identified on rhododendrons and azaleas, but those affecting florist azaleas have not. It is possible, even likely, that more than one species can be involved. Further, some species/strains can be more pathogenic than others (Weiland et al. 2013). There can be many sources of inoculum of Pythium species that can invade the florist azalea production system: (1) Used and recycled containers have been shown to harbor Pythium species, even after rigorous washing. Baiting residual contaminated medium from propagation flats or larger containers reveals that Pythium spp. are present. It generally is impossible to predict just how pathogenic each might be without inoculation experiments. The variability of species or strains within a crop has been demonstrated, however (Weiland et al. 2013). (2) Contaminated media is another potential source of inoculum leading to Pythium root rot. Most growers rely on the soilless medium they buy or mix themselves being pathogen-free. However, Pythium has been found in some peat mosses, albeit infrequently. (3) Irrigation water is a likely source of Pythium where water is being recycled in the summer months. However, often well water is used in propagation, and it is believed to be pathogenfree. Water in irrigation reservoirs should be treated in order to prevent inoculum from being delivered into production pots. Generally chlorine compounds, such as hypochlorite or chlorine dioxide, or even chlorine gas have been used for that purpose. (4) Insect vectors, such as fungus gnats, have been suggested to carry propagules of Pythium from an infected pot to healthy pots. However, there is some

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degree of controversy in this regard, in that some studies show that the pathogen does not carry over from the larval to the adult stage that flies. However, there is no doubt that the larvae transmit the pathogen from root to root within a pot. Pythium root rot often occurs in combination with some environmental stress, such as high soluble salts in the medium, over-watering, under-watering (drought), or poor drainage. Water saturation of the medium can allow the pathogen to produce sporangia and then zoospores spread from root to root. Those conditions also stress plants in a physiological manner that makes them more susceptible, even for cultivars that generally are thought to be somewhat resistant. Having covered all of the primary sources of Pythium spp. of potting medium, used containers, plant propagules, or irrigation water, it is still essential to prevent the introduction of Pythium thereafter. Pythium spp. can be lurking around, just waiting for some lack of sanitation. So it is important to disinfect all bench surfaces, potting benches, tools, and equipments that will contact the potting mix. Management. Preventing Pythium root rot should be the primary management strategy. Eliminating the pathogen from the major sources of irrigation water, the growth medium, the containers, and even the cuttings can go a long way to managing this disease. Thereafter, application of preventative chemical fungicides provides some protection against invasion from a number of sources, including greenhouse workers. The chemicals used for Pythium control are the same as for Phytophthora control, and they generally target only those genera and some other fungus-like microorganisms, but not fungal pathogens. They are preventative chemicals, such as products containing etridiazole, mefenoxam, cyazofamid, fenamidone, and phosphonates. However, some species of Pythium are not sensitive to mefenoxam (Aegerter et al. 2002), so rotation or combinations with other chemicals should be practiced. In addition to chemical agents, there are some biological control agents that target Pythium spp., such as species of Trichoderma and Streptomyces. There are examples of suppressive soils that prevent or limit some root diseases, often created by incorporating organic materials, such as composts, that contain mixtures of microbes with great diversity. Within that microbial population are members with specific capacity to block the development of specific diseases. Some have the capacity to block sporangium development and therefore reduce zoospore release. Some are able to lyse the pathogen mycelium, while others produce other antibiotic compounds that inhibit growth of the pathogen in soil.

2.3

Cylindrocladium Blight and Wilt (Cylindrocladium scoparium Morgan)

Geographic occurrence and impact. A serious disease of evergreen, florist azaleas occurred in the USA and in European countries in the mid-1950s; within 10 years it appeared in major proportions in many parts of the USA and other countries. The cause of the disease was Cylindrocladium scoparium, a fungus previously known to attack primarily conifer and other tree species in seedling

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nurseries. Subsequently, the disease occurred on several other major nursery crops. The diseases varied in symptom expression, including root rot, stem lesions, leaf spots and flower blight, and wilt. In recent years to the present, serious Cylindrocladium diseases have occurred on azalea and mini rose, as well as other ericaceous plants, primarily in the early stages of cutting propagation. Subsequent sporulation of the pathogen under greenhouse conditions and spread by water splash or air movement of spores results in devastating incidence and severity on many, often thousands, of plants. During that phase of sporulation and spread of the disease, more plants become infected, but remain symptomless, only to wilt and succumb later. Growers have sustained, and are still sustaining, severe economic losses from these diseases, often in the range of hundreds of thousands of dollars per year per grower. Symptoms/signs. There are multiple symptoms of Cylindrocladium infections of florist azaleas, depending on the production stage and environmental conditions (Linderman and Benson 2014). Leaf spot infections can occur if there is airborne inoculum in the greenhouse (Fig. 3). Often those spots go unnoticed because the infected leaf will abscise (due to the production of ethylene) and be lost on the pot surface among other leaves that had fallen for other reasons. Leaf spots are discrete spots that become necrotic as seen from the upper leaf surface. On the lower surface, however, mainly on red or pink cultivars, the veins radiating out from the spot are red. That symptom is absent or less pronounced on white cultivars. Flowers may also be infected by aerial inoculum, forming necrotic flecks that rapidly coalesce (Fig. 4). Infected fallen leaves become completely colonized by the pathogen in time as they remain moist. Internally, the pathogen forms microsclerotia in the leaf tissue

Fig. 3 Leaf spots on azalea plant caused by Cylindrocladium scoparium (left); close-up of spot on cutting during propagation (right) (R.G. Linderman # 2017. All Rights Reserved.)

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Fig. 4 Flower infections on azalea caused by inoculation with Cylindrocladium scoparium (R.G. Linderman # 2017. All Rights Reserved.)

Fig. 5 Microsclerotia of Cylindrocladium scoparium formed in infected leaves (left and middle) or roots (right) (R.G. Linderman # 2017. All Rights Reserved.)

that provide it long-term survival. Microsclerotia may also form in infected roots (Fig. 5). Under moist conditions, the sclerotia can generate abundant leaf-surface conidial sporulation that can contribute to the pathogen’s dispersal within the greenhouse (Fig. 6). Examination of such leaves, even with a hand lens, will reveal the diagnostic sporulation. Conidia released from fallen, infected leaves can wash into the medium and initiate root rot on newly formed roots. Infected roots are brown initially and then become necrotic. The extent of roots infected determines when the plant will succumb. Often, infected cuttings will die in pots where four cuttings were stuck

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Fig. 6 Photo on the left shows sporulation of Cylindrocladium spp. on abscised azalea leaf. Photo in the center is a close-up of conidial clumps generated from microsclerotia imbedded in infected leaf. Photo on the right shows perithecia of Calonectria kyotensis (teleomorph) formed on infected azalea leaf (R.G. Linderman # 2017. All Rights Reserved.)

Fig. 7 Wilt symptoms on azalea cuttings (left) and older plant (right) where one of four cuttings was infected. Note distinctive defoliation from infection in photo on right (R.G. Linderman # 2017. All Rights Reserved.)

(Figs. 7 and 8). In many cases, however, the infections do not affect the appearance of the aboveground portions of the plant, mainly because new roots are forming fast enough to maintain the plant in spite of the infected roots. At a later stage, however, mortality may occur. Often, three or four rooted cuttings are transplanted into a single pot for growing on, and only one or two of them die (Fig. 9). That is a sign that the infection began at an earlier stage in propagation. Historically, however, when only one plant was transplanted to a larger pot, that plant eventually wilted due to the

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Fig. 8 Root systems of azalea plants from one pot showing effects of root rot caused by Cylindrocladium (R.G. Linderman # 2017. All Rights Reserved.)

Fig. 9 Wilting of one azalea plant of four in the pot due to earlier infection by Cylindrocladium (left photo). Older plant wilting and death due to earlier root infection (right photo) (R.G. Linderman # 2017. All Rights Reserved.)

initial root infections on the cuttings. At that time, cutting into the lower stem would usually reveal an internal, brownish canker. That symptom alone, however, would not distinguish Cylindrocladium as the cause from cankers caused by Phytophthora, for example. Additional diagnostic tests need to be done to confirm Cylindrocladium blight.

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Biology and epidemiology. The main causal agent is Cylindrocladium scoparium Morgan, but other species with a perfect stage (teleomorph), such as Calonectria kyotensis and Calonecria theae, were also shown to be pathogenic (Alfieri et al. 1972). Those species with a Calonectria stage were shown to eject ascospores aerially when relative humidity declined (Linderman 1974a). In nearly all cases, cutting and root rot of azaleas caused by species of Cylindrocladium begins during cutting propagation and spreads thereafter, depending on the greenhouse production system. It was demonstrated experimentally (Linderman 1973, 1974b) that the pathogen produced microsclerotia in infected leaves that dropped from plants with leaf spot infections. Those infections were often not noticed, and leaf abscission occurred as a result of ethylene production (Axelrood-McCarthy and Linderman 1981). The microsclerotia imbedded in the fallen leaves generated conidiophores on the surface, and conidia then splashed to other cuttings, causing root rot (Linderman 1974a). Some cutting roots became infected, but the plant remained symptomless. In the absence of stress under ideal growing conditions, liner plants moved through the production stages were sold and shipped only to die at some later time. Management. Aerially dispersed conidia that landed on plants from which cuttings would be taken can be contaminants, not yet causing infections. Therefore, chemical sanitization by dipping cuttings in solutions of fungicides or chlorine compounds, killing the contaminating conidia, is an effective management practice (Linderman, unpublished). Failure to sanitize cuttings has led to disastrous consequences from this disease. The source of airborne conidia in a greenhouse may be unknown, but infected plants with sporulation at the lower stem could be one source (Linderman, personal observation). Fallen, infected leaves on or below the production benches also can harbor the pathogen and become a source of dispersed conidia. Removal of such debris and flaming the areas under the benches and chemically sanitizing the benches is essential to removing sources of the pathogen inoculum. Assessment of the presence of Cylindrocladium spp. in soil or soilless media might also be needed, especially if the propagation medium is to be reused. Inserting azalea leaves into the medium and incubating for a week was shown to be effective in detecting the pathogen. The leaves were selective in being only infected by Cylindrocladium spp. (Linderman 1972). The method was also effective in isolating the pathogen from the stem by inserting a leaf into the cut end. If Cylindrocladium, or any other soilborne fungal pathogen, was detected, the medium must be sanitized with heat or a fumigant like Metam sodium prior to reuse (Linderman and Davis 2008b). Application of fungicides can also provide protection of plants exposed to aerial inoculum. Frequency of application is also critical due to the rate of sporulation of the pathogen as well as development of new, unprotected leaf surface from expansion. Fungicides containing thiophanate methyl have been shown to be effective.

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2.4

Diseases of Azalea

885

Leaf and Flower Gall (Exobasidium spp.)

Geographic occurrence and impact. Leaf and flower gall disease of azaleas occurs widely throughout the world, mainly on landscape plants, but it can occur on florist azaleas under moist conditions. In the United States, the causal agent is generally considered to be Exobasidium vaccinii (Fuckel) Woronin. In China, Japan, and Korea, however, E. japonicum Shirai is considered to be the pathogen. Exobasidium rhododendri (Fuckel) C. Cramer has been reported on both azalea and rhododendron worldwide. The origin of the disease is not known, but likely first occurred on rhododendrons in their centers of origin. If enough galls appear on a plant, there can be some loss of vigor of the plant. Such severe infections rarely occur in commercial greenhouse production, however. The impact of the disease comes from the time and cost of its management, involving labor and preventative chemical applications. Symptoms/signs. Leaf gall first appears as light green swellings on the edges and undersides of leaves. Both expanding leaves and flower buds and young, developing shoots are susceptible to infection. The infected tissue expands to form gall-like structures that generally involve the entire leaf (Fig. 10). As the galls mature, they turn white from surface sporulation of the fungal pathogen. The galls can be small or large and bladder shaped. The fungus causes the plant cells to divide and enlarge, making the tissue soft and succulent. Once galls turn white from sporulation, the pathogen can spread very rapidly by water splash or air movement to other plants, initiating new, secondary infections on leaves and even flowers. After sporulation occurs, the leaf galls slowly turn brown, shrink, and become quite hard. Biology and epidemiology. Spores of the pathogen apparently overwinter in bud scales. As buds break, the spores germinate and infect the young bud tissue as it expands. Humidity level in the greenhouse is a critical factor in the epidemiology of the leaf gall disease. High relative humidity and free water on the leaves and flowers

Fig. 10 Azalea leaf gall caused by Exobasidium vaccinii. Left: young galls; Middle: pathogen sporulation on gall; Right: older, necrotic galls (R. Rosetta # 2017. All Rights Reserved.)

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is required for spore germination and initiation of new infections, so irrigation scheduling is very important. Overhead watering must be done early in the day so the foliage is not wet through the night. Because of the variation in relative humidity, often the lower leaves, where humidity is higher, are more susceptible than upper leaves where air circulation is better. Once infection has begun, the fungus grows in the intercellular spaces of the surface tissue, producing haustoria that penetrate cells to derive its nutrition. Eventually, often within a few days, the pathogen sporulates on the gall surface, causing it to turn white. Spores produced on a gall can be released and spread to other expanding buds, causing secondary infections. Some of the spores settle into protected sites in forming buds and are in position to initiate new infections. Management. The main strategy for managing azalea leaf gall is to diligently scout production plants for the first signs of gall development. Small galls must be removed before they turn white with sporulation. Prevention of galls also generally involves application of fungicides as buds break and new foliage appears. Plants should be sprayed before and after removing galls with chemicals such as mancozebcontaining fungicides, copper-containing materials, quaternary ammonium materials, and even oxidizing agents such as hydrogen peroxide. A good sticker may be needed to help retain the applied chemical. Application of any of these materials is strictly preventative, not curative.

2.5

Rhizoctonia Root Rot and Web Blight (Rhizoctonia spp.)

Geographic occurrence and impact. Rhizoctonia root rot and subsequent blighted foliage occurs occasionally on florist azaleas in the United States (Copes and Benson 2014). The disease also has been reported in Australia. This disease can occur on many different woody ornamental plants, including rhododendrons. Most often, the disease occurs during cutting propagation where moist, warm conditions prevail. Economic losses result from the loss of affected rooted cuttings, but some infections aren’t expressed until later when canopy growth is affected and some dieback/blight occurs. Symptoms/signs. When Rhizoctonia root rot occurs during propagation of cuttings, the affected cuttings fail to root or the new roots become infected, causing the entire cutting to collapse (Fig. 11). Often the infection site will expand and develop, spreading to adjacent cuttings. Webbing of aerial hyphae occurs, looking like a spider web moving from plant to plant. The pathogen moves both in the rooting medium and from leaf to leaf above the medium surface. Biology and epidemiology. Some propagators root cuttings under a reemay cover in order to retain heat and to reduce the frequency of misting from above. The high humidity under the reemay allows the fungus to move from one infected cutting to

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Fig. 11 Hyphal webbing site of azalea cuttings in propagation caused by Rhizoctonia solani. Note discrete spots on leaf in lower right (R.G. Linderman # 2017. All Rights Reserved.)

adjacent cuttings, often leading to a patch of infected cuttings. The occurrence of a web-blight pattern where the fungus becomes aerial and infects the lower leaves results in a distinct leaf spot at the infection site. The primary causal agent, Rhizoctonia solani J. G. Kuhn, is a nonsporulating Deuteromycete, so how the pathogen invades the propagation system is unknown. The possible sources could be the medium, reused rooting flats, or some cuttings becoming contaminated during cutting collection. The incidence of this disease is very infrequent and spread is by vegetative means. Spread may also occur in particulate material moved by air currents. Management. The infrequent occurrence of Rhizoctonia root rot and cutting death supports the idea that no active management is required, other than removal of affected cuttings at an infection site, including adjacent cuttings that usually show no signs of infection. Often that means removal of the entire affected plug flat. Addressing the possible sources of the pathogen, however, is in order. If propagation flats are being reused, then their sanitization with aerated steam (Linderman and Davis 2008b) is required to kill any carryover of the pathogen from a previous crop, even crops other than azalea. If the rooting medium is suspected as the source, then a decision would be needed as to steam treatment that is generally not a course of action unless other diseases have occurred that would also be curbed by medium pasteurization. Since R. solani can colonize organic components of the medium as a saprophyte, rigid sanitation in the propagation area is called for to prevent that from happening. Affected areas of the propagation area should also be treated chemically to avoid spread and carryover. Chemicals that specifically target Rhizoctonia pathogens should be used; broader spectrum chemicals might be used if other causal agents that can cause similar

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symptoms are known to occur, such as species of Pythium or Phytophthora. Since sporulation is not involved in disease initiation or spread, cuttings are less likely to be contaminated, so their sanitation generally is not necessary.

3

Genetic Abnormalities

3.1

Witches’ Broom

Geographic occurrence and impact. Some cultivars of florist azaleas have a tendency to develop a cluster of small leaved branches in a tight configuration characterized as a witches’ broom. The occurrence of these structures has little impact on the growth of the plant, and plants that exhibit them are generally discarded as being offtype or the affected branch simply pruned off (Linderman 2014). Symptoms/signs. The witches’ broom branches occur generally on one location on the plant, often near the base of the plant. The tight branching and greatly reduced leaf size distinguish it from the rest of the plant (Fig. 12). Biology and epidemiology. Evergreen azaleas and rhododendrons occasionally exhibit abnormal, miniaturized vegetative growth in the form of a witch’s broom. Researchers generally agree that the condition is the result of a genetic change that occurred, and that no biotic causal agent is involved. Therefore, there is no risk of transmission from affected plants to adjacent plants. Management. Since no biotic agent is involved in the development of witches’ broom, no management is required other than to either prune off the broom, or simply discard the plant as being off-type. Fig. 12 Witches’ broom on azalea (R.G. Linderman # 2017. All Rights Reserved.)

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References Aegerter BJ, Greathead AS, Pierce LE, Davis RM (2002) Mefenoxam – resistant isolates of Pythium irregulare in an ornamental greenhouse in California. Plant Dis 86(6):692 Alfieri SA Jr, Linderman RG, Morrison RH, Sobers EK (1972) Comparative pathogenicity of Calonectria theae and Cylindrocladium scoparium to leaves and roots of azalea. Phytopathology 62:647–650 Axelrood-McCarthy PE, Linderman RG (1981) Ethylene production by cultures of Cylindrocladium floridanum and C. scoparium. Phytopathology 71:825–830 Copes WE, Benson DM (2014) Rhizoctonia damping-off and root rot; Rhizoctonia web blight. In: Linderman RG, Benson DM (eds) Compendium of rhododendron and azalea diseases and pests, 2nd edn. APS Press, St. Paul, pp 14–19 Ivors KL (2014) Pythium damping-off and root rot. In: Linderman RG, Benson DM (eds) Compendium of rhododendron and azalea diseases and pests, 2nd edn. APS Press, St. Paul, pp 13–14 Jones RK, Benson DM (eds) (2001) Diseases of woody ornamentals and trees in nurseries. APS Press, St. Paul Linderman RG (1972) Isolation of Cylindrocladium from soil or infected azalea stems with azalea leaf traps. Phytopathology 62:736–739 Linderman RG (1973) Formation of microsclerotia of Cylindrocladium spp. in infected azalea leaves, flowers, and roots. Phytopathology 63:187–191 Linderman RG (1974a) Ascospore discharge from perithecia of Calonectria thea, C. crotalariae, and C. kyotensis. Phytopathology 64:567–569 Linderman RG (1974b) The role of abscised Cylindrocladium-infected azalea leaves in the epidemiology of Cylindrocladium wilt of azalea. Phytopathology 64:481–485 Linderman RG (2014) Witches’ broom. In: Linderman RG, Benson DM (eds) Compendium of rhododendron and azalea diseases and pests, 2nd edn. APS Press, St. Paul, p 68 Linderman RG, Benson DM (eds) (2014) Compendium of rhododendron and azalea diseases and pests, 2nd edn. APS Press, St. Paul Linderman RG, Davis EA (2008a) Evaluation of chemical agents for the control of Phytophthora ramorum and other species of Phytophthora on nursery crops [Online]. Plant Health Prog. doi:10.1094/PHP-2008-0211-01-RS Linderman RG, Davis EA (2008b) Eradication of Phytophthora ramorum and other pathogens from potting medium or soil by treatment with aerated steam or fumigation with metam sodium. HortTechnology 18(1):106–110 Linderman RG, Moore LW, Baker KF, Cooksey DA (1983) Strategies for detecting and characterizing systems for biological control of soilborne plant pathogens. Plant Dis 67:1058–1064 Weiland JE, Beck BR, Davis A (2013) Pathogenicity and virulence of Pythium species obtained from forest nursery soils on Douglas-fir seedlings. Plant Dis 97:744–748

Diseases of Begonia

30

Cristina Rosa and Gary W. Moorman

Abstract

Begonias are susceptible to a wide variety of fungi, bacteria, and viruses, as well as nematodes and abiotic diseases. The systemic nature of some of the pathogens makes it likely that the diseases they cause can be found wherever vegetatively propagated begonias are shipped. Management of these pathogens is paramount for specialty propagators while growers purchasing plants must inspect incoming plants for symptoms and understand the biology of the pathogens involved in order to manage them effectively. Keywords

Fusarium wilt • Botrytis blight • Powdery mildew • Pythium root rot • Bacterial spot • Viruses • Foliar nematodes • Abiotic diseases

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Fusarium Wilt (Fusarium foetens) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Gray Mold (Botrytis Blight; Botrytis cinerea Pers.: Fr.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Powdery Mildew (Oidium begoniae, Asexual Stage) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Pythium Root Rot (Pythium sp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Bacterial and Phytoplasma Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Bacterial Spot (Bacterial Blight; Xanthomonas campestris pv. begoniae (Takimoto) Dye) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Viral Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Clover Yellow Mosaic Potexvirus (ClYMV) Described by Johnson (1942) . . . . . . . . 4.2 Cucumber Mosaic Virus (CMV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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C. Rosa (*) • G.W. Moorman (*) Department of Plant Pathology and Environmental Microbiology, The Pennsylvania State University, University Park, PA, USA e-mail: [email protected]; [email protected] # Springer International Publishing AG 2018 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florists’ Crops Diseases, Handbook of Plant Disease Management, https://doi.org/10.1007/978-3-319-39670-5_30

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4.3 Impatiens Necrotic Spot Virus (INSV) and Tomato Spotted Wilt Virus (TSWV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4 Tobacco Mosaic Virus (TMV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5 Tobacco Necrosis Necrovirus (TNV) and Carnation Mottle Carmovirus (CarMV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6 Tobacco Ringspot Virus (TRSV) and Arabis Mosaic Nepovirus (ArMV) . . . . . . . . . . 4.7 Broad Bean Wilt Virus (BBWV) Described by Stubbs (1947) . . . . . . . . . . . . . . . . . . . . . . 4.8 Zucchini Yellow Mosaic Virus (ZYMV) Described by Lisa et al. (1981) . . . . . . . . . . . 5 Nematode Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 Foliar Nematode (Aphelenchoides fragariae) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 Abiotic Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Introduction

Begonias are a very diverse and complex group of plants. The name “Begonia” was first used by Charles Plumier, a Franciscan monk and botanist, in 1703. Linnaeus described all the species known to him in 1753, and by the end of the 1700s, begonias were being cultivated widely in Europe. By the early 1800s, hybrids were being developed. Based on their dominant horticulture characteristics, begonias can be grouped in categories including cane-like, shrub-like, semperflorens, rhizotamous, rex cultorum, tuberous, and trailing-scandent (Thompson and Thompson 1981).

2

Fungal and Fungus-Like Diseases

2.1

Fusarium Wilt (Fusarium foetens)

Geographic occurrence and impact. This disease has been found in Europe, Japan, and North America causing wilt and sometimes stem rot (Elmer and Vossbrinck 2004; Schroers et al. 2004; Sekine et al. 2008; Tian et al. 2012; Tschope et al. 2007; Van der Gaag and Raak 2010). Symptoms/signs. Leaves become dull green and the leaf veins may yellow. The leaves wilt and die. The vascular tissue of infected plants turns brown. A basal stem rot may develop as the disease process proceeds (Fig. 1). Large numbers of pale orange spores develop on the dying tissue, particularly if the tissue is placed on a wet surface in a container and incubated. Biology and epidemiology. Fusarium foetens is related to the Fusarium oxysporum complex that contains many form species responsible for vascular wilts of various plants. It is thought that the pathogen was introduced to Europe on plant material brought there for breeding (Schroers et al. 2004), indicating that the pathogen may be spread long distances associated with plant shipments.

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Fig. 1 Fusarium wilt on begonia (Wade Elmer # 2017. All Rights Reserved.)

Management. Infected plants should be destroyed. The various types and cultivars of begonia differ in relative susceptibility (Brand and Wienberg 2005; Tian et al. 2012; Tian and Zheng 2012). The biological control agents Bacillus subtilis, Streptomyces griseoviridis, Gliocladium catenulatum, Streptomyces lydicus, and Trichoderma harzianum have been reported to provide protection of begonias against Fusarium foetens in greenhouse experiments (Tian and Zheng 2013). To reduce the presence of the fungus in production systems, chlorine bleach, hydrogen peroxide, and quaternary ammonia sanitizers have been found effective (Elmer 2008).

2.2

Gray Mold (Botrytis Blight; Botrytis cinerea Pers.: Fr.)

Geographic occurrence and impact. This disease is very common and occurs wherever plants are grown if the environmental conditions are conducive to its development. The pathogen, Botrytis cinerea, can infect almost any plant in a greenhouse including begonias. Symptoms/signs. Cuttings rot at their base. Tan spots develop on leaves. Established plants rot at the crown or a dry shriveled area develops along a branch (Tompkins 1950). Abundant, dusty, gray masses of spores form on infected tissue if the humidity is high or if infected tissue is placed in a container with moisture. Biology and epidemiology. Seedlings and all above ground parts of mature plants are susceptible. The fungus can infect intact tissue or through wounds. Infection occurs most readily when the fungus has a food base from which it attacks, such as fading flowers or senescent leaves that have fallen onto healthy leaves. High relative humidity, wetness on the plant, and temperatures between 18  C and 25  C greatly favor Botrytis infection (Jarvis 1980). The spores are readily spread by air currents (Hausbeck and Pennypacker 1991).

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Management. Maintain low humidity within the crop canopy by spacing plants well and venting in a greenhouse setting to improve air circulation. Remove dead and dying flowers and leaves from the area around the plants, from the potting soil surface, and from the plant. Remove this debris from the greenhouse promptly or put it in a closed waste container. Avoid unnecessarily damaging plants. Apply a fungicide preventively but note that the exclusive use of one class of chemical, particularly systemic chemicals, can result in the selection of fungicide resistance to the chemical class. Resistance to benzimidazole fungicides is very widespread in Botrytis populations Do not rely on only one chemical class (Elad et al. 1992; Gullino and Garibaldi 1987; Hausbeck and Moorman 1996; Moorman and Lease 1992).

2.3

Powdery Mildew (Oidium begoniae, Asexual Stage)

Geographic occurrence and impact. This disease is widespread, probably found wherever begonias are grown, and it significantly damages the aesthetic quality of affected plants. The obligate plant pathogenic organism exhibits dimorphism. That is, the asexual phase of growth looks significantly different from the sexual stage (Bélanger et al. 2002). The asexually reproducing phase of the pathogen is named Oidium begoniae. The sexual stage is thought to be a member of the genus Microsphaera because the conidia of the asexual stage are not formed in chains (Quinn and Powell 1981), but the structures formed as a result of sexual reproduction (chasmothecia; formerly cleistothecium) are rarely found associated with begonia. It is known that Sphaerotheca fuliginea (Powell 1985) and Golovinomyces cichoracearum (formerly Erysiphe cichoracearum) (Sammons et al. 1982) may also cause powdery mildew in begonia. Symptoms/signs. White, mealy fungal growth develops on leaves, flowers, and stems (Fig. 2). Tissue beneath the fungus may die. Fig. 2 Powdery mildew on begonia (The Pennsylvania State University Department of Plant Pathology and Environmental Microbiology # 2017. All Rights Reserved.)

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Biology and epidemiology. Disease develops most rapidly when temperatures are 20–21  C and is greatly inhibited if temperatures are above 28  C (Quinn and Powell 1982). Unlike essentially all other fungal plant pathogens, wetness has little influence on disease development of this powdery mildew. In fact, it flourishes when leaf surfaces are relatively dry. Severe powdery mildew outbreaks should be anticipated when cool, damp night conditions alternate with warm, dry days. Management. Examine plants carefully and frequently to detect the onset of disease. Maintain a fungicide program to protect plants. It is known that horticultural oil can kill powdery mildew fungi when it comes in direct contact with the pathogens. Once the oil dries on the plant surface, it has no activity against powdery mildew. That is, it has no residual activity. Applied under the wrong conditions or when the plant is wilting, oils are phytotoxic.

2.4

Pythium Root Rot (Pythium sp.)

Geographic occurrence and impact. Several species of Pythium have been reported to cause root rot and lower stem rot of begonia around the world including P. ultimum (Globisporangium ultimum) and P. splendens (Globisporangium splendens) (Farr et al. 1989). Phytopythium helicoides (formerly, Pythium helicoides) has also been found on begonias (Yang et al. 2013). Symptoms/signs. Infected seedlings die. Shiny tan, water-soaked areas develop on the stems and petioles of established plants at or just above the soil line as plants collapse and die (Middleton 1942). Under very high humidity conditions, the pathogen may be seen as a fluffy, white mass near the soil line but this is unusual to see. Biology and epidemiology. Pythium species are often found in field soil, sand taken from streams and rivers, and can be found in pond and lake sediments, and dead roots of previous crops (Ivors and Moorman 2014). Excessively wet potting mixes greatly favor the development of root rot. Management. Plant in pasteurized, not sterilized, potting media. If the potting mix has been sterilized (killing all living organisms in it) by heating it to too high a temperature or heating it too long, a biological vacuum will have been created. If Pythium then contaminates the sterile potting mix, it can cause very severe crop losses, because it has no competition and no natural inhibition by other microbes. Keep hose ends off the ground in order to avoid picking up Pythium-contaminated soil and then spraying it onto the crop. Use a well-drained potting mix and do not over-water the plants, particularly if plants are not utilizing a great deal of

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water because of prevailing weather conditions. Apply a fungicide or biological control agent at planting for the best protection. If the crop is grown for several months, any chemical or biological control agent will have to be applied repeatedly. Listed below are additional fungus or fungus-like organisms which have been associated with Begonia (Farr et al. 1989). For the most up-to-date listing, search http://nt.ars-grin.gov/fungaldatabases/fungushost/fungushost.cfm: Aecidium begoniae Agrobacterium tumefaciens Alternaria sp. Alternaria tenuis (Alternaria alternata) Armillaria mellea Bartalinia begoniae Botryosphaeria sp. Botryotinia fuckeliana (Botrytis cinerea) Ceratobasidium sp. Cercospora begoniae Cercospora sigesbeckiae Choanephora cucurbitarum Cladosporium inconspicuum Cladosporium sphaerospermum Coleosporium begoniae Colletotrichum capsici (Colletotrichum truncatum) Colletotrichum gloeosporioides Colletotrichum sp. Corticium solani (Rhizoctonia solani) Corynespora cassiicola Curvularia inaequalis Erysiphe begoniae Erysiphe begoniicola Erysiphe communis (Erysiphe pisi var. pisi) Erysiphe orontii (Golovinomyces orontii) Erysiphe polygoni Erysiphe polyphaga (Golovinomyces orontii) Fusarium begoniae Fusarium equiseti Fusarium roseum Fusarium solani Gloeosporium begoniae Glomerella cingulata (Colletotrichum gloeosporioides) Golovinomyces orontii Helminthosporium sp. Lasiodiplodia theobromae

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Diseases of Begonia

Macrophoma sp. Macrophomina phaseoli (Macrophomina phaseolina) Marssonina mali Meliola begoniae Microsphaera begoniae (Erysiphe begoniicola) Moniliopsis aderholdii (Rhizoctonia solani) Mycena citricolor Myrothecium roridum Oidium begoniae (Golovinomyces orontii) Oidium ellipticum Penicillium bacillisporum (Talaromyces bacillisporus) Pestalotiopsis sp. Phoma sorghina (Epicoccum sorghi) Phoma sp. Phomopsis sp. Phyllachora begoniae Phyllosticta begoniae Phyllosticta sp. Phyllostictina sp. Phytophthora cactorum Phytophthora cryptogea Phytophthora cryptogea f. sp. begoniae Phytophthora nicotianae var. nicotianae (Phytophthora nicotianae) Phytophthora niederhauserii Phytophthora parasitica (Phytophthora nicotianae) Pseudoidium sp. Pucciniastrum boehmeriae Pythium aphanidermatum Pythium debaryanum (Globisporangium debaryanum) Pythium intermedium (Globisporangium intermedium) Pythium irregulare (Globisporangium irregulare) Pythium vexans (Phytopythium vexans) Rhizoctonia solani Sclerotinia sclerotiorum Sclerotium rolfsii (Athelia rolfsii) Septonema sp. Septoria begoniae Shrungabeeja begoniae Sphaeropsis begoniicola Stemphylium sp. Thanatephorus cucumeris (Rhizoctonia solani) Thielaviopsis basicola Verticillium albo-atrum Verticillium dahliae

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Bacterial and Phytoplasma Diseases

Where possible, the scientific names used are those accepted by the International Society of Plant Pathology Committee on the Taxonomy of Plant Pathogenic Bacteria (Bull et al. 2010).

3.1

Bacterial Spot (Bacterial Blight; Xanthomonas campestris pv. begoniae (Takimoto) Dye)

Geographic occurrence and impact. This disease, one of the most damaging to begonias, was first described in the 1920s (Powell 1985) (Fig. 3). It causes a leaf spot or blighting of some cultivars but can be systemic in Rieger elatior begonias (Harri et al. 1977). In some cases such as a slight infection, the disease is easily overlooked. Thus, the pathogen can be inadvertently shipped with plants if infection goes undetected. Symptoms/signs. Water-soaked circular lesions surrounded by yellow halos develop on leaves. In wax begonias (semperflorens types), the spots may be very small (1 mm diameter) (Daughtrey et al. 1995). Plants slowly die one leaf at a time. If infection begins at the leaf margin, wedge-shaped brown dead areas develop with the wide part of the wedge at the margin and the point of the wedge pointing toward the plant stem. When infected tissue is cut, placed in a clear drop of water on a microscope slide, and observed with a bright field microscope with the iris diaphragm mostly closed, bacteria can be seen to stream from the tissue. When Rieger begonias are infected systemically, they wilt and die. Biology and epidemiology. The bacteria can enter injured roots or leaves and become systemic in some cultivars of Rieger begonias. Eventually wilting and death of the plant occurs. In some other begonias such as Rex, the pathogen remains

Fig. 3 Bacterial blight on begonia (The Pennsylvania State University Department of Plant Pathology and Environmental Microbiology # 2017. All Rights Reserved.)

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localized in leaf spots (Jodon and Nichols 1974). The pathogen can survive for extended periods of time if the leaf debris is dry. The pathogen can be spread from plant to plant on workers’ hands, clothing, and tools and has been shown to be disseminated in recycled irrigation water (Atmatjidou et al. 1991). Management. Purchase plants free of the pathogen. Discard infected plants, particularly Rieger-types which are systemically infected. Remove infected leaves from Rex and tuberous types because they are not systemically infected. Irrigate plants in a manner that keeps water off the foliage. Do not propagate from infected plants. Remove the debris from infected plants from the greenhouse or place it in a closed container. Workers should wash their hands thoroughly periodically when performing tasks on the plants and should disinfect any tools that come in contact with the plants. Additional Diseases with Bacterial Pathogens Crown gall (Rhizobium tumefaciens, formerly Agrobacterium tumefaciens) (Powell 1985) Bacterial fasciation (Rhodococcus fascians) (Powell 1985) Soft rot (Pectobacterium carotovorum, formerly, Erwinia carotovora) (Powell 1985) Additional Diseases with Phytoplasma Pathogens Phytoplasmas induce symptoms such as phyllody, shoot proliferation, small leaves and flowers, and stunted plant growth. These bacteria are vectored primarily by leafhoppers, planthoppers, and psyllids. “Candidatus Phytoplasma asteris” A phytoplasma in the 16SrI group was reported to infect begonia (Powell 1985). Shoot proliferation. A phytoplasma belonging to the 16SrIII group was associated with shoot proliferation in begonia in Brazil in 2006 (Ribeiro et al. 2006).

4

Viral Diseases

Viruses of begonia are associated with mild to severe mosaic and mottling, ringspots, leaf malformation, stem necrosis, chlorosis, and stunting. Generally viruses can be controlled by using clean propagative material, by discarding diseased plants and isolating surrounding plants and controlling vector populations for viruses that are vector transmitted. Clean potting soil needs to be used to control viruses whose vectors are soil inhabitants. The presence of viruses can be diagnosed by use of molecular tests such as reverse transcriptase PCR or immune-based assays such as ELISA or rapid immunostrips, or by indicator hosts. The EPPO Panel on “Certification of Ornamentals” developed procedures for the production of healthy carnation, pelargonium, lily, narcissus, chrysanthemum, tulip, crocus, iris, begonia, impatiens, rose, freesia, hyacinth, kalanchoe, and petunia.

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In the USA, certification programs are managed by the major commercial propagators. Pelargonium and other major flower crops are tested for the presence of the most prevalent viruses either by in house plant pathologists or by testing services contracted with by the propagators. In the USA, phytosanitary certificates are required for intra- and interstate plant movement. Data on pathogens of high consequence, including viruses, are recorded by the National Plant Diagnostic Network. Where available, information was taken from Adams and Antoniw (2006).

4.1

Clover Yellow Mosaic Potexvirus (ClYMV) Described by Johnson (1942)

Geographic occurrence and impact. It is common in the Western USA and Canada (Agrawal et al. 1962; Pratt 1961). It has occasionally being imported to the UK, but it has not established in the EPPO region. Symptoms. ClYMV induces color variations and poor vigor. Biology and epidemiology. ClYMV is in the family Alphaflexiviridae. ClYMV is transmitted by infected sap but no vector is known. Reports indicate that it is also seed transmitted (Hampton 1963). Clover yellow mosaic virus is serologically distantly related to White Clover mosaic virus. Management. Use of healthy propagative material and sanitation is recommended.

4.2

Cucumber Mosaic Virus (CMV)

Geographic occurrence and impact. CMV is found worldwide, has an extremely broad host range, and can infect species in more than 100 plant families (Zitter and Murphy 2009). Symptoms/signs. Symptoms observed on infected begonia are mosaic, leaf deformation and curling, and vein clearing. Biology and epidemiology. CMV (Jacquemond 2012; Palukaitis and GarcíaArenal 2003) belongs to the family Bromoviridae. CMV is a ss + RNA virus with a tripartite genome, is aphid transmitted in a noncirculative (nonpersistent) manner (Hoggan 1933; Simons 1955; Watson and Roberts 1939), and can be seed transmitted in some plant species (Neergaard 1977). Management. Control for this virus is particularly difficult, since CMV is a common virus on many plant Families and it can be transmitted by multiple aphid species. Because aphids transmit CMV readily during probing, aphid control is

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partially efficacious at reducing the spread of the virus (Zitter and Murphy 2009). Mineral oil has been proposed to delay viral symptoms in different crops (Simons and Zitter 1980).

4.3

Impatiens Necrotic Spot Virus (INSV) and Tomato Spotted Wilt Virus (TSWV)

Geographic occurrence and impact. INSV and TSWV are prevalent in many plant families and species, making their control particularly challenging. INSV can infect around 800 plant species and TSWV more than 1,000. Symptoms/signs. Symptoms associated with INSV and TSWV are mosaic, mottling, stem necrosis, ringspots on leaves, and leaf deformation (Fig. 4). Biology and epidemiology. INSV (Law and Moyer 1990) and TSWV (Samuel et al. 1930) are in the Family Bunyaviridae. These ssRNA viruses have tripartite genomes, with negative or ambisense orientation. INSV and TSWV are transmitted by thrips in a circulative and propagative manner (Ullman et al. 1993; Wijkamp et al. 1993); thus, they can replicate in their plants as well as in their insect hosts. Management. One of the best ways to manage these viruses is to control thrips numbers, since thrips need a relatively long acquisition and transmission time, and since the viruses encounter a latent period in the vector prior to becoming transmissible. For instance, in the Netherlands, where INSV and TSWV have been reported on begonia, the incidence of disease has decreased significantly thanks to the implementation of a rigorous thrips management program. The Netherlands has also established a certification program for many ornamental plants, including begonia. To complicate matters, many thrips are insecticide resistant (Brødsgaard 1994; Zhao et al. 1995); thus, care needs to be taken to rotate different classes Fig. 4 Impatiens necrotic spot virus on begonia (The Pennsylvania State University Department of Plant Pathology and Environmental Microbiology # 2017. All Rights Reserved.)

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of insecticides. Weed management is also necessary (Bond et al. 1983; Cho et al. 1986; Kobatake et al. 1984), since weeds can serve as reservoirs for tospoviruses and thrips.

4.4

Tobacco Mosaic Virus (TMV)

Geographic occurrence and impact. TMV can infect a variety of plants in 30 Families worldwide (Shew and Lucas 1991). Symptoms/signs. Symptoms on begonia are reported to be yellowing, necrosis, mosaic, leaf distortion, and plant stunting. Biology and epidemiology. TMV belong to the Virgaviridae family. The virus has a ss + RNA genome. Symptoms attributed to TMV are often caused by other Tobamoviruses, and many Tobamovirus species were once classified as strains of TMV. TMV is occasionally transmitted by chewing insects, but most commonly it is mechanically spread (Harris and Bradley 1973; Lojek and Orlob 1969), in fact TMV virions are extremely stable. TMV can persist in the soil, probably on plant debris, and can infect roots. It can penetrate wounded embryos from infected seed coats (Broadbent 1965). TMV is a special concern in greenhouses, where it can be very hard to eradicate (Broadbent and Fletcher 1963). Management. Sanitation is the best TMV control method.

4.5

Tobacco Necrosis Necrovirus (TNV) and Carnation Mottle Carmovirus (CarMV)

Geographic occurrence and impact. TNV (Price 1940) can experimentally infect 88 species in 37 plant families and CarMV can infect members in more than 9 families of plants worldwide. Symptoms/signs. On most hosts TNV causes necrotic lesions and rarely systemic symptoms. TNV can support coinfection with satellite viruses. Carnation mottle virus can infect Begonia elatior and Begonia x cheimantha where it is associated with symptoms of vein clearing, leaf curling, and flower breaking. Biology and epidemiology. These viruses are in the Family Tombusviridae. Members of this family have a ss + RNA monopartite genome. TNV (described first by Smith and Bald 1935 and more recently by Fraenkel-Conrat 1988) is transmitted by the zoospores of the fungus-like microorganism Olpidium brassicae (Kassanis and MacFarlane 1964; Teakle 1962; Teakle and Gold 1963), is not transmitted by seed or by pollen, and can be transmitted mechanically. CarMV is not known to be transmitted by a vector or by seed, but it is probably transmitted by plant to plant contact and mechanically during cultural practices. The

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virus is also stable in water and can be transmitted through irrigation. An unconfirmed report from India indicates that aphids could be vectors of this virus. Management. To control TNV, care should be taken in using clean irrigation water and potting soil, and in avoiding diseased propagative material. Control of CarMV consists of using certified virus-free material, strict sanitation, and care in handling infected plants.

4.6

Tobacco Ringspot Virus (TRSV) and Arabis Mosaic Nepovirus (ArMV)

Geographic occurrence and impact. ArMV (Smith and Markham 1944) has been detected in 13 States in the USA where it has been the object of quarantine, but it is not widespread or established outside Europe. ArMV has a wide host range. TRSV (Fromme et al. 1927) geographic distribution is mainly in northern USA and China, but it has also been found in Europe and Australia. More than 17 plant families are susceptible to TRSV (Price 1940). Symptoms/signs. Begonia yellow spot, caused by TRSV, shows symptoms of chlorotic local lesions and yellow mottle patterns, sometimes resembling natural variegation. Biology and epidemiology. ArMV and TRSV belong to the family Secoviridae. Their genomes consist of two linear ss + RNA segments. ARMV is nematode and seed transmitted and can be transmitted by Cuscuta sp. Nematodes lose the virus during molting and do not transmit it to their progeny. TRSV is transmitted by nematodes (McGuire 1964), pollen, and seeds as well as by insects (Dunleavy 1957; Komuro and Iwaki 1968; Messieha 1969; Schuster 1963) and mites (Thomas 1969) in a nonspecific manner. TRSV does not replicate in its nematode vector, is lost during molting, and is not transmitted to the nematode progeny. Management. TRSV is controlled by using clean potting soil, discarding infected propagative material, and by using clean seed.

4.7

Broad Bean Wilt Virus (BBWV) Described by Stubbs (1947)

Geographic occurrence and impact. The virus has been reported worldwide (Lisa and Boccardo 1996). The virus was reported in the USA in New York state, South Carolina and Minnesota in the early 1980s. BBWV was subdivided in two species, BBWV-1 and BBWV-2, in 2000. The two species are serologically distinct, with BBWV-1 more prevalent in Europe, while BBWV-2 is more prevalent in North America, Asia, and Australia. BBWV has a broad host range (Edwardson and Christie 1991).

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Symptoms/signs. Symptoms associated with BBWV on begonia are leaf mottling, faint ringspots, color-break in flowers, and generalized stunting. Symptoms tend to be more severe in winter. Biology and epidemiology. As ArMV and TRSV, BBWV is in the family Secoviridae, and its genome consists of two ss + RNA segments. In contrast to ArMV and TRSV, BBWV is aphid transmitted in a noncirculative (nonpersistent) manner (Lisa and Boccardo 1996). Management. Control of BBWV is difficult because its host range is broad and it is transmitted in a nonpersistent manner; thus, exclusion of aphids is advised, as well as of weeds that serve as virus reservoirs.

4.8

Zucchini Yellow Mosaic Virus (ZYMV) Described by Lisa et al. (1981)

Geographic occurrence and impact. The virus was detected on begonia in Taipei in 2008. As for CMV, another nonpersistently aphid transmitted virus, several plant families are susceptible to ZYMV and the virus is widely distributed, making its control difficult. Symptoms. Symptoms in begonia consist of faint ringspots on leaves at the early stage of infection; the spots become chlorotic and coalesce during disease progression. Biology and epidemiology. ZYMV is a potyvirus in the family Potyviridae. Its genome consists of a ss + RNA filament and is monopartite. The virus is aphid transmitted in a noncirculative (nonpersistent) manner. Management. While the use of insecticides is not recommended to control nonpersistent viruses and not many experiments have been reported on control of ZYMV in begonia, strategies used to control this virus on other crops would be probably appropriate. These strategies are: the use of resistant material, the use of clean propagative material and adoption of good hygiene practices, the use of mineral oil, and the exclusion of aphids from greenhouses.

5

Nematode Diseases

5.1

Foliar Nematode (Aphelenchoides fragariae)

Geographic occurrence and impact. Aphelenchoides affects a large number of ornamental plants, including begonia. In some cases, the nematode invades the

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vascular tissue and the begonia remains mostly symptomless. Thus, the nematode can be inadvertently shipped from place to place in infected plants. Severe damage can occur in begonias. Symptoms/signs. Plants may be stunted. In some cases, excessive red pigmentation develops in infected leaves. Bronzed or water-soaked areas develop on leaves of some cultivars. Fibrous-rooted cultivars have small brown leaf spots. Some cultivars exhibit no symptoms despite heavy infection. If leaf or stem tissue is placed in a drop of water in a clear dish and teased into small pieces, numerous colorless nematodes can be seen with some magnification, vigorously moving in a snake-like motion in the water. The nematodes will tend to settle to the bottom of the dish. Biology and epidemiology. This nematode can enter stomata or wounds, continue to move in the plant, and feed on cells inside the plant (migratory endoparasitic life style). Or the nematode can remain on the surface of the plant, continue to move at will if there is water on the plant tissue, and feed on surface cells (migratory ectoparasitic life style). There is some suggestion that the nematode may have the ability to enter intact leaf surfaces directly (Riedel 1985). Peirson (1974) found that the “Aphrodite Rose” cultivar of Rieger begonia can harbor 12,000 larvae of this nematode per gram of fresh weight of leaves, yet exhibit no symptoms. Management. Purchase nematode-free plants. Irrigate plants in a manner that keeps water off the foliage so that movement of the nematodes on the leaf surface is inhibited and localized dispersal among closely spaced plants by splashing is avoided. Discard infected plants. A very high value stock plant can be heat-treated to eliminate nematodes (Guba and Gilgut 1938). Some experimentation is required to determine the best temperature and exposure duration for eliminating the nematodes while not killing the plant. An effective hot water dip may range from 1 min at 49  C (120  F) to 5 min at 45  C (115  F), depending upon the cultivar. A treated plant may be severely damaged, but it will recover over time and be free of the nematode. Additional Diseases with Nematode Pathogens Root-knot (Meloidogyne spp.) (Powell 1985)

6

Abiotic Diseases

A variety of leaf symptoms develop as a result of nutrient deficiencies in begonia. If yellowing is the predominant symptom and the yellowing is between the veins, iron or magnesium may be lacking. Uniformly yellowing leaves indicates nitrogen or calcium deficiency. Dead tissue at the leaf margin may indicate potassium deficiency, while stunting of an otherwise green plant could be due to lack of phosphorus. A lack of boron causes the foliage to russet and cracks may develop in the leaf petioles and the plants may be very brittle (Nelson et al. 1977).

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Quinn JA, Powell CC (1981) Identification and host range of powdery mildew of begonia. Plant Dis 65:68–70 Quinn JA, Powell CC (1982) Effects of temperature, light, and relative humidity on powdery mildew of begonia. Phytopathology 72:480–484 Ribeiro LFC, Mello APD, Bedendo IP, Gioria R (2006) Phytoplasma associated with shoot proliferation in Begonia. Sci Agric 63(5):475–477 Riedel RM (1985) Nematode problems. Diseases of floral crops. In: Strider DL (ed) Diseases of floral crops, vol 1. Praeger Scientific, New York, pp 295–312 Sammons B, Rissler JF, Shanks JB (1982) Development of gray mold of poinsettia and powdery mildew of begonia and rose under split night temperatures. Plant Dis 66:776–777 Samuel G, Bald JG, Pittman HH (1930) Investigations on ‘spotted wilt’ of tomatoes. Aust Counc Sci Ind Res Bull 44:8–11 Schroers HJ, Baayen RP, Meffert JP, de Gruyter J, Hooftman M, O’Donnell K (2004) Fusarium foetens, a new species pathogenic to begonia elatior hybrids (Begonia  hiemalis) and the sister taxon of the Fusarium oxysporum species complex. Mycologia 96(2):393–406 Schuster MF (1963) Flea beetle transmission of tobacco ringspot virus in the Lower Rio Grande Valley. Plant Dis Rep 47:510–511 Sekine T, Kanno H, Aoki T (2008) Occurrence of a leaf and stem rot caused by Fusarium foetens in begoia elatior hybrids (Begonia  hiemalis). Jpn J Phytopathol 74:164–166 Shew HD, Lucas GB (1991) Compendium of tobacco diseases. APS Press, St. Paul Simons JN (1955) Some plant-vector-virus relationships of southern cucumber mosaic virus. Phytopathology 45:217–219 Simons JN, Zitter TA (1980) Use of oils to control aphid-borne viruses. Plant Dis 64:542–546 Smith KM, Bald JG (1935) A description of a necrotic virus disease affecting tobacco and other plants. Parasitology 27:231–245 Smith KM, Markham R (1944) Two new viruses affecting tobacco and other plants. Phytopathology 34:324–329 Stubbs IL (1947) A destructive vascular wilt virus disease of broad bean (Vicia faba L) in Victoria. J Dept Agric Vic 46:323–332 Teakle DS (1962) Transmission of tobacco necrosis virus by a fungus, Olpidium brassicae. Virology 18:224–231 Teakle DS, Gold AH (1963) Further studies of Olpidium as a vector of tobacco necrosis virus. Virology 19:310–315 Thomas C (1969) Transmission of tobacco ringospot virus by Tetranycus sp. Phytopathology 59:633–636 Thompson ML, Thompson EJ (1981) Begonias: the complete reference guide. Times Books, New York Tian X, Zheng Y (2012) Species susceptibility and biological control of Fusarium wilt of Hiemalis begonias in Canada. Can J Plant Pathol 34:345–346 Tian X, Zheng Y (2013) Evaluation of biological control agents for Fusarium wilt in Hiemalis begonia. Can J Plant Pathol 35:363–370 Tian XL, Dixon M, Zheng YB (2012) Susceptibility of various potted begonias to Fusarium foetens. Can J Plant Pathol 34:248–254 Tompkins CM (1950) Botrytis stem rot of tuberous-rooted begonia. Hilgardia 19:401–410 Tschope B, Hey M, Wohanka W, Hennig F (2007) Characterisation and identification of Fusarium foetens, causative agent of wilting and stem rot of begonia elatior hybrids (Begonia  hiemalis) by its volatile compounds. Eur J Hortic Sci 72(4):152–157 Ullman DE, German TL, Sherwood JL, Westcot DM, Cantone FA (1993) Tospovirus replication in insect vector cells: immunocytochemical evidence that the nonstructural protein encoded by the S RNA of tomato spotted wilt tospovirus is present in thrips vector cells. Phytopathology 83:456–463

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Van der Gaag D, Raak M (2010) Pest risk assessment Fusarium foetens. Plant Protection Service, Ministry of Agriculture, Nature and Food Quality, Wageningen. Bulletin 11-16495 Watson MA, Roberts FM (1939) A comparative study of the transmission of Hyoscyamus virus 3, potato virus Y and cucumber virus 1 by the vector Myzus persicae (Sulz.), M. circumflexus (Buckton) and Macrosiphum gei (Koch). Proc R Soc Ser B 127:543–576 Wijkamp I, van Lent J, Kormelink R, Goldbach R, Peters D (1993) Multiplication of tomato spotted wilt virus in its insect vector, Frankliniella occidentalis. J Gen Virol 74:341–349 Yang X, Richardson PA, Olson HA, Hong CX (2013) Root and stem rot of begonia caused by Phytopythium helicoides in Virginia. Plant Dis 97(10):1385 Zhao G, Liu W, Brown JM, Knowles CO (1995) Insecticide resistance in field and laboratory strains of western flower thrips (Thysanoptera: Thripidae). J Econ Entomol 88:1164–1170 Zitter TA, Murphy JF (2009) Cucumber mosaic. Plant Health Instructor. doi:10.1094/PHI-I-20090518-01

Diseases of Coleus

31

Blair R. Harlan and Mary K. Hausbeck

Abstract

Coleus (Plectranthus scutellarioides), an herbaceous bedding plant, has been prized by gardeners for its bright colorful foliage since Victorian times and in recent years has seen a resurgence in popularity. At retail greenhouses, coleus can be purchased as flats of mixed cultivars, hanging baskets, or individually potted plants. The most serious pathogen of coleus is downy mildew. Downy mildew on coleus, identified as a Peronospora sp. based on morphological characteristics, was first reported in the United States in 2005. Coleus are not considered to be susceptible to the pathogens commonly associated with other bedding plants. However, Rhizoctonia crown and stem blight, Botrytis gray mold, and select viruses have been observed. Management of coleus diseases in the greenhouse and landscape usually centers on the control of downy mildew and includes chemical and cultural control and host resistance. Keywords

Downy mildew • Botrytis • INSV • Pythium • Rhizoctonia • Scutellarioides • Herbaceous

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Botrytis Blight (Botrytis cinerea) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Downy Mildew (Peronospora sp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Pythium Root Rot (Pythium spp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Rhizoctonia Root and Crown Rot (Rhizoctonia solani) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

912 912 912 914 916 918

B.R. Harlan (*) • M.K. Hausbeck Department of Plant, Soil and Microbial Sciences, Michigan State University, East Lansing, MI, USA e-mail: [email protected]; [email protected] # Springer International Publishing AG 2018 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florists’ Crops Diseases, Handbook of Plant Disease Management, https://doi.org/10.1007/978-3-319-39670-5_32

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3 Viral Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Bromoviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Bunyaviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Potyviridae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4 Betaflexivirdae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Viroid Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Pospiviroidae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Introduction

Coleus (Plectranthus scutellarioides L. (Codd)) is a species of flowering plant in the family Lamiaceae. Coleus is a popular foliage plant grown in gardens, containers, and interiorscapes throughout the United States. In recent years, coleus has become increasing popular with gardeners who are drawn to its vibrantly colored foliage and variegation patterns. The 2014 wholesale value of coleus in the United States was $14.3 million (Anonymous 2015). Coleus plants are considered to be tropical to subtropical, preferring temperatures ranging between 16  C and 24  C (61  C and 75  F) (Pedley and Pedley 1974). Temperatures higher than 24  C will not negatively affect growth, although temperatures lower than 10  C/50  F result in decreased vigor. Coleus can survive in most sunlight conditions; however, foliage becomes more vibrant when shaded. Coleus is commercially propagated with vegetative cuttings or by seed. Prior to the emergence of downy mildew as a pathogen, coleus was relatively easy to grow with minimal chemical inputs needed during cultivation. Coleus grown in the United States may be treated with fungicides to prevent and limit downy mildew infection.

2

Fungal and Fungus-Like Diseases

2.1

Botrytis Blight (Botrytis cinerea)

Geographic occurrence and impact. This disease has been officially reported in Alaska; however, due to the endemic nature of this pathogen, it is likely much more widely distributed (Anonymous 1960). Symptoms/signs. Beginning as small water-soaked spots on leaves, stems, or blossoms, the spots coalesce rapidly, affecting large portions of tissue (Hausbeck and Moorman 1996). Leaf blight may be initiated when infected and senescent tissue falls onto healthy leaves. Stem blight typically begins in a broken or cut stem surface and progresses downward, causing a dieback of the entire stem; in severe cases, the blight extends into the base of the plant and kills it. B. cinerea readily sporulates on necrotic and diseased tissue, appearing as a fuzzy or powdery gray mold. This sporulation is quite different when compared to downy mildew sporulation, which tends to be sparser and only on the underside of the leaf surface. When sporulation of

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B. cinerea is extensive over the plant, a cloud of gray conidia typically can be observed if the plant is physically disturbed. Biology and epidemiology. Conidia, ascospores, mycelial fragments, and sclerotia all contribute to dispersal of B. cinerea (Jarvis 1980). In the majority of cases, conidia serve as the initial inoculum for B. cinerea infection, and although conidia can land on plant surfaces and invade immediately, they may remain dormant before germinating (Hausbeck and Moorman 1996). Hyphae may spread the disease when infected senescent plant parts fall onto healthy plant parts. Chlamydospores are thick-walled spores that can remain dormant in crop debris and give rise to new infections when conditions are favorable. A rise and subsequent decrease in relative humidity have been correlated with high counts of atmospheric B. cinerea conidia. Once the conidia land on a suitable host, under moist conditions, germ tubes are produced that can penetrate the plant surface leading to infection. Management • Cultural control – Shorten the intervals between scouting events during periods of high relative humidity. Be vigilant in watching for stem infections after pinching or pruning. Watering in the morning so that the foliage can dry rapidly is one way to minimize Botrytis. Reducing relative humidity by spacing plants farther apart and providing good air circulation can be helpful (Westcott 1979). Removing leaves with the sporulating pathogen from plants may reduce the amount inoculum for future infections. • Sanitation – Sanitation is important and includes removing fading flowers and blighted foliage (Westcott 1979). Plant debris should not be allowed to accumulate on greenhouse benches and floors. This plant debris supports sporulating B. cinerea and allows continued release and dispersal of conidia (Hausbeck and Moorman 1996). Keep waste containers that contain old plant material closed and away from greenhouse intake vents. • Fungicides and biocontrols – Due to the relative infrequency of Botrytis blight on coleus, fungicide treatments may not be necessary. However, if Botrytis is observed on coleus, fungicides can effectively limit the spread of the pathogen. For best results, apply fungicides at the highest label rate. In a trial conducted on geraniums by Hausbeck and Harlan (2010), applications of effective fungicides significantly reduced infection compared to untreated plants. The most effective active treatments included chlorothalonil, fenhexamid, thiophanate-methyl, fludioxonil/cyprodinil, and pyraclostrobin/boscalid. However, resistance to benzimidazole and dicarboximide fungicides has been well documented and should be avoided as standalone treatments (Moorman and Lease 1992). The biopesticide polyoxin D zinc salt has been shown to be highly effective in controlling Botrytis in greenhouse crops (Hausbeck et al. 2002). Other biopesticides have shown varying degrees of efficacy; Bacillus subtilis was effective in limiting Botrytis lesions in a trial conducted by Webster and Hausbeck (2004). • Resistance – At this time there is no published information on the resistance of coleus to B. cinerea.

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Downy Mildew (Peronospora sp.)

Geographic occurrence and impact. This disease has been reported in Japan, the USA (Louisiana, New York, Michigan, Florida, Tennessee), the United Kingdom (Daughtrey et al. 2006; Denton et al. 2015; Harlan and Hausbeck 2011; Ito et al. 2015; Palmateer et al. 2007; Rivera et al. 2016), and Germany (Anonymous 2008). Symptoms/signs. Downy mildew has been observed on both seed-propagated and vegetatively propagated coleus. Symptoms of downy mildew infection include plant stunting, leaf distortion, and leaf abscission (Harlan and Hausbeck 2011). Necrotic lesions may be present on infected leaf tissue. Under high relative humidity, downy mildew-like growth may be observed on the undersides of infected leaves, often prior to any signs of necrosis (Fig. 1). Leaf abscission may occur on older infected leaves, giving the plant a “palm tree” shape. Downy mildew can be difficult to detect in some cases as the pathogen may become latent within an infected plant. These latently infected plants may appear healthy until environmental conditions become favorable for the growth and sporulation of the pathogen (Harlan and Hausbeck 2013).

Fig. 1 Downy mildew signs and symptoms: The most recognizable sign of downy mildew on coleus is sporulation observed on the underside of leaves (top left). Infected leaf tissue often displays necrotic lesions (top left, top right, lower right). Symptoms of infection include stunting (bottom left) and leaf abscission (bottom right) (B.R. Harlan # 2017. All Rights Reserved.)

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Biology and epidemiology. The causal agent of downy mildew on coleus has been classified as a Peronospora sp., from the group Peronosporaceae. Downy mildew is an obligate pathogen, meaning that the organism must live and reproduce on living plant material. Peronospora produces sporangia, which germinate by germ tubes (Shaw 1981). In most cases sporulation occurs on the underside of the leaf surface; however, under favorable environmental conditions, it can be observed on the upper leaf surface and petioles. The environment plays a crucial role in the development of downy mildew. Studies have concluded that infection and sporulation will occur readily at 15  C and 20  C (59  F and 68  F). When temperatures reached 25  C/77  F, infection and sporulation was significantly reduced. Infection and sporulation was not observed on any of the plants incubated at 30  C/86  F (Harlan et al. 2012). The prevalence of atmospheric downy mildew sporangia in relation to environmental conditions has been well documented. The release of Peronospora sp. sporangia has been correlated to long dew periods and periods of leaf wetness in field-grown ornamentals (Byrne et al. 2005). Greenhouse studies have shown increased levels of coleus downy mildew sporangia in the atmosphere when high relative humidity (>95%) was followed by lower relative humidity. Like other Peronospora spp. (Pinckard 1942), sporangial release of coleus downy mildew followed a daily periodicity with sporangial counts highest between 1000 and 1500 h. Management • Cultural control – Coleus should be scouted on a regular basis for signs and symptoms of downy mildew. Infected plants should be kept separate and isolated from healthy plants, as the sporangia can be airborne. Intervals between scouting events should be shortened during periods of high relative humidity. Environmental conditions that may limit downy mildew include keeping relative humidity below 85% and temperatures at or over 25  C/77  F. Limiting extended leaf wetness periods by watering in the morning or early afternoon will ensure that the foliage does not remain wet overnight. Increased plant spacing will ensure better ventilation and shorter leaf wetness periods. • Sanitation – Removing downy mildew-infected plant material is essential (Harlan and Hausbeck 2011). Although Peronospora is an obligate pathogen, needing live plant material to reproduce, the sporangia on abscised leaves may be viable for a short time. • Fungicides and biocontrols – If effective fungicides are used, downy mildew on coleus can be adequately controlled (Harlan and Hausbeck 2010). Fungicide active ingredients that have been shown to be highly effective in limiting Peronospora infection on coleus include mefenoxam, dimethomorph, fluopicolide, azoxystrobin, fosetyl-al, mandipropamid, fluoxastrobin, and fenamidone (Harlan and Hausbeck 2010). Research on other Peronospora species has shown that drench applications of mefenoxam, fluopicolide, and oxathiapiprolin have offered long-term protection from infection. In particular, these applications, when applied to greenhouse plants, have offered protection from downy mildew long after transplanting into the landscape (Harlan and

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Hausbeck 2016). Due to the prevalence of the coleus downy mildew pathogen in recent years, preventive fungicide applications are warranted. Biocontrol products have been tested against Peronospora spp. with limited success. An extract of Reynoutria sachalinensis, when applied as a preventive spray, has shown some efficacy against coleus downy mildew (Harlan and Hausbeck 2010; Ivors et al. 2011). In a trial conducted by Warfield et al. (2008), spray applications of a phosphorous acid salts product were highly effective. • Resistance – With the vast number of different coleus cultivars commercially sold and the varying colors and shapes observed among these cultivars, it is not surprising that a wide range of susceptibility to downy mildew has been observed. Daughtrey et al. (2014) tested 147 cultivars over a multiyear study and rated 32% of the cultivars with low susceptibility, while 27.2% were rated as highly susceptible to downy mildew. In particular, all 11 cultivars in the Wizard series were rated as having medium to high susceptibility, while seven of the nine Fairway series cultivars were rated as having a low or medium susceptibility. When growing highly susceptible cultivars, it is recommended that preventive fungicides be applied prior to disease development.

2.3

Pythium Root Rot (Pythium spp.)

Geographic occurrence and impact. This disease has been reported in Florida, Mississippi, California, and Maryland (Alfieri et al. 1984; Anonymous 1960; Parris 1959). Symptoms/signs. Pythium is a water mold that can “nibble” the feeding roots of plants, resulting in wilting and stunted growth. This reduced plant vigor can eventually affect marketability (Garzon et al. 2011). In severe cases, stems of seedlings or recent transplants collapse at the soil line and die (Pscheidt and Ocamb 2016). Plants that survive the initial infection phase are likely to remain stunted. Both cutting- and seed-propagated cultivars of coleus are susceptible. If Pythium root rot is suspected on a plant showing aboveground symptoms, the root system should be inspected. Infected roots will be brown or water-soaked and sparse when compared to a healthy root system (Fig. 2). Biology and epidemiology. Pythium species are soilborne oomycete organisms that prefer wet conditions for infection and reproduction. This preference for wet conditions explains Pythium’s characterization as a water mold. Three distinct spore types are produced by Pythium: oospores, sporangia, and zoospores. Oospores are thick-walled sexually produced spores that can lay dormant for extended periods. Oospores can withstand long periods of drying, and once environmental conditions are conducive, or a suitable host is presented, they can germinate and infect via fungal-like strands. Sporangia and zoospores are the most common spore type associated with Pythium epidemics as they are produced in large numbers.

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Fig. 2 Pythium root rot symptoms: Infected roots are brown, water soaked, and sparse (M. Daughtrey # 2017. All Rights Reserved.)

Zoospores, in particular, are associated with the spread of the pathogen in ebb-andflow irrigation systems as this particular spore type is motile in water (Koike and Wilen 2009). Management • Cultural control – Overwatering, creating soil conditions that are overly saturated for extended periods, favors Pythium development. In particular, soil moisture conditions of 70% or higher are conducive to infection by Pythium. Keep fungus gnats and shore flies under control as they can spread the pathogen (Koike and Wilen 2009). • Sanitation – This pathogen can become a greenhouse “resident” that hibernates on dirty plant containers, benches, hoses, and greenhouse walkways (Hausbeck and Harlan 2013). The reuse of flats, pots, and containers is not recommended; however, if they are reused, they should be thoroughly disinfected with a cleaning agent such as quaternary ammonium chloride salts or chlorine bleach. (Refer to ▶ Chap. 9, “Sanitation for Management of Florists’ Crops Diseases” for additional information about disinfectants.) Plants with a suspected Pythium infection should be isolated from healthy plants to reduce the further spread of the pathogen. • Fungicides and biocontrols – Unfortunately, there are very few effective fungicides registered against Pythium root rot. When attempting to control root rot pathogens with fungicide drenches, it is important to make applications to the soil. Applications of the low risk fungicide mefenoxam can be highly effective in limiting Pythium infection. However, resistance of the pathogen to mefenoxam is an issue (Del Castillo Munera and Hausbeck 2016). Etridiazole is another product that is highly effective against Pythium. Other products, including fluopicolide, fenamidone, and phosphorous acid salts, have shown promise in managing Pythium (Hausbeck and Harlan 2013).

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Although using biocontrol against Pythium in a greenhouse can be helpful, studies have shown the level of control to be inconsistent. For instance, in a study published by Daughtrey and Tobiasz (2001), four popular biocontrol products were tested against Pythium, and all received ratings statistically similar to the untreated control plants. However, Baysal-Gurel and Miller (2013) found applications of biocontrol products effective in limiting Pythium damping-off in the greenhouse. • Resistance – At this time there is no published information on the resistance of coleus to Pythium spp.

2.4

Rhizoctonia Root and Crown Rot (Rhizoctonia solani)

Geographic occurrence and impact. This disease has been reported in Florida, Illinois, New York, and Texas (Alfieri et al. 1984; Anonymous 1960). Symptoms/signs. Rhizoctonia can cause pre- and postemergence damping-off of coleus. Younger plants are more susceptible (Wright 2013). Infected stems will appear blackened and water-soaked (Pscheidt and Ocamb 2016). In severe cases, Rhizoctonia can move into the plant canopy, causing spots on leaves and petioles. Root rot symptoms are similar to those caused by Pythium, and a diagnostic clinic may be necessary to differentiate with which pathogen you are dealing. Biology and epidemiology. Rhizoctonia is a saprophytic fungal pathogen in the phylum Basidiomycota that is associated with several greenhouse crops throughout the world. Rhizoctonia is not host-specific, meaning one isolate can infect several different species of plants (Wright 2013). Unlike many of the other pathogens associated with greenhouse production, spores are not the usual method of dissemination for Rhizoctonia. Mycelia, the hairlike fungal strands produced by Rhizoctonia, are spread by the movement of infested soil particles, often by workers, which grow and infect nearby hosts. Sclerotia, the resting structure produced by Rhizoctonia, can survive for extended periods in soil particles. Rhizoctonia can thrive under almost all greenhouse conditions, although warmer, more humid conditions may enhance its spread. Prolonged periods of high humidity can result in the pathogen growing from the soil, up the stem, and into the foliage canopy. Management • Cultural control – One of the primary sources of Rhizoctonia inoculum in the greenhouse are infested trays (Gutierrez et al. 2001). The reuse of trays, pots, and other containers is not recommended; however, if they are reused, they should be thoroughly disinfected with a cleaning agent such as quaternary ammonium chloride salts or chlorine bleach. • Sanitation – Between crops, clean soil, and organic material from benchtops and floors as this material can harbor the pathogen for extended periods. Because Rhizoctonia does not produce spores, one of the more common methods of

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transmission is by workers. All pruning tools should be disinfected between crops and foot baths should be installed between greenhouses. Greenhouse carts should also be thoroughly cleaned. • Fungicides and biocontrols – Moorman and Lease (1999) conducted a study looking at soil drenches to control Rhizoctonia rot on coleus and found that azoxystrobin and thiophanate-methyl were effective at limiting infection. Other products that have consistently reduced Rhizoctonia on ornamental plants include fludioxonil, PCNB, metconazole, and pyraclostrobin. The type of symptoms observed should determine the fungicide application method; drench applications should be used for root and crown rots, while high volume sprays or “sprenches” should be used to control foliar symptoms. The biopesticide, polyoxin D zinc salt, is highly effective against Rhizoctonia (Hausbeck and Harlan 2015). Using biocontrols in potting mixes may be helpful in managing damping-off when the plants are still young and more susceptible to the pathogen. • Resistance – At this time there is no published information on the resistance of coleus to Rhizoctonia.

3

Viral Diseases

3.1

Bromoviridae

Geographic occurrence and impact. Viruses in the Bromoviridae are some of the most economically important and widely distributed viruses in the world. One member of this group has been reported in coleus: Cucumber mosaic virus (CMV, genus: Cucumovirus). Cucumber mosaic virus has been reported in Louisiana (Holcomb and Valverde 1991). Symptoms/signs. Symptoms of CMV infection of coleus include ring-spotting, mosaic, and oak-leaf symptoms. Biology and epidemiology. CMV is spread by more than 80 aphid species (Hemiptera: Aphididae) in a non-circulative (nonpersistent) manner. When these insects feed on CMV-infected plants, the virus moves into the mouthparts of the vector and can spread when the insect moves and feeds on adjacent plants. Although seed transmission has been reported on other crops (Neergaard 1977), it has not been reported or studied on coleus. Management. There is no cure or chemical control for plant viruses. New shipments of plants should be inspected for symptoms of viruses upon arrival. Vector control is the most important management aspect associated with CMV on coleus. This management plan should include monitoring insect populations and applying chemical controls when needed. Insect populations can be tested for CMV infection.

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Stock plants should be tested regularly for CMV, and infected plants should be discarded. Refer to ▶ Chap. 4, “Insect Management for Disease Control in Florists’ Crops” for additional virus management information.

3.2

Bunyaviridae

Geographic occurrence and impact. Viruses belonging to the Bunyaviridae include those most commonly associated with ornamental crops. Impatiens necrotic spot virus (INSV, genus: Tospovirus) has been reported on coleus in New York (Catlin 2014), Michigan (Byrne 2008), and France (Lambert 2005). Symptoms/signs. Ring spots are the most typical symptom associated with INSV on coleus (Fig. 3). Other symptoms of INSV include leaf mottle, stem or petiole lesions, irregularly shaped necrotic lesions, or plant stunting (Catlin 2014). Biology and epidemiology. INSV is spread by the western flower thrips (Frankliniella occidentalis) as well as two closely related species F. fusca and F. intonsa. Unlike viruses in the Bromoviridae family, INSV enters the gut of the vector where it replicates. Thrips must be in their juvenile stage (first and early second larval stages) to acquire the virus, and only immature thrips that acquire these viruses or adults derived from such immatures are vectors. Management. There is no cure or chemical control for plant viruses. New shipments of plants should be inspected for symptoms of viruses upon arrival. Western

Fig. 3 Symptoms of INSV: ring spots and irregularly shaped necrotic spots (left, L. Pundt © 2017. All Rights Reserved.; right, M. Daughtrey # 2017. All Rights Reserved.)

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flower thrips populations should be closely monitored. In particular, flowers should be inspected, as thrips are likely to reside in such protected areas. If thrips are detected, chemical controls might be necessary. Insect populations can be tested for INSV incidence. Since almost all herbaceous plants associated with horticulture can harbor INSV, greenhouses should be scouted regularly for signs of infection. Any plants showing symptoms of INSV should be disposed of immediately.

3.3

Potyviridae

Geographic occurrence and impact. Potyviruses are fairly common and infect many crops. One member of this group has been recently discovered in coleus: Tobacco etch virus (TEV, genus: Potyvirus). TEV has been reported on coleus in Missouri and Minnesota (Lockhart et al. 2010). Symptoms/signs. Symptoms of TEV are similar to those associated with coleus vein necrosis virus and include foliar and veinal lesions. Biology and epidemiology. At this time, a number of aphid species are the only known vectors of TEV (Sikora 2004). The virus has a broad host range (>120 species in 19 dicot families) including a number of solanaceous crops such as pepper, tomato, and tobacco. Management. There is no cure or chemical control for plant viruses. New shipments of plants should be inspected for symptoms of viruses upon arrival. Greenhouses should be scouted regularly for aphids. If aphids are detected, chemical controls might be necessary. Due to the rarity of this virus on coleus, plants showing symptoms of a viral infection should first be tested for INSV and CMV.

3.4

Betaflexivirdae

Geographic occurrence and impact. Of the virus families associated with coleus, viruses in the Betaflexivirdae are the least common and economically important. A new virus was observed on coleus in 2005 and was named coleus vein necrosis virus (CVNV, genus: Carlavirus) by Mollov et al. (2007). Symptoms/signs. Symptoms of CVNV on coleus include abnormal leaf coloration, vein necrosis, and ring patterns (Fig. 4). Biology and epidemiology. It is unknown how this virus is spread in the environment. In laboratory studies, mechanical viral transmission was successful. However, when aphids (Myzus persicae) were tested as a possible vector, transmission of CVNV was not observed (Mollov et al. 2007).

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Fig. 4 Symptoms of CVNV: abnormal leaf coloration and ring patterns (M. Daughtrey # 2017. All Rights Reserved.)

Management. There is no cure or chemical control for plant viruses. New shipments of plants should be inspected for symptoms of viruses upon arrival. Due to the rarity of CVNV, if virus symptoms are observed on coleus, testing for INSV and CMV should be completed as they are much more common. At this time, there is not a commercially available test for CVNV.

4

Viroid Diseases

4.1

Pospiviroidae

Geographic occurrence and impact. Viroids are small, single-stranded sections of RNA that do not have protein coats. The coleus blumei viroids (CbVd, genus: Coleviroid) are widely distributed and have been classified as six different species, CbVd-1 through CbVd-6. CbVd-1 was first detected in Brazil in 1989 and is the most common species and has been reported worldwide (Fonseca et al. 1989; Tsushima and Sano 2015). CbVds has been reported in Germany, China, India, Indonesia, and Japan (Spieker et al. 1990; Hou et al. 2009; Jiang et al. 2013; Tsushima and Sano 2015). Symptoms/signs. A wide range of symptoms can be observed on CbVd-infected coleus. In fact, CbVd is often detected on plants that appear to be symptomless and healthy (Fu et al. 2011). Stunting has been identified as a symptom, although yellowing or chlorosis-type symptoms are the most common (Tsushima and Sano 2015). This yellowing can be observed on the leaves in patches, often irregular in shape (Adkar-Purushothama et al. 2013). Biology and epidemiology. Very little is known on how this pathogen infects and spreads on coleus. Jiang et al. (2014) did show that seed transmission is possible and vegetative cuttings are likely to spread the viroid.

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Management. There is no cure or chemical control for plant viroids. Getting seed and cuttings from stock plants that have been tested for CbVd will reduce the spread of the disease.

References Adkar-Purushothama CR, Nagaraja H, Sreenivasa MY, Sano T (2013) First report of Coleus blumei viroid infecting coleus in India. Plant Dis 97:149 Alfieri SA Jr, Langdon KR, Wehlburg C, Kimbrough JW (1984) Index of plant diseases in Florida. Bulletin No 11 (revised). Florida Department of Agriculture and Consumer Services, Division of Plant Industry, Gainesville, p 389 Anonymous (1960) Index of plant diseases in the United States. USDA handbook no 165. Washington, DC, p 531 Anonymous (2008) Coleus downy mildew. Plant Clinic News, July. Central Science Laboratory. http://fera.co.uk/news/resources/documents/PCN%20-%20plantClinicNews0708.pdf. Accessed 25 Mar 2016 Anonymous (2015) Floriculture crops 2014 summary. USDA National Agricultural Statistics Service. http://usda.mannlib.cornell.edu/MannUsda/viewDocumentInfo.do?documentID=1072. Accessed 7 Oct 2016 Baysal-Gurel F, Miller SA (2013) Evaluation of fungicides and biorational products for management of Pythium and Rhizoctonia damping-off in greenhouse-produced vegetables. Phytopathology 103(Suppl 2):S2.13 Byrne J (2008) Diagnostic update for greenhouse samples. Michigan State University Extension News for Agriculture-Floriculture, Mar 28. http://msue.anr.msu.edu/news/diagnostic_update_ for_greenhouse_samples. Accessed 7 Oct 2016 Byrne JM, Hausbeck MK, Sconyers LE (2005) Influence of environment on atmospheric concentrations of Peronospora antirrhini sporangia in field-grown snapdragon. Plant Dis 89:1060–1066 Catlin N (2014) INSV on coleus. e-GRO Alert 3(37). http://www.e-gro.org/pdf/337.pdf. Accessed 7 Oct 2016 Daughtrey ML, Tobiasz M (2001) Biocontrol effects against Pythium root rot of seedling geraniums, 2001. Biol Cult Tests 17:O09 Daughtrey ML, Harlan B, Linderman S, Hausbeck MK (2014) Coleus cultivars and downy mildew. Special research report #136. American Floral Endowment, Disease Management. http://endowment.org/wp-content/uploads/2013/03/136-ColeusDM-Cv-2014.pdf. Accessed 7 Oct 2016 Daughtrey ML, Holcomb GE, Eshenaur B, Palm ME, Belbahri L, Lefort F (2006) First report of downy mildew on greenhouse and landscape coleus caused by a Peronospora sp. in Louisiana and New York. Plant Dis 90:1111 Del Castillo Munera J, Hausbeck MK (2016) Characterization of Pythium species associated with greenhouse floriculture crops in Michigan. Plant Dis 100:569–576 Denton GJ, Beal E, Denton JO, Clover G (2015) First record of downy mildew, caused by Peronospora belbahrii, on Solenostemon scutellarioides in the UK. New Dis Rep 31:14 Fonseca MEN, Boiteaux LS, Singh RP, Kitajima EW (1989) A small viroid in Coleus species from Brazil. Fitopatol Bras 14:94–96 Fu FH, Li SF, Jiang DM, Wang HQ, Liu AQ, Sang LW (2011) First report of Coleus blumei viroid 2 from commercial coleus in China. Plant Dis 95:494 Garzon CD, Molineros JE, Yanez JM, Flores FJ, Del Mar Jimenez-Gasco M, Moorman GW (2011) Sublethal doses of mefenoxam enhance Pythium damping-off of geranium. Plant Dis 85:1233–1238

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Gutierrez WA, Shew HD, Melton TA (2001) Rhizoctonia diseases in tobacco greenhouses. North Carolina State University Plant Pathology Extension TB07, Tobacco Disease Note 7. http:// www.ces.ncsu.edu/depts/pp/notes/Tobacco/tdin007/tb07.html. Accessed 7 Oct 2016 Harlan BR, Hausbeck MK (2010) Evaluation of foliar sprays and soil drenches of fungicides for the control of downy mildew of coleus, 2009. Plant Dis Manag Rep 4:OT018 Harlan BR, Hausbeck MK (2011) Is coleus downy mildew here to stay? Michigan State University Extension News for Agriculture-Floriculture, Feb 8. http://msue.anr.msu.edu/news/is_coleus_ downy_mildew_here_to_stay. Accessed 7 Oct 2016 Harlan BR, Hausbeck MK (2013) Understanding coleus downy mildew. Special research report #134. American Floral Endowment, Disease Management. http://endowment.org/wp-content/ uploads/2014/03/134ColeusDMDisease2013.pdf. Accessed 7 Oct 2016 Harlan BR, Granke L, Hausbeck MK (2012) Epidemiology and management of downy mildew, a new pathogen of coleus in the United States. Acta Hortic 952:813–818 Harlan BR, Hausbeck MK (2016) Epidemiology and management of impatiens downy mildew in the United States. Acta Hortic 952:813–818 Hausbeck MK, Harlan BR (2010) Evaluation of registered and unregistered fungicides for control of Botrytis blight of geranium, 2009. Plant Dis Manag Rep 4:OT011 Hausbeck MK, Harlan BR (2013) Pythium root rot in the greenhouse. Michigan State University Extension News for Agriculture-Floriculture, Oct 7. http://msue.anr.msu.edu/news/pythium_ root_rot_in_the_greenhouse. Accessed 7 Oct 2016 Hausbeck MK, Harlan BR (2015) Greenhouse disease control update. In: Greenhouse session summaries, proceedings of the 2015 Michigan Greenhouse Expo, Grand Rapids, pp 1–7. http://glexpo.com/summaries/2015summaries/Greenhouse_DiseaseControl.pdf. Accessed 7 Oct 2016 Hausbeck MK, Moorman GW (1996) Managing Botrytis in greenhouse-grown flower crops. Plant Dis 80:1212–1219 Hausbeck MK, Quackenbush W, Linderman SD (2002) Evaluation of biopesticide for control of Botrytis blight of geranium, 2002. Fungicide Nematicide Tests 58:OT028 Holcomb GE, Valverde RA (1991) Identification of a virus causing a mosaic on coleus. Plant Dis 75:1183–1185 Hou WY, Li SF, Wu ZJ, Jiang DM, Sano T (2009) Coleus blumei viroid 6: a new tentative member of the genus Coleviroid derived from natural genome shuffling. Arch Virol 154:993–997 Ito Y, Takeuchi T, Matsushita Y, Chikuo Y, Satou M (2015) Downy mildew of coleus caused by Peronospora belbahrii in Japan. J Gen Plant Pathol 81:328–330. http://link.springer.com/ article/10.1007/s10327-015-0601-3. Accessed 25 Mar 2016 Ivors K, Lacy LW, Milks DC (2011) Evaluation of fungicides for the control of downy mildew on coleus, 2010. Plant Dis Manag Rep 5:OT019 Jarvis WR (1980) Epidemiolgy. In: Coley-Smith JR, Verhoeff K, Jarvis WR (eds) The biology of Botrytis. Academic, London, pp 219–250 Jiang DM, Li SF, Fu FH, Wu ZJ, Xie LH (2013) First report of Coleus blumei viroid 5 from Coleus blumei in India and Indonesia. Plant Dis 97:561 Jiang D, Rui Gao R, Qin LV, Wu Z, Xie L, Hou W, Li S (2014) Infectious cDNA clones of four viroids in Coleus blumei and molecular characterization of their progeny. Virus Res 180:97–101 Koike ST, Wilen CA (2009) Pythium root rot – UC IPM pest management guidelines: floriculture and ornamental nurseries. UC ANR publication 3392. http://www.ipm.ucdavis.edu/PMG/ r280100211.html. Accessed 7 Oct 2016 Lambert L (2005) Virus: plantes sensibles. Reseau D’Avretissements Phytosanitaires Bulletin d’information Cultures En Serres No 2. https://www.agrireseau.net/horticulture-serre/docu ments/68143. Accessed 7 Oct 2016 Lockhart BEL, Mason SL, Johnson DA, Mollov DS (2010) First report of tobacco etch virus infection in coleus in the United States. Plant Dis 94(7):921 Mollov DS, Hayslett MC, Eichstaedt KA, Beckman NG, Daughtrey ML, Lockhart BE (2007) Identification and characterization of a carlavirus causing veinal necrosis of coleus. Plant Dis 91:754–757

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Moorman GW, Lease RJ (1992) Benzimidazole- and dicarboximide-resistant Botrytis cinerea from Pennsylvania greenhouses. Plant Dis 76:477–480 Moorman GW, Lease RJ (1999) Control of Rhizoctonia root rot of coleus, 1999. Fungicide Nematicide Tests 56:OT5 Neergaard P (1977) Seed pathology, vol 1. MacMillan, London, p 839 Palmateer AJ, Harmon PF, Schubert TS (2007) Downy mildew of coleus (Solenostemon scutellarioides) caused by Peronospora sp. in Florida. New Dis Rep 16:11 Parris GK (1959) A revised host index of Mississippi plant diseases. Mississippi State University, Botany Department Miscellaneous Publication 1, pp 146 Pedley R, Pedley K (1974) Coleus: a guide to cultivation and identification. Bartholomew and Son, Edinburgh, p 116 Pinckard JA (1942) The mechanism of spore dispersal in Peronospora tabacina and certain other downy mildew fungi. Phytopathology 32:505–511 Pscheidt JW, Ocamb CM (2016) Pacific northwest plant disease management handbook. http:// pnwhandbooks.org/plantdisease/. Accessed 7 Oct 2016 Rivera Y, Salgado-Salazar C, Windham AS, Crouch JA (2016) Downy mildew on coleus (Plectranthus scutellarioides) caused by Peronospora belbahrii sensu lato in Tennessee. Plant Dis. doi:10.1094/PDIS-10-15-1120-PDN Shaw CG (1981) Taxonomy and evolution. In: Spencer DM (ed) The Downy mildews. Academic, San Francisco, pp 17–29 Sikora EJ (2004) Tobacco etch virus. Alabama cooperative extension ANR-869. http://www.aces. edu/pubs/docs/A/ANR-0869/ANR-0869.pdf. Accessed 7 Oct 2016 Spieker RL, Haas B, Charng Y-C, Freimuller K, Sanger HL (1990) Primary and secondary structure of a new viroid ‘species’ (CbVd1) present in the Coleus blumei cultivar ‘Bienvenue.’. Nucleic Acids Res 18:3998 Tsushima T, Sano T (2015) First report of Coleus blumei viroid 5 infection in vegetatively propagated clonal coleus cv. ‘Aurora Black Cherry’ in Japan. New Dis Rep 32:7 Warfield CY, Sugar JC, Sugar KJ (2008) Evaluation of fungicides for the control of downy mildew on coleus, 2007. Plant Dis Manag Rep 2:OT004 Webster BJ, Hausbeck MK (2004) Evaluation of reduced risk fungicides and biopesticides for control of Botrytis blight of geranium, 2004. Fungicide Nematicide Tests 60:OT009 Westcott C (1979) Westcott’s plant disease handbook, 4th edn. Van Nostrand Reinhold Company, New York, p 112 Wright J (2013) Greenhouse diseases 101: Rhizoctonia. Greenhouse grower. http://www.greenhou segrower.com/production/crop-inputs/greenhouse-diseases-101-rhizoctonia/. Accessed 2 Feb 2016

Diseases of Gardenia

32

A. J. Palmateer and A. R. Chase

Abstract

Gardenias are a very important crop for both potted flowers for the florist trade and the landscape industries. Gardenias are susceptible to root rot, and the most common and damaging pathogens include Phytophthora and Rhizoctonia. One of the most devastating diseases affecting gardenias is canker caused by Diaporthe gardeniae, which can lead to unsightly cankers and galls on the lower stem causing plant decline and eventual death. Several leaf spot pathogens are reported on gardenia with Myrothecium and Xanthomonas being among the most common. Disease management practices for gardenia depend on good sanitation, judicious use of pesticides, and cultural practices. Keywords

Gardenia • Phytophthora • Myrothecium • Pythium • Rhizoctonia • Gardenia canker • Xanthomonas

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Gardenia Canker (Diaporthe gardeniae = Phomopsis gardeniae) . . . . . . . . . . . . . . . . . . 2.2 Myrothecium Leaf Spot (Myrothecium roridum) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Powdery Mildew (Erysiphe polygoni, Oidium sp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Rhizoctonia Root Rot (Rhizoctonia solani, Rhizoctonia sp.) . . . . . . . . . . . . . . . . . . . . . . . .

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A.J. Palmateer (*) Department of Plant Pathology, Tropical Research and Education Center, University of Florida, Homestead, FL, USA e-mail: ajp@ufl.edu A.R. Chase Chase Agricultural Consulting LLC, Cottonwood, AZ, USA e-mail: [email protected] # Springer International Publishing AG 2018 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florists’ Crops Diseases, Handbook of Plant Disease Management, https://doi.org/10.1007/978-3-319-39670-5_33

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2.5 Phytophthora Root and Stem Rot (Phytophthora cinnamomi and P. nicotianae) . . . 2.6 Pythium Root Rot (Pythium splendens, P. spinosum) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Bacterial and Phytoplasma Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Bacterial Leaf Spot (Xanthomonas maculifoliigardeniae) . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Abiotic Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Iron Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Bud Drop . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Introduction

Gardenia jasminoides is a popular flowering shrub in the Rubiaceae family that can be found growing in tropical and semitropical areas in the landscape worldwide (Fig. 1). It is also grown as a potted flowering crop by the florist industry in temperate climates using protected culture. Patio gardenias are normally not grafted and cuttings are grown in pots. Some cultivars are also grown as standards where the apical meristem is pruned to form a head. Landscape plantings are mostly grafted to the root stock Gardenia thunbergia for resistance to root-knot nematodes and Rhizoctonia spp.

2

Fungal and Fungus-Like Diseases

2.1

Gardenia Canker (Diaporthe gardeniae = Phomopsis gardeniae)

Geographic occurrence and impact. This disease has been reported within the continental USA; gardenia canker has been reported in California (Hansen and Barrett 1938), Washington, Nebraska, Kansas, Ohio, Massachusetts, and Florida, and has also been reported in Europe (Italy) (Alfieri 1967) and South America. Fig. 1 Healthy Gardenia jasminoides with showy white blooms (A.J. Palmateer # 2017. All Rights Reserved.)

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Symptoms/signs. Initial symptoms of gardenia canker include wilting, yellowing, shriveling, and defoliation of leaves and frequent premature bud drop of unopened flowers. The wilting can affect a portion of the plant or the entire plant where it can appear as a sudden wilt. The cankers are at first small, circular to oblong, swollen brownish lesions on the main stem and branches. Cankers can become as large as 5 cm (2 in.) in diameter and cause partial to complete girdling of stems and branches. Even partial girdling can result in stunting and wilt. The diseased area decays and begins to separate from trunk or branch tissue exposing the wood. In some cases, the periphery of the canker becomes calloused and appears rough and corrugated. The cankers become overgrown with corky callous tissue extending longitudinally in both directions, increasing in size and forming galls. These galls typically increase two to three times in size and produce deep longitudinal cracks. Branches in close proximity to galls begin to lose vigor compared to those further away or originating from unaffected stem tissue. Stem cankers are more common at the base of the plant affecting the crown, but they can occur anywhere on the plant especially where wounded from mechanical injury. The foliage of older infected plants appears dull eventually turning yellow or drying out and becoming brown before defoliating. Flower buds will abscise before opening. Disease severity and wilting is usually more pronounced under cooler temperatures. During warmer conditions diseased plants may continue to live, but remain stunted and appear unhealthy. Sometimes signs of the imperfect stage of the fungus (Phomopsis gardeniae) are present consisting of black fruiting bodies on the lesions (pycnidia) that contain both alpha and beta conidia. Biology and epidemiology. The fungal pathogen is widely considered to be a wound parasite; however, once infection occurs and the fungus becomes established in host tissue, disease develops and progresses forming mature cankers and subsequent pycnidia. Under conditions of high relative humidity and warm temperatures, an abundance of spores forms within the partially submerged pycnidia and is easily spread by windblown rain and splash dispersion in irrigation water. The fungus is most prevalent on or near cankers and overwinters from one season to the next on the diseased portions of infected plants. The fungus is notorious for entering gardenias through leaf joints at the base of cuttings. Thus, freshly cut leaf bases act as an excellent point of entry for the fungus. Management • Cultural practices – Disease-free stock plants should be used for propagation. Pathogen elimination through sanitation by steam-pasteurizing rooting media, sand, and peat is highly recommended. All cuttings should be taken with sharp blades avoiding rough and jagged edges where spores of the pathogen are more likely to gain entry. All tools contacting plant material should be properly disinfected to minimize contamination and potential spread of the pathogen. Plant injury due to careless cultural practices should be avoided. The productivity of affected plants has been shown to be enhanced or lengthened by piling soil up

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around the stem base covering basal cankers to encourage new root formation above the diseased portions of stems. • Fungicides and biological controls – Preventative applications of fungicides containing chlorothalonil (FRAC M5), mancozeb (FRAC M3), propiconazole (FRAC 3), or thiophanate methyl (FRAC 1) ensuring excellent coverage of all plant parts may minimize disease outbreaks. In addition, the combination of pyraclostrobin (FRAC 11) and boscalid (FRAC 7) has been effective on similar pathogens on other crops. It is important to rotate fungicides from different classes in order to help prevent the development of resistance. Scientific literature reports good efficacy against Phomopsis on other plant species using biological control products containing Gliocladium virens. These products were applied as a soil drench and as a wound treatment.

2.2

Myrothecium Leaf Spot (Myrothecium roridum)

Geographic occurrence and impact. This disease was found and described from Pennsylvania in 1941 (Fergus 1957) and has since been found in California (Barrett and Hardman 1947) and Florida. Symptoms/signs. The leaf spots are mostly circular and vary in size up to 2 cm (~1 in.) in diameter (Fergus 1957). Initial spots appear water soaked and turn brown with age. Often the centers of the lesions fall out giving a shot hole appearance. Under wet conditions, the spots coalesce and large portions of the leaves can rot away. Characteristic greenish-black pillow-shaped sporodochia with white setal margins form on necrotic tissue and can be seen on both the upper and lower leaf surface (Fig. 2). Biology and epidemiology. Myrothecium leaf spot on gardenia is most common under humid conditions such as those found when attempting to root cuttings. Fig. 2 Myrothecium roridum on gardenia leaves (A.J. Palmateer # 2017. All Rights Reserved.)

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Pathogenicity of M. roridum isolates is reported to vary greatly with most isolates being wound pathogens on gardenia. Management • Cultural practices – Monitor nutritional programs closely, because overfertilization has been shown to increase leaf spots and rots caused by Myrothecium in some tropical foliage plants. Imbalanced fertilization programs, damage from pesticides, heat or cold injury, and leaf necrosis caused by water stress should be avoided. Wounded or weakened tissue allows Myrothecium to form a colony in the canopy, which serves as the focal point for disease spread. • Fungicides and biological control – Apply fungicides containing azoxystrobin or pyraclostrobin (FRAC 11), chlorothalonil (FRAC M5), fludioxonil (FRAC 12), iprodione (FRAC 2), or mancozeb (FRAC M3). Good coverage is very important as this fungus can colonize both sides of the leaf surface. No effective biocontrol agents have been reported for this disease.

2.3

Powdery Mildew (Erysiphe polygoni, Oidium sp.)

Geographic occurrence and impact. This disease has been reported in Texas and tropical America but is not common. Symptoms/signs. Powdery mildew of gardenia primarily affects young leaves and shoots where symptoms of infection include leaf yellowing, slight deformation of leaf and bud tissue, and leaf drop. Powdery mildew fungi typically grow on the surface of plants and infect plant cells using a haustorium to absorb nutrients. Signs of infection include a white to gray colored fungal growth that appears like powder and is most often found on the upper leaf surface but may occur on the underside of leaves when conditions are highly favorable. Biology and epidemiology. Powdery mildew fungi can survive in leaf litter. High relative humidity and moderate to warm temperatures favor outbreaks of powdery mildew. Shade especially during periods of high relative humidity and moderate temperatures are most favorable conditions for powdery mildew. Management • Cultural practices – Proper location of plants in full to partial sun and avoiding heavily shaded areas will help to prevent outbreaks of powdery mildew. Space plants apart to allow for good air movement through the plant canopy. Overhead irrigation may actually reduce the spread of powdery mildew, because the water, unlike with other fungi, actually inhibits spore germination. Closely monitor plants and examine new shoot growth for signs of powdery mildew. • Fungicides and biological controls – The use of fungicides may be necessary in production situations when favorable conditions persist. Horticultural oil or plantbased oil such as neem oil can be used to control powdery mildew. Be certain

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not to apply oil within 14 days of a sulfur spray or plant injury may occur. Never apply oil when ambient temperatures exceed (30  C/86 F) or when plants are water-stressed. Sulfur-based products are commonly used for controlling powdery mildew and should be used only as a preventative before signs of powdery mildew appear. Fungicides with efficacy for controlling powdery mildew include thiophanate methyl (FRAC 1); copper (FRAC M1); myclobutanil, propiconazole, triadimefon (FRAC 3); azoxystrobin, pyraclostrobin, trifloxystrobin (FRAC 11); chlorothalonil (FRAC M5); and Bacillus spp. (FRAC 44).

2.4

Rhizoctonia Root Rot (Rhizoctonia solani, Rhizoctonia sp.)

Geographic occurrence and impact. This disease has been reported wherever gardenia is grown and is considered to be one of the most common and economically important diseases affecting gardenia. Specific reports from the USA (California, Florida, New Jersey), Greece, and Japan have been made. Symptoms/signs. Common symptoms in cutting beds include damping-off, hypocotyl rot, root rot, stunting, yellowing, and death. On older plants, Rhizoctonia causes the roots to turn light to dark brown and macerates the tissue; but the pathogen has been reported to affect all portions of the plant. Light to dark brown irregularly shaped spots form anywhere on the stem and foliage during propagation (Fig. 3) or in the landscape. Mycelium of the fungal pathogen is frequently present and looks like a light brown web colonizing affected tissue giving rise to the common name web blight. Biology and epidemiology. Rhizoctonia can survive in soil and plant debris as mycelium and sclerotia that can be stimulated to germinate by plant exudates. Rhizoctonia stem rot and aerial blight or leaf spot occurs in the warmer months especially during the rainy season when there is an abundance of moisture. Such

Fig. 3 Darkly stained stem cankers on gardenia in propagation due to Rhizoctonia solani (A.J. Palmateer # 2017. All Rights Reserved.)

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favorable conditions can occur at anytime throughout the year when plants are produced in covered structures that employ overhead irrigation. Under favorable conditions severe outbreaks are common and can occur in as little 7 days. Management • Cultural practices – Prevention is the most important step for managing Rhizoctonia. Start with high quality, healthy cuttings, use new or sterilized pots and new potting media. Placing plants on hard surfaces, ground clothes, or benches so that they are not in direct contact with soil is crucial. Due to the quick onset of this disease it is recommended to scout plants weekly for early symptom detection. • Fungicides and biological controls – Specific trials on fungicides and biocontrol agents have not been reported on gardenia for Rhizoctonia. On other woody ornamental crops, strobilurins (FRAC 11); fludioxonil (FRAC 12); and thiophanate methyl (FRAC 1) are among the most effective fungicides. Additionally, the biocontrol agent Trichoderma harzianum T22 can be very effective against root rot but less so against stem rot or cutting rot.

2.5

Phytophthora Root and Stem Rot (Phytophthora cinnamomi and P. nicotianae)

Geographic occurrence and impact. Phytophthora cinnamomi is typically the most common species associated with container and field grown gardenias in the Southeastern USA, but other species have been widely reported to cause root and stem rot of gardenia. P. nicotianae has been reported from the USA in North Carolina (Olson and Benson 2011), California, and Florida and also Greece. Symptoms/signs. Phytophthora usually reduces the overall volume of roots that are present, thus greatly inhibiting the plant’s ability to take up water and nutrients for proper growth. The roots of diseased plants appear brittle and brown to reddish brown in color (Fig. 4a). The pathogen usually colonizes the crown often girdling the stem at or just above the soil line. A brown to reddish brown discoloration of the tissues occurs just below the bark layer and may extend up the stem above the soil. Affected tissue is most often wet in appearance. Aboveground symptoms can easily be confused with those caused by a nutritional disorder, overwatering, drought stress, and other abiotic disorders. Slight yellowing of the leaves followed by wilting and possibly plant death (Fig. 4b) are common symptoms associated with Phytophthora. Landscape or field grown gardenias may show symptoms of general decline for up to 1 year or more before succumbing to root rot caused by Phytophthora. Biology and epidemiology. Phytophthora species are fungus-like water molds or Oomycetes that most often survive as resting structures consisting of chlamydospores, oospores, and mycelia in diseased roots, crowns, and other plant debris. These structures are released into the soil or potting medium from infested crop

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Fig. 4 (a) Brittle and brown diseased roots due to infection by Phytophthora nicotianae. (b) Necrotic and wilted leaves on gardenia with stem infection from Phytophthora nicotianae (A.J. Palmateer # 2017. All Rights Reserved.)

debris and can be easily spread throughout propagation and container production areas in windblown plant and potting material and by splashing rain and irrigation water. Heaviest losses occur in poorly drained areas where there is standing water. Overwatering in the nursery and landscape are contributing factors to losses from Phytophthora. Management • Cultural practices – An integrated approach is necessary for success in controlling Phytophthora root rot. Prevention and sanitation are key, because once symptoms appear it is often too late to adequately control the disease. Providing for proper plant establishment and following good horticultural practices including adequate nutrition and irrigation will help to reduce potential losses from this disease. Recycling irrigation water does not appear to be a significant source of disease in gardenias. • Fungicides and biological controls – Fungicides should be used as a preventative measure, and those used for managing Phytophthora include fosetylaluminum, phosphorous acid (FRAC 33); fluopicolide (FRAC 43); mefenoxam (FRAC 4); dimethomorph, mandipropamid (FRAC 40); chlorothalonil (FRAC M5); azoxystrobin, pyraclostrobin, fenamidone (FRAC 11); etridiazole (FRAC 14); propamocarb hydrochloride (FRAC 28); cyazofamid (FRAC 21); oxathiapiprolin (FRAC U5); and Bacillus spp. (FRAC 44).

2.6

Pythium Root Rot (Pythium splendens, P. spinosum)

Geographic occurrence and impact. Pythium root rot is common in most gardenia production areas although losses are usually minimal and damage is not as severe when compared to that caused by Phytophthora. Symptoms/signs. The aboveground symptoms often include yellowing of the oldest leaves first followed by defoliation and in severe cases wilting (Fig. 5a).

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Fig. 5 (a) Pythium root rot exhibiting severe leaf chlorosis and wilting caused by P. splendens (A.J. Palmateer # 2017. All Rights Reserved.). (b) Pythium root rot on gardenia showing typical sparse, necrotic, and discolored roots (A.R. Chase # 2017. All Rights Reserved.). (c) Outer cortex of roots infected with Pythium spp. often deteriorates leaving the light colored central core (A.R. Chase # 2017. All Rights Reserved.)

The plants typically appear unhealthy for an extended period of time and may not respond to fertilization due to Pythium attacking the feeder roots and inhibiting the plant’s ability to take up water and nutrients. Healthy roots are white whereas diseased roots appear brown and discolored, or may be completely missing (Fig. 5b). The outer cortical tissue can be easily removed when pulling on the roots (Fig. 5c), leaving behind the threadlike core or stele of the water conducting tissue. Biology and epidemiology. The biology and epidemiology is very similar to Phytophthora. Disease development is highly favored by any factor that encourages wet soil conditions, including poor drainage and over watering. Planting gardenias too deep also contributes to the disease. Management • Cultural practices – An integrated approach is necessary for success in controlling Pythium root rot. Closely monitoring irrigation and using a potting medium that allows for adequate drainage or aeration is key. Providing for proper plant

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establishment and following good horticultural practices including adequate nutrition and irrigation will help to reduce potential losses from this disease. Recycling irrigation water does not appear to be a significant source of disease in gardenias. • Fungicides and biological controls – Fungicides should be used as a preventative measure and those used for managing Pythium include fosetyl-aluminum, phosphorous acid (FRAC 33); fluopicolide (FRAC 43); mefenoxam (FRAC 4); dimethomorph, mandipropamid (FRAC 40); chlorothalonil (FRAC M5); azoxystrobin, pyraclostrobin, fenamidone (FRAC 11); etridiazole (FRAC 14); propamocarb (FRAC 28); cyazofamid (FRAC 21); oxathiapiprolin (FRAC U5); and Bacillus spp. (FRAC 44).

3

Bacterial and Phytoplasma Diseases

3.1

Bacterial Leaf Spot (Xanthomonas maculifoliigardeniae)

Geographic occurrence and impact. This disease was reported for the first time in California in 1941 (Burkholder and Pirone 1941) on greenhouse grown gardenias. It was later described by Ark and Barrett in 1946. It is commonly found in production and the landscape in Florida and California. Symptoms/signs. Very young tender leaves are the first to develop symptoms which consist of small yellow spots. The yellow spots are angular to circular and gradually increase in size where the center initially appears water soaked. As the leaves age, the center portion of the spots turns reddish brown and necrotic. Lesions are most often surrounded by a wide chlorotic halo (Fig. 6). Spots eventually coalesce and produce larger necrotic areas. Premature leaf drop is common on severely affected plants. Biology and epidemiology. Conditions providing high relative humidity and temperature, especially in greenhouse production where gardenias are induced to bloom, favor disease development. Symptoms may appear rather suddenly on gardenias growing outdoors during warm and wet periods. Management • Cultural practices – Overhead irrigation should be avoided or timed for periods when leaves dry quickly as bacterial cells are readily dispersed in irrigation water increasing the incidence of diseased plants and plant parts. The disease is most severe during periods of prolonged leaf wetness. • Bactericides and biological controls – As a preventative measure apply a bactericide containing copper hydroxide, copper oxychloride, copper sulfate, or other forms (FRAC M1); mancozeb (FRAC M3); or streptomycin sulfate (FRAC 25). The quaternary ammonium (QA) product DDAC also shows good control of

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Fig. 6 Xanthomonas leaf spot on gardenia caused by Xanthomonas maculifoliigardeniae (A.J. Palmateer # 2017. All Rights Reserved.)

bacterial diseases on other ornamentals. Possible biological control agents include species of Bacillus such as B. amyloliquifaciens and B. subtilis which have both shown good efficacy against related bacterial pathogens on other ornamentals and agronomic crops. These are less common diseases that are known to affect gardenia: Botrytis cinerea – California, Florida Hawaii, US Virgin Islands, Puerto Rico, Greece, Saudi Arabia, and China Capnodium spp. (sooty mold) – California, Georgia, US Gulf Coast States, and Venezuela Cercospora (and Pseudocercospora) – Indonesia, Venezuela, and Florida Colletotrichum spp. (anthracnose) – Hong Kong, Cina, India, and Florida Pestalotia (and Pestalotiopsis) – Alabama, Florida, Cuba, Japan, and China Phyllosticta gardeniae – Japan, Republic of Georgia, India, and the USA (Michigan, North Carolina, New Jersey, and Texas) Phytoplasma – China (Sun and Zhao 2012) Tomato spotted wilt virus – USA

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4

Abiotic Disorders

4.1

Iron Deficiency

Geographic occurrence and impact. Gardenias are an acid loving plant, so when soil pH is greater than 6.0 the potential for micronutrient deficiencies increase. The most common nutrient deficiency affecting gardenias is iron, and when soil pH is above 7 the iron may be in a form that is not readily plant-available. Symptoms/signs. Most often iron deficient gardenias are stunted and have pale green to yellow leaves. Symptoms are most obvious in younger leaves which turn completely yellow except for dark green veins. Older leaves may only turn yellow along the edges (Fig. 7). Management • Cultural practices – Anything that can be done to acidify (lower) the pH of the soil will help. Some relevant soil amendments include aluminum sulfate, iron sulfate, or wettable sulfur. Gardenias growing in soils with a high buffering capacity such as calcareous soils require foliar applications of iron chelate or ferrous sulfate (FeSO4.2H20) can be very effective for correcting iron deficiency in gardenias. Allow the soil to remain evenly moist but not completed saturated.

4.2

Bud Drop

Geographic occurrence and impact. Bud drop is a prevailing issue with gardenias and is especially common after a change in location or growing conditions. It can occur in both container and landscape gardenias.

Fig. 7 Iron deficiency shows stunted chlorotic leaves when plants are subjected to high pH (A.J. Palmateer # 2017. All Rights Reserved.)

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Fig. 8 Irregular watering or temperature fluctuations causes premature bud abortion and drop (A.R. Chase # 2017. All Rights Reserved.)

Symptoms/signs. When gardenias are under stress unopened flower buds may prematurely drop off from the plant (Fig. 8). Numerous things can cause stress, but some of the most common include insects feeding, diseases including root feeding nematodes, inadequate fertilization, irrigation and light, and temperature fluctuations such as unusually cold or hot conditions. Management • Cultural practices – Anything that can be done to minimize plant stress. Gardenias do not respond well to changes in growing conditions, so avoid moving or relocating plants that are thriving. Monitor irrigation so that the soil remains evenly moist, but not saturated.

References Alfieri SA Jr (1967) Gardenia canker, vol 54, Division of plant industry circular. Florida Department of Agriculture, Gainesville Ark PA, Barrett JT (1946) A new bacterial leaf spot of greenhouse-grown gardenias. Phytopathology 36:865–868 Barrett JT, Hardman DA (1947) Myrothecium leaf spot and canker of Gardenia. Phytopathology 37:360 Burkholder WH, Pirone PP (1941) Bacterial leaf spot of gardenia. Phytopathology 31:192–194 Fergus CL (1957) Myrothecium roridum on Gardenia. Mycologia 49:124–127 Hansen HN, Barrett JT (1938) Gardenia canker. Mycologia 30:15–19 Olson HA, Benson DM (2011) Characterization of Phytophthora spp. on floriculture crops in North Carolina. Plant Dis 95:1013–1020 Sun XC, Zhao WJ (2012) First report of a group 16Sri phytoplasma associated with Gardenia jasminoides in China. Plant Dis 96:1576

Diseases of Geranium

33

Cristina Rosa and Gary W. Moorman

Abstract

Geraniums are susceptible to a wide variety of fungi, bacteria, and viruses as well as nematodes and abiotic diseases. The systemic nature of some of the pathogens makes it likely that the diseases they cause can be found wherever vegetatively propagated geraniums are shipped. Management of these pathogens is paramount for specialty propagators, while growers purchasing plants must inspect incoming plants for symptoms and understand the biology of the pathogens involved in order to manage them effectively. Keywords

Botrytis cinerea • Pythium Spp. • Ralstonia solanacearum • Xanthomonas campestris • Virus • Edema • Culture Indexing • Virus Indexing • Certification

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Alternaria Leaf Spot (Alternaria Sp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Black Root Rot (Thielaviopsis basicola (Berk. And Br) Ferris) . . . . . . . . . . . . . . . . . . . . . 2.3 Cercospora Leaf Spot (Cercospora Sp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Gray Mold (Botrytis Blight; Botrytis cinerea Pers.: Fr.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Pythium Root Rot and Blackleg (Pythium Sp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 Rhizoctonia Root and Crown Rot (Thanatephorus cucumeris (Rhizoctonia solani Kuhn) Rhizoctonia Sp.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Rust (Puccinia pelargonii-zonalis Doidge, Puccinia granularis, Puccinia morrisoni) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

942 943 943 944 945 945 947 949 950

C. Rosa (*) • G.W. Moorman Department of Plant Pathology and Environmental Microbiology, The Pennsylvania State University, University Park, PA, USA e-mail: [email protected]; [email protected] # Springer International Publishing AG 2018 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florists’ Crops Diseases, Handbook of Plant Disease Management, https://doi.org/10.1007/978-3-319-39670-5_34

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2.8 Sclerotinia Crown Rot or Cottony Stem Rot (Sclerotinia sclerotiorum (Lib.) de Bary) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.9 Verticillium Wilt (Verticillium albo-atrum Reinke and Berthe or V. dahliae Kleb.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Bacterial and Phytoplasma Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Bacterial Blight (Xanthomonas hortorum pv. pelargonii; Formerly, Xanthomonas campestris pv. pelargonii) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Bacterial Fasciation (Rhodococcus fascians, Formerly Known as Corynebacterium fascians, Bacterium fascians, and Phytomonas fascians) . . . . . . . . 3.3 Bacterial Wilt or Southern Wilt (Ralstonia solanacearum, Formerly Known as Burkholderia solanacearum, and Pseudomonas solanacearum) . . . . . . . . . . . . . . . . . 3.4 Crown Gall (Rhizobium tumefaciens, Formerly Agrobacterium tumefaciens) . . . . . . 3.5 Pseudomonas Leaf Spot (Bacterial Leaf Spot; Pseudomonas cichorii) . . . . . . . . . . . . . 3.6 “Candidatus Phytoplasma Asteris” . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Viral Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Arabis Mosaic Virus (ArMV), Tobacco Ringspot Virus (TRSV), Tomato Ringspot Virus (ToRSV), and Artichoke Italian Latent Virus (AILV) . . . . . . . . . . . . . . . 4.2 Beet Curly Top Virus (BCTV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Impatiens Necrotic Spot Virus (INSV) and Tomato Spotted Wilt Virus (TSWV) . . . 4.4 Pelargonium Zonate Spot Virus (PZSV), Cucumber Mosaic Virus (CMV) . . . . . . . . . 4.5 Tobacco Mosaic Virus (TMV) and Tobacco Rattle Virus (TRV) . . . . . . . . . . . . . . . . . . . . 4.6 Tomato Bushy Stunt Virus (TBSV), Tobacco Necrosis Virus (TNV), Moroccan Pepper Virus (MPV), and Pelargonium Leaf Curl Virus (PLCV) . . . . . . . . . . . . . . . . . . . 4.7 Pelargonium Flower Break Virus (PFBV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Nematode Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 Abiotic Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1 Edema . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2 Heat Stress . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1

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Introduction

Species of Geranium (described by Linnaeus) are often grown as herbaceous perennials outdoors, while Pelargonium (described by L’Héritier) are generally grown indoors for long periods of time or as annuals or perennials outdoors. Emphasis in this chapter is placed on the pathogens and diseases of Pelargonium graveolens, P. peltatum, and the complex hybrids Pelargonium X domesticum and Pelargonium X hortorum. Common names for these Pelargonium include florists, garden, bedding, zonal, ivy, horseshoe, rose, and scented and regal geranium, among others depending upon the species. It is thought that Pelargonium was brought into cultivation in Europe from plants initially obtained in South Africa in the early 1600s and are well documented as having been grown in many European gardens by the mid-1700s (Laughner 1993). At one time, Pelargonium was the mainstay of the bedding plant industry, and the majority of plants were propagated vegetatively. While vegetatively propagated varieties are still grown extensively, many other varieties are grown from seed. Disease susceptibility varies greatly with the variety and the pathogen involved, and some pathogens pose more of a problem in greenhouse production than when plants are grown outdoors. The geographic distribution

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of particular pathogens reported in the literature may provide clues to the environmental conditions that favor the disease, but the worldwide trade in millions of Pelargonium cuttings greatly increases the potential for any pathogen to be found anywhere Pelargonium is grown. Crucial to the production of Pelargonium varieties through vegetative propagation is the establishment of stock plants that are free of the plant pathogens that infect systemically or stay closely associated with plant tissues including Verticillium, Xanthomonas campestris pv. pelargonii, Ralstonia solanacearum, Agrobacterium tumefaciens, Rhodococcus fascians, and all the viruses noted here. Culture indexing is used to detect those plant pathogens that readily grow in broth containing required nutrients, and virus indexing employs various methods of testing plant tissues for the presences of specific viruses (De Boer et al. 1996; Huttinga 1996; Oglevee-O’Donovan 1993). Plants free of selected pathogens are then maintained as stock plants from which numerous cuttings are taken. This “culture/virus indexing” procedure has been central in the production of “pathogen-free” Pelargonium, but note that they are “free” of selected pathogens. Other pathogens may be present in or on the plants.

2

Fungal and Fungus-like Diseases

For a list of pathogens of this plant in the USA, see Farr, et al. (1989). For the most upto-date listing, search http://nt.ars-grin.gov/fungaldatabases/fungushost/fungushost.cfm.

2.1

Alternaria Leaf Spot (Alternaria Sp.)

Geographic occurrence and impact. This disease occurs when conditions are not well suited to geranium growth, particularly if the temperatures are too high or low or when other factors favor plant senescence. Generally, economic losses are minor (Engelhard 1993; Munnecke 1956). Symptoms/signs. Water-soaked spots initially formed on the underside of the leaf enlarge to 5–10 mm (1/4–1/2 in.). Leaves may have numerous spots, and some spots may have a yellow halo. Concentric dark rings form in the spots creating a target-like pattern where the fungus has formed dark brown, club-shaped, multi-celled spores on the surface of the leaf (Fig. 1). The spots may merge and occupy most of the leaf. Diagnosis is facilitated if infected leaves are placed in a container with moisture, and the spores are allowed to develop (Engelhard 1993). Biology and epidemiology. Warm, wet conditions greatly favor infection and development of symptoms particularly on older leaves. The fungus survives on dead leaves on the soil surface, and spores are dispersed by air currents. The disease generally begins on older leaves, particularly if they are senescing, and progresses upward on the plant.

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Fig. 1 Alternaria leaf spot (The Pennsylvania State University Department of Plant Pathology and Environmental Microbiology # 2017. All Rights Reserved.)

Management. Irrigate plants in a manner that keeps water off the foliage. Ensure good air circulation among plants so that the humidity within the canopy is kept low. Remove and destroy infected leaves and any crop debris in pots, or on benches, and walkways. Apply a fungicide to protect plants.

2.2

Black Root Rot (Thielaviopsis basicola (Berk. And Br) Ferris)

Geographic occurrence and impact. This disease can occur in Pelargonium and Geranium production (Linderman 1993). In Pelargonium, seed-type geraniums vary in susceptibility. While individual growers may sustain serious losses, the occurrence of this disease appears to be very sporadic and rare, particularly with the wide use of Thielaviopsis-free, peat-based potting mixes that have replaced field soilbased mixes. Symptoms/signs. Infected plants may be symptomless or may be stunted and have yellowed older foliage. Older infected roots may be very dark brown, almost black, while infected young roots may be only slightly darkened or may appear healthy unless observed microscopically. Thielaviopsis forms very dark brown, thickwalled, cigar-shaped spores in root cells that are readily found through microscopic observation and a thin-walled colorless spore on the surface of plant tissues (Linderman 1993). Biology/epidemiology. Thielaviopsis survives well in soil. If infested soil is brought into contact with plants, infection is likely to occur if environmental conditions and the physiological status of the plant favor disease development. It appears that when plants are under high temperature and low soil moisture stress, disease is likely to develop. Though rare, Thielaviopsis has been found in peat-based potting mixes (Graham and Timmer 1991). Management • Cultural practices – Start with healthy, pathogen-free cuttings and seedlings. If the potting mix contains soil, pasteurize it before use. Black root rot is less severe

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in soil with a pH of 5–5.5, but growers should exercise caution when lowering the pH outside of the optimal range for plant growth. • Fungicides – They are available for suppressing Thielaviopsis but should be used routinely for this pathogen only if the operation has a history of crop losses from it. Discard infected plants.

2.3

Cercospora Leaf Spot (Cercospora Sp.)

Geographic occurrence and impact. The pathogen is reported to infect Pelargonium in North America and the Philippines (Daughtrey et al. 1995) and has been found on Geranium. Symptoms/signs. Initially small (1–5 mm; 1/8–1/4 in.) light green spots enlarge and become gray (Fig. 2). Dark fruiting structures develop within the gray area. As the spots enlarge, they may have a yellow area surrounding them, and the spots may merge. Infected flowers often do not open (Engelhard 1993). Biology and epidemiology. Little is known of the disease cycle of this pathogen in Pelargonium, but it is known that the pathogen is wind disseminated and that infection is greatly favored by wetness of the foliage (Daughtrey et al. 1995). Management. Irrigate plants in a manner that keeps water off the foliage. Remove infected foliage. Apply a fungicide to protect plants. The chlorothalonil and mancozeb fungicides are effective against Cercospora diseases. Check production guides for your region.

2.4

Gray Mold (Botrytis Blight; Botrytis cinerea Pers.: Fr.)

Geographic occurrence and impact. This disease is probably the most common and widespread one in Pelargonium production, and the pathogen infects many Fig. 2 Cercospora leaf spot on Pelargonium (Robert McGovern # 2017. All Rights Reserved.)

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additional plants (Daughtrey et al. 1995). In fact, the very high susceptibility of Pelargonium results in Pelargonium being a significant harbor of Botrytis for other greenhouse crops (Hausbeck and Pennypacker 1991a; Hausbeck and Pennypacker 1991b). Symptoms/signs. Flowers turn dark and fall prematurely. Where infected petals land on leaves, Botrytis grows from this nutrient source, and an irregular leaf spot forms on the green leaf (Fig. 3). Cuttings develop a dark brown to black rot (Fig. 4) near the base (also see Pythium root rot, blackleg). Abundant, dusty, gray masses of spores form on infected tissue if the humidity is high or if infected tissue is placed in a container with moisture. Biology and epidemiology. All aboveground parts of the plant are susceptible, and the fungus can infect intact tissue or through wounds. Seedlings and mature plants are susceptible. Infection occurs most readily when the fungus has a food base from which it attacks, such as fading flowers or senescent leaves. High relative humidity, moisture on the plant, and temperatures between 18  C and 25  C (64  F and 77  F) greatly favor Botrytis infection (Jarvis 1980). The spores are readily spread by air Fig. 3 Botrytis infected petal has fallen on a leaf. Botrytis grows from this food base into the healthy leaf (The Pennsylvania State University Department of Plant Pathology and Environmental Microbiology # 2017. All Rights Reserved.)

Fig. 4 Botrytis-infected Pelargonium cuttings (The Pennsylvania State University Department of Plant Pathology and Environmental Microbiology # 2017. All Rights Reserved.)

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currents, and any activity in the greenhouse that changes the relative humidity quickly or disturbs the plants can result in the release of spores (Hausbeck and Pennypacker 1991a, Hausbeck and Pennypacker 1991b). Management • Cultural practices – Heat and ventilate plants in a greenhouse setting, and space plants to ensure good air circulation and low humidity within the crop canopy. It is best to not crowd plants and to not hang plants above Pelargonium if those plants tend to drop faded flowers or if irrigating the pots above will result in excess irrigation water dripping onto the geraniums. Avoid unnecessarily damaging plants. Do not leave large stubs of tissue on the stock plant when taking cuttings. Remove and destroy fading flowers and leaves from the plant, the surface of the potting soil, from benches and greenhouse floors. Remove this material promptly from the greenhouse or place it in a closed container. • Fungicides – Apply a fungicide. Exclusive use of one class of chemical, particularly systemic chemicals, can result in the selection of resistance to the chemical class. Resistance to benzimidazole fungicides is very widespread in Botrytis populations. Do not rely on only one chemical class (Elad et al. 1992; Gullino and Garibaldi 1987; Hausbeck and Moorman 1996; Moorman and Lease 1992).

2.5

Pythium Root Rot and Blackleg (Pythium Sp.)

Geographic occurrence and impact. Many species of Pythium are known to be pathogens of Pelargonium. P. aphanidermatum, P. cryptoirregulare, P. irregulare, P. myriotylum, and P. ultimum are recovered the most frequently, but P. complectens (Phytopythium vexans), P. debaryanum (Globisporangium debaryanum), P. deliense, P. mamillatum (Globisporangium mamillatum), P. splendens (Globisporangium splendens), and P vexans (Phytopythium vexans) have reported. These species can be found worldwide, but a particular species may be more important in a given region than other species. Pythium root rot and blackleg (rot of the base of cuttings) can cause extensive crop losses. Symptoms/signs. Blackleg, the coal black rot of cuttings, first develops as brown water-soaked rot at the cut end of the tissue. The rot proceeds up the stem and kills the cutting rapidly. Seedling geraniums yellow and collapse when root rot occurs (Fig. 5). In the case of root rot of established plants, root tips appear translucent and water-soaked. The outer layers of root tissue strip off when plants are pulled from soil leaving the central core of vascular tissue bare. Usually, spherical spores can be found in root cells when examined microscopically. If Pythium is suspected but no spherical spores are observed in the tissue, plate the tissue on plain water agar, and Pythium usually grows out within 24 h. There are simple kits available commercially that can be used easily to detect the presence of Pythium sp. These immunoassay kits provide a result in less than 30 min.

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Fig. 5 Pythium root and blackleg (Robert McGovern # 2017. All Rights Reserved.)

Biology/epidemiology. Pythium species are often found in field soil, sand taken from streams and rivers and can be found in pond and lake sediments, and dead roots of previous crops (Ivors and Moorman 2014). While some species are a problem in hydroponic systems, species of Pythium present in the clear water (not the sediment) of ebb and flood irrigation systems may not all be serious plant pathogens (Lanze 2015). It is important to know what species is present. Pelargonium most susceptible are those being grown at high fertilizer levels (Gladstone and Moorman 1989, Gladstone and Moorman 1990). Management • Cultural practices – Pythium root rot is difficult to control once rot has begun. Every effort should be directed toward preventing the disease before it begins by using a heat-pasteurized potting mix. The entire pile must be heated to 75 –80  C (165 –180  F) and held at that temperature for 30 min. Potting mixes heated for very long times or temperatures higher than 80  C will result in the death of most beneficial organisms in the soil and create a “biological vacuum” that is readily filled by Pythium if the mix becomes contaminated in some way. If the commercial mix is purchased in sealed bags or bales, try to keep the covering on until used in order to prevent contamination. Pythium is easily introduced into pasteurized soil or soilless mixes by using dirty tools, dirty pots or flats, and dirty loading equipment, by walking on or allowing pets to walk on the mixes, and by dumping the mixes on benches or potting shed floors that have not been thoroughly cleaned and disinfected. Discard infected cuttings since affected rooted cuttings later develop root rot. Keep hose ends off the ground to prevent picking up contaminated soil. If pond or stream water is used for irrigation, be certain the intake pipe is well above the bottom so that sediment is not drawn in. If a known plant pathogenic species is found in the water, treatment may be required (refer to ▶ Chap. 9, “Sanitation for

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Management of Florists’ Crops Diseases,” and pertinent chapters in Hong et al. (2014)). For example, slow sand filtration has been shown to be an effective, simple, and inexpensive method for removing Pythium from water. Heat, ultraviolet light, ozone, and chlorination can also be effective. The best of these methods is the one that can be used consistently and efficiently in an operation. Disinfect all bench surfaces, potting benches, tools, and equipment that will contact the potting mix. Periodically, thoroughly clean and disinfect ebb and flood reservoirs, benches, and flood and drain floors. • Fungicides and biocontrols – In a greenhouse operation with a history of Pythium root rot, apply a fungicide or a biological control agent as early in the cropping cycle as possible. Biological agents are generally applied to the potting mix before, during, or immediately after transplanting. However, read the label of the product to obtain information on exactly when the agent should be applied. In some cases, the agent must be in the potting mix for several days before plants are put in the mix in order to avoid phytotoxicity. Some biological control agents can be applied to plants in plug trays before transplanting. If chemical fungicides are also to be used, a general guideline is to not apply any chemical pesticides to the potting mix 10 d before and 10 d after applying the biological control agent. Biological control agents and fungicides may have to be applied more than once in order to maintain adequate protection for several weeks. They do not cure a plant once the plant is infected. Populations of some Pythium species have resistance to fungicides (Moorman and Kim 2004; Moorman et al. 2002).

2.6

Rhizoctonia Root and Crown Rot (Thanatephorus cucumeris (Rhizoctonia solani Kuhn) Rhizoctonia Sp.)

Geographic occurrence and impact. Although the pathogen is well known worldwide, this disease is not widespread in Pelargonium production, particularly with the use of soilless potting mixes. Symptoms and signs. A brown, dry rot develops at the base of cuttings (Manning et al. 1973) or seedlings are killed (Powell 1993). When infected tissue is incubated on a moist surface in a closed container, threads of fungal hyphae grow from the infected tissue directly down to the moist surface in 24 h. Biology and epidemiology. Little is known of this disease. It is reported that drought and high soluble salts renders plants more susceptible to the pathogen (Powell 1993). This species infects a wide range of host plants. Management. Every effort should be directed toward preventing the disease before it begins by using a heat-pasteurized potting mix. The entire pile must be heated to 75 –80  C (165 –180  F) and held at that temperature for 30 min. If this pathogen

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has been a problem before in production, the application of a fungicide is recommended in order to protect plants not yet infected.

2.7

Rust (Puccinia pelargonii-zonalis Doidge, Puccinia granularis, Puccinia morrisoni)

Geographic occurrence and impact. The pathogen was first found in South Africa (Doidge 1926) but now can be found sporadically wherever Pelargonium is grown. However, because it is readily recognized and prevented, it is generally not a major problem. Symptoms/signs. Chlorotic specks on the upper leaf surface appear directly opposite pustules of rust-colored spores on the underside of leaf. Spores erupt in concentric rings forming a “target” spot (Fig. 6). Biology/epidemiology. This rust requires only one host (autoecious; there is no alternate host plant), and it produces only two different types of spores: brown singlecelled urediniospores and pale brown, two-celled teliospores. When temperatures are between 16  C and 21  C (61  F and 70  F) and there is water on the leaves, the fungus can continuously produce urediniospores that germinate and infect plants through the stomata (Harwood and Raabe 1979). The urediniospores can be spread by wind or splashing or on workers’ hands and clothing. Apparently, teliospores form rarely. Management. Purchase rust-free cuttings. Irrigate plants in a manner that keeps water off the foliage. Discard unwanted geraniums at season’s end unless they are to be kept under observation and treated with fungicides if necessary. When infected plants are found, discard them and treat the remaining plants with a fungicide. Fig. 6 Geranium rust on Pelargonium (The Pennsylvania State University Department of Plant Pathology and Environmental Microbiology # 2017. All Rights Reserved.)

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2.8

Diseases of Geranium

951

Sclerotinia Crown Rot or Cottony Stem Rot (Sclerotinia sclerotiorum (Lib.) de Bary)

Geographic occurrence and impact. This pathogen infects many different bedding plants in the seeding stage, including Pelargonium, where growing conditions are hot and humid. It is generally not a problem in greenhouses in temperate regions. Symptoms/signs. White, cottony fungal growth develops quickly near the soil line. As the mycelium ages, black sclerotia (up to 2–3 mm in size) develop on the infected tissue. Biology and epidemiology. The pathogen is carried in soil or on infected plants. It survives as sclerotia in crop debris and in soil. Management. If soil is a component of the potting mix, the potting mix should be heat pasteurized first. Benches, potting areas, and equipment used to move potting mix should be cleaned and disinfected periodically. Crop debris should be collected and removed from the growing area, particularly between cropping cycles. Infected plants should be discarded (Strider 1985).

2.9

Verticillium Wilt (Verticillium albo-atrum Reinke and Berthe or V. dahliae Kleb.)

Geographic occurrence and impact. This is a problem primarily where plants are grown outdoors exposed to soil. But if infected plants or infested soil are shipped from place to place, this is an avenue for long distance spread to wherever Pelargonium is grown. Symptoms/signs. Middle and upper leaves yellow, collapse, dry, and fall. Vascular tissue of affected stems is browned or blackened. Symptoms are readily confused with those of bacterial blight. Sometimes, infected plants are severely stunted before any other symptoms are exhibited (Strider 1985). Biology and epidemiology. The pathogen survives well in soil and in the tissues of many species of infected plants. Management. Purchase culture-indexed cuttings. Use pasteurized potting mixes. Destroy infected plants. Fungus or fungus-like organisms reported to be associated with Pelargonium [see Farr et al. (1989) and Farr, and Rossman Fungal Databases, Systematic Mycology and Microbiology Laboratory, ARS, USDA. Retrieved from http://nt.ars-grin. gov/fungaldatabases/]:

952

Aecidium pelargonii. Alternaria alternata. Alternaria pelargonii. Alternaria tenuis (Alternaria alternata). Alternaria tenuissima. Armillaria fuscipes. Armillaria heimii. Armillaria mellea. Ascochyta sp. Aspergillus fischeri. Aspergillus fischerianus (Aspergillus fischeri). Bipolaris maydis. Botryosphaeria berengeriana (Botryosphaeria dothidea). Botryosporium pulchrum. Botryotinia fuckeliana (Botrytis cinerea). Botrytis pelargonii. Calonectria morganii. Cladosporium fumago (Fumago salicina). Cladosporium sphaerospermum. Coleroa circinans. Colletotrichum gloeosporioides. Coniella australiensis. Coniothyrium trabutii. Cryptovalsa ampelina. Cylindrocarpon olidum (Thelonectria olida). Cylindrocladiella camelliae. Cylindrocladiella parva. Cylindrocladium scoparium. Cyphella pelargonii. Diaporthe medusae (Diaporthe rudis). Diaporthe rudis. Discohainesia oenotherae (Pilidium lythri). Drechslera setariae (Bipolaris setariae). Erysiphe communis (Erysiphe pisi var. pisi) Fibroidium pelargonii. Fusarium oxysporum. Fusarium pelargonii. Fusarium semitectum (Fusarium incarnatum). Gloeosporium pelargonii. Glomerella cingulata (Colletotrichum gloeosporioides). Helicobasidium purpureum. Leptosphaeria elaoudi. Leptosphaeria pelargonii. Macrophomina phaseoli (Macrophomina phaseolina). Macrosporium geraniaceae. Macrosporium pelargonii.

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Oidium sp. Patellaria atrata (Lecanidion atratum). Peroneutypa heteracantha. Pestalotia pelargonii. Pestalotia versicolor (Pestalotiopsis versicolor). Pestalotiopsis sp. Phaeosolenia pelargonii. Phyllosticta geraniicola. Physalospora geranii. Phytophthora cactorum. Phytophthora drechsleri. Phytophthora palmivora var. palmivora (Phytophthora palmivora) Phytophthora x pelgrandis. Pilidium concavum (Pilidium lythri). Pleosphaerulina sp. Pleospora herbarum. Pleospora phaeocomoides. Rhizoctonia solani. Schizophyllum commune. Sclerotiopsis sp. Sclerotium sp. Septoria canberrica. Septoria geranii. Septoria pelargonii. Sphaeropsis sp. Sphaerotheca fugax (Podosphaera fugax). Sphaerulina pelargonii. Sporotrichum epiphyllum. Stemphylium solani. Tuberculina pelargonii. Uredo pelargonii.

3

Bacterial and Phytoplasma Diseases

Where possible, the scientific names used are those accepted by the International Society of Plant Pathology Committee on the Taxonomy of Plant Pathogenic Bacteria (Bull et al. 2010).

3.1

Bacterial Blight (Xanthomonas hortorum pv. pelargonii; Formerly, Xanthomonas campestris pv. pelargonii)

Geographic occurrence and impact. At one time, this was the most serious disease of Pelargonium worldwide. The pathogen is spread in Pelargonium production and throughout the world wherever infected stock plants or cuttings are shipped.

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Losses can approach 100% in some cultivars. Since first reported in the early 1900s, this disease has increased or decreased depending upon the vigilance of growers, diligence in eliminating infected stock plants, and acceptance of growing plants from culture-indexed cuttings. Over the decades of Pelargonium production, outbreaks of this disease have severely damaged the reputations of specialty propagators. Seedling geraniums are susceptible to the pathogen but are free of the disease unless grown in close proximity to vegetatively propagated Pelargonium. Although perennial Geranium is not severely affected; they can be an ongoing source of the pathogen for Pelargonium (Nameth et al. 1999). Symptoms/signs. (Figs. 7, 8, and 9). Small spots [less than 6.4 mm (1/4 in.)] develop on the underside of the leaf and become sunken and well defined. The leaf wilts and dies as the bacteria spread through water-conducting vessels of veins and petioles. V-shaped areas form with the wide part of V on the leaf margin and point of the V on veins. These symptoms are seen on both Pelargonium and Geranium. On Pelargonium, the leaf spotting may occur late in disease development. Prior to leaf spotting, lower leaves may wilt at the margins, while the blade and petiole remain turgid. As wilt progresses, the entire leaf collapses. The vascular tissue of the main stem on the side of the affected portion of the plant is discolored gray to brown. Lower leaves die and fall. Note that although many of these symptoms can be confused with those caused by Ralstonia (bacterial or southern wilt), Ralstonia does not cause leaf spots in Pelargonium. There are simple, rapid test immunological kits commercially available for testing plants for both Xanthomonas hortorum pv. pelargonii and for Ralstonia solanacearum, and there are molecular methods for identifying the presence of Xanthomonas hortorum pv. pelargonii in plant tissues (Sulzinski et al. 1996; Sulzinski et al. 1997). Note that other species are known to cause leaf spots on Pelargonium but that they are relatively minor problems (Rockey et al. 2015).

Fig. 7 Bacterial blight leaf spot (Xanthomonas) on Geranium (The Pennsylvania State University Department of Plant Pathology and Environmental Microbiology # 2017. All Rights Reserved.)

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Fig. 8 Bacterial blight on ivy geranium (Pelargonium peltatum) (The Pennsylvania State University Department of Plant Pathology and Environmental Microbiology # 2017. All Rights Reserved.)

Fig. 9 Bacterial blight – vascular discoloration (Xanthomonas) in Pelargonium (The Pennsylvania State University Department of Plant Pathology and Environmental Microbiology # 2017. All Rights Reserved.)

Biology and epidemiology. The pathogen survives in infected plants and therefore is primarily a problem in Pelargonium propagated vegetatively. Disease develops most rapidly when temperatures are between 20  C and 30  C (68  F and 86  F). When the pathogen is splash dispersed and enters leaves through hydathodes, bacteria enter the vascular tissue and move throughout the plant. The only tissues not infected are the meristem tips where the vascular tissue is not fully differentiated. Viable bacteria survive on moist plant surfaces and within infected tissue and in the vascular tissue. At temperatures between 10  C and 15  C (50  F and 50  F), bacterial activity is greatly suppressed, and infected plants may remain symptomless. This can lead growers to believe that stock plants are not infected. However, as temperatures rise, symptoms become apparent. In the meantime, if cuttings are taken from infected stock plants, the cuttings may root but will eventually exhibit symptoms. In addition to splash dispersal, the bacteria can be spread on workers’ hands and on knives used for taking cuttings (Munnecke 1954).

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Management. Purchase culture-indexed cuttings or grow plants from seed. Immediately discard infected plants after a positive diagnosis is made. Irrigate plants in a manner that keeps water off the foliage. Discard all unwanted geraniums at season’s end if they will not be examined periodically for symptoms. If cuttings are taken with knives, the knives should be washed and thoroughly disinfested after use on one plant and before use on another plant. If branches to be used for cuttings are broken off plants by hand, workers should wash their hands frequently or wear disposable gloves and disinfect them after use on one plant and before use on another plant.

3.2

Bacterial Fasciation (Rhodococcus fascians, Formerly Known as Corynebacterium fascians, Bacterium fascians, and Phytomonas fascians)

Geographic occurrence and impact. Although the bacteria can persist in infested soil, the probable main source of the pathogen in Pelargonium production is through the propagation from infected stock plants. For that reason the disease may appear anywhere Pelargonium is produced vegetatively propagation is grown (Strider 1985). Symptoms/signs. Short, thick, fleshy, leafy galls formed at base of main stems at or below soil level are pale green or green-yellow (Fig. 10). The rest of the plant appears healthy. Biology and epidemiology. Rhodococcus fascians survives associated with live plants or in soil. Many plants in addition to Pelargonium are susceptible to this pathogen. It is believed that Rhodococcus fascians does not require a wound to enter the plant. The pathogen is very difficult to grow in culture from infected plant tissues. Management. Purchase culture-indexed plants. Discard infected stock plants and cuttings and infested media. Fig. 10 Bacterial fasciation on Pelargonium (The Pennsylvania State University Department of Plant Pathology and Environmental Microbiology # 2017. All Rights Reserved.)

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3.3

Diseases of Geranium

957

Bacterial Wilt or Southern Wilt (Ralstonia solanacearum, Formerly Known as Burkholderia solanacearum, and Pseudomonas solanacearum)

Geographic occurrence and impact. Ralstonia solanacearum attack almost 200 plant species in 33 different plant families. The common name for the diseases this organism causes varies with the host that is attacked and geographic location. It is sometimes called southern wilt (in the northern hemisphere) or bacterial wilt. This bacterium is noted for diseases caused outdoors in land areas bounded by 45 N and 45 S latitudes where rainfall averages above 100 cm/year (39 in./year), the average growing season exceeds 6 months, the average winter temperatures are not below 10  C (50  F), the average summer temperatures are not below 21  C (70  F), and the average yearly temperature does not exceed 23  C (72  F) (Lucas 1975). Symptoms/signs. Lower leaves wilt, yellow, and fall. Vascular tissue of affected stems turns brown or black; roots may be discolored and rotted (Figs. 11 and 12). Note that although these symptoms can be confused with those caused by Xanthomonas (bacterial blight), Ralstonia does not cause leaf spots in Pelargonium. Slimy, sticky ooze forms tan-white to brownish beads where the vascular tissue is cut. When an infected stem is cut across and the cut ends held together for a few seconds, a thin thread of ooze can be seen as the cut ends are slowly separated. If one of the cut ends is suspended in a clear container of clean water, bacterial ooze will form a thread in the water. Back lighting of the container helps reveal the ooze. Biology and epidemiology. Although the primary location of survival in the environment is in crop and weed hosts, the bacteria can also survive in soil. They can be readily spread through the movement of contaminated soil and infected vegetatively propagated plants, in contaminated irrigation water, and on the surfaces of tools (cutting knives) and equipment used to work with the plants, and on soiled Fig. 11 Bacterial wilt (Ralstonia) (Robert McGovern # 2017. All Rights Reserved.)

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Fig. 12 Bacterial wilt (Ralstonia) (Robert McGovern # 2017. All Rights Reserved.)

clothing. Populations within this genus and species can be divided into races and biovars based on differing host ranges, biochemical properties, susceptibility to bacteria-infecting viruses (phages), and serological reactions. Race 1 is endemic to North America where it attacks many floricultural and vegetable bedding plant crops including Pelargonium, Catharanthus, Impatiens, Ageratum, Chrysanthemum, Gerbera, Tagetes, Zinnia, Salvia, Capsicum, Lycopersicon, Nicotiana, Petunia, Solanum melongena (eggplant), Tropaeolum (nasturtium), and Verbena. Race 3 is tropical in distribution and does not occur naturally in North America. Race 3 biovar 2 infects potato (Williamson et al. 2002) and other hosts including Pelargonium, tomato, peppers, eggplant, bean, and beet. Weed hosts include black nightshade, climbing nightshade, horsenettle, Jimson weed, purslane, mustards, lamb’s-quarters, and bitter gourd. The bacteria can infect through roots and through any fresh wounds. The bacterium can be difficult to work with in the laboratory because it quickly loses pathogenicity and viability in artificial culture. Race 3 biovar 2 is considered a quarantinable pathogen. Management. Growing and propagating from pathogen-free plant material is the main way to avoid problems with Ralstonia, regardless of the race and biovar involved. It is imperative that propagators use pathogen-free potting soil or other media, establish stock plants that are tested and known to be free of the bacteria, train workers handling the stock plants in methods and procedures that prevent the pathogen from contaminating the potting soil or coming in contact with the stock plants, and then maintaining this system throughout the propagation phase of crop production. Do not bring ground-planted geraniums into the production area or propagate from them. Destroy infected plants. There are no chemicals or biological agents that adequately control Ralstonia. Infected plants MUST be discarded as soon as possible. The purchaser of cuttings or prefinished plants should isolate all new, incoming plants as if the health of the plants were unknown, even if the plants have been certified as healthy. New plants must not be commingled or dispersed among

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other plants in the greenhouse from other sources. This procedure is crucial because keeping plants originating from one source together limits the area that may need to be quarantined, sanitized, or isolated should Ralstonia solanacearum be found. If the pathogen is found in plants being irrigated using a recycling water system (such as ebb and flood), all susceptible plants in the system are at risk to infection. New, incoming plants should not be put into such a system being used for any other plants until their health is verified, and it is known that they do not harbor the pathogen.

3.4

Crown Gall (Rhizobium tumefaciens, Formerly Agrobacterium tumefaciens)

Geographic occurrence and impact. The bacteria can persist in infested soil, but the probable main source of the pathogen in Pelargonium is through the propagation of new plants from infected stock plants. Although this disease may appear anywhere Pelargonium are produced by vegetative propagation, it is usually recognized by the propagator and infected stock plants, and cuttings are discarded. Thus, the disease is very seldom a problem except where a local grower maintains the pathogen in a favorite stock plant (Strider 1985). An understanding of this pathogen’s biology and its disease cycle has significantly contributed to modern agriculture in the area of biotechnology and genetic engineering (Nester et al. 2005). Symptoms/signs. Spherical white- to light tan-colored galls develop near the soil line on the roots and stems of infected plants. Biology and epidemiology. Rhizobium tumefaciens survives in infested soil and in infected plants. The pathogen is spread through the movement of these materials from place to place. Wounding of plants through taking cuttings allows entry of the pathogen. Management. Growing and propagating from pathogen-free stock plants is the main way to avoid crown gall. Propagators must use pathogen-free potting soil or other media, establish stock plants that are known to be free of the bacteria, and destroy infected plants. Ground-planted geraniums should not be brought into the production area nor should cuttings be taken from them.

3.5

Pseudomonas Leaf Spot (Bacterial Leaf Spot; Pseudomonas cichorii)

Geographic occurrence and impact. This disease was first described on Pelargonium X hortorum being grown in Florida, USA, under warm, very humid conditions (Engelhard et al. 1983). In greenhouses or outdoors, the disease can be very damaging particularly if plants are irrigated in a manner that puts water directly on the foliage. Long distance spread of the pathogen occurs when infected plants are shipped from place to place (Daughtrey et al. 1995). The pathogen can infect many different plants in addition to Pelargonium.

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Fig. 13 Pseudomonas leaf spot on Geranium (The Pennsylvania State University Department of Plant Pathology and Environmental Microbiology # 2017. All Rights Reserved.)

Symptoms. Water-soaked spots, 5–10 mm (1/4–1/2 in.) in diameter, form on leaves (Fig. 13). Spots become dark brown to black and irregularly shaped. A yellow halo may surround each spot (Engelhard et al. 1983). Under ideal conditions, spots merge, and the entire leaf may be affected. Infected flower buds and sometime the entire inflorescence are blackened. Biology and epidemiology. Spotting severity increases with temperature between 16  C and 28  C (61  F and 82  F) but is greatly inhibited at temperatures above 28  C. High relative humidity greatly favors infection and spot enlargement (Jones et al. 1984). Sprinkler irrigation and rain splash the pathogen among leaves and flowers and provide the water on the plant tissues required by the bacteria for infection. Management. Purchase pathogen-free cuttings. It is of utmost importance to water plants in a manner that keeps leaf surfaces dry and to maintain low relative humidity among the plants. Discard infected plants and treat the remaining plants with a bactericide.

3.6

“Candidatus Phytoplasma Asteris”

Yellows diseases caused by phytoplasma can occur on Pelargonium. For instance, Candidatus phytoplasma asteris (Group 16SRI) was reported to infect geranium in Pakistan for the first time by Fahmeed et al. (2009).

4

Viral Diseases

The EPPO Panel on “Certification of Ornamentals” developed procedures for the production of healthy carnation, pelargonium, lily, narcissus, chrysanthemum, tulip, crocus, iris, begonia, impatiens, rose, freesia, hyacinth, kalanchoe, and petunia. A

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combination of thermotherapy, meristem tip culture and virus indexing can be used in geranium to eliminate viruses (Horst et al. 1977; Horst and Klopmeyer 1993). In the USA, certification programs are managed by the major commercial propagators. Pelargonium and other major flower crops are tested for the presence of the most prevalent viruses either by in-house plant pathologists or by companies that provide testing services. In the USA, phytosanitary certificates are required for intraand interstate plans movement. Data on pathogens of high consequence, including viruses, are recorded in the USA by the National Plant Diagnostic Network (https:// www.npdn.org/home). Where available, references for viruses are from Adams and Antoniw (2006).

4.1

Arabis Mosaic Virus (ArMV), Tobacco Ringspot Virus (TRSV), Tomato Ringspot Virus (ToRSV), and Artichoke Italian Latent Virus (AILV)

Geographic occurrence and impact. ArMV (Smith and Markham 1944) has been detected in 13 states in the USA where it has been the object of quarantine, but it is not widespread or established outside Europe. ArMV has a wide host range. TRSV (Fromme et al. 1927) geographic distribution is mainly in northern USA and China, but it has been found in Europe and Australia. More than 17 plant families are susceptible to the virus (Price 1940). ToRSV (Price 1936) is originally from the west coast of the USA (Frazier et al. 1961), and it is probably distributed to ornamental plants in many parts of the world. It can infect plants in 35 families. AILV (Majorana and Rana 1970; Vovlas et al. 1971) is found in Southern Italy and Bulgaria. AILV can infect a number of woody and herbaceous plants, including weeds. Symptoms/signs. ArMV is associated with mosaic symptoms in pelargonium. Pelargonium ringspot is a disease associated with both TRSV and ToRSV, but little evidence is found to confirm if the disease is the result of double or single infections. In fact, since the two viruses can infect the same hosts in the same geographic area, they have been often confused. ToRSV has also been associated with ringspots and mosaic symptoms in pelargonium. AILV in Pelargonium zonale produces severe leaf malformations and reduction in size, elongation of petioles, and stunted plant growth (Vovlas 1974). Biology and epidemiology. These three viruses are Nepoviruses in the family Secoviridae. Their genomes consist of two linear ss+RNA segments. ArMV is nematode transmitted (Fritzsche and Schmidt 1963; Harrison and Cadman 1959; Jha and Posnette 1959) and seed transmitted (Lister and Murant 1967, Murant and Lister 1967) and can be transmitted by dodder (Cuscuta sp.). Nematodes lose the virus during molting and do not transmit it to their progeny. TRSV is transmitted by the nematode Xiphinema americanum (McGuire 1964), by pollen and seeds, as well as by many insects (Dunleavy 1957; Komuro and Iwaki 1968; Messieha 1969; Schuster 1963) and mites (Thomas 1969) in a nonspecific

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manner. TRSV does not replicate in its nematode vector, is lost during molting, and is not transmitted to the nematode progeny. TRSV and ToRSV are not serologically related, and serological tests can be used to confirm their identification. ToRSV is transmitted by the nematode Xiphinema americanum (Téliz et al. 1966), but seed transmission has also been reported in some plant species. Small ssRNA satellite viruses have been associated with some strains of ToRSV, but scant information is available on the presence of these satellite viruses in pelargonium. AILV is transmitted by the nematode Longidorus apulus (Rana and Roca 1976), and it is not serologically related to any other Nepovirus. Management. Management of these viruses involves the use of virus-free seed and other propagative material, nematode avoidance through soil and water treatment where necessary, and elimination of weedy hosts.

4.2

Beet Curly Top Virus (BCTV)

Geographic occurrence and impact. The virus originated from the Eastern Mediterranean basin and is common in the Western part of the USA from Mexico to Canada (Bennett and Tanrisever 1957). Today it is present also in Africa, Asia, and Central and South America. BCTV has a broad host range and can infect plant species in 44 families (Bennett 1971). Symptoms/signs. The disease caused by BCTV on pelargonium, called leaf cupping, is very severe, with pronounced tissue yellowing, leaf curling, and distortion. Biology and epidemiology. BCTV is a member of the family Geminiviridae with an ss+DNA genome contained in geminate particles. Two species of leafhoppers from the arid and semi-arid regions, Circulifer tenellus described by Ball (1909) and C. opacipennis (Kheyri and Alimoradi 1969; Stahl and Carsner 1923), are vectors of the virus, and the virus is phloem limited and does not multiply in its insect vectors. The virus is also not transmitted through the vector eggs and requires a latent period before transmission to plants. The virus is present in the seed but cannot invade the embryo. Management. Control of this virus consists of controlling the insect vector, excluding infected plants, and using clean propagative material. Usually plants infected with this virus do not survive and thus are not found commercially.

4.3

Impatiens Necrotic Spot Virus (INSV) and Tomato Spotted Wilt Virus (TSWV)

Geographic occurrence and impact. INSV (Law and Moyer 1990) and TSWV (Samuel et al. 1930) together with their thrips vectors are prevalent worldwide in

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Fig. 14 Symptoms of INSV (Jay Pscheidt # 2017. All Rights Reserved.)

many plant families and in hundreds plant species, making their control particularly challenging. Symptoms/signs. Symptoms associated with INSV and TSWV are mosaic, mottling, stem and leaf necrosis, ringspots on leaves, leaf deformation, and stunted growth (Fig. 14). Symptomatology is driven by plant phenology and environmental conditions. Biology and epidemiology. INSV and TSWV are in the family Bunyaviridae. These ssRNA viruses have tripartite genomes, with negative or ambisense orientation. INSV and TSWV are transmitted by thrips in a propagative (persistent manner) (Ullman et al. 1993; Wijkamp et al. 1993); thus, they can replicate in their plants as well as in their insect hosts. Management. One of the best ways to manage these viruses is to exclude thrips from greenhouses and to reduce their numbers, since thrips need a relatively long acquisition and transmission time, and since the viruses encounter a latent period in the vector prior to becoming transmissible. Unfortunately, thrips are now resistant to many insecticides (Brødsgaard 1994; Zhao et al. 1995). Weed management is also necessary (Bond et al. 1983; Cho et al. 1986; Kobatake et al. 1984). (For additional information, refer to ▶ Chap. 4, “Insect Management for Disease Control in Florists’ Crops”).

4.4

Pelargonium Zonate Spot Virus (PZSV), Cucumber Mosaic Virus (CMV)

Geographic occurrence and impact. PZSV is geographically restricted to Southern Italy. The virus can infect tomato and artichoke and is common in weeds. CMV is found worldwide, has an extremely broad host range, and can infect species in more than 100 plant families (Zitter and Murphy 2009).

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Symptoms/signs. Symptoms induced by PZSV are leaf malformations, puckering, and zonate yellow bands (Martelli and Cirulli 1969; Quacquarelli and Gallitelli 1979). CMV in geranium causes flower breaking disease. Biology and epidemiology. PZSV and CMV (described by Doolittle 1916; Jagger 1916, and more recently by Jacquemond 2012; Palukaitis and Garcia-Arenal 2003) have tripartite ss+RNA genomes. Both viruses are in the family Bromoviridae. PZSV belong to the Anulavirus genus (Gallitelli 1982) and CMV to the Cucumovirus genus (Wildy 1971). PZSV is seed and pollen transmitted but apparently not vector-transmitted. Thrips can serve as carriers of the infected pollen from host to host. CMV is aphid transmitted (Kennedy et al. 1962) in a non-propagative (nonpersistent) manner (Hoggan 1933; Simons 1955; Watson and Roberts 1939), and it can be seed transmitted in some plant species (Neergaard 1977). Management. PZSV can be controlled using clean propagative material. Control for CMV is particularly difficult, since the virus is common on many plant families, and it can be transmitted by multiple aphid species. Because aphids transmit CMV readily during probing, aphid control is particularly important for reducing the spread of the virus. In open field, the use of mineral oil to discourage aphid feeding has been used especially in Europe, but thus far its use in a greenhouse setting has been limited.

4.5

Tobacco Mosaic Virus (TMV) and Tobacco Rattle Virus (TRV)

Geographic occurrence and impact. These viruses can infect a variety of plants in 30 families for TMV (Shew and Lucas 1991) and more than 50 (Horváth 1978; Noordam 1956; Schmelzer 1957; Uschdraweit and Valentin 1956) for TRV in Europe, Japan, New Zealand, and North America. Symptoms/signs. Symptoms attributed to TMV (Siegel and Wildman 1954) are often caused by other tobamoviruses, and many tobamovirus species were once classified as strains of TMV. Symptoms on pelargonium are reported to be mosaic and small light colored or brown lesions on leaves. Biology and epidemiology. TMV and TRV belong to the Virgaviridae family. These viruses have an ss+RNA genome. TMV is occasionally transmitted by chewing insects, but most commonly, it is mechanically spread (Harris and Bradley 1973; Lojek and Orlob 1969); in fact TMV virions are extremely stable. TMV can persist in the soil, probably on plant debris, and can infect roots. It can penetrate wounded embryos from the infected seed coat (Broadbent 1965). TMV is a special concern in greenhouses, where it can be very hard to eradicate (Broadbent and Fletcher 1963). TRV is transmitted by nematodes, Paratrichodorus spp. and Trichodorus spp., (Taylor and Brown 1997), and pelargonium can become infected with this virus if grown in soil infested with viruliferous nematodes. TRV is also transmitted mechanically and by seed in certain hosts.

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Management. Sanitation is the best TMV and TRV control method. Sanitation practices include the use of clean tools and pots, disinfected soil, virus-free seeds, and prompt removal of infected plants. Control of nematodes is essential to limit TRV spread; thus, the use of clean irrigation water and sanitized soil is recommended.

4.6

Tomato Bushy Stunt Virus (TBSV), Tobacco Necrosis Virus (TNV), Moroccan Pepper Virus (MPV), and Pelargonium Leaf Curl Virus (PLCV)

Geographic occurrence and impact. TBSV has a restricted natural host range and causes damage primarily on agricultural crops, while it can experimentally infect more than 120 plant species. It is geographically distributed in Europe, the Americas, and North Africa. TNV has a natural broad host range (Price 1940) that include tobacco, cucumber, melon, European strawberry, citrus, apple, and pear. It can experimentally infect more than 37 plant families, and it is distributed worldwide. PLCV (Hollings and Stone 1965) is common in the USA and Mediterranean region. It was detected in India in 2013 (Kumar et al. 2013). Susceptible host species are Pelargonium zonale, common bean, Datura, and Chenopodium. Some of these species are considered experimental hosts. MPV is present in Morocco, Iran, USA, Europe, and Central Asia (Wintermantel and Hladky 2013). Its host range includes tomato, lettuce, pepper, pelargonium, jimson weed, and lisianthus. Symptoms/signs. These viruses induce Pelargonium leaf curl or crinkle, and Pelargonium necrotic spot, with stunted growth, leaf deformation and splitting, extensive spots first hyaline and then necrotic, and yellowing and necrosis of leaves. Symptoms are strongly influenced by temperature. In pelargonium, TNV can cause symptomless infection that turns symptomatic under different light and temperature conditions. PLCV is the causal agent of leaf curl disease. The disease symptoms are leaf curl and yellow stellate spots that are more pronounced in winter and at lower temperatures and diminish in summer at higher temperatures. Biology and epidemiology. These are monopartite ss+RNA viruses and are members of the Tombusvirus genus in the family Tombusviridae. TBSV (Ainsworth 1936, Bawden and Pirie 1938 Smith 1935) is soilborne and is transmitted by infected seed and soil. Defective interfering RNAs (DI-RNAs) and satellite RNAs have been associated with TBSV in its naturally infected plants (Galetzka et al. 2000, Gallitelli and Hull 1985), but were not investigated in pelargonium. TNV (Babos and Kassanis 1963; Bawden 1941; Smith and Bald 1935) is a soilborne virus and is transmitted by zoospores of the chytrid fungus Olpidium brassicae (Kassanis and MacFarlane 1964, Teakle 1962, Teakle and Gold 1963). TNV can support the multiplication of satellite viruses in some plants and serologically can be divided into groups, but group serology does not coincide with symptomatology. PLCV is

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serologically related to TBSV. MPV (Makkouk et al. 1981) is distantly serologically related to TBSV. PLCV and MPV were described to be associated with soil infested by Olpidium spp. in Iran in 2011 (Rasoulpour and Izadpanah 2011). PLPV is an unassigned species in the Tombusviridae. Management. Virus control requires the use of clean propagative material and soil, avoiding the spread of the vectoring zoospores by removing infested soil and water, and good sanitation practices. Research on virus-resistant varieties is needed.

4.7

Pelargonium Flower Break Virus (PFBV)

Geographic occurrence and impact. Virus described by Stone and Hollings (1973). It infects six plant families and has limited host range. The virus was reported to be the most common viral pathogen of Pelargonium sp. in Western Europe (Krczal et al. 1995). Symptoms/signs. It can infect Pelargonium domesticatum and can cause flowerbreaking in several cultivars, but it remains usually symptomless on P. peltatum and P. zonale. Other symptoms can include flower streaking, stunting, and chlorotic spotting (Fig. 15). Seedlings of pelargonium-type “Nittany Lion” appear to be immune to PFBV (Kemp 1969). Biology and epidemiology. PFBV is an ss+RNA virus belonging to the genus Carmovirus in the family Tombusviridae. PFBV is transmitted mechanically and by vegetative propagation, but does not seem to be transmitted by seed. Transmission of the virus in recirculating irrigation systems and by thrips via pollen has been demonstrated (Krczal et al. 1995). The virus is not serologically related to other viruses. Fig. 15 Symptoms of PFBV (Robert Wick # 2017. All Rights Reserved.)

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Management. Use of clean propagative material and discarding of infected plants is recommended. Hollings and Stone (1974) reported that Pelargonium can be freed from PFBV by thermotherapy, but plant survival rate was low. Other viruses associated with Pelargonium: Tombusviridae family: Pelargonium chlorotic ring pattern virus. Pelargonium leaf curl virus. Pelargonium line pattern virus. Pelargonium necrotic spot virus. Pelargonium vein banding virus is a virus related to viruses in the Badnavirus genus, not yet approved as a species. Pelargonium vein clearing is in the genus Nucleorhabdovirus. It has a monopartite ss-RNA genome and replicates in the plant cell nucleus. Other viruses: Cherry leaf roll virus. Lilac chlorotic leaf spot virus. Tomato black ring virus.

5

Nematode Diseases

Nematodes, including dagger nematode (Xiphinema sp.), foliar nematode (Aphelenchoides sp.), and root-knot nematode (Meloidogyne sp.), have been recorded to occur on Pelargonium but do not appear to be a widespread problem (Strider 1985).

6

Abiotic Diseases

A myriad of factors including air pollutants, misuse of plant growth regulators, and individual cultivar responses to environmental conditions can result in symptoms on Pelargonium (Freeman 1993).

6.1

Edema

Geographic occurrence and impact. This problem can occur wherever Pelargonium is grown. It is most damaging to ivy geranium (Pelargonium peltatum). (Balge et al. 1969; Digat and Albouy 1976) Symptoms/signs. Small water-soaked pimples or blisters form on the underside of lower leaves (Fig. 16). The blisters become corky and brown as they enlarge. Severely affected leaves fall.

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Fig. 16 Edema on ivy geranium (Pelargonium peltatum) (The Pennsylvania State University Department of Plant Pathology and Environmental Microbiology # 2017. All Rights Reserved.)

Biology and epidemiology. Edema development is favored when the weather is cool, cloudy, and humid while the soil is wet, particularly if mites are feeding on the plants. Although over the decades scientists have referred to this as a physiological disorder and have not linked it to a biotic pathogen, it is interesting to note that symptoms can be induced in otherwise healthy plants by injecting them with sap taken from a plant with edema (Digat and Albouy 1976). Management. Spacing plants to provide good air circulation, using a well-drained potting mix, not overwatering during cool, cloudy weather, and mite suppression, are reported to help in preventing the development of edema.

6.2

Heat Stress

Geographic occurrence and impact. This problem can occur wherever Pelargonium is grown. Sensitivity to high temperatures varies greatly among cultivars (Strider 1985). Symptoms/signs. Upper leaves and stems completely lose chlorophyll. Sometimes, only the center of the leaf is bleached. Usually only a few leaves are affected, but in severe cases, most of the leaves and stems may appear bleached. Biology and Epidemiology. Pelargonium exposed to temperatures above about 28  C/82  F for more than 12 h develop symptoms. While some cultivars exhibit severe symptoms, others in the same greenhouse may be completely free of symptoms. Management. Monitor temperatures in the area where cultivars known to be heat sensitive are grown. If it appears that temperatures could persist above 28  C for more than 12 h, mist the sensitive cultivars so that evaporative cooling occurs.

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However, many of the pathogens detailed elsewhere in this section are favored when moisture is on the foliage. Apply only enough moisture to slightly dampen the leaf surface and then evaporate completely in minutes. Do not mist to the extent that water begins to drip off the foliage.

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Smith KM (1935) A new virus disease of the tomato. Ann Appl Biol 22:731–741 Smith KM, Bald JG (1935) A description of a necrotic virus disease affecting tobacco and other plants. Parasitology 27:231–245 Smith KM, Markham R (1944) Two new viruses affecting tobacco and other plants. Phytopathology 34:324–329 Stahl CF, Carsner E (1923) A discussion of Eutettix tenellus Baker as a carrier of Curly top of sugar beets. J Econ Ent 16:476–479 Stone OM, Hollings M (1973) Some properties of pelargonium flower-break virus. Ann Appl Biol 75:15–23 Strider DL (1985) Geranium. In: Strider DL (ed) Diseases of floral crops, vol 2. Praeger Scientific, New York, pp 111–187 Sulzinski MA, Moorman GW, Schlagnhaufer B, Romaine CP (1996) Characteristics of a PCR-based assay for in planta detection of Xanthomonas campestris pv pelargonii. J Phytopathol 144:393–398 Sulzinski MA, Moorman GW, Schlagnhaufer B, Romaine CP (1997) A simple DNA extraction method for PCR-based detection of Xanthomonas campestris pv. pelargonii in geraniums. J Phytopathol 145:213–215 Taylor CE, Brown DJF (1997) Nematode vectors of plant viruses. CAB International, Wallingford Teakle DS (1962) Transmission of tobacco necrosis virus by a fungus, Olpidium brassicae. Virology 18:224–231 Teakle DS, Gold AH (1963) Further studies of Olpidium as a vector of tobacco necrosis virus. Virology 19:310–315 Téliz D, Grogan RG, Lownsbery BF (1966) Transmission of tomato ringspot, peach yellow bud mosaic, and grape yellow vein diseases by Xiphinema americanum. Phytopathology 56:658–663 Thomas C (1969) Transmission of tobacco ringspot virus by Tetranychus sp. Phytopathology 59:633–636 Ullman DE, German TL, Sherwood JL, Westcot DM, Cantone FA (1993) Tospovirus replication in insect vector cells: immunocytochemical evidence that the nonstructural protein encoded by the S RNA of tomato spotted wilt tospovirus is present in thrips vector cells. Phytopathology 83:456–463 Uschdraweit HA, Valentin H (1956) Das Tabakmauchevirus an Zierpflanzen. Nachrichtenbl Dtsch Pflanzenschutzdienst 8:132–133 (In German) Vovlas C (1974) Le malformazioni fogliari, una nuova virosi del geranio. Phytopathol Mediterr 13:139–142 (In Italian) Vovlas C, Martelli GP, Quacquarelli A (1971) Le virosi delle piante ortensi in Puglia. Il complesso delle maculature anulari della cicoria. Phytopathol Mediterr 10:244–254 (In Italian) Watson MA, Roberts FM (1939) A comparative study of the transmission of Hyoscyamus virus 3, potato virus Y and cucumber virus 1 by the vector Myzus persicae (Sulz.), M. circumflexus (Buckton) and Macrosiphum gei (Koch). Proc R Soc Ser B 127:543–576 Wijkamp I, van Lent J, Kormelink R, Goldbach R, Peters D (1993) Multiplication of tomato spotted wilt virus in its insect vector, Frankliniela occidentalis. J Gen Virology 74:341–349 Wildy P (1971) Classification and Nomenclature of Viruses. First Report of the International Committee on Nomenclature of Viruses. Monographs in Virology no. 5. Basel: Karger. Basel, pp. 33–34 Williamson L, Nakaho K, Hudelson B, Allen C (2002) Ralstonia solanacearum race 3, biovar 2 strains isolated from geranium are pathogenic on potato. Plant Dis 86(9):987–991 Wintermantel WM, Hladky LL (2013) Complete genome sequence and biological characterization of Moroccan pepper virus (MPV) and reclassification of Lettuce necrotic stunt virus as MPV. Phytopathology 103(5):501–8 Zhao G, Liu W, Brown JM, Knowles CO (1995) Insecticide resistance in field and laboratory strains of western flower thrips (Thysanoptera: Thripidae). J Econ Entomol 88:1164–1170 Zitter TA, Murphy JF (2009) Cucumber mosaic. The Plant Health Instructor. doi:10.1094/PHI-I2009-0518-01

Diseases of Holiday Cacti: Schlumbergera and Hatiora

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Robert L. Wick

Abstract

The most important diseases of Schlumbergera truncata, the Thanksgiving cactus, are Fusarium basal stem rot, Phytophthora root and stem rot, Bipolaris blight, and bacterial soft rot. Impatiens necrotic spot virus (INSV) and Tomato spotted wilt virus (TSWV) are a problem because infected plants cannot be sold. Hatiora gaertneri (Regel) Barthlott, the Easter cactus is relatively free of important diseases except for Fusarium stem rot which is very destructive to Hatiora. Keywords

Fusarium oxysporum • Bipolaris cactivora • Phytophthora nicotianae • Pectobacterium carotovora • Pythium • INSV • TSWV

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Basal Stem and Root Rot [(Fusarium oxysporum Schlecht); (Fusarium oxysporum f. sp. opuntiarum)] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Bipolaris Blight Bipolaris cactivora (Petr.) Alcorn.; Previously Drechslera cactivora (Petr.) M. B. Ellis and Helminthosporium cactivorum Petr. . . . . . . . . . . . . . . 2.3 Pythium Root and Stem Rot [Pythium aphanidermatum (Edson) Fitzp. Pythium irregulare Buisman, and Pythium cryptoirregulare Garzon, Yanez, and Moorman] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Phytophthora Root and Stem Rot Phytophthora nicotianae Breda de Haan; Formerly Phytophthora parasitica Dastur. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Other Minor Diseases Caused by Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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3 Bacterial Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Bacterial Soft Rot [Pectobacterium carotovorum subsp. carotovorum (Jones) Hauben et al. emend. Gardan et al.)]. Formerly Erwinia carotovora subsp. carotovora . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Virus Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Tomato Spotted Wilt Virus (TSWV) and Impatiens Necrotic Spot Virus (INSV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Introduction

Schlumbergera truncata (Haworth) Moran, Thanksgiving cactus (also called Christmas cactus), is the most common Schlumbergera species under cultivation (Boyle 2007). The interspecific hybrid S. x buckleyi (Buckley) Tjaden (= S. russelliana x S. truncata) blooms a month later and is commonly called the Christmas cactus (Boyle 2007; Daughtrey et al. 1995). Schlumbergera truncata was formerly Zygocactus truncatus (Haw.) K. Schum. Members of the Cactaceae are native to Brazil; they are widely cultivated as a flowering potted plant along with Hatiora gaertneri (Regel) Barthlott, the Easter cactus. Hatiora gaertneri was previously known as Rhipsalidopsis gaertneri (Regel) Moran.

2

Fungal and Fungus-Like Diseases

2.1

Basal Stem and Root Rot [(Fusarium oxysporum Schlecht); (Fusarium oxysporum f. sp. opuntiarum)]

Geographic occurrence and impact. Fusarium basal stem and root rot of S. truncata was first reported in a proceedings in 1975 (Miller 1975) and formally published in 1980 (Moorman and Klemmer 1980). It has been reported in the USA, Argentina (Petrone et al. 2007), Italy (Lops et al. 2013), the Netherlands (Baayen et al. 2000), and Germany (Baayen et al. 2000; Gerlach 1972). Inoculation trials with F. oxysporum isolated from S. truncata failed to cause disease on H. gaertneri (Chase 1982). However subsequent research showed that an F. oxysporum isolated from S. truncata caused stem and root rot on several cultivars of H. gaertneri similar in extent to several cultivars of S. truncata (Mitchell 1987). Nevertheless there is a paucity of information regarding the occurrence of F. oxysporum root and stem rot on H. gaertneri. Fusarium oxysporum f. sp. opuntiarum was described from Schlumbergera in Germany in 1972. Subsequent reports of F. oxysporum on cactus did not include the f. sp. opuntiarum epithet; however, it is not known if F. oxysporum cited in the USA and Argentina is different or the same as F. oxysporum f. sp. opuntiarum reported in Europe. Fusarium basal stem and root rot is one of the most destructive diseases of Schlumbergera.

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Symptoms/signs. The disease can manifest itself as a root rot, basal stem rot, and cladophyll lesion. Basal rot can occur on wounded or unwounded stems and is at first water soaked in appearance with reddish-brown margins. As the lesion ages, the tissue may dry to a papery tan (Moorman and Klemmer 1980). Lesions on wounded inoculated cladophylls of S. truncata can become up to 50 mm in diameter in 10 days (Chase and Yuen 1993). Lesions on inoculated H. gaertneri cladophylls ranged from about 5 to 20 mm in diameter. Biology and epidemiology. F. oxysporum is the most widely dispersed of Fusarium spp. and occurs throughout the world in many types of soils, mostly as a soil saprophyte (Domsch et al. 1980; Leslie and Summerell 2006). A true soil-borne fungus survives in the soil by producing resting structures, chlamydospores, and active growth. Since Hatiora and Schlumbergera are vegetatively propagated, the widespread occurrence of F. oxysporum on cactus is probably due to movement of infected plant material and infested soil. Fusarium oxysporum is best known for the destructive nature of the vascular wilt diseases it causes on a wide variety of crops (Agrios 2005); more than 100 forma speciales and races have been described, most of which have a very narrow host range, often restricted to a single plant species or cultivar. While the descriptions of F. oxysporum disease of cactus often include wilt, wilt is apparently due to root rot and basal stem canker as opposed to vascular wilt. No one has demonstrated that F. oxysporum is a vascular pathogen of Schlumbergera. Management • Cultural Practices – New plants coming into the greenhouse should be carefully inspected for root rot, basal canker, and lesions on cladophylls. Diseased plants and plant debris should be removed from the presence of healthy plants. When growing plants in soil-less media, take care not to introduce field soil by hands, tools, and hose ends dropped on the floor. If the growing medium contains field soil, it must be pasteurized. Air drying of excised cladophylls up to 72 h before propagation did not prevent Fusarium from infecting (Mitchell 1987). With other Fusarium diseases, the application of nitrate as opposed to ammonium forms of nitrogen can suppress Fusarium. In addition, acid soils 80 reported) of aphid species in a non-circulative (nonpersistent) manner. This means that the virus can be acquired and inoculated by the aphid during probing. CMV can be seed

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Fig. 11 Mosaic pattern on leaves associated with CMV infection (Wageningen University and Research # 2017. All Rights Reserved.)

transmitted in some hosts. Mechanical transmission through contaminated knives, machines, or water has been reported (Conijn 2014). Management. Starting with virus-free planting material is of primary importance. Control measures consist of reduction of the transmission by aphids through monitoring and control of aphids and plant treatments with mineral oil and pyrethroids. Aphid transmission can be decreased with reflective mulches or by the use of fine mesh nets or screens. Remove virus sources by rogueing crops and thorough weed control.

4.4

Lily mottle virus (LMoV, Genus: Potyvirus)

Geographic occurrence and impact. LMoV is the most common virus in lily and has spread worldwide. The host range is mostly restricted to the Liliaceae family (Lisa et al. 2002). Symptoms/signs. Symptoms can include vein clearing, leaf mottle, leaf mosaic, chlorotic and yellow streaking, leaf curling, and narrowing. Some cultivars can show color breaking of the flower (Fig. 12).

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Fig. 12 Flower break associated with LMoV infections (Wageningen University and Research # 2017. All Rights Reserved.)

Biology and epidemiology. LMoV is transmitted by aphids in a nonpersistent way (Asjes 2000; Conijn 2014). Management. Use virus-free or certified plant material in combination with sprays of mineral oil plus pyrethroid insecticides, to prevent virus spread. Resistance to LMoV occurs in some cultivars of the Asiatic hybrid group.

4.5

Lily symptomless virus (LSV, Genus: Carlavirus)

Geographic occurrence and impact. LSV is the most harmful virus in lily and has spread worldwide. It has a host range restricted to the Liliaceae family and is transmitted by aphids nonpersistently (Fischer 2013). Symptoms/signs. LSV can occur symptomlessly in lily and Alstroemeria. The name Lily symptomless virus is perhaps unfortunate because other lily viruses can also sometimes infect lilies symptomlessly, and LSV itself causes symptoms in lilies in certain genotypes under certain environmental conditions. Symptoms can include mild, pale veinclearing and mottle on leaves in lilies grown for cut flowers under glasshouse conditions (Fig. 13). The leaves of LSV-infected plants generally turn yellow fairly quickly (Boontjes 1983). Biology and epidemiology. LSV is transmitted by the aphids Macrosiphum euphorbiae, Myzus persicae, Aphis gossypii and others species in a nonpersistent way (Asjes 2000; Conijn 2014). In the Netherlands, aphid flights with concomitant virus spread gradually increase in April/May, with a bigger increase in June and July (Asjes 1997, 2000).

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Fig. 13 Mild veinclearing and mottle on leaf (right) associated with LSV infection (Wageningen University and Research # 2017. All Rights Reserved.)

Management. Use of virus-free planting material in combination with sprays of mineral oil plus pyrethroid insecticides to prevent virus spread.

4.6

Lily virus X (LVX, Genus: Potexvirus)

Geographic occurrence and impact. LVX has a very narrow host range. It infects lily and some weeds (Chenopodium quinoa and Tetragonia expansa under experimental conditions). Symptoms. LVX infects lily without causing symptoms (latent infection). Biology and epidemiology. LVX is transmitted via contact (mechanically). There is no known vector for LVX, but vector transmission is suspected as the use of insecticides (pyrethroids) has limited transmission (Asjes 2000).

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Management. Start with virus-free plant material. Hygienic measures and awareness of virus sources (infected plant material, water in dipping baths) are extremely important for contact-transmitted viruses, like potexviruses.

4.7

Plantago asiatica mosaic virus (PlAMV, Genus: Potexvirus)

Geographic occurrence and impact. PlAMV has a very broad host range, in both monocots and dicots. It was discovered in lily in recent years (Anonymous 2010). Hosts of the virus are lily, various weeds, and even shrubs like Nandina domestica. Symptoms/signs. Rust-brown necrotic spots occur mostly on the backside of the leaf (Fig. 14). On the upper side grayish spots are sometimes observed. Later, brown necrotic spots occur along the veins on the upper side of the leaf (mostly at the end of the vegetative phase). Leaves of infected plants are often brittle. PlAMV can also be symptomless in lily. Mixed infections of PlAMV with LSV or LMoV can cause much more severe symptoms including dwarfism.

Fig. 14 Rust-brown necrotic spots leaves from PlAMVinfected plant (Wageningen University and Research # 2017. All Rights Reserved.)

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Biology and epidemiology. Transmission is mostly via contact with infested water in dipping baths. Also, transmission via soil, without a vector, has been reported (De Kock et al. 2013). Management. Start with virus-free plant material. Hygienic measures and awareness of virus sources (infected plant material, water in washing and dipping baths) are extremely important for contact-transmitted viruses, like potexviruses. Avoid plots where PlAMV-infected plants have been grown; plots can remain contaminated with this virus through overwintering weeds and infected volunteers.

4.8

Strawberry latent ringspot virus (SLRSV, Not Assigned to a Genus)

Geographic occurrence and impact. SLRSV has a broad host range in both monocots and dicots. Various ornamental, vegetable, and fruit crops can be infected by SLRSV. Known flower bulb hosts are lily and daffodil. Symptoms/signs. SLRSV in lily is symptomless; however, there is one report of asymmetric opening of flowers in the Oriental hybrid cultivar ‘Stargazer’ (Cohen et al. 1995). Biology and epidemiology. SLRSV is transmitted by the free-living nematodes Xiphinema diversicaudatum and X. coxi. Also seed transmission in various crops was reported, including lily (Verbeek and Stijger 2016). Management. Start with virus-free plant material on Xiphinema-free plots, and also control weeds. Nematodes can be controlled chemically with nematicides and also by biological measures such as planting certain Tagetes spp. and by biofumigation. Be aware of possible seed transmission in seed-propagated lilies. (Additional information on nematode management may be found in Sect. 5 below and in ▶ Chap. 8, “Soil/Media Disinfestation for Management of Florists’ Crops Diseases”).

4.9

Tobacco rattle virus (TRV, Genus: Tobravirus)

Geographic occurrence and impact. TRV has a very broad host range in more than 50 monocotyledonous and dicotyledonous plant families. Various food crops, ornamentals, and weeds are infected by TRV. Among the flower bulbs, lily, tulip, hyacinth, crocus, and daffodil are hosts of this virus. Symptoms/signs. Chlorotic spots and veinal chlorosis, small necrotic spots and twisted growth, and leaf curling.

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Biology and epidemiology. TRV is transmitted by nematodes from the genera Trichodorus and Paratrichodorus (family Trichodoridae). Both adult and juvenile nematodes can transmit the virus, but the virus is probably lost when the nematodes molt. Nonfeeding nematodes can retain the virus for several months. Seed transmission was reported for some weeds (shepherd’s purse, poppy). Management. Use virus-free plant material and nematode-free plots. Control weeds. Nematodes can be controlled chemically with nematicides and also by biological measures such as planting certain Tagetes spp. and by biofumigation. (Additional information on nematode management may be found in Sect. 5 below and in ▶ Chap. 8, “Soil/Media Disinfestation for Management of Florists’ Crops Diseases”.

4.10

Tulip breaking virus (TBV, Genus: Potyvirus)

Geographic occurrence and impact. TBV is the most common virus in tulip but is also found in lily. The virus has a host range restricted to the Tulipa and Lilium. Symptoms/signs. TBV causes mild to moderate mottling in the leaves and has been associated with a brown ring formation in the Asiatic hybrid cultivar “enchantment” (Derks 1976). Biology and epidemiology. TBV is transmitted by aphids in a nonpersistent way (Asjes 2000; Conijn 2014). Management. Use virus-free or certified plant material in combination with sprays of mineral oil plus pyrethroid insecticides to prevent virus spread. Resistance to TBV occurs in Tulipa fosteriana (Marasek-Ciolakowska et al. 2012).

4.11

Tulip virus X (TVX, Genus: Potexvirus)

Geographic occurrence and impact. Hosts of the virus are tulip, lily, and various weeds. Symptoms/signs. Symptoms: sunken, light green or yellow spots in between the veins, and lighter or darker stripes at the outside of the flowers (Fig. 15). Biology and epidemiology. Transmission mostly via contact with infested water in dipping baths. Mite transmission is suspected. Transmission via soil, without a vector, has been reported (De Kock et al. 2013). Management. Start with virus-free plant material. Hygienic measures and awareness of virus sources (infected plant material, water in washing and dipping baths)

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Fig. 15 Sunken, light green to yellow spots in between veins on a TVX-infected plant (Wageningen University and Research # 2017. All Rights Reserved.)

are extremely important for contact-transmitted viruses like potexviruses. Avoid plots where TVX-infected plants have been grown; plots can remain contaminated with this virus through overwintering weeds, etc.

5

Nematode Diseases

5.1

Root Lesion Nematode (Pratylenchus penetrans (Cobb, 1917) Filipjev and Schuurmans Stekhoven, 1941)

Geographic occurrence and impact. P. penetrans is mainly found in temperate areas including the USA, Europe, Australia, Canada, Egypt, India, Japan, New Zealand, Peru, the Philippines, Zimbabwe (Rhodesia), Russia, South Africa, and Tunisia (Goodey et al. 1965; Siddiqui et al. 1973). It has a wide host range and more than 350 hosts have been recorded including apple, cherry, other fruit trees, conifers, roses, tomato, potato, corn, and sugar beets. It is considered the most economically important plant-parasitic nematode in the northeastern USA and field-grown Easter lilies in California and Oregon and in the Netherlands when lilies are grown on light and sandy soils (Conijn 2014; Westerdahl et al. 2003). Symptoms/signs. Disease symptoms include a general devitalization of the plant: retarded top growth, chlorotic foliage, and restricted root growth (Overman 1961) (Fig. 16). Shoots may not be able to emerge from bulbs. In less serious cases, the symptoms are not evident until late in the growing season. Because these symptoms could be indicative of problems other than nematodes, soil and root samples should be sent to a nematode diagnostic laboratory for evaluation. Biology and epidemiology. P. penetrans is a migratory endoparasite, with all life cycle stages found either within roots or in the soil. Reproduction is sexual. Females lay eggs singly in roots or in soil. Second-stage juveniles hatch from eggs, feed, and undergo three molts to the adult stage. The complete life cycle takes 30–86 d,

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Fig. 16 Stunting and yellowing of plants due to root lesion nematode (Becky B. Westerdahl # 2017. All Rights Reserved.)

depending on temperature, and is shortest at 30  C/86  F. Adults, J2, J3, and J4, can all invade roots. If conditions in the root become unfavorable, any juvenile stage or the adult nematodes may leave the root and invade other nearby roots. Invasion usually takes place in the region of root elongation. Root lesion nematodes feed upon cells in the root cortex. Cells are killed and in many instances, small roots are killed. The migratory parasitism of this nematode opens up roots to secondary invasion by other soil microorganisms such as fungi and bacteria (Corbett 1973). Management. The number of nematodes in the soil, soil characteristics, and the nematode populations in the roots of planted bulbs affects the degree of damage caused by the root lesion nematode. Conijn (2014) reports that researchers at Wageningen University Research (WUR) are developing a computer-based, decision-support system named NemaDecide to predict the probability of damage by nematodes (Elberse et al. 2012). Hot water treatment of recently harvested bulbs and a subsequent storage period at 2  C/28  F for 2 mo before planting is an effective method to control Pratylenchus penetrans in lily planting stock (Conijn 1966, 2014). This approach prevents nematode damage to bulb roots and also reduces their spread to uninfested fields. Marigolds (Tagetes spp.), which are known to produce the nematicidal compound thiophene α-therthienyle (Oostenbrink et al. 1957; Conijn 1994; Pudusaini et al. 2006; Gommers and Bakker 1988), are sometimes used by Dutch lily growers who sow them directly after harvesting tulips or other spring-flowering bulb crops and have been shown to reduce populations of nematodes in the soil by 90% after 105 d (Pudasaini et al. 2006). Easter lily growers in Oregon and California face a number of challenges in managing root lesion nematodes. Field-grown bulbs are sold to greenhouse operations for forcing to produce potted, flowering plants at Easter. The quality of field-grown bulbs is based on bulb circumference and appearance, with only bulbs with white scales that have plentiful roots being saleable. Nematode management is complicated because bulbs must be grown for 2–4 y before they are large enough for sale. Land is prepared for fumigation in May, fumigated in July, bulblets are planted from August

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through October with a planting treatment of a nonfumigant nematicide, and bulbs are harvested the following August through October (D’Herde et al. 1960; Roberts et al. 1985) (Fig. 17). Bulbs not large enough for sale are replanted for an additional growing season. Avoidance of the root lesion nematode problem in Easter lilies by crop rotation or planting site selection is problematic. The nematode has a very wide host range and is supported by the pasture grasses that are commonly grown as an alternative to bulb production (Giraud et al. 2011; Westerdahl et al. 1993, 2003).

5.2

Foliar Nematode: (Aphelenchoides spp.)

Geographic occurrence and impact. Foliar nematodes have a wide host range and are found in temperate and tropical regions. A. fragariae has an extensive host range, including fern, lily, begonia, African violet, strawberry, and many aquatic plants (Goodey et al. 1965; Siddiqui et al. 1973). Symptoms/signs. Foliar nematodes produce discolored streaks in lily foliage and will cause the bulbs to decline (Gill et al. 2006). Infested plants are stunted and distorted (Fig. 18). Infested flower buds may not bloom or flowers may be distorted (Siddiqi 1975). Because these symptoms could be indicative of problems other than nematodes, soil and plant samples should be sent to a nematode diagnostic laboratory for evaluation. Biology and epidemiology. Foliar nematodes are migratory endoparasites in leaves, but also feed ectoparasitically on leaf and flower buds. Nematodes enter leaves through stomata or by penetrating tissues. They have been shown to move long distances on infested plant material (Handoo 2012). They reproduce sexually. The life cycle is completed in 10–13 d and females produce about 30 eggs. Unlike Fig. 17 Growth of Easter lilies treated with a nonfumigant nematicide (left) at the time of planting in a field infested with root lesion nematode (Easter Lily Research Foundation # 2017. All Rights Reserved.)

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Fig. 18 Severely stunted plant (center) infested with foliar nematodes (Department of Entomology and Nematology, UC Davis, Slide Collection # 2017. All Rights Reserved.)

most plant parasites that appear sedentary in water, foliar nematodes are rapid swimmers. They survive in soil for about 3 mo, with longer survival in plant tissue. Weeds can harbor the foliar nematodes that infect lily. The nematode overwinters in dead plant tissues in soil and also in buds and growing points. It thrives best in moist, cool situations (Siddiqi 1975). Management. Cultural practices include rogueing plants with symptoms, controlling weeds, using irrigation practices that reduce the incidence and duration of leaf wetness and splash dispersal of the nematode, hot water treatments of planting stock (Jagdale and Grewal 2002, 2004), burning infested material, starting/replacing with healthy stock, and general sanitation. Foliar or soil treatments with systemic chemicals are also effective (Kohl 2011; Siddiqi 1975).

5.3

Stem and Bulb Nematode: (Ditylenchus dipsaci (Kühn, 1857) Filipjev, 1936)

Geographic occurrence and impact. Stem nematode is widely distributed in temperate regions. It is a common crop pest with more than 450 hosts (Goodey et al. 1965; Siddiqui et al. 1973). Host range determination is complicated by there being as many as 25 host races or biotypes, some with a limited host range (Greco et al. 1991). The bulb race infests most bulbs including hybrid lilies, daffodil, narcissus, and tulip (Hooper 1972). Symptoms/signs. Distortion and swelling of leaves and bulbs are common symptoms (Greco et al. 1991). The nematode causes individual bulb scales to rot and eventually can kill the bulb (Gill et al. 2006). Secondary invasion by bacteria and fungi may contribute to bulb rot. Because these symptoms could be indicative of problems other than nematodes, soil and plant samples should be sent to a nematode diagnostic laboratory for evaluation.

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Biology and epidemiology. D. dipsaci is a migratory endoparasite, meaning that all life stages can be found either within a plant or in the soil. At the beginning of the crop season, fourth-stage juveniles enter young tissues through stomata or by direct penetration. The nematode is thought to secrete a pectinase enzyme that breaks down plant tissues. Migration on plant parts aboveground requires free water and may occur after rain or sprinkler irrigation. Wet conditions and mild temperatures (15–20  C/59–68  F) favor infestation. Under dry conditions, the nematode survives in plant tissues as a quiescent fourth-stage juvenile, and this aids spread in plant materials (Greco et al. 1991). Management. Hot water treatment of bulbs has been a common practice for managing this nematode. However, a hot water bath is also an effective means to spread fungi and bacteria so that an additive is needed to control these pests (Qiu et al. 1993). Soil treatments with nematicides will control nematodes present in the soil but not those contaminating planting stock. Rotation for 2–3 y to nonhost crops plus weed control is an effective cultural practice. Using drip rather than overhead irrigation will help to minimize spread of the nematode within a crop (Greco et al. 1991).

5.4

Root-Knot Nematode: (Meloidogyne spp.)

Geographic occurrence and impact. Root-knot nematodes are found worldwide but are more common in temperate, subtropical, and tropical areas (Goodey et al. 1965; Mitkowski and Abawi 2003; Siddiqui et al. 1973; Taylor and Sasser 1978). Several species including M. incognita, M. chitwoodi, and M. fallax have been reported as being pathogenic to lilies (Den Nijs et al. 2004). Root-knot nematodes are a quarantine pest in many countries. The cosmopolitan distribution of root-knot nematode is the result of the movement of infested plants, water, soil, and equipment. Symptoms/signs. Plants exhibit poor growth and roots infected with root-knot nematodes show round swellings, or galls (root knots), typically most noticeable on the small feeder roots. There is poor top growth and the foliage is frequently chlorotic (yellow) because essential elements are not taken in and transported by the impaired root system. Severe infections cause wilting of the foliage and the plants require more frequent irrigations. Because these symptoms could be indicative of problems other than nematodes, soil and root samples should be sent to a nematode diagnostic laboratory for evaluation. Female nematodes are visible under a microscope in cross sections of the root swellings and nematode infestations will usually appear as circular patches in the field. Biology and epidemiology. Root-knot nematodes typically have very broad host ranges with some differences evident between species. M. incognita, for example, has more than 700 hosts. These include most cultivated crops and ornamentals.

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Root-knot nematodes are sedentary endoparasites. Some species are parthenogenic, not requiring males for reproduction. Second-stage infective juveniles hatch from the eggs. These invade roots in the region of elongation near the root cap. They migrate between and through cells and position themselves with the head in the vascular tissues. Cell damage occurs as a result of the migration and if several juveniles enter the root tip, cell division stops and there is no root elongation. As feeding continues, several cells near the head begin to enlarge and become multinucleate. These are called giant cells and there are usually 3–6 associated with each female nematode. The formation of giant cells and galls is the result of cell enlargement and of increased numbers of cells. Giant cells are induced by salivary secretions that are introduced into cells and surrounding tissues during feeding by the nematode. During this process, the xylem vessels become disrupted and the roots cannot function normally with respect to water and nutrients. During the process of gall formation, the nematodes undergo second, third, and fourth molts to reach the adult stage. Mature females are saccate (pear-shaped) and lay eggs into a gelatinous matrix. This matrix may protrude from the surface of small roots or may be entirely within the gall. Eggs hatch in about 7 d. The length of the life cycle is dependent on species as well as soil temperature. For M. incognita, the entire life cycle is completed in 20–25 d at 21  C (70  F). A subsequent generation can be borne in as few as 2 wk, depending on species and environment, and eggs can remain viable in the soil for a year (Mitkowski and Abawi 2003). Males, when present, remain vermiform (Westerdahl and Frate 2007). Management. Preplant treatment of soil with nematicides can reduce disease risk in areas known to contain root-knot nematodes (Mitkowski and Abawi 2003). Adjusting planting dates to cooler times of the season when nematodes are less active can help to reduce damage. Providing optimal conditions for plant growth including sufficient irrigation and soil amendments can help make plants more tolerant to nematode infestation.

5.5

Needle Nematode: (Longidorus spp.)

Geographic occurrence and impact. India, Israel, South Africa, the USA, and Zimbabwe (Goodey et al. 1965; Siddiqui et al. 1973). Symptoms/signs. Needle nematode is devastating to seedling root systems. Plants are stunted and galls are formed at root tips (Radewald et al. 1969). Because these symptoms could be indicative of problems other than nematodes, soil and root samples should be sent to a nematode diagnostic laboratory for evaluation. Biology and epidemiology. Needle nematodes are migratory ectoparasites with all stages being found outside of roots in the soil. They prefer coarse, well-drained soils. The nematode feeds at root tips (Fig. 19). Males are rare and reproduction is probably by parthenogenesis. A first-stage juvenile hatches from the egg. For

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Fig. 19 Ectoparasitic nematode feeding on root tip (Department of Entomology and Nematology, UC Davis, Slide Collection # 2017. All Rights Reserved.)

L. africanus, the life cycle from adult to adult is 7 to 9 wk at 28  C/82  F, and it can survive in fallow soil for at least 3 mo at 25  C/77  F (Radewald et al. 1969). This genus is capable of transmitting Strawberry latent ringspot nepovirus and other viruses to plants (Adekunle et al. 2006), and this can result in asymmetrical opening of flowers (Cohen et al. 1995). Management. Preplant nematicides have been shown to be effective. For L. africanus, timing of planting of crops to avoid temperature conditions (above 16  C/61  F) favorable to the nematode has been shown to reduce damage (Kolodge et al. 1979; Radewald et al. 1969). Additional information on nematode management may be found in ▶ Chap. 8, “Soil/Media Disinfestation for Management of Florists’ Crops Diseases”.

6

Physiological Disorders

Physiological disorders are those not associated with pathogens such as bacteria, fungi, or viruses. They are sometimes referred to as abiotic disorders and are caused by numerous factors (nonoptimal light levels, temperatures, soil aeration, nutrition, water availability, or presence of toxins) that adversely affect plant growth. For example, the presence of ethylene in the greenhouse atmosphere may cause flower abortion, a physiological or abiotic disorder. Or suboptimal root zone aeration may interfere with normal root function, leading to reduced micronutrient uptake that is expressed as a nutrient deficiency. One must be careful, however, as it is conceivable that a root disease organism could cause similar symptoms as a result or reduced root functionality from the infection. Thus, diagnosis of abiotic or physiological disorders should include, as much as possible, the elimination of biotic causes. Below is a summary of several specific physiological disorders that occur on lilies.

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Upper Leaf Necrosis (ULN)

This disorder has long been present in forced (greenhouse grown) lilies, most especially in the Oriental hybrid group. It is characterized by necrotic, usually twisted young upper leaves associated with the buds (Fig. 20). The disorder was specifically named ULN by Chang and Miller (2003), and their extensive research has led to a much clearer understanding of this disorder (Chang and Miller 2004, 2005a; Chang et al. 2004). It is also seen on lilies in the Asiatic and LA-hybrid groups, where it presents significantly different symptoms of white-gray bands across the leaves in the upper third of the plant, with the leaf tips remaining green (Berghoef 1986) (Fig. 21). The disorder appears during greenhouse production of cut flowers or potted plants and is a calcium deficiency disorder. Chang and Miller’s work demonstrated the relationship to calcium in a number of ways and included analysis of symptomatic and non-symptomatic tissue (Chang and Miller 2005a), alterations in transpiration of young leaves (Chang and Miller 2004), applications of exogenous calcium (Chang and Miller 2005b), demonstration of increased symptom severity in plants growing from bulbs depleted in calcium (Chang and Miller 2003), reduction of the problem as a result of daily foliar calcium applications (Chang et al. 2004), the relationship of severity of the disorder to bulb size (Chang and Miller 2005b), and very significant differences in cultivar susceptibility (Chang et al. 2008). Chang’s research was key in dispelling several “industry myths” about this disorder, namely, that it was caused by a warm, sunny day following an extended period of dark, cloudy weather. Onset of the disorder bore no relationship to prior weather conditions (Chang and Miller 2005a). A key realization was that plants growing from larger bulbs, which have much greater sensitivity to the disorder, have more buds and young, developing leaves associated with them. It is these very leaves that suffer from reduced transpiration and calcium uptake that eventually show calcium deficiency, expressed as upper leaf necrosis. While foliar sprays of calcium salts (nitrate or chloride) can be effective, daily sprays were necessary (Chang et al. 2004) and are mostly impractical. Berghoef Fig. 20 Upper leaf necrosis (ULN) on an Oriental hybrid lily cultivar (William B. Miller # 2017. All Rights Reserved.)

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Fig. 21 Calcium deficiency, essentially the same disorder as ULN, as expressed on an Asiatic hybrid lily cultivar (William B. Miller # 2017. All Rights Reserved.)

(1986) similarly reported the potential for phytotoxicity as a result of the required calcium sprays.

6.2

Leaf Scorch

“Leaf scorch” is a term that should be used carefully when discussing lilies as much Dutch literature will refer to ULN (see above) as “leaf scorch”. Here, however, we take this term to mean leaf injury caused by fluoride toxicity from root uptake. Specifically, this injury presents as necrotic areas on the leaf tip or margin, with a surrounding yellow (chlorotic) halo. This problem was commonly seen in Easter lily (Lilium longiflorum) crops in the 1960s and 1970s before Marousky and Woltz (1975) Woltz and Marousky (1975) demonstrated that fluoride in fertilizer solutions (commonly seen in superphosphate fertilizer) caused leaf scorch in lily if applied in sufficient amounts to plants grown in acid soils (pH 5.0). There are significant cultivar differences (e.g., the now-extinct cultivar “Ace” was more susceptible to the disorder than is “Nellie White”), and similar differences probably exist in modern cultivars. Fluoride-induced leaf scorch is readily controlled by manipulation of root zone pH, with a more alkaline reaction reducing fluoride availability and uptake.

6.3

Lower Leaf Yellowing

Whether grown as pot plants or cut flowers, many lilies are susceptible to chlorosis and yellowing of the lower leaves. There are two main classes of leaf yellowing in Lilium (Miller 2013). The first is yellowing and gradual senescence of lower leaves during crop growth in the greenhouse (Fig. 22). This is essentially a nutritional disorder (principally nitrogen deficiency) that can be caused by suboptimum fertilizer applications or by root injury from any cause, whether biotic (root pathogens) or abiotic (cold root zone, hypoxic root zone). The symptoms develop over several

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Fig. 22 Lower leaf yellowing, of the “gradual” or “greenhouse” type (William B. Miller # 2017. All Rights Reserved.)

weeks and can usually be corrected by increasing fertilizer application, using a different root substrate and altering (usually reducing) irrigation frequency. A more interesting lower leaf disorder is one that leads to rapid loss of chlorophyll from most if not all of lower to mid-leaves, usually in the retail or consumer phase, which has been described in detail and has been termed “catastrophic leaf yellowing” (Fig. 23) (Miller 2013). This disorder is exacerbated by prolonged cold storage which commonly occurs in the potted plant or cut flower postharvest chain. The disorder gained prominence with Easter lily in the late 1980s in the USA, when retailers reported that plants turned yellow within days of being placed in the retail environment. Research ultimately indicated that in addition to postharvest cold storage, the problem is exacerbated by anti-gibberellin growth regulators (used for reducing height for potted plant production) (Ranwala et al. 2000), and by growing plants in “negative DIF” environments, where night temperatures are warmer than day temperatures, also a height control technique (Ranwala et al. 2000). While the initial work on this problem focused on Lilium longiflorum, subsequent research has examined lower leaf yellowing-related disorders on hybrid lilies. Ranwala and Miller (2005) found significant differences in hybrid lily cultivar susceptibility to cold-storage-induced lower leaf yellowing. Practical solutions based on exogenous application of gibberellin4+7 (GA4+7) have been developed for the industry to reduce this problem (Fig. 24). Gibberellin4+7 sprays are highly effective in reducing or eliminating lower leaf yellowing in the postharvest phase. In various regions of the world, registered GA products include Fascination (Valent BioSciences), Fresco (Fine Americas), BVB (Chrysal), and Bulb 100 (Floralife). While these products contain equal concentrations of GA4+7 and benzyladenine, research has shown that benzyladenine is essentially inert against leaf yellowing and nearly all the activity is from the GA4+7 (Ranwala and Miller 1998, 1999, 2000). For pot plant production, sprays of 50–100 ppm are applied one to two times during plant production. The details of how this is done are quite involved and beyond the scope of this review. The essential concerns are that the gibberellin has the potential to cause stem elongation (undesirable for pot plants); furthermore, elongation

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Fig. 23 Leaf yellowing of the “catastrophic” type, after 2 wk in a 4  C/39  F dark cooler prior to being placed in a postharvest evaluation room. There was no leaf yellowing at the start of the cold treatment (William B. Miller # 2017. All Rights Reserved.)

Fig. 24 These plants were treated with GA4+7 and held in a 4  C/39  F cooler along with the plants in Fig. 23. The gibberellin inhibits the development of catastrophic leaf chlorosis (William B. Miller # 2017. All Rights Reserved.)

can come from GA sprayed on the aboveground part of the plant or through root uptake as a result of spray contacting the soil surface (Ranwala et al. 2003). Successful use of this product on Easter lilies in North America is predicated on careful spraying to only contact lower leaves and applying enough material to uniformly contact all leaf surfaces (to avoid green or yellow islands) while minimizing spray volume so as to reduce to runoff into the root zone. For cut flowers, while plants may be sprayed with GA4+7 before harvest with excellent results, (Ranwala and Miller 2005), a more practical treatment is after cutting, while stems are being held in liquid before packing. In this case, stems are held in buckets with dilute GA4+7 and the GA moves into the stem and leaves with the xylem flow. For both potted plant and cut flower systems, the GA4+7 treatment has an added benefit of extending individual flower life and life of the entire stem (inflorescence) (Ranwala et al. 2000; Ranwala and Miller 2002, 2005).

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Flower Bud Abortion and Abscission

The basis of most terminology related to floral abortion and abscission is derived from De Hertogh et al. (1971), Miller (1993), and Beattie and White (1993). Essentially, cessation of normal growth of a bud leading to its death at any stage is “abortion.” Bud or flower “blast” is abortion, but only after all floral parts are visible. In common use, North American growers tend to refer to bud abortion as that occurring at a very early stage, when only a small remnant of the aborted bud is present (Fig. 25), and to “blasted” buds as those that die when larger than ca. 0.5 cm/ 0.2 in. (Fig. 26). Abscission is the straightforward detaching of the bud from the plant. Many abiotic and biotic factors cause bud abortion, blasting, or abscission, and the hybrid groups show broad differences in their susceptibility to each factor. For example, Asiatic hybrid lilies are broadly susceptible to flower abscission and bud blast (buds usually less than 2 cm/0.8 in. long) from ethylene, suboptimal light, and supraoptimal temperatures in early crop growth (Durieux 1975; Durieux et al. 1982; Elgar et al. 1999). Oriental hybrid cultivars are much less susceptible to bud abortion and abscission overall, but small buds (less than ca. 1–3 cm/0.4–1.2 in. long) are readily injured by ethylene. L. longiflorum cultivars are susceptible to bud abortion and blast especially in early-Easter forcing years when growers are growing plants at warmer temperatures early in the crop (essentially in early January) when light levels are very low. Bud abscission is not commonly seen in L. longiflorum. There are a number of management practices that help reduce the risk of bud abortion and abscission. These include avoiding forcing temperatures above 21  C (71  F) (especially for Asiatic hybrid lilies), forcing lilies under glass instead of plastic, and providing supplemental (assimilation) lighting during periods of low light. Greenhouse-scale treatments of the gaseous anti-ethylene compound 1-methylcyclopropene (1-MCP) are financially impractical but would likely be effective. Sprayable formulations of 1-MCP are available in agriculture but are not presently registered for use in North America or other areas of the world. Postharvest

Fig. 25 An Asiatic hybrid lily cultivar showing an aborted bud, barely visible at the base of the two top leaves. Two “blasted” buds are visible immediately beneath these two leaves (William B. Miller # 2017. All Rights Reserved.)

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Fig. 26 Buds of Lilium longiflorum showing injury from ethylene exposure in a commercial greenhouse. Four buds are already dead or dying and a fifth (leftmost bud of the two upward-facing buds) may still fail to open (William B. Miller # 2017. All Rights Reserved.)

treatment of cut flowers with silver thiosulfate (STS) or 1-MCP also helps reduce the effects of ethylene and extends the vase life and quality of cut flowers.

6.5

Other Miscellaneous Disorders

Chemical Toxicity There are a few specific toxicities known in lilies. One is leaf tip injury from the fungicide metalaxyl. In L. longiflorum, this is expressed as a whitening of the tip (ca. last 1 cm/0.4 in.) of leaves. Forced lilies have been injured by root uptake of the herbicide glyphosate in certain subirrigation regimes (Weller and Hammer 1984). Another is fluoride-induced leaf scorch (see Sect. 6.2). Premature Flower Opening Atmospheric ethylene contamination is a common cause of prematurely-opening buds in a number of lily cultivars (Fig. 27). Bud “Split” This is an uncommon disorder, with no clearly defined cause. Aphids have been attributed to this problem (Post 1941), yet split buds are typically observed to be insect-free. Leaf Epinasty Downward-curling leaves can be caused by a number of factors including negative DIF growing environments (i.e., warmer nights and cool days, as used for height management in pot plant crops), by ethylene (Blankenship et al. 1993) and by growing crops at very low temperatures. Epinasty is also commonly seen in mechanically-stimulated plants, a technique which is also potentially useful for height control. Epinasty is generally reversible by eliminating its cause.

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Fig. 27 Two buds on this Lilium longiflorum plant are opening prematurely as a result of ethylene exposure in a commercial greenhouse (William B. Miller # 2017. All Rights Reserved.)

Stems Not Growing (“No Shows”) Stems may fail to grow out of the bulbs for a number of reasons, some related to preplant storage and others to conditions after planting. In the last 15–20 years, the lily industry has adopted frozen storage of bulbs to allow year-around planting and flower availability. Young shoots can be killed by too cold temperatures or by freezing the bulbs the wrong time (Gude and Kok 2005). Similarly, as the cold temperature (vernalization) tends to improve uniformity of sprouting of the population, suboptimal conditions during vernalization can lead to very slow and irregular emergence of the sprouts. Usually, the causes would be lack of oxygen, non-moist conditions or too short of a vernalization period (Miller 1992). During cooling, sprouts can grow out of the bulb and they are very susceptible to physical breakage. Malformed Buds Developing flower buds sometimes shows abnormalities such as malformed, twisted petals and longitudinal openings along the valve of the bud. This can be caused by high temperatures in the greenhouse after extended periods of frozen storage (Lee and Roh 2001). Postharvest Bud Necrosis Certain Oriental hybrid cultivars rapidly develop necrotic patches of blotches on unopened buds when placed in cold storage after harvest; essentially, chilling injury (Fig. 28). The problem is exacerbated by elevated greenhouse temperatures and high light levels before harvest and also by cold storage temperatures below approximately 4  C/39  F (Kim and Miller 2008).

Black Sprouts “Black sprouts” is a problem in lily flower production especially for Oriental hybrid cultivars forced at late dates. During freezer storage, the sprouts in the lily bulb can be killed, become black, and do not flower. Delaying freezer storage

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Fig. 28 Typical symptoms of “postharvest bud necrosis” which develops during cold storage of susceptible Oriental hybrid lily cultivars (William B. Miller # 2017. All Rights Reserved.)

until the bulbs are mature and respiration is low reduces this problem (Gude and Dijkema 2010).

Nutrient Deficiency A number of nutrient deficiencies can occur in lily pot and flower production (IBC 2008). Barnes et al. (2011) concluded that symptoms of nitrogen (N), sulfur (S), boron (B), and iron (Fe) deficiencies and boron (B) toxicity are the first disorders to be manifested in Easter and hybrid lilies. In their experiments, they did not get visual symptoms when plants were grown under phosphate (P), potassium (K), magnesium (Mg), copper (Cu), manganese (Mn), molybdenum (Mo), and zinc (Zn) deficient conditions. Deficiency symptoms may not appear in some instances because the bulb may provide a source of these elements. The climate conditions in the greenhouse, moisture deficit, pH, and electrical conductivity (EC) of the soil are also important (Kok 2013). Lilies are sensitive to salt. Soil with an EC higher than 1.2 can give salt damage and hinder the absorption of nutrients. Iron and Mg are hard for lily plants to absorb in a high pH soils, so deficiency can occur. High relative humidity reduces mineral uptake and transport, exacerbating certain deficiencies, especially calcium (Ca, see Sect. 6.1).

References Adekunle OK, Kulshrestha S, Prasad R, Hallan V, Raikhy G, Verma N, Ram R, Kumar S, Zaidi AA (2006) Plant parasitic and vector nematodes associated with Asiatic and oriental hybrid lilies. Bioresour Technol 97:364–371 Aegerter BJ, Greathead AS, Pierce LE, Davis RM (2002) Mefenoxam-resistant isolates of Pythium irregulare in an ornamental greenhouse in California. Plant Dis 86(6):692 Agrios GN (2005) Plant pathology, 5th edn. Elsevier Academic Press, Burlington Alabouvette C, Schippers B, Lemanceau P, Bakker PAHM (1998) Biological control of Fusarium wilts: toward development of commercial products. In: Boland GJ, Kuykendall LD (eds) Plantmicrobe interactions and biological control. Marcel Dekker, New York, pp 15–36

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Alabouvette C, Olivain C, L’Haridon F, Aimé S, Steinberg C (2005) Using strains of Fusarium oxysporum to control wilts: dream or reality? In: Vurro M, Gressel J (eds) Novel biotechnologies for biocontrol agent enhancement and management. Springer, Dordrecht, pp 157–177 Alabouvette C, Olivain C, Migheli Q, Steinberg C (2009) Microbiological control of soil-borne phytopathogenic fungi with special emphasis on wilt-inducing Fusarium oxysporum. New Phytol 184:529–544 Alfieri SA, Langdon KR, Kimbrough JW, El-Gholl NE, Whelburg C (1984) Diseases and disorders of plants in Florida, Florida Department of Agriculture and Consumer Services, Division of Plant Industry Bulletin # 14, Gainesville Allen TC (1974) Control of viruses in lilies. In: Lilies 1974 and other Liliaceae. Royal Horticultural Society, London, pp 37–45 Allen TC (1975) Viruses of lilies and their control. Acta Hortic 47:69–75 Allen TC, Anderson WC (1980) Production of virus-free ornamental plants in tissue culture. Acta Hortic 110:245–251 Allen TC, Fernald K (1972) Elimination of viruses in hybrid lilies. Lily Yb N Am Lily Soc 25:53–55 Allen TC, Linderman RG (1976) Rate of infection of virus-free lilies with lily symptomless virus in the field. Acta Hortic 59:37–38 Allen TC, Lyons AR (1969) Electron microscopy of lily symptomless virus and cucumber mosaic virus within fleck diseased lilies. Phytopathology 59:1318–1322 Alvarez MG (1976) Primer catalogo de enfermedades de plantas Mexicanas. Fitofilo 71:1–169 Amiri A, Heath SM, Peres NA (2013) Phenotypic characterization of multifungicide resistance in Botrytis cinerea isolates from strawberry fields in Florida. Plant Dis 97:393–401 Anonymous (1960) Index of plant diseases in the United States. USDA Agric Handb 165:1–531 Anonymous (2009) Floriculture and ornamental nurseries – Easter lily (Lilium longiflorum) disease control outlines. How to manage pests. UC pest management guidelines. http://www.ipm. ucdavis.edu/PMG/r280111311.html. Accessed 1 Jan 2016 Anonymous (2010) Plantago asiatica mosaic virus on Lilium spp. Pest report – The Netherlands. Plant Protection Service of the Netherlands, Wageningen, pp 1–2 Arens P, Shahin A, Van Tuyl JM (2014) (Molecular) breeding of Lilium. Proc. IIIrd IS on the genus Lilium. Acta Hortic 1027:113–128 Ark PA, MacLean NA (1951) Botrytis spot and blight of tuberoses in California. Plant Dis Rep 35:45–46 Asjes CJ (1974) Soil-borne virus diseases in ornamental bulbous crops and their control in The Netherlands. Agric Environ 1:303–315 Asjes CJ (1976) Some developments of the virus pathology in lilies in the Netherlands. Lily Yb N Am Lily Soc 29:120–126 Asjes CJ (1991) Control of air-borne field spread of tulip breaking virus, lily symptomless virus and lily virus X in lilies by mineral oils, synthetic pyrethroids, and a nematicide in the Netherlands. Neth J Plant Path 97:129–138 Asjes CJ (1997) Virus in bloembollen in kaart gebracht. Bloembollencultuur 108(4):50 Asjes CJ (2000) Control of aphid-borne lily symptomless virus and lily mottle virus in Lilium in The Netherlands. Virus Res 71:23–32 Asjes CJ, Segers LC (1983) Incidence and control of necrotic leaf mosaic caused by Arabis mosaic virus in Lilium tigrinum splendens in the Netherlands. J Phytopathol 106:115–126 Asjes CJ, Vos de NP, Van Slogteren DHM (1973) Brown ring formation and streak mottle, two distinct syndromes in lilies associated with complex infections of lily symptomless virus and tulip breaking virus. Neth J Plant Path 79:23–35 Aycock R (1966) Stem rot and other diseases caused by Sclerotium rolfsii, North Carolina Agricultural Experiment Station Technical Bulletin no 174, 202pp Baayen RP, Förch MG (2001) Transfer cell formation reveals a biotrophic phase in bulb rot of lilies infected by Fusarium oxysporum f.sp. lilii. In: Summerell BA, Leslie JF, Backhouse D, Bryden WL, Burgess LW (eds) Fusarium: Paul E. Nelson memorial symposium. APS Press, St. Paul, 408p

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Baayen RP, Rijkenberg FHJ (1999) Fine structure of the early interaction of lily roots with Fusarium oxysporum f.sp. lilii. Eur J Plant Path 105:431–443 Baayen RP, Forch MG, Waalwijk C, Bonants PJM, Loffler HJM, Roebroeck EJA (1998) Pathogenic, genetic and molecular characterization of Fusarium oxysporum f.sp. lilii. Eur J Plant Pathol 104:887–894 Bald JG (1979) Stem lesion of Easter lilies – a complex disease. Calif Agr 33:12–13 Bald JG, Chandler PA (1957) Reduction of the root rot complex on ‘Croft’ lilies by fungicidal treatment and propagation form bulb scales. Phytopathology 47:285–291 Bald JG, Solberg RA (1960) Antagonism and synergism among organisms associated with scale tip rot of lilies. Phytopathology 34:966–975 Bald JG, Paulus AO, Lenx JV, Chanadler PA, Susuki T (1969) Disease control with pathogen-free bulb stocks for Easter lily improvement. Calif Agr 23(11):6–8 Bald JG, Paulus AO, Lenx JV (1983) Control of field root and bulb diseases of Easter lilies. Plant Dis 67:1167–1172 Balode A (2009) Breeding for resistance against Botrytis in lily. Acta Hortic 836:143–148 Balode A (2011) Effect of biological product trihodermin B-J on Botrytis suppression and lily bulb development. Acta Hortic 900:89–94 Barnes J, Whipker BE, McCall I, Frantz J (2011) Characterization of nutrient disorders of Lilium longiflorum ‘Nellie White’ and Lilium hybrid ‘Brunello’. Acta Hortic 900:205–209 Bastiannse C, Koster ATJ, van der Meer LJ, van den Ende DJE, Pennock I, Buurman FPM (1997) A disease-forecasting system of botrytis blight (‘fire’) in lily. Acta Hortic 430:657–660 Beattie DJ, White JW (1993) Lilium – hybrids and species. In: De Hertogh AA, Le Nard M (eds) Physiology of flower bulbs. Elsevier, Amsterdam, pp 423–454 Beers CM, Barba-Gonzalez R, Van Silfhout AA, Ramanna MS, Van Tuyl JM (2005) Mitotic and meiotic polyploidization in lily hybrids for transferring Botrytis resistance. Acta Hortic 673:449–452 Beijersbergen JCM, van der Hulst CTC (1980) Detection of lily symptomless virus (LSV) in bulb tissue of lily by means of ELISA. Acta Hortic 109:487–493 Benschop M, Kamenetsky R, Le Nard M, Okubo H, De Hertogh A (2010) The global flower bulb industry: production, utilization, research. In: Horticultural reviews. John Wiley & Sons, Hoboken, pp 1–115 Berghoef J (1986) Effect of calcium on tipburn of Lilium ‘Pirate’. Acta Hortic 177:433–438 Bertaccini A, Kamińska M, Botti S, Martini M (2002) Molecular evidence for mixed phytoplasma infection in lily plants. Acta Hortic 568:35–41 Bertaccini A, Fránová J, Botti S, Tabanelli D (2005) Molecular characterization of phytoplasmas in lilies with fasciation in the Czech Republic. FEMS Microbiol Lett 249:79–85 Bishop CD, Cooper RM (1983) An ultrastructural study of root invasion in three vascular wilt diseases. Physiol Mol Plant Pathol 22:15–27 Blankenship SE, Bailey DA, Miller JE (1993) Effects of continuous, low levels of ethylene on growth and flowering of Easter lily. Sci Hort 53:311–317 Bobev, S. (2009) Reference Guide for the Diseases of Cultivated Plants, 466 pp. [http://nt.ars-grin. gov/fungaldatabases.] Boerema GH, Hamers MEC (1988) Check-list for scientific names of common parasitic fungi. Series 3a: fungi on bulbs: Liliacea. Neth J Plant Pathol 94(suppl 1):1–32 Bollen GJ (1971) Resistance to benomyl and some chemically related compounds in strains of Penicillium speices. Neth J Plant Path 77:187–193 Bollen GJ (1977) Pathogenicity of fungi isolated from stems and bulbs of lilies and their sensitivity to benomyl. Neth J Plant Pathol 83(suppl 1):317–329 Boontjes J (1983) Virusvrije lelies zijn groter areaal waard. Vakblad voor de Bloemisterij 38(18):36–39 Brent KJ (1995) Fungicide resistance in crop pathogens: how can it be managed? FRAC monograph no 1. GIFAP, Brussels, 49p Byther RS, Chastagner GA (1993) Diseases. In: DeHertogh AA, LeNard M (eds) The physiology of flower bulbs. Elsevier, Amsterdam, pp 71–99

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Mordue JEM (1974b) Sclerotium rolfsii, CMI descriptions of pathogenic fungi and bacteria no 410. CAB International, Wallingford Mordue JEM, Holliday P (1976) Sclerotinia sclerotiorum, CMI descriptions of pathogenic fungi and bacteria no 513. CAB International, Wallingford Mullen J (2001) Southern blight, southern stem blight, White mold. Plant Health Instr. doi:10.1094/ PHI-I-2001-0104-01 Muller PJ, Vink P, Van Zaayen A (1988) Flooding causes loss in viability and pathogenicity of sclerotia of Rhizoctonia tuliparum. Neth J Plant Path 94:45–47 Nesi B, Trinchello D, Lazzereschi S, Grassotti A (2009) Production of lily symptomless virusfree plants by shoot meristem tip culture and in vitro thermotherapy. Hortscience 44(1): 217–219 Núñez de Cáceres González F, Davey M, Cancho Sanchez E, Wilson Z (2015) Conferred resistance to Botrytis cinerea in Lilium by overexpression of the RCH10 chitinase gene. Plant Cell Rep 34 (7):1201–1209 Okabe I, Matsumoto N (2003) Phylogenetic relationship of Sclerotium rolfsii (teleomorph Athelia rolfsii) and S. delphinii based on ITS sequences. Mycol Res 107:164–168 Okubo H (2014) History of Lilium species in Asia. Acta Hortic 1027:11–26 Oostenbrink M, Jacob JJ, Kuiper K (1957) Tagetes als Feindpflanzen von Pratylenchus-Arten. Nematologica 2:424–433 Orieux L, Felix S (1968) List of plant diseases in Mauritius. Phytopathol Pap 7:1–48 Overman AJ (1961) Pre-storage treatment of lily bulbs with nematicides. Proc Fla State Hortic Soc 74:386–388 Overy DP, Frisvad JC, Steinmeier U, Thrane U (2005) Clarification of the agents causing blue mold storage rot upon various flower and vegetable bulbs: implications for mycotoxin contamination. Postharvest Biol Technol 35:217–221 Palmer C, Vea E (2010) IR-4 ornamental horticulture program Phytophthora efficacy: Phytophthora cactorum, Phytophthora cinnamomi, Phytophthora citricola, Phytophthora cryptogea, Phytophthora dreschleri, Phytophthora nicotianae/Phytophthora parasitica, Phytophthora palmivora, Phytophthora ramorum, Phytophthora syringae, Phytophthora tropicalis. The IR-4 Project, Rutgers University. http://ir4.rutgers.edu/Ornamental/SummaryReports/ PhytophthoraDataSummary2010.pdf. Accessed 10 Dec 2015 Palmer C, Vea E (2014) IR-4 ornamental horticulture program Botrytis efficacy: A literature review. The IR-4 Project, Rutgers University. http://ir4.rutgers.edu/Ornamental/SummaryReports/ BotrytisEfficacySummary_2014.pdf. Accessed 10 Dec 2015 Parris GK (1959) A revised host index of Mississippi plant diseases. Mississippi State Univ Bot Dept Misc Publ 1:1–146 Paulitz TC, Bélanger RR (2001) Biological control in greenhouse systems. Annu Rev Phytopathol 39:103–133 Pennycook SR (1989) Plant diseases recorded in New Zealand, 3 vol, Plant Diseases Division. DSIR, Auckland Plakidas AG (1944) Black scale: a disease of Easter lily bulbs. Phytopathology 34:556–571 Poncarová-Vorácková Z, Franová ZJ, Valová P, Mertelik BP, Navrátil M, Nebesárová J (1998) Identification of phytoplasma infecting Lilium martagon in the Czech Republic. J Phytopathol 146:609–612 Post K (1941) Problems in forcing Easter lilies. Proc Am Soc Hortic Sci 39:415–418 Prince TA, Cunningham MS, Peary JS (1987) Floral and foliar quality of potted Easter lilies after STS or phenidone application, refrigerated storage, and simulated shipment. J Am Soc Hortic Sci 112:469–473 Pscheidt JW, Ocamb CM (2015) Pacific Northwest plant disease control handbook. Oregon State University. http://pnwhandbooks.org/plantdisease/. Accessed 10 Dec 2015 Pudasaini MP, Viaene N, Moens M (2006) Effect of marigold (Tagetes patula) on population dynamics of Pratylenchus penetrans in a field. Nematology 8(4):477–484

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Putnam ML, Miller ML (2007) Rhodococcus fascians in Herbaceous Perennials. Plant Dis 91:1065–1076 Qiu J, Westerdahl BB, Giraud D, Anderson CA (1993) Evaluation of hot water treatments for management of Ditylenchus dipsaci and fungi in daffodil bulbs. J Nematol 25:686–694 Raabe RD (1985) Rust diseases. In: Strider DL (ed) Diseases of floral crops. Praeger, New York, pp 173–227 Raabe RD, Hurlimann JH (1971) Control of Easter lily root rots with fungicides. Lily Yb N Am Lily Soc 24:11–13 Radewald JD, Osgood JW, Mayberry KS, Paulus AO, Shibuya F (1969) Longidorus africanus a pathogen of head lettuce in the Imperial Valley of southern California. Plant Dis Rep 53:381–384 Ranwala AP, Miller WP (1998) Gibberellin4+7, benzyladenine and supplemental light improve postharvest leaf and flower quality of cold-stored ‘Stargazer’ hybrid lilies. J Am Soc Hort Sci 123:563–568 Ranwala AP, Miller WP (1999) Timing of gibberellin4+7 + benzyladenine sprays influences efficacy against foliar chlorosis and plant height in Easter lily. Hortscience 34:902–903 Ranwala AP, Miller WP (2000) Preventive mechanisms of gibberellin4+7 and light on dark low temperature-induced leaf senescence in Lilium cv. Stargazer. Postharvest Biol Technol 19:85–92 Ranwala AP, Miller WP (2002) Effects of gibberellin treatments on flower and leaf quality of cut hybrid lilies. Acta Hortic 570:205–210 Ranwala AP, Miller WP (2005) Effects of cold storage on postharvest leaf and flower quality of potted oriental-, Asiatic- and LA-hybrid lily cultivars. Sci Hortic 105:383–392 Ranwala AP, Legnani G, Miller WB (2003) Minimizing stem elongation during spray applications of gibberellins4+7 and benzyladenine to prevent leaf chlorosis in Easter lilies. HortScience 38:1210–1213 Ranwala AP, Miller WP, Kirk TI, Hammer PA (2000) Ancymidol drenches, reversed greenhouse temperatures, post-greenhouse cold storage and hormone sprays affect post-harvest leaf chlorosis in Easter lily. J Am Soc Hort Sci 125:248–253 Ribeiro OK (1978) A source book of the genus Phytophthora. CAB, Wallingford, 417pp Ribeiro OK (2013) A historical perspective of Phytophthora. In: Lamour K (ed) Phytophthora: a global perspective. CAB International, Wallingford, pp 1–10 Roberts AN, Stang JR, Wang YT, McCorkle WR, Riddle LJ, Moeller FW (1985) Easter lily growth and development. Technical Bulletin 148. Agricultural Experiment Station, Oregon State University, Corvallis, 74p Schnitzler WH (2004) Pest and disease management of soilless culture. Acta Hortic 648:191–203 Shahin A, Arens P, van Heusden S, Van Tuyl JM (2009) Conversion of molecular markers linked to Fusarium and virus resistance in Asiatic lily hybrids. Acta Hortic 836:131–136 Shahin A, Arens P, Van Heusden AW, Van der Linden G, Van Kaauwen M, Khan N, Schouten H, Van de Weg WE, Visser RGF, Van Tuyl JM (2010) Genetic mapping in Lilium: mapping of major genes and QTL for several ornamental traits and disease resistances. Plant Breed 130(3):372–382 Shaw CG (1973) Host fungus index for the Pacific Northwest – I. Hosts. Washington State Univ Agric Exp Sta Bull 765:1–121 Sholberg PL, Conway WS (2016) Postharvest Pathology. In: Gross KC, Wang CY, Saltveit M, Lester G (eds) The commercial storage of fruits, vegetables, and florist and nursery crops, USDA Agriculture Handbook 66. USDA, Agricultural Research Service, Beltsville. http:// www.ba.ars.usda.gov/hb66/pathology.pdf. Accessed 27 Mar 2016 Siddiqi MR (1975) Aphelenchoides fragariae, C.I.H. descriptions of plant-parasitic nematodes, set 5, no 74. Commonwealth Institute of Parasitology, CAB International, 4 pages Siddiqui IA, Sher SA, French AM (1973) Distribution of plant parasitic nematodes in California. State of California Department of Food and Agriculture, Division of Plant Industry, Gainesville, 324p. Slate GL (1936) Disease among the lilies. Hortic Boston 14:96–97

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Slate GL (1972) Diseases of garden lilies and their control. North American Lily Society, Owatonna, MN. 20pp Sobers EK (1958) Colletotricha associated with lily bulbs. Their pathogenicity and host range. PhD dissertation, Louisiana State University, Baton Rouge, 92p Sobers EK (1980) Black scale of lily bulbs. Plant pathology circular no 33. Florida Department of Agriculture, March 1965, Division of Plant Industry (Revised July), 2p Sobers EK, Plakidas AG (1962) Colletotrichums associated with lily bulbs. Phytopathology 52:884–887 Sochacki D, Gabryszewska E, Kamińska M (2012) The application of ribavirin for elimination of viruses in lily plants growing in vitro. In: Bioforsk FOKUS 7(9), The 13th international symposium on virus diseases of ornamental plants, Norway, June 24–29. Staats M, van Baarlen P, van Kan JAL (2005) Molecular phylogeny of the plant pathogenic genus Botrytis and the evolution of host specificity. Mol Biol Evol 22:333–346 Staats M, van Baarlen P, van Kan JAL (2007) AFLP analysis of genetic diversity in populations of Botrytis elliptica and Botrytis tulipae from the Netherlands. Eur J Plant Pathol 117:219–235 Stone OM (1980) Two new potexviruses from monocotyledons. Acta Hortic 110:59–63 Straathof TP, van Tuyl JM (1995) Genetic variation in resistance to Fusarium oxysporum f.sp. lilii in the genus Lilium. Ann Appl Biol 125:61–72 Straathof TP, Jansen J, Loffler HJM (1993) Determination of resistance to Fusarium oxysporum in Lilium. Phytopathology 83:568–572 Summerell BA, Leslie JF, Backhouse D, Bryden WL, Burgess LW (eds) (2001) Fusarium: Paul E. Nelson memorial symposium. APS Press, Minnesota, 408pp Taylor AL, Sasser JN (1978) Biology, identification, and control of root-knot nematodes (Meloidogyne species). North Carolina State University Graphics, Raleigh Terhem RB, Staats M, van Kan JAL (2015) Mating type and sexual fruiting body of Botrytis elliptica, the causal agent of fire blight in lily. Eur J Plant Pathol 142:615–624 Tompkins CM, Hansen HN (1950) Glower blight of Stephanotis floribunda caused by Botrytis elliptica and its control. Phytopathology 38:114–117 Trolinger JC, Strider DL (1985) Botrytis diseases. In: Strider DL (ed) Diseases of floral crops, vol 1. Praeger Scientific, New York, pp 17–101 Tsukiboshi T (2002) Botrytis elliptica. NIAES, Microbial Systemics Lab. http://www.maes.affrc. go.jp/inventry/microorg/eng/zl6e-Botery.html. Accessed 15 Jan 2016 Turlier M-F, Eparvier A, Alabouvette C (1994) Early dynamic interactions between Fusarium oxysporum f.sp. lini and the roots of Linum usitatissimum as revealed by transgenic GUS-marked hyphae. Can J Bot 72:1605–1612 Van Aartrijk J, Van der Linde PCG (1986) In vitro propagation of flower bulb crops. In: Ammirato PV, Evans DR, Sharp WR, Bajaj YPS (eds) Handbook of plant cell culture. McGraw-Hill, New York, pp 317–331 Van Baarlen P, Staats M, van Kan JAL (2004) Induction of programmed cell death in lily by the fungal pathogen Botrytis elliptica. Mol Plant Pathol 5:559–574 Van Baarlen P, Legendre L, Van Kan JAL (2007) Plant defense compounds against Botrytis infections. In: Elad Y et al (eds) Botrytis: biology, pathology and control. Springer, Dordrecht, pp 143–161 Van den Ende JE, Pennock JG (1977) Influence of temperature and wetness duration on infection of lily by Botrytis elliptica. Acta Bot Neerl 46:332 Van den Ende JE, Pennock-Vos IMG (1997) Primary sources of inoculum of Botrytis elliptica in lily. Acta Hortic 430:591–595 Van den Ende JE, Pennock-Vos MG, Bastiaansen C, Koster ATJ, van der Meer LJ (2000) BoWaS: a weather-based warning system for the control of Botrytis blight in lily. Acta Hortic 519:215–220 Van Heusden AW, Jongerius MC, Van Tuyl JM, Straathof TP, Mes JJ (2002) Molecular assisted breeding for disease resistance in lily. Acta Hortic 572:131–138 Van Kesteren HA (1966) Bolrot van lelies, veroorzaakt door Sclerotium wakkeri (zwartbenigheid). Neth J Plant Path 72:314–316

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Van Tuyl JM, Arens P (2011) Lilium: breeding history of the modern cultivar assortment. Acta Hortic 900:223–230 Van Tuyl JM, Van Holsteijn HMC (1996) Lily breeding research in the Netherlands. Acta Hortic 414:35–45 Van Tuyl JM, Arens P, Marasek-Ciolakowska A (2012) Chapter 6, Breeding and genetics of ornamental geophytes. In: Kamenetsky R, Okubo H (eds) Ornamental geophytes: from basic science to sustainable production. CRC Press, Boca Roca, pp 131–158 Van Zaayen A, Asjes CJ, Bregnan D, Koster ATJ, Muller PJ, van der Valk GGM, Vos I (1986) Control of soil-borne diseases, nematodes and weeds in ornamental bulb cultivation by means of flooding. Acta Hortic 177:524 Vantomme R, Elia S, Swings J, DeLey J (1982) Corynebacterium fascians (Tilford 1936) Dowson 1942 the causal agent of leafy gall on lily crops in Belgium. Parasitica 38:138–192 Verbeek M, Stijger CCMM (2016) Strawberry latent ringspot virus in Lily is seed transmitted and localised in the embryo. In: 13th International Plant Virus Epidemiology Symposium, 6–10 June 2016, Avignon, p 154 Vidaver AK, Lambrecht PA (2004) Bacteria as plant pathogens. Plant Health Instr. doi:10.1094/ PHI-I-2004-0809-01 Vink P (2011a) Inundation kills sclerotia of Crown rot (in Dutch). BloembollenVisie 224:23 Vink P (2011b) Cause of stem rot of lilies can be Rhizopus mold (in Dutch). Bloembollen-Visie 226:21 Vrind TA (2005) The Botrytis problem in figures. Acta Hortic 669:99–102 Ward HM (1888) A lily disease. Ann Bot 2:319–382 Weintraub PG, Zeidan M, Spiegel S, Gera A (2007) Diversity of the known phytoplasmas in Israel. Bull Insectol 60(2):143–144 Weller SC, Hammer PA (1984) Susceptibility of Easter lily to glyphosate injury. Hortscience 19:698–699 Westerdahl BB, Frate CA (2007) Parasitic nematodes in alfalfa. Irrigated Alfalfa Manag 11:1–12. University of California Division of Agriculture and Natural Resources, Publication 8297 Westerdahl BB, Giraud D, Radewald JD, Anderson CA, Darso J (1993) Management of Pratylenchus penetrans on oriental lilies with drip and foliar applied nematicides. J Nematol 25(4S):758–767 Westerdahl BB, Giraud D, Etter S, Riddle LJ, Radewald JD, Anderson CA, Darso J (2003) Management options for Pratylenchus penetrans in Easter lily. J Nematol 35:443–449 Wiehe PO (1948) The plant diseases and fungi recorded from Mauritius. Mycol Pap 24:1–39 Woltz SS, Marousky FJ (1975) Fluoride leaf scorch of lily and gladiolus: soil acidity, superphosphate and diagnostic techniques. Florida State Horticultural Society, Gainesville, pp 609–612. Wu XW, Tian M, Wang LH, Cui GF, Yu RP, Lu ZH, Jia WJ, Qu SP, Gui M, Wang JH (2014) Native species of the genus Lilium in China. Acta Hortic 1027:27–40 Xu Z, Gleason ML, Mueller DS, Esker PD, Bradley CA, Buck JW, Benson DM, Dixon PM, Montiero JEBA (2008) Overwintering of Sclerotium rolfsii and S. rolfsii var. delphinii in different latitudes of the United States. Plant Dis 92:719–724 Yang XM, Wang JH, Qu SP, Wang L-H (2007) First report of southern blight caused by Sclerotium rolfsii on lily in China. Plant Dis 91:109 Yang XM, Wang JH, Qu SP, Wang LH, Peng LC (2010) First report of Phytophthora blight of lily caused by Phytophthora nicotianae in China. Plant Dis 94:782 Yunis H, Elad Y (1989) Survival of Botrytis cinerea in plant debris during summer in Israel. Phytoparasitica 17:13–21 Zehnder GW, Murphy JF, Sikora EJ, Kloepper JW (2001) Application of rhizobacteria for induced resistance. Eur J Plant Pathol 107:39–50 Zhang Z (2006) Flora Fungorum Sinicorum, vol 26, Botrytis, Ramularia. Science Press, Beijing, 277 pages

Diseases of Gladiolus

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Wade H. Elmer and Kathryn K. Kamo

Abstract

Gladiolus (Gladiolus spp.) are in the Iridaceae family and are native to South Africa, and over 200 species have been described. Gladiolus have become a major crop in the florist industry. Growers of gladiolus plant their corms in the spring and harvest the flower spikes during the summer and early fall. Although the crop can be propagated sexually from seeds, most of the industry is based on movement of corms and cormels, which leads to many diseases being disseminated with the crop. Fusarium corm rot, Gladiolus rust, and Curvularia spot are the most limiting fungal diseases, whereas Cucumber mosaic virus and Bean yellow mosaic virus emerge as the more threatening viral diseases affecting gladiolus. Keywords

Botrytis cinerea • Fusarium spp. • Potyvirus • Integrated disease management

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Botrytis Blight (Botrytis spp.): Botrytis gladiolorum Timm and B. cinerea Pers. . . 2.2 Curvularia Spot (Curvularia gladiolus Boerema and Hamers) . . . . . . . . . . . . . . . . . . . . . 2.3 Fusarium Yellows and Fusarium Corm Rot (Fusarium oxysporum f. sp. gladioli) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Gladiolus Rust (Uromyces transversalis Thum) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Stromatinia Rot [Stromatinia gladioli (Drayt.) Whet.] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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W.H. Elmer (*) Department of Plant Pathology and Ecology, The Connecticut Agricultural Experiment Station, New Haven, CT, USA e-mail: [email protected] K.K. Kamo USDA-ARS, Beltsville, MD, USA e-mail: [email protected] # Springer International Publishing AG (outside the USA) 2018 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florists’ Crops Diseases, Handbook of Plant Disease Management, https://doi.org/10.1007/978-3-319-39670-5_47

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3 Bacterial and Phytoplasma Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Bacterial Scab Burkholderia gladioli pv. gladioli (Formerly Pseudomonas gladioli pv. gladioli) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Aster Yellows (“Candidatus Phytoplasma asteris”) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Viral Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Bean yellow mosaic virus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Cucumber mosaic virus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Introduction

Gladiolus (Gladiolus spp.) have a high economic value and play a major role in the global flower bulb industry (Benschop et al. 2010). Gladiolus spp. belong to the Iridaceae family, together with Iris and Crocus (Meerow 2012). Over 260 species have been described and all originate from Africa, Madagascar, and Eurasia (Goldblatt et al. 2008). Hybridization of gladiolus began in the 1800s to produce new colors when the Colvillei hybrids were first produced (Hartline 2015). Since then, hundreds of crosses have been developed. Today, gladiolus are a major crop in the florist industry. Growers of gladiolus plant their corms in the spring and harvest the flower spikes during the summer and early fall. The crop is propagated sexually from seeds, and asexually from corms and cormels, but commercial flower spikes are obtained from mature corms. In most northern climates, the corms and cormels are usually dug and stored in the fall. Where data are available, the management of major gladiolus diseases is presented in the following sections. We advise the reader to inform him/herself with the general management strategies found in the introductory chapters on integrated disease management.

2

Fungal and Fungus-Like Diseases

2.1

Botrytis Blight (Botrytis spp.): Botrytis gladiolorum Timm and B. cinerea Pers.

Geographic occurrence and impact. Botrytis blight develops on all florist crops. The fungi are ubiquitous and can attack gladiolus in the field when environmental conditions are conducive, but it causes more problems reducing shelf life as a postharvest disease. The first major report of the disease was in Florida in 1940 (Dimock 1940). Symptoms/signs. Symptoms of Botrytis blight are characterized as three distinct spots: (1) very small, rust-colored spots that appear on only one side of the leaf; (2) small, yellowish brown spots that develop reddish brown margins; and (3) the large, oval spots that develop long, reddish margins (Magie 1956) (Fig. 1). Pinpoint water-soaked lesions also appear on flowers if the flower spike stays wet for at least 14 h (Magie 1956). Flower lesions enlarge and are first watery then dry and turn light

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Fig. 1 Botrytis blight of Gladiolus (R.D. Milhollland # 2017. All Rights Reserved.)

brown unless the relative humidity is very high, and then the whole lesion becomes wet and slimy. Latent infections are common and cause postharvest losses during transit to market (Magie 1956). The fungus causes basal stem infections which can extend downward to the corm causing a neck rot. In severe cases, the fungus infects the corms and continues to rot the tissue in cold storage conditions. Biology and epidemiology. There is much known about the Botrytis blight fungi. The fungi survive in soil as sclerotia and as mycelium on plant debris. The fungus requires very high humidity to infect and sporulate, so weather plays a key role in the development of the disease. The optimal temperature for disease development was found to be 20
 10
C (68  18
F), and disease severity will increase with increasing periods of leaf wetness reaching the maximum at 96 h (Sehajpal and Singh 2014). Management • Cultural practices – Reduction of the relative humidity and leaf wetness are critical for suppression of Botrytis blight. In field plants, growers should space plants in the row to allow for maximum airflow. Plants should be watered in the morning to allow time for leaves to dry. • Sanitation – Careful inspection of corms for lesions can reduce the damage of Botrytis blight. Remove and discard senescing blossoms and plant debris in packing sheds and during postharvest. • Fungicides – Botrytis blight is suppressed very effectively by fungicides. Care should be taken to rotate chemicals in different classes to delay the development of resistance. Many fungicides are effective against the disease. The fungicides chlorothalonil, iprodione, mancozeb, and strobilurins are all effective (Singh et al. 2005, 2008, 2011). The benzimidazoles may also be effective against certain species/strains, but resistance has emerged in many strains of B. cinerea. Copper oxychloride, triadimefon, ziram, difenoconazole, penconazole, hexaconazole, and tridemorph have also been shown to be effective. Sprays should be started before the appearance of the disease, preferably before the plants reach the flowering stage (Singh et al. 2005).

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Curvularia Spot (Curvularia gladiolus Boerema and Hamers)

Geographic occurrence and impact. The disease was first noted in Florida, USA, in 1948 (Magie 1948) and in Canada in 1954 (Parmelee 1954). It was identified as Curvularia lunata (Parmelee 1956), renamed as C. trifolii f. sp. gladioli, and renamed again in 1989 as C. gladioli Boerema and Hamers (Boerema and Hamers 1989). It has been reported in North America (Magie 1948), South America (Torres et al. 2013b), China, Europe (Parmelee 1954), India (Singh 1968), and Philippines (Mendiola-Ela 1952). There is little information on the economic significance of Curvularia leaf spot, but some cultivars are highly susceptible (Torres et al. 2013b). Symptoms/signs. The disease affects leaves, stems, and petals. Symptoms usually begin on leaves first as light to dark brown, oval spots. The symptomatic tissues show leaf spots that are oval to circular, brown with dark edges, and surrounded by a yellow halo. Often the lesions become necrotic, and the leaves acquire a dry and wilted appearance (Torres et al. 2013a). The fungus produces a black, powdery mass of spores in the center of the spot. Torres et al. (2013b) noted symptoms were a function of the host resistance. For example, on G. grandiflorus var. Red Beauty, the spots were more irregular and necrotic and surrounded by yellow halos. In other species, such as G. callianthus, Curvularia spots were round and always surrounded by yellow halos. More tolerant cultivars of G. grandifloras, such as Amsterdam, Friendship Rose, Tradehorn, Veronica, and Gold Yester, reacted differently to Curvularia leaf spot and exhibited light to dark brown spots. One report observed vascular discoloration (Forsberg 1957) Biology and epidemiology. C. gladioli is favored by warm, wet conditions. Infection occurs after a 13-h dew period, but actual lesions may take 5 days after infection to appear. Torres et al. (2013a) observed lesions on susceptible cultivars after 3 days. Optimum fungal growth is from 24 to 29
C (75 to 85
F). Leaf spots may show up 4–5 days after infection. The fungus can survive on infected corms or in soil for at least 3 years. Management • Cultural – Corms should be inspected for lesions and discarded. After harvest, infected leaves should be removed from the field when practical by deep plowing or raking and burning leaves. All infected corms should be destroyed. • Fungicides – Consult local recommendation guides for approved fungicides. Sprays are most effective when applied preventively during warm, wet weather when the spores are actively being dispersed. Fungicides known to be effective as preventative contact sprays are chlorothalonil, mancozeb, and benzimidazoles. Magie (1948) reported that mancozeb was superior to other ethylene-bis-dithiocarbamates. • Resistance – Many cultivars listed above have moderate resistance, but Picardy is very susceptible. No cultivar is immune but a wide range of reactions are known to occur (Torres et al. 2013a). Growers should combine resistance to augment other strategies for suppressing the disease (Singh et al. 2006)

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2.3

Diseases of Gladiolus

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Fusarium Yellows and Fusarium Corm Rot (Fusarium oxysporum f. sp. gladioli)

Geographic occurrence and impact. Fusarium corm rot is the most common fungal disease on gladiolus and can be one of the most destructive (Nelson et al. 1981). The disease occurs wherever gladiolus is grown and can be lethal on certain cultivars. Characteristic symptoms of corm rot were first reported in 1912, but it took another 16 years before the disease and the pathogen were formerly described (Massey 1928). The pathogen is the fungus Fusarium oxysporum f. sp. gladioli, which inhabits many soils as different biotypes. As a result, the disease occurs wherever gladiolus is grown, in part, due to resident populations of the pathogen and due to latent corm infections that are frequently present (Nelson et al. 1981). About 30% annual losses have been estimated in Germany (Bruhn 1955), whereas 60–80% annual losses were noted in India (Protsenko 1958 as cited by Chandel and Deepika 2010). Symptoms/signs. Due to the wide diversity in the pathogen populations, symptoms can be diverse and may include stunting and chlorosis (Fig. 2) and, in severe cases, wilt and death (Fig. 3). Along with the rotting of corms (Fig. 4), the disease can cause vascular discoloration. The corm rot is usually a dry corky rot that stays localized until the entire corm becomes infected. Biology and epidemiology. The most common means for long-distance dissemination is via latent infections on the corms and cormels. However, once the pathogen is established in the soil, the fungus survives well as persistent chlamydospores and as mycelium on plant debris. The fungus is very adept at surviving in the absence of gladiolus for many years. The fungus can also endophytically colonize the roots of non-symptomatic plants. New infections result from hyphae that emerge from soil inoculum consisting primarily of chlamydospores or mycelium. The fungus invades roots intracellularly causing root rot as it advances toward the corm. The corm may

Fig. 2 Fusarium corm rot of Gladiolus showing yellowing and stunting (Wade Elmer # 2017. All Rights Reserved.)

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Fig. 3 Fusarium corm rot of Gladiolus showing wilt and death (Wade Elmer # 2017. All Rights Reserved.)

Fig. 4 Fusarium corm rot of Gladiolus (Wade Elmer # 2017. All Rights Reserved.)

be able to compartmentalize invasion, or the infection continues becoming systemic in the vascular tissues. It is not clear whether the resistance of the host or virulence of the pathogenic biotype of F. oxysporum f. sp. gladioli governs whether vascular invasion occurs. Disease incidence is a linear function of soil inoculum and soil temperatures above 25
C (77
F) (Chandel 2004; Sharma and Tripathi 2008). Management • Cultural – While other cultural strategies are limited in their efficacy, sanitation should be highlighted as the most significant approach. Hot water treatment of

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corms and cormels (30 min at 57
C/135
F) followed by biological and inorganic amendments at planting can be effective (Sharma and Tripathi 2008), but temperature baths should be accurately monitored to provide the best disinfestation and to prevent heat damage (Chandel 2004; Magie 1971). Gladiolus requires a well-drained soil with a pH of about 6.5 since acidity promotes Fusarium corm rot. Soil tests should routinely be performed and soil limed appropriately to achieve a soil pH of around 6.5–7.0. Although numerous papers on other plant have shown that Fusarium suppression occurs with nitrate fertilization (Elmer 2012; Woltz 1958), excessive nitrogen can be deleterious and result in more disease (Ashour and Gamal 1966). Fertilization with nitrate-N should be used instead of ammonium-N, which lowers soil pH and tends to promote Fusarium wilt in other crops like tomato (Woltz and Jones 1973). McClellan and Stuart (1947) found organic N promoted disease compared to inorganic N. Addition of increasing amounts of calcium superphosphate decreased corm rot (Ashour and Gamal 1966). A survey of 18 fields in Pakistan found higher levels of disease were associated with low soil potassium, manganese, and zinc indicating the importance of micronutrient availability (Riaz et al. 2009). • Fungicides – Ram et al. (2004) found that soaking corms in acidified solutions of benomyl for 60 min gave the best protection against corm rot. However, others have found that benzimidazoles were not effective probably due to tolerant strains being transported on the corms (Magie 1974). Mishra et al. (2000) found exposing corms to combinations of Trichoderma virens and carboxin provides superior results in suppressing corm rot. Ramos-Garcia et al. (2009) reported that Biorend ® (a biodegradable polymer) applied at 1.5% accelerated corm emergence, the number of flowers, and the vase life. Preplant corm treatments have received much attention as they reduce the cost, labor, and environmental concern compared with soil drenches. Elmer (2006) demonstrated that triflumizole and fludioxonil gave season-long suppression when applied as a corm soak. Many fungicides applied singularly or combined with other chemicals and/or biological products have often provided more suppression than when applied individually. • Biological control – Several researchers have found biological products to be effective (Mishra et al. 2000; Mohamed and Gomaa 2000; Riaz et al. 2010), while others found no control (Elmer 2006). However, given the differences of cultivars, inoculum, and field versus greenhouse, the data are too fractionated to make consistent claims for most products. Nevertheless, a consensus has emerged demonstrating that Trichoderma spp. show greater efficacy than any other biocontrol product. Other reports have shown disease reduction by rhizobacteria and nonpathogenic strains of Fusarium. Talc-based formulations that combined two plant growth-promoting rhizobacterial strains (Bacillus atrophaeus and Burkholderia cepacia) reduced vascular wilt by 48.6% and corm rot incidences by 46.1%, respectively, when compared to the non-treated control (Shanmugam et al. 2011). These treatments were comparable to the fungicide carbendazim that provided a 51.5% reduction in vascular wilt and a 47.1% reduction in corm rot incidence. An increase in the number of spikes (58.3%) and corms (27.4%) were

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also noted with this treatment when compared to control. Magie (1980) conducted field studies where corms were treated with certain isolates of F. subglutinans Snyd. and Hans and F. solani and found disease protection was equal to that obtained with benomyl as a corm dip treatment. Biologicals should be used preventatively. • Resistance – The management of Fusarium corm rot has been difficult. Although partial resistance has been identified in some lines (Straathof et al. 1998), the wide diversity of biotypes of the pathogen and the lack of persistent screening for resistance have hindered efforts to identify highly resistant cultivars. Growers are advised to check with current information regarding resistance/tolerance of cultivars in their regions.

2.4

Gladiolus Rust (Uromyces transversalis Thum)

Geographic occurrence and impact. Gladiolus rust is a disease of quarantine significance in the USA since it was first detected in Florida in 2006 (Schubert et al. 2007). The disease apparently evolved along with the host in Southern Africa. The fungus spread to Southern Europe in the 1960s and reached England in 1996. Since then, major eradication efforts have been initiated to slow the spread of gladiolus rust in the USA. Reports of gladiolus rust have been made from California (Blomquist et al. 2007), South and Central America (Rodríguez-Alvarado et al. 2006; Valencia-Botín et al. 2013), and Australia (Beilharz et al. 2001). Symptoms/signs. Symptoms are very distinct. The orange uredinia form transverse lines across gladiolus foliage. Under severe inoculum pressure, lesions can be observed on flower spikes (Fig. 5). Biology and epidemiology. U. transversalis is an autoecious rust that has no alternate host. It produces only two spore types, urediniospores and teliospores. It infects Gladiolus spp. and Anomatheca, Crocosmia, Melasphaerula, Tritonia, and Watsonia spp. The rust spores are airborne (urediniospores) and infect and cause

Fig. 5 Gladiolus rust (Tom Gulya. USDA.)

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profuse sporulation on the leaf surface. Severely infected plants cannot photosynthesize sufficiently. They remain stunted and will not produce flower spikes. The fungus overwinters as teliospores on the foliage. Management • Sanitation – Currently, gladiolus rust is a quarantinable pathogen in the USA. Strict rust management guidelines and fallow host-free periods need to follow the appearance of the disease. • Fungicides – In Mexico, where the disease was more common, fungicide trials found that the triazole fungicides, cyproconazole, difenoconazole, epoxiconazole, myclobutanil, propiconazole, and tebuconazole gave excellent control.

2.5

Stromatinia Rot [Stromatinia gladioli (Drayt.) Whet.]

Geographic occurrence and impact. The disease was recognized in 1883 in England and known to occur wherever gladiolus is grown. The fungus was first described as Sclerotinia gladioli (Hawker et al. 1944) but was changed to Stromatinia gladioli. The disease was reported along the USA Gulf Coast (Magie 1954; Drayton 1934). Symptoms/signs. Symptoms appear as a dry rot of the corm (Fig. 6). Individual lesions may appear as superficial small, round, reddish brown spots that can have brown to black centers. Lesions more frequently appear at the point of the husk attachment. The actual husk can appear dark colored, brittle, and shredded. One diagnostic characteristic of Stromatinia rot is the presence of black sclerotia in rotted cortical tissue of the roots (McRitchie and Leahy 1988). Biology and epidemiology. The fungus overwinters in the soil in the form of sclerotia, which can last in the absence of a host for 10 years (Nelson 1948). The disease is more severe when cool, wet conditions prevail. Young corms (1–2 years) Fig. 6 Stromatinia rot of Gladiolus (R.D. Milholland # 2017. All Rights Reserved.)

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tend to be more susceptible than older ones. Wounds are not required for infection (Hawker et al. 1944). The disease spreads faster when plants are planted in close proximity. Inoculation of the top of an old corm is more effective than that of the side or base. The percentage of diseased new corms produced from infected parent corms varies from 0% to 100% according to soil conditions. Disease is favored by wet soil. Management • Cultural practices – Crop rotation should be practiced at 3–4-year intervals to reduce soil densities of the sclerotia. Any volunteer gladiolus plants should be rogued as they can serve as sources of inoculum. Since cool, wet soil favors the disease, delaying the time of planting and improving drainage can lessen the loss due to Stromatinia rot. Removal of “husks” (leaf bases) increases the susceptibility of corms planted in contaminated soil, but the removal of husks is not an effective control. Dehusking did not increase the number of diseased young corms in infected stock planted in new soil. No variety was strikingly more resistant than the rest. • Fungicides – Corm and soil treatment with fungicides should be done where there is a history of the disease. Good control was obtained when corms of a diseased stock were treated with mercuric chloride, mercurous chloride (calomel), or the proprietary mercury fungicide Aretan. Calomel gave the most consistent results. Various other treatments were tested in small-scale trials with some success. Red copper oxide, formalin, and the proprietary mercury compound, Ceresan, were harmful. Sterilization of contaminated soil with formalin, mercuric chloride, Aretan, or Uspulun gave good but not complete control. Formalin was the least effective.

3

Bacterial and Phytoplasma Diseases

3.1

Bacterial Scab Burkholderia gladioli pv. gladioli (Formerly Pseudomonas gladioli pv. gladioli)

Geographic occurrence and impact. Bacteria scab is not a major problem for gladiolus growers. Unlike the fungal corm rot diseases, bacterial scab does not kill the affected plants or affect the flower spikes. Symptoms/signs. The lesions are very dark and appear on the surface of the corm. Scab lesions are sunken with a concentric border. Most lesions develop a shiny black layer over the surface of the lesion. Bacterial scab can be differentiated from Stromatinia by where the lesions tend to appear. Bacterial scab lesions appear more frequently at the base of corm, whereas Stromatinia lesions appear more commonly on the top of the corm (Forsberg 1965).

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Biology and epidemiology. The bacteria are ubiquitous and can be introduced on previously infected corms or through wounds made from handling or insect feeding. The bacterium also infects a number of other horticultural crops including lisianthus. Management. Inspect corms for lesions and discard infected corms. Avoid planting in areas where the disease has appeared. Efforts to suppress bulb mites will also reduce infection sites.

3.2

Aster Yellows (“Candidatus Phytoplasma asteris”)

Geographic occurrence and impact. Aster yellows has been reported in Gladiolus in the USA, and it infects a wide range of plant species (Treeful and Ash 2000). Symptoms/signs. Gladiolus plants mature early, have an arrested root development, and have small or few corms (Fig. 7) during the season when they are first infected. The next year, multiple shoots that are thin and weak develop from the

Fig. 7 Phytoplasma-infected Gladiolus cultivars showing (a, b) stunting and spike twisting; (c, d) virescence; (e, f) malformation and reduction of floret size; and (h) small corms with poor root systems as compared to (g, i) healthy plants collected from the gardens of NBRI, Lucknow, India (Raj et al. 2011) – http://link.springer.com/article/10.1007/s10327-010-0259-9

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corm giving the appearance of their nickname “grassy top” (Koike et al. 2007) (Fig. 7). The flower spike grows in a spiral pattern (Fig. 7). Biology and epidemiology. This disease is caused by phytoplasmas formerly known as mycoplasma-like organisms and not by a virus. Phytoplasmas are microscopic organisms similar in size and composition to bacteria (Treeful and Ash 2000). Transmission is by Macrosteles sexnotatus leafhoppers (Koike et al. 2007). An infected leafhopper will remain infected throughout its life and will inject phytoplasma into the phloem cells of a plant when feeding (Missouri Botanical Garden). Plants will exhibit symptoms in 10–40 days. Aster yellows has been reported to occur in many floral crops and in a number of weed species (Convolvulus, Capsella, and Cirsium) (Vicchi and Bellardi 1988). Aster yellows spreads more readily in cool, wet summers because phytoplasmas and leafhoppers do not thrive as well in dry, hot weather (Missouri Botanical Garden). Massive spread of aster yellows is rare (Stein 1995). Management. Infected plants should be destroyed (Koike et al. 2007). Weeds, particularly dandelions and plantains, should be controlled in the area where Gladiolus is being grown. Monitor plants for leafhoppers. Fine mesh fabrics and strips of aluminum foil between rows of plants can be used to keep leafhoppers away from the plants (Missouri Botanical Garden).

4

Viral Diseases

Twenty viruses have been reported to infect Gladiolus (Stein 1995; Asjes 1997). The predominant viruses that infect Gladiolus are Bean yellow mosaic virus (BYMV) and Cucumber mosaic virus (CMV). Other viruses that have been reported in Gladiolus are listed in Table 1. A number of companies offer testing services and/or ELISA kits to detect various gladiolus viruses. Nine Gladiolus viruses can be detected immunologically by Agdia, Inc., Elkhart, IN, USA, in their “Gladiolus Screen” (www.agdia.com/testing-services/Gladiolus.cfm). Bioreba AG, Reinach, Switzerland, can test for the presence of six gladiolus viruses by ELISA (http:// www.bioreba.ch/files/Tecnical_Info/TS_Broschuere_2011_e_CHF.pdf). Neogen Europe, Ltd., Auchincruive, Scotland, UK, manufactures ELISA kits which can detect a number of gladiolus viruses. The viruses in Gladiolus differ for each country, but two viruses, BYMV and CMV, are by far the most prevalent viruses found worldwide in Gladiolus.

4.1

Bean yellow mosaic virus

Geographic occurrence and impact. Almost all Gladiolus plants are infected with Bean yellow mosaic virus (BYMV) as reported for plants grown in the USA (Zettler and Abo El-nil 1977), Ontario Canada (Berkeley 1953), the Netherlands

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Table 1 Viruses reported in Gladiolus other than BYMV and CMV Virus Arabis mosaic virus Bearded iris mosaic virus (=Iris severe mosaic virus) Broad bean wilt virus Clover yellow vein virus Cowpea mosaic virus Cycas necrotic stunt virus Impatiens necrotic spot virus

Occurrence in Gladiolus Italy, India

Transmission Xiphinema (nematode)

Symptoms ND

USA, Netherlands

Aphids

ND

Korea

Aphids

Symptomless

Park et al. 1998

Korea

Aphids

ND

Park et al. 2002

USA

Beetles

ND

Japan

Nematodes and seed Thrips

ND ND

Brierley and Smith 1962 Hanada et al. 2006 Marchoux et al. 1992; Ghotbi et al. 2005; Louro 1996 Stein 1995; Asjes 1997 Kaur et al. 2011

France, Iran, Portugal

References Bellardi et al. 1986; Katoch et al. 2003 Brierley and Smith 1948; Brunt et al. 1988

Narcissus latent virus Ornithogalum mosaic virus

Netherlands

Aphids

ND

India, Netherlands

Aphids

Plum pox virus Soybean mosaic virus Strawberry latent ringspot virus Tobacco mosaic virus

Germany USA

Aphids Aphids

Foliar mosaic, chlorotic spots, floral deformation ND ND

Italy

Xiphinema (nematode)

ND

Bellardi et al. 1984

Japan, Netherlands, Canada Russia, Lithuania

Mechanical

ND

Olpidium brassicae (fungus)

ND

Netherlands, Israel, Egypt, Poland, India, Korea, Japan, Czech Republic, Lithuania

Tools used during planting, low rate of transmission in Gladiolus seeds, nematodes of the family Trichodoridae

Notched leaf blades, severely distorted plant, chlorotic, brown or dead stripes and spots on leaves, crumpled tissue between veins, stem and

Berkeley 1951; Fukumoto et al. 1982 Stein 1995; Navalinskiene and Samiutiene 2010 Navalinskiene and Samuitiene 2001; Univ. Illinois Extension 1983; Stein 1995; Asjes 1997; Yamaji et al. 1998; Park et al. 2002;

Tobacco necrosis virus

Tobacco rattle virus

Kroll 1978 Stein 1995

(continued)

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Table 1 (continued) Virus

Occurrence in Gladiolus

Transmission

Symptoms

References

buds may stop growing

Katoch et al. 2004; Duraisamy and Pokorný 2009 Brierley 1952; Brierley and Smith 1960; Berkeley 1953; Randles and Francki 1965; Fukumoto et al. 1982; Kaniran and Izadpanah 1982; Univ. Illinois Extension 1983; Katoch et al. 2003; Navalinskiene and Samuitiene 2010 Bellardi et al. 1987

Tobacco ringspot virus

Japan, India, Iran, USA, Canada, Australia, Lithuania

Cutting tools (low rate of cross contamination), Xiphinema americanum (nematode), thrips, mites, grasshoppers, tobacco flea, and beetles

Typically symptomless in gladiolus, sometimes necrotic or chlorotic ringspot patterns on leaves

Tobacco streak virus

Italy

Pollen, thrips

Tomato black ring virus

Poland, Italy (not reported outside of Europe) USA, Iran

Longidorus (nematode)

Stunted flowers, necrotic spots on leaves Symptomless

Tomato ringspot nepovirus

Tomato spotted wilt tospovirus Tomato aspermy virus

USA, France, S. Australia, Portugal India

Xiphinema americana (nematode), low rate by Gladiolus seeds

Thrips

ND

Stunted growth, abnormally short flower spikes with only 4–5 florets. Leaves smaller, stiffer, and more erect than on healthy plants Chlorosis with degeneration Severe chlorosis along midrib

Kaminska 1978; Bellardi and Pisi 1985a, b; Stein 1995 Bozarth and Corbett 1958; Univ. Illinois Extension 1983; Stein 1995; Ghotbi et al. 2005

Lee et al. 1979; Stein 1995; Louro 1996 Raj et al. 2011

(Brunt 1970; Nagel et al. 1983), Israel (Stein 1995), Argentina (Arneodo et al. 2005), Iran (Kaniran and Izadpanah 1982; Dorrigiv et al. 2013), Italy (Bellardi and Pisi 1985a), Japan (Wada et al. 2000), Korea (Park et al. 2002), Russia (Gnutova et al. 1989), the Czech Republic (Duraisamy and Pokorný 2009),

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Belarus (Voinilo and Burgansky 1999), Lithuania (Navalinskiene and Samuitiene 2001), New Zealand (Fry 1953), and India (Katoch et al. 2002). Bean yellow mosaic virus is also known as the Pea mosaic virus. Symptoms/signs. BYMV-infected Gladiolus plants are typically symptomless or have mild symptoms. It is difficult to attribute viral symptoms to a specific virus because Gladiolus is generally infected with BYMV and a second virus (Stein 1995). The symptoms of BYMV have been described as faint, inconspicuous pencil-stripe break patterns that are lighter in color than the normal flower although it may be conspicuous on specific cultivars (Univ. Illinois Extension 1983). The leaves and flower stems may have light or dark green mottling seen in the early summer (Univ. Illinois Extension 1983). Symptoms will be affected by environmental conditions (Asjes 1997). Dry, high-temperature conditions will mask the symptoms (Univ. of Illinois Extension 1983). Katoch et al. (2002) examined 32 cultivars of Gladiolus and reported a BYMV incidence of 0–100% based on visual symptoms, but almost 100% were found to have BYMV as indicated by ELISA. Biology and epidemiology. BYMV is of the genus Potyvirus and belongs to the Potyviridae family, which is the largest and most economically important group of viruses as it infects many species of vegetables, forage, fruit, ornamental, and field crops (Jordan and Hammond 1991). BYMV infects many legumes, and the virus overwinters in perennial legumes such as alfalfa, clovers, or vetch. Hampton et al. (2005) verified the non-circulative (nonpersistent) transmission of BYMV from clover, Trifolium repens, to an adjacent field of snap beans, Phaseolus vulgaris, by four aphid species (Myzus persicae, Acyrthosiphon pisum, Aphis fabae, and Nearctaphis bakeri). Over 20 species of aphids serve as vectors for BYMV transmission. Transmission of BYMV through bean seeds of legumes has been reported (Latham et al. 2000). BYMV particles are flexuous and filamentous (750 nm in length and 15 nm in diameter) (Guyatt et al. 1996). They consist of single-stranded, positive sense RNA. Various strains of BYMV have been identified serologically using both polyclonal and monoclonal antisera (Nagel et al. 1983; Jordan and Hammond 1991). Virus is readily detectable in leaves and flowers by ELISA (Stein 1995). The low level of BYMV in corms requires that the corm be cut, and then virus can be detected by ELISA in the callus tissue that forms at the wounded areas (Stein 1995). Immunological detection of BYMV in corms of Gladiolus has been possible using a tissue blotting technique (Lin et al. 1990). Polymerase chain reaction (PCR) amplification of BYMV RNA has also been used to detect BYMV in corms (Rosner et al. 1992). Another method to increase the sensitivity of detection (105 higher than when using ELISA) was achieved by hybridizing PCR products with a radioactive 32P-labeled viral riboprobe (Vunsh et al. 1991). Management. Because aphid species spread BYMV, growing Gladiolus plants under fine mesh nets was found to greatly reduce virus infection (Stein 1995). BYMV as well as CMV, but not Tobacco ringspot virus, were found to be

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transmitted on tools used when harvesting flowers and corms of Gladiolus (Brierley 1962). Tools should be disinfested with 70% alcohol between cuts to prevent cross contamination (Univ. Illinois Extension 1983). Logan and Zettler (1985) developed an efficient tissue culture system that had the potential to produce 50,000 gladiolus plants from a single shoot tip during a 30-week period. Gladiolus plants free from BYMV can be produced using meristem culture (Stein 1995). Propagating with the apical meristem of a corm grown under sterile conditions in vitro can eliminate 60–100% of the virus. Removal of three viruses from Gladiolus was achieved by performing meristem culture twice, adding ribavirin (5–20 ppm) to the culture medium, and treating with high (38–40
C) temperature (Li et al. 2003). Regeneration of plants from callus was reported to be more effective than culturing shoot tips from corms for virus elimination (Park et al. 2002). Callus induced from cormels infected with BYMV and CMV was cultured on a medium containing ribavirin (virazole, 1-β-D-ribofuranosyl-a, 2, 4-triazole-3-carboxamide), and after 6–8 weeks, plants were regenerated and certified as virus-free (Singh and Dubey 2007). Virusfree plants must be grown in insect-proof conditions if the Gladiolus plants are to remain free of virus or else 80–100% of the crop will become infected with BYMV within one growing season (Stein 1995; Asjes 1997). It is recommended (Univ. Illinois Extension 1983; Moran 1996) that virus-free plants be grown at least 1 km from virus-infected Gladiolus plants and legumes, that effective insecticides be used to control aphids on the crop plants and on corms during their storage, and that plants showing viral symptoms be rogued and destroyed by burning or composting. Weeds in the vicinity of Gladiolus plants should be eradicated as they may serve as reservoirs of viruses and insects that can be vectors for viruses (Univ. Illinois Extension 1983). Aluminum strips placed between rows or Gladiolus plants can be used to repel aphids and will also decrease the number of weeds present.

4.2

Cucumber mosaic virus

Geographic occurrence and impact. Cucumber mosaic virus (CMV) is in the genus Cucumovirus and is a member of the family Bromoviridae. CMV consists of three spherical particles that are each 28 nm in diameter. Its genome consists of three single-stranded messenger RNA molecules in the sense orientation. There are two subgroups, I and II, of CMV that are distinguished either by their serological response to antibodies or by their nucleotide sequences (Chen 2003; Zitter and Murphy 2009). The CMV that infects Gladiolus plants belongs to subgroup I and has the highest nucleotide sequence homology based upon its coat protein to the CMV strain Fny (Chen 2003; Dubey et al. 2010). CMV infects 1,200 plant species that include economically important vegetable crops such as cucumbers, squash, and melons and most floral crops. CMV occurs throughout the world and has been reported in Gladiolus in the USA (Brierley 1952; McWhorter 1957); Iran (Dorrigiv et al. 2013); Ontario, Canada (Berkeley 1953); Tasmania (Wade 1984); Israel (Stein 1995); India (Dubey et al. 2010); the Czech Republic (Duraisamy and Pokorný 2009); and Lithuania (Navalinskiene and Samuitiene 2001).

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Fig. 8 Symptoms of viral infection in Gladiolus are (a–c) color break on flower petals. In b the left spike is a healthy spike, and the right spike shows color break of petals indicative of virus infection. (d) The two inner plants are stunted in their growth as compared to the two outer, healthy plants. (e) Viral symptoms on leaves (streaking or (f) blotching. (g) Corms infected with CMV are deformed (Kathy Kamo. USDA.)

Biology and epidemiology. The worldwide occurrence of CMV in Gladiolus can be attributed to the international trade of corms and to the virus’ wide host range. When only 3% of the corms are initially infected with CMV, after one growing

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season, 60% of the plants may become infected (Stein 1995). CMV transmission occurs by many aphid species in a non-circulative (nonpersistent) manner. Most commercial cultivars of Gladiolus are susceptible to CMV. The cultivars Peter Pears, Spic and Span, and White Friendship are readily infected by CMV whereas Trader Horn and Eurovision are more resistant (Aly et al. 1986). Symptoms/signs. Generally, Gladiolus plants with CMVoften appear symptomless until flowering when there may be spotting on petals; obvious color break that is more readily visible on purple, lilac, pink, and red flowers than on white or yellow flowers (Fig. 8a–c); and a reduced size of the floret accompanied by a change in the color of the petals, thickened petals, twisted and deformed florets, and flowers that will not fully open. Flowers may have crinkled petals, open slowly, imperfectly, and fade early (Univ. Illinois Extension 1983). Flower bracts can be yellow and wither early in the season. Other symptoms can be deformity of the whole plant. Stunting of infected plants often occurs (Fig. 8d) with severely affected leaves that have gray or yellow-green streaking or spots that are either gray, yellow, brown, or reddish (Fig. 8e, f). Young leaves may have a mosaic or chlorophyll break. Some cultivars have pitting and discoloration of the corms or corms that are deformed and have a warty appearance (Univ. of Illinois Extension 1983; Stein 1995). Management. If possible, it is recommended that CMV-tested, certified corms and resistant cultivars be planted. A number of diagnostic companies manufacture rapid detection ELISA kits for CMV. The use of fine mesh nets coated with mineral oil spray or by directly spraying mineral oil on the plants in the absence of nets has been found to help with controlling spread of CMV (Aly et al. 1986). Controlling the aphid population and rogueing infected plants during flowering when symptoms are most obvious on the flowers also help in the field to prevent a large outbreak of CMV. It is advised that Gladiolus should not be grown near cucumbers, melons, and tomatoes because of the virus transmission that can occur between CMV-infected crops (Univ. Illinois Extension 1983). The strategies to manage CMV are the same as for BYMV (rogueing, aphid control, eliminate weeds).

References Aly R, Stein A, Levy S, Raccah B, Loebenstein G (1986) Spread and control of Cucumber Mosaic Virus in gladiolus. Phytoparasitica 14:205–217 Arneodo JD, Breuil SD, Lenardon SL, Conci VC, Conci LR (2005) Detection of Bean yellow mosaic virus and Cucumber mosaic virus infecting gladiolus in Argentina. Agriscientia 22:87–89 Ashour WE, Gamal El-Din IF (1966) Effect of fertilization on corm diseases of Gladiolus in the United Arab Republic. Phytopathol Mediterr 115–121 Asjes CJ (1997) Incidence and control of viruses in Gladiolus in the Netherlands. Acta Hortic 430:699–708

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Beilharz V, Parbery DG, Pascoe IG (2001) Gladiolus rust (caused by Uromyces transversalis) in eastern Australia. Australas Plant Pathol 30(3):267–270 Bellardi MG, Pisi A (1985a) Survey of gladiolus viruses in Italy. Revista della Ortoflorofrutticoltura Italiana 69:133–144 Bellardi MG, Pisi A (1985b) Identification of three isolates of tomato black ring virus (TBRV) from gladiolus. Revista della Ortoflorofrutticoltura Italiana 69:299–309 Bellardi MG, Canova A, Tacconi R, Gelli C (1984) Flower colour-break of Gladiolus associated with strawberry latent ringspot virus. Atti Giornate Fitopatologiche Sorrento 26–29:303–312 Bellardi MG, Canova A, Gelli C (1986) Comparative studies on gladiolus isolates of Arabis mosaic virus (ArMV). Phytopathol Mediterr 26:73–80 Bellardi MG, Vicchi V, Gelli C (1987) Stunting of gladiolus flower spike associated with tobacco streak virus (TSV). Phytopathol Mediterr 26:73–80 Benschop M, Kamenetsky R, Le Nard M, Okubo H, De Hertogh A (2010) The global flower bulb industry: production, utilization, research. Hortic Rev 36:1–115 Berkeley GH (1951) Gladiolus viruses. Phytopathology 41:3–4 Berkeley GH (1953) Some viruses affecting Gladiolus. Phytopathology 43:111–115 Blomquist CL, Thomas SL, McKemy JM, Nolan PA, Luque-Williams M (2007) First report of Uromyces transversalis, causal agent of gladiolus rust, in San Diego County, California. Plant Dis 91(9):1202 Boerema GH, Hamers ME (1989) Check-list for scientific names of common parasitic fungi. Series 3b: Fungi on bulbs: Amaryllidaceae and Iridaceae. Netherlands J Plant Pathol 95(3):1–29 Bozarth RF, Corbett MK (1958) Tomato ringspot virus associated with stunt or stub head disease of Gladiolus in Florida. Plant Dis Rep 42:217–221 Brierley P (1952) Evidence on the significance of Cucumber mosaic and Tobacco ringspot viruses in Gladiolus. Gladiolus Mag 16(28–29):36–37 Brierley P (1962) Transmission of some Gladiolus viruses on tools in harvesting flowers and corms. Plant Dis Rep 46:505 Brierley P, Smith FF (1948) Two additional mosaic diseases of iris. Phytopathology 38:574–575 Brierley P, Smith FF (1960) Transmission of Tobacco ringspot virus on tools during harvesting flowers and corms of Gladiolus. Plant Dis Rep 44:463–464 Brierley P, Smith FF (1962) Three Cowpea mosaic viruses from Gladiolus. Plant Dis Rep 46:335–337 Bruhn C (1955) Untersuchungen a uber die Fusarium Krankeit der gladiolen. Phytopathol Z 25:31–38 Brunt AA (1970) Gladiolus. Annual Report Glasshouse Crops Research Institute 1970:150 Brunt AA, Derks AFLM, Barnett OW (1988) Iris severe mosaic virus. CMI/AAB Descriptions Plant Viruses 388:4 Chandel S (2004) Management of Corm Rot and Wilt of Gladiolus Combining Chemical and Non Chemical Approaches Annual Progress Report Chandel S, Deepika R (2010) Recent advances in management and control of Fusarium yellows in Gladiolus species. J Fruit Orn Plant Res 18(2):361–380 Chen YK (2003) Occurrence of Cucumber Mosaic Virus in Ornamental Plants and Perspectives of Transgenic Control. PhD thesis, van Wageningen Universiteit, p 151 Dimock AW (1940) Epiphytotic of Botrytis blight on Gladiolus in Florida. Plant Dis Rep 24 (8):159–161 Dorrigiv R, Jafarpour B, Rastegar MF (2013) Detection of some virus pathogens of gladiolus in Iran. Int J Agric Crop Sci 5:1653–1658 Drayton FL (1934) The gladiolus dry rot caused by Sclerotinia gladioli (Massey) n. comb. Phytopathology 24:397–404 Dubey VK, Aminuddin, Singh VP (2010) Molecular characterization of Cucumber mosaic virus infecting Gladiolus, revealing its phylogeny distinct from the Indian isolate and alike the Fny strain of CMV. Virus Genes 41:126–134

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Duraisamy GA, Pokorný R (2009) Survey of virus pathogens in gladiolus, iris, and tulip in the Czech Republic. Acta Universitatis Agriculturae Silviculturae Mendelianae Brunensis 57:79–86 Elmer WH (2006) Efficacy of preplant treatments of gladiolus corms with combinations of acibenzolar-S-methyl and biological or chemical fungicides for suppression of fusarium corm rot [Fusarium oxysporum f. sp. gladioli]. Can J Plant Pathol 28(4):609–614 Elmer WH (2012) Cultural management of Fusarium. In: Garibaldi A, Katan J, Lodovica Gullino M (eds) Fusarium wilts of greenhouse vegetable and ornamental crops. APS Press, ST. Paul, pp 67–84 Forsberg JL (1957) A vascular form of the Curvularia disease of gladiolus. Phytopathology (abstr) 47:12 Forsberg JL (1965) The relationship of Pseudomonas marginata, Stromatinia gladioli, bulb mites, and chemical soil treatments to the occurrence and control of scab and Stromatinia rot of gladiolus. Phytopathology 55:1058 Fry PR (1953) Two virus diseases of gladiolus. N Z J Sci Technol A34:460–464 Fukumoto F, Ito Y, Tochihara H (1982) Viruses isolated from gladiolus in Japan. Ann Phytopathol Soc Jpn 48:68–71 Ghotbi T, Sharaeen N, Winter S (2005) Occurrence of tospoviruses in ornamental and weed species in Markazi and Tehran provinces in Iran. Plant Dis 89:425–429 Gnutova RV, Kakareka NN, Tolkach VF, Chuyan AX, Sibiryakova II, Rubleva NV, Krylov AV (1989) Some properties of a bean yellow mosaic-virus, identified in the south of the Far East. Izvestiya Akademii Nauk Sssr Seriya Biologicheskaya 3:442–449 Goldblatt P, Rodriguez A, Powell MP, Davies TJ, Manning JC, Van der Bank M, Savolainen V (2008) Iridaceae “Out of Australasia”? Phylogeny, biogeography, and divergence time based on plastid DNA sequences. Syst Bot 33:495–508 Guyatt KJ, Proll DF, Menssen A, Davidson AD (1996) The complete nucleotide sequence of bean yellow mosaic potyvirus RNA. Arch Virol 141:1231–1246 Hampton RO, Jensen A, Hagel GT (2005) Attributes of bean yellow mosaic potyvirus transmission from clover to snap beans by four species of aphids (Homoptera: Aphididae). J Econ Entomol 98:1816–1823 Hanada K, Tanaka Y, Iwanami T, Fukumoto F, Kusunoki M, Kameya-Iwaki M (2006) Cycas necrotic stunt virus isolated from gladiolus plants in Japan. J Gen Plant Pathol 72:383–386 Hartline C (2015) Gladiolus history, Chapter 18 in. how to grow glorious gladiolus. North American Gladiolus Council. http://www.gladworld.org/Chapter%20Eighteen.pdf Hawker LE, Bray RJ, Burrows TW (1944) Diseases of the gladiolus: II. Experiments on dry rot disease caused by Sclerotinia gladioli Drayt. Ann Appl Biol 31:211–218 Jordan R, Hammond J (1991) Comparison and differentiation of potyvirus isolates and identification of strain-, virus-, subgroup-specific and potyvirus group-common epitopes using monoclonal antibodies. J Gen Virol 72:25–36 Kaminska M (1978) Some properties of isolates of tomato black ring virus. Zesz Probl Postep Nauk Rol 214:109–117 Kaniran R, Izadpanah K (1982) Isolation and identification of bean yellow mosaic and tobacco ringspot viruses from gladiolus in Shiraz. Iranian J Plant Pathol 17:1–10 Katoch M, Ram R, Zaidi AA, Garg ID (2002) Status of BYMVon Gladiolus. Crop Prot 21:861–865 Katoch M, Ram R, Zaidi AA (2003) First report of Tobacco ringspot virus occurring in gladiolus in India. Plant Pathol 52:789 Katoch M, Ram R, Zaidi AA (2004) First report of Tobacco rattle virus occurring in gladiolus in India. Plant Pathol 53:236 Kaur C, Raj SK, Snehi SK, Goel AK, Roy RK (2011) Natural occurrence of Ornithogalum mosaic virus newly reported on gladiolus in India. New Dis Rep 24:2 Koike ST, Wilen CA, Raabe RD, McCain AH, Grebus ME (2007) UC IPM pest management guidelines: floriculture and ornamental nurseries, US ANR Publication 3392. http://www.ipm. ucdavis.edu/PMG/r280111611.html

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Kroll J (1978) On infection of gladiolus (Gladiolus spp.) with plum pox virus. Arch Phytopathol Pflanzenschutz 14:415–416 Latham LJ, Jones RAC, McKirdy SJ (2000) Cucumber mosaic cucumovirus infection of coolseason crop, annual pasture, and forage legumes: susceptibility, sensitivity, and seed transmission. Aust J Agric Res 52:683–697 Lee TC, Francki RIB, Hatta T (1979) A serious disease of gladiolus in Australia caused by tomato spotted wilt virus. Plant Dis Rep 63:343–348 Li C, Guo T, Jiao P, Zhang Y, Du L (2003) Studies on virus-free and rapid reproduction in cultural plant of gladiolus and its effect. Acta Hort Sinica 30:358–360 Lin NS, Hsu YH, Hsu HT (1990) Immunological detection of plant viruses and a mycoplasmalike organism by direct tissue blotting on nitrocellulose membranes. Phytopathology 80:824–828 Logan AE, Zettler FW (1985) Rapid in vitro propagation of virus-indexed Gladioli. Acta Hortic 164:169–180 Louro D (1996) Detection and identification of tomato spotted wilt virus and Impatiens necrotic spot virus in Portugal. Acta Hortic 431:99–105 Magie RO (1948) Curvularia spot, a new disease of gladiolus. Plant Dis Rep 32:11–13 Magie RO (1954) Stromatinia disease of gladiolus. Florida State Hort Soc Proc 67:313–317 Magie RO (1956) Gladiolus botrytis control. Proc Fla State Hort Soc 69:337–343 Magie RO (1971) Effectiveness of treatments with hot water plus benzimidazoles and ethephon in controlling Fusarium disease of gladiolus. Plant Dis Rep 55:82–85 Magie RO (1974, April). Tolerance of Fusarium oxysporum f. sp. gladioli to benzimidazole fungicides. In: II International Symposium on Flower Bulbs 47, pp 107–112 Magie RO (1980) Fusarium disease of gladioli controlled by inoculation of corms with nonpathogenic Fusarium spp. Proc Fla State Hort Soc 93:172–175 Marchoux G, Gebre-Selassie K, Nono-Wonidim R, Gogzalons P, Berling A, Dufouro, Pivot Y (1992) Epidemiology and variability of tospoviruses in France. Abstracts 6th International Congress of Plant Pathology, Montreal, p 321 Massey LM (1928) Dry rot of Gladiolus corms. Phytopathology 18:519–529 McClellan WD, Stuart NW (1947) The influence of nutrition on Fusarium basal rot of narcissus and on Fusarium yellows of gladiolus. Am J Bot 34:88–93 McRitchie JJ, Leahy RM (1988) Stromatinia dry rot of gladiolus. Plant pathology circular/Florida. Department of Agriculture and Consumer Services. Division of Plant Industry McWhorter FP (1957) A localized occurrence of Cucumber mosaic virus in Gladiolus. Plant Dis Rep 41:141–143 Meerow AW (2012) Taxonomy and phylogeny. In: Kamenetsky R, Okubo H (eds) Ornamental geophytes: from basic science to sustainable production. CRC Press, Boca Raton, pp 17–56 Mendiola-Ela V (1952) The Curvularia disease of gladiolus in the Philippines. Philipp Agric 35:517–533 Mishra PK, Mukhopadhyay AN, Fox RTV (2000) Integrated and biological control of gladiolus corm rot and wilt caused by Fusarium oxysporum f. sp. gladioli. Ann Appl Biol 137:361–364 Missouri Botanical Garden. Aster Yellows. http://www.missouribotanicalgarden.org/gardens-gar dening/your-garden/help-for-the-home Mohamed FG, Gomaa AO (2000) Effect of some bio agents and agricultural chemicals on Fusarium wilt incidence and growth characters of gladiolus plants. Annals Agric Sci 38:883–906 Moran J (1996) Virus diseases of gladiolus. http://agriculture.vic.gov.au/agriculture/pests-diseasesand-weeds/plant-diseases/flowers-and-ornamentals/virus-diseases-of-gladiolus Nagel J, Zettler FW, Hiebert E (1983) Strains of bean yellow mosaic virus compared to clover yellow vein virus in relation to Gladiolus production in Florida. Phytopathology 73:449–454 Navalinskiene M, Samuitiene M (2001) Viral diseases of flower plants 15. Identification of viruses affecting gladiolus (Gladiolus L). Biogeosciences 1:31–35 Navalinskiene M, Samuitiene M (2010) Identification of viral and phytoplasmal diseases affecting gladioli (Gladiolus L.). Acta Biol Univ Daugavp Suppl 2:137–144

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Nelson R (1948) Diseases of gladiolus. Mich State College Agr Exp Sta Spec Bull 350. 63 pp Nelson PE, Horst RK, Woltz SS (1981) Fusarium diseases of flowering bulb crops. In: Nelson PE, Toussoun TA Cook, RJ (eds) Fusarium: diseases, biology and taxonomy. Penn State Press, University Park, pp 121–129 Park I, Kim K, Kyun H, Chang M (1998) The viruses in Gladiolus hybridus cultivated in Korea. 2. Broad bean wilt virus, cucumber mosaic virus and tobacco rattle virus. Korean J Plant Path 14:83–91 Park IS, Choi JD, Goo DH, Kim KW (2002) Elimination of viruses from virus-infected gladiolus plants through cormel tip and callus culture. J Korean Soc Hort Sci 43:531–535 Parmelee JA (1954) Curvularia on gladiolus in Canada. Plant Dis Rep 38:515–517 Parmelee JA (1956) The identification of the Curvularia parasite of Gladiolus. Mycologia 48 (4):558–567, http://doi.org/10.2307/3755336 Protsenko EP (1958) Premature yellowing of gladioli. Bull Centr Bot Gdn Moscow 30:78–84 Raj SK, Kumar S, Verma DK, Snehi SK (2011) First report on molecular detection and identification of Tomato aspermy virus naturally occurring on gladiolus in India. Phytoparasitica 39:303–307 Ram R, Manuja S, Dhyani D, Mukherjee D (2004) Evaluations of fortified fungicide solutions in managing corm rot disease of gladiolus caused by Fusarium oxysporum. Crop Prot 23:783–788 Ramos-García M, Ortega-Centeno S, Hernández-Lauzardo AN, Alia-Tejacal I, Bosquez-Molina E, Bautista-Baños S (2009) Response of gladiolus (Gladiolus spp.) plants after exposure corms to chitosan and hot water treatments. Sci Hortic 121(4):480–484 Randles JW, Francki RIB (1965) Some properties of a tobacco ringspot virus isolate from South Australia. Aust J Biol Sci 18:979 Riaz T, Khan SN, Javaid A (2009) Correlation between soil nutrient status and corm-rot disease of gladiolus in Rawalpindi and Islamabad, Pakistan. Pakistan J Phytopathol 21(2):148–151 Riaz T, Nawaz Khan S, Javaid A (2010) Management of corm-rot disease of gladiolus by plant extracts. Nat Prod Res 24(12):1131–1138 Rodríguez-Alvarado G, Fernández-Pavía SP, Valenzuela-Vázquez M, Loya-Ramírez JG (2006) First report of gladiolus rust caused by Uromyces transversalis in Michoacán, México. Plant Dis 90(5):687–687 Rosner A, Stein A, Levy S (1992) Transcription amplification of polymerase chain reaction products of bean yellow mosaic virus RNA extracted from gladioli corms. Ann Appl Biol 121:269–276a Schubert TS, Leahy RM, Davison DA, Silagyi AJ, Killgore EM (2007) Gladiolus rust caused by Uromyces transversalis makes first necrotic appearance in Florida. Plant Dis 91(9):1202–1202 Sehajpal PK, Singh PJ (2014) Effect of temperature, leaf wetness period, light and darkness on development of Botrytis blight (Botrytis gladiolorum Timm.) of gladiolus (Gladiolus grandiflorum L.). Int J Res Appl Nat Soc Sci 2:211–218 Shanmugam V, Kanoujia N, Singh M, Singh S, Prasad R (2011) Biocontrol of vascular wilt and corm rot of gladiolus caused by Fusarium oxysporum f. sp. gladioli using plant growth promoting rhizobacterial mixture. Crop Prot 30(7):807–813 Sharma N, Tripathi A (2008) Integrated management of postharvest Fusarium rot of gladiolus corms using hot water, UV-C and Hyptis suaveolens (L.) Poit. essential oil. Postharvest Biol Technol 47:246–254 Singh RN (1968) Curvularia disease of gladiolus in India. Plant Dis Rep 52:552 Singh BR, Dubey VK (2007) Inhibition of mosaic disease of Gladiolus caused by Bean yellow mosaic- and Cucumber mosaic viruses by virazole. Sci Horticult 114:54–58 Singh PJ, Sidhu GS, Kuma R (2005) Effect of pre-and post-inoculative sprays of fungicides on blight of gladiolus caused by Botrytis gladiolorum. J Orn Horticult 8:137–139 Singh PJ, Aulakh JS, Thind TS, Arora JS (2006) Management of Curvularia trifolii f. sp. gladioli of gladiolus through chemicals and host resistance. Pl Dis Res-Ludhiana 21:106

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Singh PJ, Kumar R, Sidhu GS (2008) Efficacy of fungicides against Botrytis gladiolorum of gladiolus. Plant Dis Res 23:19–23 Singh PJ, Sidhu GS, Kumar R, Thind TS (2011) Superior performance of kresoxim-methyl (Stroby) and trifloxystrobin (Flint) against Botrytis blight (Botrytis gladiolorum) of gladiolus (Gladiolus x hortulanus Bailey). Plant Dis Res 26:101–105 Stein A (1995) Gladiolus. In: Loebenstein G, Lawson RH, Brunt AA (eds) Virus and virus-like diseases of bulb and flower crops. Wiley, New York, pp 281–292 Straathof TH, Roebroeck EJA, Löffler HJM (1998) Studies on Fusarium-Gladiolus interactions. J Phytopathol 146:83–88 Torres DP, Silva MA, Furtado GQ (2013a) Infection of Curvularia gladioli on different gladiolus genotypes. Trop Pl Pathol. 38: doi.org/10.1590/S1982-56762013000600011 Torres DP, Silva MA, Pinho DB, Pereira OL, Furtado GQ (2013a) First report of Curvularia gladioli causing a leaf spot on Gladiolus grandiflorus in Brazil. Plant Dis 97:847 Treeful L, Ash C (2000) Aster yellow. http://www.extension.umn.edu/garden/yard-garden/flowers/ aster-yellows University of Illinois Extension (1983) RPD No. 612. http://ipm.illinois.edu/diseases/rpds/612.pdf Valencia-Botín AJ, Jeffers SN, Palmer CL, Buck JW (2013) Fungicides used alone, in combinations, and in rotations for managing gladiolus rust in Mexico. Plant Dis 97:1491–1496 Vicchi V, Bellardi MG (1988) The role of weeds in the epidemiology of Gladiolus viruses and MLO. Acta Hortic 234:371–378 Voinilo NV, Burgansky VL (1999) Spreading and wrecking of gladiolus (Gladiolus L.) viruses in Belarus. Vestsi Natsyyanal’nal Akademii Navuk Belarusi. Seryya Biyalagichnykh Navuk 4:12–15 Vunsh R, Rosner A, Stein A (1991) Detection of bean yellow mosaic virus in gladioli corms by the polymerase chain reaction. Ann Appl Biol 119:289–294 Wada Y, Iwai H, Ogawa Y, Arai K (2000) Comparison of pathogenicity and nucleotide sequences of 30 -terminal regions of bean yellow mosaic isolates from gladiolus. J Gen Plant Pathol 66:345–352 Wade GC (1984) Gladiolus diseases. Tasmania J Agric 19:36–40 Woltz SS (1958) Fertilization of gladiolus. Proc Fla State Hort Soc 69:347–351 Woltz SS, Jones JP (1973) Interactions of nitrogen fertilizer and liming procedure in the control of fusarium wilt of tomato. HortSci 8:137–138 Yamaji Y, Horikoshi K, Yamashita H, Matsumoto T (1998) Detection of tobacco rattle virus on gladiolus by RT-PCR. Res Bull of the Plant Protection Service Japan 0 (34):107–111 Zettler FW, Abo El-nil MM (1977) Bean yellow mosaic-virus infections of Gladiolus in Florida. Plant Dis Rep 61:243–247 Zitter TA, Murphy JF (2009) Cucumber mosaic. The Plant Health Instructor. APSnet. doi:10.1094/ PHI-I-2009-0518-01

Diseases of Tulip

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Robert J. McGovern and Wade H. Elmer

Abstract

Tulip is the most important ornamental geophyte worldwide, grown for bulbs, cut flowers, potted plants, and landscape use. Major increases in consumer demand in the first half of the twentieth century were associated with the identification of many new disease problems. Tulip is susceptible to a number of diseases caused by fungi, bacteria, viruses, and nematodes including Botrytis tulipae, Fusarium oxysporum f. sp. tulipae, Pectobacterium carotovorum, Tulip breaking virus, and Ditylenchus dipsaci that can significantly reduce flower and bulb production. Since the plant is propagated vegetatively, this factor can facilitate the spread of disease if pathogen-free propagative material is not used and integrated disease management is not followed. Keywords

Botrytis tulipae • Fusarium oxysporum f. sp. tulipae • Tobacco rattle virus • Tulip breaking virus • Ditylenchus dipsaci • Paratrichodorus spp. • Trichodorus spp. • Integrated disease management

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1314 2 Fungal and Fungus-Like Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1314 2.1 Tulip Fire [Botrytis tulipae (Lib.)] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1314

R.J. McGovern (*) NBD Research Co. Ltd., Lampang, Thailand Department of Entomology and Plant Pathology, Chiang Mai University, Chiang Mai, Thailand e-mail: [email protected] W.H. Elmer Department of Plant Pathology and Ecology, The Connecticut Agricultural Experiment Station, New Haven, CT, USA e-mail: [email protected] # Springer International Publishing AG 2018 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florists’ Crops Diseases, Handbook of Plant Disease Management, https://doi.org/10.1007/978-3-319-39670-5_49

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2.2 Fusarium Bulb Rot, Fusarium Basal Rot (Fusarium oxysporum (Schlecht.) f. sp. tulipae Act.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Penicillium Bulb Rot, Blue Mold (Penicillium spp.; Penicillium corymbiferum Westling) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Pythium Root and Bulb Rot (Pythium spp. Pythium ultimum, Pythium ultimum Trow var. ultimum, P. irregulare Buisman, P. spinosum Sawada) . . . . . . . . . . . . . . . . . . 2.5 Rhizoctonia Rot and Blight, Gray Bulb Rot [Rhizoctonia tuliparum (Whetzel and Arthur)] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Bacterial and Phytoplasma Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Bacterial Soft Rot [Pectobacterium carotovorum (Erwinia carotovora); Dickeya didantii (Erwinia chrysanthemi)] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Viral Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Nematode Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 Stem and Bulb Nematode [Ditylenchus dipsaci (Kühn)] . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2 Stubby Root Nematode (Trichodorus spp., Paratrichodorus spp.) . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1

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Introduction

Tulip (Tulipa gesneriana and Tulipa hybrids), a Eurasian and North African perennial plant, has been associated with the Netherlands since the late sixteenth century, is synonymous with Spring, and is the most important ornamental geophyte in the world. Tulip is grown for bulbs, cut flowers, potted plants, and landscape use. The Netherlands dominates tulip production on land accounting for 88% of the area devoted to the crop worldwide; Japan, France, Poland, Germany, and New Zealand are distant competitors (Buschman 2005). The country produced an estimated 4.32 billion tulip bulbs (Buschman 2005). The total sales value of tulip cut flowers in the USA in 2012 was $65.3 million (Anon. 2014). Tulip is susceptible to a number of diseases caused by fungi, bacteria, viruses, and nematodes including Botrytis tulipae, Fusarium oxysporum f. sp. tulipae, Pectobacterium carotovorum, Tulip breaking virus, Tobacco rattle virus, Ditylenchus dipsaci, Paratrichodorus spp., and Trichodorus spp. that can significantly reduce flower and bulb production. Since the plant is propagated vegetatively, this factor can facilitate the spread of disease if pathogen-free propagative material is not used and integrated disease management is not followed.

2

Fungal and Fungus-Like Diseases

2.1

Tulip Fire [Botrytis tulipae (Lib.)]

Geographic occurrence and impact. Tulip fire is a very common and serious disease wherever tulips are grown. The pathogen can cause extensive losses in both flower and bulb production and landscape plantings.

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Symptoms/signs. All parts of the tulip may be affected. As its name implies, the disease can move very rapidly through a tulip crop given a conducive environment. Yellow to brown round lesions may be seen on bulbs when the outer scales are removed. Small, black long-lasting resistive structures (sclerotia) of B. tulipae may form on the outer scales. Initial symptoms on leaves are small, yellowish, elongated spots that become grayish-brown blotches, which coalesce as they enlarge (“fire”); sclerotia may form in necrotic tissue (Fig. 1). Leaves may fail to emerge. Similar but more depressed lesions form on stalks and may cause stalk breakage. Shoot distortion and stunting may occur. The fungus causes brown lesions on light-colored flowers and whitish lesions on darker-colored flowers (Fig. 2); complete blighting of flowers is possible. Plant death can occur depending on the environment and tulip genotype. In general, T. fosteriana is more susceptible to B. tulipae than T. gesneriana (Reyes et al. 2005). Fig. 1 Tulip fire on leaves caused by Botrytis tulipae; note the presence of black sclerotia (The Pennsylvania State University Department of Plant Pathology and Environmental Microbiology # 2017. All Rights Reserved.)

Fig. 2 Tulip fire on flowers caused by Botrytis tulipae (R. Wick # 2017. All Rights Reserved.)

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Biology and epidemiology. Botrytis tulipae survives in infected bulbs and sclerotia in the soil and in association with plant tissue. In spring, sclerotia germinate as hyphae or give rise to conidiophores and conidia, which disseminate the fungus via water splash and air currents to foliar tissue. Germination of sclerotia is greatest at 5  C (41  F) (Coley-Smith and Javed 1972). Infection by the fungus is enhanced by cool soil temperatures. Doornik and Bergman (1971) found that a soil temperature of 5  C (41  F) and the resultant slower flowering led to higher shoot infection by B. tulipae than a soil temperature of 15  C/59  F, which resulted in faster flowering. In addition, more new bulb infections occurred at a soil temperature of 9  C (48  F) than 18  C (64  F) (Doornik and Bergman 1973). High humidity and rainfall and moderate temperatures favor the blight phase of tulip fire, and a fuzzy gray mold consisting of conidiophores and conidia may form in diseased tissue. The host range of B. tulipae appears to be limited to tulip (Schönbeck 1976). Management • Cultural practices – Use vigorous, blemish-free, and pathogen-free bulbs. Rotate to another production site each year. Researchers at Washington State University are investigating the effectiveness of growing cover crops in rotation with tulip to reduce tulip fire (MacKenzie 2015). Avoid high rates of nitrogen. Use a proper plant spacing and rigorous weed control to increase air circulation and decrease plant wetness. Harvest bulbs during dry weather and process and store them properly to avoid injury. • Sanitation – Discard/burn infected bulbs and other diseased plant tissue when plant surfaces are dry. • Soil disinfestation – Use a labeled fumigant or, where feasible, steam treatment to reduce the inoculum of B. tulipae in the soil (refer to ▶ Chap. 8, “Soil/Media Disinfestation for Management of Florists’ Crops Diseases”). • Fungicides – A large number of fungicides are labeled in the USA for management of Botrytis on tulip (Pscheidt and Ocamb 2016). (Also refer to ▶ Chap. 7, “Fungicides and Biocontrols for Management of Florists’ Crops Diseases”). Resistance to benzimidazole and dicarboximide fungicides has been identified in B. tulipae (Hsiang and Chastagner 1990). It is important to rotate fungicides with different modes of action to prevent the development of fungicide resistance; refer to the Fungicide Resistance Action Committee website (http://www.frac. info/home) for more information. • Resistance – Absolute resistance was found in T. tarda, but this species cannot be crossed with the various T. gesneriana. Partial resistance was found in some cultivars of T. gesneriana and T. kaufmanniana (Straathof et al. 2002).

2.2

Fusarium Bulb Rot, Fusarium Basal Rot (Fusarium oxysporum (Schlecht.) f. sp. tulipae Act.)

Geographic occurrence and impact. Fusarium bulb rot was described in the 1950s when the disease was reproduced using strains of F. oxysporum (Slootweg 1955). In 1958, Apt (1958) assigned the formae specialis, F. oxysporum f. sp.

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tulipae, to the pathogen. Although the disease can occur wherever tulips are grown, most damage is observed during bulb production and storage (Gould and Miller 1970; Schenk and Bergman 1969). There was a striking increase in the incidence of disease in the 1960s following consumer demands for increased production (Schenk and Bergman 1969). Dutch growers opted for shorter rotations that favored a buildup of inoculum. In addition, the advent of mechanization, which led to increased wounding during harvest, combined with poor grading to remove damaged bulbs favored a widespread increase in Fusarium bulb rot. Symptoms/signs. The disease was first coined by Slootweg (1955) as “zuur” meaning sour disease due to the rank smell that infected bulbs produced. Infected bulbs tend to be whitish and the infection is clearly demarcated as dry, firm, sunken areas (Fig. 3). Depending on the relative humidity, the mycelium can be observed under the outer husk at the base of the bulb (Fig. 4) (Apt 1958; Bergman 1965). Frequently, a tan gummosis exudes, hardens, and can glue scales and occasionally entire bulbs together in storage (Liou et al. 2006). In the field, infected bulbs sprout poorly and their leaves usually turn red, wilt, and eventually die (Bergman 1965). Occasionally, atypical symptoms are associated with Fusarium bulb rot; Schenk and Bergman (1969) reported mycelium only between the parenchymatous tissues in tulip bulbs that had been forced in the greenhouse after a precooling at 5  C/41  F. Although variation can appear in strains of F. oxysporum f. sp. tulipae, molecular assays found close relatedness between all isolates from tulip and from lily suggesting a common progenitor (Baayen et al. 2000). Biology and epidemiology. Latent infections by Fusarium oxysporum f. sp. tulipae commonly occur in commercial tulip production and can affect the resulting health of the daughter bulbs harvested in the following year (Bergman and Bakker-Van Der Voort 1979). This mode is also the major means for disseminating the pathogen

Fig. 3 Tulip bulbs infected with Fusarium oxysporum f. sp. tulipae (Oregon State University Plant Disease Clinic # 2017. All Rights Reserved.)

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Fig. 4 Mycelium of Fusarium oxysporum f. sp. tulipae colonizing between the leaf scales of tulip bulbs (Oregon State University Plant Disease Clinic # 2017. All Rights Reserved.)

abroad (Schenk 1972). The pathogen can remain viable in the bulb during dry storage for several months but, once established in the soil, can persist for longer periods (Price 1974). Under greenhouse conditions, the pathogen can penetrate through the roots into the tissue of the planted bulbs and colonize the basal plate of the young bulb in the field. Infection more frequently occurs on the fleshy outer scale of the new bulb. This was noted to occur during the weeks before the bulb skin turns brown presumably due to the warmer soil temperatures that favor disease (Bergman and Noordermeer-Luyk 1973). Another problem comes from the fact that the Fusarium infection tends to produce ethylene, a plant hormone (Jarecka et al. 2008). Ethylene can cause flower abortion, stunting, poor rooting, and excessive gummosis. Infection of young growing bulbs by the pathogen occurs mainly during the last few weeks before harvest. Beijersbergen and Lemmers (1972) found that extracts of the bulb skin contained an antifungal compound called tulipalin (α-methylene butyrolactone). The compound appears to decrease antifungal activity as soon as the tissue dries. Bergman and Beijersbergen (1971) speculated that variations in the compound may influence the wide range of reactions to Fusarium bulb rot among different cultivars at different times. Observations by researchers have concluded that the fungus is not a typical vascular parasite but tends to colonize nonvascular tissue resulting in only bulb rots (Schenk and Bergman 1969; Suazo 2012). Management • Cultural practices – Minimizing damage to the bulb during harvest followed by careful culling of damaged bulbs can significantly reduce disease incidence. The distinct smell that is present with infected bulbs should be employed as a tool to cull damaged bulbs. In addition, avoiding excessive nitrogen fertilizing and making an attempt to not delay harvest since warmer soil temperatures will increase disease incidence is important. A number of reports have shown that

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fertilizing with nitrate nitrogen (as opposed to the ammonium form) can reduce Fusarium disease of ornamental bulbs (Gullino et al. 2015). • Resistance – Considerable progress has been made in screening and breeding for resistance (Eijk et al. 1979; Straathof et al. 1996), which provides the most effective management strategy for Fusarium bulb rot of tulip. Van Tuyl and van Creij (2007) discovered almost complete resistance in the T. gesneriana assortment, but since seedling selection produced some susceptible plants, plants still need to be rescreened at the clonal level. Growers should inquire whether or not resistance is known to occur in cultivars they seek to purchase. • Fungicides – Strategies useful for suppression of Fusarium diseases on Narcissus (refer to ▶ Chap. 40, “Diseases of Daffodil (Narcissus)”) would be appropriate for Fusarium bulb rot of tulip (Hanks 1996) However, growers should be aware of fungicide-resistant strains of F. oxysporum f. sp. tulipae (Duineveld and Beijersbergen 1975).

2.3

Penicillium Bulb Rot, Blue Mold (Penicillium spp.; Penicillium corymbiferum Westling)

Geographic occurrence and impact. The disease is a storage rot problem and common to most production facilities. Changes in modified atmospheric conditions for the storage of tulips have favored an increase in the disease (Prince et al. 1982). It is reported in the USA and Europe (Dugan et al. 2014). The disease is caused by Penicillium corymbiferum. However, the pathogen has recently been subdivided into seven taxa based on pathogenicity (Overy et al. 2005a). From the original “corymbiferum group,” the species P. hirsutum, P. radicicola, P. tulipae, and P. venetum were found to be the predominant pathogens on tulip Tulipa gesneriana. Symptoms/signs. Bulbs in storage initially develop brown spots. The blue mold develops on the bulb basal root plate, and/or between the scales. Internally, the disease can discolor the young leaf initials. Depending on the severity of the disease, the entire bulb or only a portion of it shows symptoms. When infections are severe, the bulb will frequently dry and become very hard. Sprouted bulbs can have reddish stunted foliage. Biology and epidemiology. These fungi generally attack bulbs that are stressed or injured and then put into storage under moist conditions. The pathogens in the P. corymbiferum group are unique among Penicillium species in that most can grow at 0  C (32  F) but can proliferate at 5  C (41  F) (Overy et al. 2005b). Management • Cultural practices – The pathogen has been reported on other bulb crops including crocus, gladiolus, iris, and onion (Dugan et al. 2014), so field rotations to nonsusceptible crops should be practiced to limit inoculum buildup. Growers should dig their bulbs at the proper time and not delay. Care should be exercised in

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harvesting bulbs to avoid bruising. Bulbs should be stored with moderate humidity and sufficient air circulation. All damaged bulbs should be destroyed. • Fungicides – Fungicide bulb soaks can be effective. Thiabendazolein in water at 13  C (55  F)–24  C (75  F) for 15–30 min or thiophanate-methyl-based products applied at 27  C (80  F)–29  C (85  F) have been recommended (Prince et al. 1986, 1988). Bulbs should be treated within 48 h after digging. Growers should be aware that benzimidazole-resistant strains of the pathogen have been reported (Duineveld and Beijersbergen 1975).

2.4

Pythium Root and Bulb Rot (Pythium spp. Pythium ultimum, Pythium ultimum Trow var. ultimum, P. irregulare Buisman, P. spinosum Sawada)

Geographic occurrence and impact. The disease was first reported in England, Denmark, and Holland (Moore and Buddin 1937). Although different species were implicated, the disease appears to be the same (Moore and Buddin 1937). In Japan, the species P. ultimum var. ultimum, P. irregulare, and P. spinosum were isolated the most frequently (Ichitani et al. 1991). The disease can be devastating if plants are grown in heavy wet soils. Symptoms/signs. Pythium root rot causes a browning and collapse of the roots. It can frequently be encountered with Rhizoctonia infection (Fig. 5). In addition, symptoms on the aboveground growth can frequently be observed as stunting, dieback of leaf tips, weak or distorted growth, and yellowing (Fig. 6). Biology and epidemiology. Pythium species are ubiquitous soil species and can be devastating when soil conditions are optimal. Cool wet soil favors disease. The pathogen(s) may also be spread on bulbs into new areas. Fig. 5 Root rot symptoms of Pythium and Rhizoctonia on tulip (Tom Creswell # 2017. All Rights Reserved.)

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Fig. 6 Aboveground symptoms of Pythium root rot showing leaf tip dieback (Tom Creswell # 2017. All Rights Reserved.)

Management • Cultural practices – Careful culling of damaged bulbs will avoid creating potentially susceptible tissue. Improving soil water drainage is paramount to managing Pythium root rot. Packing material can frequently be contaminated with Pythium, so recycling of packing materials should be avoided if they cannot be steam disinfested (Van Bruggen and Duineveld 1993). • Fungicides – Soil treatments with etridiazole gave good results (Humphreys-Jones and de Rooy 1974). In Holland, the compounds etridiazole, fenaminosulf, and prothiocarb are widely used although results are not always satisfactory. In glasshouse production of tulips, furalaxyl and metalaxyl along with dichloropropene + etridiazole have been used with varying success (Koster and De Rooij 1980). • Biological control – Weststeijn (1990) explored using fluorescent Pseudomonas isolates against P. ultimum and found suppression with one isolate when it was mixed into the soil.

2.5

Rhizoctonia Rot and Blight, Gray Bulb Rot [Rhizoctonia tuliparum (Whetzel and Arthur)]

Geographic occurrence and impact. The pathogen Rhizoctonia tuliparum is closely related to R. solani and has been observed in Europe and the USA (Whetzel and Arthur 1924). R. tuliparum has seriously threatened flower bulb production at different times. In 1974, a severe outbreak of gray bulb rot caused major losses on tulips in Lincolnshire, England.

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Symptoms/signs. Infected bulbs rarely emerge and ones that do emerge are distorted, stunted, and rarely flower. Bulbs and roots turn grayish in color and progressively dry and shrivel at the site of infection. Roots and bulbs become discolored and collapse. If the soil is very wet, it is not uncommon to have Pythium infections along with Rhizoctonia (Figs. 5 and 6). Biology and epidemiology. Rhizoctonia tuliparum overwinters as sclerotia and as mycelium on plant debris. One study documented that about 10% of the sclerotia were still viable after 10 years. Where severe outbreaks of disease have occurred, longer intervals between susceptible crops are suggested. Growers should avoid hosts that are also susceptible to R. tuliparum such as lilies and onions. R. solani is closely related to R. tuliparum and has been reported to cause bulb rot. Many times a mixture of genetically related strains may be found. In the Netherlands, the pathogens causing Rhizoctonia disease on tulips may comprise a mixture of distinct strains (Dijst and Schneider 1996). Cooler soil temperature may play a role in dictating which strains of R. solani dominate (Schneider et al. 2001). Management • Cultural practices – Care should be exercised when harvesting bulbs to minimize damage. Any damaged bulbs should be discarded. • Sanitation – When tulips are forced in the greenhouse, sanitation should be practiced and clean disinfested soil or soilless potting mixes should be used. Field plants should be placed in areas where other susceptible plants have not been grown, e.g., lilies and onions. Once bulbs are infected, they must be discarded. • Fungicides – Dip bulbs in PCNB after harvest. On potted plants, drench applications of thiophanate methyl are also recommended. Other fungal or fungus-like organisms reported to be associated with Tulipa spp. (See Farr DF and Rossman AY Fungal Databases, Systematic Mycology and Microbiology Laboratory, ARS, USDA. Retrieved 23 May 2016 from http://nt.arsgrin.gov/fungaldatabases/): Aecidium tulipae Alternaria sp. Ascochyta tulipae Aspergillus niger Aspergillus sp. Botrytis cinerea Botrytis parasitica (Botrytis tulipae) Botrytis sp. Cercospora sp. Cercosporella sp. Colletotrichum dematium Colletotrichum fioriniae

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Diseases of Tulip

Colletotrichum sp. Corticium solani (Rhizoctonia solani) Corticium vagum (Haplotrichum curtisii) Curvularia sp. Drechslera sp. Fusarium oxysporum Fusarium poae Fusarium roseum Fusarium sp. Gibberella avenacea (Fusarium avenaceum) Gloeosporium thuemenii Gloeosporium thumenii f. tulipae Ilyonectria sp. Nectria inventa (Acrostalagmus luteoalbus) Nectria sp. Oidium sp. Penicillium cyclopium (Penicillium aurantiogriseum) Penicillium verrucosum var. cyclopium Phytophthora cactorum Phytophthora citricola Phytophthora cryptogea Phytophthora erythroseptica var. erythroseptica Phytophthora hedraiandra Phytophthora porri Phytophthora sp. Puccinia prostii Puccinia sp. Puccinia tulipae Pythium intermedium (Globisporangium intermedium) Pythium irregulare (Globisporangium irregulare) Pythium oopapillum Pythium ultimum (Globisporangium ultimum) Rhizopus necans Rhizopus sp. Rhizopus stolonifer Sclerotinia fuckeliana (Botrytis cinerea) Sclerotinia sativa Sclerotinia sclerotiorum Sclerotinia sp. Sclerotium perniciosum Sclerotium rolfsii (Athelia rolfsii) Sclerotium tulipae (Botrytis tulipae) Sclerotium tuliparum (Rhizoctonia tuliparum) Septoria sp. Synchytrium laetum

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Thanatephorus cucumeris (Rhizoctonia solani) Thielaviopsis basicola Urocystis sp. Uromyces erythronii Ustilago heufleri (Vankya heufleri) Ustilago vaillantii (Antherospora vaillantii) Vankya heufleri Verticillium lateritium (Acrostalagmus luteoalbus)

3

Bacterial and Phytoplasma Diseases

3.1

Bacterial Soft Rot [Pectobacterium carotovorum (Erwinia carotovora); Dickeya didantii (Erwinia chrysanthemi)]

Geographic occurrence and impact. The disease has been reported in Australia, Belarus, China, Japan, the Netherlands, Turkey, and the USA (Boyraz et al. 2006; Fahy and Persley 1983; Fujimura et al. 2004; Gill et al. n.d.; Guolan and Zhang 1998; Gvozdyak et al. 1985) and can cause production and postharvest losses. Symptoms/signs. Mature bulbs are most frequently attacked by soft-rotting bacteria. Infected bulb scales initially appear water-soaked and light yellow to light brown or light gray to white. As the infection advances throughout the bulb, affected tissue becomes soft and watery (macerated) due to the bacterium’s pectolytic enzymes (Fig. 7). A dark, foul-smelling liquid exudes from diseased bulbs. Pectobacterium carotovorum has also been reported to cause small, ellipsoidal water-soaked spots which enlarge to blight leaves (Fig. 7) and bud neck rot in tulip in Konya, Turkey (Boyraz et al. 2006). The incidence of bacterial leaf blight in that area was about 27% and 17% in 2002 and 2003, respectively.

Fig. 7 Bulb soft rot (left) and advanced leaf blight (right) caused by Pectobacterium carotovorum (Boyraz et al. 2006)

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Biology and epidemiology. Pectobacterium and Dickeya spp. attack the soft, fleshy tissue of many plant hosts. They survive in infected propagative material and in soil in association with decaying plant material and are spread by water; insects that feed on declining plant tissue; tools, hands, and clothing of workers; equipment; and airborne particles and aerosols (Bradbury 1977). The bacteria enter plants through natural openings (stomata and lenticels) and wounds. The disease is favored by poorly drained soil and air circulation. Management • Cultural – Use vigorous, blemish-free, and pathogen-free bulbs. Avoid planting in poorly drained areas and use adequate spacing to increase air circulation. Harvest bulbs during dry weather and process and store them properly to avoid injury. Control possible insect vectors. • Sanitation – Discard/burn infected bulbs and other diseased plant tissue when dry. Workers should disinfest hands, tools, and equipment after processing rotted plant tissue; 10% household bleach, 70% alcohol, or a commercial material may be used to disinfest tools and equipment (Gill et al. n.d.) (refer to ▶ Chap. 9, “Sanitation for Management of Florists’ Crops Diseases”). • Bactericides – refer to ▶ Chap. 7, “Fungicides and Biocontrols for Management of Florists’ Crops Diseases” for general information on bactericides. Other bacteria reported to be associated with Tulipa spp. include: Bacillus sp. (Gvozdyak et al. 1985) Burkholderia andropogonis (syn. Pseudomonas andropogonis) (Morikawa et al. 1996) Burkholderia gladioli (syn. Pseudomonas gladioli) (Morikawa et al. 1996) Curtobacterium flaccumfaciens pv. oortii (syn. Corynebacterium flaccumfaciens pv. oortii) (Fox 2014) Rhodococcus fascians (Tilford) Goodfellow (syn. Corynebacterium fascians (Tilford) Dowson.) (van Aartrijk et al. 2000)

4

Viral Diseases

Geographic occurrence and impact. Viruses are among the most damaging pathogens of tulips because of the reductions in quantity and quality of flowers and bulbs that they cause. In addition, viruses are extremely difficult to control in a vegetatively propagated crop such as tulip. A wide range of viruses have been reported in tulip. The most important viruses are Arabis mosaic virus (ArMV), Cucumber mosaic virus (CMV), Lily symptomless virus (LSV), Tobacco necrosis virus (TNV), Tobacco rattle virus (TRV), and particularly Tulip breaking virus (TBV). The geographic occurrence and host range of these viruses are indicated in Table 1. Other viruses reported to infect Tulipa spp. include those in the following genera (Asjes 1994; Lesnaw and Ghabrial 2000; Mowat 1972):

Mechanical; the green peach aphid (Myzus persicae)

Mechanical; the fungus Olpidium brassicae (Chytridiales)

South Korea, the Netherlands, Poland

The Czech Republic, New Zealand, the Netherlands, Poland

Lily symptomless virus (LSV) Carlavirus Tobacco necrosis virus (TNV) Necrovirus Depending on the strain: Chenopodium amaranticolor, C. quinoa; Cucumis sativus (cucumber); Nicotiana clevelandii, N. tabacum (tobacco); Phaseolus vulgaris (bean)

Lilium (lily)

A very large host range including many horticultural crops and weeds

Mechanical; > 60 aphid spp.

The Czech Republic, Japan, South Korea, New Zealand, the Netherlands, Poland

Cucumber mosaic virus (CMV) Cucumovirus

Other natural hosts Many species of wild and cultivated monocotyledonous and dicotyledonous plants

Transmission Mechanical; dagger nematodes (Xiphinema spp.); seed of some plants

Geographic occurrence The Netherlands

Virus, genus Arabis mosaic virus (ArMV) Nepovirus

Table 1 Selected tulip viruses

Brown necrotic streaks in leaves and stems

Color breaking in flowers and leaf mottling (Fig. 9)

Symptoms Oval gray or brown ring spots primarily at the bases of lower leaves (Fig. 8); whole plants appear brittle, dwarfed and twisted, and may die Sunken brown spots or rings in bulbs which produce distorted and stunted plants; color breaking in flowers

Asjes 1994; Pearson et al. 2009; Sochacki 2013; Teakle and Brunt 1982/1990

Asjes 1994; Duraisamy and Pokorny 2009; Francki and Habili 1980/ 1987; Kim et al. 1995; Pearson et al. 2009; Sochacki 2013; Yamamoto 1971 Asjes 1986, 1994; Kim et al. 1995; Sochacki 2013

References Asjes 1976; Murant 1984;

1326 R.J. McGovern and W.H. Elmer

New Zealand, the Netherlands, Poland, South Africa

Worldwide, wherever tulips are grown

Japan, New Zealand, the Netherlands, Norway, Poland, Sweden, UK

Tobacco rattle virus (TRV) Tobravirus

Tulip breaking virus (TBV) Potyvirus

Tulip virus X (TVX) Potexvirus

Mechanical; a number of aphid spp. including the green peach aphid (Myzus persicae) and the cotton aphid (Aphis gossypii) Mechanical

Mechanical; stubby root nematodes (Paratrichodorus, Trichodorus spp.); seed of some plants

Only reported in tulip

Lilium (lily)

At least nine families including Brassicaceae, Cucurbitaceae, Leguminosae, and Solanaceae

Chlorotic or necrotic gray-brown streaks in leaves; intensified colored or necrotic streaks in flowers

Symptoms in Tulipa spp. and Darwin hybrids include color breaking in flowers and leaf chlorosis

Mottling and lightcolored streaks in leaves

Dees et al. 2011; Mowat 1982, 1990; Sochacki 2013; Ward et al. 2008

Asjes 1994; Duraisamy and Pokorny 2009; Harrison and Robinson 1984; Pearson et al. 2009; Sochacki 2013; Whitlock 1984 Lesnaw and Ghabrial 2000; Philips 1986

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Ampellovirus – Tulip severe mosaic virus (TSMV) Ophiovirus – Tulip mild mottle mosaic virus (TMMMV) Potyvirus – Lily mottle virus (LMoV), Rembrandt tulip-breaking virus (ReTBV), Tulip band-breaking virus (TBBV), Tulip chlorotic blotch virus (TCBV), Tulip top-breaking virus (TTBV) Tombusvirus – Tomato bushy stunt virus (TBSV) Symptoms/signs. Virus symptoms in tulip vary depending on the virus, virus strain, mixed viral infections, and tulip variety. In addition, different viruses can cause similar symptoms in the same tulip variety making diagnosis of virus infection by symptoms alone unreliable. Besides TBV, other potyviruses such as TBBV, TTBV, ReTBV, and LMoV, as well as non-potyviruses such as ArMV, CMV, and LSV, can cause some degree of color-breaking symptoms (Dekker et al. 1993; Lesnaw and Ghabrial 2000). Historically, color breaking of flowers induced by viruses was viewed as an attractive and desirable trait. However, this virusbased characteristic is unstable, and virus infection leads to reduction in the quality and quantity of tulip flowers and bulbs (Bos 1970; Yamamoto 1971). Today, for the most part, flower variegation in tulips is the result of stable genetic traits produced by plant breeding. Common symptoms of selected viruses in tulip are presented in Table 1. Biology and epidemiology. As obligate pathogens, viruses need a living host, either tulip or an alternate, for their survival and replication. A number of important tulip viruses, including all those listed in Table 1, are mechanically transmissible in plant sap. This factor is very important for virus spread in a vegetatively propagated crop such as tulip. In addition, most of these viruses also have vectors which disseminate them to new plants: CMV, LSV, and TBV are vectored in a non-propagative (nonpersistent) manner on the stylets of aphids; ArMV and TRV are spread by dagger and stubby root nematodes, respectively, while TNV is vectored by the fungus Olpidium brassicae. Management. Once infected by a virus, unlike with other pathogens, only demanding tissue culture procedures can free plants of viruses, and even these stringent measures are not always successful. Therefore, management of tulip viruses must be based on prevention of infection by integrating appropriate practices: • Cultural practices – Use certified virus-free propagative material. Isolate foundation stock and exclude aphids with fine mesh screening. Harvest early to reduce exposure to increasing aphid populations (Chastagner et al. 2002). Thomsen (1980) reported that the spread of TRV in sandy soil was greater when the bulbs were grown at 15 cm than at 7.5 cm. [The highest densities of T. similis, a TRV vector, in sandy and sandy-loam soils are found at a depth of 20–30 cm (Decraemer and Geraert 2006).] Spent mushroom compost added in-furrow at planting and fodder radish as a preceding crop reduced the infection percentage in

43

•

•

•

•

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tulip under favorable conditions for TRV infection (Zoon et al. 2002). Late planting of TRV susceptible cvs. may be helpful (van Aartrijk et al. 2000). Sanitation – Routinely scout fields and remove and dispose of/burn diseased and volunteer bulbs. Control weeds, which may provide reservoirs of viruses and their vectors. Disinfest cutting tools, containers, and equipment (refer to ▶ Chap. 9, “Sanitation for Management of Florists’ Crops Diseases”). Insecticides – Asjes (1981) found that weekly sprays of mineral oil were more effective in reducing spread of the aphid-vectored TBV than the insecticides pirimicarb or permethrin. It was further demonstrated that the combination of mineral oil with a pyrethroid was more effective than either insecticide alone in reducing the spread by aphids of TBV and LSV in lily (Asjes 1991). Soil disinfestation – Use an appropriate preplant soil fumigant or other soil disinfestation practice to reduce the density of nematode and fungal vectors of ArMV, TNV, and TRV (refer to ▶ Chap. 8, “Soil/Media Disinfestation for Management of Florists’ Crops Diseases”). Resistance – Three tulip cvs., Apeldoorn, Kees Nelis, and Lustige Witwe, appeared to be susceptible to TBV, and four, Cantata, Juan, Madame Lefeber, and Princeps, showed a high degree of resistance to the virus (Romanow et al. 1990).

5

Nematode Diseases

5.1

Stem and Bulb Nematode [Ditylenchus dipsaci (Kühn)]

Geographic occurrence and impact. The nematode occurs in most temperate areas (Europe and the Mediterranean region, North and South America, northern and southern Africa, Asia and Oceania), but it does not appear to establish in tropical regions except at higher elevations that have temperate climates (Anon. 2017). Ditylenchus dipsaci infects and causes damage to a wide range of other cultivated hosts including Scilla spp., snowdrops (Galanthus spp.), Iris spp., Hyacinth spp., Narcissus spp., onions (Allium spp.), oat (Avena sativa), phlox, and strawberry (Fragaria  ananassa) (Agrios 2005; Slootweg 1960). Symptoms/signs. General symptoms of D. dipsaci include swellings and twisting of leaves and stems and necrosis or rotting of stem bases, bulbs, tubers, and rhizomes; seedling death may also occur. In tulip, aboveground parts exhibit light yellow to white spots or swellings which may coalesce, and cracking of the epidermis occurs with white, ragged edges. Especially characteristic symptoms in tulip flowers include swellings, which become warty growths; often a portion of the petal above the stem infection site remains wholly or partly green (Fig. 10). Affected bulbs frequently dry out during storage and are secondarily affected by Penicillium and mites (van Aartrijk et al. 2000).

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Fig. 8 Oval ring spots caused by ArMV (Asjes 1976)

Fig. 9 Color breaking in flowers (left) and leaf mottling (right) caused by LSV (Horst 2008)

Biology and epidemiology. Ditylenchus dipsaci is a migratory endoparasite of bulbs, stems, and leaves and normally completes its life cycle in these tissues. The nematode has a five-stage life cycle and the ability to enter into a dormant stage. Ditylenchus dipsaci enters the soil only during unfavorable conditions such as when extensive bulb decay occurs. Juveniles then move out of plant tissue and accumulate around the basal plates of dried bulbs as grayish-white, cottony masses called “nematode wool,” through which they can remain dormant for a number of years. Nematodes may also survive in many alternate cultivated crops such as those cited above and weeds such as chickweed [Stellaria spp.], groundsel [Senecio vulgaris], shepherd’s purse [Capsella bursa-pastoris], grasses, and wild onions (Allium spp.)

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Fig. 10 Tulip flower on left infected with stem nematodes; note lesion on the stem (lower arrow) and the persistence of green coloration in the petal nearest to the infection site (upper arrow) (Gratwick 1992)

(Slootweg 1960). However, D. dipsaci occurs in more than 20 biological races, some of which have a limited host range (Anon. 2017).The nematode is spread in infected bulbs and by contaminated soil moved by water, wind, and machinery and on the clothing of workers (van Aartrijk et al. 2000). Management • Cultural practices – Use nematode-free bulbs, and rotate with a nonhost. Eliminate weed hosts. Flooding of infested sandy soil for about 10 weeks at a water temperature of about 17  C/62  F resulted in the elimination of D. dipsaci (Muller and van Aartrijk 1989). • Sanitation – Remove and discard/destroy infected plants and bulbs. Disinfest equipment and bulb containers (refer to ▶ Chap. 9, “Sanitation for Management of Florists’ Crops Diseases”). Tulip bulbs are assumed to be sensitive to hot water treatments above 43  C/ 109  F (van Dam 2013). However, research evaluating various pretreatment temperatures (PTT) and hot water treatments (HWT) indicated that a HWT of 4 h at 47  C/116.6  F preceded by PTT at 27, 30, or 33  C (80.6, 86, or 98.6  F) for 1 week resulted in an average yield of 39, 64, and 92%, respectively, compared to the control treatment (van Dam 2013). Total elimination of D. dipsaci occurred with 4 h at 47  C/116.6  F, after PTT at 30  C/86  F for 1 week. • Nematicides – Nøhr Rasmussen and Lindhardt (1980) found that a bulb dip in oxamyl and in-furrow application of aldicarb or carbofuran at planting provided satisfactory control. Multiple field application of aldicarb or oxamyl significantly

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reduced the number of infected tulip bulbs but did not eradicate D. dipsaci from highly infected bulbs (Windrich 1985). • Soil disinfestation – Use a preplant soil disinfestation procedure such as steam, fumigation, anaerobic soil disinfestation, etc. (Weststeijn and De Rooy 1974) (refer to ▶ Chap. 8, “Soil/Media Disinfestation for Management of Florists’ Crops Diseases”).

5.2

Stubby Root Nematode (Trichodorus spp., Paratrichodorus spp.)

Geographic occurrence and impact. These nematodes are of global significance and can cause direct damage to the roots of a large and very diverse array of plant hosts including agronomic crops, fruit and forest trees, grasses, ornamentals, vegetables, and weeds; a number of trichorid nematodes also cause extensive damage as virus vectors (Anon. n.d.). Symptoms/signs. Symptoms include stubby and reduced roots, chlorosis, wilting, and stunting. Biology and epidemiology. Trichodorus and Paratrichodorus are migratory ectoparasitic nematodes that feed on the epidermal cells at or near the root tip region, but never wholly enter the root tissue. The nematode lays eggs in the soil, which hatch to produce juveniles and then adults, completing their life cycle in 20–45 days under favorable conditions. All free juvenile stages and adults can attack and feed on plants (Agrios 2005; Decraemer and Geraert 2006). Stubby-root nematodes are obligate plant parasites; they must feed on plants in order to survive and reproduce (Crow 2014). As mentioned previously, a number of Trichodorus and Paratrichodorus species vector TRV. However, there is a pronounced specificity between virus serotype and vector. For example, the PRV serotype of TRV from Europe is only transmitted by P. pachydermus (Brown et al. 2004). Adults and juveniles can transmit TRV, but the virus is probably not retained through the molt and is not transovarial (Robinson and Harrison 1989). Management. Refer to control practices for Trichodorus and Paratrichodorus indicated in the Viral Diseases management section above and ▶ Chap. 8, “Soil/ Media Disinfestation for Management of Florists’ Crops Diseases.” Other nematodes reported to attack tulip: Aphelenchoides subtenuis (Cobb) (van Aartrijk et al. 2000) Ditylenchus destructor (Thorne) (van Aartrijk et al. 2000) Pratylenchus penetrans (Cobb) (van Aartrijk et al. 2000)

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Index

A Abiotic, 18 diseases, 905 disorders, 938–939 Absolute resistance, 109 Aceria, 709 Acervuli, 286, 717 Achromobacter, 261 Acibenzolar-S-methyl, 269, 596, 1108 Acidic solutions, 273 Acidifiers, 273, 1139 Acidovorax anthurii, 284, 305–307 Acidovorax avenae subsp. cattleyae, 648 Acidovorax leaf spot, 650 Acinetobacter calcoaceticus, 261 Aconitum latent virus, 526 Acquisition, 74 Acrostalagmus luteoalbus, 1323, 1324 Acute testing, 141 Acyrthosiphon (Aulacorthum) solani, 484 Additive gene action, 110 Adiantum raddianum, 260, 271 Aecia, 730 Aecidium A. kalanchoe, 1013 A. narcissi, 1166 A. tulipae, 1322 A. umbilici, 1013 Aerated steam, 421 Aerial endoparasitic nematodes, 364–365 Ageratum yellow vein virus, 614 Aglaonema, 295 Agrobacterium, 267 A. radiobacter, 354, 734 A. radiobacter strain K84, 479

A. rhizogenes, 735 A. tumefaciens, 4, 353–354, 477, 520, 573, 760 Agrobacterium-mediated transformation, 609 Agrocin, 84, 479 Airborne conidia, 884 Airborne inoculum, 880 Air circulation, 510 Albugo bliti, 396 Albugo tragopogonis, 818–820 Alcaligenes sp., 261 Alcohols, 209 Aldehydes, 212 Aldicarb, 180 Alfalfa mosaic virus, 523, 685, 1004 Alfamovirus, 523 Allyl isothiocyanate, 181 Alphaflexiviridae, 527, 1174–1175 AlSO4, 245 Alstroemeria peruviana, 272 Alternaria sp., 257, 258, 262, 269, 443, 790–791, 982, 1000–1001, 1023–1025, 1086, 1322 A. alternata, 257, 262, 381, 425, 443, 534–536, 563–564, 639, 759 A. axiaeriisporifera, 572 A. carthami, 258 A. celosiicola, 381 A. dianthi, 258, 321–322, 572 A. dianthicola, 258, 322–323, 572 A. gomphrenae, 381 A. gypsophillae, 563, 564 A. helianthi, 790 A. juxtiseptata, 563, 564 A. subelliptica, 563, 564

# Springer International Publishing AG 2018 R.J. McGovern, W.H. Elmer (eds.), Handbook of Florists' Crops Diseases, Handbook of Plant Disease Management, https://doi.org/10.1007/978-3-319-39670-5

1339

1340 Alternaria sp. (cont.) A. tenuissima, 443 A. zinniae, 790, 841–843 Alternaria blight, 321–322, 841–843 Alternaria flower blight, 322–323 Alternaria leaf spot, 425, 534–536, 670–671 Alternaria leaf spot and blight, 381–382, 443–444, 790 Aluminum, 248 strips, 1304 sulfate, 818 tris, 459 tris (O-ethyl phosphonate), 1241 Amblyseius swirskii, 79 Amendments, 127 Ametoctradin + dimethomorph, 459 Ammoniacal sources of N, 1108 Ammonium compound, 262 Ammonium fertilization, 1115 Ammonium-N, 1295 Ampelomyces, 267 Amphobotrys blight, 1025–1026 Amphobotrys ricini, 1075–1076 Anaerobes, 259 Anaerobic environment, 1118 Anaerobic soil disinfestation, 192 Angular spots on the leaves, 861–862 Animal and Plant Health Inspection Service (APHIS), 6 Antagonistic microorganisms, 128 Antagonistic plants, 1094 Antheridium, 513, 755 Anther smut, 320–321 Antherospora vaillantii, 1324 Anthracnose, 123, 286, 426, 444–445, 674, 745, 996, 1026 Anthracnose-resistant anthurium, 90 Anthurium, 90–91, 259 A. andraeanum, 301 A. andreanum, 259 A. antioquiense, 301 Anthurium cultivars, 91 Anthurium decline, 308–312 Antibiosis, 267 Antimicrobial agent, 270 Antimicrobial compounds, 270 Antimicrobial properties, 272 Antirrhinum, 750 A. majus, 272 (see also Snapdragons) Aphelenchoides, 365, 490, 656, 861–862, 1123 A. besseyi, 4 A. fragariae, 4, 365, 904, 1263 A. ritzemabosi, 489 A. subtenuis, 1195–1196, 1332

Index Aphid(s), 403, 429, 524, 1123, 1168, 1170, 1173, 1175, 1177, 1179, 1180, 1182, 1184, 1256, 1306, 1328 inoculation, 111 population, 1306 Aphid-transmitted viruses, 1170 Aphis A. craccivora, 1090 A. gossypii, 577, 607, 616, 1091, 1256, 1327 A. spiraecola, 406, 616 Aphlenchoides fragariae, 489 Apical chlorosis, 820 Apion, 801 Apothecia, 344, 528, 812 Apple mosaic virus (ApMV), 736 Arabis mosaic nepovirus, 903 Arabis mosaic virus (ArMV), 489, 526, 736, 1168, 1177, 1181, 1183, 1252, 1253, 1301, 1325 Argyranthemum frutescens, 452 Armillaria, 982 A. gallica, 705 A. luteobubalina, 705 A. mellea, 705, 716, 1166 Armillaria root rot, 704 ArMV, see Arabis mosaic virus (ArMV) Arthrobacter sp., 334 Artificial substrates, 125 Ascochyta A. aquilegiae, 509–510 A. asteis, 426 A. chrysanthemi, 445 A. tulipae, 1322 Ascochyta leaf spot, 426 Ascochyta ray blight, 445–446 Ascomycete, 528 Ascospores, 344, 506, 812 Asiatic hybrid lily, 103 Aspergillus, 466 A. niger, 1075, 1322 A. parasiticus, 1075 Aspergillus tuber rot, 1075 Aster yellows, 355–356, 426–427, 471–472, 520–521, 575, 824, 854–855, 1299–1300 Asymptomatic. diagnosis, 1121 Athelia rolfsii, 343–344, 426, 1082–1085, 1323 Autoecious fungi, 730 Autoecious rust, 1296 Availability of Mn, 244 Avsunviroidae, 483 Axillary buds, 326 Azotobacter, 586 Azoxystrobin, 268, 392, 444, 461, 462, 470, 587, 594, 934, 936, 990, 1243

Index B Bacillus sp., 261, 267, 586, 596, 728, 795, 1148, 1325 B. amyloliquifaciens, 937 B. atrophaeus, 1295 B. subtilis, 261, 268, 298, 325, 448, 462, 637, 937, 990, 1105 B. subtilis QST 713, 385, 587 B. subtilis var. amyloliquefaciens, 388 Bacteria, 471–480, 518 Bacterial black spot, 429 Bacterial blight, 3, 90, 92, 284, 777–781 Bacterial blight and soft rot, 474–475 Bacterial canker, 1054–1055 Bacterial fasciation/leafy gall, 475–476 Bacterial leaf spot, 479, 548–549, 820, 852–853, 1001–1002 blight, 1087 bud blight, 476–477 of carnation, 348–349 Bacterial occlusions, 260 Bacterial rot, 258, 1167 Bacterial slow wilt of carnation, 349–351 Bacterial soft rot, 982, 1014–1015, 1088, 1324–1325 Bacterial spot, 898–899 Bacterial streak, 1167 Bacterial stunt of carnation, 349–351 Bacterial wilt, 284, 428–429, 479 Bacterial wilt and stem cracking of carnation, 351–353 Bacteria scab, 1298–1299 Bactericides antibiotics, 159 biocides, 159 inorganics, 158–159 Bacterium tumefaciens, 353–354 Barriers, 203 Basal rot, 104, 121, 328–330, 564–566, 1132, 1155, 1169, 1173, 1191, 1193, 1194, 1198, 1207, 1208, 1210, 1214 Basal stem segment, 260 Basidiophora entospora, 426 Basidiospores, 449 Batcheleromyces, 695 B biotype, 6 Bean yellow mosaic virus (BYMV), 615, 652, 1300–1304 Bearded iris mosaic virus, 1301 Beet curly top virus, 404, 526 Begomovirus, 613 Begonia(s), 892 Begonia  semperflorens-cultorum, 260 Belonolaimus, 830

1341 Bemisia tabaci, 6, 614 Benalaxyl, 591 Bench design, 124 Beneficial nutrients, 239 Benomyl, 421, 423, 977 Benzimidazole(s), 262, 1291, 1292, 1295 fungicides, 1234 resistance to, 587 Benzovindiflupyr, 461 Betaflexiviridae, 483–484, 486, 526, 1175 Bicarbonate(s), 448, 462, 1105 Bicarbonate formulations, 587, 588 Bidens mottle virus (BiMoV), 433, 826 Bioantagonists, 1041 Biocide, 270, 1133, 1136, 1138, 1140, 1159, 1186, 1194, 1195, 1208, 1210, 1214 Biocontrol, 267 Biocontrol products, 222 Biofumigation, 181, 813, 818 Biological(s), 138, 139, 142, 158 agents, 267 control, 77, 432, 592, 596, 600, 601, 609, 893, 1037, 1059, 1136, 1141, 1148, 1159, 1193 control agents, 79, 80, 877 Biome, 19 Biopesticide(s), 139, 267, 385 Biorational material, 1037 Biorend ®, 1295 Biotechnological tools, 89 Biotrophic prokaryotes, 398 Biotypes, 1293 Bipolaris sp., 726, 978, 1086 B. cactivora, 978 B. setariae, 456, 758 Bipolaris blight, 978 Bitertanol, 286 Black leaf spot, 518 Black leg, 519 Black root rot, 1050–1054 Black rot, 634–637 Black sclerotia, 698 Black slime disease, 1165 Black spot, 105, 510, 717 Black spot resistance, 106 Black sprouts, 1274 BLAST search, 30 Blight, 505–507, 746–748 Blossom blight, 323–326, 781–783 Blue mold, 1319 Bordeaux mixture, 138, 262 Boron, 245 Boscalid, 447, 587, 930, 1244 Botryosphaeria, 696–697 Botryosporium sp., 1086

1342 Botryotinia, 1143 B. fuckeliana, 791 B. narcissicola, 1155 B. polyblastis, 1143 Botrytis, 3, 264, 726, 1026–1030, 1143 B. cinerea, 4, 98, 255, 262, 264, 266, 268, 323–326, 382–385, 420–422, 446, 536–538, 564, 638, 719–720, 746–747, 791–793, 843–844, 893, 912–913, 945–947, 992–994, 1026–1030, 1075–1076, 1103, 1104, 1155, 1158, 1233, 1291, 1322, 1323 B. elliptica, 256, 1233 B. narcissicola, 1149, 1151, 1155, 1167 B. paeoniae, 665 B. parasitica, 1322 B. polyblastis, 1143, 1144 B. sphaerosperma, 1233 B. squamosa, 1233 B. tulipae, 109, 1314, 1322, 1323 Botrytis blight, 323–326, 382, 446–448, 505, 536–538, 585–587, 697–698, 745, 791, 843–844, 893, 912–913, 1076, 1232, 1290–1291 Botrytis blight/gray mold, 420–422, 1075–1076 Botrytis gray mold, 564, 665–667 Bradysia spp., 593, 1110 Branch-inducing phytoplasma, 1059–1060 Branch rot, 328–330 Brassica cover crops, 813, 818 Brassicaceae, 768 Breeding, 88 Brevicoryne brassicae, 616 Bright colored clothing, 223 Broad bean wilt virus (BBWV), 526, 615, 903, 1181, 1301 Bromoviridae, 429, 484–486, 523, 606, 855–856, 1122, 1175–1176 Brunneosphaerella, 695 Buckskin rot, 1249 Bud death, 1203 drop, 938 rot, 323–326 split, 1273 Buffer zones, 176 Bulb basal rot, 102 Bulb mite, 1133, 1136, 1158, 1199, 1208, 1299 Bulb-scale mite, 1136, 1156, 1189, 1202, 1208, 1210 Bullhead, 1203

Index Bunyaviridae, 404, 429–433, 486–489, 606–613, 760, 856, 1120 Burkholderia B. andropogonis, 348–349, 1325 B. caryophylli, 94, 349, 351–353, 363, 604 B. cepacia, 1045, 1295 B. gladioli, 4, 650–651, 1325 Burkholderia wilt (Burkholderia caryophylli), 351–353, 604 BYMV, see Bean yellow mosaic virus (BYMV) C CaCl2, 243 Caladium, 92, 168, 180 Caladium virus X (CalVX), 1089–1090 Calcium, 243 deficiency, 1029 nitrate, 720 sulfate, 266 Calendula, 244 Calendula officinalis, 260 Calibration, 144 Calla lily, 190, 262 Callistephus chinensis chlorosis virus (CCCV), 433 Callistephus chinensis (L.) Nees, 420 Camarosporium eriogonii, 426 Camarosporium leaf spot, 426 Candida, 267 Candidatus, 1251 Phytoplasma sp., 3, 651 Phytoplasma asteris, 355–356, 398, 471, 520–521, 854–855, 899 (see also Aster yellows) Phytoplasma aurantifolia, 471 Phytoplasma solani, 471 Canker, 351, 1023 Ca(NO3)2, 243 Capitphorus elaegni, 827 Capsicum chlorosis virus (CaCV), 652 Captafol, 700 Captan, 461, 791, 1241 Carbendazim, 262, 421, 423 Carlavirus, 358, 526, 1175, 1256, 1326 Carmovirus, 358–359, 652, 1185 Carnation(s), 95, 189, 261 bacterial and phytoplasma diseases, 348–356 bacterial wilt, 94 fungal and fungus-like diseases, 320–348 nematode diseases, 362–365 viral diseases, 356–362

Index Carnation etched ring virus (CERV), 356–357 Carnation Italian ringspot virus (CIRV), 357 Carnation latent virus (CLV), 358, 1175 Carnation mottle carmovirus, 902 Carnation mottle virus (CarMV), 357–359, 361, 615, 652 Carnation necrotic fleck virus (CNFV), 360 Carnation ringspot virus (CRV), 360–361 Carnation vein mottle virus (CVMoV), 361–362 Carnation yellow fleck virus (CYFV), 362 Carotovorum, 259 Carthamus tinctorius L., 258 Caryophyllaceae, 320, 356, 358, 360, 563 Caryophylloseptoria spergulae, 345–346 CaSO4, 243 Cations, 51 Cattleya, 636 Caulimoviridae, 856 Caulimovirus, 356–357 CChMVd, see Chrysanthemum chlorotic mottle viroid (CChMVd) Celery calico, 523 Celosia argentea, 386 Celosia mosaic virus, 402–406 Ceratobium mosaic virus (CerMV), 652 Cercospora spp., 645, 726, 982, 1248 C. antirrhini, 747–748 C. chrysanthemi, 456 C. delphinii, 510 C. dianthi, 348 C. eustomae (see Pseudocercospora eustomatis (Cercospora eustomae)) C. hydrangeae, 990–992 C. zinniae, 844–845 Cercospora blight, 747–748 Cercospora leaf spot, 385, 587–588, 844–845, 1008 Cercosporella sp., 1248, 1322 Certification, 185, 961 Chaetomium, 586 Chamelaucium axillare, 267 Chamelaucium floriferum, 267 Chamelaucium uncinatum, 256, 267 Charcoal rot, 386, 387 Chasmothecia, 461, 727, 1035 Chemical(s), 139, 142, 143 control, 263, 1042 toxicity, 1273 Chemigation, 601 Chenopodium quinoa, 1257 Cherry leaf roll virus, 526, 1003 China, 5

1343 China aster diseases bacterial and phytoplasma diseases, 426–429 fungal and fungus like diseases, 420–426 nematode diseases, 433 viral diseases, 429–433 China aster seed, 421 Chinese Narcissus Potyvirus, 1188 Chitosan, 271 Chlamydospore(s), 337, 452, 458, 507, 753, 875, 1051, 1107, 1293 Chloride, 244 Chloride salts, 1108 Chlorine, 244 containing compound, 266 compounds, 272–273, 878 dioxide, 272, 1139, 1194, 1246 releasing disinfestant, 204 Chlorobenzenes, 262 Chloropicrin, 178, 423, 596, 601 Chlorothalonil, 262, 444, 445, 447, 461, 469, 586–588, 748, 791, 930, 934, 936, 977, 1105, 1109, 1235, 1237, 1291, 1292 Chlorothalonil thiophanate methyl, 587 Chlorotic spots, 357, 1090, 1091 Choanephora cucurbitarum, 1030, 1086 Chocolate spot, 1199 Christmas cactus, 976 Chronic testing, 141 Chrysanthemum(s), 96, 260–262, 424, 430, 431, 442 bacterial and phytoplasma diseases, 471 brown rust, 448–449 flower types, 441 fungal and fungus-like diseases, 443–471 mild mosaic virus, 484 nematode diseases, 489–491 phloem necrosis, 472–474 virus and viroid diseases, 479 white rust, 449–451 Chrysanthemum x morifolium, 441 Chrysanthemum chlorotic mottle viroid (CChMVd), 483 Chrysanthemum stem necrosis virus (CSNV), 429–431, 486, 487, 608 Chrysanthemum stunt viroid (CSVd), 96, 479 Chrysanthemum virus B (CVB), 483 Cicadellidae family, 605 Cichoracearum, 426 Ciculifer, 404 Cisgenic black spot resistance, 107 Cissus rhombifolia, 1103 Citric acid, 272, 273

1344 Cladosporium spp., 726, 1248 C. cladosporioides, 269 C. cyclaminis, 1113 C. echinulatum, 327–328 C. herbarum complex, 327 Cleistothecia, 390, 753 Climate change, 1133, 1171 Climate management, 122 Clonostachys rosea, 469 Closteroviridae, 857 Closterovirus, 360, 362, 652 Clover yellow mosaic potexvirus (ClYMV), 900 Clover yellow vein virus (ClYVV), 652, 1301 CMV, see Cucumber mosaic virus (CMV) Cochliobolus setariae, 456 Cockscomb, 380 Coleosporium C. asterum, 426 C. helianthi, 808–810 C. tussilaginis, 1166 Coleroa, 695 Coleus, 912 Coleus blumei viroids (CbVd), 922 Coleus vein necrosis virus (CVNV), 921 Collar rot, 336–337, 567–569 Colletotrichum sp., 699, 1086 C. antirrhini, 745 C. capsici, 396, 1244 C. dematium, 1244, 1322 C. destructivum, 745 C. dianthi, 347 C. fioriniae, 1322 C. gloeosporioides, 288–290, 426, 646–647, 745, 996–997, 1101, 1244 C. lilii, 1244 C. spaethianum, 1244 C. truncatum, 1244 Colletotrichum Blight (Colletotrichum acutatum), 588–589 Colletotrichum tip dieback, 699 Colmanara mosaic virus (CoIMV), 652 Colocasia, 295 Colombia, 4 Colombian datura virus (CDV), 652 Coloradoa rufomaculata, 484 Commercial biocontrol products, 813 Common yellow leaf spot, 345–346 Comoviridae, 857–858 Competition, 267 Competitive saprophytic ability, 1114 Complete resistance, 105 Compost, 132, 175, 1045

Index Computerized systems, 122 Conidia, 258, 323, 335, 383, 514, 1101 Conidial concentration, 1035 Conidial sporulation, 881 Coniothyrium, 267, 813 C. minitans, 469 C. wernsdorffiae, 720–721 Consolida, 504 Containers, 217 Contaminated media, 878 Copper, 245, 932 Copper-based dips, 262 Copper-based products, 1235 Copper hydroxide, 936 Copper oxychloride, 936, 1291 Copper sulfate, 936, 1118 Copper sulfate pentahydrate, 444, 587 Corms and cormels, 1290 Corrosiveness, 217 Corticium solani, 1323 Corticium vagum, 1323 Corynebacterium fascians, 354–355, 475, 1249, 1325 Corynebacterium flaccumfaciens pv. oortii, 1325 Corynespora, 1031 C. cassiicola, 997–998 Cottony rot, 768–769 Cover crops, 193 Covering material, 121 Cowpea aphid, 1090 Cowpea mosaic virus, 1301 Cracking, 351 Cracks, 350 Criconemella, 188 Criconemella curvata, 364 Cristulariella moricola, 1076–1077 Critical concentration, 45 Critical production points, 203 Crop losses, 2, 5 Crop rotation, 387, 408, 1234 Cross protection, 606 Crotalaria juncea, 1094 Crotalaria spectabilis, 1094 Crown and root gall disease of Gypsophila, 574–575 Crown canker, 875 Crown gall, 353–354, 477–479, 520, 573, 734, 760 Crown rot, 509–510, 564–566, 1043–1046, 1076, 1166 Cryptocline cyclaminis, 1101 Cryptosporella umbrina, 721

Index CSNV, see Chrysanthemum stem necrosis virus (CSNV) CSVd, see Chrysanthemum stunt viroid (CSVd) Cucumber mosaic virus (CMV), 362, 399–403, 429, 485, 523–524, 549, 576–577, 606, 652, 685, 761, 855, 900, 919, 1122–1123, 1175, 1177, 1252, 1254–1255, 1304–1306, 1325 Cucumovirus, 362, 429, 485, 523–524, 576–577, 652, 1122, 1175, 1254, 1326 Cull piles, 594 Cultivar selection, 1142, 1159 Cultivation practices, 696 Cultural factors, 1152 Cultural practices, 124, 1137, 1145, 1148, 1159, 1164, 1165, 1193 Culture-indexed stock, 397 Culture indexing, 347, 943 Culture media, 25 Cup fungus, 528 Curative activity, 143 Curtobacterium, 1054–1055 Curtobacterium flaccumfaciens pv. oortii, 1325 Curtovirus, 404, 526 Curvularia sp., 1323 Curvularia eragrostidis, 639 Curvularia leaf spot, 1292 Cuscuta sp., 762 C. epithymum, 473 C. subinclusa, 398 Cut flowers, 2, 4, 5, 168, 176, 193, 255, 260, 664 Cutting propagation, 880, 884 Cutting rots, 328–330, 723–724 Cutting tools, 594 CVB, see Chrysanthemum virus B (CVB) Cyathia, 1023 Cyazofamid, 392, 459, 934, 936, 1241 Cycas necrotic stunt virus, 1301 Cyclamen persicum, 256, 1100, 1108 Cyclamen fatrense, 1114 Cyclamineus, 1211 Cylindrocarpon, 1236 Cylindrocladiella peruviana, 1086 Cylindrocladium spp., 874, 884 C. scoparium, 723, 879 Cylindrocladium blight and wilt, 879 Cylindrocopturus spp., 801 Cylindrosporium chrysanthemi, 456 Cymbidium ringspot virus (CymRSV), 652 Cymoxanil + dithianon, 591 Cyproconazole, 800, 1297 Cyprodinil, 447, 469

1345 Cyprodinil+fludioxonil, 444 Cyrtanthus elatus Virus A, 1188 Cyst, 512, 1123–1124 Cyst nematodes, 1124 D Daffodil bacterial and phytoplasma diseases, 1167–1168 brown ring symptoms, 1189 bulb rot, 1134, 1153 bulb-scale mite, 1190 bulb with sclerotia, 1158 bullhead, 1203 Carlton, 1204 chocolate spot, 1199 with curled leaves, 1156 damage due to HWT, 1211, 1212 dead buds, 1202 flower bud death, 1202 foliage senescence, 1144 foliar symptoms, 1168 fungal diseases, 1132–1167 grassiness in, 1201 herbicide damage, 1207 hot-water treatment, 1207 leaf scorch, 1147, 1148 leaf symptoms, 1144 lesions of white mould, 1162 melting, 1204 neck rot, 1150, 1151 nematode diseases, 1187–1199 Penicillium sp., 1154 physiological disorders, 1199 reddish leaf-tip lesions, 1146 Rhizopus soft rot, 1161 rust, 1200 stem nematodes, 1191, 1192 tepals due to smoulder, 1157 virus diseases, 1168 yellow stripes and chlorosis, 1170 Dagger nematodes, 740, 1326 Dahlia, 257, 259, 260 D. variabilis, 260 Dalotia coriaria, 79 Damping off, 121, 777 pathogens, 877 of seedlings, 1239 Dasheen mosaic virus (DsMV), 652, 1090–1091 Daylilies, 247 Dazomet, 172, 592, 596

1346 Decay, 568 Decision-support system, 1159, 1164 Deep plowing, 813 Defence response, 268–269 Deficiency, 42, 45, 49, 50, 52, 54, 60, 62, 63 Degenerate primers, 30 Degradation, 140, 142 Delphinium, 195, 504 Delphinium ringspot, 522 Delphinium vein clearing virus, 527 Demand load, 214 Dendranthema grandiflora, 260 Dendrobium, 271, 636 Dendrobium mosaic virus (DeMV), 652 Dendrobium ringspot virus (DRV), 652 Dendrobium vein necrosis virus (DVNV), 652 Dianthovirus, 360–361 Dianthus, 195 D. capitatus ssp. andrzejowskianus, 94 D. caryophyllus, 260, 264, 272, 423 (see also Carnation) Diaporthe spp., 802 D. gardeniae, 928–930 Diaporthe blight, 506–507 DICA, 272, 273 Dicarboximides, 262, 1234 resistance to, 587 Dichlorofluanid, 586 Dichloroisocyanuric acid, 272 1,3-Dichloropropene, 178, 423 Dichloropropene + etridiazole, 1321 Dichotomophthora, 982 Dickeya sp., 254, 259, 428–429 D. chrysanthemi, 3, 474, 649, 1014, 1015, 1056–1057 D. dadantii, 258, 1088–1089, 1324 D. dianthicola, 349–351 D. dieffenbachiae, 649 Didecyl dimethyl ammonium chloride, 1246 Dieback, 874 Dieback and wilt, 875 Dieffenbachia maculata, 260 Difenoconazole, 800, 805, 1291, 1297 Dimethomorph, 459, 934, 1241 Dimethyl disulfide (DMDS), 180, 1094 Dinitroaniline, 806 Dip, 1139, 1141, 1145, 1152, 1166, 1194, 1196, 1210 Diplocarpon rosae, 105, 717–719 Diplodia seriata, 696 Diplodina delphinii, 509–510

Index Diplodina disease, 509–510 Dipping, 1135, 1136, 1138, 1167 Dipsticks, 27 Discarded plant material, 227 Discovery, 139–140 Disease(s), 70, 75, 77 Disease-forecasting, 1159, 1163 Disease-free cuttings, 1048 Disease-free stock plants, 225 Disease impact direct, 5 indirect, 5–6 Disease management, 139, 158, 243, 247, 248 Disease ratings, 102 Disease-resistant cultivars, 88 Disease triangle, 202 Disinfectant, 216, 1137, 1139, 1194, 1210 Disinfestant(s), 203, 216, 1053 Disorders of flower development, 1203–1207 Dithiocarbamates, 388 Ditylenchus, 1123, 1187 D. destructor, 1332 D. dipsaci, 365, 528, 1136, 1187, 1207, 1208, 1264, 1329 Diurus virus Y (DVY), 652 D-limonene, 421 Dodder, 473, 762 Dominant gene, 108 Double-antibody sandwich ELISA (DAS-ELISA), 26 Double-tent, 174 Downy growth, 749 Downy mildew, 107, 326–327, 426, 516–518, 589–591, 745, 769–772, 796–797, 914–916 Drechslera sp., 978, 1323 Drip irrigation, 586 Drying, 219, 1136, 1137, 1141, 1152, 1159, 1161 Dry-scale rot, 1166 E Ebb-and-flood, 1025, 1039 Echeveria, 1009 Ecotoxicology, 140, 142 Ectoparasitism, 657 Ecuador, 4 Edema, 968 Efficacy, 140–141 Eggs, 1124

Index Electrical conductivity (EC), 64, 127 Elicitors, 268–269 Elsinoe, 700 E. rosarum, 730 Emilia fosbergii, 424 Endoconidia, 1051, 1115 Endoparasitism, 656 Endophytic, 19 Enterobacter agglomerans, 261 Entomopathogenic nematodes, 80 Entyloma E. polysporum, 799 E. winteri, 518 E. wyomingense, 518 Environmental parameters, 120 Environmental stress, 879 Environment control, 325 Enzyme-linked immunosorbent assays (ELISA), 26, 1300, 1303 Epicoccum sp., 268, 269 Epiphytic, 19 Epipremnum, 295 Epoxiconazole, 1297 Equipment, 218 Erica gracilis, 1103 Erwinia, 169, 822 E. carotovora, 474, 1088–1089, 1324 E. caratovora subsp. atroseptica, 519 E. carotovora subsp. carotovora, 1167 E. chrysanthemi, 396–397, 428, 474, 1088–1089, 1324 E. chrysanthemi pv. dianthicola, 349–351 Erysiphe E. aquilegiae var. ranunculi, 514 E. buhrii, 3, 335–336, 566–567 E. celosiae, 389–391 E. cichoracearum, 461, 753–754, 804, 847–849 E. poeltii, 989 E. polygoni, 101, 514, 931–932, 989 E. sedi, 1011 Erythonium grandiflorum var. pallidium, 1233 Escherichia, 261 Essential element, 42 Ethiopia, 4 Ethylene, 264, 271, 1104 Etridiazole, 392, 637, 934, 936, 1321 Eumerus, 1136 Euphorbia pulcherrima, 256 European Plant Protection Organization (EPPO), 6

1347 Eustoma grandiflorum, 351 Exclusion, 605 Exobasidium spp., 885 F Fabavirus, 526, 1181 Fairy-ring leaf spot, 327–328 Famoxadone + cymoxanil, 1241 Fasciation, 354–355, 573–574, 763 Feathering, 1090 Feathery tip rot, 1249 Fenamidone, 392, 459, 753, 936, 1241 Fenaminosulf, 1110, 1321 Fenhexamid, 447, 587, 1105, 1235 Fenheximid-resistant Botrytis pseudocinerea, 666 Fertilization tube, 513 Fertilizer(s), 243, 244, 1135, 1214 Filiation, 224 Fine mesh screening, 484, 605, 606, 608 Fire, 1143–1146 First juvenile stage (J1), 762 Fishy odor, 1117 Flavobacterium, 261 Flecks, 360 Floater/sinker method, 1138 Flood-floor, 1025 Flooding, 818 Flooding of fields, 813 Floriculture, 131 Florists crops, loss estimation, 3 Flower blight, 256 Flower bud abortion, 1272 Flower bud setting, 876 Flower distortion, 360 Flower spikes, 1290, 1297, 1298, 1300 Fluazinam, 586, 1244 Fludioxonil, 292, 394, 444, 447, 462, 466, 469, 587, 596, 748, 933, 1105, 1116, 1235, 1237, 1295 Fludioxonil + cyprodinil, 1244 Fluopicolide, 459, 753, 934, 936, 1241 Fluoxastobin, 444 Fluoxastrobin + myclobutanil, 1241 Flutolanil, 292, 470, 1243 Fod, 331 Foliar nematode (Aphelenchoides ritzemabosi), 433, 489–490, 656, 686, 904–905, 1123, 1124, 1263 Folpet, 262

1348 Food security, 5 Foot baths, 221 Foot rot, 336–337, 774–776 Forced into flower, 874 Formaldehyde, 596, 1136, 1140, 1152, 1165, 1194, 1195, 1209, 1210, 1213 Formalin, 1136 Fosetyl-aluminum, 591, 636, 936 Fourth stage juvenile (J4), 762 Frankliniella, 405, 1176 F. bispinosa, 487, 610 F. cephalica, 487, 610 F. fusca, 487, 610, 920 F. gemina, 610 F. intonsa, 610, 920 F. occidentalis, 430–431, 487, 607, 610, 920 (see also Western flower thrips) F. schultzei, 487, 607, 610 Freesia, 255, 259, 265, 270 F. hybrida, 256 Frost, 1205 Fuchsia hybrida, 256 Fulgoroidea, 605 Fumigants, 176, 365, 1172, 1195 Fumigation, 508, 816, 818, 1196 Fungi, 70, 73, 674 Fungicide(s), 222, 262, 506, 1031, 1132, 1136, 1138, 1141, 1143, 1145, 1149, 1152, 1153, 1155, 1159, 1160, 1164, 1186, 1194, 1196, 1208, 1210, 1214 benzimidazoles, 156 biofungicides, 158 dicarboximides, 156 dithiocarbamates, 155 FRAC, 145 inorganics, 146–155 phenylamides, 157 pthalimides, 156 resistance, 384, 667, 1141, 1155 sterol inhibitors, 156 strobilurins, 157–158 substituted aromatics, 155 triazoles, 156–157 Fungicide Resistance Action Committee (FRAC), 422, 543, 731, 930–934, 936, 982, 1023, 1025, 1029, 1031, 1036, 1041, 1042, 1045, 1046, 1050, 1059 classification by mode of action, 145–146 group 1 fungicides, 932 group 2 fungicides, 931 group 3 fungicides, 719, 720, 728, 731, 930 group 4 fungicides, 725 group 5 fungicides, 720, 722, 930, 931

Index group 7 fungicides, 930 group 11 fungicides, 933 group 12, 724, 931, 933 group 14, 934 group 28, 934 group 33, 934 group 44, 934 Fungus gnats (Bradysia spp.), 73–74, 79, 593, 600, 1053, 1110, 1118 Fungus-like microorganisms, 338 Furalaxyl, 1110, 1321 Fusarium, 169, 179, 180, 268, 507–509, 642, 723, 756, 846, 1032, 1132, 1293–1296 F. acuminatum, 798 F. avenaceum, 4, 328–330, 592, 1323 F. bulbigenum, 1132, 1149 F. celosiae sp. nov, 387 F. culmorum, 328–330, 564, 566 F. equiseti, 798 F. foetens, 892 F. graminearum, 328–330, 564–566 F. lateritium f. sp. celosiae, 387–388 F. moniliforme, 564–566 F. oxysporum, 178, 642, 706–707, 798, 977, 1086, 1167, 1323 F. oxysporum f. sp. callistephi, 422–423 F. oxysporum f. sp. cattleyae, 643 F. oxysporum f. sp. chrysanthemi, 452, 538–540 F. oxysporum f. sp. cyclaminis, 1105 F. oxysporum f. sp. dianthi, 95, 330, 337, 340, 347, 363 F. oxysporum f. sp. dianthus, 423 F. oxyxporum f. sp. eustomae, 594, 595 F. oxysporum f. sp. gladioli, 3, 100, 242, 1293 F. oxysporum f. sp. lilii, 1235–1238 F. oxysporum f. sp. mathioli, 772–774 F. oxysporum f. sp. narcissi, 104, 1132–1143, 1149–1153, 1192, 1207 F. oxysporum f. sp. tracheiphilum, 452, 538–540 F. oxysporum f. sp. tulipae, 109, 1316 F. poae, 347, 1323 F. proliferatum, 328–330, 564–566, 642 F. roseum, 329, 1323 F. solani, 3, 92, 451, 540–541, 591, 642, 1032, 1077–1079, 1296 F. sporotrichioides, 347, 798 F. subglutinans, 642, 1296 F. verticillioides, 328–330, 564–566, 798 Fusarium basal rot, 1316 Fusarium bulb rot, 1316–1319

Index Fusarium corm rot, 1293–1296 Fusarium crown and root rot, 124, 591–592 Fusarium crown and stem rot, 592–594 Fusarium root rot, 422–424, 846 Fusarium Screen Analysis Software, 110 Fusarium stem rot, 328–330, 451, 564–566 Fusarium tuber rot, 93, 1077 Fusarium wilt, 95, 128, 330–335, 422, 451–455, 538–540, 594–596, 707, 772–774, 892–893, 1105–1109 Fusarium yellowing and corm rot, 100–101

G GA3, 266 Galling, 1092 Galls, 354, 363, 574, 734 Gamma irradiation, 265 Gardenia canker, 928–930 Gardenia jasminoides, 928 Geminiviridae, 404, 526, 615–617, 858 GenBank ®, 29 General combining ability, 99 Genetic abnormalities, 888 Genetic diversity, 1058 Genetic engineering, 613 Geraldton waxflower, 268, 269 Geranium, 239, 257, 265, 942 Gerbera, 98, 255, 257, 262, 265, 424, 456 Alternaria leaf spot, 534–536 bacterial leaf spot, 548–549 Botrytis blight, 536–538 cultivars, 99 daisy, 534 flowers, 272 Fusarium root and stem rot, 540–541 Fusarium wilt, 538–540 powdery mildew, 544–547 Phytophthora crown rot, 541–543 Pythium root rot, 547–548 viral disease, 549–552 Gerbera jamesonii, 256, 272, 452, 534 Gibberella avenacea, 592, 1323 Gibberella baccata, 388 Gibberella zeae, 330, 566 Gladiolus bacterial and phytoplasma diseases, 1298–1300 breeding, 100 fungal and fungus-like diseases, 1290–1298 rust, 1296–1297 viral diseases, 1300–1306

1349 Gliocladium, 466, 813 G. catenulatum, 268, 385 G. roseum, 268, 469 Globalization, 1040 Globisporangium G. intermedium, 1323 G. irregulare, 1323 G. splendens, 895 G. ultimum, 895, 1323 Globodera pallida, 182 Globodera rostochiensis, 182 Gloeosporium sp., 1086 G. thuemenii, 1323 G. thumenii f. tulipae, 1323 Glomerella cingulata, 426, 646 Glomus intraradices, 592, 601 Golovinomyces, 426 G. cichoracearum, 461, 804, 847–849 G. ichoracearum, 544–547 G. orontii, 753–754, 988, 1012 Grassiness, 1201 Grassy tuber, 1085–1086 Gray mold, 323–326, 382, 697, 843–844, 1027 Greasy canker, 1055–1056 Green island, 724 Green island mosaic, 1015 Green peach aphid, 75–76, 1090 Greenhouse crops, 120 Grey bulb rot, 1166, 1321 Grey mold, 98, 791–793, 1158 Groundnut ringspot virus (GRSV), 429, 431, 608 Growth distortions, 356, 575 Guignardia spp., 644 Gummosis, 1317 Gypsophila bacterial and phytoplasma diseases, 573–576 fungal and fungus-like diseases, 563–572 nematode diseases, 577–578 viral diseases, 576–577 Gypsophilaelegans, 563 Gypsophila gall, 574–575 Gypsophila paniculata, 352, 563, 577 H Habenaria mosaic virus (HaMV), 652 Haematonectria haematococca, 1077–1079 Hail, 1205 Hairy root, 735 Haplolaimus, 1196 Haplotrichum curtisii, 1323

1350 Hardwood bark, 131 Harpin protein, 636, 640 Hatiora, 977, 978 Haustoria, 727, 753, 886 Haustorium, 1035 Heating, 121, 173 Heat units, 187 Helicotylenchus, 188, 830 H. varicaudatus, 364 Heliothis, 806 Helminthosporium sp., 978, 1087 Hemiptera, 403, 404, 406, 606 Hemlock, 994 Herbicides, 1207 Heritability, 103 Heterodera, 363 H. daverti, 362 H. radicicola, 1123 H. schachtii, 578 H. trifolii, 362, 363 Heterorhabditis spp., 80 Heterosporium spp., 1248 Heuchera sanguinea, 271 Hexaconazole, 262, 1291 Hibiscus, 257 High humidity, 1104, 1291 Highly pathogenic, 229 Hilling, 722 Hinomyces moricola, 1076–1077 Histiostoma, 1136 Honey fungus, 1166 Horizontal resistance, 106 Hose-end holders, 221 Host defense, 246, 247 Hot-water treatment (HWT), 657, 1075, 1084, 1132, 1135, 1137, 1138, 1140, 1141, 1148, 1149, 1152, 1160, 1164, 1166, 1187, 1193–1195, 1202, 1207–1213, 1262, 1265 Hot weather, 1113 Humid conditions, 1103, 1115 Humid weather, 1113 Husbandry, 1132, 1136, 1152, 1171, 1172, 1199, 1203 HWT, see Hot-water treatment (HWT) Hyacinthus, 259 Hyaloperonospora parasitica, 769–772 Hybridization breeding, 89 Hybridoma, 26 Hydrangea, 247 bacterial diseases, 1001 fungal diseases, 988–1001 viral diseases, 1002–1004

Index Hydrangea macrophylla, 101, 256, 988 Hydrangea mosaic virus, 1002 Hydrangea ringspot virus, 1003 Hydroponics, 125, 715 Hydroxyquinoline sulphate, 270 Hygiene, 1167, 1193, 1196 Hyperplastic growth, 354, 574 Hypoaspis aculeifer, 80 I ICM, see Integrated crop management (ICM) Ilarvirus, 827 Ilyonectria sp., 70, 1323 I. destructans, 1196, 1198 Immobile elements, 48 Immunity, 108 Immunodiagnostics, 26 Immunofluorescent, 27 Impatiens necrotic spot virus (INSV), 3, 71, 72, 78, 362, 486, 487, 524–525, 550, 608, 760–761, 901, 920, 983, 1017, 1121, 1301 Impermeable films, 177 Improving disease resistance, 88 Inactivate, 203 Increased plant spacing, 586 Incubation period, 747 Indian, 5 Indicator plant, 488 Infected liner plants, 875 Injecting aerated steam, 222 Inoculum reservoirs, 1122 Inorganic anion, 244 Insect(s) aphids and whiteflies, 75–77, 80 exclusion, 220 fungus gnats, 73–74, 79 shore flies, 74–75, 80 western flower thrips, 71–73, 77–79 Insecticide(s), 71, 77, 78, 80, 1171, 1176 Inspection, 1149, 1150, 1167, 1189, 1193, 1196 INSV, see Impatiens necrotic spot virus (INSV) Integrated crop management (ICM), 1136, 1193, 1214 Integrated disease management, 231, 302, 703, 1123, 1290, 1314 Integrated disease management of viruses, 606, 613, 614 Integrated management, 861 Integrated pest management (IPM), 1053 Integrated strategies, 341 Integration, 335

Index Integrative strategies, 613 International trade, 2, 5 Interspecific crosses, 101, 103 Interspecific hybridization, 89 Interspecific hybrids, 110 Ionizing irradiation, 265 IPM, see Integrated pest management (IPM) Iprodione, 262, 447, 466, 587, 588, 791, 1105, 1235, 1244, 1291 Iris, 259 Iris severe mosaic virus, 1301 Iris yellow spot virus (IYSV), 486, 487, 608 Iron deficiency, 938 Irrigation water, 1105, 1111 Isothiocyanates, 813 Itersonilia perplexans, 455 Itersonilia petal blight (Itersonilia perplexans derx), 424–425, 455–456 J Japanese hornwort mosaic virus, 1091 Jasmonic acid, 268–269 Jonquil cultivars, 1174 Jonquil daffodils, 1179 Jonquilla cultivars, 1211 J-shaped canal, 230 K K2SO4, 243 Kalanchoe, 259, 1008, 1014 Cercospora, 1008 mosaic virus, 1015 Phytophthora, 1010 Stemphylium, 1009 top-spotting virus, 1016 Tospoviruses, 1017 Kalanchoe gastonis-bonnierii, 259 Kalanchoe latent virus, 1015 KCl, 243 Kenya, 4 Kluyveronzyces fragilis, 261 KNO3, 243 Konjac mosaic virus (KoMV), 1091–1092 L Labels, 144 Larkspur, 504 Larval transmission, 74 Lasidiplodia theobromae, 696 Late blight, 123

1351 Latent expression of symptoms, 1106 Latent infections, 1028, 1257, 1293, 1317 Latent period, 256 Lateral flow devices, 27 Lathyrus odoratus, 1103 Leaf abscission, 884 Leaf and flower gall, 885 Leaf blight, 256 Leaf crinkling, 576 Leaf epinasty, 736, 1273 Leaf gall, 885 Leafhopper(s), 356, 398, 404, 471, 521, 575, 605, 825 Leafhopper feeding, 1119 Leafminer, 476 Leaf mottle, 360 Leaf scorch, 1136, 1143, 1145–1149, 1269 Leaf smut, 799–800 Leaf spot(s), 284, 321, 322, 348, 456–457, 509–510, 563–564, 644–646, 648–649, 745, 748, 750, 759, 1009, 1101, 1248 Colletotrichum gloeosporioides, 1102 diseases, 694 phyllosticta, 1109 ramularia, 1112 septoria, 1114 Leafy gall, 354 Lesion nematode, 491 Lethal dose response curve, 215 Leucanthemella, 449 Leucodendron, 705 Leucospermum, 699, 701, 706 Leveillula, 1033 Leveillula taurica (Oidiopsis taurica), 389, 597, 804 Light, 120 Light intensity, 125 Ligneous defense barriers, 246 Lilies basal rot, 1236 blue mold, 1245 Botrytis blight, 1232–1235 gray bulb rot, 1248 leaf rust, 1247–1248 leafy gall, 1249 nematode diseases, 1261–1267 southern blight/crown rot, 1242–1243 physiological disorders, 1267–1275 phytoplasma diseases, 1251 Pythium root rot, 1239–1240 Rhizoctonia stem rot, 1238–1239 scale rots, 1249 soft rot, 1246

1352 Lilies (cont.) stump rot, foot rot and stem blight, 1240–1242 viral diseases, 1252–1261 white stem rot, 1243 Lilium Asiatic hybrid, 271 Lilium L. auratum, 1231 L. bulbiferum, 1231 L. dauricum, 103, 1231 L. davidii, 1231 L. henryi, 103 L. longiflorum, 1231, 1237 L. maculatum, 1231 L. regale, 103, 256, 1231 L. speciosum, 1231 L. speciosum var. rubrum, 256 L. tigrinum, 256 Lilium Oriental hybrid, 271 Lily, 102–104 Lily mottle virus (LMoV), 1188, 1252, 1255, 1328 Lily symptomless virus (LSV), 1252, 1256, 1325 Lily virus X, 1252, 1257 Lime, 1108 Lisianthus diseases bacterial and phytoplasma diseases, 603–605 fungal and fungus like diseases, 585 nematode diseases, 619–621 viral diseases, 606 Lisianthus necrosis virus (LNV), 617, 652 Lisianthus necrotic ringspot virus (LNRV), 608 Longidorus sp., 1182, 1184, 1266 L. africanus, 1267 L. elongatus, 361 L. macrosoma, 361 Lonicera japonica, 256 Lose activity, 214 Lower leaf yellowing, 1269–1271 Lower nitrogen fertilization, 803 Low temperature injuries, 273

M Macluravirus, 1177 Macroconidia, 507 Macroconidium, 1107 Macrophomina phaseolina, 386, 793–794 Macrosiphoniella (Pyrethromyzus) sanborni, 484

Index Macrosiphum euphorbiae, 484, 1256 Macrosteles fascifrons, 427, 471 Magnesium, 243 Magnolia M. grandiflora, 260 M. macrophylla, 260 M. soulangeana, 260 M. tripetala, 260 Major and common pathogens, 231 Malaysia, 4 Malformation, 576 Malformed buds, 1274 Mancozeb, 262, 286, 445, 461, 587, 588, 640, 647, 700, 791, 930, 936, 977, 1109, 1235, 1291, 1292 Mancozeb fungicides, 1103 Mandipropamid, 459, 753, 934, 936, 1241 Manganese, 245 Mapping population, 108 Marafivirus, 1061 Marigolds (Tagetes spp.), 1198, 1262 Marker-assisted selection, 103 Maternal effects, 103 Matthiola incana, 272, 768 Mechanical inoculation, 111 Mechanically transmissible, 1328 Medium pasteurization, 887 Mefenoxam, 392, 459, 636, 753, 795, 934, 936, 1240, 1241 Mefenoxam + mancozeb, 590 Melanagromyza, 806 Meliola sp., 1087 Meloidogyne sp., 169, 180, 363, 406, 491, 577–578, 761–762, 827, 862, 1092–1094, 1123 M. arenaria, 180, 362, 363, 577, 1092 M. chitwoodi, 1265 M. fallax, 1265 M. floridensis, 1092 M. hapla, 363, 525, 740 M. hispanica, 363 M. incognita, 93, 189, 195, 362, 363, 407, 525, 577, 1092, 1094, 1265, 1266 (see also Root knot nematode (Meloidogyne incognita)) M. javanica, 362, 363, 525, 577 Melting, 1204 Meristem culture, 576, 1304 Meristem-tip culture, 358, 1061 Merodon equestris, 1136 Metabolomics, 19 Metagenomics, 19 Metalaxyl, 393, 1110, 1240, 1241, 1321

Index Metam sodium, 423, 592, 596 Metconazole, 596 Methyl bromide, 176, 178, 180, 189, 195, 423, 592 Methyl bromide + chloropicrin, 189 1-Methylcyclopropane, 264–265 Methyl isothiocyanate, 173 Methyl jasmonate, 268–269 Microbiome, 19 Microbotryum dianthorum, 320–321 Microcephalothrips abdominalis, 607 Micrococcus, 261 Microconidia, 507, 1107 Micronutrients, 238 Microsclerotia, 386, 395, 731, 758, 817, 880 Microscopy, 16, 24 Microsphaera, 894 Migratory endoparasites, 830, 1124 Migratory endoparasitic nematodes, 364 Mildew, 748–750, 753–754 downy, 516–518 powdery, 514–516 Mild mottle virus, 358 Milk, 1123 Mineral nutrition, 120 Mineral oil, 1091, 1329 Minimedusa, 1141 Mixed infection, 360 Mobile elements, 48 Mode of inheritance, 101 Moist chamber, 25 Molasses, 195 Molecular assays, 1120 Molecular cloning, 107 Molecular markers, 89 Molybdenum, 245 Monoclonal antibodies, 26 Mono-potassium phosphate (MKP), 242 Morning glory, 247 Moroccan pepper virus (MPV), 617 Mosaic, 1090, 1306 Mosaic persicae, 616 Mother blocks, 482 Moth flies (Psychoda spp.), 594 Mottling, 361, 521 MRI sensor, 1138 Mucor, 1138 Multi locus sequence analysis (MLSA), 30 Multiple pond system, 230 Muscari, 259 Mustard seed meal, 193 Mycelia, 340, 511

1353 Mycelium, 1107 Myclobutanil, 425, 444, 456, 462, 932, 1297 Mycoparasites, 339, 570 Mycorrhizae, 1103 Mycorrhizal fungi, 242 Mycosphaerella sp., 426, 695, 1248 M. dianthi, 327–328 M. ligulicola, 445 Mycosphaerella leaf spot, 695 Mycostop, 994 Myrothecium gall, 726 Myrothecium sp., 1087 M. roridum, 758, 930, 999 Myzus persicae, 357, 361, 403, 429, 484, 486, 577, 607, 827, 1090, 1256, 1326 N Nandina domestica, 1258 Na penicillin G, 474 Narcissus sp., 423, 1174 N. broussonetti, 105 N. jonquilla, 105, 1174 N. obvallaris, 1199 N. pseudonarcissus, 1174 N. tazetta, 1148, 1173 N. telamonius var. plenus, 1199 Narcissus common latent virus (NCLV), 1188 Narcissus degeneration virus (NDV), 1168, 1173, 1177 Narcissus flies, 1136, 1190, 1202 Narcissus late season yellows virus (NLSYV), 1168, 1169 Narcissus latent virus (NLV), 1168, 1177–1178, 1301 Narcissus mosaic virus (NMV), 1168, 1174 Narcissus mottling-associated virus (NMAV), 1188 Narcissus poeticus ‘Flore Pleno’, 1203 Narcissus Q virus, 1188 Narcissus silver stripe virus, 1168 Narcissus symptomless virus, 1188 Narcissus tip necrosis virus (NTNV), 1168, 1172, 1185 Narcissus white streak virus (NWSV), 1168, 1169, 1172, 1178, 1180 Narcissus white stripe-associated virus, 1188 Narcissus yellow stripe virus (NYSV), 1168, 1171, 1178–1179 Natriphene, 649 NDV, see Narcissus degeneration virus (NDV) Neck rot, 1149, 1152, 1160 Necrotic flecks, 356

1354 Necrotic lesions, 257 Necrotic spots, 1090 Necrotroph, 382 Necrovirus, 1326 Nectria haematococca, 540–541 Nectria inventa, 1323 Needle nematode (Longidorus spp.), 621, 1266 Neem, 385, 387, 408 Neem seed oil, 990 Nematicidal, 1171 Nematicide(s), 159, 528, 829, 1138, 1195 Nematode(s), 169, 180, 188, 192, 362–365, 1170, 1172, 1181, 1185, 1187 diseases, 740–741 resistance, 94 wool, 1191 Neofusicoccum, 696 Neoseiulus cucumeris, 79, 432, 609 Nepoviridae, 489 Nepovirus, 489, 526, 527, 576, 652, 1181–1184, 1253, 1326 Nerine latent virus, 1188 Nerium oleander, 273 Netherlands, 4 Next Generation Sequencing (NGS), 32 (NH4)2SO4, 474 NH4-N, 239 Nickel, 247 Niponanthemum, 449 Nitrate(s), 1108 Nitrate fertilization, 1295 Nitrification, 238 Nitrogen, 238 Nitrogen deficiency, 1114 NLV, see Narcissus latent virus (NLV) NMV, see Narcissus mosaic virus (NMV) NO3-N, 239 Nonpathogenic Fusarium oxysporum, 1108 Non-persistent way, 1256 North American Plant Protection Organization (NAPPO), 6 NTNV, see Narcissus tip necrosis virus (NTNV) Nucleorhabdovirus, 433 Nutrient(s), 1028 deficiencies, 1275 translocation, 46 Nutrition, 238 NWSV, see Narcissus white streak virus (NWSV) NYSV, see Narcissus yellow stripe virus (NYSV)

Index O Oakleaf hydrangea, 1001 Obligate parasites, 326, 335, 356 Observational skills, 18 Oidiopsis taurica, see Leveillula taurica (Oidiopsis taurica) Oidium sp., 390, 544–547, 597, 753–754, 1323 O. asteris-punicei, 426 O. begoniae, 894 O. dianthi, 335–336, 566–567 Olpidium sp., 618 O. brassicae, 1172, 1186, 1326, 1328 Oncidium, 636 Onion yellow dwarf virus (OYDV), 1179 Oogonium, 513, 755 Oomycetes, 590, 1037 Oospore(s), 74, 327, 337, 339, 458, 753, 797, 875, 1039 Ooze, 351 Oozing lesions, 1117 Ophiostoma narcissi, 1167 Orchidaceae, 634 Organic acids, 195 Organic amendments, 128 Organic and inorganic material removal, 219 Organic wastes, 131 Orius spp., 79, 433, 609 Ornamental plants Alternaria spp., 257–258 bacterial species, 258–261 Botrytis cinerea, 255–257 Ornithogalum Mosaic Virus, 1188, 1301 Orthophenylphenol, 649 Osteospermum sp., 452 Overhead irrigation, 586 Overwinters, 1109 Ovularia delphinii, 510 Oxalic acid, 511 Oxathiapiprolin, 934, 936 Oxycarenus, 709 Oxytetracline-HCl, 474 OYDV, see Onion yellow dwarf virus (OYDV) Ozone, 213 P Packaging, 264 Paecilomyces, 466 Paeonia, 664, 665 Pantoea agglomerans pv. gypsophilae, 574–575 Papaya leaf curl guangdong virus, 614

Index Paper daisy, 247 Paphiopedilum, 640 P. venustum, 640 Paraconiothyrium P. fuckelii, 723 P. minitans, 469 Parasitic nematode control, 193 Paratrichodorus sp., 830, 1187, 1260, 1327, 1332 Paratrichodorus minor, 188, 433 Paratylenchus, 364, 830 P. curvitatus, 364 P. dianthi, 362, 364 P. hamatus, 364 P. projectus, 364 Partial black spot resistance, 106 Partial resistance, 96, 100 Partial saturation, 1110 Passalora rosicola, 722 Pasteuria, 184 Pasteurization, 818 Pasteurized soil, 813 Pathogen(s), 73, 75, 77 Pathogen-free growth media, 874 Pathogen-free ornamentals, 224 Pathogen-free procedure, 943 Pathogen-free seed, 397 Pathogen spread, 6 Pathotypes, 331 Pathovar of X. campestris, 1001 PCNB, 1243, 1322 PCR, 16, 19, 28, 29–30, 351, 353, 355, 573, 575 PCR-RFLP, 30 Pea mosaic virus, see Bean yellow mosaic virus (BYMV) Peat, 131 Pecteilis mosaic Virus (PcMV), 652 Pectic enzymes, 259, 261 Pectin, 1117 Pectobacterium, 169, 259 P. atrosepticum, 519, 824 P. cacticidum, 824 P. carotovorum, 259, 474, 824, 1056–1057, 1088–1089, 1324 P. carotovorum pv. dianthicola, 349–351 P. carotovorum subsp. carotovorum, 982, 1014, 1167 P. chrysanthem, 649 Pectolytic enzymes, 261 Pelamoviroid, 482, 483 Pelargonium, 942 P. domesticum, 257

1355 P. peltatum, 257 P. scented, 257 P. zonale, 256, 1103 Pelargonium x hortorum, 256, 265 Pelargonium zonate spot virus (PZSV), 826 Pellicularia filamentosa, 393 Pellicularia rolfsii, 511 Penconazole, 805, 1291 Penicillium, 262, 1087, 1138, 1140, 1141, 1151, 1153–1155, 1245 P. albocoremium, 1245 P. aurantiogriseum, 1245, 1323 P. corymbiferum, 1245, 1319 P. cyclopium, 1245, 1323 P. gladioli, 1245 P. hirsutum, 1245, 1319 P. jensenii, 423 P. radicicola, 1319 P. tulipae, 1245, 1319 P. venetum, 1319 P. verrucosum var. cyclopium, 1323 Penicillium bulb rot, 1319 Penstemon, 752 Pentachloronitrobenzene, 466, 468, 470 Pepper ringspot virus (PepRSV), 617 Pepper veinal mottle virus (PVMV), 615 Perithecia, 330, 506 Peritrichous flagella, 259 Periwinkle, 189 Permethrin, 1329 Peronospora sp., 914–916 P. antirrhini, 4, 748–750 P. chlorae, 589–591 P. dianthi, 326–327 P. dianthicola, 326–327 P. ficariae, 516–518 P. parasitica, 769–772 P. sparsa, 107, 724–726 Peroxygen compounds, 206 Pesta, 1045 Pest dissemination, 2, 5, 6 Pestalotiopsis, 695 Petal blight, 639 Petal specking, 269 Petal spot, 322–323 Peyronellaea, 1145 Peyronellaea (Stagonospora) curtisii, 1149, 1151, 1154, 1167 Peziza repanda, 528 pH, 120, 1053 Phaius flavus, 640 Phalaenopsis, 635

1356 Phalaenopsis chlorotic spot virus (PhCSV), 652 Phenol(s), 211 Phenol metabolism, 246 Phenylalanine ammonia lyase (PAL), 265 Phenylpyrroles, 388 Phialophora asteris, 426, 817 Phialophora cinerescens, 346–347 Phialophora wilt, 426 Phialospores, 1051 Phoma black stem, 800–802 Phoma sp., 514, 998 P. caryophylli, 348 P. dianthi, 348 P. gypsophilae, 572 P. leveillei, 1167 P. macdonaldii, 800–802 P. pinoggii, 572 Phomopsis, 506, 982 P. callistephi, 426 P. gardeniae, 928–930 Phomopsis stem and leaf blight (Phomopsis sp.), 597 Phomopsis stem canker, 426, 802–803 Phosphites, 459 Phosphorus, 239 Phosphorous acid, 459, 936, 1242 Photoperiod, 125 Phragmidium mucronatum, 729–730 Phyllactinia poinsettiae, 1032–1037 Phyllocoptes fructiphilus, 738 Phyllody, 356, 397, 471, 472, 520, 575, 651, 709, 1118, 1119 Phyllosticta sp., 695, 1248 P. antirrhini, 750–752 P. celosiae, 396 P. chrysanthemi, 456 P. cyclaminis, 1109 P. delphinii, 510 P. dianthi, 348 Phyllosticta blight, 750–752 Phyllosticta leaf spot, 572 Phylogenetic trees, 30 Phymatotrichopsis omnivora, 426 Phymatotrichopsis root rot, 426 Phymatotrichum omnivorum, 514, 759 Physical management, 1136 Physiological disorders, 1199–1207 Physiological neck rot, 1150 Phytopathology, 21 Phytophthora sp., 4, 169, 175, 179, 180, 242, 243, 248, 338, 511, 634, 732,

Index 777, 794–795, 847, 875, 1041, 1167, 1235, 1323 P. cactorum, 336–337, 457, 567–569, 637, 752–753, 1241, 1323 P. capsici, 336–337, 567–569 P. chrysanthemi, 457 P. cinnamomi, 702, 933, 1240 P. citricola, 702, 1323 P. citrophthora, 702 P. cryptogea, 336–339, 426, 457, 541–543, 567–569, 702, 752–753, 1323 P. drechsleri, 388, 541–543, 567–569 P. erythroseptica var. erythroseptica, 1323 P. hedraiandra, 1323 P. megasperma, 336–337 P. multivora, 702 P. nicotianae, 4, 336–337, 388, 457, 567–569, 637–638, 702, 933, 980, 1240 P. nicotianae var. parasitica, 288, 388, 567, 569 P. niederhauserii, 1010 P. palmivora, 336–337, 541–543 P. parasitica, 248, 336–337, 388, 567–569, 637, 752–753 P. parasitica var. nicotianae, 567, 569 P. porri, 336–337, 1323 P. tentaculata, 457 P. tropicalis, 288 Phytophthora blight, 672 Phytophthora crown rot, 541–543, 567–569 Phytophthora root, 426 and crown rot, 847 rot, 336–337, 777, 875 and stem rot, 457–459, 752–753, 1010 Phytophthora wilt, 99 Phytoplasma spp., 471 Phytoplasma, 471–474, 681–683, 824, 1059–1060, 1168, 1300 aurantifolia, 399 diseases, 397, 575–576, 605, 709 mali, 1251 trifolii, 399 Phytopythium helicoides, 462, 895 Phytopythium vexans, 337–339 Phytosanitary certification, 17 Phytosanitary regulations, 224 Phytosanitary requirements, 23 Phytotoxicity, 42, 763, 1108 Pirimicarb, 1329 Pith necrosis, 479 Plant breeding, 1136, 1142, 1173, 1193 Plant collapse, 1083

Index Plant covers, 608 Plant debris, 330, 340 Plant disease clinics, 229 Plant disease diagnosis, 15 Plant health, 238, 242, 244 Plant health regulatory organizations, 224 Plant pathogenic fungi, 74 Plant pathogens, 681 Plant problem diagnosis, 15, 16 Plant protection, 77, 79 Plant resistance, 231 Plant tissue wetness, duration of, 586 Plantago asiatica mosaic virus, 1253, 1258 Planthoppers, 605 Plasmid, 353, 354, 573 Plasmopara halstedii, 796–797 Plastic inserts, 217 Plectosporium blight/leaf spot/cutting rot, 459–461 Plectosporium tabacinum, 459 Plectranthus scutellariodes, 912 Pleospora, 257 P. tarda, 426 Plum pox virus, 1301 Podosphaera sp., 544–547 delphinii, 514 fusca, 546 pannosa, 108, 727–728 xanthii, 98, 546, 804 Poetaz cultivars, 1142 Poeticus, 1211 Poinsettia, 1023 bacterial diseases, 1054–1060 branch-inducing phytoplasma, 1059–1060 cryptic virus, 1062 fungal and fungus-like diseases, 1023–1054 viruses, 1060–1062 Poinsettia latent virus (PnLV), 1062 Poinsettia mosaic virus, 1060–1061 Polianthes tuberosa, 1233 Polyclonal antibodies, 26 Polyethylene, 187 Polygalacturonase, 261 Polygenic mode of inheritance, 105 Polyoxin D, 394, 448, 1105 Polyoxins, 1116 Polyphenol oxydase (PPO), 265 Pospiviroid, 479, 482 Pospiviroidae, 479–483 Post-harvest, 324, 665 bud necrosis, 1274 deterioration, 256

1357 losses, 424 pathogens, 263, 269 Post-lifting sprays, 1141 Potassium, 242 bicarbonate, 425, 456, 587, 990 peroxymonosulfate, 262 peroxymonosulfate plus sodium chloride, 210 Potato cyst nematodes, 180, 182 Potato spindle tuber viroid (PSTVd), 479 Potato virus, 527 Potato virus X (PVX), 489 Potato virus Y (PVY), 489 Potexvirus, 489, 527, 1174, 1257, 1258, 1260, 1327 Potted flowers, 2 Potting media, 172 Potyviridae, 433, 489, 527, 615, 858–860, 1177–1180, 1303 Potyvirus, 361–362, 433, 527, 615, 652, 739, 783, 827, 1177, 1178, 1180, 1255, 1260, 1303, 1327 Powdery mildew, 121, 247, 335–336, 426, 461–462, 514–516, 544–547, 566–567, 597, 671, 745, 847–849, 894–895, 988–990, 1011, 1032–1037, 1112 Pratylenchus, 364, 829–830, 1123 P. coffeae, 364 P. penetrans, 364, 491, 740, 1196, 1261, 1332 P. pratensis, 364, 528 P. vulnus, 740 Predatory nematodes, 185 Predisposing conditions, 20 Premature flower opening, 1273 Pre-soaking, 1209, 1213 Preventive activity, 143 Pre-warming, 1209, 1213 Primulaceae, 1100 Prochloraz, 262, 286, 586, 700, 1140 Proctolaelaps, 709 Procymidone, 791 Production, 2 Proliferation, 354 Promocarb hydrochloride, 636 Propagation, 329 Propagative, 17 Propagules, 202 Propamocarb, 936 Propamocarb hydrochloride, 591 Propiconazole, 286, 425, 456, 462, 805, 930, 932, 1297

1358 Propineb oxadixyl, 591 Proteaceae Armillaria root rot, 704 Botryosphaeria leaf blight and stem cankers, 696 Botrytis blight, 697 Colletotrichum tip dieback, 699 Fusarium wilt, 706 Ilyonectria black foot rot, 707–708 leaf spot diseases, 694–696 Phytophthora root and collar rot and sudden death disease, 702–704 phytoplasma diseases, 709 Pyrenophora blight, 701–702 Pythium root rot, 704 Rhizoctonia root rot, 706 Prothiocarb, 1321 Prunus necrotic ringspot virus (PNRSV), 736 Pseudocercospora sp., 1248 P. eustomatis (Cercospora eustomae), 587 P. puderi, 722 Pseudoidium P. cyclaminis, 1112 P. kalanchoes, 1012 P. poinsettiae, 1032–1037 Pseudomonas, 260, 261, 267, 268, 586, 596, 1167, 1249, 1321 P. andropogonis, 1325 P. antirrhini, 759–760 P. carotovorum, 262 P. cepacia, 261 P. cichorii, 260, 476, 548–549 P. corrugata, 479 P. fluorescens, 259, 261, 425, 1108, 1148 P. gladioli, 650, 1325 P. putida, 261 P. solanacearum, 479, 604, 1118 P. syringae, 260, 397, 479, 732 P. syringae pv. delphinii, 518 P. syringae pv. helianthi, 820 P. syringae pv. maculicola, 781–783 P. syringae pv. morsprunorum, 733 P. syringae pv. tabaci, 396 P. syringae pv. tagetis, 820 P. viridiflava, 429, 479, 1055–1056 Psychoda spp., 594 Puccinia sp., 1247 P. antirrhini, 756–758 P. arenariae, 572 P. basiporula, 1247 P. caniculata, 809 P. chrysanthemi, 448 P. delphinii, 518 P. enceliae, 808

Index P. helianthi, 807–808 P. horiana, 3, 97, 449 P. massalis, 808 P. narcissi, 1166 P. prostii, 1323 P. recondita, 518 P. schroeteri, 1166 P. sporoboli, 1247 P. sporoboli var. sporoboli, 1247 P. tulipae, 1323 P. xanthii, 809 Pucciniastrum hydrangea, 994 Pulse treatment, 271 Purple specks, 327 Pustula helianthicola, 818–820 Pustules, 341 Pycnidia, 506, 720, 751, 1109, 1114 Pyracantha, 247 Pyraclostrobin, 462, 469, 587, 930, 934, 936, 1242 Pyraclostrobin + boscalid, 444 Pyrenophora, 701 Pyrethroid, 1329 Pyrimethanil, 262, 587 Pyroclostobin, 447 Pythium sp., 169, 175, 179, 189, 242, 462, 547–548, 634, 754–755, 777, 846–847, 877, 916–918, 947–949, 979, 1012–1013, 1037–1041, 1080–1082, 1167, 1235 P. acanthicum, 339, 570 P. aphanidermatum, 337–339, 391, 462–464, 569–571, 755, 795, 1109 P. cryptogea, 795 P. debaryanum, 392, 704, 795, 1109, 1239 P. drechsleri, 795 P. intermedium, 1323 P. irregulare, 337–339, 391, 548, 569–571, 594, 599, 755, 795, 1109, 1239, 1320, 1323 P. myriotylum, 93, 569–571, 599, 1080–1082 P. oligandrum, 339, 424, 570 P. oopapillum, 1323 P. paroecandrum, 569–571 P. periplocum, 339, 570 P. rostratum, 795 P. spinosum, 392, 599, 704, 934, 1320 P. splendens, 288, 426, 795, 895, 934, 1239 P. sylvaticum, 569–571 P. ultimum, 239, 337–339, 391, 426, 569–571, 754, 755, 895, 1239, 1323 P. ultimum var. ultimum, 1320 P. vexans, 337–339, 704

Index Pythium basal rot, 569–571 Pythium root, 426 Pythium root and bulb rot, 1320–1321 Pythium root rot, 93, 180, 337–339, 547–548, 569–571, 599–600, 754–755, 846–847, 879, 895–897, 916–918, 1080 Pythium root rot resistance, 93 Q Quantitative trait, 98 Quantitative trait loci, 89 Quaternary ammonium chloride, 1237 Quaternary ammonium compounds (QAC), 207 Quaternary ammonium (QA) product, 936 Quiescent infections, 257 R Race, 330 Race 3 biovar 2 (R3bv2), 5 Race-specific resistance, 105 Radophilis similis, 284 Ralstonia solanacearum, 5, 284, 303–305, 479, 853–854, 957–959, 1118 Ralstonia wilt, 853–854 Ramularia sp., 695, 1162, 1248 R. cyclaminicola, 1113 R. delphinii, 510 R. vallisumbrosae, 1162 Raspberry ring spot virus (RpRSV), 527, 1182 Recirculating subirrigation, 1037 Records, 232 Recycled containers, 876 Recycled irrigation water, 876 Recycling containment basin, 230 Redhead leaf spot, 1076 Reduction of RH, 586 Reflective mulch(es), 484, 605, 606, 608, 827 Registration, 142–143 Regulations, 203 Regulatory agencies, 218 Relative humidity (RH), 120, 1035 Rembrandt tulip-breaking virus (ReTBV), 1328 Removal of diseased plant tissue, 227 Reniform nematode (Rotylenohulus reniformis), 185, 433 Replant disease, 728–729 Reservoirs, 77 Resistance, 145–146, 214, 334, 587, 594, 609, 797

1359 to anthracnose, 90 to bacterial blight, 92 to CBW, 94 to CSVd, 97 to F. solani, 92 to Fusarium, 100 gene, 89 to grey mold, 98 to insecticides, 71 mechanisms, 96 to Phytophthora, 99 to powdery mildew, 101 to systemic Xad infection, 91 Resistant carnation cultivars, 96 Resistant strains, 1105 Resistant to Fusarium wilt, 95 Resting spores, 339 Reverse-transcriptase, 32 Reynoutria sachalinensis, 385 Rhabdoviridae, 433 Rhabdovirus, 652 Rhizobium, 586 R. radiobacter, 477, 734 Rhizoctonia sp., 642, 886, 1237 R. solani, 189, 290, 393–394, 425, 464, 514, 571–572, 706, 755–756, 774–776, 794–795, 849, 918–919, 932, 1043–1046, 1082, 1235, 1238, 1248, 1323, 1324 R. solani (Thanatephorus cucumeris), 600 R. tuliparum, 1166, 1248, 1321, 1323 Rhizoctonia basal stem rot, 755–756 Rhizoctonia crown, and stem rot, blight, 600 Rhizoctonia cutting, 339–341 Rhizoctonia damping-off, 174 Rhizoctonia root, 464–466 and crown rot, 918–919 rot, 849, 886 rot and blight, 1082 Rhizoctonia rot, 571–572 Rhizoctonia rot and blight, 1321–1324 Rhizoctonia stem and cutting rot, 464–466 Rhizoctonia stem and root rot, 425–427 Rhizoctonia stem base rot, 571–572 Rhizoctonia stem rot, 336 Rhizoctonia web blight, 464–466 Rhizoglyphus, 1136 Rhizomes, 264 Rhizomorphs, 716 Rhizophagus intraradices, 592, 601 Rhizopus, 805, 1046–1047, 1160, 1247 R. necans, 1323 R. stolonifer, 1087, 1323 Rhizopus head rot, 805–806

1360 Rhodococcus fascians, 354–355, 475, 573–574, 1249, 1325 Rhodotorula glutinis, 268 Rhynchostylis, 635 Ringspots, 360, 521, 1121 Rinsing, 219 Risk assessment, 142 Root and collar rot, 322 Root and crown rots, 745 Root ectoparasitic nematodes, 364 Root-knot, 619–621, 862, 1092–1094 Root-knot nematodes, 93, 180, 181, 189, 362, 407, 433, 491, 525–528, 686, 740, 761–762, 827–829, 1078, 1124, 1265 Root-lesion nematode(s), 687, 740, 829–830, 1196, 1261 Root plate rot, 1198 Root rot(s), 123, 247, 512, 567–569, 976, 980, 981, 1012, 1043–1046, 1081, 1196–1199 Phytophthora, 1110–1112 Pythium, 1109 Thielaviopsis black, 1115, 1116 Root rotting, 351 Root sloughing, 1081 Rosa hybrida, 255, 264, 272 Rose, 247, 255, 257, 261, 268, 271, 273 Rose leaf curl, 736 Rosellinia necatrix, 1164 Rose mosaic, 736 Rose rosette, 737–738 Rose spring dwarf, 738–739 Rosewarne hybrids, 1142 Rose yellow mosaic virus, 739 Rot, 350 Rotations, 141, 144, 145, 157, 1036 Rotylenchulus macrodoratus, 364 Rotylenchulus reniformis, 185, 364 Rubiaceae, 928 Ruscus aculeatus, 256 Rust, 247, 341–343, 426, 518, 572, 648, 756–758, 994, 1013, 1201 S Septoria leucanthemi, 456 Sadwavirus, 527 Safflower, 258 Salicyclic acid, 269 Sanitation, 77, 79, 1040, 1114, 1137, 1145, 1148, 1159, 1164, 1208 of containers, 874 practices, 202, 384

Index Sanitizing cutting propagation material, 874 Sanitizing station, 220 Sanitizing tools, 216 Sap, 1172, 1175, 1179, 1180, 1182, 1185 Sarcochilus virus Y (SVY), 652 Scab, 1023, 1047–1050 Scale tip rot, 1249 S-carvone, 267 Scatella sp, 593 Schefflera, 260 S. arboricola, 260 Schlumbergera, 977, 979, 981, 983 Scirtothrips, 405 Sclerotia, 291, 324, 340, 383, 447, 469, 505, 642, 665, 719, 932, 1025, 1083, 1113, 1114, 1243, 1291, 1297, 1315, 1322 Sclerotinia, 768–769 S. bulborum, 1165 S. fuckeliana, 1323 S. gladioli, 1297 S. minor, 759, 810–813 S. narcissicola, 1155 S. polyblastis, 1143 S. sativa, 1166, 1323 S. sclerotiorum, 3, 179, 344–345, 396, 426, 466, 572, 759, 810, 850, 1087, 1243–1244, 1323 S. serica, 572 Sclerotinia cottony stem rot, 466–469 Sclerotinia flower rot, 344–345 Sclerotinia rot, 426 Sclerotinia stem rot, 344–345, 466–469, 850 Sclerotium S. batatitcola, 386 S. delphinii, 511, 1242 S. narcissi, 1166 S. perniciosum, 1323 S. rolfsii, 181, 343–344, 396, 426, 469, 511, 758, 815, 850–851, 1082–1085, 1166, 1242, 1323 S. tulipae, 1323 S. tuliparum, 1323 S. wakkeri, 1248 Sclerotium blight, 426, 815, 1082 Sclerotium soil-line rot, 343–344 Sclerotium stem blight, 850–851 Sclerotium stem rot, 343–344, 640–642 Sclerotium rolfsii, 640 Scorch, 1009 Scouting, 1037 Screening anthurium, 90 Screening gladiolus seedlings, 100 Screening procedure, 101

Index Screening protocols, 102 Scrophulariacea, 760 Secondary infections, 885 Secondary invaders, 877 Second juvenile stage (J2), 762 Secoviridae, 526, 527, 615, 1180–1185 Sedentary endoparasites, 1124 Sedentary endoparasitic nematodes, 362–363, 828 Sedum, 1009 Seed, 1175, 1176, 1180, 1181, 1183, 1184, 1187 borne, 791 infection, 421, 423, 425 transmission, 606 treatments, 791, 797 Seedling blight, 759–760 Semi-endoparasitic nematodes, 364 Semiselective culture media, 25 Senescence, 1030 Septoria sp., 645, 1323 S. atropurpurea, 426 S. callistephi Gloyer, 426 S. chrysanthemi, 456 S. delphinella, 510 S. dianthi, 345–346 S. helianthi, 814 S. helianthina, 814 S. spergulae, 345–346 Septoria leaf spot, 345–346, 426 Septoria leaf spot and blight, 814 Sequencing, 19 Serological or molecular assays, 1120 Serratia liquefaciens, 334 Serratia marcescens, 1108, 1114 Settling tower, 108 Shading, 125 Shallot Virus X, 1188 Sheet-steaming, 596 Shikimic acid pathway, 246 Shore flies, 74–75, 80, 593, 1053 Shrubs, 988 Siderophores, 245 Signalling proteins, 246 Silicon, 127, 247 Silver nano-particles, 270–272 Silver nitrate, 270–272 Silver thiosulphate, 270–272 Simple sequence repeat, 95 Single pond system, 230 Single-spore isolates, 105 Skin diseases, 1140, 1167 SLRSV, see Strawberry latent ringspot virus (SLRSV)

1361 Smolder, 1143, 1145, 1149, 1155–1160 Snapdragons, 189 anthracnose, 745 blight, 750–752 Botrytis blight, 746–747 Cercospora blight, 747–748 downy mildew, 748–750 INSV, 760–761 nematode diseases, 761–762 nonparasitic disorders, 762–763 Phyllosticta blight, 750–752 Phytophthora stem rot and wilt, 752–753 powdery mildew, 753–754 Pythium root rot, 754–755 Rhizoctonia basal stem rot, 755–756 rust, 756–758 seedling blight, 759–760 Verticillium wilt, 758 SNP markers, 110 Soaps and detergents, 223 Sodium chloride, 262 Sodium hydroxide plus formaldehyde, 482 Sodium hypochlorite, 266, 272, 423, 482 Soft crown rot, 519 Soft rot, 649, 1056–1057, 1160–1162 Soil, 340 acidification, 818 disinfestation, 816 drenches, 1108 floors, 221 fumigants, 222 fumigation, 333, 813, 1165, 1198 line, 343 moisture, 120 nitrification, 244 pH, 238, 602, 1115 reductive sterilization, 333 saprophyte, 977 sickness, 1196 solarization, 187, 333, 423, 425, 433, 1084 treatment, 1195, 1198 Soil-borne diseases, 132 Soil-borne nematodes, 1124 Soil-borne pathogens, 874 Soil-borne plant pathogens, 73, 75, 176 Solar radiation, 122 Solarization, 174–175, 337, 508, 735, 758, 795, 813, 816, 1094 Source of inoculum, 1113 Southern blight, 343–344, 469–470, 758, 850–851, 1082–1085 Soybean mosaic virus, 1301 Soybean oils, 385

1362 Specific combining ability, 101 Sphaceloma poinsettiae, 1047–1050 Sphaerotheca fuliginea, 804 Sphaerotheca humili var. fuliginea, 514 Sphenospora kevorkianii, 648 Spikkels, 1189 Spiranthes mosaic virus2 (SpiMV2), 652 Spiranthes mosaic virus3 (SpiMV3), 652 Sporangia, 326, 512, 749, 875, 1042, 1111 Sporangiophores, 517, 749 Spores, 223, 506 Sporodochia, 330 Sporulation, 880 Spots, 255, 360 Stagonospora, 1145 S. curtisii, 1145 Standardized protocol, 102 Steam, 173, 174, 182, 592 Steam plough, 596 Steinernema spp., 80 Steinernema feltiae, 79, 433 Stem and bulb nematode, 1264–1265, 1329–1332 Stem and collar rot, 339–341 Stem canker, 507, 843–844 Stem-infesting insects, 800 Stem lesion, 1249 Stem nematode, 1132, 1133, 1136, 1138, 1169, 1179, 1187–1195, 1207, 1210, 1213, 1214 Stem rot, 256, 758, 979, 981, 1012, 1056–1057 Stemphylium, 1009 S. bolickii, 1009 S. botryosum, 426 S. floridanum, 348 S. lycopersici, 426 S. solani, 1009 S. xanthosomatis, 1009 S. laticeps, 1136, 1156, 1189 Stemphylium leaf spot, 426 Stephanotis floribunda, 1233 Sticky cards, 609 Stock, 768 Stock-scion grafts, 473 Storage, 1135, 1137, 1145, 1155, 1159, 1167, 1180, 1187, 1192, 1195, 1211, 1213 Straminipiles, 326 Stratiolaelaps scimitus, 79, 80 Strawberry latent ringspot virus (SLRSV), 527, 736, 1177, 1182, 1252, 1259, 1301 Streptomyces sp., 267, 596, 1141 S. griseoviridis, 325, 385

Index S. lydicus, 325, 385, 388 Streptomycin, 649 Streptomycin sulfate, 477, 936 Stripes, 348 Strobilurin(s), 394, 447, 466, 748, 933, 1103, 1105, 1116, 1291 Strobilurin fungicides, 795, 797, 803 Stromata, 385 Stromatinia gladioli, 1166, 1297–1298 Stubby root nematodes, 1327, 1332 Stub dieback, 328–330 Stunted, 326, 338, 360 Stunting, 349, 357, 360, 363, 521, 576, 1085 Styrofoam transplant trays, 217 Subchronic testing, 141 Suberinization, 1078 Sub-irrigation, 590 Sub-Saharan Africa, 4 Substrates, 171 Sulfur, 244, 245 Sun scorch, 1206 Sunflower, 424, 456 Sunflower chlorotic mottle virus (SuCoMoV), 826 Sunflower mosaic virus (SuMV), 825–826 Sunflower stem weevil, 794 Suppressive, 334 Suppressiveness, 128 Suppressive soils, 877 Sweetpotato whitefly, 76–77 Symptomless, 360 cuttings, 329 endophyte, 595 strain, 483 Synchytrium S. aureum, 510 S. laetum, 1323 Syngonium, 259, 295 S. podophyllum, 259 Systemic acquired resistance, 1030 Systemic infection, 748 Systemic vascular occlusion, 346 Systems approach, 231 T Tagetes spp., 1094 T. patula, 1237 Tank mixes, 141, 144, 145 TAV, see Tomato aspermy virus (TAV) Tazetta, 1143, 1167, 1173, 1175, 1177, 1179, 1181, 1183, 1187, 1188, 1195, 1204, 1206

Index Tebuconazole, 1297 Teliospores, 320, 341, 449, 994 Temperature, 120 Teratosphaeria, 695 Test kits, 1123 Tetracycline, 649, 1060 Tetracycline-HCl, 474 Tetragonia expansa, 1257 Thanatephorus cucumeris, 339–341, 393, 571–572, 775, 1082, 1324 Thanksgiving cactus, 976 Thermal fog, 1026 Thiabendazole, 1237 Thielaviopsis, 248 T. basicola, 601–603, 759, 1050–1054, 1324 T. thielavioides, 716 Thielaviopsis black root rot (Thielaviopsis basicola), 601–603 Thiophanate(s), 292, 388, 588 Thiophanate-methyl, 268, 286, 387, 394, 423, 425, 445, 461, 462, 466, 468, 647, 748, 930, 932, 933, 977, 1116, 1235, 1237, 1244 fungicide, 1103 resistance frequency, 421 Thiophanate-methyl-based products, 1246, 1320, 1322 Thiophene α-therthienyle, 1262 Thiram, 586, 791, 1237 Third stage juvenile (J3), 762 Thripinema nicklewoodi, 433 Thrips, 486, 606, 1120, 1172 Thrips sp., 1176 T. palmi, 487, 610 T. parvispinus, 607 T. setosus, 487, 610 T. tabaci, 430, 431, 487, 607, 610, 1184 Thrips trap crop, 488 Thysanoptera:Thripidae, 405, 430, 606 Tigergrass, 247 Tip blight, 762 Tissue blot immunoassay, 111 Titanium dioxide, 1059 Tobacco etch virus (TEV), 921 Tobacco mosaic virus (TMV), 617, 902, 1301 Tobacco necrosis necrovirus (TNV), 902, 1186, 1301, 1325 Tobacco rattle virus (TRV), 3, 433, 652, 683, 1168, 1186, 1252, 1259, 1301, 1325 Tobacco ringspot virus (TRSV), 489, 857, 903, 1003, 1168, 1183–1184, 1302, 1303 Tobacco streak virus (TSV), 606, 826, 1302

1363 Tobamoviruses, 652 Tobravirus, 652, 1259, 1327 Tolerance, 70, 78 Tomato aspermy virus (TAV), 485, 1302 Tomato black ring virus (TBRV), 1177, 1184, 1302 Tomato bushy stunt virus (TBSV), 652, 1328 Tomato chlorotic spot virus (TCSV), 429, 431, 608 Tomato mosaic virus (ToMV, Tobamovirus), 617 Tomato ringspot nepovirus, 1302 Tomato ringspot virus (ToRSV), 489, 527, 576, 652, 1003, 1184–1185 Tomato spotted wilt tospovirus, 1302 Tomato spotted wilt virus (TSWV), 3, 362, 404, 429, 431, 486, 488, 524–525, 550, 551, 608, 685, 760–761, 856, 901, 983, 1003, 1017–1018, 1120, 1172, 1176 Tomato yellow leaf curl virus (TYLCV, Begomovirus), 613 Tombusviridae, 615–617, 1185–1186 Tombusvirus(es), 357, 577, 652 ToRSV, see Tomato ring spot virus (ToRSV) Tospovirus(es), 71, 362, 404, 429–431, 486, 524, 550, 551, 606, 652, 760, 1120, 1176 Toxicity, 49, 53, 63 Toxicology, 140–142 Toxoptera citricida, 406, 616 Transcriptome, 18 Transgressive segregation, 110 Transmission, 72, 79 Trap crops, 829, 1094 Triadimefon, 444, 462, 805, 932, 1291 Triage, 20 Triazole(s), 748 Triazole fungicides, 1297 Tribasic copper, 477 Trichoderma sp., 176, 267, 268, 466, 469, 596, 795, 813, 1141, 1295 T. hamatum, 341, 469 T. harzianum, 268, 325, 334, 586, 637, 644, 933, 1234 T. lignorum, 600 T. polysporum, 423 T. virens, 328, 1295 T. viride, 423, 1234 Trichodoridae, 1260 Trichodorus sp., 433, 1187, 1260, 1327, 1332 Tricyclazole, 262 Tridemorph, 1291

1364 Trifloxystrobin, 268, 636 Triflumizole, 444, 447, 462, 466, 596, 1105, 1116, 1295 Triticonazole, 444 TRSV, see Tobacco ring spot virus (TRSV) TRV, see Tobacco rattle virus (TRV) Tsuga canadensis, 994 TSWV, see Tomato spotted wilt virus (TSWV) Tuber rot, 1075–1076 Tulip, 109–111, 1314 Tulipa gesneriana, 423, 1314 Tulipa hybrids, 1314 Tulip band-breaking virus (TBBV), 1328 Tulip breaking virus, 111, 1260 Tulip chlorotic blotch virus (TCBV), 1328 Tulip fire, 1314–1316 Tulip mild mottle mosaic virus (TMMMV), 110, 1328 Tulips, 261 Tulip severe mosaic virus (TSMV), 1328 Tulip top-breaking virus (TTBV), 1328 Tulip virus X, 1253, 1260 Tumor(s), 353, 573 Turnip mosaic virus (TuMV), 615, 652, 783 Twist, 1249 Tylenchida: Heteroderidae, 406 Tylenchorhynchus, 830 U Ulocladium altrum, 268 Universal primers, 30 Upper leaf necrosis, 1268 Uredinia, 1296 Urediniospore, 757 Urediospores, 341, 448 Urocystis sp., 1324 U. colchici, 1166 U. narcissi, 1166 U. sorosporioides, 518 Uromyces spp., 1247 U. aecidiiformis, 1247 U. caryophyllinus, 341–343 U. dianthi, 341–343 U. erythronii, 1247, 1324 U. himalaicus, 1247 U. transversalis, 1296 USA, 5 USDA/APHIS, 6 Ustilago heufleri, 1324 Ustilago vaillantii, 1324 UV-B radiation, 589 UV-C irradiation, 265

Index UV light, 122 V Vallota speciosa virus, 1188 Value, 138, 143 Vanda, 635 Vanilla necrosis virus (VanNV), 652 Vankya heufleri, 1324 Vascular blockage, 261, 270, 271 Vascular discoloration, 351, 1101, 1106 Vascular occlusions, 260 Vascular system, 331 Vascular tissue, 349 Vase solutions, 270–273 Vectors, 70, 72, 77–80, 321, 356, 1328 Veitchii, 992 Ventilation, 121 Vertical resistance, 106 Verticillium sp., 178, 179, 248 V. albo-atrum, 394–396, 426, 758 V. cinerescens, 346–347 V. dahliae, 193, 394, 731–732, 758, 776, 816–818 V. lateritium, 1324 V. lecanii, 343 V. zaregamsianum, 776 Verticillium leaf mottle/wilt, 816–818 Verticillium wilt, 346–347, 426, 758, 776 Viburnum tinus, 271, 273 Vinca, 257 Vinclozolin, 268, 791 Viral diseases, 356–362, 1002 Alfalfa mosaic virus, 523 Cucumber mosaic virus, 523–524 Delphinium ringspot virus, 522 Tomato spotted wilt virus, 524–525 Virescence, 471, 472, 520, 1118 Virgaviridae, 433, 617–618, 1186 Viroids, 480–489 Virus(es), 70, 72, 75, 78, 479, 651–656, 899, 901, 1168 acquisition, 72 diseases, 1168–1187 indexing, 943 indicator plants, 609 Virus-free plants, 1304 Virus-indexed, 1174 arabis mosaic, 961–962 Artichoke Italian latent, 961 beet curly top, 962 cucumber mosaic, 963 cultivars, 1173

Index impatiens necrotic spot, 962 Moroccan pepper, 965 Pelargonium flower break, 966 Pelargonium leaf curl, 965 Pelargonium zonate spot, 963–964 stock, 1173 tobacco mosaic, 964 Tobacco necrosis virus, 965 tobacco rattle, 964 tobacco ringspot, 961 tomato bushy stunt, 965–966 tomato ringspot, 961 tomato spotted wilt, 962 Virus-tested stock plants, 358, 1173 Virus/viroid-indexing, 482 Visual diagnosis, 43 Volatilization, 177 Volcano-shaped lesions, 256 W Warm, 1292 Warm-storage, 1194, 1209 Washing hands, 223 Watering practices, 79 Watermelon mosaic virus (WMV), 615 Water molds, 389, 1037 Water pH levels, 230 Water potential, 244 Water treatment system, 228 Water-soaked, 324, 336, 574 Waxflowers, 267 Weather, 1205–1206 Weed(s), 187, 1165, 1172, 1173, 1176, 1181, 1183, 1184, 1186, 1193 Weevils, 801 Western flower thrips, 71–73, 77–79, 488, 920 Wet conditions, 1292 Wet soil conditions, 1109 White mold, 344–345, 572, 1143, 1149, 1160, 1162–1164 White root rot, 1164–1165 White rust, 97, 818–820 White rust-resistant cultivars, 97 Wilt(s), 121, 247, 507, 567–569, 752–753, 758 Wilt diseases, 259 Wilting, 331, 363 Wirestem, 775 Witches’ brooms, 397, 471, 575, 709, 888 Woody ornamentals, 1111 Wool, 1192, 1194, 1209, 1213 Woolflower, 380 Worldwide, 323

1365 Wounds, 330 X Xanthomonas sp., 254, 259, 396, 1057–1059 X. axonopodis pv. dieffenbachiae, 3, 90, 92, 169, 284, 292–302, 1087 X. axonopodis pv. poinsettiicola, 1057–1059 X. campestris, 259, 953–956, 1001 X. campestris pv. begoniae, 898 X. campestris pv. dieffenbachiae, 259, 284 X. campestris pv. incanae, 777–781 X. campestris pv. syngonii, 259 X. campestris pv. zinniae, 852–853 X. hortorum pv. pelargonii, 3 X. maculifoliigardeniae, 936–937 Xanthomonas leaf spot, 1057–1059 Xanthosoma, 295 Xiphinema sp., 489, 830, 1181, 1183, 1185, 1326 X. americanum, 1183 X. coxi, 1259 X. diversicaudatum, 361, 740, 1259 Xylem, 260 tissues, 331 vessels, 261, 575 Y Yellow halo, 1076, 1087 Yellowing, 356, 575 Yellow mottling, 362 Yield, 1133, 1137, 1139, 1143, 1145, 1149, 1152, 1155, 1160, 1162, 1166, 1170, 1171, 1173, 1176, 1177, 1179, 1180, 1183, 1209, 1213 Z Zambia, 5 Zantedeschia, 259 Z. elliottiana, 262 Zantedeschia mosaic virus, 1091 Zinc, 245 Zinnia sp., 247, 257 Z. acerosa, 258 Z. angustifolia, 840 Z. elegans, 840, 862 Ziram, 1291 Zoospores, 337, 512, 755, 875, 1039, 1110, 1111 Zucchini yellow mosaic virus (ZYMV), 489, 904 Zygote, 513