Handbook of diseases of banana, abacá and enset 9781780647197, 1780647190, 9781780647210, 9781780647203

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Handbook of Diseases of Banana, Abacá and Enset

Handbook of Diseases of Banana, Abacá and Enset

Edited by

David R. Jones

CABI is a trading name of CAB International CABI Nosworthy Way Wallingford Oxfordshire OX10 8DE UK Tel: +44 (0)1491 832111 Fax: +44 (0)1491 833508 E-mail: [email protected] Website: www.cabi.org

CABI 745 Atlantic Avenue 8th Floor Boston, MA 02111 USA Tel: +1 (617)682-9015 E-mail: [email protected]

© CAB International 2019. All rights reserved. No part of this publication may be reproduced in any form or by any means, electronically, mechanically, by photocopying, recording or otherwise, without the prior permission of the copyright owners. A catalogue record for this book is available from the British Library, London, UK. Library of Congress Cataloging-in-Publication Data Names: Jones, D. R. (David Robert), 1946- editor. Title: Handbook of diseases of banana, abaca and enset / edited by David   R. Jones. Description: Boston, MA : CABI, [2018] | Includes bibliographical references   and index. Identifiers: LCCN 2018034301| ISBN 9781780647197 (hardback) | ISBN   9781780647210 (epub) | ISBN 9781780647203 (ePDF) Subjects: LCSH: Bananas--Diseases and pests. Classification: LCC SB608.B16 H36 2018 | DDC 634/.772--dc23 LC record   available at https://lccn.loc.gov/2018034301 ISBN-13: 9781780647197 (Hardback) 9781780647203 (ePDF) 9781780647210 (ePub) Commissioning editors: Rachael Russell and Rebecca Stubbs Editorial assistants: Alexandra Lainsbury and Tabitha Jay Production editor: Ali Thompson Indexer: Cath Topliff Typeset by SPi, Pondicherry, India Printed and bound in the UK by Bell & Bain Ltd, Glasgow

Contents

Contributors

xi

The Editor: David Jones

xiii

Preface

xv

1  INTRODUCTION TO BANANA, ABACÁ AND ENSET 1 D.R. Jones and J.W. Daniells The Genera Musa and Ensete 1 Banana 2 Abacá 32 Enset 34 2  FUNGAL DISEASES OF THE FOLIAGE 41 Sigatoka Leaf Spots 41 Overview – D.R. Jones 41 Black Leaf Streak – M. Guzmán, L. Pérez-Vicente, J. Carlier, C. Abadie, L. de Lapeyre de Bellaire, F. Carreel, D.H. Marín, R.A. Romero, F. Gauhl, C. Pasberg-Gauhl and D.R. Jones 41 Sigatoka Leaf Spot – D.R. Jones 115 Eumusae Leaf Spot – D.R. Jones, J. Carlier and X. Mourichon 127 Other Leaf Spots 132 Black Cross Leaf Spot – D.R. Jones 132 Cordana Leaf Spot – D.R. Jones 135 Deightoniella Leaf Spot – D.R. Jones, E.O. Lomerio, M. Tessera and A.J. Quimio 138 Eyespot – D.R. Jones, M. Tessera and A.J. Quimio 141 Malayan Leaf Spot – D.R. Jones 142 Pestalotiopsis Leaf Spot – D.R. Jones 144 Phaeoseptoria Leaf Spot – D.R. Jones 145 Pyricularia Leaf Spot – M. Tessera, A.J. Quimio and D.R. Jones 148 Speckles, Freckle and Rust – D.R. Jones 150 Cladosporiun Leaf Speckle 150 Mycosphaerella Leaf Speckle 156 Taiwan Leaf Speckle 159 v

vi Contents

Tropical Leaf Speckle 161 Freckle 166 Rust 171 3  FUNGAL DISEASES OF THE ROOT, CORM AND PSEUDOSTEM 207 Fusarium Wilt – R.C. Ploetz 207 Armillaria Corm Rot – D.R. Jones 228 Cylindrocladium Root Rot – D.R. Jones 229 Rosellinia Root and Corm Rot – D.R. Jones 232 Damping-off of Musa seedlings – D.R. Jones 233 Deightoniella Pseudostem Rot of Abacá – E.O. Lomerio 233 Fungal Root Rot – D.R. Jones and R.H. Stover 234 Marasmiellus Pseudostem and Root Rot – D.R. Jones and E.O. Lomerio 235 Pseudostem Heart Rot – D.R. Jones and E.O. Lomerio 238 Ceratocystis Corm Rot – D.R. Jones 239 Sclerotium Corm and Pseudostem Rot – D.R. Jones 240 Cephalosporium Infloresence Spot of Enset – M. Tessera and A.J. Quimio 241 4  FUNGAL DISEASES OF BANANA FRUIT 255 Preharvest Diseases – D.R. Jones and R.H. Stover 255 Overview 255 Anthracnose Fruit Rot 255 256 Brown Spot Cigar-end Rot 257 261 Tip-end Rot Deightoniella Fruit Speckle 264 Diamond Fruit Spot 266 Freckle 267 Peduncle Rot 268 Pitting 269 Sooty Mould and Sooty Blotch 270 Postharvest Diseases – D.R. Jones and I.F. Muirhead 271 Overview 271 Crown Rot 272 Anthracnose 280 285 Fungal Scald Stem-end Rot 285 Main-stalk Rot 286 Botryodiplodia Finger Rot 286 Squirter 286 Other Diseases 287 5  DISEASES CAUSED BY BACTERIA AND PHYTOPLASMAS 296 Bacterial Wilt Diseases 296 Xanthomonas Bacterial Wilt – G. Blomme and W. Ocimati 296 Moko Bacterial Wilt and Bugtok – S.J. Eden-Green 314 Blood Bacterial Wilt – S.J. Eden-Green 323 Abacá Bacterial Wilt – D.R. Jones and S.J. Eden-Green 328 Javanese Vascular Disease – D.R. Jones 329 Rhizome and Pseudostem Bacterial Rots – D.R. Jones 329 Overview 329 330 Bacterial Rhizome Rot and Tip-Over Bacterial Soft Rot of Rhizome and Pseudostem 334



Contents

vii

Pseudostem Wet Rot 336 Heart Rot of Abacá 339 Bacterial Leaf Blight and Leaf Spot – D.R. Jones 339 Leaf Blight 339 Black Leaf Spot 339 Bacterial Diseases of Fruit – D.R. Jones 339 Bacterial Finger-tip Rot 339 Phytoplasma Diseases – D.R. Jones 342 6  DISEASES CAUSED BY VIRUSES 362 Banana Bunchy Top – J.E. Thomas 362 Abacá Bunchy Top – J.E. Thomas 376 Bract Mosaic – J.E. Thomas, R. Selvarajan, M.-L. Iskra-Caruana, L.V. Magnaye and D.R. Jones 378 Banana Mosaic – B.E.L. Lockhart, D.R. Jones and J.E. Thomas 384 Abacá Mosaic – J.E. Thomas and L.V. Magnaye 391 Banana Streak – M.L. Iskra-Caruana, J.E. Thomas and M. Chabannes 393 Enset Streak – M. Tessera, A.J. Quimio and D.R. Jones, 409 Banana Mild Mosaic – J.E. Thomas, B.E.L. Lockhart, M.L. Iskra-Caruana, K.S. Crew and M. Sharman 410 Banana Virus X – K.S. Crew and J.E. Thomas 412 Ampeloviruses – K.S. Crew, S. Massart and J.E. Thomas 413 7  NEMATODE PATHOGENS 429 D.L. Coyne and S. Kidane Introduction 429 Burrowing Nematode 429 Root-lesion Nematodes 438 Spiral Nematodes 442 443 Root-knot Nematodes Other Nematodes 448 8  NON-INFECTIOUS DISORDERS OF BANANA 462 E. Lahav, Y. Israeli, D.R. Jones and R.H. Stover Overview 462 Plant Disorders 462 Heart Leaf Unfurling Disorder of Plantain 462 462 High Mat Leaf-edge Chlorosis 463 Roxana Disease 463 Yellow Mat 464 Fruit Disorders 465 Alligator Skin 465 466 Dark Centre of Ripe Fruit Break Neck 466 Hard Lump 466 467 Malformed Fingers and Hands Maturity Stain 468 469 Mixed Ripe Fruit Neer Vazhai 469 Physiological Finger Drop 469 470 Premature Field Ripening Sinkers 470

viii Contents

Split Peel Uneven De-greening Withered Pedicels Yellow Pulp

471 471 472 472

9  MINERAL DEFICIENCIES OF BANANA 475 E. Lahav and Y. Israeli Introduction 475 Nitrogen Deficiency 475 Phosphorus Deficiency 477 Potassium Deficiency 478 Calcium Deficiency 478 Magnesium Deficiency 479 Sulfur Deficiency 481 Manganese Deficiency 481 482 Iron Deficiency Zinc Deficiency 482 Boron Deficiency 484 484 Copper Deficiency 10  INJURIES TO BANANA CAUSED BY ADVERSE CLIMATE AND EXTREME WEATHER 487 Y. Israeli and E. Lahav Introduction 487 488 Seasonal Winds and Windstorms Low-Temperature 492 Chilling and Frost 498 Hail 502 Heat 503 506 Solar Radiation Drought 507 512 Excess Water Lightning Strike 514 Under-peel Discoloration 515 Global Climate Change 516 11  CHEMICAL INJURY TO BANANA 527 E. Lahav and Y. Israeli Introduction 527 Injury Caused by Agricultural Chemicals 527 Fertilizers 527 Pest and Disease Control Chemicals 527 Herbicides 530 Disinfectants 535 De-suckering Agents 535 Injury Caused by Elements in Toxic Concentration 536 536 Sodium and Chlorine Calcium 539 Manganese 539 Iron 539 Boron 540 Copper 541 Aluminium 541

Contents

ix

Arsenic 541 Fluorine 541 12  GENETIC ABNORMALITIES OF BANANA 546 Y. Israeli and E. Lahav Introduction 546 Variations in Stature 547 Abnormal Foliage 551 Variations in Pseudostem Pigmentation 556 Chimeras 558 Inflorescence and Fruit Variations 559 Concluding Remarks 561 13  QUARANTINE AND THE SAFE MOVEMENT OF MUSA GERMPLASM 567 D.R. Jones Introduction 567 Movement of Pathogens 569 571 Aspects of Spread of some Major Health Problems Principles of the Safe Movement of Musa Germplasm 577 Development of a System for the Safe Movement of Musa Germplasm 578 582 Current Recommendations for the Safe Movement of Musa Germplasm Virus Detection and Therapy 583 586 Results of Virus Indexing at the Virus Indexing Centres Index of Musa and Ensete Species, Banana Subgroups and Clone Sets, and Banana, Abacá and Enset Cultivars

593

General Index

599

Contributors

C. Abadie, CIRAD, UMR BGPI, F-97130, Capesterre Belle-Eau, Guadeloupe, France (Email: catherine. [email protected]). G. Blomme, Bioversity International – Productive and Resilient Farms, Forests and Landscapes, c/o ILRI, P.O. Box 5689, Addis Ababa, Ethiopia (Email: [email protected]). J. Carlier, CIRAD, UMR BGPI F-34398 Montpellier, France (Email: [email protected]). F. Carreel, CIRAD, UMR AGAP, F-34398 Montpellier, France (Email: [email protected]). M. Chabannes, CIRAD, Biology and Genetics of Plant-Pathogen Interactions (UMR BGPI), F-34098 Montpellier, France (Email: [email protected]). D.L. Coyne, International Institute of Tropical Agriculture, Kasarani, P.O. Box 30772-00100, Nairobi, Kenya (Email: [email protected]). K.S. Crew, Agri-Science Queensland, Queensland Department of Agriculture and Fisheries, Ecosciences Precinct, Level 2C West, GPO Box 267, Brisbane, Queensland 4001, Australia (Email: kathy.crew@ daf.qld.gov.au). J. Daniells, Queensland Department of Agriculture and Fisheries, South Johnstone, Queensland 4859, Australia (Email: [email protected]). L. de Lapeyre de Bellaire, CIRAD, UPR GECO, F-34398 Montpellier, France (Email: luc.de_lapeyre_ [email protected]). S.J. Eden-Green, EG Consulting Larkfield, Kent ME20 6JA, United Kingdom (Email: sedengreen@ gmail.com). E. Fouré, c/o CIRAD, UPR GECO, F-34398 Montpellier, France. GECO, Univ Montpellier, CIRAD, Montpellier, France (Email: [email protected]). F. Gauhl, FG-Inter-Agro-Consult, Obere Jungenberggasse 18, 1210 Vienna, Austria (Email: [email protected]). M. Guzmán, CORBANA, Plant Pathology Department, P.O. Box 390-7210, Guápiles, Limón, Costa Rica (Email: [email protected]). M.L. Iskra-Caruana, CIRAD, Biology and Genetics of Plant-Pathogen Interactions (UMR BGPI), F-34098 Montpellier, France (Email: [email protected]). Y.L. Israeli, Jordan Valley Banana Experiment Station, Zemach 15132, Israel (Email: yisraeli7@ gmail.com). D.R. Jones, 58 Watts Road, Callala Beach, New South Wales 2540, Australia (Email: bananadoctor@ msn.com). S. Kidane, International Institute of Tropical Agriculture, Kasarani, P.O. Box 30772-00100, Nairobi, Kenya (Email: [email protected]). xi

xii Contributors

E. Lahav, Regional Experiment Station, Akko 25212, Israel (Email: [email protected]). P. Lepoivre, Laboratoire de Phytopathologie, Faculté Universitaire des Sciences Agronomiques, Passage des Déportés 2, 5030 Gembloux, Belgium (Email: [email protected]). B.E.L. Lockhart, Department of Plant Pathology, University of Minnesota, 495 Borlaug Hall, 1991 Upper Buford Circle, St Paul, Minnesota, MN 55108, USA (Email: [email protected]). E.O. Lomerio, c/o Fiber Industry Development Authority (FIDA), Department of Agriculture, Bicol University Compound, Legazpi City, Philippines. W. Ocimati, Bioversity International – Productive and Resilient Farms, Forests and Landscapes, Plot 106, Katalima Road, P.O. Box 24384, Kampala, Uganda (Email: [email protected]). A.J. Quimio, c/o Department of Plant Pathology, University of the Philippines at Los Baños, College, Laguna, 3720, Philippines (Email: [email protected]). L.V. Magnaye, c/o Bureau of Plant Industry, Department of Agriculture, Bago-Oshiro, 8000 Davao City, Philippines. D.H. Marín, Consultores MC, PO Box 709-2250, Tres Ríos, Cartago, Costa Rica (Email: douglashmarin@ gmail.com). S. Massart, Laboratory of Integrated and Urban Phytopathology, Gembloux Agro-Bio Tech, University of Liège, Passage des déportés, 2, 5030 Gembloux, Belgium (Email: sebastien. [email protected]). X. Mourichon, c/o CIRAD, UMR BGPI F-34398 Montpellier, France (Email: xaviermourichon@ gmail.com). I.F. Muirhead, 12 Mondra Street, Kenmore Hills, Queensland 4069, Australia (Email: i.muirhead@ bigpond.com). C. Pasberg-Gauhl, FG-Inter-Agro-Consult, Obere Jungenberggasse 18, 1210 Vienna, Austria ([email protected]). L. Pérez-Vicente, Plant Health Research Institute (INISAV) Ministry of Agriculture. P.O. Box 11300, Havana, Cuba (Email: [email protected]). R.C. Ploetz, University of Florida, Tropical Research & Education Center, 18905 SW 280th Street, Homestead, Florida 33031-3314, United States of America (Email: [email protected]). R.A Romero, Del Monte Fresh Produce, 241 Sevilla Avenue, Coral Gables, Florida 33134, United States of America (Email: [email protected]) R. Selvarajan, Molecular Virology Lab, ICAR- National Research Centre for Banana, Thogamalai Road, Thayanur Post, Tiruchirapalli-620102, TN, India (Email: [email protected]). M. Sharman, Agri-Science Queensland, Queensland Department of Agriculture and Fisheries, Ecosciences Precinct, Level 2C West, GPO Box 267, Brisbane, Queensland 4001, Australia (Email: [email protected]). R.H. Stover† M. Tessera, c/o Ethiopian Institute of Agricultural Research, PO. Box 2003, Addis Ababa, Ethiopia (Email: [email protected]). J.E. Thomas, Agri-Science Queensland, Queensland Department of Agriculture and Fisheries, Ecosciences Precinct, Level 2C West, GPO Box 267, Brisbane, Queensland 4001, Australia (Email: [email protected]).

The Editor: David Jones

Dr David R. Jones was born in Birmingham and grew up in nearby Dudley in the UK’s West Midlands region. He graduated in botany from the University of Hull in 1968 and obtained an MSc degree in plant pathology from the University of Exeter in 1970 before undertaking research on rust fungi at the University of Keele. Awarded a PhD in 1973, he became a post-doctorate research fellow working with Dr Ed Ward at Agriculture Canada’s research laboratory in London, Ontario, on resistant and susceptible interactions between various fungi and Capsicum annum. Dr Jones was next awarded a post-doctorate research fellowship to work with Professor Brian Deverall at the University of Sydney on the interactions between the Lr20 gene in wheat and the rust fungus Puccinia recondita. From 1977 to 1987, Dr Jones held the position of quarantine plant pathologist with the Queensland Department of Primary Industries in Brisbane, Australia. During these years, he xiii

xiv The Editor: David Jones

inspected and indexed imported plant germplasm, ranging from ornamental orchids to crop plants, as part of a national programme to prevent the introduction of exotic pathogens into Australia. He was the first pathologist to travel to the Torres Strait region and discovered black leaf streak and freckle diseases on local banana plants. Dr Jones later formulated guidelines for the safe introduction of Musa species into Australia. Subsequently, as a senior plant pathologist, he worked on banana leaf diseases and postharvest problems. He also visited Papua New Guinea and Southeast Asia as part of missions to collect valuable Musa germplasm and participated in a banana improvement project in the South Pacific. In 1992, Dr Jones joined the International Network for the Improvement of Banana and Plantain (INIBAP) in Montpellier, France, as Crop Protection Research Coordinator. Later, as Scientific Research Coordinator, he played a role in developing INIBAP’s plan of action. During this period, he also served as co-editor of INIBAP’s magazine Infomusa and led a global project to test the resistance of new banana hybrids and other clones to significant diseases. Dr Jones also undertook banana disease surveys in Thailand and Malaysia. His collections of specimens led to the identification of banana bract mosaic virus in India and Sri Lanka and to the recognition of a new banana leaf spot disease in India, Sri Lanka, Malaysia and Thailand now known to be caused by Pseudocercospora eumusae. Leaving INIBAP in 1996, Dr Jones took part in a European Union (EU)-funded project to improve the banana industry of the Windward Islands. He became well known on St Vincent as ‘the banana doctor’ as a result of extension broadcasts on local radio and television. Later, on his return to the UK, Dr Jones served as editor for the CAB International publication entitled Diseases of Banana, Abacá and Enset, which was published in 1999. The next year, he joined the Central Science Laboratory of the Department of the Environment, Food and Rural Affairs near York in England as a pest risk analyst formulating guidelines for the safe movement of plant material into the EU. Afterwards, he became a consultant for the European Food Safety Authority (EFSA) in Parma, Italy. Retiring in 2008, Dr Jones kept his interest in banana diseases and has also recently published a book on the Crimean War of the 1850s. He presently lives with his wife on the shores of Jervis Bay on the south coast of New South Wales, Australia.

Preface

Three to four years ago, I became aware that almost all copies of the CABI Publishing book entitled Diseases of Banana, Abacá and Enset, which I had edited and which was released at the turn of the century, had been sold and inflated prices were being asked by online retailers for the few that remained. Not long after this discovery, I approached CABI Publishing and offered to edit an updated version of the book in my retirement. After an investigation into the need for a new issue of the book, I was given the go ahead to begin work on finding authors for profiles of the various diseases and disorders of the crops involved. I was informed by CABI Publishing that the title of this new book would be Handbook of Diseases of Banana, Abacá and Enset. Published results of research in certain areas seems to have grown exponentially in recent years. Since the previous publication, much work has been undertaken on the most important diseases of banana, such as black leaf streak, Fusarium wilt, Xanthomonas bacterial wilt, bunchy top and banana streak. This is reflected in the increase in the number of pages devoted to these particular problems. More is also known about some of the pathogens causing less serious and widespread diseases. The nomenclature of many fungi affecting banana has changed or is changing as a result of genetic sequencing. Taxonomic mycologists have now abandoned the dual nomenclature system (sexual and asexual morphs) in favour of a natural classification based on one name for a fungal species (Wingfield et al., 2012). If the chosen name is the sexual morph, then the asexual morph name becomes a synonym and vice versa. This approach has not been without its problems and, although the determination of which scientific name to use is based on the principle of priority and for the sexual morph, situations exist in which applying these principles strictly does not contribute to the nomenclatural stability of fungi (Rossman, 2014; Wingfield et al., 2012). While these changes are in progress, it has been problematic to decide which fungal name to include in this publication. Most fungal names are those that are considered to be current in Mycobank and/or Index Fungorum. It will be for the editor or editors of the third edition of this book to resolve issues arising from future decisions made by the Nomenclature Committee for Fungi. As for virus species names, John Thomas has determined when they should and when they should not be written in italics. Should the name of a disease that contains the genus of the causal fungal pathogen change when the fungus is accommodated in a different genus? I was faced with this dilemma in light of reclassifications of the causal agent and the new ‘one name for one fungal species’ nomenclature. For the sake of continuity, I have mostly kept the old disease names in this book even if the fungal genus in the name is no longer appropriate. Again, it is for the editor or editors of the next edition to determine if these diseases need renaming. xv

xvi Preface

The book again follows the format laid down by the late Harry Stover in his 1972 publication entitled Banana, Plantain and Abacá Diseases. As before, chapters cover diseases caused by various groups of causal agents and disorders caused by unknown and known factors. By far the biggest chapter is devoted to fungal diseases of the foliage. The increase in information on diseases that has become available since 1999 means far more of the book discusses these problems. There are no chapters dedicated to conventional banana breeding and genetic engineering for disease resistance as in the first edition. Instead, these topics are covered under the sections on ‘control’ for the various diseases. The revised book is now complete and I would like to thank those who contributed, all of whom are authorities in their fields and many my personal friends. I found my disease descriptions to be much more time consuming than I had anticipated before I started, mainly because I had underestimated increases in the relevant information available since the last edition. I would like to thank Rachael Russell and Alexandra Lainsbury of CABI Publishing for their patience with me, especially when I encountered setbacks. While most authors provided their own illustrations, some additional material was obtained from Tony Cooke and Jeff Daniells of the Queensland Department of Agriculture and Fisheries (QDAF). I would like to thank all contributors for sending me digital images of disease symptoms, most of which appear in this book. Roger Shivas of QDAF is also especially thanked for valuable discussions on fungal taxonomy and the new fungal nomenclature. I am also very grateful to Yu Pie Tan of QDAF for additional mycological information. This book, like the last, is for all ‘banana doctors’ around the world. I hope that it serves as a useful field and laboratory guide plus a source of information to all those investigating problems of the banana, abacá and enset crops. David R. Jones Callala Beach, New South Wales December 2017

References Rossman, A.Y. (2014) Lessons learned from moving to one scientific name for fungi. IMA Fungus 5(1), 81–89. Wingfield, M.J., Wilhelm de Beer, Z., Slippers, B., Wingfield, B.D., Groenewaild, J.Z., Lombard, L. and Crous, P.W. (2012) One fungus, one name promotes progressive plant pathology. Molecular Plant Pathology 13(6), 604–613.

Publisher’s note: Distribution Maps of Plant Diseases Within this volume, Distribution Maps of Plant Diseases have been mentioned as the specific version referenced by the chapter author. These maps are the authoritative source for accurate data on the worldwide distribution of plant diseases of economic or quarantine importance. They were first published in 1942 and created in association with the European and Mediterranean Plant Protection Organization (EPPO). CABI has now published over 1000 Disease Maps. Please visit https://www. cabi.org/dmpd/ to access the latest Distribution Maps of Plant Diseases.

1  Introduction to Banana, Abacá and Enset

D.R. Jones and J.W. Daniells

The Genera Musa and Ensete Banana, abacá and enset belong to the Musaceae family of flowering plants in the order Zingerberales. The wild Musaceae grow mainly in tropical environments from the South Pacific to West ­Africa, but most diversity is found in the Southeast Asia–New Guinea region. Characteristics of the Musaceae that differentiate this family from others in the same order are that the leaves and bracts are spirally arranged, male and female or hermaphrodite flowers are separated within one inflorescence and the fruit is a many-seeded berry (Stover and Simmonds, 1987). The family has two genera: Musa and Ensete. Banana and abacá belong to genus Musa and enset to the genus Ensete. Some species within the Musaceae are utilized as ornamentals, but banana, abacá and enset are the three most economically important crop plants. Species within Musa and Ensete are large herbs with pseudostems of leaf sheaths. New leaves are formed from a meristem near ground level and push up through the pseudostem in a tight roll. Lamina are large, usually oblong, with a stout midrib and numerous parallel veins extending to the margin. The meristem later initiates a terminal inflorescence, which is thrust up the centre of the pseudostem by elongation of the true stem. Flowers are borne in clusters,

each cluster in the axil of a large spathaceous bract. The perianth consists of one compound lobed tepal and one narrow free tepal. The ovary is inferior with three locula, each loculus with numerous ovules in axile placentation. Seeds have a thick, hard testa, a straight embryo and copious endosperm. Pseudostems die after the fruiting phase (Purseglove, 1972; Stover and Simmonds, 1987). Musa spp. have tightly clasping leaf sheaths and a slightly swollen pseudostem base. Basal flowers are generally female only and distal flowers male. Flowers and bracts are inserted independently on the peduncle and, except for functional female ovaries in basal hands, are commonly deciduous by abscission. Bracts are usually reddish, purple or violet due to anthocyanins. Suckers arise freely from an underground rhizome. Pollen grains are finely granular and seed is 7 mm or less in diameter (Purseglove, 1972). In contrast, Ensete spp. have lax leaf sheaths and a pseudostem base which is usually markedly swollen. Basal flowers are usually hermaphrodite. Flowers and bracts are integral with each other and with the axis and are persistent. Suckers are not produced unless induced by humans. Pollen grains are warty and seed is 6 mm or more in diameter (Purseglove, 1972). Species in the genus Musa were originally split into four groups known as sections, based

© D.R. Jones 2019. Handbook of Diseases of Banana, Abacá and Enset (D.R. Jones)

1

2

Chapter 1

on chromosome number and phenotypic characters (Cheesman, 1947):

• •

Eumusa and Rhodochlamys – contain species that have 22 (n = 11) chromosomes. Australimusa and Callimusa – contain species that have 20 (n = 10) chromosomes.

Simmonds and Weatherup (1990) used numerical methods of classification and recognized the same four sections, with Eumusa divided into two distinct subgroups numbered 1 and 2. De Langhe (2000) considered that the characteristics known to be more stable in species evolution in the study of Simmonds and Weatherup (1990) should receive greater weighting and re-analysed their data accordingly. Whilst he found evidence of Eumusa and Rhodochlamys being in the same section, the same was not true for Australimusa and Callimusa. Almost 70 Musa species have now been recognized (Häkkinen, 2013). Some have been found that have anomalous characters and are intermediate between sections. More recently, the results of amplified fragment length polymorphism analyses of DNA extracted from leaf tissue of Musa species and restriction fragment length polymorphism analysis of DNA from organelles of Musa species have provided evidence for the merger of the Eumusa and Rhodochlamys sections (Wong et  al., 2002; Nwakanma et  al., 2003). In addition, Wong et al. (2002) believed from their work that the Australimusa and Callimusa sections could also be combined. As a result of this and other work, Hakkinen (2013) proposed two sections called Musa and Callimusa. Musa with 33 species is a combination of Eumusa and Rhodochlamys while Callimusa with 37 species is a combination of Callimusa and Australimusa, but also contains Musa ingens, which had been proposed by Argent (1976) as belonging to a new section called Ingentimusa. This new arrangement has oversimplified the taxonomic problem. The two sections cannot be defined in either geographical or morphological terms. Furthermore, the ‘new’ Callimusa section still contains species with chromosome numbers other than 20. Additional research is required to resolve the issue. Further discussion in this chapter is based on the original four sections proposed by Cheesman (1947). Eumusa is the largest and most wide-ranging section of the genus and contains Musa acuminata

Colla and Musa balbisiana Colla, which are the principal progenitors of most edible banana cultivars. Another, much smaller group of edible Musa, comprising the Fe’i banana cultivars, is derived from wild species in the Australimusa series, representatives of which are found mainly on the island of New Guinea. Abacá is the species Musa textilis Née, which is also in the Australimusa section. M. textilis is native to the Philippines, but has been introduced to other countries for cultivation. The genus Ensete comprises six to seven species, which are divided between Asia and ­Africa. In Africa, Ensete ventricosum (Welw.) Cheesm. (syn. Musa ensete Gmel.; Ensete edule (Gmel.) Horan.), which is extremely variable, is the most widely distributed and is found between Ethiopia and the Central African Republic, Sudan and South Africa. In Ethiopia, edible cultivars of ­enset have been selected and developed from E. ventricosum.

Banana Facts and figures Banana is one of the most important, but undervalued, food crops of the world. It is grown predominantly in gardens and smallholdings in some 120 countries in the tropics and subtropics and provides sustenance to millions of people. Total world production of bananas in 2013 was estimated to be about 145 million tonnes (FAO, 2015). About 44% comes from the Asian–Pacific region, 31% from Africa and 25% from the Americas. Most fruit is consumed locally. The highest consumption rates are in the Great Lakes region of East Africa, where bananas occupy a large proportion of the diet and amount to 200–250 kg/ person/year. This compares with consumption figures of 8–18 kg/person/year in Europe and North America (FAO, 2014). Bananas for export account for about 14% of total production and are mostly grown in Latin America and the Philippines. Banana is an attractive perennial crop for farmers in developing countries. The fruit can be produced all year round, thus providing a steady cash income and/or supply of nutritious food. Bananas for domestic consumption are harvested



Introduction to Banana, Abacá and Enset 3

from a multitude of cultivars, which are grown on a wide variety of soils and in many different situations. These cultivars can be grown on holdings that range from small plantations to garden plots and jungle clearings. The number of different clones has been estimated to be about 1000 (INIBAP/IPGRI, 2000). Bananas can be divided into two main categories: dessert bananas and cooking bananas. Dessert bananas, which constitute 59% of world production (Lescot, 2015), are eaten raw when ripe, as they are sugary and easily digestible. There are many different types of dessert bananas, but fruit from cultivars in the Cavendish subgroup are the most common and account for 46% of the world’s total production. In some tropical countries, it is the custom to cook dessert bananas. However, cooking bananas, which account for the other 41% of world banana production (Lescot, 2015), are usually starchy when ripe and need to be boiled, fried or roasted to make them palatable. They are important in the diets of many peoples throughout the tropics and are mostly consumed locally. Plantains, the most well known of the cooking types, form 15% of the world’s total production of bananas (Lescot, 2015). As well as being eaten as a fresh or cooked fruit, ripe bananas can be sun- or oven-dried to make ‘figs’ or sliced and fried when unripe to make ‘chips’. Bananas can also be turned into flour, brewed to make beer and, in the Philippines, even form the basis of a ‘tomato’ ketchup. In some countries in Southeast Asia, the male bud of the banana is eaten after the removal of the fibrous outer bracts. Banana is often grown in association with other crops, affording protection and shade. Leaves of banana are utilized as packaging for other foods and serve as plates, tablecloths and decorative items for religious ceremonies. Chopped pseudostems, peduncles and fruit peel can be fed to animals. The banana is a very versatile plant.

Botany The banana is a large, herbaceous plant consisting of a branched underground stem (a rhizome in the strict botanical sense, but also often called a corm) with roots and vegetative buds, and an

erect pseudostem composed of tightly packed leaf bases (Fig. 1.1). The apical meristem, which is located in the centre of the pseudostem at about soil level, gives rise to a succession of leaf primordia. Each primordium grows upwards and differentiates into a leaf base, a petiole and a lamina. The petiole and tightly rolled lamina emerge at the top of the pseudostem in the centre of the cylinder formed by older leaf bases. The lamina begins to unfold when fully emerged, the lefthand side, when viewed from above and from the petiole end, unrolling before the right-hand side. Initially, the leaf forms a tunnel where rainwater or dew can collect and it is at this stage when infection by leaf pathogens can occur. Leaves emerge at different rates depending on the cultivar and environmental conditions. With cultivars in the Cavendish subgroup growing in Honduras, this varies from 3.5–3.8 leaves/month in summer to 2.5–3.0 leaves/month in winter. In winter in the cool subtropics, the leaf emergence rate can fall to 0.1–1.2 leaves/month. A banana plant generally has ten to 15 functional leaves at flowering and five to ten at harvest, but numbers can be less due to environmental and disease constraints. Unless removed, old, senescing leaves hang down the pseudostem. At a certain stage of plant development, usually after about 25–50 leaves have been produced, the apical growing point stops producing leaves and develops an inflorescence. Long photoperiods may promote the change to flowering, but do not seem to be essential for its initiation (Fortescue et  al., 2011). The inflorescence is forced out through the top of the plant as the true stem elongates. The portion of the true stem with the inflorescence that protrudes beyond the base of the uppermost leaf at flowering is known as the transitional peduncle. The bracts on the inflorescence lift to expose double layers of female nodes with tightly packed fruit known as hands. Individual fruit that arise from each flower are often referred to as fingers. Bracts later dry and fall off. This is followed along the peduncle by two or three clusters of neutral flowers, which commonly abscise. Clusters of male flowers then follow to the end of the male peduncle where the male bud is located, which contains the remaining male flowers tightly enclosed in their bracts. The male bud is sometimes called the bell and the male peduncle has been commonly known as the rachis. The underground rhizome, the aerial

4

Chapter 1

Leaf lamina

Midrib Petiole

Unfurling cigar leaf

Bunch with hands of fingers

Pseudostem (tightly packed leaf sheaths)

Inflorescence

Peduncle

Bract Male bud

Sucker

Cross-section of pseudostem Rhizome or corm True stem Roots Fig. 1.1.  Diagrammatic representation of a fruiting banana plant with suckers (from Champion, 1963).

stem (to which is attached the bases of the leaves present within the pseudostem) and the peduncle (to which is attached the inflorescence) are now recognized in banana as all being parts of the true stem. The bunch is the collective name for the hands of fruit attached to the female peduncle. Bunch size and weight depend on plant vigour and health. In the tropics bunches can be harvested between 85 and 110 days after flowering, but in the cool subtropics fruit development can take up to 210 days. After harvest, the pseudostem dies back and is therefore usually cut down. The plant propagates by forming a sucker, which is an outgrowth of a vegetative bud on the

rhizome. A young sucker emerging from the ground is called a peeper. Several suckers can arise from each rhizome. One sucker is usually selected by the farmer as a follower to grow on and regenerate the plant. Other suckers are ­either removed physically or their apical meristems are destroyed chemically. If all suckers are allowed to grow, a clump consisting of plants of various sizes all arising from the same connected rhizome system or mat eventually develops. Primary roots, which are about 5–8 mm in diameter, usually arise in groups of three or four from the fleshy rhizome. When healthy and vigorous, primary roots are white in colour, but later turn grey or brown before dying. A system



Introduction to Banana, Abacá and Enset 5

of secondary and tertiary roots develops from the primary roots. The root system is shallow, seldom penetrating much below 600 mm. Horizontal spread can be as far as 5 m, but more commonly 1–2 m. Banana roots are sensitive to adverse conditions, such as waterlogging, desiccation and compacted soil (Wardlaw, 1961; Robinson and Galán Saúco, 2010).

Cultivation and trade Banana is believed to have been grown initially in Southeast Asia and Melanesia in small permanent gardens around settlements (Plate 1.1), in forest clearings by shifting cultivators (Plate 1.2) and in gardens with other crops (Plate 1.3). Many cultivars with differing genetic backgrounds and different responses to pathogens and pests were most probably cultivated together. Disease problems are unlikely to have been severe enough under these agrosystems to have influenced the selection of disease-resistant cultivars over susceptible ones. Horticultural characteristics were

probably the main considerations in the selection, propagation and spread of early landraces. The first crop after planting is known as the plant crop and the second cycle as the ratoon crop or first ratoon. This is followed by the second ratoon, third ratoon, etc. Traditional planting material is either the sucker (Plate 1.4) or bit, which is a piece of rhizome with a vegetative bud. However, since the 1980s, plants derived from tissue culture have also been utilized in commercial operations. Tissue-cultured plants have the advantage of a higher establishment rate producing a more uniform crop, which typically gives a higher yield in the first crop cycle, and they also have the potential of being completely free of pests and diseases when planted. However, the use of this type of planting material is not without problems. Extra care needs to be taken during establishment and in the subsequent selection of followers. Young plants also have a greater susceptibility to pests and diseases (Daniells, 1997; Smith et al. 1998). In addition, some plants may be adversely affected as a result of somatic mutations occurring during the in vitro multiplication process (Daniells et al., 1999).

Plate 1.1.  Banana plots close to houses in Kasaka village, East New Britain, Papua New Guinea (photo: D.R. Jones, QDPI).

6

Chapter 1

Plate 1.2.  Banana growing in a temporary clearing in the rainforest near Kiunga, Western Province, Papua New Guinea (photo: D.R. Jones, QDPI).

Plate 1.3.  Banana growing together with taro, cassava, sweet potato and sugarcane in a garden on Badu Island in the Torres Strait region of Australia (photo: D.R. Jones, QDPI).



Introduction to Banana, Abacá and Enset 7

Plate 1.4.  A sucker of ‘Robusta’ (AAA, syn. ‘Giant Cavendish’) selected for planting in St Vincent, Windward Islands (photo: D.R. Jones, SVBGA).

Banana plants need ample supplies of water if they are to yield well. This has been estimated at 25 mm/week for satisfactory growth in the tropics (Purseglove, 1972). Banana grown in drier areas has to be irrigated to maintain optimal growth. Plants also usually require inputs of nutrients, such as nitrogen and potassium, to prevent deficiency problems. Banana cultivated as a monocrop on a commercial scale is only a relatively recent phenomenon. Large plantation-scale methods of cultivation began in the late 19th century, after the advent of steamships and cool storage techniques, by a great increase in demand for the fruit in ­Europe and North America. ‘Gros Michel’ was the first export banana cultivar to be planted over thousands of hectares, but fruit for export today is mainly produced by cultivars in the Cavendish subgroup. In Latin America and the Philippines, export fruit is usually produced on extensive plantations on relatively flat coastal plains with

alluvial soils. In contrast, export bananas produced in the Windward Islands in the Caribbean are produced on smallholdings, which are often found on steeply sloping hillsides. The cultivation of genetically similar banana plants on vast tracts of land is not confined to areas where fruit is grown for export. Some popular banana cultivars whose fruit is produced for domestic markets can also be grown contiguously over large areas of land. Examples are ‘Kluai Namwa’ (ABB, Pisang Awak subgroup) in Thailand, ‘Pisang Berangen’ (AAA, Lakatan subgroup) in Malaysia and East African highland banana cultivars (AAA, Lujugira–Mutika subgroup) in Uganda (Plate 1.5). Export bananas need to be as blemish-free as possible at markets to obtain premium prices. The bunch is therefore usually covered with a perforated, polyethylene sleeve, cover or bag (Plate 1.6) soon after emergence to protect the fruit from rubbing leaves and wind-blown debris. Bagging can also shorten the fruit maturation

8

Chapter 1

Plate 1.5.  Contiguous small plots of East African highland banana cultivars occupy large swathes of the countryside in the Rutoto Valley in the Bushenyi district of Uganda (photo: D.R. Jones, INIBAP).

period by 3–4 days and increase the weight of the bunch by 1.8–2.3 kg. If impregnated with insecticide, the bag offers protection from thrips and other fruit-attacking insects that cause damage. At about the time of bagging, the male bud is snapped off because in some circumstances this practice can also increase bunch weight. The bag is usually tied to the peduncle above the scar formed by the upper spathaceous bract. In windy areas, the bag is also tied at the bottom to prevent the polyethylene rubbing on the fingers, which causes scarring. If the bottom of the bag is tied, the knot is positioned at one side to allow rainwater to drain freely from the bag. This is because a build-up of humidity within the bag can lead to fungal disease problems on the fruit. Rough handling of bananas leads to bruising, scarring and other damage. Export plantations have developed methods of handling bananas to minimize this wastage, including establishing cableways for moving bunches to packing sheds where fruit is de-handed, washed, treated with fungicide to reduce postharvest losses and placed into cardboard cartons for transport.

Mature banana fruit produces ethylene, which triggers ripening. Ripening involves the conversion of starch to sugar, a softening of the fruit texture and usually a change in peel colour from green to yellow. These physiological changes can also stimulate the development of postharvest diseases. Locally consumed fruit can be left on the plant until ripening commences. However, bananas for export or consumption some distance away from growing areas are usually harvested at the mature hard green stage. Timing of harvest is important in these circumstances. This can be determined by tagging bunches on emergence and then calculating approximate harvesting dates. Usually, coloured ribbons are tied around the peduncle (Plate 1.6), the colour of the ribbon signifying the particular week of emergence. When the bunch reaches harvesting age, maturity or grade of the fruit is checked by measuring the diameter of the middle finger on certain hands with a caliper. The bunch is harvested at the desired grade. Export fruit has to be held and transported at cool temperatures to minimize the respiration rate of fruit and associated ethylene release and



Introduction to Banana, Abacá and Enset 9

For many years, most banana scientists worked predominantly for multinational companies or for governments supporting export growers. However, in the past few decades, banana has been recognized as an important local fruit crop by developing countries in the Americas, Africa and Asia and local improvement programmes have been initiated. Associated with these programmes has been a need to identify constraints to local banana production, including pests and diseases. In countries where Cavendish is not the most popular type of banana, new disease problems are emerging as local cultivars are beginning to be grown as a monocrop and on a scale much larger than before.

Origin of the edible banana

Plate 1.6.  A protective polyethylene bag covering a bunch of a cultivar in the Cavendish subgroup growing in an export plantation in Ecuador. The bunch has been tagged with a blue ribbon, which identifies its week of emergence (photo: J.W. Daniells, QDAF).

so delay the onset of ripening. Even so, premature ripening can occur if fruit is over-mature, damaged physically or harvested from plants with high levels of leaf spot. Mixed ripe fruit is often rejected at wholesale markets. Controlled atmosphere conditions on ships transporting bananas can reduce the incidence of ripe and turning fruit. At markets, bananas can be stored under cool conditions until needed and then ripened artificially with ethylene. Disease and pest problems have the potential to be more serious when plants with an identical genetic make-up are cultivated on a large scale. With banana, this first happened when ‘Gros Michel’ was grown on plantations in the Latin American–Caribbean region. In the course of time, Fusarium wilt destroyed the viability of this first major export cultivar. After 1960, the banana trade in the region was dominated by Cavendish cultivars and the problems associated with clones in this subgroup, such as Sigatoka leaf spot and burrowing nematode, became limiting factors requiring much research.

Wild Musa species are seedy, generally opportunistic and in nature grow in forest clearings and along watercourses. Simmonds (1962) believed that early agriculturists initially selected plants with high levels of parthenocarpy, which is the ability to form fruit without fertilization. This fruit would have had fewer seeds, thus increasing edibility. At the same time, selection would also have been for female sterility, which would also lower the number of seeds. Selected plants would have been propagated by suckers, thus maintaining their genetic composition. The long-distance movement of vegetative planting material by humans enabled edible banana cultivars from Asia eventually to reach other regions of the world (Simmonds, 1962).

Evolution of edible banana cultivars from Musa species in the Eumusa section Simmonds (1962) postulated that the evolution of edible cultivars derived from Musa species in the Eumusa section most probably began with wild M. acuminata subspecies, which occur naturally in an area stretching from South Asia to Australasia (Fig. 1.2). Another key event was believed to have been hybridization with M. balbisiana, a more drought tolerant species that now occurs in an arc from the Indian subcontinent through the southern China region to the Philippines and New Guinea (Fig 1.3). This would have

10

Chapter 1

CHINA INDIA

anic ides n a o rm ic bu ann m r bu

siamea

truncata a

arp

c cro

mi

errans

banksii

malaccensis

zebrina

AUSTRALIA Fig. 1.2.  Approximate natural distribution of Musa acuminata subspecies (adapted from Simmonds, 1962; Horry et al., 1997).

led to the generation of cultivars that could be grown in drier areas. Both M. acuminata and M. balbisiana are diploids. The genome of wild M. acuminata is represented by AAw and wild M. balbisiana by BBw. Fruit of these two wild species is not usually consumed as it is seedy. However, bananas from one selection of M. balbisiana known as ‘Bhimkol’ or ‘Arthiyakol’ are eaten in Assam, India, along with the tender inner pseudostem and male buds (Borborah et al., 2016). Diploid cultivars derived solely from M. acuminata are designated by AA and diploid hybrids with M. balbisiana by AB. Recent analyses of biotechnological taxonomic data by Perrier et al. (2011) indicated that edible AA cultivars were, as Simmonds (1962) suggested, derived from hybridizations between different M. acuminata subspecies. This is believed to have occurred in the islands of Southeast Asia and western Melanesia as a result of the movement by humans of M. acuminata subspecies and primitive AA cultivars to new areas. Their work also indicated that the structural heterozygosity of these first hybrids, caused by

chromosomal rearrangements between parental subspecies of M. acuminata, contributed to gamete sterility and that this sterility, in association with human selection for pulp enhancement, led to parthenocarpic fruits and edibility. Perrier et al. (2011) believed that a consequence of the hybrid status of the AA cultivars was erratic meiosis occasionally producing diploid gametes and that the fusion of diploid gametes with haploid gametes would have generated sterile triploid (AAA) genotypes. Further, they argued that this spontaneous triploidization occurred in almost all diploid cultivars. Diploid cultivars can often be distinguished from triploids because of their more slender pseudostems and more upright leaves (Fig. 1.4). Triploid banana plants are typically bigger, sturdier plants with increased fruit size. This latter characteristic would have encouraged their adoption into farming systems. The integration of cultivars derived from M. acuminata subspecies with genetic components from M. balbisiana is thought by Perrier et  al. (2011) to have occurred when the latter was spread by humans south to the



Introduction to Banana, Abacá and Enset 11

CHINA

INDIA balbisiana

MALAYSIA

INDONESIA

AUSTRALIA Fig. 1.3.  Approximate natural distribution of Musa balbisiana (adapted from Simmonds, 1962). Argent (1976) believed that the species was introduced to New Guinea. Perrier et al. (2011) considered that the species was also introduced to the Philippines. Simmonds (1962) thought M. balbisiana to be indigenous to both locations.

Diploid

Triploid

Fe’i

Fig. 1.4.  General appearance of diploid and triploid cultivars in the Eumusa series of edible banana and Fe’i cultivars in the Australimusa series of edible banana (Bourke, 1976).

12

Chapter 1

Philippines and New Guinea from southern ­China. A scheme illustrating the various steps in the evolution of the main genomic groups is shown in Fig. 1.5. Human selection would have then led to the diversity of modern cultivated triploids, including pure M. acuminata cultivars (AA, AAA) and interspecific M. acuminata × M. balbisiana cultivars (AB, AAB, ABB). Vegetative propagation of the popular diploid and triploid cultivars over long periods of time resulted in the appearance of variants with different phenotypic characteristics. This process of natural somatic mutation would have increased morphological diversity (Noyer et al., 2005). Only a few natural cultivars have been recognized as belonging to the AAAA, AAAB, AABB and ABBB genomic groups (Richardson et al., 1965; Shepherd and Ferreira, 1984). These tetraploids, which have robust pseudostems and leaves that tend to droop, are believed to have arisen from the fertilization of triploid egg cells by haploid pollen. ‘Kluai Teparot’, a cultivar long believed to have been an ABBB (Simmonds and Shepherd, 1955; Richardson et al., 1965; Silayoi and Sompen, 1991), was found by flow cytometry in association with chromosome counting and molecular analysis in the late 1990s to be an ABB (Jenny et  al., 1997; Horry et  al., 1998). Similarly, ‘Pisang Jambe’ was discovered to be an AAA and

not an AAAA as previously thought (Horry et al., 1998). However, ‘Kluai Ngoen’, which was identified as an AAB from phenotypic characteristics, has now been found to be an AAAB (Horry et al., 1998). Clearly, some cultivars need to be investigated thoroughly before their ploidy can be determined with any degree of certainty. Tetraploid hybrids have been artificially bred for disease resistance and are becoming important in some countries. Classification of banana cultivars derived from Musa species in the Eumusa section The genome of banana cultivars derived from Musa species in the Eumusa section has in the past been deduced from an analysis of morphological characters including pseudostem colour, shape of the petiolar canal and bract features (Simmonds and Shepherd, 1955). This taxonomic method has been proved over time to give a good approximation of the genetic composition generally correlating well with the results of modern molecular techniques. However, as noted under the previous subheading, more modern techniques are necessary to confirm ploidy. Within each genomic group, morphological characters in addition to those used to define Hybrids

M. acuminata Wild species

M. balbisiana BBw

AAw

Selection of parthenocarpy Primitive, fertile diploid cultivars

AA

Selection of sterility Diploid cultivars

AA

Triploid cultivars

AAA

AB

AAB

ABB

Fig. 1.5.  Evolution of the main genomic groups of edible banana cultivars of the Eumusa series (after Simmonds, 1962; Carreel, 1995).



Introduction to Banana, Abacá and Enset 13

genomic group are used to identify clones. Many clones have mutated over time to form morphotypes that differ in certain characters, such as fruit and bunch morphology, pigmentation and height. Dwarfness is a common mutation and has occurred in many locations. Clones that are thought to have arisen originally from a base clone by mutation form subgroups. Major subgroups with large numbers of clones have formed in centres of secondary diversification. It is not always possible to identify the original base clone of major subgroups. The same clone can have different names in different locations. This is especially true in Papua New Guinea, a country with over 700 languages, where the names of cultivars can vary between villages. In this publication, the synonym most frequently used by banana workers has been adopted to signify the clone. This ‘best-known synonym’ is in many cases the one applied to the clone in the Caribbean where much of the early taxonomic work was undertaken. Thus ‘Silk’, a synonym of West Indian origin, gives its name to the clone that is known in India as ‘Rasthali’, in the Philippines as ‘Latundan’ and in Australia as ‘Sugar’. Cultivar names in some countries begin with the local word for banana. For example, names in Malaysia and Indonesia begin with ‘Pisang’, in Thailand with ‘Kluai’, in Vietnam with ‘Chuo͂́i’, in Hawaii with ‘Mai’a’ and in ­Samoa with ‘Fa’i’. Some workers believe this word to be redundant and accessions in germ­ plasm collections have been registered without using the prefix. However, the local word for ­banana has been retained in this publication ­because it is considered an integral part of the cultivar name and also gives an indication as to the original source of the material. Cultivars are placed in subgroups if they are deemed to have been derived by mutation from a single original or base clone. Mutations affect pigmentation, height and other morphological characters. Over time, multiple changes could have occurred increasing diversity. Subgroups are named after the best-known synonym of the most important clone in the subgroup (e.g. Gros Michel and Bluggoe subgroups), the cultivars first described to identify the main characteristics of the subgroup (e.g. Iholena and Lujugira–Mutika subgroups) or a generic term (e.g. Cavendish and Plantain subgroups). In the case of the Plantain subgroup, it is believed that

more than one base clone exists (Stover and Simmonds, 1987). To help the reader relate the cultivars mentioned in this publication to the subgroup or clone, cultivar names are followed in parentheses by the genomic group and then by either the subgroup or best-known synonym.

Subgroups and important clones of banana cultivars derived from Musa species in the Eumusa section AA and AB genomic groups Well-known AA and AB clones and some of their synonyms are listed in Table 1.1. The most important AA clone is ‘Sucrier’, which has exceptional fruit quality. It is quite common in Southeast Asia and is present elsewhere in the world. ‘Sucrier’ is exported from the Philippines and Latin America. ‘Pisang Lilin’ is another popular AA clone, but is not grown widely outside Malaysia and Thailand. Fruit of both ‘Sucrier’ and ‘Pisang Lilin’ can often be seen on sale in Thailand (Plates 1.7 and 1.8). Other AA diploids are cultivated in Southeast Asia, but are not as common. For example, ‘Inarnibal’ is found in Malaysia, Indonesia and the Philippines. There are also many AA clones in Papua New Guinea with more than 100 listed by Arnaud and Horry (1997). In general, they are not very productive, but are grown for their eating qualities and ethnic preferences. Some produce a bunch quickly while others soften rapidly when cooked compared with the widespread ‘Kalapua’ (ABB). ‘Pisang Lilin’ and ‘Paka’ from East Africa and ‘Pisang Jari Buaya’ from Malaysia have been used in breeding programmes because of their resistance to disease. The only AB clones of note are ‘Ney Poovan’ (Plate 1.9) and ‘Kunnan’. ‘Ney Poovan’ is not common, but is found in many countries. In Uganda, it is used for making beer. AAA genomic group The AAA genomic group contains some of the most productive clones, which are widely grown and constitute about half of the world’s total output of banana fruit (Lescot, 2015). Major AAA subgroups are presented in Table 1.2.

14

Chapter 1

Table 1.1.  AA and AB genomic groups – important subgroups. Subgroup AA genomic group Sucrier

Pisang Lilin Inarnibal

Pisang Jari Buaya AB genomic group Ney Poovan Kunnan

Clones/Synonyms

Location

Main use of fruit

‘Sucrier’/‘Figue Sucrée’ ‘Pisang Mas’ ‘Kluai Khai’ ‘Amas’ ‘Lady’s Finger’ ‘Orito’ ‘Pisang Lilin’ ‘Kluai Lep Mu Nang’ ‘Inarnibal’ ‘Pisang Lemak Manis’ ‘Pisang Lampung’ ‘Pisang Jari Buaya’

West Indies Indonesia, Malaysia Thailand Philippines Hawaii Ecuador Malaysia Thailand Philippines Malaysia Indonesia Indonesia, Malaysia

Dessert

‘Ney Poovan’/‘Safet Velchi’ ‘Kisubi’ ‘Kunnan’

India Uganda India

Dessert Brewing Dessert

Dessert Dessert

Dessert

Plate 1.7.  Fruit of ‘Kluai Khai’ (AA, Sucrier subgroup) for sale at roadside stall near Kamphaeng Phet, north-western Thailand (photo: D.R. Jones, INIBAP).

‘Gros Michel’ is a tall clone and was the first banana of the export trades. Despite its susceptibility to Fusarium wilt, it is still grown for local consumption in many countries around the world because of its superior flavour and golden yellow appearance when ripe. ‘Gros Michel’ produces a few seeds when pollinated. As a

consequence, ‘Gros Michel’ and its shorter variants (‘Cocos’, ‘Highgate’ and ‘Lowgate’) have been utilized in breeding programmes. Cavendish cultivars may have originated in Indochina (Perrier et  al., 2011). Two ‘Dwarf Cavendish’ plants were sent to the UK from Mauritius in 1829 having been collected in China



Introduction to Banana, Abacá and Enset 15

Plate 1.8.  Fruit of ‘Kluai Lep Mu Nang’ (AA, Pisang Lilin subgroup) for sale in Bangkok, Thailand (photo: D.R. Jones, INIBAP).

Plate 1.9.  Bunch of ‘Ney Poovan’ (AB) on a plant growing in the germplasm collection at the South Johnstone Research Station, North Queensland, Australia (photo: J.W. Daniells, QDAF).

2–3 years previously. One was grown and propagated in a glasshouse at Chatsworth House belonging to William Cavendish, the 6th Duke of Devonshire. These plants were described as Musa Cavendishii by Paxton (1837). From Chatsworth, suckers were taken to Samoa and then other ­Pacific Islands. The Cavendish subgroup is responsible for 46% of the world’s production of banana fruit (Lescot, 2015). Stover and Simmonds (1987) recognize four major clone sets distinguished on height: ‘Dwarf Cavendish’ types are the shortest in stature, ‘Grande Naine’ types are medium dwarfs, ‘Giant Cavendish’ types are taller and ‘Pisang Masak Hijau’ types the tallest (Stover and Simmonds, 1987). This classification is somewhat arbitrary as in reality there is an almost continuous gradation in height from ‘Extra-Dwarf Cavendish’, which is shorter than ‘Dwarf Cavendish’, upwards to ‘Pisang Masak Hijau’. Each clone set contains different cultivars, some of which are listed in Table 1.2. As well as height, clones in the Cavendish subgroup differ in other morphological characters such as petiole length, leaf length/breadth ratio, bract persistence, bunch grade and pseudostem colour. Other characteristics, such as leaf emergence rate and length of crop cycle, have also been used to differentiate cultivars (Daniells, 1990b).

16

Chapter 1

Table 1.2.  AAA genomic group – Gros Michel, Cavendish and Lujugira–Mutika subgroups. Subgroup

Clone

Clones/synonyms

Location

Main use of fruit

Gros Michel

‘Gros Michel’

West Indies Malaysia Thailand Sri Lanka Hawaii Indonesia

Dessert

‘Cocos’ ‘Highgate’ ‘Lowgate’

‘Gros Michel’ ‘Pisang Embun’ ‘Kluai Dok Mai’ ‘Anamala’ ‘Bluefields’ ‘Pisang Ambon Putih’ ‘Cocos’ ‘Highgate’ ‘Lowgate’

Honduras Jamaica Honduras

Dessert Dessert Dessert

Subgroup

Clone set

Some cultivars in clone set

Location

Main use of fruit

Cavendish

Extra-Dwarf Cavendish Dwarf Cavendish

Dessert

‘Zhong Ba’ ‘Poyo’ ‘Pei-Chiao’ ‘Chuõí Tieu’ ‘Veimama’ ‘Nañicao’ ‘Valery’ ‘Pisang Masak Hijau’

Australia Israel Australia India, Pakistan French Antilles Latin America French Antilles Philippines General India, Windward Islands Australia, South Africa China West Africa Taiwan Vietnam Fiji Brazil Latin America Malaysia

‘Buñgulan’ ‘Pisang Ambon Lumut’ ‘Lacatan’

Philippines Indonesia Latin America

Some cultivars in clone set

Location

Main use of fruit

Uganda

Cooking

Uganda

Cooking

Uganda

Cooking

Uganda

Cooking

Grande Naine

Giant Cavendish

‘Dwarf Parfitt’ ‘Dwarf Nathan’ ‘Dwarf Cavendish’ ‘Basrai’ ‘Petite Naine’ ‘Grand Nain’ ‘Grande Naine’ ‘Umalag’ ‘Giant Cavendish’ ‘Robusta’ ‘Williams’

Pisang Masak Hijau

Subgroup

Clone set

Dessert

Dessert

Dessert

Dessert

Lujugira–Mutika Musakala

Nakabululu Nakitembe Nfuuka

‘Musakala’, ‘Enyoya’, ‘Mudwale’, ‘Mukazialanda’ ‘Nakabululu’, ‘Butobe’, ‘Kibuzi’, ‘Mukite’ ‘Nakitembe’, ‘Mbwazirume’, ‘Nasaala’, ‘Waikova’ ‘Nfuuka’, ‘Enyeru’, ‘Nakhaki’, ‘Kasenene’



Introduction to Banana, Abacá and Enset 17

‘Dwarf Cavendish’ (Plate 1.10) was the basis of local banana trades in the subtropics, but suffered from ‘choke’ (impeded bunch emergence) in the cooler weather. Here and elsewhere, the clone has been progressively replaced by medium dwarf and giant Cavendish cultivars, which are higher yielding and better meet market specifications. The medium dwarf ‘Grande Naine’, which is less prone to wind damage, but more susceptible to drought than other Cavendish cultivars, has replaced ‘Valery’ (‘Giant Cavendish’-type) as the most popular cultivar in international trade in tropical areas. The next most important clone is ‘Williams’. ‘Robusta’ (‘Giant Cavendish’-type), which is less susceptible to a water deficit than ‘Grande Naine’, is still grown extensively in the Windward Islands where irrigation is not widely available and seasonal droughts occur. In addition to being the principal banana types grown for export fruit in the Latin American– Caribbean, African and Southeast Asian regions, Cavendish cultivars form the backbone of domestic banana industries in such places as

Plate 1.10.  ‘Dwarf Cavendish’ (AAA) growing in a back-garden in Abepura, Papua, Indonesia. Note the persistent bracts on the peduncle just above the male bud (photo: J.W. Daniells, QDAF).

Australia, China, Egypt, India, Pakistan, South Africa and Vietnam (Plate 1.11). They have higher yields than all other natural clones (Robinson and Galán Saúco, 2010). Cultivars in Lujugira–Mutika subgroup are the predominant banana grown in the Great Lakes region of East Africa. They are also often referred to as belonging to the East African highland banana (EAHB) subgroup, though other AAA cultivars, including those in the Ilalyi subgroup, and the AA cultivars ‘Mshare’ and ‘Muraru’ are found in the same environment and have been considered part of a EAHB complex (De Langhe et al., 2001; Karamura et al., 2016). It has been estimated from figures published by Lescot (2015) that approximately 10% of all banana fruit produced globally comes from clones of EAHB subgroup. ‘Lujugira’ and ‘Mutika’ are the cultivars that represent the two main subdivisions proposed for this subgroup by Shepherd (1957). This division is based on fruit morphology, with further differentiation related to other characteristics (Stover and Simmonds, 1987). A numerical taxonomic study of over 200 Ugandan accessions in the subgroup distinguished five different clone sets (Karamura, 1999). However, a recent genetic analysis of 90 cultivars has shown only a limited amount of variation that could not be correlated with previous divisions based on morphological characteristics (Kitavi et al., 2016). The fruit of the ‘Lujugira–Mutika’ subgroup, like fruit from other AAA clones, is sweet when ripe. However, East Africans either cook the fruit as ‘matooke’ or use it for brewing to make ‘banana beer’. The pulp qualities of the fruit determine if a cultivar is a cooking or brewing type. Some cooking cultivars can change spontaneously to become beer cultivars. This change occurs with more frequency above 1400 m. A checklist of East African highland types in Uganda lists 145 cooking cultivars and 88 beer cultivars together with synonyms (Karamura and Karamura, 1994). Fruit from these cultivars are very important in the diets of people in the eastern part of the Democratic Republic of the Congo, Rwanda, Burundi, Uganda, Tanzania and western Kenya. Bunch features of this subgroup are illustrated in Plate 1.12. The AAA group also contains a number of other subgroups, some of which are listed in Table 1.3. ‘Red’, and its common sport ‘Green

18

Chapter 1

Plate 1.11.  Bunches of ‘Chuõí Tieu’ (AAA, syn. ‘Giant Cavendish’) being transported to market near Pho Ho, Vietnam (photo: J.W. Daniells, QDAF).

Plate 1.12.  Bunches from cultivars in the Lujugira–Mutika subgroup (AAA) at a village in the Bushenyi district of Uganda awaiting transportation to market in Kampala (photo: D.R. Jones. INIBAP).



Introduction to Banana, Abacá and Enset 19

Table 1.3.  AAA genomic group – Red, Lakatan and Ibota subgroups. Subgroup

Clones/synonyms

Location

Main use of fruit

Red

‘Red’ – ‘Green Red’ ‘Pisang Raja Udang’ – ‘Pisang Mundam’ ‘Morado’ – ‘Moradong Puti’ ‘Ratambala’ – ‘Galanamalu’ ‘Chenkadali’ – ‘Venkadali’ ‘Red Dacca’ – ‘Green Dacca’ ‘Lakatan’ ‘Pisang Berangan’ ‘Pisang Barangan’ ‘Ibota Bota’ ‘Kluai Khom Bao’, ‘Kluai Khai Thong Ruang’ ‘Pisang Saripipi’ ‘Yangambi Km 5’

West Indies Malaysia

Dessert

Lakatan

Ibota

Red’, are widely distributed tall garden clones. The difference between the two cultivars is in their pigmentation, especially in the fruit peel. Immature fruit of ‘Red’ is red in colour (Plate 1.13) while fruit of ‘Green Red’ is green. ‘Red’ has dwarf forms, such as ‘Figue Rose Naine’. ‘Lakatan’, a banana with an excellent flavour, is very popular in Southeast Asia (Plate 1.14). ‘Ibota Bota’ is a clone highly resistant to disease. A semi-dwarf variant has been described (Daniells and Bryde, 1995). Many other AAA clones, which are important locally, have been identified (Stover and Simmonds, 1987). ‘Pisang Nangka’ is found in Malaysia and Indonesia and in the Philippines as ‘Nangka’. ‘Pisang Susu’ is grown in Indonesia. ‘Paji’ from Zanzibar is resistant to black leaf streak disease. ‘Pisang Ambon’ and ‘Orotava’ accessions collected in Indonesia in the 1980s are different from clones in the Gros Michel and Cavendish subgroups (Carreel, 1995). Ten distinct AAA clones were collected in Papua New Guinea in 1988/89 (Arnaud and Horry, 1997). AAB genomic group Four of the major subgroups recognized within the AAB genomic group are presented in Table 1.4. The Plantain subgroup is very important as plantain cultivars provide food for many millions

Philippines Sri Lanka India Australia Philippines Malaysia Indonesia Dem. Rep. of the Congo Thailand

Dessert

Dessert

Indonesia Widely distributed accession

of people in the West and Central Africa and Latin American–Caribbean regions. Cultivars are also found in East Africa and South and Southeast Asia. A total of 15% of the world’s production of banana fruit comes from plantain (Lescot, 2015). The term plantain is often used for all cooking banana types, but in this publication it refers only to those cultivars belonging to the Plantain subgroup within the AAB genomic group of banana (Shepherd, 1990). Cultivars in the Plantain subgroup are placed in four main clone sets, which are distinguished on bunch and inflorescence characteristics. French plantain types have many hands with comparatively small fingers and an inflorescence axis covered with persistent hermaphrodite and male flowers. The large male bud is also persistent. The bunch features of French plantain are illustrated in Plate 1.15. At the other extreme, Horn plantain types have few hands of very large fingers, no hermaphrodite flowers and no male axis. French Horn and False Horn plantain types are intermediate classification categories between French and Horn plantain. The male bud is absent at maturity in both of these types, but there are many hermaphrodite flowers on French Horn cultivars and few on False Horn cultivars (Tezenas du Montcel and Devos, 1978; Swennen, 1990). Tezenas du Montcel (1979) later proposed six

20

Chapter 1

Plate 1.13.  Bunches of ‘Cevvazhai’/‘Sevazhai’ (AAA, Red subgroup) for sale at a wholesale market in Tiruchirappalli, Tamil Nadu State, India (photo: J.W. Daniells, QDPI).

Plate 1.14.  Fruit of ‘Pisang Barangan’ (AAA, Lakatan subgroup) is very popular in Indonesia. (photo: J.W. Daniells, QDAF).

subdivisions (Giant French, Average French, Dwarf French, French Horn, False Horn and True Horn) after an analysis of many different characteristics ranging from the colour and circumference of the pseudostem to orientation of the fingers and male bud characteristics. West and Central Africa is an important centre of secondary diversification of plantain. De Langhe (1961), who recognized 56 cultivars in the Democratic Republic of the Congo, believed that they all arose from one clonal source. Recent investigations of over 200 plantain cultivars indicate that genetic diversity is very low. No correlation has been found between the diversity that has been found and morphological characteristics (Christelová et  al., 2016). ‘Agbagba’, a False Horn



Introduction to Banana, Abacá and Enset 21

Table 1.4.  AAB genomic group – Plantain, Pome, Mai’a Maoli-Popoulu and Iholena subgroups. Subgroup

Clone set

Clones/synonyms

Location

Main use of fruit

Plantain

French

‘Obino I’Ewai’ ‘Njock Kon’ ‘Nendran’ ‘Dominico’ ‘French Sombre’ ‘Mbang Okon’ ‘Batard’ ‘3 Vert’ ‘Agbagba’ ‘Orishele’ ‘Dominico-Hartón’ ‘Cuerno’ ‘Barraganete’ ‘Ihitisim’ ‘Pisang Tandok’ ‘Tindok’

Nigeria Cameroon India Colombia Cameroon Nigeria Cameroon Cameroon Nigeria Nigeria Colombia Central America Ecuador Nigeria Malaysia Philippines

Cooking

French Horn

False Horn

Horn

Cooking

Cooking

Cooking

Subgroup

Clones

Synonyms

Location

Main use of fruit

Pome

‘Pome’

‘Pome’ ‘Prata’ ‘Virupakshi’, ‘Vannan’, ‘Sirumalai’ ‘Kijakazi’ ‘Brazilian’ ‘Prata Anã’ ‘Prata Catarina’ ‘Pacovan’ ‘Pachanadan’ ‘Lady Finger’

Canary Islands Brazil India

Dessert

Zanzibar Hawaii Brazil Brazil Brazil India Australia

Subgroup

Clones/synonyms

Location

Main use of fruit

Mai’a Maoli-Popoulu

‘Mai’a Maoli’, ‘Mai’a Ele’ele’ ‘Mei’a Ma’ohi Hai’, ‘Mei’a Mao’i Maita’ ‘Comino’, ‘Pompo’, ‘Maqueño’ ‘Pacific Plantain’ ‘Mai’a Ka’io’, ‘Mai’a Huamoa’ ‘Mei’a Po’u Hu’amene’, ‘Mei’a Po’upo’u’ ‘Popoulou’ ‘Mai’a Ha’a’,

Hawaii

Cooking

‘Prata Anã’ ‘Pacovan’ ‘Pachanadan’

Iholena

‘Mai’a Kapua’, ‘Mai’a Lele’ ‘Mei’a Ore’a ‘Ute ‘Ute’ ‘Maritú’

Dessert Dessert

French Polynesia Ecuador Australia Hawaii French Polynesia CRBPa Hawaii

French Polynesia Colombia

Germplasm collection of the Centre régional bananiers et plantains, Cameroon.

a

Dessert

Cooking and Dessert

22

Chapter 1

Plate 1.15.  Bunches of ‘Dominico’ (AAB, French Plantain-type) for sale near Guayaquil, Ecuador (photo: J.W. Daniells, QDAF).

plantain, has been reported to revert occasionally to French plantain (Tezenas du Montcel et  al., 1983). The Linnaean name Musa paradisiaca has been applied to French plantain, but should not be used today (Stover and Simmonds, 1987). Cultivars in the Pome subgroup are important dessert banana types in India and Brazil, where their subacid flavour is much appreciated. The taste is also popular in Australia and ­Hawaii. However, cultivars are generally not very productive. Uma and Sathiamoorthy (2002) recognized 17 distinct cultivars of the Pome subgroup in India. ‘Prata Anã’ and a selection named ‘­Prata Catarina’ are synonymous dwarf variants of ‘Prata’ from Brazil. ‘Santa Catarina Prata’ (Plate 1.16) from Hawaii originated in Brazil. ‘Pacovan’ is a large fruited variant from Brazil. Both ‘Prata Anã’ and ‘Pacovan’ have been used in breeding programmes. The ‘Lady Finger’ cultivar grown commercially in Australia may be synonymous with ‘Pachanadan’ of India. Traditional cooking-banana cultivars found on islands in the Pacific Ocean have been placed in the Mai’a Maoli-Popoulu subgroup of the AAB genomic group. The Mai’a Maoli and Popoulu types of Hawaii differ in aspects of fruit shape and bunch appearance (Pope, 1926; Kepler and Rust, 2011), and isozyme studies by Lebot et al. (1994)

Plate 1.16.  ‘Santa Catarina Prata’ (AAB, Pome subgroup) photographed at the South Johnstone Research Station in North Queensland came from Hawaii, but originated in Brazil, where it is popular as are other ‘Prata Anã’ selections. (photo: J.W. Daniells, QDAF).



Introduction to Banana, Abacá and Enset 23

suggested that the two types may have had independent origins. However, their separation into two distinct subgroups does not appear to be supported by simple sequence repeat (SSR) marker studies (Christelová et al., 2016). Furthermore, there are several cultivars from the subgroup in the western Pacific that are intermediate between Mai’a Maoli and Popoulu types in their characteristics, such as ‘Pacific Plantain’ grown in Australia (Plate 1.17). The original base clones of the Mai’a Maoli and Popoulu types, which may have originated in the New Guinea area (Daniells, 1990a; Lebot et al., 1994), were probably carried by Polynesians on their migrations across the Pacific. Iholena is the name given to a third type of AAB banana found in the Pacific, but mainly in Hawaii. The fruit is sweet, but is primarily used for cooking (Plate 1.18). Cultivars, which differ in some morphological characteristics, are also thought to have arisen from a base clone carried by migrating Polynesians from the New Guinea area (Lebot et  al., 1994). The occurrence of a few representatives of the Iholena and Mai’a

Plate 1.17.  ‘Pacific Plantain’ (AAB, Mai’a Maoli-­ Popoulu subgroup), seen here at East Palmerston in Queensland, is grown commercially on a small scale in Australia (photo: J.W. Daniells, QDAF).

Maoli–Popoulu subgroups in the Andes region of South America has led to speculation that banana may have been introduced to the continent from the Pacific in pre-Columbian times (Langdon, 1993). Some other important clones in the AAB genomic group are listed in Table 1.5. ‘Silk’, which has fruit with a sweet-acid taste, is a very popular dessert cultivar in South and Southeast Asia, East Africa and Latin American–­ Caribbean regions (Plate 1.19). Uma and Sathiamoorthy (2002) recognized five distinct variants in India. ‘Silk’ has been described under the Linnaean name Musa sapientum, but this and all other Latin names for banana should be disregarded (Stover and Simmonds, 1987). ‘Mysore’ is grown on a large scale in South Asia (Plate 1.20) and like ‘Silk’ has sweet-acid fruit. However, outside India and Sri Lanka, it is usually only occasionally encountered. Exceptions are Trinidad, where it is used to shade cocoa, and Samoa. Only a few sports of ‘Mysore’ have been identified. ‘Pisang Raja’ is a clone found in Malaysia and Indonesia, but is rare elsewhere (Plate 1.21). ‘Pisang Kelat’ is a dessert banana from Malaysia.

Plate 1.18.  Bunch of ‘Uzakan’ (AAB, Iholena subgroup) in a germplasm collection at Mbarara, Uganda (photo: J.W. Daniells, QDAF).

24

Chapter 1

Table 1.5.  AAB genomic group – Silk, Mysore, Pisang Raja, Pisang Kelat, Sukali Ndizi and Laknau subgroups. Subgroup

Clones/synonyms

Location

Main use of fruit

Silk

‘Silk Fig’ ‘Rasthali’ ‘Kolikutt’ ‘Pisang Rastali’ ‘Pisang Raja Sereh’ ‘Latundan’ ‘Maçã’, ‘Manzano’ ‘Pukusa’ ‘Sugar’ ‘Mysore’ ‘Poovan’ ‘Embul’ ‘Fa’i Misiluki’ ‘Pisang Keling’ ‘Kikonde Kenya’ ‘Pisang Raja’ ‘Grindy’ ‘Pisang Kelat’ ‘King’ Sukali Ndizi ‘Laknau’ ‘Pisang Raja Talong’ ‘Kune’

West Indies India Sri Lanka Malaysia Indonesia Philippines South America Zanzibar Australia West Indies, Australia India Sri Lanka Samoa Malaysia Zanzibar Malaysia, Indonesia Windward Islands Malaysia Trinidad Uganda Philippines Malaysia Papua New Guinea

Dessert

Mysore

Pisang Raja Pisang Kelat Sukali Ndizi Laknau

Dessert

Dessert and cooking Dessert Dessert Cooking

Plate 1.19.  Fruit of ‘Rasthali’ (AAB, Silk subgroup) for sale in Tiruchirappalli, Tamil Nadu State, India (photo: D.R. Jones, INIBAP).



Introduction to Banana, Abacá and Enset 25

Plate 1.20.  Bunches of ‘Poovan’ (AAB, Mysore subgroup) for sale at the wholesale banana market in Tiruchirappalli, Tamil Nadu State, India (photo: D.R. Jones, INIBAP).

‘Laknau’ is a plantain-like cultivar that has been used for breeding. ‘Sakali Ndizi’ is popular in East Africa (Plate 1.22). One AAB clone is an oddity. ‘Pisang Seribu’ has hundreds of small, tightly packed fingers on the bunch and has been described under the Latin name Musa chiliocarpa Backer (Plate 1.23). ABB genomic group

Plate 1.21.  Bunch of ‘Pisang Raja’ (AAB) in the MARDI germplasm collection at Serdang, Selangor State, West Malaysia. Note the persistent male flowers (photo: D.R. Jones, INIBAP).

ABB cultivars are generally hardy and disease-­ resistant. Most produce starchy fruit, which is cooked. The popular cultivar ‘Bluggoe’ gives its name to a major ABB subgroup (Table 1.6). and is regarded as the base clone. ‘Silver Bluggoe’, a common, waxy-fruited variant, is also important. Both are widely distributed. A number of other variants have been described from India, the likely origin of the subgroup and centre of diversity. Dwarfing to create the ‘Dwarf Bluggoe’ type is rare and may have only occurred in the western hemisphere. Fruit of ‘Bluggoe’ has a distinctive angular shape (Plate 1.24). Other important clones in the ABB genomic group are listed in Table 1.7. ‘Monthan’ gives its name to another ABB subgroup that seems likely to have originated in India. Clones are morphologically similar to cultivars in the Bluggoe subgroup. ‘Monthan’ and

26

Chapter 1

Plate 1.22.  Fruit of ‘Sakali Ndizi’ (AAB) being transported to market near Jinja, Uganda (photo: D.R. Jones, INIBAP).

Plate 1.23.  Bunch of ‘Kluai Roi Wi’ (AAB, syn. ‘Pisang Seribu’), which has hundreds of small fingers, at the Surat Thani Research Station in southern Thailand (photo: D.R. Jones, INIBAP).

‘Bluggoe’ distinguished by the pronounced ­cylindrical apex on fruit of the former compared with the tapering apex of fruit of the latter. Uma and Sathiamoorthy (2002) recognized six clones in the Monthan subgroup. There are a number of ABB cooking-­ banana cultivars in the Philippines, typified by ‘Saba’, which forms a distinct subgroup. Valmayor et  al. (2002) recognized 13 cultivars. They include the waxy-fruited sport ‘Abuhon’, the larger-­ fruited ‘Cardaba’ and ‘Gubao’, the fused-­­ fingered ‘Inabaniko’, the small-fruited ‘Turangkog’ and its waxy-fruited mutant ‘­Sabang Puti’. A bunch of ‘Pisang Kepok’, an Indonesian cultivar in the Saba subgroup, is illustrated in Plate 1.25. ‘Pisang Awak’ is a widely disseminated, high-yielding cooking/dessert cultivar, which is also used as a beer banana in East Africa. It is very common in Thailand, Vietnam and elsewhere in Indochina. Subclones in Thailand differ in fruit pulp colour. ‘Kluai ­Namwa’ (syn. ‘Pisang Awak’)

has yellow pulp, ‘Kluai Namwa Khao’ has white pulp and ‘Kluai Namwa Daeng’ has pink pulp. ‘Kluai Namwa Khom’ is a dwarf form (Silayoi and Chomchalow, 1987). One variant, which has been collected in West Malaysia, has sweeter fruit containing many more seeds and an apparent increased susceptibility to freckle disease. ‘Pelipita’, which is known as ‘Pelipia’ in the Philippines, has been planted in Central America as a Moko bacterial wilt-resistant ­cultivar. ‘Ney Mannan’, which is a clone from South Asia, is very popular in Sri Lanka, where it is known as ‘Alukehel’ or ‘Ash Plantain’. ‘Pata Sina’ in the Ney Mannan subgroup is shown in Plate  1.26. ‘Peyan’ is another South Asian clone. ‘Kalapua’ is a common clone in Papua New Guinea with several variants, including a dwarf form. ‘Pitogo’ is a cultivar from the ­Philippines with almost round fruit (Plate 1.27). ‘Kluai Teparot’, the fruit of which is also shown in Plate 1.27, is now included in the ABB group following the finding that it is not a tetraploid.



Introduction to Banana, Abacá and Enset 27

Table 1.6.  ABB genomic group – Bluggoe subgroup. Clone

Synonyms

Location

Main use of fruit

‘Bluggoe’

‘Bluggoe’, ‘Moko’ ‘Pisang Abu Keling’ ‘Nalla Bontha’ ‘Fa’i Pata Samoa’ ‘Kidhozi’, ‘Kivuvu’ ‘Matavia’ ‘Kluai Som’ ‘Largo’ ‘Square Cooker’, ‘Mondolpin’ ‘Silver Bluggoe’, ‘Silver Moko’ ‘Kluai Hakmuk’ ‘Thella Bontha’ ‘Katsila’ ‘Chamaluco Enano’

West Indies Malaysia India Samoa East Africa Philippines Thailand Hawaii Australia West Indies Thailand India Philippines Puerto Rico

Cooking

‘Silver Bluggoe’

‘Dwarf Bluggoe’

Cooking

Cooking

Genomic groups with S and T components Shepherd and Ferreira (1984) and Arnaud and Horry (1997) identified some cultivars in Papua New Guinea believed to contain genetic elements of Musa schizocarpa, another wild species in the Eumusa section. Genomic groups were designated AS, AAS and ABBS, where S indicates hybridization with M. schizocarpa. Other cultivars were thought to have genetic components from species in the Australimusa section. AAT, AAAT and ABBT genomic groups have been recognized, where T indicates hybridization with an Australimusa species. Research using molecular taxonomic methods has indicated that M. schizocarpa and one or more species in the Australimusa section did play a role in the origin of some Eumusa cultivars present in Papua New Guinea (Carreel, 1995). However, cultivars with these unusual combinations of genomic constituents seem to be only occasionally found in cultivation. Plate 1.24.  Bunch of ‘Bluggoe’ (ABB) at South Johnstone, Queensland, Australia (photo: J.W. Daniells, QDAF).

The male axis is occasionally absent from this robust clone, which is found in many countries in Southeast Asia (Stover and Simmonds, 1987).

BBB genomic group There is no evidence to suggest that parthenocarpy occurred in M. balbisiana as it did in M.  ­acuminata. Although ‘Saba’-type cultivars have been classified as having a BBB genome using the scoring method of Simmonds and ­ Shepherd (1955) (Pascua and Espino, 1987), this has been challenged on morphological grounds

28

Chapter 1

Table 1.7.  ABB genomic group – Monthan, Saba, Pisang Awak, Pelipita, Ney Mannan, Peyan, Kalapua and Kluai Teparot subgroups. Subgroup

Clones/synonyms

Location

Main use of fruit

Monthan Saba

‘Monthan’, ‘Saba’ ‘Pisang Kepok’ ‘Pisang Abu Nipah’ ‘Pisang Awak’ ‘Kluai Namwa’ ‘Katali’ ‘Chuõí Tay’ ‘Karpuravalli’ ‘Kayinja’ ‘Ducasse’ ‘Pelipita’ ‘Pelipia’ ‘Ney Mannan’ ‘Alukehel’, ‘Ash Plantain’ ‘Blue Java’ ‘Ice Cream’ ‘Peyan’ ‘Kalapua’ ‘Kluai Teparot’ ‘Tiparot’ ‘Pisang Abu Siam’

India Philippines Indonesia Malaysia Malaysia Thailand Philippines Vietnam India East Africa Australia Central America Philippines India Sri Lanka Fiji, Australia Hawaii India Papua New Guinea Thailand Philippines Malaysia

Cooking Cooking

Pisang Awak

Pelipita Ney Mannan

Peyan Kalapua Kluai Teparot

Plate 1.25.  Young bunch of ‘Pisang Kepok’ (ABB, Saba subgroup) at Besum, Papua, Indonesia (photo: J.W. Daniells, QDAF).

Cooking and Dessert

Cooking Cooking

Cooking Cooking Cooking

Plate 1.26.  Bunch of ‘Pata Sina’ (ABB, Ney Mannan subgroup) near Apia, Upolu, Samoa (photo: J.W. Daniells, QDAF).



Introduction to Banana, Abacá and Enset 29

Plate 1.27.  Fruit of ‘Kluai Teparot’ (left), an ABB clone with angular fingers found in a number of Southeast Asia countries and ‘Pitogo’ (right), an ABB clone with small rounded fingers grown in the Philippines (photo: D.R. Jones, INIBAP).

(­Shepherd, 1990) and on molecular evidence (Jarret and Litz, 1986; Carreel, 1995). However, ‘Kluai Lep Chang Kut’ from Thailand may be a BBB as it has no M. acuminata characteristics and resembles M. balbisiana morphologically. This clone may have arisen from a cross between an ABB cultivar, such as ‘Kluai Teparot’, and M. balbisiana (Shepherd, 1990; J.P. Horry, Montpellier, 1999, personal communication). For more information on subgroups and clones, see Stover and Simmonds (1987) and Daniells et  al. (2001). Consult Robinson and Galán Saúco (2010) for data on bunch weights and yields/ha of various banana cultivars. Advances in knowledge on the origin of cultivars derived from Musa species in the Eumusa section M. acuminata has a number of subspecies and each has its own area of distribution (see Fig. 1.2). The Malayan peninsula was suggested by Simmonds (1962) as the location of the origin of edible banana because M. acuminata ssp. malaccensis was believed by him to be the primary source of edibility. However, this hypothesis was challenged by Carreel (1995). With the aid of restriction fragment length polymorphism (RFLP)

markers, she compared DNA from chloroplasts (inherited through the female parent), mitochondria (inherited through the male parent) and the nucleus of many Musa species, subspecies and landraces. Her work gave an insight into the wild species and subspecies that have contributed to the genetic make-up of cultivars derived from Musa species in the Eumusa section. This in turn has given clues as to probable location of origin of cultivars. The wild Musa species and subspecies implicated in the ancestry of the cultivated Eumusa banana cultivars are listed in Table 1.8. As discussed earlier, the first crucial step in banana domestication is believed to be hybridization between geographically isolated subspecies of M. acuminata found in Southeast Asia and western Melanesia (Perrier et al., 2011). Three regions where this hybridization is likely to have occurred have been identified. One southern contact zone stretched from New Guinea to Java, one eastern contact zone from New Guinea to the Philippines and another northern contact zone from the Philippines to mainland Southeast Asia (Perrier et al., 2011). In the southern contact zone, M. acuminata ssp. banksii (Plate 1.28), zebrina and microcarpa may have interbred with ssp. banksii and errans crossing in the eastern contact zone and ssp. microcarpa, malaccensis and errans

30

Chapter 1

Table 1.8.  Wild Musa implicated in the ancestry of the Eumusa series of edible banana cultivars (Carreel, 1995). Species

Subspecies

Geographical distributiona

Musa acuminata

banksiib erransb burmannicad (burmannicoides) siamead malaccensis microcarpa zebrina

New Guinea, north-east Queensland (Australia), Samoac Philippines Myanmar

Musa acuminata

Musa acuminata Musa acuminata Musa acuminata Musa balbisiana Musa schizocarpag Australimusa speciesh

Thailand, Indo-China Southern Thailand, West Malaysia, Sumatra (?) North Borneo Java Indochina, northern Myanmar, India, Sri Lanka, Philippines, New Guinea,e Malaysia,f Thailandf New Guinea New Guinea, Solomon Islands

Based on information from Simmonds (1962), Shepherd (1990), Carreel (1995) and Daniells et al. (2016). The nuclear genomes of M. acuminata ssp. banksii and ssp. errans are similar, but the cytoplasmic genomes are different. c Musa acuminata ssp. banksii is believed to have been introduced to Samoa (Simmonds, 1962). d The nuclear, chloroplastic and mitochondrial genomes of M. acuminata ssp. burmannica, burmannicoides and siamea accessions in international collections are similar (Carreel, 1995). e Simmonds (1962) believed M. balbisiana to be indigenous to Papua New Guinea, but Argent (1976) and Perrier et al. (2011) thought the species was introduced. f Musa balbisiana was introduced to Malaysia and Thailand, where it was cultivated. g This species has contributed to the genome of some cultivars found in Papua New Guinea. h One or more species within the Australimusa section may be contributing to the genome of some cultivars in Papua New Guinea. a b

in the northern contact zone. Hybridization with M. balbisiana (Plate 1.29) is believed to have occurred in the eastern and southern contact zones after its dissemination south from the Asian mainland. India is thought to be another area where M. balbisiana hybridized with introduced cultivars (Perrier et al., 2011). The work of Carreel (1995) indicated that M. acuminata ssp. banksii or ssp. errans contributed genetically to very many edible cultivars. Numerous cultivated parthenocarpic diploids are now thought to have arisen as a result of crosses between these subspecies (or their fertile derivatives) disseminated by humans to new areas and the local subspecies. Other interesting information has emerged from Carreel’s study. Cooking and beer-making cultivars in the Lujugira–Mutika subgroup (AAA), which are common in a secondary centre of diversity in the highlands of East Africa, contain genetic components of M. acuminata ssp. banksii and ssp. zebrina. It is believed that the progenitor of the distinct East African highland banana types had its origin in south-eastern Indonesia where genetically close AAA clones are

still cultivated (Perrier et  al., 2011). Banana is considered by some historians to have reached the east coast of Africa at about the same time as other Southeast Asian food crops and to have been transported by Indonesian voyagers Simmonds, 1962). Carreel (1995) also con(­ firmed that both the A genomes of AAB Plantain and Mai’a Maoli-Popoulu subgroups were derived from M. acuminata ssp. banksii, as had been suggested by earlier workers (Horry and Jay, 1990; Lebot et al., 1994). From a plant pathological perspective, it is interesting to note that M. acuminata ssp. banksii and ssp. errans, which are genetically close, are recognized as having considerably more disease problems than other Musa species and subspecies of M. acuminata (Vakili, 1965, 1968; Argent, 1976). If almost all edible banana cultivars in the Eumusa section have inherited some component of their genetic make-up from M. acuminata ssp. banksii and ssp. errans, it may explain their susceptibility to certain diseases to varying degrees. Current research is throwing more light on the origin of today’s cultivars. Greater knowledge of genetic backgrounds will ultimately



Introduction to Banana, Abacá and Enset 31

Plate 1.28.  Musa acuminata ssp. banksia (AAw), pictured here growing wild in north-western Papua New Guinea, is the probable progenitor of the first edible banana cultivars in the Eumusa series (photo: D.R. Jones, QDPI).

explain the reactions of landraces to disease and assist breeding programmes. Edible banana cultivars derived from Musa species in the Australimusa section A second distinct group of edible cultivars called Fe’i banana is found in eastern Indonesia, New Guinea and islands in the Pacific. Cultivars in this group have mostly upright fruit bunches (see Fig. 1.4), orange fruit when ripe and sap varying from pink through to purple (Plate 1.30). These similarities to Musa maclayi, a wild Australimusa species found in Papua New Guinea, led Simmonds (1962) to believe that M. maclayi must have played a major role in the evolution of Fe’i cultivars, though he did not rule out an interspecific origin. Cheesman (1947) noted similarities between the Fe’i banana and Musa lolodensis, another

Plate 1.29.  Bunch of a black-pigmented Musa balbisiana (BBw, accession ‘Pisang Klutuk Wulung’) growing in a germplasm collection in Guadeloupe (photo: J.W. Daniells, QDAF).

species in the Australimusa section from New Guinea. Close links have been demonstrated between three Fe’i cultivars and M. lolodensis, using RFLP analysis. This led Jarret et al. (1992) to speculate that the section may be derived solely from this species. However, work by Carreel (1995) indicated that the Fe’i banana may have an interspecific origin as the nuclear genome of some of the cultivars she analysed were close to M. maclayi, some close to M. lolodensis and some close to Musa peekelii, which is yet another species in the Australimusa section found in Papua New Guinea. She also discovered that some cultivars are diploid whilst others are triploid. It seems very likely that Fe’i cultivars originated in the New Guinea–Solomon Islands area and spread eastwards across the Pacific with migrating Polynesians (MacDaniels, 1947). The Fe’i cultivars seem highly resistant to diseases of the foliage, but little is known about their reaction to root diseases or their response to viruses and bacteria. This lack of knowledge is a reflection of the relatively unimportant role that these types now play as foodstuff in most

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Plate 1.30.  ‘Pisang Tongka Langit Alifuru’ (Fe’i cultivar with upright bunch) growing on Seram Island, Maluku Province, Indonesia (photo: J.W. Daniells, QDAF).

countries where they are still cultivated. However, in recent years there has been a revival of interest after the fruit of some clones was discovered to contain high levels of provitamin A carotenoids (Englberger et al., 2003). High concentrations correlated with orange-coloured pulp. The fruit of the cultivar ‘Utin Iap’ of Pohnpei was determined to have levels of provitamin A 100 times greater than Cavendish (Englberger et al., 2006). Importance of banana classification to banana pathologists Often in the past, research has been conducted on incorrectly identified germplasm, which has led to much confusion in the scientific literature. It is important that banana germplasm that is the subject of research be accurately identified by scientists and agriculturists. Only then can valid comparisons be made between the results of work undertaken at different times and at different locations.

One of the main challenges facing banana taxonomists today is to resolve synonymy among the many different names for banana clones and develop a system whereby cultivars can be easily identified. This would enable research workers to define their material more accurately. It is just as important for plant pathologists to know the correct identity of material being tested or screened for disease resistance as it is for them to know the correct identity of the pathogen. Photographs of the banana host being assessed for disease would help taxonomic specialists to confirm the clone’s identity. With this knowledge, the reaction of a particular clone (with a particular genetic composition) to a disease can be put into perspective vis-à-vis the reaction of other clones. An attempt has been made in this volume to provide information on the reaction of the different types of banana to each disease. Inevitably, more information is available on the responses of cultivars to serious and widespread diseases that have been well studied than on host responses to minor diseases of limited distribution. As mentioned previously, to help the reader place germplasm named in the text of this publication into perspective, the name of the genomic group, followed by the best-known synonym or subgroup, has been placed in parentheses after the cultivar name at appropriate places throughout the text. Bioversity International has recently developed an online Banana Cultivar Checklist on its ProMusa website that has the names of cultivars, their synonyms and subgroups.

Abacá Abacá is indigenous to the Philippines, where most is grown, and is often called ‘Manila hemp’, a name given to it by early European traders, who found it for sale in the Manila market. Abacá produces the strongest of the cordage fibres and is used to make the best grades of commercial cables and ropes. Because it has a high degree of resistance to sea water and a low degree of swelling when wet, it is particularly suited to marine cordage. Large amounts are also pulped and used to make high-quality paper and specialties such as tea bags, paper sacks and movable walls



Introduction to Banana, Abacá and Enset 33

for Japanese houses (Purseglove, 1972). More recently, it is being used in the automobile industry for dashboards and car interiors. Abacá grows best in the wet tropics where the annual precipitation is 2000–3200 mm, spread evenly throughout the year. The average annual temperature should be about 27°C, but not below 21°C, and the relative humidity around 80%. Abacá thrives in well-drained, deep, fertile soil, rich in humus and potash. Although these soils have relatively high fertility levels, continuous cropping of abacá leads eventually to yield declines because of the rapid rate of nutrient uptake by plants. In the Philippines, the crop is usually grown on land below 500 m (Berger, 1969; Purseglove, 1972). The fruit of abacá, which is about 7.5 cm in length, is inedible, containing numerous black seeds. Bunches are also small (Plate 1.31). The plant has an underground rhizome with numerous small roots, which do not penetrate far into the soil. Erect pseudostems arise from the rhizome and reach a height of 5–8 m. As with banana, pseudostems consist of thickened, clasping leaf bases. The fibre from the outer sheaths is coarser,

Plate 1.31.  Bunch of Musa textilis (abacá) in a germplasm collection in Guadeloupe (photo: J.W. Daniells, QDAF)

stronger and darker in colour than the innermost fibre, which is whiter and weaker. Planting material may be either suckers or whole corms or pieces of corm with a vegetative bud. However, suckers are rarely used, because of the difficulty of transport. True seed can also be planted, but plants take longer to mature. ‘Seed’ pieces, the equivalent of banana bits, are planted 2.5–3.0 m apart in holes/furrows and covered with 5–10 cm of soil. Young plants may be partially shaded for protection from excessive heat. The date of the first harvest depends on the cultivar, soil conditions and climate, but fullgrown pseudostems can be harvested from 18 to 24 months after planting. After the first harvest, two or four pseudostems can be harvested from each mat every 4–6 months. Yields are initially small, but reach a maximum after 4–5 years, when 12.5 t of dry fibre/ha can be obtained. Yields decline after 7–8 years and the crop is replanted after 10–15 years. Optimum cutting time is just before the flag leaf emerges. The flag leaf is the rudimentary and very small leaf that precedes the appearance of the inflorescence and its emergence can be anticipated because plant growth slows and leaf blades gradually shorten. Abacá is harvested by first trimming off the leaves and then cutting the pseudostem close to the ground. The percentage of fibre in a pseudostem is 1.5–3.0% of the weight. Fibre is extracted as soon after harvest as possible by hand stripping, spindle stripping or using a decorticating machine. Extracted fibre is either sun-dried in the open or air-dried in sheds. It is then tied in bundles, graded and baled for export. Abacá fibre consists of collections of sclerenchyma cells about 2.5–12 mm long. Strands of fibre can reach a length of 1.5–3.0 m. They are three times as strong as cotton and twice as strong as sisal fibres. Low-quality fibre comes from plants harvested too young or too old. Abacá fibre deteriorates during prolonged storage. The main fungi responsible have been identified as Aspergillus fumigatus and Chaetomium funicolum (Purseglove, 1972). Abacá is traditionally a crop of the Philippines where Musa textilis occurs in nature. Here, the bulk of the crop is grown in the eastern Visayas and Bicol regions (E.O. Lomerio, Legazpi City, 1996, personal communication). In 2013, 65,000 t were produced on 138,250 ha. Ecuador was the next largest producer, with 36,500 t

34

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grown on 25,300 ha. Minor suppliers are Costa Rica with 1225 t, Indonesia with 600 t and Equatorial Guinea with 250 t (FAO, 2016). More than 100 cultivars of abacá have been identified in the Philippines, but only about 20 are of commercial significance (Anunciado et al., 1977). These are listed in Table 1.9. Cultivars are distinguished by the colour and shape of their flowers and pseudostem, yield and quality of fibre and resistance to disease. Some, like ‘Itom’, are natural hybrids of M. textilis × M. balbisiana and are used to make ‘Canton fibre’ (Purseglove, 1972).

Enset Enset (Fig. 1.6) is cultivated in Ethiopia to the south and southwest of Addis Ababa (Fig. 1.7) contributing to the food security of many in this densely populated area. The enset plant provides a starchy foodstuff and animal feed and is a source of fibre for making rugs, sacks, bags and ropes. In addition, fresh leaves are used to wrap food and serve as plates. Dried petioles and midribs are burnt as fuel and their pulp is utilized as cleaning rags and brushes, baby nappies and cooking-pot stands. Certain cultivars and parts of enset are used medicinally to treat humans and livestock to heal wounds and bone fractures, as a treatment for some childbirth problems and as a cure for diarrhoea. As an abortifacient, it is also utilized in birth control (ECA, 1996; Brandt et al., 1997).

Enset forms an important part of the diet of about 17.5 million people, which represents nearly 20% of the Ethiopian population. Enset farming is believed to be one of Africa’s few surviving examples of indigenous, sustainable agriculture, which has evolved over hundreds of years. Enset is probably one of the oldest useful plants in Africa and has magic–religious significance in many areas. The Ari people preserve enset genetic resources by a highly ritualized, ex situ conservation of wild plants, which maintains the genetic diversity of landraces through a constant gene flow between wild and cultivated plants (Shigeta, 1990). The system of cultivation, by planting enset mainly around the homestead, coupled with a continuous use of manure and household refuse, enables farmers to grow enset in the same plot for generations without the aid of chemical fertilizers. Enset is resistant to climatic and environmental fluctuations and produces edible products that can be stored for months, if not years. It is a high-yielding crop. Only 42 mature plants grown on a small plot of land can support the annual food demand of a household of seven (Alemu and Sandford, 1991; Kippe, 2002).

Midrib Leaf

Table 1.9.  Abacá cultivars recommended for growing in the Philippines (information from E.O. Lomerio, Legazpi City, 1996, personal ­communication). Region

Cultivar

Visayas Mindanao

‘Inosa’, ‘Linawaan’, ‘Linino’, ‘Linlay’ ‘Tangoñgon’, ‘Maguindanao’, ‘Bongolanon’ ‘Itom’, ‘Sogmad’, ‘Tinawagan Pula’, ‘Tinawagan Puti’, ‘Lausigon’, ‘Abaub’, ‘Casilihon’, ‘Lausmag 24’, ‘Itolaus 45’, ‘M. textilis 52’, ‘M. textilis 51’, ‘M. textilis 50’ ‘Tinawagan Pula’, ‘Tinawagan Put’i’, ‘Sinibuyas’, ‘Putian’

Bicol

Southern Tagalog

Infloresence

Pseudostem

Leaf sheath

Corm Root

Fig. 1.6.  Diagrammatic representation of enset (Ensete ventricosum) (from Brandt et al., 1997).



Introduction to Banana, Abacá and Enset 35

Addis Ababa

xx

ETHIOPIA

xx

GURAGE

OROMO

HADIYA

xxx

x

L. Zway L. Abiyata

xxxxx

L. Lanxx xxxxxx L. Awassa Awassa

KAMBATA

Co

jeb

SHEKO

WOLAYTA

L. Abaya

xxx

SIDAMA GAMO ARI L. Gramo

n Weil x xx

Om

o

Areas of enset cultivation

0

100

200 km

L. Chew Bahir L. Turkana

Fig. 1.7.  Enset-growing areas in southwest Ethiopia with names of enset-cultivating ethnic groups (from Brandt et al., 1997).

Enset is planted at altitudes ranging from 1200 m to 3100 m, but grows best at elevations of 2000–2750 m. The average temperature where it is grown varies from 10°C to 21°C, with a relative humidity range of 63–80%. Like ­banana, enset does not tolerate freezing and frost damage is often observed above 2800 m. The annual rainfall of most growing areas is 1100–1500 mm, with the majority falling between March and September. Irrigation is used to supplement rainfall at altitudes below 1500 m, where rainfall decreases and there is a greater evaporative demand. Enset grows in most soil types if they are sufficiently fertile and well drained. Cattle manure is used as the main o ­ rganic fertilizer.

Ideal soils for enset cultivation have a pH of 5.6–7.3 and contain 2–3% organic matter (Brandt et al., 1997). Enset development is similar to banana in that leaves are successively produced by an apical meristem until flowering. The plant is normally 5–7 m in height and the pseudostem has a circumference of 0.5–3.0 m at maturity. It has a shallow rooting system, which extends 2–3 m. It takes about 6–7 years at the optimum ­altitude for growth for an enset plant to reach maturity and form an inflorescence. The fruit, which is seedy and leathery, is inedible and varies in size and weight between clones. After ­fruiting, the plant declines.

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Although enset only reproduces by seed in nature, the Ethiopian people have developed a technique by which vegetative shoot formation is induced, thus enabling superior plants to be clonally propagated. The central apical meristem at the top of the rhizome of a 3–4-year-old plant is hollowed out and the rhizome is then usually split into two equal parts, which are buried and covered with manure. About 40– 200 buds are produced by this method and the suckers emerge from the soil after 4–6 weeks (Demeke, 1986). The suckers are later severed from the corm and planted in nurseries. Management varies considerably after this stage, depending on the ethnic community and household requirements. Plants may be transplanted up to four times at ever wider spacing. Some may be harvested young, while others are left for harvesting when mature. Generally, a leaf canopy is maintained that covers the soil for most of the year. Enset may grow alone or in mixed plantings with other crops, such as maize, cabbage, coffee and citrus (Brandt et al., 1997).

As the developing inflorescence uses up starch reserves in the plant (Purseglove, 1972) farmers usually harvest enset at flower emergence (Yemataw et  al., 2012). The rhizome is grated using either a special hardwood implement or a bamboo scraper (Plate 1.32). After decortication, the pulp of the leaf sheaths that make up the pseudostem is also removed. Enset fibre is a by-product of this process. A fermentation process that lasts for a month or more takes place in a pit lined with enset leaves and transforms the rhizome and leaf sheath scrapings into a soft starchy mass called kocho. This is the most important of the different food products made from enset and it can be kept for many months or several years. The amount of kocho obtained is dependent on the number of transplants before harvesting and the space between each plant, with a maximum experimental yield of 33.2 t/ha/year ­being reported by Tsegaye and Struik (2001). These authors calculated that the edible yield of enset in terms of weight and energy was

Plate 1.32.  Women, who traditionally process enset (Ensete ventricosum) in Ethiopia, abrading decorticated rhizomes with hardwood graters (foreground) and a leaf sheath with a bamboo scraper (background) near Jima in the Kefa district (photo: G. Blomme, BI).



Introduction to Banana, Abacá and Enset 37

much higher than those of other carbohydrate-­ rich root and tuber crops or cereals cultivated in Ethiopia. Another food product is bulla, which is a porridge, pancake or dumpling made from dehydrated juice obtained from the scrapings of the leaf sheath, peduncle and grated corm. Amicho is the boiled rhizome, usually from a young plant (Brandt et al., 1997).

Acknowledgements The advice given by Edmond De Langhe, JeanPierre Horry, Rony Swennen and Deborah Karamura on some of the finer points of banana ­classification is much appreciated. Guy Blomme, Mulugeta Diro, Zerihun Yemataw and Temesgen Addis are thanked for providing new information on the cultivation of enset in Ethiopia.

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De Langhe, E., Karamura, D. and Mbwana, A. (2001) Tanzania Musa expedition 2001. INIBAP, Kampala, Uganda, 106 pp. Demeke, T. (1986) Is Ethiopia’s Ensete ventricosum crop her greatest potential food? Agriculture International 38, 362–365. ECA (1996) Manual on Production and Utilization of Enset (Ensete ventricosum) in South and Southwestern Ethiopia, Vol. 2. Joint publication of the Southern Nations, Nationalities and Peoples’ Regional Government and the Economic Commission for Africa. [In Amharic language with English summary.] United Nations Development Program/Economic Commission for Africa, Addis Ababa, Ethiopia. Englberger, L., Schierle, J., Marks, G.C. and Fitzgerald, M.H. (2003) Micronesian banana, taro, and other foods: Newly recognized sources of provitamin A and other carotenoids. Journal of Food Composition and Analysis 16, 3–19. Englberger, L., Schierle, J., Aalbersberg, W., Hofmann, P., Humphries, J. et  al. (2006) Carotenoid and ­vitamin content of Karat and other Micronesian banana cultivars. International Journal of Food and Nutrition. 57, 399–418. FAO (2014) Banana market review and banana statistics 2012–2013 from FAO website. FAO, Rome. Available at: www.fao.org/docrep/019/i3627e/i3627e.pdf (accessed 30 May 2018). FAO (2015) Statistics from FAO website. Available at : http://faostat3.fao.org/. FAO (2016) Statistics from FAO website. Available at: http://faostat3.fao.org/download/Q/QC/E). Fortescue, J.A., Turner, D.W. and Romero, R. (2011) Evidence that banana (Musa spp.), a tropical monocotyledon, has a facultative long-day response to photoperiod. Functional Plant Biology 38, 867–878. Hakkinen, M. (2013) Reappraisal of sectional taxonomy in Musa (Musaceae). Taxon 62(4), 809–813. Horry, J.P. and Jay, M. (1990) An evolutionary background for bananas as deduced from flavonoids diversification. In: Jarret, R.L. (ed.) Identification of Genetic Diversity in the Genus Musa, Proceedings of an International Workshop held at Los Baños, Philippines, 5–10 September 1988. INIBAP, Montferriersur-Lez, France, pp. 41–55. Horry, J.P., Ortiz, R., Arnaud, E., Crouch, J.H., Ferris, R.S.B. et al. (1997) Banana and plantain. In: Fuccillo, D., Sears, L. and Stapleton, P. (eds) Biodiversity in Trust, Conservation and Use of Plant Genetic Resources in CGIAR Centres. Cambridge University Press, Cambridge, UK, pp. 67–81. Horry, J.P., Doležel, J., Doleželova, M. and Lysak, M.A. (1998) Do natural A × B tetraploid bananas exist? Infomusa 7(1), 5–6. INIBAP/IPGRI (2000) Bananas/Les Bananes. International Network for the Improvement of Banana and Plantain/International Plant Genetic Resources Institute. INIBAP, Montpellier, France. Available at: http://www.bioversityinternational.org/e-library/publications/detail/bananasles-bananes/ (accessed 30 May 2018). Jarret, R.L. and Litz, R.E. (1986) Determining the interspecific origins of clones within the ‘Saba’ cooking banana complex. Horticultural Science 21, 1433–1435. Jarret, R.L., Gawel, N., Whittemore, A. and Sharrock, S. (1992) RFLP based phylogeny of Musa species, Papua New Guinea. Theoretical and Applied Genetics 84, 579–584. Jenny, C.F., Carreel, F. and Bakry, F. (1997) Revision on banana taxonomy: ‘Klue Tiparot’ (Musa sp.) reclassified as a triploid. Fruits 52, 83–91. Karamura, D.A. (1999) Numerical taxonomic studies of the East African highland Bananas (Musa AAA – East Africa) in Uganda. Submitted PhD thesis, Department of Agricultural Botany, University of Reading, January 1998. INIBAP, Montpellier, France, 192 pp. Karamura, D.A. and Karamura, E.B. (1994) A Provisional Checklist of Banana Cultivars in Uganda. INIBAP, Montpellier, France, 28 pp. Karamura, D., Kitavi, M., Nyine, M., Ochola, D., Ocimati, W., Muhangi, S., Talengera, D. and Karamura, E. (2016) Genotyping the local banana landrace groups of East Africa. Acta Horticulturae 1114, 67–73. Kepler, A. K. and Rust, F.G. (2011) The world of bananas in Hawaii: then and now – Traditional Pacific & global varieties, cultures, ornamentals, health & recipes. Pali-O-Waipiʻo Press, Haiku, Hawaii, 586 pp. Kippe, T. (2002) Five Thousand Years of Sustainability? A Case Study of Gedeo Land Use Southern Ethiopia. Tree Book 5, Treemial Publishers, Heelsum, The Netherlands. Kitavi, M., Downing, T., Lorenzen, J., Karamura, D. Onyango, M., Nyine, M. Ferguson, M. and Spillane, C. (2016) The triploid East African highland banana (EAHB) genepool is genetically uniform arising from a single ancestral clone that underwent population expansion by vegetative propagation. Theoretical and Applied Genetics 139, 547–561. Langdon, R. (1993) The banana as a key to early American and Polynesian history. Journal of Pacific History 28, 15–35.



Introduction to Banana, Abacá and Enset 39

Lebot, V., Meilleur, B.A. and Manshardt, R.M. (1994) Genetic diversity in Eastern Polynesian cultivated bananas. Pacific Science 48, 16–31. Lescot, T. (2015) Genetic diversity of the banana. FruiTrop, 231, 98–102. MacDaniels, L.H. (1947) A study of the Fe’i banana and its distribution with reference to Polynesian migrations. Bernice P. Bishop Museum Bulletin 190, 3–56. Noyer, J.L., Causse, S., Tomekpe, K., Bouet, A. and Baurens, F.C. (2005) A new image of plantain diversity assessed by SSR, AFLP and MSAP markers. Genetica 124, 125–136. Nwakanma, D.C., Pillay, M., Okoli, B.E. and Tenkouane, A. (2003) Sectional relationships in the genus Musa L. inferred from the PCR-RFLP of organelle DNA sequences. Theoretical and Applied Genetics 107, 850–856. Pascua, O.C. and Espino, R.R.C. (1987) Taxonomic classification of Philippine Bananas. In: Persley, G.J. and De Langhe, E.A. (eds) Banana and Plantain Breeding Strategies. Proceedings of an International Workshop held at Cairns, Australia, 13–17 October 1986. ACIAR Proceedings No. 21, Australian Centre for International Agricultural Research, Canberra, Australia, pp. 161–164. Paxton, J. (1837) Musa Cavendishii. In: Paxton’s Magazine of Botany and Register of Flowering Plants, Volume the Third. W.S. Orr and Co., London, pp. 51–52. Perrier, X., De Langhe, E., Donohue, M., Lentfer, C., Vrydaghs, L. et al. (2011) Multidisciplinary perspectives on banana (Musa spp.) domestication. Proceedings of the National Academy of Science of the United States of America 108, 11311–11318. Pope, W.T. (1926) Banana culture in Hawaii. Bulletin No. 55, Hawaii Agricultural Experiment Station, Kapaa, Hawaii. Purseglove, J.W. (ed.) (1972) Musaceae. In: Tropical Crops – Monocotyledons. Longman, London, pp. 343–384. Richardson, D.L., Hamilton, K.S. and Hutchison, D.J. (1965) Notes on bananas. I. Natural edible tetraploids. Tropical Agriculture (Trinidad) 42, 125–137. Robinson, J.C. and Galán Saúco, V. (2010) Bananas and Plantains. Crop Production Science in Horticulture 19, CAB International, Wallingford, UK, 311 pp. Shepherd, K. (1957) Banana cultivars of East Africa. Tropical Agriculture (Trinidad) 34, 277–286. Shepherd, K. (1990) Observations on Musa taxonomy. In: Jarret, R.L. (ed.) Identification of Genetic Diversity in the Genus Musa: Proceedings of an International Workshop held at Los Baños, Philippines, 5–10 September 1988. INIBAP, Montferrier-sur-Lez, France, pp. 158–165. Shepherd, K. and Ferreira, F.R. (1984) The PNG Biological Foundation’s Banana Collection at Laloki, Port Moresby, Papua New Guinea. IBPGR Regional Committee for Southeast Asia Newsletter 8(4), 28–34. Shigeta, M. (1990) Folk in-situ conservation of ensete (Ensete ventricosum [Welw.] E.E. Cheesman): towards the interpretation of indigenous agricultural science of the Ari, southwestern Ethiopia. Kyoto University, African Studies Monographs 10(3), 93–107. Silayoi, B. and Chomchalow, N. (1987) Cyotaxonomic and morphological studies of Thai banana cultivars. In: Persley, G.J. and De Langhe, E.A. (eds) Banana and Plantain Breeding Strategies. Proceedings of an International Workshop held at Cairns, Australia, 13–17 October 1986. ACIAR Proceedings No. 21, Australian Centre for International Agricultural Research, Canberra, Australia, pp. 157–160. Silayoi, B. and Sompen, P. (1991) Chromosome number and karyotypes of some Thai bananas. Kasetsart Journal (Natural Science) 25, 400–407. Simmonds, N.W. (1962) The Evolution of the Bananas. Longmans, Green, London, 170 pp. Simmonds, N.W. and Shepherd, K. (1955) The taxonomy and origins of the cultivated bananas. Journal of the Linnean Society of London 55, 302–312. Simmonds, N.W. and Weatherup, S.T.C. (1990) Numerical taxonomy of the wild bananas (Musa). New Phytologist 115, 567–571. Smith, M.K., Whiley, A.W., Searle, C., Langdon, P.W., Schaffer, B. and Pegg, K.G. (1998) Micropropagated bananas are more susceptible to Fusarium wilt than plants grown from conventional material. Australian Journal of Agricultural Research 49, 1133–1139. Stover, R.H. and Simmonds, N.W. (1987) Bananas, 3rd edn. Longman Scientific and Technical, Harlow, UK, 468 pp. Swennen, R. (1990) Limits of morphotaxonomy: names and synonyms of plantains in Africa and elsewhere. In: Jarret, R.L. (ed.) Identification of Genetic Diversity in the Genus Musa: Proceedings of an International Workshop held at Los Baños, Philippines , 5–10 September 1988. INIBAP, Montferrier-­ sur-Lez, France, pp. 172–210.

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Tezenas du Montcel, H. (1979) Les plantains du Cameroon. Propositions pour leur classification et dénominations vernaculaires. Fruits 34, 83–97. Tezenas du Montcel, H. and Devos, P. (1978) Proposal for establishing a plantain determination card. Paradisiaca (Ibadan, Nigeria) 3, 14–17. Tezenas du Montcel, H., De Langhe, E. and Swennen, R. (1983) Essai de classification des bananiers plantain (AAB). Fruits 6, 461–474. Tsegaye, A. and Struik, P.C. (2001) Enset (Ensete ventricosum (Welw.) Cheesman) kocho yield under different crop establishment methods as compared to yields of other carbohydrate-rich food crops. Netherlands Journal of Agricultural Science 49, 81–94. Uma, S. and Sathiamoorthy, S. (2002) Names and Synonyms of Bananas and Plantains of India. ­National Research Centre for Banana, Indian Council of Agricultural Research (ICAR), Tiruchirappalli, India, 62 pp. Vakili, N.G. (1965) Fusarium wilt resistance in seedlings and mature plants of Musa species. Phytopathology 55, 135–140. Vakili, N.G. (1968) Response of Musa acuminata species and edible cultivars to infection by Mycosphaerella musicola. Tropical Agriculture (Trinidad) 45, 13–22. Valmayor, R.V., Espino, R.R.C. and Pascua, O.C. (2002) The Wild and Cultivated Bananas of the Philippines. Philippine Agriculture and Resources Research Foundation Inc. (PARRFI) and the Bureau of Agricultural Research (BAR), Los Baños, Philippines, 242 pp. Wardlaw, C.W. (1961) Banana Diseases Including Plantains and Abaca. Longmans, Green, London, 648 pp. Wong, C., Kiew, R., Argent, G., Set, O., Lee, S.K. and Gan, Y.Y. (2002) Assessment of the validity of the sections in Musa (Musaceae) using AFLP. Annals of Botany 90, 231–238. Yemataw, Z., Mohamed H., Diro M., Addis T., Blomme G. (2012) Genetic variability, inter-relationships and path analysis in Enset (Ensete ventricosum) clones. The African Journal of Plant Science and ­Biotechnology 6, 21–25.

2 

Fungal Diseases of the Foliage

Sigatoka Leaf Spots Overview D.R. Jones Three similar major leaf spot diseases have been recognized on banana. These are black leaf streak, Sigatoka leaf spot and eumusae leaf spot (Crous et al., 2003; Jones, 2003). They are described in this publication under the heading Sigatoka leaf spots. All would appear to have arisen in the Southeast Asian–Australasian region, which is also the centre of diversity of banana. All have been found to be phylogenetically distinct, but derived from a common ancestor (Arzanlou et al., 2010). The first of these diseases to have been documented was called Sigatoka leaf spot. This became important in the 1930s when it began to spread around the world. The second, which eventually replaced Sigatoka leaf spot in most regions as the most serious problem on banana leaves, was black leaf streak. This disease began to spread globally from Asia and the Pacific region in the 1960s. The third, known as eumusae leaf spot, was only identified as a different leaf spot, which was present mainly in parts of South and Southeast Asia, in the 1990s. It seems likely that it may have been misidentified as Sigatoka leaf spot on symptomology

and on morphological characteristics, which are close, for many years (Crous et  al., 2003; Jones, 2003). This disease has also been found outside its suspected centre of origin. Chang et al. (2016) presented images of the symptoms of the three leaf spot diseases. They also sequenced and analysed the genomes of the causal agents of eumusae leaf spot and Sigatoka leaf spot and compared them with the available genome sequence of the pathogen that causes black leaf streak. The results of this work are reported below in the causal agent section of the description of eumusae leaf spot. In the dual nomenclature system with different names for the sexual and asexual morphs, the three Sigatoka leaf spot pathogens were placed in the Mycosphaerella and Pseudocercospora genera. The move to a one species name for a fungus rather than two has resulted in the three pathogen species now being accommodated in the genus Pseudocercospora with their Mycosphaerella sexual morphs becoming synonyms.

Black Leaf Streak M. Guzmán, L. Pérez-Vicente, J. Carlier, C. Abadie, L. de Lapeyre de Bellaire, F. Carreel, D.H. Marín, R.A. Romero, F. Gauhl, C. Pasberg-Gauhl and D.R. Jones

© D.R. Jones 2019. Handbook of Diseases of Banana, Abacá and Enset (D.R. Jones)

41

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Introduction L. Pérez-Vicente, M. Guzmán, C. Pasberg-Gauhl, F. Gauhl and D.R. Jones Black leaf streak, or black Sigatoka, as it is known in the Latin American–Caribbean region, is regarded as the most economically important leaf disease of banana and one of the most serious threats to global food security (Pennisi, 2010). Symptoms are similar to Sigatoka leaf spot, but the pathogen is more virulent and affects a wider range of banana genotypes. Black leaf streak develops much more rapidly than Sigatoka leaf spot and causes more severe defoliation. Symptoms of black leaf streak are also similar to eumusea leaf spot, making identification difficult. In many coastal locations and at altitudes of up to 1000 m above sea level, black leaf streak has replaced Sigatoka leaf spot as the predominant disease within a few years from its introduction in these areas. Black leaf streak destroys banana leaves (Plate 2.1), which leads to a reduction in yield and quality and induces premature ripening

of the fruit. Since it was first recognized in the South Pacific, black leaf streak has spread to most of the major banana-growing regions of the world and is still spreading. Countries where the disease occurs are listed in Table 2.1. The year of the first record in Table 2.1 is not necessarily the year of introduction. For instance, the first published report of black leaf streak in Papua in Indonesia was 2000, but it is likely to have been present from at least the 1960s and perhaps even earlier. Distribution in Asia, Australia and the Pacific Black leaf streak was first recognized in February 1963 in the Sigatoka district of the island of Viti Levu in Fiji, where it was said to be spreading rapidly (Rhodes, 1964). The disease was also described as replacing Sigatoka leaf spot wherever it became established (Leach, 1964a, b). Surveys conducted between 1964 and 1967 showed black leaf streak to be widespread in the Pacific and parts of the Pacific Rim. According to Johnston (1965) and Graham (1968), the disease was present in the Federated States of Micronesia, New Caledonia, Papua New Guinea, Philippines,

Plate 2.1.  ‘Grande Naine’ (AAA, Cavendish subgroup) in centre foreground with no functional leaves in a plantation in Costa Rica left unprotected by fungicides. Its bunch has been drastically pruned in an attempt to accelerate fruit development and filling (photo: M. Sánchez and M. Guzmán, CORBANA).



Fungal Diseases of the Foliage

43

Table 2.1.  Countries where black leaf streak has been found and year of first official record. Region/country Africa   Benin   Burundi   Cameroon     Southeast     Southwest    Central African Republic   Comoros    Congo (Democratic Republic of the)     Highlands     Lowlands    Congo (Republic of the)    Côte d’Ivoire   Ethiopia   Gabon   Ghana   Kenya   Malawi   Mayotte   Nigeria   Rwanda    São Tomé   Tanzania     Pemba     Zanzibar   Togo   Uganda   Zambia Latin America and Caribbean   Bahamas   Belize   Bolivia   Brazil   Colombia    Costa Rica   Cuba   Dominica    Dominican Republic   Ecuador    El Salvador   Grenada   Guadeloupe   Guatemala   Guyana   Haiti   Honduras   Jamaica   Martinique   Mexico

Year of detection

Reference

1993 1987

Jones and Mourichon (1993) Sebasigari and Stover (1988)

1980 1983 1996 1993

Tezenas du Montcel (1982) Wilson and Buddenhagen (1986) X. Mourichon, France, 1996, pers. comm. Jones and Mourichon (1993)

1987 1988 1985 1985 2016 1978 1986 1988 1990 1993 1986 1986 1980 1987 1987 1987 1988 1990 1973a

Sebasigari and Stover (1988) Mobambo and Naku (1993) Mourichon (1986) Mourichon and Fullerton (1990) Gurmu et al. (2017) Frossard (1980) Wilson (1987) Kung’U et al. (1992) Ploetz et al. (1992) X. Mourichon, France, 1996, pers. comm. Wilson and Buddenhagen (1986) Sebasigari (1990) Frossard (1980) Sebasigari and Stover (1988) Dabek and Waller (1990) Sebasigari and Stover (1988) Mourichon and Fullerton (1990) Tushemereirwe and Waller (1993) Raemaekers (1975)

2004 1975 1996 1998 1981 1979 1990 2012 1996

Ploetz (2004) Stover (1980b) Tejerina (1997) Cordeiro et al. (1998) Merchan (1990) Jaramillo (1979), Stover (1980b) Vidal (1992) Pérez-Vicente (2012b) X. Mourichon, France, 1996, pers. comm.; Polanco et al. (2002) Mourichon and Fullerton (1990) Mourichon and Fullerton (1990) IPPC (2007) Hubert (2012) Stover (1980b) ISID (2009a), (2010a) Pollard (1998) Stover and Dickson (1976) A. Johanson, Chatham, 1996, pers. comm. Ioos et al., (2011) Stover and Simmonds (1987) Continued

1986 1990 2006 2012 1977 2008 1998 1972 (1969)b 1995 2010 1980

44

Chapter 2

Table 2.1. Continued. Region/country

Year of detection

Reference

  Nicaragua   Panama   Peru    Puerto Rico    St Vincent & The Grenadines    St Lucia    Trinidad & Tobago    (USA) Florida   Venezuela Asia   Bhutan   China     Hainan     Guangdong      Yunnan   Indonesia     Halmahera     Java     Sumatra     Kalimantan

1979 1981 1994 2004 2009 2010 2003 1998 1991

Stover (1980b) Stover (1987) Fujimori and Vázquez (1995) Almodóvar (2004), Irish et al., 2006) ISID (2009b) ISID (2010b) Fortune (2005) Ploetz and Mourichon (1999) INIBAP (1994)

1985

Peregrine (1989)

1980 1990 1993

Stover and Simmonds (1987) Mourichon and Fullerton (1990) Jones and Mourichon (1993)

1970 1969 1993 1996

    Papua   Malaysia      West Malaysia        Johore        Langkawi      East Malaysia (Sarawak)

2000

Stover (1978) Reddy (1969) Jones and Mourichon (1993) X. Mourichon, France, 1996, pers. comm. Davis et al. (2000)

  Philippines     Luzon     Mindanao   Singapore   Taiwan   Thailand   Vietnam Australasia/Oceania    American Samoa   Australia      Torres Strait Islands     Cape York Peninsula      Tully Valley (eradicated)    Cook Islands   Fiji    French Polynesia    Hawaii (USA)   Micronesia    New Caledonia   Niue    Norfolk Island    Papua New Guinea    Solomon Islands

1965 1993 1995 1996

Graham (1968) Jones and Mourichon (1993) Infomusa (1995) X. Mourichon, France, 1996, pers. comm.

1964 1965c 1964–1967 1927c 1969 1993

Hapitan and Reyes (1970) Stover (1978) Graham (1968) Stover (1978) Reddy (1969) Jones and Mourichon (1993)

1975 1981 1981 2001 1976 1963 1964–1967 1958d 1964–1967 1964–1967 1976 1980 1957 (1951)d 1957c

Firman (1975) Jones and Alcorn (1982) Jones and Alcorn (1982) Peterson et al. (2003) Firman (1975) Rhodes (1964) Graham (1968) Stover (1978) Graham (1968) Graham (1968) Stover (1976) Jones (1990b) Stover (1976) Meredith (1970) Stover (1976) Continued



Fungal Diseases of the Foliage

45

Table 2.1. Continued. Region/country

Year of detection (1946) 1965 1964–1967 1996 d

  Tonga   Vanuatu    Wallis and Futuna Islands    Western Samoa

1965

Reference Meredith (1970) Johnston (1965) Graham (1968) X. Mourichon, France, 1996, pers. comm. Johnston (1965)

Authenticity of this record has been challenged. Symptoms of black leaf streak on colour transparencies. c Date from herbarium material examined. d Date black leaf streak recognized in hindsight. pers. comm. = personal communication. a b

Western Samoa, Singapore, Solomon Islands, Tahiti, Taiwan, Tonga, Vanuatu and West Malaysia. Meredith and Lawrence (1969) identified black leaf streak in the Hawaiian Islands and Stover (1976) reported the presence of the disease in the additional Pacific locations of the Cook ­Islands and Niue. The widespread distribution of black leaf streak in and around the Pacific suggested that it had been present in the region for some considerable time before its discovery in Fiji in 1963 (Meredith, 1970; Stover 1978; Long, 1979). Trujillo and Goto (1963) described a variant of Sigatoka leaf spot in the Hawaiian Islands that had been observed as early as 1958. This description was very similar to that of black leaf streak, prompting Meredith and Lawrence (1969) and Meredith (1970) to speculate that the disease may have initially been misdiagnosed as Sigatoka leaf spot after its arrival. In 1970, black leaf streak was found throughout the Philippines on many different cultivars (Hapitan and Reyes, 1970). Evidence suggested that the disease had been present around Los Baños on the island of Luzon in the Philippines since 1964, but had most probably arrived even earlier. After examining old herbarium specimens of banana leaf spot, Stover (1976) believed that black leaf streak was present in Papua New Guinea in 1957 and in Taiwan as early as 1927. After analysing information on distribution and probable times of introduction to various countries, he believed that black leaf streak may have originated in the New Guinea–Solomon Islands area (Stover, 1978). Black leaf streak was first described in Australia in garden plots on islands in the Torres

Strait and at Bamaga on the tip of Cape York Peninsula in 1981 during the first plant disease survey of the area (Jones and Alcorn, 1982; Jones, 1990a). However, it seems likely that black leaf streak may have been present in this remote part of Queensland, which is in close proximity to Papua New Guinea, for many years prior to its discovery. Since 1981, the disease has been detected in stands of banana at a few isolated locations further south on Cape York Peninsula. In April 2001, black leaf streak was discovered in a commercial plantation near Tully in North Queensland, a region where approximately 95% of Australia’s bananas are produced. A successful and expensive eradication programme was undertaken and no new cases were reported after March 2002. This resulted in the reinstatement of the black leaf streak-free status for the region in 2005. The success of the eradication campaign was attributed to the very early detection of the initial outbreak, regular intensive surveys of plantation for symptoms, extensive de-leafing operations in the outbreak area and the unseasonably dry conditions experienced between ­August 2001 and April 2002. Other key factors were the full cooperation and participation of the local growers, and the deployment of a polymerase chain reaction (PCR)-based molecular diagnostic assay, which allowed large numbers of suspect leaf spot samples to be processed quickly (Peterson et al., 2003; Henderson et al., 2006). In 1980, black leaf streak was observed together with Sigatoka leaf spot on Hainan Island in China (Stover and Simmonds, 1987). Mourichon and Fullerton (1990) isolated the fungal pathogen from leaf samples collected in the Guangdong region and the disease was later

46

Chapter 2

confirmed in Yunnan Province (Jones and Mourichon, 1993). These reports indicate that black leaf streak is now distributed throughout southern China. Black leaf streak was first identified in Vietnam in 1993 (Jones and Mourichon, 1993), but it may have been present much earlier. It was again identified from specimens collected in 1994 when severe symptoms were observed on cultivars in the Cavendish subgroup in some, but not all, locations (Infomusa, 1994a). The disease has not yet been confirmed in Cambodia and Laos. The occurrence of black leaf streak was reported from Bhutan in 1985 (Peregrine, 1989), but it has not yet been recorded in Myanmar, ­India or Bangladesh. In Indonesia, the disease was first reported in the late 1960s from leaf spots seen on banana in the Bogor-Bandung area of Java (Reddy, 1969). However, the characteristic streak symptoms of the disease were not noticeable in the same area of Java in 1988. The first authenticated report of black leaf streak in Java was in 1996 (X. Mourichon, France, 1996, personal communication). The causal agent of black leaf streak has also been identified from leaf spot specimens collected on Halmahera Island in the Maluku group (Stover, 1978), in West Sumatra (Infomusa, 1993), in Kalimantan (X. Mourichon, France, 1996, personal communication) and in Papua (Irian Jaya) (Davis et al., 2000). The disease was recently reported as common in Sumatra (Sahlan and Soemargono, 2011). Reports in the literature suggest that black leaf streak has been present in Singapore, West Malaysia and Thailand for some time (Reddy, 1969; Stover, 1978). However, black leaf streak was not observed near Kuala Lumpur and ­Melaka in West Malaysia or near Bangkok and Pakchong in Thailand in 1988 (Jones, 1990b). Dur­ alaysia in ing a banana disease survey of West M 1993, black leaf streak was identified in leaf spot specimens collected in Johore State (Infomusa, 1993). The disease was later confirmed as present on Langkawi Island off the west coast of West Malaysia near the Thai border (Infomusa, 1995) and in Sarawak State in East Malaysia (X.  Mourichon, France, 1996, personal communication). However, black leaf streak was not identified from leaf spot specimens ­collected during a banana disease survey of ­Thailand in 1994 (X. Mourichon, France, 1996, personal communication) and was not recorded

during a more recent survey to identify leaf spot pathogens in banana plantations in northern Thailand (Kaewjan et al., 2012). Black leaf streak has not become the dominant leaf spot in Asia, as it has in the Pacific, ­Central and South America, the Caribbean and equatorial Africa. This phenomenon may be related to the varying degrees of resistance to black leaf streak in the diverse local banana ­cultivars and the competition from other leaf d ­ iseases, particularly eumusae leaf spot. The complex indigenous, non-pathogenic microbial communities of epiphytes of the banana phyllosphere, which would vary between locations, could also play a role in determining which leaf spot pathogen predominates in any one area (Blakeman and Fokema, 1982; Lindow and Brandl, 2003; Ceballos et al., 2012). Distribution in the Latin American–Caribbean region The history of the detection and spread of black leaf streak is well documented in the Americas. The first appearance of the disease outside Asia and the Pacific was in Honduras in 1972 (Stover and Dickson, 1976). It was found together with Sigatoka in the germplasm collection of the United Fruit Company at La Lima. This collection was used for Sigatoka reaction studies and was always left unsprayed. After detection, a survey of local banana fields was undertaken. Symptoms of black leaf streak, which were always mixed with Sigatoka spots, were present in 100 ha of banana under cultivation close to La Lima (Stover and Dickson, 1976). The first severe outbreak of black leaf streak occurred in December 1973–February 1974 and covered an area of 1200 ha. By August 1974, the disease, which became known in Honduras as Sigatoka negra or black Sigatoka, was present in scattered locations for 51 km along the Ulúa River. Stover and Dickson (1976) and Buddenhagen (1987) postulated that, when black leaf streak first became established in the Ulúa valley, the pathogen did not have a sufficient level of virulence to replace Sigatoka. However, there are no scientific data to support this hypothesis. As well as export banana cultivars in the Cavendish subgroup, cultivars in the Plantain subgroup (AAB) were attacked in coastal lowlands of Honduras. This characteristic distinguished



Fungal Diseases of the Foliage

black leaf streak from Sigatoka, which does not attack plantain at low altitudes. Between 1973 and 1980, serious epidemics of the disease occurred throughout Latin America. Black leaf streak appeared in Belize in 1975, Guatemala in 1977 and El Salvador, Nicaragua and Costa Rica in 1979 (Jaramillo, 1979; Stover, 1980a, b). In Costa Rica, the most probable source of inoculum for the black leaf streak outbreak in the San Carlos region was banana leaf trash from Honduras (Woods, 1980). At the time, plantains and green reject bananas from export plantations in Central America were regularly transported by road across international boundaries (Stover, 1980b). Trash banana and plantain leaves were used for padding and shading the shipments to reduce bruising and sunburn. Leaves affected by black leaf streak were thus moved from one country to another, spreading the disease. By 1981, black leaf streak was present in all countries between southern Mexico and Panama (Stover, 1987). Also in 1981, the disease reached the Uraba export banana production area in the lowlands of Colombia, where all banana plantations were attacked (Merchan, 1990). From there, the disease spread along the Pacific and Atlantic coasts of Colombia to where plantain is grown in association with coffee or cocoa at altitudes of between 1000 and 1700 m above sea level. Black leaf streak was first observed in ­plantain districts in 1988 at 1500 m (Merchan, 1990) and later at 1600 m (Belalcázar, 1991). In 1986, black leaf streak was detected in northern Ecuador and within 4 years was found in the southern banana production areas (Mourichon and Fullerton, 1990). The disease reached western Venezuela in 1991 (INIBAP, 1994) and by 1997 had spread to the eastern part of the country (Martinez et al., 1998). In 1994, black leaf streak reached Peru (Fujimori and Vázquez, 1995), where it spread rapidly along the Ucayali River and other Amazon tributaries, which served as trade routes facilitating the movement of fruit and suckers (L. Pérez-Vicente, Cuba, 1996, personal communication). Later, the disease was ­recorded in Bolivia (Tejerina, 1997) and Brazil (Cordeiro et  al., 1998). In Guyana, the disease was first found on the Essequibo Islands in West Demerara in 2008 (ISID, 2010a) and ­after 2 years it had spread to all parts of the country (R. Singh, Guyana, 2012, personal communication). Black

47

leaf streak was also reported to have first arrived in Guyane in 2008 (de Lapeyre de Bellaire et al., 2010). It is almost certainly present in Suriname, though there are no official records in the scientific literature. From first-record information and phylogenetic studies (Robert et  al., 2012), it has been determined that black leaf streak followed two distinct invasion pathways through the Caribbean. One began in Cuba (Vidal, 1992), where the ­disease was first noticed in December 1990. It spread quickly to the eastern part of the country, but took 3 years to invade the central and western areas. The disease was next reported in ­Jamaica in 1995 (A. Johansen, Jamaica, 1995, personal communication; Conie, 2001), then in Dominican Republic (X. Mourichon, France, 1996, personal communication; Polanco et  al., 2002), Florida in the USA (Ploetz and Mourichon, 1999), Bahamas (Ploetz, 2004), Haiti (Pollard, 1998) and Puerto Rico (Almodóvar, 2004; Irish et al. 2006). Spread along the second invasion pathway was more rapid, beginning in 2003 in Trinidad and Tobago, where it was probably introduced by the movement of infected material from Venezuela (Fortune et al., 2005). It was then found in Grenada (IPPC, 2007), St Vincent and Grenadines (ISID, 2009b; Richards, 2009), St Lucia (ISID, 2010b), Martinique (Ioos et al., 2011), Dominica (Pérez-Vicente, 2012b, c) and Guadeloupe (Hubert, 2012). Distribution in Africa The earliest report of black leaf streak in Africa was on a plantation at Mununshi in the Luapula Valley, Zambia in 1973 (Raemaekers, 1975). Symptoms resembled black leaf streak, but its identity could not be confirmed from specimens sent to the UK for positive identification (Dabek and Waller, 1990) and there was still no official confirmation of the disease in Zambia (Blomme et al., 2013). The first authenticated report of black leaf streak in Africa was in Gabon in 1978, on ‘Horn’ (AAB, Plantain subgroup) in a commercial plantation at Ntoum, where it was believed that the disease might have been introduced on planting material from Asia (Frossard, 1980; Blomme et al., 2013). Frossard (1980) also ­reported that symptoms of black leaf streak had been observed in São Tomé. In 1980, black leaf streak was

48

Chapter 2

­ bserved in southern Cameroon, close to the boro ders of Equatorial Guinea and Gabon (Tezenas du Montcel, 1982). In 1985, the disease spread from Gabon to the Republic of the Congo (Mourichon, 1986; Wilson and Buddenhagen, 1986). It was also present in Côte d’Ivoire in the same year (Mourichon and Fullerton, 1990). In 1988, black leaf streak was reported from the lowlands of the Democratic Republic of the Congo (Mobambo and Naku, 1993). The presence of black leaf streak was confirmed in Nigeria and Ghana in 1986 (Wilson and Buddenhagen, 1986; ­Wilson, 1987) and Togo in 1988 (Mourichon and ­Fullerton, 1990). In East Africa, the first confirmed report of black leaf streak came from Pemba in 1987 (Dabek and Waller, 1990). From Pemba, the black leaf streak spread rapidly to the nearby island of Zanzibar and then to the Tanzanian mainland, reaching coastal Kenya in 1988 (Kung’U et  al., 1992). A survey the next year showed black leaf streak to be present in the Central and Eastern Provinces of Kenya (Kung’U et al., 1992). During a banana disease evaluation mission in the Great Lakes region of East Africa in 1987, Sebasigari and Stover (1988) observed black leaf streak in the highlands of eastern Democratic Republic of the Congo, Rwanda, where symptoms had been seen in 1986 (Sebasigari, 1990), and Burundi. At that time, the disease was not observed in Uganda. However, in 1990, the disease was recorded in Uganda (Tushemereirwe and Waller, 1993) and in Malawi (Ploetz et al., 1992). Black leaf streak is also present in the Comoros Islands (Jones and Mourichon, 1993). More recently, the disease has been reported in southwest Ethiopia (Gurmu et al., 2017). Economic importance Black leaf streak is a major constraint to banana production (Stover, 1983a, 1986a; Fouré, 1985; Fullerton, 1987; Stover and Simmonds, 1987; Marín et  al., 2003; de Lapeyre de Bellaire et  al., 2010; Churchill 2011; Guzmán et  al., 2013; Ploetz et al., 2015). After the first occurrence of black leaf streak in an area, the disease builds up and often reaches epidemic levels in a few years (Fullerton and Stover, 1990; Belálcazar, 1991). Chemical control costs and crop losses are well documented for export crops (Stover, 1986a, 1990). Losses on smallholders’ plots that produce locally consumed bananas are harder to estimate.

Black leaf streak does not kill plants immediately, but crop losses increase gradually with the age of the planting. The decrease in functional leaf area caused by the disease results in a reduction in the quality and quantity of fruit (Stover, 1983a; Stover and Simmonds, 1987; Pasberg-Gauhl, 1989; Mobambo et  al., 1993, 1996b). Fruit from infected plants ripens prematurely and does not fill properly. Bananas for export are sometimes harvested at a lower grade (younger age) in order to reduce the risks of premature ripening in transit to overseas markets (Stover and Simmonds, 1987). Until the 1970s, the common leaf diseases of cultivars in the Plantain subgroup (AAB) were not considered economically important. This changed when black leaf streak spread to areas where the crop was extensively grown. All over the tropics, plantain is cultivated and fruit consumed by smallholders. In many areas, black leaf streak has caused a considerable decrease in the availability of fruit for local consumption and this has resulted in a substantial increase in their market price. Smallholders growing plantain in the Americas either go out of business, because they cannot cover the high costs of chemical control, or form cooperatives so that their limited resources can be pooled to fight the disease. Black leaf streak is endangering the food security of resource-poor people. Africa alone contributes about 50% of the world plantain production and the demand for plantain is steadily increasing (Wilson, 1987). All traditionally grown plantain cultivars (Fouré, 1985; Mobambo et  al., 1996a) are susceptible to black leaf streak and are severely defoliated by the disease. Plants in the ratoon crop are weaker than in the first cycle and thus more affected by wind damage. On poor sandy soils in West Africa, Mobambo et al. (1996b) estimated that yield losses due to black leaf streak were 33% and 76% during the first and second cropping cycle, respectively. However, in intensively cropped backyard or home garden systems, cultivation is not so seriously affected (Mobambo et al., 1994b). In Nigeria, Mobambo et al. (1993) estimated that black leaf streak caused 39% of yield losses on plantains. Under marginal conditions, plantain production is often abandoned due to low yields. Plantain is not the only smallholder banana to be affected in Africa. The disease also causes serious damage to East African highland cultivars



Fungal Diseases of the Foliage

in the AAA Lujugira–Mutika subgroup. In Uganda, Tushemereirwe (1996) and Tushemereirwe et al. (2000) reported yield losses of 37% in the cultivar ‘Mbwazirume’ because of the effects of a leaf spot complex consisting mainly of black leaf streak and Cladosporium leaf speckle. Although the number of hands and fingers was not affected, fruit from untreated plants were smaller in girth, shorter and underweight in comparison with those of treated plants. Black leaf streak has had a devastating effect on the production of export bananas in the South Pacific. Firman (1972) noted that only 49% of unsprayed Cavendish cultivars produced fruit that reached the export quality standard. Fiji ceased exporting bananas in 1974 and Western Samoa in 1984. Exports also dropped in Tonga and the Cook Islands, because producers had problems maintaining fruit quality standards for their markets in New Zealand. Black leaf streak control had become the single largest production cost (Fullerton, 1987). In 1974, the production of dessert bananas and plantains in Central America was seriously affected by hurricane Fifi, which was also thought to be responsible for the wind-borne spread of black leaf streak to new areas. Before 1974, Honduras exported 500,000 boxes of plantain each year, but afterwards exports dropped to below 1000 boxes (Stover, 1983a). In 1978, the export of plantains from Honduras to the USA was curtailed because of the shortage of fruit with the required quality (Bustamente, 1983). Plantain exports only resumed in 1985 when black leaf streak was controlled by the aerial application of fungicides (Stover, 1987). After the identification of black leaf streak in Costa Rica in 1979 (Jaramillo, 1979), the government initiated a quarantine programme (Woods, 1980). This programme consisted of the destruction of host plants in the affected area and the establishment of roadside quarantine stations strategically located to stop movements of banana and plantain leaves, which were used for padding and shading fruit. About 3000 ha of plantain were destroyed in 1979 and early 1980. The Costa Rican government paid about US$3 million for the eradication programme, but the spread of the disease could not be stopped (Woods, 1980). By 1982, the Ministry of Agriculture in Costa Rica estimated that black leaf streak alone reduced plantain production by

49

40% (Romero, 1986). In general, yields of plantains in well-maintained fields on rich, fertile soils in Central America may have fallen by 20–50% (Stover, 1983a; Pasberg-Gauhl, 1989). Black leaf streak was first detected in Panama in 1980. Bureau (1990) estimated that plantain production in Panama decreased by 69% between 1979 (100,910 t) and 1984 (31,134 t). During this period, the price of plantains rose by up to 50% in local markets. Jaramillo (1987) ­reported that the area planted with plantain ­decreased by 22%, from 7432 ha to 5800 ha ­between 1982 and 1985. About 34% of growers were believed to have abandoned their holdings, leading to a decrease in production of 47%. Colombia is one of the largest plantain producers in Latin America, with 400,000 ha u ­ nder cultivation and an estimated yearly production of 2.5 million tonnes. About 96% of plantains are consumed locally, the remainder being exported (Belálcazar, 1991). About 88% of the plantain is grown in association with coffee by smallholders. Only 12% of the crop is grown in monoculture in larger plantations. After the introduction of black leaf streak, this staple food became scarce and much higher prices were demanded in the market (Belálcazar, 1991). ­ ­Because of the high cost of plantains, consumers changed to other, cheaper food crops. This in turn had a negative effect on plantain production. Black leaf streak has thus had a significant impact on Colombian agriculture and the eating habits of a nation. In neighbouring Venezuela, Zabala and Bermúdez (1999) reported that black leaf streak had caused a 25% decline in plantain production in the main growing area of the country. Control measures substantially increased growers’ costs. Table 2.2 shows the impact of black leaf streak introduction on banana cultivation in various Latin American and Caribbean countries. Usually, there were rises in the costs of crop protection and production, reductions in areas cultivated and losses in yields and fruit quality. This led to more export fruit being rejected and the loss of export certifications as well as changes in consumption habits. Growers also lost heart and turned to other crops. In susceptible banana cultivars, black leaf streak’s impact on the foliage can cause between 20% and 40% reduction of fruit weight and 100% reduction in export quality. In untreated

50

Chapter 2

Table 2.2.  The impact of black leaf streak disease on the production of bananas and plantains in some Latin American and Caribbean countries. Country

Year

Observation of the impact

References

Honduras

1973

Costa Rica

1978–1980

Stover and Dickson (1976); Stover (1986a) González and Jaramillo (1979)

Costa Rica Panama Colombia

1980–1986 1980–1984 1991

Fruit in 10–20 % of bunches seen to be ripening early on a 200 ha plantation Loss of more than 3000 ha of plantain in San Carlos area 40% reduction of plantain production 40% reduction in banana production Plantain production reduction with a large increase in prices for consumers Destruction of 2000 ha of banana in Tabasco: 50–100% fruit losses 50% reduction of land under banana (5000 ha) in Colima Production costs increase by 40–45%. 25% reduction in plantain production and a substantial increase in production costs in the southern part of Maracaibo Lake, the main plantain production area A threefold increase in fungicide costs to control leaf spot in Cavendish plantations Land growing Cavendish reduced from 14,000 to 600 ha with land growing plantain reduced from from 45,000 to less than 12,000 ha Changes of consumption habits Reduction in export crop from more than 85,000 tonnes to little more than 10,000 tonnes Area under banana in Mato Grosso state reduced by 63% 50–100% production losses in Amazonia forcing dietary changes No plantains exported between 2009 and 2012. Exports reduced from 8400 t in 1990 to zero in 2012 by the combined effect of black leaf streak and hurricanes Farms abandoned and an important banana cropping area forced out of production In 1990, 80,000 t of bananas were exported from the island of St Vincent alone. In 2011, only 25,000 t were exported from all five Windward Island countries. Export banana and plantain production declines by 90–100% with trade practically ceasing Reluctance of growers to invest in banana as an export crop. Significant reduction in production area

Mexico

1980 1989–1991

Venezuela

1987–1990 1992–1995

Cuba

1990–1995

Jamaica

1995–2005

Brazil

1999–2004 2000–2002

Guyana

2009–2012

Grenada

2005–2012

Windward Islands

2005–2012

pers. comm. = personal communication

Romero (1986) Bureau (1990) Belalcázar (1991) Orozco-Santos (1998); Orozco-Santos et al. (2001) Orozco-Santos et al. (1996); Orozco-Santos (1998) Martínez et al. (2002) Zabala and Bermúdez (1999)

Pérez-Vicente et al. (2000a, b)

Pérez-Vicente (2012a)

J. Conie, Jamaica, 2006 (pers. comm.) da Silveira et al., (2016) L. Gasparoto, Brazil, 2003, (pers. comm.) Pérez-Vicente et al. (2016). Pérez-Vicente (2012d)

Pérez-Vicente et al. (2016)



Fungal Diseases of the Foliage

plants, the disease significantly affects the filling, length and weight of fruit, but not the number of hands per bunch. The impact of the disease on production and quality is more evident in soils with low fertility or under conditions of nutrition deficiency, as shown in Table 2.3 (M. Guzmán and R. Villalta, Costa Rica, 2015, unpublished results). Similar results were reported by Tushemereirwe et  al. (2000) in the first ratoon crop of an East African highland banana cultivar. In addition, the physiology of fruit from plants with high levels of black leaf streak disease changes so that maturation is accelerated (Plate 2.2). It has been found that fruit from disease-affected plots produces 300% more carbon dioxide and 60% more ethylene at its climacteric peak than fruit from disease-free plots (Castelan et  al., 2012). It has also been shown that black leaf streak increases

51

sensitivity of banana to the postharvest diseases crown rot and anthracnose. This effect of increased sensitivity to postharvest diseases was not observed in fruit from plants affected with Sigatoka leaf spot (Ewané et al., 2013). Cost of control Black leaf streak can be chemically controlled on plantations, but requires twice or more than twice the number of fungicide applications that are necessary to control Sigatoka leaf spot. Strict cultural practices also have to be implemented (Stover, 1990; Marín et al., 2003). In the humid tropics, up to 36 spray cycles/year may be ­required for plantations producing dessert bananas for export and up to 19 cycles/year for commercial plantings of plantain (Stover, 1980b, 1990;

Table 2.3.  Effect of black leaf streak disease on fruit produced by ‘Grande Naine’ (AAA, Cavendish subgroup) growing on soils with low or high fertility in Costa Rica (M. Guzmán and R. Villalta, Costa Rica, 2015, unpublished results). Bunch weight (kg)

Number of hands per bunchb

Treatmenta With chemical control Without chemical controle Difference (%) P > Ff

Fruit diameterc Fruit lengthd second hand Fruit diameterc second hand (mm) last hand (mm) (cm)

Fruit lengthd last hand (cm)

Soil with low fertilityg (average of two crop cycles) 24.1

6.7

34.4

32.8

23.9

20.5

16.9

6.4

31.1

29.3

20.7

18.2

–29.9 < 0.0001

–4.5 0.7665

–9.6 < 0.0001

–10.7 < 0.0001

–13.4 < 0.0001

–11.2 < 0.0001

Soil with high fertilityh (average of four crop cycles) With chemical control Without chemical controle Difference (%) P > Ff

29.2

8.0

36.0

33.3

24.0

19.8

21.9

7.8

33.1

30.0

22.1

18.1

–25.0 < 0.0001

–2.5 0.4454

–8.1 < 0.0001

–9.9 < 0.0001

–7.9 < 0.0001

–8.6 0.0002

Each treatment was applied to 40 plants (four replicates with ten plants/replicate). Hands counted after pruning to remove two true last hands just before bunches were bagged. Fruit diameter measured on central finger in the outer line of fingers. d Fruit length measured on central finger in the outer line of fingers. e Plants without chemical control have no functional leaves at harvest (90±2 days after flower emergence). f T-test (P = 0.05). g Low soil fertility = extractable acidity 3.38 cmol/l; Ca at 1.84 cmol/l; Mg at 0.63 cmol/l; K at 1.22 cmol/l; P at 20 ppm; Fe at 119 ppm; Cu at 4 ppm; Zn at 3.5 ppm; Mn at 34 ppm; organic matter at 6.5%. h High soil fertility = extractable acidity 0.23 cmol/l; Ca at 23.17 cmol/l; Mg at 10.04 cmol/l; K at 0.62 cmol/l; P at 110 ppm; Fe at 315 ppm; Cu at 9 ppm; Zn at 45 ppm; Mn at 44 ppm; organic matter at 1.8%. a b c

52

Chapter 2

Plate 2.2.  Poor control of black leaf streak in commercial Cavendish plantations can induce field-ripening of fruit (upper), the fruit condition known as ‘creamy pulp’ (lower left) or premature ripening in transit (lower right). Such fruit is discarded (photos: M. Sánchez and M. Guzmán, CORBANA).

Fouré, 1983, 1988a, b; Belalcázar, 1991; Gauhl, 1994; Romero and Sutton, 1997a). The cost of black leaf streak control is much higher than the cost of Sigatoka leaf spot control. Stover and Simmonds (1987) reported that 27% of production costs in dessert banana plantations went towards controlling black leaf streak. From 1972 until 1985, the estimated cost of black leaf streak control in Central America, Colombia and Mexico was more than US$350 million (Stover and Simmonds, 1987). In 1993, chemical control measures costs were calculated to be US$400–1400/ha/year (Infomusa, 1993). During the 1980s, the cost of black leaf streak control in export banana crops in Costa Rica was estimated at approximately US$17.5 million/year (Stover and Simmonds, 1987). Between 1985 and 1994, the area under banana cultivation increased from approximately 21,000 ha to

52,737 ha (Serrano and Marín, 1998). As consequence, black leaf streak control costs in 1995 were estimated to have increased to US$49 million/year (Romero and Sutton, 1997a). This cost continued to increase because of a build-up of resistance to fungicides and favourable environmental conditions for disease development. In 2015, it was estimated at US$79 million for the 43,000 ha under cultivation (M. Guzmán, Costa Rica, 2016, personal communication). In Cuba, leaf spot protection costs for plantations of Cavendish cultivars increased to US$2336/ha/year after black leaf streak became established in the early 1990s. Total costs of protection rose to US$2 million/year (Pérez-­Vicente et al., 2002a). This led to a reduction in the area planted to susceptible Cavendish cultivars and their replacement by resistant cultivars, especially hybrids bred for resistance to black leaf



Fungal Diseases of the Foliage

streak at the Fundación Hondureña de Investigación Agrícola (FHIA). After black leaf streak was introduced to Jamaica, the average number of spray treatments with fungicide rose to 22/ year (J. Conie, Banana Board, Jamaica, 2006, personal communication) and control costs varied between US$1076/ha/year and US$1555/ ha/year, which amounted to 8.1–18.0% of total production costs.

Symptoms D.R. Jones, C. Pasberg-Gauhl, F. Gauhl, E. Fouré, M. Guzmán and L. Pérez-Vicente In Hawaii, Meredith and Lawrence (1969) observed the development of symptoms of black leaf streak on mature, pre-flowering plants of susceptible ‘Dwarf Cavendish’ (AAA, Cavendish subgroup). They described first symptoms, which were faint reddish-brown specks less than 0.25 mm in diameter, as being visible on the lower surface of the leaf at an ‘initial speck stage’. These specks then elongated, becoming slightly wider, to form a narrow, reddish-brown streak with dimensions of 20 mm × 2 mm, the long axis of the streak being parallel to leaf veins. At this ‘first streak stage’, streaks were more clearly visible on the lower leaf surface than on the upper. The streaks could be densely aggregated in places and frequently overlapped to form larger, compound streaks. At the ‘second streak stage’, the colour of streaks, which were clearly visible on the upper leaf surface, changed to a very dark brown, almost black, colour. The entire leaf could blacken at this stage if streaks were numerous, but, if less densely congregated, streaks broadened and water-soaked borders, which were seen best after rain or dew, appeared. At this ‘first spot stage’, lesions became fusiform or elliptical in outline. At the ‘second spot stage’, the dark brown or black centres of spots became slightly depressed and leaf tissue immediately surrounding the more pronounced water-soaked border yellowed slightly. At the ‘third or mature spot stage’, the centre of the spot dried, becoming light grey or buff in colour. Each mature spot had a well-defined dark brown or black border and surrounding tissue was often bright yellow. Where spots coalesced, whole sections of leaves

53

became necrotic. After the leaf withered, spots remained visible because of their light-coloured centres and dark borders (Meredith and Lawrence, 1969). Fouré (1987), who worked with black leaf streak in West Africa, identified six main stages of symptom development (Plate 2.3). His ‘stage 1’, which precedes the initial speck stage of Meredith and Lawrence (1969), is characterized by small whitish or yellow specks that appear on the underside of the leaf. In the second phase of this stage, specks turn rusty brown in colour. These specks are less than 1 mm long and are not visible in translucent light. The specks grow into characteristic narrow, reddish-brown or dark brown streaks, which Fouré called ‘stage 2’. Streaks, which are generally 2–5 mm long, are visible in translucent light and are easy to recognize from a distance of 1–2 m. On some clones, brown streaks are visible on the underside of the leaf with corresponding yellow streaks on the upper surface (Plate 2.3). The colour of the upper surface streak changes through brown to black, but the brown colour is retained on the underside. Streaks become longer in ‘stage 3’ and can reach 20–30 mm in length. If conditions are favourable and inoculum potential is high, streaks in stages 2 or 3 coalesce and cause leaf necrosis. In Fouré’s ‘stage 4’, individual streaks that have not coalesced broaden to form a spot, have a brown colour on underside of leaf and are often black on the upper leaf surface. At ‘stage 5’, spots turn elliptical and black on both sides of the leaves, usually surrounded by a yellow halo, and the centre begins to flatten out. During ‘stage 6’, the centre of the spot dries and fades to clear grey, and is usually surrounded by a well-defined black border which in turn is surrounded by a water-soaked or yellow halo. The grey spot with dark border remains visible after the leaf has died and dried (Plate 2.3). It can be difficult to distinguish symptoms of black leaf streak from those of other leaf diseases, especially Sigatoka leaf spot and eumusae leaf spot. Experience with banana diseases is ­required to make a correct diagnosis on visual symptoms and even then a microscopic examination and/or molecular tests are needed for confirmation. Plate 2.4 shows the typical symptoms of Sigatoka leaf spot and black leaf streak and the early characteristic symptoms of both in a mixed infection. The symptoms that occur when Cordana leaf spot begins to develop in

54

Chapter 2

Plate 2.3.  Stages of black leaf streak symptom development according to Fouré (1987). Stage 1: characterized by small whitish, yellow or rusty brown specks of less than 1 mm in length that appear on the underside of the leaf (upper left). Stage 2: specks grow into narrow reddish-brown or dark brown streaks, which are generally 2–5 mm long (upper centre). Stage 3: streaks become wider and longer reaching 20–30 mm in length accompanied by a colour change from brown to dark brown or black (upper right). Stage 4: streaks broaden to form spots that are dark brown on lower leaf surfaces and black on upper leaf surfaces (lower left). Stage 5: development of an elliptical spot, which is black on both sides of the leaf, and usually surrounded by a yellow halo with the lesion being slightly depressed (lower centre). Stage 6: the depressed centre of the spot, which is surrounded by a well-defined black border, dries and turns whitish to clear grey. The scale is in millimetres (photos: M. Sánchez and M. Guzmán, CORBANA).

f­ungicide-arrested lesions of black leaf streak is also shown to demonstrate how closely lesions resemble those of eumusae leaf spot. The pattern of first symptoms is determined by the stage at which the unfurling leaf is infected. As the unfurling leaf is a constantly expanding funnel, new susceptible tissue is being gradually exposed to the inoculum. If wind-dispersed spores are deposited on the lower surface of the apical third of the largely furled heart leaf, a distinct line of specks, which later develop into streaks and spots, appears along the left edge of the leaf, particularly towards the tip (as viewed looking down on the upper leaf surface from the base towards the apex). When conditions are extremely favourable for infection, these early speck symptoms may appear on leaf 2 (counting down the plant from the first fully opened leaf). However, they are usually first seen on leaves 3 and 4. Symptoms that develop due to later infections

are generally observed at the distal end of the right lamina margin, followed by the middle section and successively towards the base of the leaf (Stover, 1980a; Gauhl, 1989). On a growing susceptible plant, streaks are usually present on the third, fourth and fifth youngest fully opened leaves and both streaks and spots on the fifth and older leaves. If the plant is stressed and slow growing, advanced symptoms may be seen on the second or even the first fully opened leaf. On juvenile leaf tissue, such as on water-suckers and young plants derived from tissue culture, initial specks quickly develop into oval or circular brown spots without showing the streak stage (Plate 2.4). It usually takes 3–4 weeks after symptoms first appear for a leaf to die (Stover, 1972; Gauhl, 1994). However, when young streaks at ‘stage 2’ coalesce, large brown areas are formed, which quickly turn black and the whole leaf may die



Fungal Diseases of the Foliage

55

Plate 2.4.  Symptoms of Sigatoka leaf spot on a leaf of a Cavendish cultivar in Martinique (upper left); black leaf streak on a leaf of a Cavendish cultivar in Costa Rica (upper right); Cordana leaf spot on a leaf of Musa balbisiana in Costa Rica (middle left); and black leaf streak disease on a leaf of a young plant of a Cavendish cultivar in Costa Rica (middle right). Early symptoms of a mixed infection of black leaf streak (blue arrows) and Sigatoka leaf spot (white arrows) on a Cavendish cultivar in the EMBRAPA germplasm collection at Cruz das Almas, Brazil in September 2016 (lower left). Cordana leaf spot developing in lesions of black leaf streak following treatment with a systemic fungicide (lower right) (photos: M. Guzmán, CORBANA).

within 1 week. Sometimes, because of the intensity of infection, fewer than six living leaves may be present on growing plants in the vegetative stage (see Plate 2.1). On more resistant cultivars, symptoms are only seen on the older, lower leaves, due to slower symptom development. Black leaf streak looks more severe on plants with bunches, because leaf production ceases and symptoms appear on leaves higher up the plant. If disease pressure is great, it is not uncommon for a very susceptible cultivar to have no viable leaves at harvest (see Plate 2.1). The

number of functional leaves at harvest has been used to gauge the resistance of cultivars to black leaf streak. Streaks and spots have been noted as being different in respect of colour, size and shape on different cultivars (Meredith and Lawrence, 1970a). Many accessions of wild Musa acuminata subspecies and some cultivars in the AA and AAA groups are highly resistant to the pathogen. These usually respond to infection by producing small yellow, brown or chlorotic dots, streaks or spots (Plates 2.5), which do not develop further.

56

Chapter 2

Different patterns of streaks and lesions are observed on leaves depending on the mode of infection. When infection is caused by conidia distributed by water drops running across the

unfurled leaves, distinct lines of lesions are produced that run diagonally from the tip of the leaf to the midrib (Plate 2.6). Ascospores carried to the leaf by air currents result in a random distribution of lesions on the exposed unfurling leaf tissue (Plate 2.6). During times of prolonged rainfall, such as in wet seasons, conspicuous necrotic spots develop along the leaf midribs of plants treated with systemic fungicides and oil. This is caused by the movement of the active ingredients of the fungicide to leaf margins leaving the leaf tissue near the midrib unprotected. Because of their position close to vascular tissue, coalescing necrotic lesions along the midrib usually result in the leaf lamina dying.

Causal agent Plate 2.5.  Very small necrotic lesions formed on the leaves of Musa acuminata ssp. burmannica (AAw, accession ‘Calcutta 4’) after inoculation with Pseudocercopsora fijiensis. This expression of high resistance (HR) is characterized by an early plant reaction at the infection site and the blockage of further symptom development at the early streak stage (photo: M. Sánchez and M. Guzmán, CORBANA).

J. Carlier and C. Abadie Taxonomy and phylogeny The fungus that causes black leaf streak disease was first described by Leach (1964a) and given the name Mycosphaerella fijiensis (sexual morph). It is a hemibiotroph and heterothallic ascomycete

Plate 2.6.  Characteristic infection patterns associated with Pseudocercospora fijiensis conidia (left) and ascospores (right) under natural conditions (photo: M. Sánchez and M. Guzmán, CORBANA).



Fungal Diseases of the Foliage

with a bipolar mating system. The asexual morph of M. fijiensis is Pseudocercospora fijiensis (Crous et al., 2003). When black leaf streak was first found in Latin America, it was called black Sigatoka and the name M. fijiensis var. difformis was assigned to the pathogen (Stover, 1974). This name was later validated on the basis of an examination of specimens from La Lima, Honduras (Mulder and Stover, 1976). The taxonomic characteristic used to separate M. fijiensis var. difformis from M. fijiensis was the presence of sporadic stroma, which gave rise to dense and loose fascicles of conidiophores. This feature had not been described for M. fijiensis. However, a later, comprehensive taxonomic study of isolates of M. fijiensis and M. fijiensis var. difformis revealed that the two were synonymous (Pons, 1987, 1990). This synonymy was later confirmed by molecular studies (Carlier et al., 1994; Crous et al., 2003). The Mycosphaerellacaea family, to which Mycosphaerella belongs, has been under taxonomic revision based on the results of genetic sequencing studies (Arango-Isaza et  al., 2016). The name of the pathogen under the new onename-for-one-species taxonomic system is Pseudocercospora fijiensis (asexual morph) with Mycosphaerella fijiensis being a synonym. Conidiophores of P. fijiensis first develop in initial brown flecks or early streaks on the lower surface of the leaf and continue to be produced until the second spot stage of Meredith and Lawrence (1969) or stage 4 of Fouré (1982). They emerge singly or in small groups from stomata within the boundary of the lesion. Most are produced on the lower surface, though a few arise on the upper. Conidiophores are pale to medium olive-brown, becoming slightly paler towards the tip. They are straight or bent, often with geniculations and sometimes with a basal swelling up to 8 μm in diameter, 0–5-septate, 16.5–62.5 × 4–7 μm, usually slightly narrower, but occasionally wider, at the tip. One or more scars are present near the tip of the conidiophore, either flat against the apex or on the side, or on a slightly sloping shoulder (Fig. 2.1). Conidia are formed singly at the apex of the conidiophore, later becoming lateral as the conidiophore develops. Fouré et  al. (1984) estimated that three to four conidia are produced on each conidiophore with up to four mature conidia attached at any one time. Conidia are pale green or olivaceous, obclavate to cylindro-obclavate,

57

C

10 μm

B

A

Fig. 2.1.  Stroma (A), conidiogenous cells (B) and conidia (C) of Pseudocercospora fijiensis (from Pons, 1987).

1–10-septate (commonly 5–7-septate), straight or curved, obtuse at the apex, truncate or rounded at the base, with a visible and slightly thickened hilum, 30–132 × 2.5–5 μm, the broadest point being at the base (Fig. 2.1; Plate 2.7). Spermogonia develop at the stage when streaks turn into spots and are more abundant on the lower surface of the leaf. They are consistently associated with conidiophores. Spermogonia are hourglass-shaped, oval or almost globose and measure 55–88 × 35–50 μm (Plate 2.7). The ostiole is slightly prominent and protrudes through the stoma pore. Mature spermatia are hyaline, rod-shaped and measure 2.5–5.0 × 1.0–2.5 μm. The sexual fruiting body has been referred to as perithecium (Meredith and Lawrence, 1969), but the ascocarp of P. fijiensis is known as a pseudothecium because asci are not regularly organized into a hymenium and are bitunicate, having a double wall which expands when it

58

Chapter 2

Plate 2.7.  Pseudocercospora fijiensis. Pseudothecia and spermogonia on diseased leaf tissue of ‘Grande Naine’ (AAA, Cavendish subgroup) (upper left); ascospores (upper centre); and ascospores germinating on water-agar (upper right). Colonies, each derived from one ascospore, after 21 days on potato dextrose agar (lower left). Conidium produced on an in vitro culture (lower centre). Colonies, each derived from one conidium, after 18 days on potato dextrose agar (lower right) (photos: F. Alfaro and M. Guzmán, CORBANA).

takes up water and shoots the enclosed spores out suddenly (Crous 2009; Crous et  al., 2009; Churchill, 2011). The shapes of the pseudothecia vary, but are mostly globose with a diameter of 47–85 μm. They are immersed in the leaf tissue (Plate 2.7) with protruding ostioles and are found on both leaf surfaces, though more abundant on the upper. The wall of the pseudothecium is dark brown with three or more layers of polygonal cells. The numerous asci are bitunicate, obclavate and without paraphyses. The ­hyaline, biseriate ascospores have dimensions of 12.5–16.5 × 2.5–3.8 μm and are two-celled, with the larger cell uppermost in the ascus. The ascospore is slightly constricted at the septum (Mulder and Holliday, 1974a). The ascostroma and asci are illustrated in Fig. 2.2. Ascospores are illustrated in Fig. 2.2 and Plate 2.7.

B 10 µm

A

Cultural characteristics Growth on culture media is slow. A single conidium seeded on potato dextrose agar forms a colony about 1 cm in diameter after 38 days of incubation at 26°C (Meredith, 1970). The optimal temperature for growth ranges from 24°C to ­

Fig. 2.2.  Acostroma with asci (A) and ascospores (B) of Pseudocercospora fijiensis (from Pons, 1987).



Fungal Diseases of the Foliage

28°C (Meredith, 1970; Stover, 1983b; MouliomPefoura and Mourichon, 1990). Cultures are raised and stromatic and have a velvety surface coloured pale grey, pink–dark grey or grey-brown (Stover, 1976). Mycelial characteristics in any one segment are not constant and may change as the colony grows. Colonies derived from conidia and ascospores are shown in Plate 2.7. Ten- to 21-day-old colonies can produce conidia (Mourichon et  al., 1987; Jacome et  al., 1991). Sporulation has been reported to be optimal on cultures grown on V8-juice agar adjusted to pH 6 and incubated at 20°C under conditions of continuous light (Abadie et al., 2008). Some adaptations based on brush harvesting mycelium and spectral lights were proposed to improve conidial production in vitro (Peraza-Echeverria et al., 2008; Sepúlveda-­Ramos et al., 2009). The optimum temperature for the germination of conidia and ascospores in culture is around 26°C (Jacome et  al., 1991). Maximum germination occurs in water and decreases as the relative humidity (RH) lowers. No conidia have been observed germinating below 95% RH and no ascospores below 98% RH under controlled conditions (­Jacome et al., 1991). Diagnostic methods Diseases caused by Pseudocercospora fijiensis, P. musae, P. eumusae and Phaeosphaeriopsis musae are difficult to distinguish on the basis of symptom expression. However, these pathogens can be separated from one another on microscopic differences between asexual stages of the fungi on leaf samples and in culture (Meredith and Lawrence, 1969, 1970a; Mourichon and Fullerton, 1990; Carlier et  al., 1999c; Crous et  al., 2003; Zapater et al., 2008a). Molecular methods have also been developed to help distinguish P. fijiensis and P. musae based on the ITS regions of the rDNA (Johanson and Jeger, 1993; Henderson et al., 2003, 2006). More recently, Arzanlou et al. (2007a) developed TaqMan real-time quantitative PCR assays based on the beta-tubulin gene that proved to be a specific and reliable molecular diagnostic tool that differentiates P. fijiensis, P. musae and P. eumusae. Reproduction and mating types Pseudocercospora fijiensis can reproduce both asexually and sexually on plants or in culture. However, because P. fijiensis is heterothallic,

59

pseudothecia, asci and ascospores can be produced in culture only by crossing isolates having complementary mating types (Mourichon and Zapater, 1990). The idiomorphs mat1-1 and mat1-2 were sequenced by Conde-Ferráez et al. (2007). This information will be useful in determining the mating type of isolates, which can then be crossed in vitro to study the genetic inheritance of different traits. The sequences of the mating type loci of P. fijiensis were compared with those of P. musae, P. eumusae and other ­fungi with Mycosphaerella as the sexual morph (Arzanlou et  al., 2010). The analysis showed that the three major pathogens of the Sigatoka leaf spot complex share a common ancestor. Genetic transformation The capability to transform and to create mutants of pathogens is today essential for understanding infection cycles and identifying pathogenic genes. The first transformation system for P. fijiensis, but also for P. musae and P. eumusae, was developed by Balint-Kurty et al. (2001) based on traditional polyethylene glycol (PEG)/protoplast-mediated methods. Later, Portal et al. (2012) ­developed a system using a restriction enzyme-­mediated integration (REMI) methodology. More recently, a transformation protocol that is simpler, much more efficient, faster and reproducible by underwater shock waves has been described (Escobar-­ Tovar et al., 2015b). Together with the available genomic sequences of both P. fijiensis and Musa acuminata, these transformation systems offer new opportunities to investigate and determine the genetic basis of pathogenicity. Genetic structure and history of populations introduction. 

Investigations on the genetic structure of fungal populations can throw light on their distribution history and means of local/ global spread. This information is crucial in the identification of durable strategies for disease management and breeding for disease resistance. Such investigations with P. fijiensis have been conducted in the past two decades using different genotyping methods.

genotyping methods. 

Restriction fragment length polymorphism (RFLP) markers based on single-­ copy DNAs were the first molecular markers ­developed for P. fijiensis (Carlier et  al., 1994, 1996). Later, a DNA fingerprinting method

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based on synthetic simple repetitive oligonucleotides was published (Muller et al., 1997). These methods were based on the use of radioactively labelled probes and were rapidly replaced by PCR-based methods that were easier to use. Neu et  al. (1999) characterized the first microsatellites markers in P. fijiensis. Zapater et al. (2004) developed PCR-RFLP markers based on single-­ copy DNAs that relied on a simple and robust protocol transferable to locations lacking sophisticated equipment. About 32 new microsatellites markers isolated from DNA libraries enriched for microsatellites motives were published (Zapater et al., 2008a). About 15 of these markers can be used in multiplex PCR analysis. Following the release of the P. fijiensis genome ­sequence, new microsatellites markers based either on di- and trinucleotide motives (Yang and Zhong, 2008) or on tetranucleotide motives (Robert et  al., 2010) were developed. Tetranucleotide microsatellite markers are less variable and thus more adapted to compare populations from distant geographical origins in multiplex panels (Robert et al., 2010, 2012). About eight sequence-based markers were developed in variable nuclear introns from the genome sequence (Robert et  al., 2012). The genome sequence was also screened for more variable number tandem repeat (VNTR) markers (Garcia et  al., 2010). PCR markers based on the distribution of repetitive elements across the P. fijiensis genome have also been recently described (Queiroz et al., 2013; Silva et al., 2014). random mating type and recombination effects in p.  fijiensis populations. 

Evidence for random mating has been found in most P. fijiensis populations analysed to date (Carlier et  al., 1996; Rivas et  al., 2004; Robert et  al., 2012). High levels of recombination in P. fijiensis lead to linkage equilibrium between loci and a high ­level of genotypic diversity. Conde-Ferraez et al. (2010) developed a multiplex-PCR method from idiomorph sequences to estimate the frequency of the mating types in P. fijiensis populations. In most of the Mexican populations tested, the results did not show a significant bias from the 1:1 ratio as expected in random mating populations. The same result was obtained when Brazilian samples were analysed (Queiroz et al., 2013). Intragenic recombination events have also been detected in some genes coding for effector

­ roteins that ­regulate biological activity (Stergip opoulos et  al., 2014). Thus genetic recombination could play a major role in the pathogen’s known ability to develop resistance to fungicides and overcome banana resistance genes. It also indicates that the fungus has a high evolutionary potential (McDonald and Linde, 2002). centre of diversity of p.  fijiensis and initial emergence of the disease. 

The structure of P. fijiensis populations worldwide suggests that a region encompassing Malaysia, Indonesia, Papua New Guinea and the Philippines could be the centre of P. fijiensis diversity (Carlier et al., 1996; Hayden and Carlier, 2003; Rivas et  al., 2004; Robert et al., 2012). It is here that a high genetic divergence of sequence markers and a high private allelic richness of molecular markers have been detected (Table 2.4). This region is also the centre of origin of the wild Musa acuminata subspecies that contribute to the genetic make-up of cultivated banana and where many banana cultivars were first selected and cultivated (Perrier et al., 2011). A coevolution with P. fijiensis could have occurred during the domestication process (Stukenbrock and McDonald, 2008). Stover (1978) suggested a narrower centre of origin of P. fijiensis corresponding to New Guinea and the Solomon Islands. This area is also considered as an important primary centre of banana domestication. However, the small number of isolates from Southeast Asia and the Pacific islands so far genotyped are not sufficient to definitively confirm this hypothesis. Populations of P. fijiensis from wild Musa species and banana cultivars need to be compared to further define the centre of origin and investigate the coevolution hypothesis. continental introduction events. 

Populations of P. fijiensis in Latin America and Africa have been found to be less genetically diverse than those in Southeast Asia (Table 2.4; Rivas et al., 2004; Robert et  al., 2012). This indicates that introduction events outside Southeast Asia were all accompanied by founder effects (a reduction in genetic diversity that results when a population is descended from a small number of individuals). It has been suggested that there may have been two independent introductions of black leaf streak into Africa (Pasberg-Gauhl et  al.,



Table 2.4.  Genetic diversity indices estimated in Pseudocercospora fijiensis for both microsatellites and sequences datasets, at the levels of populations and continents (abbreviations as in Fig. 2.3) (after Robert et al., 2012). Microsatellite markers variation Total dataset (735 ind.)

Populations dataset (629 ind.)

HE

Ar

pAr

S-E Asia (145)

0.65

8.3

4.6

Oceania (94)

0.33

2.6

0.1

America (279)

0.35

2.6

0.17

Africa (217)

0.22

2.4

0.25

Population (sample size)

% of polymorphic loci

HE

Ar

Private Ar

Ind (14) MaL1 (13) MAL2 (22) PHL (22) Png1 (24) Png2 (25) FIJ (25) Ncl (22) AUS (31) COL (23) CRI (33) H (32) H-G (25) MEX (10) PAN (32) VEN (25) Jam (25) RD (31) COM (34) UGA (34) CAM5 (21) CAM6 (24) CIV (34) Gab (24) NGA (24)

90 (19/21) 80 (17/21) 71 (15/21) 95 (20/21) 83 (15/18) 100 (21/21) 62 (13/21) 57 (12/21) 81 (17/21) 65 (13/20) 76 (16/21) 71 (15/21) 70 (14/20) 71 (15/21) 71 (15/21) 55 (11/20) 62 (13/21) 48 (10/21) 29 (6/21) 81 (17/21) 52 (11/21) 57 (12/21) 52 (11/21) 57 (12/21) 38 (8/21)

0.62 0.47 0.36 0.63 0.42 0.56 0.19 0.20 0.36 0.31 0.33 0.37 0.44 0.43 0.30 0.31 0.26 0.19 0.09 0.31 0.22 0.18 0.22 0.23 0.12

3.9 3.0 2.3 4.6 2.6 3.9 1.7 1.9 2.6 2.2 2.1 2.2 2.3 2.2 2.1 2.1 2.1 1.8 1.3 2.3 1.8 1.7 2.0 1.7 1.6

0.6 0.4 0.21 0.68 0.21 0.16 0.05 0 0.06 0.03 0.03 0 0.01 0 0 0 0.01 0.02 0 0.06 0.07 0.01 0 0 0

Geographical area (sample sizea)

No. of informative sites/total

S-E Asia (23)

70/4198

0.0069

Oceania (22)

25/4198

0.0028

0.96

11.7

America (28)

18/4198

0.0016

0.87

6.7

Africa (29)

10/4198

0.0009

0.87

3.8

π

Hd

k 28.6

61

HE, unbiased estimate of gene diversity (Nei, 1978); Ar, allelic richness corrected for sample size; pAr, private allelic richness corrected for sample size (standardization to a sample size of 15 individuals for populations and 180 individuals for continents); π, nucleotide diversity (Nei, 1987); Hd, haplotype diversity (Nei, 1987); k, average number of nucleotide differences (Tajima, 1983). a Excluding sequences with missing data not usable for calculations.

Fungal Diseases of the Foliage

Geographical area (sample size)

Sequence-based markers variation

62

Chapter 2

1999; Blomme et  al., 2013). However, the results obtained by Robert et  al. (2012) strongly indicate a single introduction of P. fijiensis from one Southeast Asian source population. Indeed, all the African isolates form an isolated and relatively homogenous genetic cluster. A more recent statistical analysis supports this theory (J. Carlier, France, 2016, personal communication) and has allowed effective population sizes to be estimated. The original P. fijiensis population in Africa was estimated to be approximately a few tens of thousands of individuals. A very strong bottleneck reduced such an effective population to a few tens of individuals or less over a period of about ten generations. In addition, the study suggested a subsequent colonization of ­Africa through random founder events across the continent as first described by Rivas et  al. (2004). Such a colonization pattern could have arisen from the long-distance dispersal of windblown ascospores or by the movement of infected material. By contrast, the results obtained by Robert et al. (2012) working with isolates of P. fijiensis from the Americas suggest that populations there are the result of interbreeding between two or more previously isolated populations from different independent locations in Southeast Asia and the Pacific. This event was shown recently to be the most probable using an Approximate Bayesian Computation (ABC) method of analysis with the size of the ancestral population estimated at a few tens of thousands of individuals (Robert et al., 2015). This was in the same order of magnitude as the one estimated in the African study. However, the intensity of the bottleneck was weaker in comparison with Africa with an order of 100 individuals for about 50 generations. An admixture from genetically diverse sources is consistent with the historical reports of the first detection of P. fijiensis in the American continent. The pathogen was identified for the first time in 1972 in a banana germplasm collection in Honduras. However, typical black leaf streak symptoms were subsequently recognized on colour transparencies taken in the collection in 1969. The disease may have been introduced even earlier and remained undetected (Stover and Dickson, 1976). The germplasm collection consisted of nearly 800 Musa species and cultivated clones obtained from South Asia, Southeast Asia and the Pacific (Papua New

Guinea, Solomon Islands and Hawaii) between 1959 and 1963 (Rosales et al., 1999). The finding that the highest level of genetic diversity of P.  fijiensis in the Americas occurs in Honduras supports the theory that the collection was the site of introduction of the pathogen into the Americas. The most recent populations of P. fijiensis to be analysed came from Brazil, where the disease was first described in 1998. This work revealed the existence of different genetic groups (Queiroz et al., 2013; Silva et al., 2014, 2016). These groups will need to be compared with others from various geographical locations before it can be determined if they arose from a bottleneck event during the spread of P. fijiensis in the Americas or were introduced separately. spread in the caribbean region.  A comprehensive population study including samples from Latin American countries and Caribbean islands has been conducted with microsatellites markers (J. Carlier, France, 2016, personal communication). This study shows that P. fijiensis spread through two different epidemic waves within the Central American–Caribbean basin. Starting from Honduras, a northern wave followed a route from Mexico to Puerto Rico and a southern wave a route from Venezuela to St Lucia. The relatively homogenous genetic background of isolates in each wave indicates a natural step-bystep dispersal process between islands. However, this study also highlighted the occurrence of more random and unpredictable events, especially in the French Indies. For instance, populations in Martinique have been found to have originated from both the northern and southern pathways. The population typical of the southern pathway most likely gained entry by natural spread from St Lucia, but the movement of infected material into Martinique is strongly suspected as the source of the population typical of the northern pathway. As a consequence of the introductions, an admixture between introduced populations has created a new population with new genetic characteristics that has spread north from Martinique to Dominique and Guadeloupe. To further understand this process and localize geographical zones where admixture ­between populations has occurred, a landscape genetic study from about 300 isolates covering the whole of Martinique and Guadeloupe is in



Fungal Diseases of the Foliage

progress (V. Roussel., M.-F. Zapater, D. Bieysse, Y. Chilin, C. Abadie and J. Carlier, France, 2016, personal communication). local genetic variation. 

A study conducted in Nigeria showed that a moderate level of genetic variability exists among local P. fijiensis populations (Zandjanakou-Tachin et al., 2009). Halkett et al. (2010) analysed samples of the pathogen collected in 10–15 sites distributed along 250– 300 km transects in Cameroon and Costa Rica using microsatellite markers. Low to moderate genetic variability was detected in both countries, with genetic isolation by distance identified in Cameroon. In contrast with genetic variability patterns observed at the continent scale, the genetic differences observed between local banana-­production areas in Cameroon in combination with historical data suggest a continuous range expansion through a gradual ­dispersal of spores. Also in Cameroon, in an area measuring 50 × 50 km next to the Nigerian ­border, two populations clearly delineated by a sharp and asymmetric genetic discontinuity were detected (Rieux et  al., 2011). This finding probably reflects the recent history of the spread of P. fijiensis in Cameroon with the area in question being the point of contact between two e­xpanding populations. A new method using rates of changes in cline shapes for neutral markers was developed which allowed a parameter related to the dispersal in P. fijiensis to be estimated indirectly (Rieux et al., 2013b). The results of this study support the estimate of spore dispersal obtained from direct epidemiological studies (Rieux et al., 2014; see section ‘Disease cycle and epidemiology’ ­below). No significant genetic differentiation has been detected in other studies conducted in Cameroon on a local scale (Rivas et  al., 2004; Fahleson et al., 2009). When Rieux et al. (2013a) undertook an analysis of isolates collected over a 50 × 80 km area in Central Cameroon, no variability patterns were detected, suggesting an expansion of populations and/or no influence from potential barriers. In addition, no differences were detected between samples taken from commercial and food-crop plantations, suggesting that a high population size and/or a high migration rate might be responsible for the failure of disease control practices to affect the

63

­ enetic make-up of populations. Finally the reg sults that have been published to date demonstrate that the genetic diversity of P. fijiensis is distributed at a low scale. For example, about 90% of the diversity detected in Cameroon is distributed at the scale of a plantation and, within a plantation, all the diversity is found at the scale of a plant (Rivas et  al., 2004). More recently Ngando et  al. (2015) revealed that 30% of lesions from which about ten conidia were cloned had a mixture of two different microsatellite haplotypes. Pathogenic variability and host selection Isolates of P. fijiensis from different locations in Papua New Guinea and elsewhere have been found to vary in their pathogenicity (Fullerton and Olsen, 1995). This variability was detected when young plants of a standard set of Musa genotypes, which had been propagated in tissue culture, were inoculated with suspensions of conidia harvested from isolates growing in culture. The reactions of the genotypes to P. fijiensis isolates were differential, suggesting specific interactions. Some genotypes considered as highly resistant or hypersensitive in the field, such as the wild banana M. acuminata ssp. burmannica (AAw, accession ‘Calcutta 4’) and ‘Yangambi Km 5’ (AAA, Ibota subgroup), were attacked by several isolates. Jacome and Schuh (1993) reported that six isolates of P. fijiensis from Honduras induced ­different levels of disease severity, which was ­defined as the percentage of a leaf target area covered by black leaf streak lesions, on ‘Grande Naine’ (AAA, Cavendish subgroup). Romero and Sutton (1997b), working with isolates from different geographical regions, showed that they varied in disease severity, with the time between inoculation and the appearance of the first ten streak symptoms also differing. These results indicated that isolates of P. fijiensis may vary in their aggressiveness. More recently, P. fijiensis isolates from various locations in Nigeria were found to have different levels of aggressiveness (Zandjanakou-Tachin et al., 2013). Fullerton and Olsen (1995) reported the sudden breakdown of the resistance of the East African cultivar ‘Paka’ (AA) and the Jamaican-­ bred tetraploid ‘T8’ (AAAA) on the island of Rarotonga in the Cook Islands after approximately

64

Chapter 2

8 years of exposure to black leaf streak. They suggested that the change from high resistance to complete susceptibility might have been because of the failure of a single gene conditioning resistance. In Cameroon, Mouliom-Pefoura (1999) described the appearance of necrotic lesions with pseudothecia on leaves of ‘Yangambi Km 5’ (AAA, Ibota subgroup), a cultivar that has been reported as highly resistant to black leaf streak. A loss of high resistance was also observed in Costa Rica (Escobar-Tovar et  al., 2015a) and Puerto Rico (Irish et  al., 2013). The susceptibility of once-­ resistant ‘Yangambi Km 5’ may be the result of the breakdown of a single gene that controls the hypersensitive-type response. Field observations and inoculations of P.  fijiensis isolates under controlled conditions demonstrated an erosion of the partial resistance of ‘FHIA-18’ (AAAB, bred hybrid) after 5 years of cultivation on a large scale in Cuba (Pérez-­ Miranda et al., 2006; Pérez et al., 2006). Erosion of the resistance of the hybrid ‘FHIA-21’ (AAAB, bred plantain-type hybrid) is also suspected in the Dominican Republic and Costa Rica. In Cuba and the Dominican Republic, some modifications of the plant nutritional status and/or some epidemiological factors linked to agronomical practices are thought to be associated with the breakdown or erosion of resistance in FHIA hybrids (A. Cavalier, L. Pérez-­Vicente, D.  Rengifo, L. Miniere, T. Lescot and C. Abadie, France, 2016, unpublished results). Experiments to evaluate the effect of fertilization programmes on the resistance of these hybrids are in progress. An adaptation of P. fijiensis populations could be also implicated in the observed erosion of resistance in FHIA hybrids. A greater knowledge of genes for resistance in banana and pathogenicity in the pathogen is needed in order to develop efficient and durable strategies of resistance management. The sequencing of the P. fijiensis genome has enabled genetic work to begin on identifying the genes involved in the change in pathogenicity of pathogen populations from Cuba and the Dominican Republic (J. Carlier, France, 2017, personal communication).

2008). From the same cross, progenies were analysed with about 320 SSR and DArT markers to generate a denser genetic map (Ferreira et  al., 2009). This map was used to help assemble the recently published P. fijiensis genome (ArangoIsaza et al., 2016) and for which version 2 was released in May 2010 on the Joint Genome Institute’s website (JGI, 2010). The genome size of P. fijiensis was estimated to be 74 Mb, making it the largest within any species in the Dothideomycetes that have been sequenced to date (Ohm et  al., 2012; Arango-Isaza et  al., 2016). This large size results from the massive expansion of long terminal repeat retrotransposons representing 50% of the genome. The estimated number of gene models is about 11,000, 1.3-fold higher than in Zymoseptoria tritici. About 170 of these genes encode small secreted proteins (SSPs), which are potential effectors. Synteny with the genome of Z. tritici suggests a core set of 12 chromosomes, 11 corresponding to the largest of the 56 main genome scaffolds. The smaller scaffolds are suspected to be dispensable chromosomes as observed in many other Dothideomycetes. Finally the genome organization of P. fijiensis was recently compared with those of the closely related banana leaf spot fungi P. musae and P. eumusae (Chang et  al., 2016). Marked differences in genome architecture were detected between species. These had likely come about from differential invasions of the genomes by transposal elements. Analysis of gene content suggested that, although the three species have a similar in size predicted arsenal of protein-coding genes, either species-specific adaptation or convergent evolution might have been implicated in pathogenic evolution of the three species.

Disease cycle and epidemiology C. Abadie, L. de Lapeyre de Bellaire, L. Pérez-Vicente, M. Guzmán and D.R. Jones Infection

Genetic and genomics A first genetic map of P. fijiensis was constructed from a cross between an African (CIRAD86) i­ solate and a Colombian (CIRAD139a) isolate using mainly AFLP markers (Manzo-Sánchez et  al.,

Spores usually germinate within 2–3 h of deposition on a moist leaf surface. The minimum, ­optimum and maximum temperatures for the development of ascospore germ tubes are 12°C, 27°C and 36°C, respectively, with no development



Fungal Diseases of the Foliage

65

both conidia and ascospores penetrate through stomata (Fig. 2.3) after 48–72 h above 20°C (Stover, 1980a; Fouré and Moreau, 1992). A film of water on the leaf surface is required for

taking place at 11°C and 38°C (Stover, 1983b; Jacome et  al., 1991; Porras and Pérez, 1997). However, ascospore germ-tube growth at 20°C is half the rate it is at 27°C. Germ tubes from

Life cycle of Pseudocercospora fijiensis Ascospore discharge period >150 days

Dispersal of ascospores

Sexual sporulation

> 100 km Pesudothecia on necrotic lesions (stage 6) Ascospores

Conidia

Latent period = 22–70 days

Spores deposition > 12 m Dispersal of conidia

Antagonist Asexual sporulation

Inoculum density

Epiphyllic growth Infection period 2–3 days

Penetration of stomata (unfurled leaves)

Symptom evolution time = conidia discharge period 10–30 days

Incubation period = 12–35 days

Conidiophores and conidia on young streaks (stage 1–3)

Fig. 2.3.  Life cycle of Pseudocercospora fijiensis (syn. Mycosphaerella fijiensis) by L. de Lapeyre de Bellaire, L. Pérez-Vicente and M. Guzmán (drawing by A.M. Granados-Cáseres).

66

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ascospore infection under controlled conditions. Water is not required for conidial infection provided that the relative humidity is high (Jacome and Schuh, 1992). It was observed that the germ tube grows epiphytically for 2–3 days before penetrating the leaf via stomata (Beveraggi et  al., 1995). Germ tubes can often be observed during this period growing over stomata near the spore deposition site and penetrating ones more distant. This phenomenon is not well understood, but is probably related to physiological events associated with spore germination. Balint-Kurti et al. (2001), using transformed strains with green fluorescent protein, observed the occurrence of a stomatopodium above the penetrated stomata. Based on experimental data and statistical inference, the estimated infection efficiency of spores for a plantain is 0.016, meaning that 62 spores are needed to create one lesion (Landry et  al., 2017). This low efficiency may be explained by a low rate of stomata penetration (< 11%) ­observed by Cavalcante et al. (2011). Although it is assumed that all leaves are equally susceptible to P. fijiensis, most infections occur on new leaves between emergence and unfurling (Stover and Simmonds, 1987; Gauhl, 1994). This is probably because the unfurling leaf is in an upright position and its cylindrical shape facilitates the deposition of air-transported ascospores. At the unfurling stage, the lower surface of the leaf, which has a greater stomatal density compared with the upper surface, is also exposed (Rodríguez et al., 2009). Incubation period The time between infection and the appearance of the first speck symptoms on the leaf is the incubation period (Fig. 2.3). On Cavendish banana cultivars, conidia can be produced as soon as the first symptoms are visible (Stover, 1980a; Fouré and Moreau, 1992). The exact time of infection in the field is not usually known, but it is assumed that penetration coincides with the time of emergence of a new rolled leaf (Fouré, 1982; Mobambo et  al., 1997). Working with this assumption, the fastest incubation period is about 10–14 days under ideal conditions for disease development. Once infection is established, one or more vegetative hyphae of P. fijiensis emerge from ­stomata on the lower leaf surface, develop into

c­ onidiophores or grow across the leaf surface parallel to the veins for distances up to 3 mm to infect adjacent stomata (Stover, 1980a; Gauhl, 1989). These epiphyllic hyphae can anastomose to form a complex network. At various intervals along the hyphae, short side-branches develop, which terminate as appressoria over stomata (Meredith and Lawrence, 1969). Movement from one stoma to another is much more common with P. fijiensis than with P. musae and this eventually results in the development of black leaf streak lesions over entire leaves (Stover, 1980a). The results of some field trials indicate that the incubation period varies with the cultivar (Meredith and Lawrence, 1970a; Firman, 1972; Fouré, 1991a). However, no significant differences in the length of the incubation period on different cultivars have been reported (Gauhl, 1994; Mobambo et  al., 1996a, 1997). The incubation period is faster on 3–5-month-old plants derived from tissue culture than on field-established plants (Mobambo et al., 1997). The incubation period can be influenced by seasonal variations in the climate. During the rainy season in Nigeria, it has been recorded as 14 days on plantain (Mobambo et  al., 1996a) with symptoms first being seen on the second youngest unfurled leaf on growing plants. During the dry season, the incubation period was 24  days (Fig. 2.4). A same fluctuation of the ­incubation period has been observed on ‘Grande Naine’ (AAA, Cavendish subgroup) in Cameroon (Fouré and Moreau, 1992). In Cuba, the incubation period varied between 14–17 days (in the summer rainy season with high diurnal and nocturnal temperatures) and 27.9 days in February (in the winter dry season with temperatures under 20º C in day and night). The difference may relate to the temperature affecting mycelial growth in the leaf t­issues (L. Pérez-­ Vicente, Cuba, 2016, unpublished results). Inoculation density also influences the incubation period. When plants of ‘Grande Naine’ were inoculated under identical controlled ­conditions with 200 conidia and ten conidia of P. fijiensis/cm2 of leaf, the incubation periods were 17.5 days and 27.0 days, respectively. A similar relationship was found between the length of the incubation period and the number of ascospores deposited on leaves. The concentration of the pathogen’s m ­ etabolites in a given



Fungal Diseases of the Foliage

67

700

35

600 30

F

25

400

Days

Rainfall (mm)

500

300

20

200 15 100

10

0 A

S

O

N

D

J

F

M

A

M

J

J

Months 1990/91 Rainfall

Symptom evolution time (SET)

Incubation time (IT)

Fig. 2.4.  The development of black leaf streak symptoms in relation to rainfall in Nigeria (from Mobambo et al., 1996a). Key: F = flowering.

area of leaf could be responsible (L.  PérezVicente, Cuba, 2016, unpublished ­results). The youngest leaf with first streaks is a parameter that is regularly monitored in commercial banana plantations. If it is assumed that the foliar emergence rate is constant over the course of the observation, then the reading is an estimate of the length of the incubation period, which is influenced by climate and the efficiency of the fungicide spray regime (Marín et  al., 2003; de Lapeyre de Bellaire et al., 2010). Symptom evolution time The number of days between the appearance of first symptoms and the appearance of mature spots with dry centres is the transition period or symptom evolution time. This period is also useful for determining the time that conidia are produced (conidial contagious period) since production starts when streaks are first visible and ends

when spots dry (Fig. 2.3). Symptom evolution time varies according to environmental conditions and the susceptibility of the cultivar. In Cameroon, climatic conditions were determined to have affected the duration of the symptom evolution time, which ranged from 11 to 30 days, on ‘Grande Naine’ (AAA, Cavendish subgroup) (Fouré and Moreau, 1992). Shade has been found to slow down symptom development, but only after speck formation. Shade can reduce the leaf area affected by black leaf streak by up to 50% (F. Gauhl and C. Pasberg-Gauhl, Costa Rica, 1999, personal observation). Gauhl (1994) in Costa Rica noted that symptoms on ‘Valery’ (AAA, Cavendish subgroup) developed faster than those on ‘Curraré’ (AAB, Plantain subgroup) after the speck stage (Table 2.5) and this led to an earlier death of leaves. Gauhl (1994) also reported that first symptoms on ‘Currare’ appeared on average about 8 days later on plants that had flowered compared with plants before flowering. Jones

68

Chapter 2

Table 2.5.  Comparison of the fastest black leaf streak symptom evolution times on ‘Valery’ (AAA, Cavendish subgroup) and ‘Curraré’ (AAB, Plantain subgroup) in Costa Rica between May and July 1986 when rainfall was fairly uniform (modification from Gauhl, 1994). Time from leaf emergence to first observed symptom Symptom

‘Valery’

‘Curraré’

Speck symptom Streak symptom Spot symptom Mature spot symptom

24 days 27 days 30 days 34 days

24 days 30 days 38 days 44 days

and Tezenas du Montcel (1994) reported transition periods in different clones that ranged from 11 to 139 days.

on Cavendish cultivars, black leaf streak spots ­appearing on leaves 5–6. It is this increased speed of development coupled with higher infection densities, because of the earlier and more abundant formation of ascospores, that makes black leaf streak so much more difficult to control than ­Sigatoka leaf spot. The factors affecting the development of symptoms of black leaf streak and Sigatoka leaf spot are presented in Fig. 2.5. In commercial banana plantations, the disease development time, as assessed by noting the YLS, is an important parameter with which to gauge the efficacy of a fungicide spray programme to control P. fijiensis. Regular YLS readings over time, again assuming that the leaf emission rate is constant, give a fairly reliable indication as to the control of the pathogen. However, for a correct interpretation of YLS, the same banana plants need to be observed over time (de Lapeyre de Bellaire et al., 2010).

Disease development time A more useful criterion for measuring the rate of disease evolution is the disease development time, which is the period between infection and the formation of mature spots. Unlike the incubation and symptom evolution times, it does not rely on the detection of subtle first symptoms that can easily be overlooked. Disease development time is often assessed as the rank of the ‘youngest leaf spotted’ (YLS). This is determined by counting the leaves down the plant from the first fully opened leaf until at least ten mature spot symptoms are seen. In practice, the youngest leaf with a large necrotic area with at least ten light-coloured dry centres has also been counted as the YLS. In this assessment, the rate of foliar emission is considered as constant, which need not always be the case. This parameter was first devised for use with Sigatoka leaf spot (Stover and Dickson, 1970; Stover, 1971b). The disease development time, assessed as the YLS, depends on the susceptibility of the cultivar, intensity of infection and environmental conditions, in much the same way as has been described for Sigatoka leaf spot (Meredith, 1970). However, the rate of onset of mature spots is much faster with black leaf streak. Under field conditions, black leaf streak spots appear on leaves 3–4 on Cavendish cultivars compared with leaves 4–5 for Sigatoka leaf spot (Stover, 1980a). On plantain, disease development is slower than

Conidiophores and conidia Conidiophores with conidia are present in lesions beginning at the initial speck or first streak stages of Meredith and Lawrence (1969) and develop under conditions of high humidity. They are produced until the appearance of necrotic spots (Blanco, 1987; Fouré and Moreau, 1992; Gauhl, 1994). The period when conidia of P. fijiensis are produced is short compared with the period that ascospores are formed (Fig. 2.3). Stover (1980a) estimated that about 1200 conidia are produced in the average 20 mm2 streak on the lower leaf surface of a Cavendish cultivar. In Cameroon, Ngando et al. (2015) counted up to 1800 conidia on stage 2 lesions (Plate 2.3) on Cavendish cultivars in commercial plantations. Conidiophore production is influenced by the weather and disease development rate. In Cameroon in 1987–1989, Fouré and Moreau (1992) counted a maximum of 44 conidiophores in each square millimetre of stage 2 streak lesions in September (rainy season), whereas no conidiophores could be found in January (dry season). Blanco (1987) observed that the production of conidia was much higher on plants before flowering than on plants after flowering. Conidia are considered to play only a minor role in the disease cycle compared with ascospores (Gauhl, 1994). However, by using spores traps in Costa Rica, Burt et  al. (1997) caught more



Fungal Diseases of the Foliage

CONIDIUM OR ASCOSPORE ON LEAF Humidity Temperature Influence of microflora

GERMINATION

PENETRATION OF STOMA

FIRST SYMPTOM Humidity Temperature Light instensity Host resistance Intensity of infection

NECROTIC SPOT

Pregermination Epiphyllic growth of germ tube

Humidity Temperature

Humidity Temperature Intensity of infection Plant vigour

69

Initial invasion of leaf tissue Dispersal of conidia affected by wind and water

Ascospore liberation affected by leaf wetness or dryness

Further invasion of leaf tissue and asexual reproduction

INCUBATION PERIOD SYMPTOM EVOLUTION TIME OR TRANSITION PERIOD

Sexual reproduction MATURE SPOT WITH GREY CENTRE

Fig. 2.5.  Stages in the development of black leaf streak symptoms and factors that influence their duration (adapted from Meredith, 1970; Fouré and Moreau, 1992).

c­ onidia than ascospores within a diseased plantation. This may have been because the amount of sexual reproduction in P. fijiensis is limited in commercial plantations because of fungicide applications and the regular removal of heavily spotted leaves. In a commercial plantation in Cameroon, Rieux et  al. (2013a) showed that P. fijiensis on plants that were sprayed with fungicide mainly produced conidia, while unsprayed plants mainly produced ascospores. An average of 32 conidia/lesion (from stage 2 to stage 4) were counted on the fungicide-treated banana plants in a commercial plantation and 14 conidia/lesion on untreated plantain. This amounted to 5000– 35,000 conidia/banana plant in the commercial plantation, which was much more than the estimated number of ascospores determined from measurements of the necrotic area of lesions (Rieux et al., 2013a; Ngando et al., 2015). Since conidia can survive on banana leaves for more than 1 month (Hanada et  al., 2002), they may play a more important role than expected in the disease cycle in fungicide-treated plantations. Recently, Ravigne et al. (2017), who used a modelling approach, suggested an important role

of conidia in disease spread. In areas where P. fijiensis has only recently been introduced, such as the French West Indies, it is believed that conidia may have played a major role in the survival of the pathogen on individual plants (C. Abadie, France, 2016, personal communication). Spermagonia and spermatia Spermatia are considered as the male gametes and are produced in spermogonia in substomatal chambers primarily on the abaxial leaf surface at the streak stage 3 or first spot stage 4 (see Plate 2.3). As are conidiophores, spermogonia are produced more abundantly on the abaxial surface of infected leaves. A growing spermogonium may displace conidiophores growing nearby (Meredith and Lawrence, 1969). Hyaline single-­celled, rod-shaped spermatia are released through the ostiole of the spermogonium, which protrudes through the stomatal pore (Churchill, 2011). Pseudothecia and ascospores Pseudothecia are abundant in lesions at the second and third or mature spot stage of Meredith

70

Chapter 2

and Lawrence (1969), which corresponds to stages 5 and 6 of Fouré (1987) (see Plate 2.3). Pseudothecia and spermogonia production is seasonally variable, especially at locations with marked rainy and dry seasons, with more developing in the humid and hot period of the year. Ascospores are forcibly ejected from the pseudothecia during periods of wet weather or, in the absence of rain, in low numbers during the night, reaching a maximum before dawn as dew settles (Meredith et al., 1973; Gauhl, 1994). Ascospores can be produced in leaves over a long period (see Fig. 2.3). Hanging banana leaves with lesions discharged ascospores for up to 21 weeks (Gauhl, 1994). In Costa Rica, Gauhl (1994) found that ascospores formation and production were usually greater in lesions on the lower surface of plantain leaves. However, more pseudothecia and ascospores were reported to be produced in lesions on the upper leaf surface of banana cultivars in the Cavendish subgroup grown in plantations in Costa Rica (Table 2.6) and Cuba (Guzmán et  al., 2005b; Villalta and Guzmán, 2005; L. Pérez-Vicente, 2008, personal communication). This did not seem dependent on whether plants were being sprayed with fungicides or not. Only one crop of ascospores is produced in each pseudothecia (Stover, 1980a), but pseudothecia continue to mature in necrotic leaves. Dead leaves, which may hang on the plant or lie

on the ground after pruning operations, are a major source of inoculum. In Costa Rica, Gauhl (1994) monitored the discharge of ascospores from dead plantain leaves on the ground and ­observed large numbers released during the first 3 weeks with further releases over the next 6 weeks. Large numbers of ascospores were ­released during the first 15 days from leaves of a Cavendish cultivar lying on the ground in a plantation without chemical control and during the first 23 days in a plantation with chemical control. Almost no ascospores were discharged after 30 days. Sporulation was greater in lesions on the upper leaf surface compared with the lower and in plantations practising chemical control compared with those without chemical control (Fig. 2.6 and Table 2.6). The shorter time of sporulation and the lesser amount of sporulation recorded in leaves on the ground of the plantation without chemical control were associated with a greater amount of degradation of leaves by saprophytic microorganisms. In plantations where plants are sprayed, these microorganisms are inhibited by fungicide residues on the leaves (Guzmán et  al., 2005b; Villalta and Guzmán, 2005). In long-term epidemiological studies on black leaf streak in Costa Rica (Blanco, 1987; Gauhl, 1994) and Nigeria (Gauhl and Pasberg-­ Gauhl, 1994), ascospores were shown to be the most common form of inoculum. These results

Table 2.6.  Numbers of ascospores of Pseudocercospora fijiensis discharged from one lesion (stage 6 of Fouré, 1987) on the upper leaf surface and one lesion on the lower leaf surface of a ‘Grande Naine’ (AAA, Cavendish subgroup) plant in plantations with and without chemical control in three different experiments in Costa Rica. Each value is the sum of nine optical field readings of a microscope at 40× in the discharge area of the lesion on a water-agar medium (R. Villalta and M. Guzmán, Costa Rica, 2004, unpublished results). Plantation with chemical control

Leaf surface Upper Lower P > Fa

Experiment 1 May–June

Experiment 2 June–July

Experiment 3 August–September

56 37 0.0940

58 42 0.3736

181 85 0.0195

Plantation without chemical control Upper Lower P > Fa T-test (P = 0.05).

a

19 10 0.0768

73 41 0.0908

114 42 0.0037



Fungal Diseases of the Foliage

71

90

200

80

175 150

60

125

50

100

40

75

30

Rainfall (mm)

Sporulation (%)

70

50

20

25

10

0

0 0

8

15

29

23

36

43

Time (days) Cumulative rainfall Upper-untreated

Upper-treated Lower-untreated

Lower-treated

Fig. 2.6.  Percentage of lesions of Pseudocercospora fijiensis sporulating on the upper and lower surface of leaves of ‘Grande Naine’ (AAA, Cavendish subgroup) lying on the ground in two plantations in Costa Rica. One plantation was regularly treated with fungicide and the other was left untreated (R. Fillalta and M. Guzmán, Costa Rica, 2014, unpublished results).

confirm observations in Colombia (Mayorga, 1985), Fiji (Firman, 1972) and Hawaii (Meredith et al., 1973). Mature pseudothecia need to be impregnated with water before ascospores can be discharged (Stover, 1976; Fouré and Moreau, 1992). Therefore, spore release can follow a seasonal pattern, with only low numbers of ascospores being released during the dry season and high numbers during periods of higher rainfall (Gauhl, 1994; Gauhl and Pasberg-Gauhl, 1994). In the Atlantic lowlands of Costa Rica, Gauhl (1994) recorded the highest concentration of ascospores (6876/m3 of air) in December, which is the wettest month (Fig. 2.7). Meredith et al. (1973) also found the highest ­daily ascospore concentrations during the rainy seasons in Hawaii and Fiji (Fig. 2.7). Ascospore counts in Nigeria also peaked during the rainy season, but were found to be much lower than in other regions of the world (Gauhl and Pasberg-Gauhl, 1994) (Fig. 2.7). The lower levels of inoculum recorded in Nigeria may have been because spores were trapped at a location remote from large banana plantations. Rainfall induces ascospore release if mature pseudothecia are present. The minimum amount of rain necessary to induce ascospore

release is as low as 0.1 mm/h (Gauhl, 1994). Thal et  al. (1992) observed in Costa Rica that rains of 0.1–0.2 mm were usually sufficient to increase spore counts. When preceded by a dry period of 24 h or longer, more ascospores were released during rains of at least 0.5–1.0 mm than during lighter rains. Ascospore counts were usually found to be significantly higher during the day compared with the night. Discharge can occur within 1 h or so after rain (Burt, 1993; Gauhl, 1994). Ascospores discharged during long periods of continuous rain can be washed out of the air and off foliage. Ascospore release in laboratory experiments starts 10 min after wetting leaf tissues and is at its maximum during the next 30 min. Discharge can last for 60–75 min (Pérez-Vicente, 1998). Temperature also influences ascospore discharge. Less ascospore production was observed during periods with daily minimum temperatures below 20°C (Gauhl, 1994). Burt (1993) reported a decrease in spore release when the temperature was under 19°C or over 33°C. In Cameroon, Fouré and Moreau (1992) found many pseudothecia in coalescing mature spots in the wet season, but very few pseudothecia in individual mature spots during the dry

72

Chapter 2

35,000 30,000

Ascospores

25,000 20,000 15,000 10,000 5,000 0 0

2

4

Costa Rice 1985 Nigeria 1993

8 6 Months

10

12

Nigeria 1992 Fiji 1969

season. Spots probably did not coalesce in the dry season because low humidity and the absence of rain may have inhibited lesion spread by epiphyllic hyphae. This, coupled with restrictions on the movement of spermatia imposed by a lack of surface moisture, may have reduced the chances of cross-fertilization necessary for the production of pseudothecia and ascospores. In Costa Rica, Blanco (1987) counted 0.3– 1.2 pseudothecia in each 1 mm2 of mature lesion on the upper leaf surface of a Cavendish cultivar and 0.6–3.7 pseudothecia on the lower surface. Although Gauhl (1994) found that there was no difference in the numbers of ascospores discharged from upper and lower leaf surfaces, Blanco (1987) demonstrated that more ascospores were discharged from the lower leaf surface than from the upper. This correlates with the number of pseudothecia he found on each surface. Also in Costa Rica, Burt et al. (1999) calculated that an average of 4.5 ascospores were discharged from each pseudothecium, which seems low given that many more ascospores would be present. They also proposed a linear model for determining numbers of ascospores that can potentially be released from each square centimetre of necrotic lesion area on a leaf.

Hawaii 1969

Fig. 2.7.  Ascospore counts (total for month) in different regions of the world (after Meredith et al., 1973; Gauhl, 1994; Gauhl and C. Pasberg-Gauhl, Costa Rica, 1999, unpublished results).

Spore dispersal Pseudocercospora fijiensis spreads via ascospores and conidia. However, the spore-release mechanisms as well as aerodynamic properties of both spores are noticeably different. These differences confer very distinct dispersal properties. Conidia become dislodged by wind and water (Stover, 1980a), but, contrary to earlier work that implicated water-splash in disease epidemiology (Meredith et al., 1973), wind has been identified as the main agent that carries conidia to nearby plants (Rutter et  al., 1998; Rieux et  al., 2014). The scars on the conidia and conidiophores at the point of attachment are believed to facilitate the wind removal of conidiospores. Ascospores are actively discharged into the air from pseudothecia before being passively wind dispersed. They are commonly found in the air above the canopy of plantations following rainfall (Thal et al., 1992; Gauhl, 1994). Air samplers or spore traps have been used to determine the relative importance of ascospores and conidia in the spread of P. fijiensis inside and outside plantations in a mountainous area of Costa Rica (Burt et  al., 1997; Rutter et al., 1998) and in the banana production area



Fungal Diseases of the Foliage

in the Atlantic coast area (Gauhl, 1994). These studies showed that both types of spore are present in the air inside a banana plantation. However, results on the prevalence of each spore type differed. Conidia were determined to be more common inside plantations in the mountainous area (Burt et  al., 1997). In the Atlantic coast area, ascospores were usually much more prevalent under the canopy. The amount of ascospores trapped in a 1-year period was found to exceed the amount of conidia by a factor of 100 (Gauhl, 1994). Gauhl (1994) concluded that conidia were of secondary importance to ascospores in the spread of the disease. Ascospores were more common outside plantations, supporting the hypothesis that ascospores might be carried over long distance whereas conidia are mainly dispersed over short distances (Burt et al. 1998). Burt et al. (1997), Rutter et al. (1998) and Amil et al. (2007) carried out studies to measure dispersal distances by spore trapping or disease gradients analysis. The dispersal distance is usually described by a dispersal kernel, which gives the probability distribution of the distance travelled by any individual spore. However, the determination of an accurate dispersal kernel was not achieved, either because external contamination could not be avoided (Burt et al., 1998) or because disease gradients were observed on several disease cycles (Rutter et  al., 1998; Amil et al., 2007). More recently, Rieux et al. (2014) made an assessment of dispersal kernels for ascospores and conidia by analysing disease levels on trap plants at different distances from specific marked sources of inoculum. In such experiments, external and internal contaminations were specifically avoided. Ascospores dispersed over an average distance of 160 m and as far as 1000 m, which was the distance from the ascospore source to the furthest trap plants. The kernel that best fitted the results was an exponential function displaying a heavy tail (high probability of long-distance dispersal events) and a very steep gradient in the first few metres. Therefore, it is likely that ascospores might ­disperse at distances longer than 1 km. Since ­ascospores are killed after 6 h exposure to ultraviolet radiation, the distance of dissemination of viable spores is determined not only by the speed of the wind, but also by cloud cover and the time of day that spore release occurs (Parnell et  al., 1998). Because of this, Parnell et  al. (1998)

73

c­oncluded that wind dispersal of viable ascospores over distances greater than a few hundred kilometres is unlikely. Rieux et  al. (2014) estimated that conidia dispersed at an average distance of 4 m and no conidia were trapped over 12.5 m from the conidial source. They concluded that conidia probably only play a major role in the establishment of fungal populations on the source plant and neighbouring plants. These results confirm earlier assumptions. An epidemiological host–­pathogen model was developed recently to simulate the dynamics of black leaf streak disease under optimum conditions for the pathogen. The model shows that infection efficiency, the incubation period and the lesion growth rate strongly influence the severity of the disease, but not the dispersal distance of conidia (Landry et al., 2017). Black leaf streak versus Sigatoka leaf spot Black leaf streak is well suited for the environmental conditions prevailing in the coastal tropics and has now virtually replaced Sigatoka leaf spot as the dominant banana leaf-attacking disease in these areas. This process took 2–3 years in Honduras (Stover, 1980a) and less than 5 years in Costa Rica (Gauhl, 1994) and St Lucia (C. Abadie, France, 2016, personal communication). In parts of Brazil, where black leaf streak is only now encroaching on areas where Sigatoka leaf spot is present, it is still possible to see symptoms of both pathogens on the same ­banana ­ aribbean, leaf (see Plate 2.4). In the French C black leaf streak progression was slowed because of a regular de-leafing programmes undertaken on plantains and Cavendish cultivars and because incoming P. fijiensis populations were highly sensitive to fungicides used for disease control while established populations of P. musae were much less sensitive (Chilin-Charles, 2010; de Lapeyre de Bellaire et al., 2015). Black leaf streak lesion expansion and ascospore production are greater in the coastal tropics than those for Sigatoka leaf spot (Mouliom-­ Pefoura et al., 1996) and this has probably given this disease a competitive advantage. However, the situation has been different at altitude. ­Sigatoka leaf spot seems more adapted to cooler environments and here lesions grow faster than those of black leaf streak in upland areas (Mouliom-­Pefoura et al., 1996).

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Chapter 2

In Costa Rica in 1985, both Sigatoka leaf spot and black leaf streak diseases were identified on the same banana leaf at 900 m, but only Sigatoka leaf spot was found at 1200 m (Romero and Gauhl, 1988). However, over time, black leaf streak is being observed at higher and higher altitudes. In 1990, in the same area of Costa Rica as was surveyed in 1985, black leaf streak was found at altitudes up to 1500 m (F. Gauhl and C. Pasberg-Gauhl, Costa Rica, 1999, unpublished). The presence of P. fijiensis at 1320 m of altitude in Costa Rica was confirmed by Arzanlou et  al. (2007a) using molecular techniques. In 1988, 7 years after its introduction to Colombia, black leaf streak was found for the first time in the plantain production areas in the Cauca valley at 1500 m (Merchán, 1990). In 1991, it had reached 1600 m (Belalcázar, 1991). This advance in range to higher altitudes is believed to be because P. fijiensis is adapting to cooler environments, which makes the pathogen more competitive at altitude (Pasberg-Gauhl et  al., 1999). However, this phenomenon could also be a consequence of climatic change or a combination of both. Climatic change models predict that, in the future, areas favourable for the development of black leaf streak today will gradually become less favourable because of an increase in the ­frequency of drought periods. Conversely, areas ­unfavourable today are likely to become more favourable because of warming. However, despite any changes that may occur, extensive areas will continue to be favourable for the occurrence of the disease (Ghini et al., 2007; de Jesús-Junior et al., 2008; Ramírez et al., 2011).

Host reaction C. Abadie, J. Carlier, F. Carreel, M. Guzmán, P. Lepoivre, X. Mourichon and, L. Pérez-Vicente F. Evaluating Musa accessions for reaction to black leaf streak Banana cultivars in the Cavendish and Plantain subgroups are known to be susceptible to black leaf streak. In commercial plantations, especially those that produce fruit for export, the disease is controlled by the frequent applications of

f­ ungicide and cultural practices. However, chemical control is neither durable nor environmentally safe. It is also very expensive. Thus, the cultivation of cultivars resistant to black leaf streak, which do not require spraying, is considered to be a much better option. Resistant cultivars would also benefit small-scale and subsistence growers, who cannot afford fungicides and consequently may suffer appreciable yield losses (Marin et  al., 2003). The evaluation of Musa accessions for their reaction to black leaf streak is primarily aimed at identifying those clones that may have useful resistance. These clones could be useful as replacements for existing susceptible cultivars or have potential as parental material in banana breeding programmes. The evaluation of reaction of germplasm to black leaf streak is especially important for banana improvement programmes (Stover and Simmonds, 1987; Rowe and Rosales, 1999; Bakry et  al., 2009). Musa accessions obtained from gene banks or collecting expeditions need to be tested to determine their suitability as parental material (Bakry and Horry, 1994) and also the resistance of hybrids derived from breeding (Jenny et  al., 1994; Vuylsteke et  al., 1995). Clones treated by non-conventional approaches for improvement, such as mutagenesis (Roux, 2004), natural somaclonal variation (Nwauzoma et  al., 2002) and genetic modification (Kovacs et al., 2013) also need to be evaluated. Methods for determining host reaction evaluation in the field. 

Numerous studies on the evaluation of wild Musa species and banana cultivars for reaction to black leaf streak in the field under conditions of natural infection have been undertaken. Sometimes, the results of such studies have been contradictory, which could be because different methodologies have been used to determine levels of ­resistance and susceptibility (Tezenas du M ­ ontcel, 1990). Differences in virulence and aggressiveness of local populations of P. fijiensis (Fullerton and Olsen, 1991, 1993, 1995) and in environmental conditions at test sites (Vuylsteke et al., 1993a) may also account for some discrepancies. Other contradictory results are not so easy to understand. Meredith and Lawrence (1970a) in Hawaii were the first to document the field reactions



Fungal Diseases of the Foliage

of cultivars attacked by P. fijiensis. Some of the ­parameters used to determine the reaction category were: (i) the number of the youngest leaf with ‘initial speck stage’ symptoms counting down from the first fully unfurled leaf; (ii) the number of the youngest leaf bearing ‘first spot stage’ symptoms; (iii) the percentage area on the oldest living leaf occupied by spots and the intensity of streaking in five randomly selected 5 cm2 areas on the youngest leaf with the ‘first spot stage’; (iv) the average area of streaks on the second youngest leaf showing symptoms; (v) the number of functional leaves at harvest; and (vi)  the number of conidiophores per square ­millimetre on the lower surface of ‘second stage streaks’. The parameters based on diseased leaf rank are strongly dependent on plant growth, which can differ between cultivars. Although an almost continuous gradation of responses was recorded, cultivars were placed into three reaction categories: ‘very susceptible’, ‘moderately susceptible’ and ‘slightly susceptible’. The evaluation showed that there was much less natural resistance to P. fijiensis among cultivars than there was to P. musae. Firman (1972), working in Fiji, used criteria similar to Meredith and Lawrence (1970a), but he also noted the number of functional leaves at flowering. He experienced difficulty in measuring the intensity of streaking and areas of individual streaks. Problems were also encountered in measuring the production of conidia and ascospores. His results were similar to those of Meredith and Lawrence (1970a). Fouré (1982, 1985, 1987, 1989) and Fouré et al. (1984) evaluated the reaction of cultivars to black leaf streak in Gabon and Cameroon using parameters focused on the disease development speed, which is independent of the leaf emission rate. These were: 1. The time taken from infection to the appearance of first symptoms (incubation period). 2. The time taken from the appearance of first symptoms to the appearance of the first mature necrotic spot or necrosis due to the merging of earlier stages. 3. The intensity of asexual and sexual ­sporulation. 4. The number of functional leaves at ­harvest. These studies revealed a gradation of reactions to P. fijiensis from complete susceptibility, to

75

slight susceptibility or partial resistance through to a high level of resistance. Mobambo et al. (1996a, b) considered that field evaluation was best undertaken in Nigeria during the rainy season with the ratoon crop, rather than the plant crop. However, these authors developed a much quicker method whereby they screened young plants derived from tissue culture after they had been planted in the field for 2 months (Mobambo et al., 1997). Five parameters were used to determine the reaction of three bred plantain hybrids to black leaf streak. In this work, Musa acuminata ssp. burmannica (AAw, ­accession ‘Calcutta 4’), which was the male parent, was used as a highly resistant control and ‘Obino l’Ewai’ (AAB, Plantain subgroup), which was the female parent, as a susceptible control. The parameters were as follows. 1. Incubation time (time from infection to appearance of first symptoms). Infection was assumed to have occurred at the time the unfurling leaf was at stage B as described by Brun (1962) (Fig. 2.8). 2. Symptom evolution time (time from appearance of first symptoms to appearance of spots with dry centres). 3. Disease development time (time from infection to appearance of spots with dry centres). 4. Number of the youngest leaf spotted (YLS) as defined earlier. 5. Life of leaf (time from cigar leaf stage B to leaf death). The results showed that the incubation time parameter was similar for all clones tested, suggesting that it is not a useful measurement for determining reaction to black leaf streak. The YLS parameter was shown to correlate well with the disease development time reading (Mobambo et  al., 1996a). However, the former was easy to score and the latter time-consuming. The YLS parameter was used by Ortiz and Vuylsteke (1994) in the evaluation of 101 hybrids. From their results, four levels of host response to black leaf streak were distinguished. ‘Susceptible’ clones were defined as having < 8 leaves without spots before or at flowering, ‘less susceptible’ clones as having 8–10 leaves without spots at flowering, and ‘partially resistant’ clones as having > 10 leaves without spots at flowering. Clones designated as ‘highly resistant’ had no spots at all. It must be borne in mind that leaf

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Chapter 2

A

B

C

D

E

Fig. 2.8.  Stages of the unfurling of the heart leaf (after Brun, 1962).

emission rates on plants need to be similar in order for results derived from the YLS reading to be meaningful and comparable. Since 1990, the International Network for the Improvement of Banana and Plantain (INIBAP), which is now part of Bioversity International, has coordinated the worldwide testing of new hybrids from the major banana breeding programmes under its International Musa Testing Program (IMTP). The four parameters chosen for the detailed evaluation of reaction to black leaf streak (Carlier et al., 2002) were as follows. 1. Disease development time (time from infection, which is assumed to occur at the B stage of leaf emergence as described by Brun (1962) (Fig. 2.8), to the appearance of ten or more discrete fully mature spots or a large necrotic area with ten or more light-coloured dry centres). 2. Number of the YLS (number of the first leaf counting down from the first fully opened leaf that had ten or more discrete, fully mature spots or a large necotic area with ten or more light-­ coloured dry centres). 3. Leaf emission rate (number of new leaves produced each week). 4. Disease severity on each upright leaf expressed as the percentage of the necrotic area/ banana leaf measured using a visual scale as ­defined by Gauhl (1994) at 6 months after planting, bunch emergence and harvest. A set of control clones, known to be susceptible, partially resistant and highly resistant, were

recommended for use in each screening trial as references. Evaluations must be undertaken during the plant’s first cycle and on the first ratoon crop (second cycle). A measurement of leaf emission rates allowed for differences in growth rates to be taken into consideration in the analysis of results. Results from different trial locations can also be compared and variations in pathogen virulence or aggressiveness detected. INIBAP has also published simplified guidelines for data collection for a less detailed assessment of reaction called ‘performance evaluation’ (Carlier et al., 2003). Only the YLS at flowering and the number of erect leaves (beginning at the youngest unfurled leaf and counting down the plant) at flowering and harvest need to be recorded. The latter information is used to calculate the ‘index of non-spotted leaves’, which is the proportion of leaves without the typical latestage symptoms of black leaf streak disease. This index provides an estimation of available photosynthetic leaf area prior to fruit filling. It also corrects the differences in the number of leaves produced by different types of banana. For meaningful results, the guidelines recommended that clones to be evaluated should be planted about 2.5 m apart in blocks arranged according to a random experimental design. A susceptible cultivar is also planted in and around the blocks to produce homogenous inoculum. evaluation by artificial inoculation. 

Mourichon et al. (1987) successfully inoculated 2–3-month-old



Fungal Diseases of the Foliage

banana plants derived from tissue culture growing in pots in the glasshouse by applying conidia, which were obtained from cultures of P. fijiensis, in a suspension of water on to lower leaf surfaces. A relative humidity level of 80–95% was ­considered essential for infection. Plants were inspected weekly and symptom evolution recorded over time. Fullerton and Olsen (1995), Romero and Sutton (1997b) and Alvarado-Capo et al. (2003) also successfully used this method. Although conidial suspensions at known concentrations are preferred as inoculum, mycelial fragments (counted as fragments per millilitre or in mg/ml) have also been utilized when conidial production in cultures has been problematic (Donzelli and Churchill, 2007; Twizeyimana et  al., 2007; Peraza-Echeverria et  al., 2008; Torres et  al., 2012; Kovacs et  al., 2013; Leiva-Mora, 2015). The lower sides of leaves are usually inoculated with the inoculum applied either in droplets, in a fine spray or by using a camel-hair brush (Donzelli and Churchill, 2007; Twizeyimana et  al., 2007; Abadie et  al., 2008; Torres et  al., 2012; Kovacs et  al., 2013; Leiva-­ Mora, 2015). Symptoms are usually more severe and evolve faster when mycelial fragments are used. Disease levels are also found correlated more with the amount of applied mycelium than the degree of fragmentation of hyphae (Donzelli and Churchill, 2007). Tween 20 or 80, Triton X-100, Silwet L-77 or gelatin are sometime used to facilitate inoculum homogenization of the solution and/or adhesion to the leaf (Donzelli and Churchill, 2007; Abadie et al., 2008). Controlled production of ascospores is difficult in the laboratory because P. fijiensis is a heterothallic fungus. Plant materials used for evaluating host reaction have ranged from plantlets derived from tissue culture to leaves taken from plants growing in the field, the latter needing to be surface sterilized. However, leaves or detached leaf pieces collected from plants growing in glasshouses and placed with their upper surfaces on an agar medium in Petri dishes and incubated in growth chambers are now more commonly used (Donzelli and Churchill, 2007; Twizeyimana et al., 2007; Abadie et al., 2008; Torres et al., 2012; Kovacs et al., 2013; Leiva-Mora, 2015). Abadie et al. (2003, 2008, 2009) inoculated pieces of detached leaves, usually with an area of 16–25 cm², cut from 3–6-month-old plants

77

derived from tissue culture. The challenge in developing detached leaf methods has been ­ to maintain the normal physiology of excised ­banana leaf pieces for the 2–3 months duration of the evaluation. Cytokinin, benzimidazole or ­gibberellic acid have all been added to the culture medium in attempts to minimize senescence, with the latter extending the survival rate of ­excised leaf fragments the greatest (Twizeyimana et al., 2007; Abadie et al., 2009). Twizeyimana et al. (2007) developed an evaluation test using in vitro plantlets growing on a culture medium in tubes. The authors found that disease development was more rapid on in vitro plants than on detached leaves, but reaction results were similar. However, more studies are needed to confirm the feasibility of this method using plants grown in tubes. In artificial inoculation work, the evaluation of host reaction to one isolate of P. fijiensis and the determination of the virulence/aggressiveness of a number of isolates of P. fijiensis to one host genotype are assessed by visual observations of disease severity and disease development over time. Disease severity is expressed as the proportion of the leaf or leaf piece area diseased; and disease development is calculated by noting the time of expression of streak and spot symptoms. Donzelli and Churchill (2007, 2009) and Carreel et al. (2013) captured symptoms of the inoculated area on a digital camera or scanner and then used image analysis software to determine disease severity. advantages and disadvantages of the different evaluation strategies. 

Whether natural infestations under field conditions or artificial infestations under in vitro conditions are used for evaluation depends mainly on scientific aims. The evaluation of host reaction on mature plants under field conditions is the most reliable, as the plant and pathogen are growing under natural conditions. However, this method is time consuming and relatively costly in terms of manpower and land. It is also unsuited for rating large numbers of clones developed by breeding programmes and infection is dependent on environmental conditions. In addition, in areas where more than one Pseudocercospora leaf spot pathogen is found, problems arise in the identification of the causal agent of the symptoms ­being noted.

78

Chapter 2

Evaluation methods in the glasshouse and laboratory using young plants and leaf pieces are more suited for the evaluation of large numbers of clones/hybrids. Also, known doses of inoculum can be reproduced and environmental conditions replicated. Inoculating leaf pieces from 4–6-month-old plants, Abadie et al. (2003) observed similar reactions on seven genotypes that had been grown under field conditions, showing that the results can be comparable. However, great care needs to be taken with artificial methods involving very young banana tissues, which may not react exactly the same as mature banana tissues. The results need to agree with those found on gemplasm with known reactions growing in the field and must be reproducible. Rapid evaluation methods are useful for preliminary screening of somaclonal variants (Nwauzoma et al., 2002), induced mutants (Roux, 2004) and transgenic material (Kovacs et  al., 2013). Abadie et  al. (2009) described the process of evaluation of new hybrids in which both field and laboratory screening methods were used, the latter allowing resistance components to be characterized. Classification of Musa–P. fijiensis interactions From field evaluations of the response of banana to black leaf streak, Fouré et al. (1990) and Fouré (1992, 1994) proposed three types of reaction to the disease. Phenotype 1 is an incompatible interaction and phenoytpes 2 and 3 are compatible interactions. interaction.  Phenotype 1. The e­ xpression of very high resistance (HR), characterized by the apparent blockage of the development of symptoms at the early streak stage (Plates 2.5 and 2.8). This reaction is believed to be close to hypersensitivity and the pathogen does not sporulate. Resistance of this type could be controlled by a single host resistance gene or a small number of host resistance genes giving the plant ‘vertical’ resistance, which may be ­easily overcome by mutations in the pathogen population. Most accessions of subspecies of ­ M. acuminata and the cultivars ‘Pisang Lilin’ (AA), ‘Tuu Gia’ (AA) and ‘Yangambi Km 5’ (AAA, Ibota subgroup) react in this way. A loss of resistance of ‘Yangambi Km 5’ in Cameroon (Mouliom ­Pefoura, 1999) and Costa Rica (Escobar-Tovar

incompatible

et al., 2015b) have been reported. Fullerton and Olsen (1995) explained the rapid change in status of ‘Paka’ (AA) and ‘T8’ (a tetraploid hybrid bred using ‘Paka) in the Cook Islands from immunity to susceptibility by a change in virulence of local populations of P. fijiensis. Some histological and biochemical studies have indicated the activation of host defence reactions soon after the pathogen penetrates the stoma. Peroxidase activity has been associated with the hypersensitivity-like reaction shown by Musa acuminata ssp. burmannica (AAw, accession ‘Calcutta 4’) (Cavalcante et al., 2011). compatible interaction.  The compatible interaction has been divided into two phenotypes. Phenotype 2 has grades of expression ranging from strong partial resistance to almost complete susceptibility. The completely susceptible reaction is known as phenotype 3. Phenotype 2. The expression of partial resistance (PR), is characterized by the normal, but slow, development of symptoms to leaf spots and necrosis (Plate 2.8). During this interaction, the pathogen produces spores. Resistance of this type may be controlled by many genes, giving the plant durable or ‘horizontal’ resistance (McDonald and Linde, 2002). Plants may have a large number of functional leaves at harvest. Cultivars showing moderate partial resistance include ‘Pisang Mas’ (AA, Sucrier subgroup), while ‘Pisang Ceylan’ (AAB, Mysore subgroup) and ‘Fougamou’ (ABB, Pisang Awak subgroup) have strong partial resistance. These categories are the equivalent of the moderately susceptible and slightly susceptible divisions of Meredith and Lawrence (1970a). In Cuba, the erosion of the partial resistance of ‘FHIA-18’ (AAAB – bred Pome-type hybrid) was observed in some locations. This may have been because of nutrition and water stresses or/and an increase of aggressiveness of local populations of P. fijiensis (Pérez-Miranda et al., 2006; Pérez et al., 2006). Phenotype 3. The expression of pronounced susceptibility (S) is characterized by the rapid development of symptoms from streaks to mature leaf spot and necrosis (Plate 2.8). The pathogen may sporulate profusely if environmental conditions are favourable. Few functional leaves, if any, remain on the plant at harvest (see Plate 2.1). All the important Cavendish cultivars (AAA),



Fungal Diseases of the Foliage

79

Plate 2.8.  Examples of the three types of reaction of Musa to Pseudocercospora fijiensis proposed by Fouré et al. (1990) and Fouré (1992, 1994). Phenotype 1 – high resistance (HR) responses shown on upper row of images: Musa acuminata ssp. burmannica (AAw, accession ‘Calcutta 4’) (upper left); M. acuminata ssp. malaccensis (AAw, accession ‘Selangor’) (upper centre); and ‘Tuu Gia’ (AA) (upper left). Phenotype 2 – partial resistance (PR) responses shown on middle row of images: ‘Pisang Ceylan’ (AAB, Mysore subgroup) (middle left); ‘Pisang Berlin’ (AA) (middle centre); and ‘Pisang Mas’ (AA, Sucrier subgroup) (middle right). Phenotype 3 – susceptible (S) responses shown on lower row of images: ‘Dominico’ (AAB, Plantain subgroup) (lower left); ‘Cocos’ (AAA, Gros Michel subgroup) (lower centre); and ‘Grande Naine’ (AAA, Cavendish subgroup) (lower right). All photographs were taken at the same location at La Rita, Pococí, Costa Rica on the same day with all plants under high inoculum pressure (photos: M. Sanchez, C. Carr, F. Alfaro and M. Guzmán, CORBANA).

such as ‘Dwarf Cavendish’, ‘Grande Naine’, ‘Williams’ and ‘Robusta’, and also all plantain cultivars (AAB) fall into this category. However, disease expression can be greatly influenced by

environmental conditions and inoculum potential. This category is the equivalent of the very susceptible division of Meredith and Lawrence (1970a).

80

Chapter 2

Reaction of Musa germplasm under field conditions Symptoms of black leaf streak have been identified on cultivars in the Eumusa series of edible banana, subspecies of Musa acuminata and M. balbisiana. The disease does not affect M. textilis (abacá) or Ensete ventricosum (enset), according to Gauhl (1994). However, enset has recently been reported as susceptible in southwest ­Ethiopia (Gurmu et al., 2017). One of the first attempts to rate banana cultivars for their reaction to black leaf streak was made by Meredith and Lawrence (1970a) in Hawaii (Table 2.7). Later, over 350 accessions, which belonged mainly to the Eumusa series of cultivars, but also included wild diploids, were tested for their reaction to black leaf streak in the field in Cameroon by Fouré. The response of these accessions is presented in Table 2.8.

Although the results presented in Table 2.8 were obtained in Cameroon in one particular ­environment and may not necessarily reflect reactions observed elsewhere, they do generally confirm the findings made earlier by other authors in the Pacific (Meredith and Lawrence, 1970a; Firman, 1972; Pearson ­ et al., 1983; Gonsalves, 1987), in Latin America (Stover, 1983a; Pasberg-Gauhl, 1990) and in West Africa (Mobambo et  al., 1993, 1994a; ­Ortiz et al., 1993, 1995; Vuylsteke et al., 1993a, 1995). The summary of results in Fig. 2.9 reveals that M. balbisiana and cultivars belonging to genomic groups containing at least one B ­component (AB, AAB, ABB) all show susceptible (phenotype 3) or partially resistant (phenotype 2) compatible reactions. The behaviour of cultivars within subgroups, with a few minor exceptions, is generally fairly uniform. The highly

Table 2.7.  Reaction of different banana clones to Pseudocercospora fijiensis (modification of Table 8 of Meredith and Lawrence, 1970a). Genomic group AA AAA

AAAA AAB

ABB

Very susceptible ‘Gros Michel’b ‘Cocos’b ‘Dwarf Cavendish’c ‘Robusta’c ‘Valery’c ‘Giant Cavendish’c ‘Lacatan’c ‘Pome’ ‘Pisang Rajah’e ‘Moongil’f ‘Platano Enano’g ‘Walha’h ‘Eslesno’i

Moderately susceptible

Slightly susceptible

‘Sucrier’ ‘Green Red’

‘Tuu Gia’a

I.C.2d ‘Silk’ Horn plantain-types Mai’a Maoli-types Popoulu-types Iholena-types ‘Father Leonore’i ‘Monthan’j ‘Ice Cream’k ‘Largo’j

‘Saba’

Cultivar from Vietnam. Cultivars in Gros Michel subgroup. c Cultivars in Cavendish subgroup. d Tetraploid from the Trinidad breeding programme. e Synonym of either ‘Pisang Raja’, ‘Celat’ (Stover and Simmonds, 1987) or ‘Nendrapadathi’ (see Table 2.8). f ‘Horn’ plantain from southern India with one hand of very large fruit. g Dwarf ‘Horn’ plantain from Puerto Rico (Stover and Simmonds, 1987). h Synonym of ‘Rajapuri’ (Stover and Simmonds, 1987). i Believed to have been introduced from West Indies and may be ABB types (Stover and Simmonds, 1987). j Cultivars in Bluggoe subgroup. k Synonym of ‘Ney Mannan’. a b



Fungal Diseases of the Foliage

81

Table 2.8.  Reaction of Musa species/subspecies, subgroups and clones to black leaf streak at Njombe, Cameroon (80 m altitude). Genome

Species/subspecies

Accessions evaluated

Reactiona

AAw

‘Madang’, ‘Banksii AF’b

S and HR

‘Calcutta 4’

HR

‘Pahang’ and ‘Kluai Pa (Musore) ×’

HR

‘Pisang Cici Alas’c

HR

‘Kluai Pa (Rayong)’, ‘Kluai Khae (Phrae)c and ‘Kluai Pa (Songkla)’c

HR

BBw

Musa acuminata ssp. banksii Musa acuminata ssp. burmannica Musa acuminata ssp. malaccensis Musa acuminata ssp. microcarpa Musa acuminata ssp. siamea Musa acuminata ssp. truncata Musa acuminata ssp. zebrina Musa balbisiana

Genomic group AA

AAA

PR*

Subgroup/cloned

Cultivars evaluated

Reactiona

Sucrier Pisang Lilin Others

‘Kirun’, ‘Figue Sucrée’ ‘Pisang Lilin’ ‘Nzumoheli’, ‘Niyarma Yik’, ‘Sow Muk’, ‘Bie Yeng’, ‘Mambee Thu’, ‘Samba’, ‘Chicame’, ‘Pallen Berry’, ‘Pisang Bangkahulu’, ‘Akondro Mainty’, ‘Galeo’, ‘Guyod’ ‘Pisang Berlin’, ‘Pisang Madu’, ‘Mak’, ‘Pisang Perecet’, ‘Pa Pathalong’, ‘Kluai Thong Det’, ‘Doumboumi’, ‘Pisang Tongat’ ‘Tuu Gia’, ‘Pisang Oli’ ‘Gros Michel’, ‘Cocos’, ‘Highgate’ ‘Petite Naine’, ‘Grande Naine’, ‘Poyo’, ‘Robusta’, ‘Valery’, ‘Americani’, ‘Lacatan’ ‘Inokatoke’, ‘Intuntu’, ‘Igitsiri’, ‘Mbwazirume’, ‘Nakitengwa’ ‘Figue Rose’, ‘Figue Rose Verte’, ‘Figue Rose Naine’ ‘Yangambi Km 5’e, ‘Kluai Khom Bao’, ‘Kluai Khai Thong Ruang’ ‘Orotava’, ‘Pisang Kayu’ ‘Kluai Hom’, ‘Pisang Sri’, ‘Kluai Hom Thong Mokho’ ‘Pisang Jambe’, ‘I.C.2’, ‘1877’ ‘Safet Velchi’, ‘Lal Kelat’ ‘Figue Pomme Ekona’ ‘Amou’, ‘French Sombre’, ‘French Clair’, ‘Kelong Mekintu’, ‘Orishele’, ‘Msinyore’ ‘Prata Anã’, ‘Foconah’, ‘Nadan’, ‘Guindy’ ‘Iho U Maoli’, ‘Popoulou’, ‘Poingo’ ‘Mattui’, ‘Maritú’ ‘Figue Pomme’, ‘Muthia’, ‘Supari’ ‘Pisang Ceylan’, ‘Goroio’, ‘Zabi’ ‘Pisang Rajah Bulu, ‘Nendrapadathi’ ‘Pisang Pulut’, ‘Pisang Kelat’ ‘Laknao’

PR HR S

Gros Michel Cavendish

Red Ibota Bota Orotava Others

AAB

PR ‘Kluai Tani’, ‘Pisang Klutuk Wulung’, ‘Pisang Batu’

Lujugira–Mutika

AAAA AB

HR

Ney Poovan Other Plantain Pome Mai’a Maoli-Popoulu Iholena Silk Mysore Pisang Raja Pisang Kelat Laknau

PR

HR S S S S HR S PR PR PR S S S S PR S and PR PR S S and PR S Continued

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Chapter 2

Table 2.8. Continued. Genomic group ABB

Subgroup/cloned

Cultivars evaluated

Reactiona

Bluggoe Pisang Awak subgroup Pelipita Peyan Kluai Teparot Ney Mannan Saba Inabaniko

‘Cacambou’, ‘Cachaco’, ‘Poteau’, ‘Monthan’ ‘Fougamou’, ‘Gia Hong’, ‘Foulah 4’, ‘Brazza 2’ ‘Pelipita’ ‘Simili Radjah’ ‘Kluai Tiparot’ ‘Pisang Abu Perak’, ‘Ice Cream’, ‘Som’ ‘Saba’ ‘Benedetta’

PR PR* and PR PR PR PR* PR PR PR

Classified using Cavendish cultivars as very susceptible reference clones. S, very susceptible; PR, partial resistance; PR*, very strong partial resistance (Fouré’s compatible interactions); HR, high resistance (Fouré’s incompatible interaction). b AF, plant derived from the self-fertilization of a Musa acuminata ssp. banksii accession. c Hybrid with dominant characteristics of Musa acuminata subspecies. d Name of best-known synonym signifies clone. e This cultivar has succumbed to black leaf streak in Cameroon (Mouliom-Pefoura, 1999), Costa Rica (Escobar-Tovar et al., 2015a) and Puerto Rico (Irish et al., 2013).

Compatible interaction

Incompatible interaction

a

AAw Musa acuminata ssp. Musa acuminata ssp. ‘Truncata’) Musa acuminata ssp. Musa acuminata ssp. Musa acuminata ssp.

AA ‘Paka’, ‘Tuu Gia’ ‘Pisang Lilin’

banksii microcarpa (‘Pisang Cici Alas’, siamea (‘Kluai Pa’) malaccensis (‘Pahang’) burmannica (‘Calcutta 4’)

AAw Musa acuminata ssp. zebrina (‘Zebrina’) AA Sucrier (‘Pisang Mas’) Several (‘Pisang Berlin’,...) AAA AAAA Orotava ‘I.C.2’ (‘Pisang Sri’) Ambon (‘Pisang Ambon’)

AAA Ibota (‘Yangambi Km 5’, ‘Khom’)

HR

BBw Musa balbisiana (‘Pisang Klutuk Wulung’, ‘Tani’)

PR

AAw Musa acuminata ssp. banksii (‘Madang’) AA Several (‘Chicame’,...) AAA Gros Michel (‘Highgate’) Cavendish (‘Grande Naine’,...) Red (‘Figue Rose’,..) Oratava (‘Pisang Kayu’) Lujugira-Mutika

S AAB Plantain (‘French Sombre’,...) Pome (‘Prata Ana’) Silk (‘Muthia’) Maia'a Maolil-Popoulu (‘Popoulou’) Pisang Raja; Laknau AABB (‘Ngoen’)

AB Ney Poovan (‘Safet Velchi’,..) ABB Bluggoe (‘Cachaco’,..) Ney Mannan (‘Som’,..) Pisang Awak (‘Fougamou’) Pelipita (‘Pelipita’) Peyan (‘Peyan’) (‘Kluai Teparot’) Saba (‘Saba’) Monthan (‘Monthan’,...) AAB AAAB Mysore (‘Pisang Ceylan’) ‘Goldfinger’ Silk (‘Figue Pomme’) Iholena (‘Maritu’) Pisang Kelat (‘Pisang Kelat’)

Fig. 2.9.  Classification of wild Musa species/subspecies, banana subgroups and banana clones, according to their reaction to black leaf streak disease under field conditions at Njombé, Cameroon (80 m altitude, 1800 mm annual rainfall). Key: HR or phenotype 1 response = high resistance; PR or phenotype 2 response = partial resistance; S or phenotype 3 response = susceptible; AAw = wild Musa acuminata subspecies; BBw – wild Musa balbasiana; AA, AAA, AAAA, AB, ABB, AAAB, AABB = genomic subgroups of cultivated banana (the names of popular cultivars in subgroups appear in parentheses) (adapted from information presented in a table that appeared in Fouré et al., 1999, by C Abadie, CIRAD, France).



Fungal Diseases of the Foliage

resistant or incompatible reaction (phenotype 1) is ­expressed in most wild M. acuminata subspecies and in some cultivars within the AA and AAA groups. Hybrids bred for resistance to black leaf streak (Craenen and Ortiz, 1998;  Abadie et  al., 2009) or by genetic engineering (Vishnevetsky et al., 2011) have also been evaluated for disease response. Further evaluations to confirm the resistance of hybrids in new environments have been conducted (Cohan et  al., 2003; Noupadja et al., 2007; Irish et al., 2013). Host–pathogen interactions introduction. 

Studies on Musa–P. fijiensis interactions began with research on the microscopic events that take place in banana tissues after fungal penetration and culminate in the expression of resistance or susceptibility (Beveraggi et  al., 1995). In the past few decades, the development of phenotyping and molecular tools plus major breakthroughs in genomics has allowed a greater understanding of the molecular ‘dialogue’ that occurs between plants and pathogens. It has also brought us closer towards ­identifying the key genes that will allow durable resistance. A significant advance that will greatly assist endeavours to increase the amount of available knowledge on Musa–P. fijiensis interactions has been the publication of the genome sequences of M. acuminata and P.  fijiensis (D’Hont et  al., 2012; Arango-Isaza et al., 2016).

histology of the interaction. 

Balint-Kurti et al. (2001) developed GFP-expressing P. fijiensis transformants that allowed a more detailed and rapid microscopic analyses of the host–­parasite interaction. Cavalcante et al., (2011) used a transformant strain to compare the interactions between susceptible ‘Grande Naine’ (AAA, Cavendish subgroup), partially resistant ‘Pisang Madu’ (AA) and highly resistant M. acuminata ssp. burmanicca (AAw, accession ‘Calcutta 4’). Their observations complemented ultrastructural observations with a transmission electron microscopy undertaken by Beveraggi et al. (1995) that compared the reactions of susceptible ‘Grande Naine’ with partially resistant ‘Fougamou’ (ABB, Pisang Awak subgroup) and highly resistant ‘Yangambi km5’ (AAA, Ibota subgroup).

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Germination and stomatal penetration.  After spore germination and variable epiphyllous growth of mycelium, P. fijiensis forms stomatopodia (hyphal swellings) above stomata on the underside of banana leaves. Thin hyphal filaments then penetrate through the stomata and grow in a circular pattern in the substomatal cavity. Mycelia later spread and branch through the intercellular spaces between mesophyll cells.

Lesion development.  The first major event after penetration is necrosis of the stomatal cells. In compatible interactions, the necrosis slowly spreads to nearby cells while other necrotic areas develop some distance away from the ­ ­initial micro-lesion. These clusters of dead lower ­hypodermal or spongy mesophyll cells are ­separated from the original micro-lesion by apparently healthy tissues. As the central lesions spread, the secondary necrotic areas increase in diameter and coalesce. Other necrotic areas regularly appear ahead of the margin of the main lesion. After the initial penetration of the fungus, epiphyllous mycelia can infect other nearby ­stomata, leading to a succession of similar re­ actions. This phenomenon accelerates the colonization rate and leaf tissue at the centre of the expanding spot dries. When the inoculum is highly concentrated, lesions caused by penetrating hyphae originating from different conidia quickly coalesce and large areas of tissue rapidly become necrotic, as is often also noted in the field. In adult banana leaves infected with P.  musae, secondary veins form a mechanical barrier to the lateral spread of the fungus within the leaf (McGahan and Fulton, 1965). The pathogen can overcome these barriers by growing epiphytically on the leaf surface, but progress is not as rapid as in leaf tissue parallel to the veins and the lesion tends to become linear in shape. In young leaves, such as those found on small water suckers, the secondary veins are less developed and the lateral growth of the pathogen is not so limited. As a consequence, necrotic lesions become more elliptical. Pseudocercospora fijiensis invades banana leaf tissue in a similar manner to P. musae. Young plants derived from tissue culture have limited vein development and black leaf streak lesions tend to be more spherical.

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Growth through banana tissue.  The course of infection has been followed in several susceptible and resistant banana cultivars inoculated with P. fijiensis (Beveraggi, 1992; ­Balint-Kurti et al., 2001; Cavalcante et al., 2011). In compatible interactions, hyphae are observed between living cells ahead of the necrotic zone, indicating that the pathogen is biotrophic during the initial phases of infection. The main ­difference between susceptible and partially resistant cultivars is the much faster growth rate of hyphae in the former. In incompatible interactions, as exemplified by the infection of the highly resistant cultivar ‘Yangambi Km 5’ (AAA, Ibota subgroup), the pathogen stops growing 2–3 days after stomatal penetration and the death of the first plant cell.

Compatible interactions.  The progress of infections in ‘Fougamou’ (ABB Pisang Awak subgroup), a cultivar with strong partial resistance, and ‘Grande Naine’ (AAA, Cavendish subgroup), a susceptible cultivar, were compared by Beveraggi et al. (1993, 1995). In ‘Fougamou’, a histological analysis of healthy tissues revealed the presence of many specialized parenchyma cells containing polyphenol-­rich vacuoles in the palisade and spongy mesophyll layers. Far fewer of these specialized cells were found in uninfected ‘Grande Naine’. An ultrastructural study of the interaction between P. fijiensis and ‘Fougamou’ revealed that, once the specialized cells became necrotic, the contents of their vacuoles were released into the intercellular spaces. This material had a high affinity for fungal cell walls and their appearance around hyphae seemed to be correlated with the slow spread of the mycelium in parenchyma tissues. Incompatible interactions.  Most of the information on incompatible reactions has been obtained from the studies of interactions involving ‘Yangambi Km 5’ (AAA) (Beveraggi et al., 1993, 1995) or Musa acuminata ssp. burmannica (AAw, accession ‘Calcutta 4’) (Cavalcante et al., 2011). The first event that occurs following penetration is the very rapid death of about three to four cells around the stoma and the expulsion of electron-dense compounds through the walls of these cells into the intercellular space. The released compounds show a high a ­ ffinity for the

walls of the infection hypha and nearby host cells. The degeneration of hyphae in contact with these compounds suggests that they have a fungitoxic action. Deposits of these compounds on host cell walls could also explain the fluorescence observed at infection sites after analine-blue staining. The ability of ‘Yangambi Km 5’ cells nearest the pathogen to die rapidly seems to be a characteristic hypersensitive reaction. In this case, it appears that host resistance is determined by changes that occur in only a few cells either in contact with or in close proximity to the pathogen. A second event, which is characterized by further, more extensive cell death in surrounding leaf tissue, follows the hypersensitive response. Cavalcante et  al. (2011) found that limited necrosis at infection sites in leaves of ‘Calcutta 4’ was associated with cells with thickened walls, peroxidase activity and H2O2 production. The authors concluded that no reaction was observed early enough to suggest that it played an important role in the defence mechanism associated with the hypersensitive response. Hypersensitive reactions typically take place in gene-for-gene relationships. There is no experimental evidence of such relationships in Musa– P. fijiensis interactions, as this requires genetic studies of the host and pathogen. As mentioned previously, some P. fijiensis isolates originating from the Cook Islands and Tonga were able to overcome the resistance of ‘Yangambi Km 5’ in laboratory tests (Fullerton and Olsen, 1993), thus suggesting that this type of host–pathogen relationship could occur. molecular determinants of pathogenicity. 

The appearance of chlorotic haloes around lesions caused by P. fijiensis led to speculation that a toxin produced by the pathogen may be killing leaf tissue in advanced of its invasion by hyphae (Molina and Krauz, 1989). This led to research that identified toxins from culture filtrates (Molina and Krauz, 1989; Upadhyay et al., 1990 a, b; Stierle et al., 1991; Strobel et al., 1993) with the hope that one could be used to mass screen tissuecultured banana germplasm for resistance or susceptibility to black leaf streak. The toxic secondary metabolites identified were fijiensin, 2,4,8-trihydroxytetralone (2,4,8-THT), juglone, 4-hydroxyscytalone, 3-carboxy-3-hydroxycinnamic acid and isoochracinic acid (Churchill, 2011).



Fungal Diseases of the Foliage

Studies have concentrated on juglone, which is very phytotoxic, and 2,4,8-THT, which is produced in high concentration. Juglone was found in extremely low concentrations in vitro and to have no host selectivity by Strobel et  al. (1993), who considered 2,4,8-THT likely to be the more suitable for use in screening germplasm for reaction to black leaf streak. However, Busogoro et  al. (2004a, b) found that chloroplasts were the target of juglone and those of the partially resistant ‘Fougamou’ (AAB Plantain subgroup) were less a ­ ffected by the toxin than those of the very susceptible ‘Grande Naine’ (AAA, Cavendish subgroup). Later, El Hadrami et al. (2005) discovered that, after treatment with juglone, an early release of active oxygen species and a quick stimulation of an antioxidant system was observed in ‘Fougamou’, as compared with ‘Grande Naine’. Hoss et  al. (2000) discovered that 2,4,8THT was produced early in interactions with ­ resistant cultivars, causing the formation of necrotic micro-lesions and eliciting defence mechanisms that led to incompatibility. In contrast, sub-lethal levels of 2,4,8-THT were produced during the early phase of host–pathogen interaction with susceptible cultivars that allowed hyphae to spread in the intercellular leaf space. Increased levels of 2,4,8-THT later in the interaction then caused extensive necrosis. Both juglone (Lepoivre et  al., 2003) and 2,4,8-THT (Okole and Schultz, 1997) have been used to screen in vitro banana cell suspensions and somatic embryos or micro-cross-sections (from which plantlets can be initiated) for tolerance to the toxins in the hope of selecting material that also has resistance to black leaf streak. Toxin-tolerant plants were generated with increased resistance to P. fijiensis in growth ­ ­chambers (Okole and Schulz, 1993), but results of field tests, which would confirm resistance, have not been published. In addition, these toxins have not been shown to be primary determinants of pathogenicity. Churchill (2011) called for the strategy of selecting resistant banana clones using P. fijiensis toxins to be discontinued until there is definitive proof of mode of action of the toxins. Stierle et  al. (1991) noted that 2,4,8-THT, juglone and 4-hydroxyscytalone are shunt metabolites of fungal melanin biosynthetic pathways.

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Beltrán-García et  al. (2014) showed that amounts of melanin in naturally infected leaf tissue could be correlated with disease expression. The authors suggested that the melanin produced by P. fijiensis generates oxygen that may function as a ‘photoactivated toxin’ triggering cell death. They concluded that further studies were needed to determine the relationship between melanin-mediated reactive oxygen species and P. fijiensis pathogenesis. Later work with a melanin-deficient mutant of P. fijiensis, which was reported by Churchill (2011), found that it was as virulent as the wild-type strain on susceptible ‘Grande Naine’ plants. This suggested that melanin is not required for P. fijiensis virulence on banana and that melanin shunt metabolites do not play a key role in black leaf streak disease (Churchill, 2011). Until 2011, the majority of secondary metabolites identified in P. fijiensis culture filtrates had been lipophilic phytotoxins with a light-­ dependent phytotoxicity. Cruz-Cruz et al. (2011) developed a method to purify hydrophilic phytotoxins, which were shown to be non-host-selective and were not light-dependent. These toxins ­acted by altering the permeability of the plasma membrane through the production of reactive oxygen species. From their results, the authors suggested that a hydrophilic phytotoxin metabolite might be involved in pathogenesis. Chuc-Uc et  al. (2011) inoculated banana leaves with proteins secreted in vitro by P. fijiensis and produced necrotic lesions. The secretome produced cell death on resistant and susceptible banana leaves, but lesions appeared earlier and were larger on the resistant M. balbisiana compared with the susceptible ‘Grande Naine’. Escobar-­ Tovar et  al. (2015a) compared in vitro and in planta secretomes produced by two isolates of P. fijiensis; one virulent on ‘Yangambi Km 5’ (AAA, Ibota subgroup) and the other avirulent. The more prevalent secretomes were proteins involved in carbohydrate transport and metabolism. One unknown protein was considered to be a virulence factor or effector candidate. Portal et al. (2011) first looked for P. fijiensis genes involved in a compatible interaction. They identified a gene induced at a late stage (37 days after infection) that encoded UDP-­ glucose pyrophosphorylase, an enzyme involved in the biosynthesis of trehalose. The authors believed that the trehalose pathway is

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likely to have been induced in response to plant antifungal genes and may act as a protectant against membrane damage. Kantún-Moreno et  al. (2013) investigated the glycosyl phosphatidylinositol-protein (GPI) family in the cell wall of P. fijiensis. They identified in silico 49 genes encoding putative GPI proteins with an N-terminal secretory signal peptide that could play a role in pathogenicity. The expression of a β-1,3-glucanosyltransferase (Gas) known as MfGas2 was found to increase during the biotrophic growth phase after infection to reach a maximum concentration at the speck symptom stage. This was followed by a progressive decrease in concentration as necrotic symptoms progressed. The authors believed that MfGas2 could be involved in the remodelling of the pathogen’s cell wall which avoided early recognition by the plant host. Couoh-Uicab et  al. (2012) looked at the role that the ATP-binding cassette (ABC) protein family that drives the transport of compounds across biological membranes might play in P. fijiensis–Musa interactions. In the closely related wheat pathogen Zymoseptoria tritici (syn. Mycosphaerella graminicola), seven ABC transporters designated MgAtr1 to MgAtr7 have been identified and a role in pathogenicity has been attributed to MgAtr4. In infected banana leaves, the expression of MfAtr4, the P. fijiensis putative ortholog of MgAtr4, was found to be higher during the first symptomatic stages in comparison with the late necrotrophic phases. The authors are currently conducting research to analyse the role of MfAtr4 in virulence and its probable role in detoxification of toxic defence compounds produced by banana. Noar and Daub (2016a) conducted an in  silico analysis of genes encoding P. fijiensis’s polyketides toxins. Among the eight polyketide synthase (PKS) gene clusters identified, three were strongly upregulated during disease development in ‘Grande Naine’. This led the authors to propose that they may encode polyketides that are important for pathogenicity. A list of P. fijiensis genes with high expression in infected leaf tissue compared with that during saprophytic growth in culture medium has been identified by Noar and Daub (2016b). Those genes including ones commonly involved in secondary metabolism mainly group in 16 gene clusters. The authors also identified two putative dispensable scaffolds in the P. fijiensis

genome with a large proportion of putative pathogenicity genes. Stergiopoulos et  al. (2010) identified homologs of the Cladosporium fulvum Avr4 and Ecp2 effectors in P. fijiensis called MfAvr4 and MfEcp2, respectively. They found that the MfAvr4 and MfEcp2 proteins produced by P. fijiensis could be recognized respectively by the Cf-4 and the Cf-Ecp2 resistance proteins in tomato. They suggested that banana genetically modified to incorporate the two tomato genes coding for the resistance proteins may be resistant to P. fijiensis. As disease resistance controlled by single dominant genes can be overcome by pathogen mutations, the authors surveyed the global population of the pathogen for allelic variation. Their results indicated that some alleles able to overcome the tomato resistance genes were already present in natural populations of the pathogen. Thus, the durability of field resistance provided by the genes in transformed banana, particular in Southeast Asia where a high diversity of P. fijiensis alleles are present, was questioned (Stergiopoulos et al., 2014). molecular determinants of resistance. 

Secondary plant metabolites with antimicrobial activity (phytoprotectants) can be present in the plant before infection (phytoanticipins) and produced as a response to infection (phytoalexins). Both phytoanticipins and phytoalexins have been found in banana. Cruz Cruz et al. (2010) purified a phytoanticipin with a saponin structure that was shown to have antifungal activity against P. fijiensis from an aqueous infusion from the susceptible cultivar ‘Grande Naine’ (AAA, Cavendish subgroup). However, only infusions from 4-monthold plants had significant antifungal activity. Phenylphenalenones are known to act as phytoalexins in banana. Hidalgo et  al. (2009) suggested that the phenylphenalenones act as a catalyst for the production of other reactive oxygen species, which may cause cell damage to the pathogen. Four phenylphenalenones and two perinaphthenone-type compounds were isolated and identified from rhizomes of the P. fijiensis-­ resistant cultivar ‘Yangambi Km 5’ (AAA, Ibota subgroup). The two perinaphthenone types showed significantly more activity against P. fijiensis than the phenylphenalenones (Otalvaro et al., 2007). Hidalgo et al. (2016) analysed the chemical responses of susceptible ‘Williams’ (AAA,



Fungal Diseases of the Foliage

Cavendish subgroup) and resistant ‘Kluai Khai Thong Ruang’ (AAA, Ibota subgroup) to P. fijiensis and showed that phenylphenalenone-type compounds constituted the majority of induced metabolites. ‘Williams’ produced a smaller number and a lower level of these metabolites than did ‘Kluai Khai Thong Ruang’. However, the authors demonstrated that a virulent strain of P. fijiensis was able to overcome plant resistance by converting phenylphenalenones to sulfate conjugates. Torres et  al. (2012) investigated enzyme ­activity in susceptible ‘Willliams’ (AAA, Cavendish subgroup) and highly resistant M. acuminata ssp. burmannica (AAw, accession ‘Calcutta 4’) for 18 days following infection by P. fijiensis. The ­activities of enzymes involved in plant defence such as peroxidase, phenylalanine ammonia lyase and β-1,3-glucanase were not induced until 6 days in ‘Williams’ compared with the first 6–18 h in ‘Yangambi Km 5’ with H2O2 being produced at 72 h. This suggested to the authors that the first 72 h are important in determining the response of the host to the disease. genes for resistance.  The search for resistance gene candidates initially concentrated on looking for recurrent DNA sequences in the banana genome. Several resistance gene analogs (RGA) have been identified (Pei et  al., 2007; Azhar and Heslop-Harrison, 2008; Miller et  al., 2008; Peraza-Echeverria et  al., 2008; Baurens et al., 2010). The largest known family of plant resistance genes encodes proteins with nucleotide-­ binding site (NBS) and C-terminal leucine-­rich repeat (LRR) domains. Working with M. acuminata ssp. malaccensis (AAw) and AAB cultivars, Lu et  al. (2011) isolated four RGAs of this type with sequences that were close to those of known resistance genes from other plant species. D’Hont et  al. (2012) screened the genome for all the sequences having homology with NBS ­domains. Their number was relatively low compared with other plant species. To date, none of the banana RGA genes so far identified has been shown to be a functional resistance gene. Portal et al. (2011) and Passos et al. (2012) have made expressed sequence tag (EST) databases of transcripts expressed or induced during compatible and incompatible interactions at different stages of infection available for further work. In a compatible interaction with ‘Grande Naine’ (AAA, Cavendish subgroup), Portal et  al. (2011) identified several banana

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genes thought to be involved in the synthesis of phenylpropanoids and detoxification compounds, as well as pathogenesis-related proteins that could be involved in the response against P. fijiensis infection. At late stages of infection, jasmonic acid and ethylene signalling transduction pathways appeared to be active. Passos et al. (2012), studying a compatible interaction with ‘Grande Naine’ and an incompatible interaction with M. acuminata ssp. burmannica (AAw, accession ‘Calcutta 4’), identified a number of expressed genes potentially involved in different pathways triggering immunity. Transcriptomes analysis in P. fijiensis-­ inoculated and uninoculated susceptible, partially resistant and highly resistant banana accessions led to the identification of several active genes (D’Hont et  al., 2012). Receptor-like protein kinase genes, genes encoding WRKY transcription factors and genes coding for PR-1, germin-like proteins, hevein-like proteins, chitinases and glutathione-S-transferases were found upregulated in the susceptible (AA, ‘Pisang Pipit’) and partially resistant (AA, ‘Pisang Madu’) interaction, but not in a highly resistant interaction. This suggested that a basal defence-oriented reprogramming was induced in the susceptible and partially resistant interactions. In the case of highly resistant M. acuminata ssp. malaccensis (AAw, accession DH-Pahang), no differential expression was detected for these gene categories, possibly due to a very early, rapid resistance ­reaction. This study was conducted with the ­detached leaf bioassay and these results remain to be confirmed on inoculated whole plants. Recently Rodriguez et al. (2016) and Timm et  al. (2016) studied early interactions with a susceptible and the highly resistant M. acuminata ssp. burmannica accession ‘Calcutta 4’ (AAw). Candidate reference genes that either constitutively expressed or strongly induced defence-­ related genes were identified in early stages of infection. Those genes were either differentially expressed compared with the susceptible cultivar or only expressed in the highly resistant cultivar. Some genes were further tested by real-time PCR after selection of housekeeping internal reference genes for expression studies under experimental Musa–P. fijiensis interaction conditions. Rodriguez et al. (2016) confirmed the early role played by some defence-related genes coding for peroxidase, phenylalanine ammonia-lyase and cinnamate hydroxylase in resistant ‘Calcutta 4’ compared with susceptible ‘Williams’ as previously

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described at the chemical level by Torres et  al. (2012) and Hidalgo et al. (2016). These results suggest that recognition within the first 72 h after inoculation is critical to the host’s ability to limit pathogen invasion. Most of the genes found regulated in those studies have been reported as components of early defence responses in other plants, such as Arabidopsis thaliana, or in banana in response to other pathogens. For black leaf streak resistance as for other characters, few genetic analyses have been undertaken on banana due to poor fertility, heterozygosity and polyploidy of many accessions. Ortiz and Vuylsteke (1994) and Craenen and Ortiz (1997) analysed the genetics of resistance on diploid and tetraploid progenies from crosses between two susceptible plantain cultivars (AAB) and the highly resistant M. acuminata ssp. burmanicca (AAw, accession ‘Calcutta 4’). They distinguished three levels of host response among hybrids based on youngest leaf spotted (YLS) readings: susceptible, less susceptible and partially resistant. Segregation ratios fitted a genetic model having a major locus with a recessive allele (bs1) and at least two independent minor loci with additive effects (bsr1 and bsr2). Nothing is known on partial resistance. Analysis of more segregating populations is needed to progress our knowledge on the genetic control of this interaction. Better genetic analyses are now possible with the increasing characterization of banana genetic resources (Perrier et al., 2011), the use of new sequencing tools for the discovery, validation and assessment of genetic markers in populations and knowledge of the banana reference genome sequence (D’Hont et al., 2012; Martin et al., 2016). The identification of quantative trait loci and genes can be attempted not only from segregating populations, but also from natural populations by classical or association genetics (Sardos et al., 2016). Work on related species or similar types of interactions will also provide useful information (Passos et  al., 2013; Rudd, 2015; Rudd et al., 2015; Yang et al., 2015). Partial or quantitative resistance is generally considered more durable than high or major gene-based resistance. However, the erosion of both types of resistance has been observed (Mundt, 2014). The partial loss of the original strong resistance found in some FHIA hybrids is strongly suspected to have been caused by a gene or genes contributing to quantitative resistance being overcome (Pérez-Miranda et  al., 2006;

A. Cavalier, L. Pérez-Vicente, D. Rengifo, L. Miniere, T. Lescot and C. Abadie, 2016, unpublished ­results). Considering the high evolutionary potential of P. fijiensis, more information on the host–pathogen interaction is required before strategies for long-term resistance can be ­devised.

Control M. Guzmán, L. Pérez-Vicente, R.A. Romero and D.H. Marín Introduction The commercial production of bananas of the Cavendish subgroup requires a rigorous black leaf streak disease management programme to meet the market requirements of quality. This is achieved mainly through a combination of cultural practices and chemical methods. Biological control options have shown potential in glasshouse and small field trials, but, as yet, have not been routinely adopted in commercial operations. Fungicides are mainly used because non-chemical alternatives alone do not provide satisfactory control and there are no resistant cultivars with the green life and taste that is required by the export market. Some hybrids with resistance to black leaf streak have been conventionally bred, but their fruit has only been acceptable on local markets and as staple foods. Today, different approaches to control, some suitable for small or large operations as well as for local or export markets, are being utilized in an integrated system of disease management. These include the adoption of useful cultural practices, disease monitoring, disease forecasting systems, spray programmes that use fungicides with different modes of action, cultivars with partial resistance if fruit is acceptable at local markets and biological control methods (Marín et  al., 2003; Ganry et  al., 2011; Pérez-­ Vicente, 2012a; Guzmán et al., 2013). Large-scale standardization of management practices Management practices are better if they are organized for large areas. This is not only possible on large export plantations, but also when neighbouring growers on smaller plots agree to pool resources and implement identical control



Fungal Diseases of the Foliage

s­trategies. As ascospores of P. fijiensis can be transported long distances by wind, it is desirable that growers and farms of a specific area of production follow the same fungicide application plan to prevent a build-up of resistance (Ganry et al., 2011; Pérez-Vicente, 2012a). The application of treatments is also more efficient coming from a professional technical service following rational guidelines rather than from individual growers.

Cultural control

M. Guzmán, D.H. Marín, L. Pérez-­Vicente and R.A. Romero sanitation.  The reduction of inoculum levels by the early removal of banana leaves with extensive necrosis caused by coalescing spots and also the excision of small areas of necrotic leaf tissue helps in the management of the disease

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(Plate 2.9). The practice of removing necrotic leaf material is commonly known as ‘leaf pruning’, ‘de-leafing’, ‘de-tipping’, or ‘surgery’, depending on the tissue excised (Marín et al., 2003; Etebu and Young-Harry, 2011). It is carried out by a specialized worker called a ‘de-leafer’ or ‘leaf pruner’. Necrotic tissue cut from the plant lies on the ground, where it decomposes (Plate 2.9). If the necrotic tissue is left on the plant, production of pseudothecia continues and ascospores can be released for up to 22 weeks. Leaves on the ground produce a reduced amount of inoculum with the period of ascospore release reduced by 6 weeks or less (Gauhl, 1994; Pérez-­ Vicente, 1998; Romero, 1999; Guzmán et  al., 2005b). In addition, the spores released from leaf tissue in the ground have a reduced likelihood of reaching the young susceptible leaves on the upper part of the plant. Frequency and intensity of sanitation is another important factor. Necrotic leaf tissue with sporulating lesions is usually removed

Plate 2.9.  Reduction of inoculum levels of Pseudocercospora fijiensis by the removal of sporulating leaf tissue. The practice is known as ‘de-leafing’, ‘de-tipping’ or ‘surgery’ depending on the amount of tissue excised (left). Diseased leaf tissue lying on the ground is a source of inoculum (upper right). Application of solutions of urea, acetic acid or specific microorganisms can accelerate leaf decomposition and reduce sporulation (lower right).

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weekly or bi-weekly as soon as spot symptoms mature (stages 5 or 6 of Fouré, 1987). In a high rainfall location of Nigeria, Emebiri and Obiefuna (1992) evaluated the effect of leaf removal and/or intercropping with cassava on the incidence and severity of black leaf streak on young ‘Agbagba’ (AAB, Plantain subgroup). They found that the removal of older leaves alone was not effective in reducing incidence and severity of black leaf streak. Disease incidence was reduced by 16% and severity by 10% when plants had only their top four to six leaves. The best results were obtained when this severe leaf removal was combined with intercropping with cassava, which reduced disease incidence by 23% and severity by 25%. Engwali et al. (2013) evaluated different frequencies of leaf pruning (3, 6, 9 and 12 days) on ‘Grande Naine’ (AAA, Cavendish subgroup) in Cameroon and obtained the best results, in terms of disease severity reduction, production and monetary yield, with a 6-day pruning cycle. In a commercial plantation of a Cavendish cultivar in Costa Rica, Calvo and Bolaños (2001)

investigated the effect of three weekly de-leafing strategies. The first was the detailed excision of all necrotic tissue, the second was the pruning of leaves with 16–33% of leaf area affected, and the third was the pruning of leaves with > 50% of affected area. With the first and second strategies, pruning costs increased up to three times, but resulted in a reduction in severity of black leaf streak and numbers of bunches rejected because of ‘creamy pulp’ (see Plate 2.2). Fruit with creamy pulp look normal, but have started to ripen as a result of black leaf streak infection and the risk of further ripening in transit is very high. The third strategy was found to be insufficient for proper disease management even with the application of fungicide. This pruning option is more appropriate for plantain, which is a little more resistant to black leaf streak than Cavendish cultivars and fruit quality requirements for local markets are not so demanding (M. Guzmán, Costa Rica, 2016, personal communication). Other work has also shown some control of the disease by pruning (Fig. 2.10) and more intensive strategies of de-leafing have prevented

60

700

55

600 500

45

400

40 300

35

Rainfall (mm)

Infection index (%)

50

200

30

100

25

0

20 18

20

22

24

26

28

30

32

Week of the year (2004) Cumulative rainfall

No sanitation

Non-intensive (mild sanitation)

Intensive-detailed sanitation

Fig. 2.10.  Effect of three levels of sanitation on black leaf streak severity on actively growing plants of ‘Grande Naine’ (AAA, Cavendish subgroup) in fungicide-treated plots at Matina in Costa Rica. Key: No sanitation = no leaves or part of leaves were removed from plants; Non-intensive (mild sanitation) = weekly pruning of highly infected leaves or the half of these; Intensive-detailed sanitation = weekly elimination of alll infected leaf portion (‘de-leafing’, ‘de-tipping’ and ‘surgery’) (R. Villalta and M. Guzmán, Costa Rica, 2004, unpublished results).



Fungal Diseases of the Foliage

the premature ripening of fruit (Guzmán et al., 2005c; Chillet et al., 2013). Variation of sanitation practices on plants in the vegetative phase of growth was reported in Colombia by Martínez-­ Acosta et al. (2006). This comprised the removal of the distal end of the third and fifth youngest leaves before symptoms develop and spores were produced. This practice, called ‘early pruning’ or ‘early cut-off ’, contributed to the reduction in severity of black leaf streak. Thus, the removal of diseased necrotic tissue must be considered a key component of integrated management of the disease (Marín et al., 2003; Chillet et al., 2013). de-leafing at flower emergence.  A new practice of removing leaf tissue that could potentially be a source of inoculum before its production has been initiated has been implemented in banana plantations of Costa Rica and elsewhere. The practice is known as ‘controlled defoliation’ or ‘early pruning of leaves’ at flower emergence (Vargas et al., 2008, 2009). It has been demonstrated that ‘controlled defoliation’, which leaves the plant with seven to eight functional leaves at flower emergence, does not affect yield, plant growth, fruit quality or ‘green and yellow life’ (Ramírez et al., 2008). This is because the practice eliminates leaves that contribute little to the overall photosynthesis of the plant (Vargas et al., 2008, 2009). In Costa Rica, it is now recommended that the three oldest leaves are removed in the first week after flower emergence, which leaves the plant with nine to ten functional leaves (Plate 2.10). Continuous use of the practice for more than 3 years in a 300 ha plantation growing a Cavendish cultivar improved disease control and reduced the amount of mineral oil needed in complementary chemical control measures (Guzmán, 2012). ‘Controlled defoliation’ has been successfully implemented on more than 3000 ha of banana plantations in Costa Rica (G. Murillo and M. Guzmán, Costa Rica, 2016, personal communication). treatment of residues of de-leafing. 

Reduction of the sources of inoculum found in leaf residues in the plantation has been also addressed in black leaf streak management programmes. The application of a 10% solution of urea (Table 2.9) and specific microorganisms to pruned leaf tissue lying on the ground can reduce inoculum levels and accelerate tissue ­

91

­ ecomposition (Plate 2.9) (Guzmán et al., 2005b; d Guzmán and Villalta, 2005, 2006; Orozco-­ Santos et al., 2013a, b; Gutiérrez-Mosalve et al., 2015). A 5–10% solution of acetic acid was reported as a strong inhibitor of P. fijiensis sporulation when applied to necrotic tissue with mature lesions. A single application reduced the number of ascospores released in spot lesions by 84–90% for at least 9 days (Gómez, 2013; Guzmán et al., 2014). Other practices, such as placing pruned leaves together in heaps, also enhance inoculum depletion (Guzmán et  al., 2005b; Villalta and Guzmán, 2005). The implementation of this practice varies with the planting systems (single or double row, hexagonal system or programmed crops) (Plate 2.11). As part of an integrated strategy, pruned leaf heaps are treated with compounds or microorganism to enhance decomposition as discussed above (Orozco-Santos et  al., 2008; Guzmán et  al., 2013). A more detailed knowledge of the conditions affecting sporulation of the pathogen in detached leaves and the discovery of compounds or substances able to block this process may be a key factor in the development of more efficient management strategies of the disease. management of the crop environment.  The reduction of relative humidity within the crop has proved an important component of black leaf streak disease management. An efficient drainage system, which rapidly removes superficial and underground water (Plate 2.12), and an appropriate plant density both facilitate drying and help reduce the humidity and duration of leaf wetness after overhead irrigation (Marín et  al., 2003). The foliage of plants should not overlap (Plate 2.13). Overlapping leaves increase humidity levels and also make it difficult to cover all leaf surfaces with fungicide when spraying. In banana-producing regions where irrigation is required, under-canopy or drip irrigation systems are preferred to an overhead system, which covers leaf surfaces with water and thus encourages spore production, release, dissemination and germination (Wielemaker, 1990; Orozco-­ Santos et al., 2008, 2013a; Guzmán et al., 2013). Periodic weed control, either chemically or mechanically, is also important in reducing levels of humidity and black leaf streak. The use of a short-statured soil cover crop in plantations

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Plate 2.10.  Pruning of leaves soon after flower emergence to reduce Pseudocercospora fijiensis inoculum. Banana plant before pruning (upper left); banana plant after pruning - arrows show where leaves were removed (upper right); lower surface of newly pruned leaf showing developing lesions (lower left); and senescing leaf after pruning showing lesion development is arrested and thus inoculum considerably reduced (lower right) (photos: M. Guzmán, CORBANA).

has been found not to influence black leaf streak levels, but it does help in soil conservation and improves crop production (Guzmán and Villalta, 2007). nutrition.  Improving general plant health and growth through an adequate nutrient supply is another important part of the integrated approach for the management of black leaf streak. Good soil fertility is necessary for rapid plant

growth and a fast leaf emergence rate. Plants growing in fertile soils generally show lower disease severity than plants in poorer soils. This is because high rates of leaf emergence mean that there is always enough photosynthetic tissue to counter the serious effects of necrosis caused by infection (Pérez-Vicente, 1997, 1998, 2006). Deficiencies in potassium and magnesium, imbalances between potassium and magnesium and imbalances between nitrogen and potassium



Fungal Diseases of the Foliage

93

Table 2.9.  Effect of an application of a solution of urea (10 %) to diseased leaf tissue on the amount of ascospores of Pseudocercospora fijiensis released by a lesion (stage 6 of Fouré 1987) from the upper leaf surfaces of ‘Grande Naine’ (AAA, Cavendish subgroup) in plantations with and without chemical control in three different experiments in Costa Rica. Discharged ascospores were counted in nine random ­microscope optical fields readings at 40x in the discharge area on the surface of a water-agar medium discharge area above lesions. The average sum of the numbers of ascospores counted above 30 lesions for each treatment are the values seen in the table (R. Villalta and M. Guzmán, Costa Rica, 2004, unpublished results). Plantation with chemical control Leaf surface

Experiment 1 May–June

Urea No Urea P > Fa

21 72 0.0001

Urea No Urea P > Fa

10 19 0.0675

Experiment 2 June–July 30 70 0.0374 Plantation without chemical control 31 83 0.0097

Experiment 3 August–September 32 234 < 0.0001

36 121 0.0009

T-test (P = 0.05).

a

Plate 2.11.  Crop production and planting systems that can help to reduce the amount of Pseudocercospora fijiensis inoculum produced by infected leaf tissue lying on the ground. Single or double row planting with excised leaves in piles between the rows (upper left); hexagon planting system with excised leaves in the centre of the hexagon (upper right); programmed harvest system where all plants are cut down after bunch removal and a sucker is selected for the next crop (lower left); and traditional cropping system where harvested plants are left standing and diseased leaves cover the ground thus increasing levels of inoculum (lower right) (photos: M. Guzmán, CORBANA).

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Plate 2.12.  A malfunctioning drainage system, which increases humidity levels under the leaf canopy and encourages black leaf streak development and infection (left); and a well-maintained drainage system allows little water to accumulate, lowers the humidity between plants and lessens the effects of black leaf streak (right).

in poor soils predispose plants to infection (Roperos and Magnaye, 1991; Méndez, 1998; Guzmán et  al., 2013). Banana plants are more prone to succumb to black leaf streak in soils deficient in potassium or with unbalanced nitrogen/potassium relationship (Holderness et al., 1999; Pérez-­ Vicente et  al., 2002a). Studies in Costa Rica showed a relationship between the annual nitrogen supply and the severity of black leaf streak (Méndez, 1998). Plants fertilized with less than 100 kg nitrogen/ha/year exhibited twice the severity of black leaf streak compared with plants that received between 200 and 500 kg/ha/year. In the same study, an inverse relationship between potassium application and black leaf streak severity was observed. Plants receiving less than 150 kg/ha/year were affected more severely by the disease than those fertilized with rates between 150 and 750 kg/ha/year (Méndez, 1998). In Cuba, it was found that severity of black leaf streak in a plantation of partially resistant ‘FHIA-18’ (AAAB, bred Pome-type hybrid) was negatively correlated with the potassium content in soil and leaves (Pérez-Vicente et al., 2003).

Proper management of organic matter is not only essential for perennial productivity, but it also reduces the amount of damage caused by black leaf streak. Composted farmyard manure, crop residues and mulches improve the organic matter and nutrient status of the soil, thereby reducing disease severity and resulting in higher yields (Etebu and Young-Harry, 2011). Working with two plantain cultivars in Nigeria, Aba et al. (2011) found that the application of poultry manure at the rate of 10 t/ha/year increased tolerance to black leaf streak and improved productivity. Leachings and suspensions of Californian worm (Eisenia foetida) compost are rich in humic and fulvic acids, mineral nutrients and other organic compounds plus a myriad of microorganisms. Foliar applications have been shown to increase plant endurance and response to black leaf streak infection with an efficacy similar to that obtained with applications of mancozeb (Pérez-Vicente, 2006). Similar results have been reported on the use of leachings of banana bunch rachises in Colombia (Alvarez et al., 2013).



Fungal Diseases of the Foliage

95

Plate 2.13.  Plants of ‘Grande Naine’ (AAA, Cavendish subgroup) growing at high density in Costa Rica (left). Plant vigour is reduced and overlapping leaves (upper right) prevent adequate fungicide coverage, resulting in typical infection patterns parallel to the central leaf vein and at the leaf base (lower right) (photos: M. Guzmán, CORBANA).

High foliar levels of potassium, calcium, boron and iron were associated with low disease severity, whereas high foliar levels of magnesium correlated with high disease severity in Colombia (Aguirre et al., 2015). Foliar applications of some elements such as silicon, copper, calcium, boron and zinc may contribute to reduce disease severity (Azofeifa et  al., 2007; Guzmán, 2012; Kablan et  al., 2012). Experiments in banana have shown reductions in black leaf streak severity of up to 15% when these elements were applied to leaves in a mixture with fungicides (Azofeifa et  al., 2010). The compound containing the element is another important consideration. For example, better results were obtained using zinc oxide than using zinc sulfate or zinc nitrate (Guzmán, 2012; Guzmán and Villalta, 2014). Performance has also been increased when secondary or minor elements are foliar a ­ ­pplied, which is

preferential to soil application (­Azofeifa et  al., 2007; Martínez et al., 2007). Chemical control

R.A. Romero, L. Pérez-Vicente and M. Guzmán introduction. 

The chemical control of leaf spot diseases in Central America began with the use of Bordeaux mixture (suspension of copper sulfate, hydrated lime and water) against Sigatoka leaf spot disease in the mid-1930s. Bordeaux was superseded by the use of mineral oil alone and dithiocarbamate fungicides. The conversion from ground spraying to aerial spraying was made possible by the change to oil. In the 1950s and early 1960s, both oil and dithiocarbamates were used together in mixtures (Guyot and Cuillé, 1954a, b, 1955, 1956). The first systemic

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fungicide used in the control of Sigatoka leaf spot was benomyl, which was in widespread use by the early 1970s. Benomyl was also found to be effective against black leaf streak after the appearance of the disease in Central America in 1972. Ten years later, tridemorph was used against black leaf streak and, in 1984, propiconazole, the first of the triazole fungicides, was introduced. Table 2.10 summarizes the evolution of the use of fungicides in banana for the control of Sigatoka leaf spot diseases. Selection of the fungicide to be used to control black leaf streak must be based on a knowledge of several key factors. These include the cropping system, the epidemiology of the disease, the sensitivity of the local pathogen population to fungicides, the management of the farm and a thorough understanding of how all

these factors interact with each other. Thus, the chemical control of black leaf streak is part of an integrated disease management strategy, which involves good sanitation practices, adequate drainage systems, ideal plant densities and optimal nutritional conditions for plant growth. fungicide application. 

Aircraft and sometimes helicopters are used to spray fungicides in most export banana plantations, but tractors or motorized backpack sprayers are used on small farms growing fruit for local markets or where aerial spraying is not permitted. However, ground sprays are less efficient than aerial sprays. In most operations in Central America, the Ayres Trush 600, Ayres Turbo Trush and Air Tractor 502 are the most widely used aircraft (G. Murillo, Costa Rica, 2016, personal communication).

Table 2.10.  Approximate dates of use of fungicides and the development of strategies for the control of Sigatoka leaf spot diseases of banana in Central America (modified from Stover, 1990 and Guzmán et al., 2013). Compounds and mixtures

Date

Disease

Bordeaux mixtures, copper and dithiocarbamates Petroleum oil Dithiocarbamates in petroleum oil–water emulsions Benzimidazoles in straight oil or in oil–water emulsion ‘Cocktails’ of benzimidazoles plus dithiocarbamates in oil–water emulsions Chlorothalonil Flowable formulations of dithiocarbamates Tridemorph in oil or in oil–water emulsion Triazoles in oil or in an oil–water emulsion Mixtures of triazoles and tridemorph in an oil–water emulsion Concentrated suspensions of dithiocarbamates formulated with petroleum oil Triple mixture of triazole, tridemorph and dithiocarbamates in an oil–water emulsion to cope with fungicide resistance build-up in the population of the pathogen Several new triazoles approved and introduced (see Table 2.11) Azoxystrobin, the first strobilurin fungicide, was introduced for commercial use in bananas Acibenzolar-S-methyl, a promoter of systemic acquired resistance is approved for use in bananas First biological fungicide based on Bacillus subtilis Pyrimethanil introduction Tea tree oil extract Boscalid, the first SDHI fungicide approved for banana Dodine, a guanidine compound Second biological fungicide based on Bacillus pumilus Diethofencarb first used

1936–1962 1956–present 1956–present 1972–1995 1977–1991

Sigatoka leaf spot

1978–present 1980–present 1981–present 1984–present 1989–present 1990–present 1994–present

1996–2009 1997–present 2001–present 2002–present 2004–present 2008–present 2012–present 2013–present 2014–present 2016–present

Black leaf streak



Fungal Diseases of the Foliage

These aircraft are equipped with turboprop motors, which allow for the fast application of fungicides during brief windows of spraying opportunities, and satellite-linked geographical positioning systems, which direct the pilot to application targets and are visible on an electronic screen inside the aircraft. This technology avoids unnecessary spray overlaps, missed paths and overspray. The number of hectares that can be sprayed by an aircraft in 1 h depends on the configuration of the farms and the distance from the mixing and loading facility. However, it is usually in the range of 90–150 ha. alternative methods of fungicide application. 

Soil treatment, injection of pseudostems and rhizomes plus deposition in the axil of the leaf have been tested as alternatives methods to fungicide applications on leaves with promising results. In the late 1970s and the 1980s, the first studies of soil applications of systemic fungicides for the control of Sigatoka leaf spot and black leaf streak were made in Taiwan, Côte d’Ivoire, Gabon, Costa Rica and Cuba, with triadimefon and its derivative triadimenol, which belong to the triazole group (Mourichon, 1982; Wybou et al., 1987; Pérez and Mauri, 1994). Triadimenol is more active and systemic than triadimefon and has been proven to be superior at doses of 0.5–1 g a.i./plant. According to these studies, one soil application was equivalent to 3–6 foliar applications with a residual effect of 2.5–4.0 months (Wybou et al., 1987; Pérez and Mauri, 1994). The treatment fully protects leaves before emergence and so its efficacy is very high. In Cuba, a granulated formulation at 0.75–1 g a.i./plant has been applied for more than 10 years in commercial Cavendish banana fields where spraying is not possible. In areas of Cuba where ‘FHIA-18’(AAAB, bred Pome-type hybrid) lost its partial resistance to black leaf streak, applications of 0.5 g triadimenol/plant at 3–4 months before flowering efficiently controlled the disease until harvest without the detection of fungicide residues in fruit (Pérez-­Vicente et al., 2009, unpublished results). Other studies have also confirmed the efficacy of triadimenol applied to the soil for the control of black leaf streak (Moulion-Pefoura and Fouré, 1988; Orozco-­Santos, 1998; Gasparotto et  al., 2005). A granulated formulation of flutriafol, another ­triazole fungicide, reduced disease severity and

97

r­ emained active in leaves for 75 days after application to soil at the base of plants (Gasparotto et al., 2005). In other experiments, results have been variable since efficacy can be affected by soil type, the health of the root system and soil moisture. Selection of active ingredients for use in soil treatments should be based on the intrinsic fungicide activity against P. fijiensis, a low octanol-­ to-water coefficient partition and properties related to adsorption to soil colloids and water leakage. The application of systemic fungicides to soil may increase the probability of P. fijiensis developing resistance and could disturb the balance of soil and root microorganisms (Pérez and Mauri, 1994). Fungicide sensitivity monitoring of P. fijiensis populations in Cavendish plantations in Cuba after soil treatments with triadimenol has shown a loss of resistance to the fungicide by a factor of 77 compared with wild-type populations (L. Pérez-Vicente, Cuba, 2008, personal communication). Injection or implantation of fungicides in the rhizome or pseudostem of the plant has also been evaluated. In Australia, Allen (1991) implanted triadimenol (0.2–2.0 g a.i./plant) directly into the rhizomes of old ‘Williams’ (AAA, Cavendish subgroup) plants and observed arrested development of Sigatoka and speckle leaf spots, but only on leaves that had already unfurled at the time of treatment. The level of control was reported to be not as good as the commercial practice of spraying the leaves with an oil–propiconazole mixture. In Brazil, flutriafol injected into the pseudostem of ‘Prata Anã’ (AAB, Pome subgroup) at the rate of 0.25 g a.i./ plant every 30 days resulted in good control of black leaf steak. Plants had more than ten functional leaves until 75 days after the last application. However, plants were predisposed to bend after the emission of the bunch. These symptoms were less severe if there were longer intervals between applications (Gasparotto et  al., 2005, 2006). Gasparotto et  al. (2004, 2005) obtained good control of black leaf streak in ‘Prata Anã’ (AAB, Pome subgroup), ‘Maçã’ (AAB, Silk subgroup) and ‘D’Angola’ (AAB, Plantain subgroup) after the application of 0.25 g a.i./plant of flutriafol or azoxystrobin every 60 days in the axil of second fully unfurled leaf counting down from the top of the plant. The fungicide applications

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began when the plants were 4 months old and continued until flowering. Applications then commenced on the suckers selected for the ratoon crop and so on (Gasparotto and Rezende, 2013). The use of systemic fungicides to the soil, the pseudostem or axil of the leaf could become control alternatives for small banana and plantain growers. In commercial banana plantations, this system could help protect areas where aerial spraying is difficult or impossible. However, more studies are necessary regarding the impact of these methods of application on the development of resistance to fungicides in the pathogen and the risk of residues being translocated into fruit. fungicides for black leaf streak control. 

Fungicides listed for black leaf streak control on ­banana are presented in Table 2.11. They are divided into protectants, often called contact fungicides, and systemics, also known as curatives. Protectants remain on the surface of the leaf and do not penetrate underlying tissues. They have a broad spectrum of activity and exert a multi-site effect on the pathogen, mostly reacting with essential thiol groups (SH-groups) of enzymes, causing a non-selective toxicity (Arauz, 1998; Agrios, 2004; Gasztonyi and Lyr, 1995). Dithiocarbamates and related fungicides available for use on banana are different formulations of mancozeb, propineb, metiram and thiram. Mancozeb, the most popular compound of the group, is a complex of zinc and maneb (zinc and manganese salts of ethylene bisdithiocarbamate) and belongs more specifically to the class of compounds known as ethylene bisdithiocarbamates (EBDCs). It is classified by the Fungicide Resistance Action Committee (FRAC) in modeof-action group M (multi site action). Mancozeb itself is not fungicidal and can effectively be considered a profungicide which, when exposed to water, breaks down to release ethylene bisisothiocyanate sulfide (EBIS), which is then converted by the action of UV light into ethylene bisisothiocyanate (EBI). Both EBIS and EBI are believed to be the active toxicants and are thought to interfere with enzymes containing sulfydryl groups. This disruption of core enzymatic processes is postulated to inhibit or interfere with at least six different biochemical processes within the fungal cell cytoplasm and mitochondria (Gullino

et  al., 2010). Some formulations of mancozeb can be used in oil–water emulsion while others can be applied only in water. Chlorothalonil belongs to the chloronitrile group of protectant fungicides. It has a low water solubility, which favours its fungitoxic activity and reduces the risk of phytotoxicity. This compound has to be sprayed in water alone, as the interaction of oil and chlorothalonil causes damage to leaf tissues. Care must be taken when oil has been used in previous treatment and when applying oil later to chlorothalonil-treated surfaces. Inorganic fungicides, such as those based on sulfur and copper, can also be applied to control black leaf streak, but they are not often used commercially, because of the availability of more efficient fungicides. Systemic fungicides have the ability to penetrate the leaf through the cuticle and translocate in xylem in the evapotranspiration stream. They accumulate in leaf sites of high stomata density and exert a toxic effect on the pathogen after infection. Most systemic fungicides have a specific mode of action, which makes them non-phytotoxic to banana. The benzimidazole group of systemic fungicides comprises several compounds, of which benomyl and thiophanate-methyl have been the most widely used in Central America and elsewhere. Benzimidazoles act in preventing microtubule (spindle fibre) assembly, which is necessary for cell division during mitosis and meiosis. They bind to the β-tubulin, a subunit of the ­tubulin protein, which is a component of the microtubules (Davidse, 1988; Davidse and Ishii, 1995; Delp, 1995). Diethofencarb is a carbamate fungicide that has been recently approved for use on black leaf streak. It belongs to the N-phenylcarbamates group of fungicides, which has been shown to have negative cross-resistance with benzimidazoles. It has been found to give good control even where local populations of P. fijiensis have a high resistance to benzimidazoles (M. Guzmán, Costa Rica, 2015, unpublished results). Exposure of pathogens to two fungicides that exhibit this negative cross-resistance should greatly reduce any resistance risk associated with either component, as a shift to resistance against one automatically confers sensitivity against the other (Brent and Hollomon, 1998). However, this



Fungal Diseases of the Foliage

99

Table 2.11.  Fungicides approved on banana for the control of black leaf streak.a Type

Chemical group

Common name

Mode of action

Protectant

Dithiocarbamates and relatives

mancozeb propineb metiram thiram chlorothalonil

Multi-site contact activity

Chloronitriles (phthalonitriles) Inorganic

Systemic

Bacillus sp. and the fungicidal lipopeptides Bacillus sp. and the fungicidal amino sugars produced Terpene ­hydrocarbons and terpene alcohols Benzimidazoles Thiophanates N-phenyl carbamates Morpholines Spiroketalamines Piperidines Triazols

Strobilurin

sulfur copper Bacillus subtilis strain QST713 Bacillus pumilus QST2808

Lipid synthesis and membrane integrity Inhibition of cell wall formation

Doseb (g a.i./ha)

Resistance risk

1050–1500  700–1400 980 840–1680 540–1260

None to low

1600 – 6.7–13.4

6.9–13.8

tea tree oil extract from Melaleuca ­alternifolia carbendazim tiophanatemethyl diethofencarb

Cell ­membrane disruption (proposed) Cytoskeleton and motor proteins: ß-tubulin assembly in mitosis

140 196

tridemorphc fenpropimorph spiroxamine

Sterol biosynthesis inhibitors (Code Group 2, FRACd)

450 444–880 320

fenpropidin propiconazole difenoconazole epoxiconazole tebuconazole tebuconazole + triadimenol hexaconazole azoxistrobin trifloxistrobin pyraclostrobin

Pyridine-­ carboxamide

boscalid

Anilino-pyrimidine

pyrimethanil

Guanidines

dodine

Sterol biosynthesis inhibitors (Code Group 1)

Inhibit respiration (Quinone outside inhibitors, QoI) Inhibit respiration (Succinate Dehydrogenase Inhibitor; SDHI) Amino acids and protein synthesis Unknown: cell membrane disruption (proposed)

Unknown

89.2–178.4

High

150

450 100 100 100 100–125 120 100 100 75–100 100 150

300–420

400

Low to medium

Medium

High

Medium to high

Medium

Low to medium

Continued

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Chapter 2

Table 2.11. Continued. Type

Chemical group

Systemic Benzo-thiadiazole acquired (BTH) resistance activator

Common name

Mode of action

acibenzolar-Smethyl

Host plant defence induction

Doseb (g a.i./ha) 40

Resistance risk Not known

This list of fungicides refers to compounds with registrations in USA, Canada or Europe for use on banana and does not include registration in the country of use. b Dose based on the label of the main trademarks. c Tridemorph´s maximum residue levels (MRL) have been dropped to the default value in Europe since 2015. d FRAC: Fungicide Resistance Action Committee. References: Marín et al., 2003; Guzmán et al., 2013; FRAC, 2016 a, b; A. Segura (Bayer Crop Science, Costa Rica, 2016, personal communication). a

f­ ungicide should be used with caution as there is a risk of selecting isolates coupling benzimidazole resistance, and insensitivity toward diethofencarb, as was reported by Faretra et  al. (1989) and Leroux and Moncomble (1994) in Botrytis cinerea. Isolates with the mutation at codon 200, characterized by a single substitution of phenylalanine to tyrosine, appeared with ­resistance to both classes of fungicides (FRAC, 2014a). Resistance to benzimidazole fungicides in populations of P. fijiensis in Honduras and the Philippines occurred 2–3 years after the fungicide was first used (Stover, 1980a). Resistance has also been reported in Western Samoa (Fullerton and Tracey, 1984), in many other countries in Central America (FRAC, 1993; Romero and Sutton, 1998), Colombia (Cañas-Gutiérez et  al., 2006) and Mexico (Aguilar-Barragan et al., 2014). The build-up of resistance has been attributed to the intensive use of benzimidazoles for black leaf streak control when these fungicides first became available. Only a single mutation in the β-tubulin gene in Venturia inaequalis and other fungal pathogens results in resistance to benzimidazoles (Koenraadt et al., 1992). It is likely that the same mutation occurs in P. fijiensis. Sequence analysis of the β-tubulin gene in Colombian isolates of P. fijiensis that appeared to have moderately and highly resistant phenotypes revealed that all had the single base substitution from GAG (coding for glutamine) to GCG (coding for alanine) at codon 198 (CañasGutiérez et  al., 2006). This same amino acid change has been described in several other plant pathogenic fungi (Koenraadt et al., 1992; Luck and Gillins, 1995).

Fenpropimorph and tridemorph are members of the morpholine (amines) group of systemic fungicides. Both compounds can be applied in oil alone or in an oil–water emulsion. They inhibit the synthesis of ergosterol, which is an important component of the cell membrane in fungi, and are grouped in the sterol biosynthesis inhibitors of group II or amines. They interfere mainly with sterol Δ8–7 isomerase and Δ14–15 reductase activity (Köller, 1992; Kerkenaar, 1995). Even though tridemorph was introduced since 1981 (see Table 2.10) and has been used extensively in Central America and other countries, there are no reports of field resistance in P. fijiensis populations (Martínez-Bolañoz et al., 2012; FRAC, 2014b). Other sterol biosynthesis inhibitors of group II (amines) currently approved for the control of black leaf streak are spiroxamine (spiro-ketalamine chemical group) and fenpropidin (piperidine chemical group). There are several compounds in the triazole group of systemic fungicides registered for use on banana. All of them inhibit a cytochrome P-450 mono-oxygenase enzyme, which catalyses the C-14 demethylation reaction in the ergosterol biosynthesis pathway (Köller, 1992). Because of their action, these fungicides are known as demethylation inhibitors (DMI) or ergosterol biosynthesis inhibitors of group I. Fungicides in this group have active ingredients that differ markedly in biological properties with regards to interactions with the pathogen and movement in the plant. In 1987, propiconazole was the first DMI fungicide registered for use on banana in Central America. Due to its high efficacy against P. fijiensis, fewer applications of this fungicide were



Fungal Diseases of the Foliage

needed to achieve satisfactory standards of control. Other triazoles registered for use on banana are listed in Table 2.11. There are differences in the efficacy of these compounds (Guzmán and Romero, 1996a) and in their systemic movement inside the leaf. An indication of the amount of translocation is given by leaf infection patterns. Differences in infection patterns became evident in the field after the build-up to resistance to triazoles in the pathogen population. Propiconazole translocates very rapidly from the midrib towards the edge of the leaf and this results in a pattern of infection along the midrib where fungicide concentrations are sub-­lethal to resistant populations. Fenbuconazole and bitertanol application are less rapidly translocated and this results in significantly less ­disease along the midrib than at the leaf edge (M.  Guzmán and R.A. Romero, Costa Rica, 1995, unpublished ­results). Triazoles can be applied in an oil–water emulsion or in oil alone. Loss of sensitivity and resistance to propiconazole has been reported in  many countries in Central America (FRAC, 1993; Romero and Sutton, 1997a), Colombia (Cañas-Gutiérez et al., 2009), Mexico (Martínez-­ Bolañoz et  al., 2012; Aguilar-Barragan et  al., 2014) and Caribbean Islands (Pérez-Vicente et  al., 2001). Other DMI fungicides have been shown to be less effective against propiconazole-­ resistant populations of P. fijiensis, indicating cross-resistance (M. Guzmán and R.A. Romero, Costa Rica, 1996, unpublished results). Strobilurin fungicides are naturally occurring compounds found in various genera of fungi belonging to the Agaricaceae, which live on decaying plant material in forests (Lange et  al., 1993). The natural products themselves have physical properties that make them unsuitable for use in agriculture, but analogues have been developed with improved stability and physical properties (Table 2.11). These compounds inhibit fungal respiration by binding strongly to a specific site on the outside of the cytochrome b molecule, which prevents electron transfer between cytochromes b and c1 (Clough et al., 1994). These fungicides belong to the QoI inhibitor group. Azoxystrobin (methoxy-acrylate), trifloxystrobin (oxymino-acetate) and pyraclostrobin (methoxy-­ carbamate) have been used to control black leaf streak (Pérez-Vicente et  al., 2002b). Resistance to QoI fungicides has been related to changes in

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amino acids in different positions in the complex bc1 (Gisi et al., 2002). Two main amino acid substitutions have been detected in the cytochrome b gene in resistant plant pathogens. One is the mutation G143A (a change from glycine to alanine at position 143) and the second is the mutation F129L (a change from phenylalanine to leucine at position 129) (Sierotzki et al., 2000 a, b; Heaney et  al., 2000; Kuck and Mehl, 2003). Both G143A and F129L are based on single ­nucleotide polymorphisms in the cytochrome b  gene and the selection process is qualitative (­single step). Based on current knowledge, resistance factors (RF = ED50* [resistant strain] / ED50 [sensitive wild-type strain]) associated with G143A and F129L are different. Resistance factors caused by F129L usually range between 10 and 50, whilst resistance factors related to G143A are in most cases greater than 100. G143A has been shown to be responsible for QoI resistance in more pathogen species than F129L. Isolates carrying G143A express high (complete) resistance. Isolates with F129L express moderate (partial) resistance. ­Differences in ­mutation sites could potentially be the cause of the recently reported lack of cross-­resistance ­between azoxystrobin and pyraclostrobin for P. fijiensis populations in Colombia (Sepúlveda-­Ramos and Torres-Bonilla, 2016). More recently, another compound that inhibits fungal respiration was introduced for black leaf streak control. Boscalid acts on a different site than do strobilurin fungicides. It inhibits succinate dehydrogenase (SDHI). Pyrimethanil (anilinopyrimidine) and dodine (guanidine) are also products available for use on banana. Pyrimethanil has a different mode of action, interfering with the secretion of enzymes from the pathogen that are necessary for pathogenesis (Milling and Daniells, 1996), while guanidines appear to disrupt cell membranes. Acybenzolar-S-methyl, a compound in the benzothiadiazole (BTH) group that induces a systemically acquired resistance (Görlach et al., 1996), is currently approved for use and has shown high efficacy in the field even under conditions of high disease pressure and resistance to systemic fungicides (Madrigal et  al., 1998; Guzmán, 2002). However, its large-scale use remains very limited, due to some observed side effects on plant physiology. More recent studies with low doses of the product have shown a

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r­ eduction in deleterious side effects and satisfactory results in controlling the disease (Guzmán et  al., 2013; M. Guzmán, 2015, unpublished ­results). mineral oil.  The use of mineral oil for the control of Sigatoka leaf spot was pioneered in the French Antilles (Guyot, 1953; Guyot and Cuillé, 1954a, b, 1955, 1956) and quickly adopted elsewhere. It was later found to be effective against black leaf streak. It works by stopping disease development in the early stages of infection, delaying the appearance of symptoms and retarding the expansion of lesions (Stover, 1990; Guzmán and Romero, 1996b). Mineral oil is fungistatic rather than fungicidal when applied to leaves with advanced disease symptoms (Pérez et al., 1981). Oil helps in the control of Sigatoka leaf spots in three ways. Firstly, because it reduces the evaporation of small drops, it allows for a better coverage of leaves of fungicides in low volume applications. Secondly, because it dissolves wax threads on the leaf surface, it allows fungicides to better contact and penetrate the leaf ­cuticle. Thirdly, depending on the rate of application, it inhibits the development of early black leaf streak symptoms from streaks to spots (Pérez-Vicente, 2012a). Oil did not affect the growth of P. musae in vitro and it has been suggested that it acts by somehow increasing the resistance of banana (Meredith, 1970). Experimental work has shown that the therapeutic effect of oil is translocated to unsprayed leaves, indicating that it may act by stimulating the banana’s natural ­defence mechanisms (Guzmán and Romero, 1996b). Today, all systemic fungicides are applied in combination with oil. Oil helps systemic fungicides penetrate leaves, thus optimizing their performance. In situations where the pathogen is highly sensitive to the fungicide and few applications are necessary for the control of disease, oil can be used on its own. Application rates range from 5 l/ha to 15 l/ha (Marín and Romero, 1992), but 10 l/ha has been recommended for commercial plantations in Central America (Guzmán and Romero, 1996b). In locations where fungicide resistance is a problem and more frequent fungicide applications are required for adequate control, the amount of oil used in sprays is reduced. This is because oil can accumulate on leaves and negatively affect yield

by interfering with the exchange of gases and photosynthesis (Israeli et al., 1993). In the final analysis, it is disease pressure that dictates the amount of oil used. The serious consequences of poor disease control in commercial plantations far outweigh yield losses that might be expected from a build-up of oil. The chemical composition of mineral oil, which consists of aromatic or paraffinic fractions, has an important influence on phytoxicity and disease control. Paraffinic oils have the higher activity as pesticides and it is desirable that they form the higher component (> 60%) in the final product. The presence of double unsaturated links in the aromatic oils make them more chemically reactive, which increases the risk of phytotoxicity and effects on human health (Jacques and Kuhlmann, 2002). Oils should have an unsulfonated residue of 90% or more and an aromatic content of less than 12% (Stover and Simmonds, 1987; Israeli et  al., 1993). Those fractions distilling below 338°C at 760 mm Hg do not control black leaf streak or Sigatoka leaf spot. Fractions distilling above 365°C control the diseases, but are more phytotoxic. Oils that combine both good disease control qualities and low phytotoxicity must have a 50% distillation range of 346–354°C (Calpouzos, 1968). Highly refined paraffinic oils are available today. Chemical control strategies fungicide application based on a disease forecasting system. 

The climatic conditions at a specific location have a significant impact on black leaf streak incidence and severity and will ultimately determine the number of fungicide treatments necessary to achieve adequate control. In the tropics, rainfall patterns are the main factors that determine disease pressure. Figure 2.11 shows the relationship between rainfall and the mean number of fungicide treatments carried out yearly (Pérez-Vicente, 2013). The regular monitoring of climatic factors is essential in order for appropriate decisions to be made concerning disease control. Climatic information obtained through the regular monitoring of local weather conditions coupled with direct observations of symptom development on plants in the field form the basis of methods used to forecast strategic fungicide applications.



Fungal Diseases of the Foliage

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70

66

Number of fungicide treatments

60

56

50

48

40

39

35

55

46

39

42 41

35 35

30

32

26

20

17 9

10

17 14

0

0 0

1000 2000 3000 Cumulative annual rainfall (mm)

4000

Fig. 2.11.  Relationship between annual rainfall and number of fungicide treatments (both systemic and protectant) required for the control of black leaf streak disease. Data obtained from the country reports of Belize, Cameroon, Colombia, Costa Rica, Cuba and Mexico (presented by L. Pérez-Vicente at a meeting held in Costa Rica in 2007).

The methods for assessing disease severity and the rate of disease development, which are used for monitoring disease levels useful in forecasting systems, were initially developed for Sigatoka leaf spot. Ganry and Meyer (1972b) recorded the position of the leaf with the most advanced stage of the symptoms (without taking into account numbers of lesions) in the five youngest leaves of the plant and the leaf emission rate. This information was used to determine the speed of disease development. Ten non-flowered plants in the field were monitored weekly. The method was widely used in Cameroon and Caribbean Islands together with Piche evaporation in simplified shelters for forecasting the appropriate times to spray fungicides (Ganry and Meyer, 1972a, b; Ganry and Laville, 1983). A simplification of this system was used in Windward Islands and Jamaica (Cronshaw, 1982). Ternisien (1985) adapted the system for use with black leaf streak by taking the various stages of disease evolution into consideration and also including a recording of symptom intensity. The data obtained for each leaf was ­adjusted by taking into account its age and a correction factor for differences in the stage of unfurling of new leaves included. Finally, the

combined data was corrected by the rate of leaf emission to obtain a variable called the ‘stage of evolution of the disease’ (Ganry and Laville, 1983; Marín and Romero, 1992). If the disease was sufficiently advanced, the crop was sprayed with fungicide. Rainfall and leaf wetness periods lasting for 10–14 days correlate significantly with the speed of disease development 4 and 5 weeks later on Cavendish and plantain cultivars (Bureau, 1990; Lescot et  al., 1998; Pérez-Vicente et  al., 2000a, b). In Cuba, a forecasting system based on biological and climatic data under integrated management control reduced fungicide treatments by 35–44% during 1990 and 1994. After the appearance of black leaf streak disease in the Windward Islands and French Antilles, this type of forecasting warning system has also been adopted. The method, with some modifications and simplifications, has been used in Cameroon (Fouré, 1990a) and Central America (Corrales and Marín, 1992; Marín and Romero, 1992). Forecasting systems depend on the ability of systemic fungicides applied in an oil suspension to arrest infections at early stages of development (Marín and Romero, 1992). The early

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warning system allows fungicides to be used only when necessary, thus reducing the cost of control of the disease. The method was used with different levels of success against Sigatoka in the French Antilles and black leaf streak in Cameroon. However, results in Central America have been disappointing. This has been attributed to the rapid build-up of resistance to benomyl and propiconazole fungicides in local populations of P. fijiensis. A loss of sensitivity significantly affected the efficacy of these fungicides to arrest early infections (Romero and Sutton, 1997a). Resistance to fungicides in the pathogen population developed faster in Central America than in Cameroon. Probably, the climate is more favourable for disease development in Central America and this allows the pathogen a greater number of sexual reproductive cycles/season than in Cameroon. Also, farms in Central America are larger and in a much greater concentration in a given geographical area. The size of populations of P. fijiensis being exposed to fungicides is also much larger. All these factors are conducive to the build-up of resistance and may explain why a loss of sensitivity took longer in Cameroon. Forecasting systems for black leaf streak on Cavendish cultivars have also been proposed by Chuang and Jeger (1987a, b) and Wielemaker (1990). The former is based on disease severity, total precipitation and days with over 90% humidity and the latter solely on weekly observations of disease symptoms on young leaves. Bureau (1990) and Lescot et  al. (1998) developed forecasting systems for black leaf streak on plantain. fungicide application based on crop growth and weather conditions.  In Costa Rica, because of the build-up and persistence of resistance to the fungicide benomyl, which has been withdrawn from control programmes, and an increasing resistance to propiconazole (Romero and Sutton, 1997a, 1998) and strobilurins, systemic fungicides alone are no longer able to provide satisfactory control of black leaf streak. A similar situation regarding resistance to these fungicides has arisen in most other countries in Central America and in Cameroon. The use of protectant fungicides is now necessary, both in alternation and in mixtures with systemic fungicides, to maintain adequate control of the disease. To achieve satisfactory control, protectant fungicides need to be sprayed at intervals that

ensure that unfurling and recently expanded leaves are covered. Therefore, during weather conditions favourable for infection, protectant fungicides are applied at intervals close to the rate of leaf emission. Emission rates vary according to the temperature, the nutritional condition of the crop and soil type. Although this strategy of application of protectants in alternation or in mixtures with systemic fungicides is referred to within the industry as a calendar-­ based schedule, in reality many factors are considered when making a decision on the time and type of fungicide treatment. These factors include a weekly analysis of disease data on plants of different ages on every farm, the rate of leaf emission, weather data on a weekly basis and the sensitivity of local populations of the pathogen to site-specific fungicides. fungicide resistance management.  As a consequence of the build-up of resistance to propiconazole, more applications of the fungicide are needed to achieve the levels of disease control required for commercial plantations. The efficacy of other DMI fungicides has also decreased significantly because of cross-resistance factors, but these too are also still used to control black leaf streak. Populations of P. fijiensis with resistance to strobilurins developed within a few years after their introduction in Central America (Sierotzki et al., 2000b; Chin et al., 2001), provoking the withdrawal or a significant limitation of the number of cycles of strobilurins in control programmes in many locations. Because of the important economic consequences of resistance to systemic fungicides, the sensitivities of local pathogen populations to the different groups of systemic fungicides are established and regularly monitored by the industry, so that any changes of sensitivity can be detected early. This is undertaken in conjunction with a fungicide application strategy. The most commonly recommended strategies to prevent or delay the development of resistance to fungicides in P. fijiensis are as follows.

1. Limit the use of fungicides with high risk of resistance development such as benzimidazoles, strobilurins and carboxamides. 2. Use mixtures of systemic fungicides with a protectant, such as mancozeb. 3. Avoid the use of DMI, benzimidazole, strobilurin or carboxamide fungicides in four consecutive



Fungal Diseases of the Foliage

months during a season or year. They should be substituted with protectant or amine fungicides during these periods. 4. If only systemic fungicides are used to control the disease, systemics with different modes of action should be sprayed alternately. 5. Use a rate of oil close to 10 l/ha to enhance its activity and also that of any systemic fungicide applied with the oil. 6. Do not use reduced rates of systemics. 7. Alternate protectant and systemic fungicides. 8. Maintain all cultural practices that reduce inoculum levels. Van den Bosch et  al. (2014) reviewed the experimental and modelling evidence on the use of mixtures of fungicides of differing modes of action as a resistance management tactic. They concluded that adding a different fungicide to one that is in danger of losing its sensitivity (without lowering the dose of the at-risk fungicide) reduces the rate of selection for fungicide resistance. This holds for the use of mixing fungicides that have either multi-site or single-site modes of action. The benefits of the use of DMI fungicides in mixtures with protectants have been demonstrated in Costa Rica (Guzmán, 2007). In experiments, the infection index of black leaf streak was found to be significantly lower after treatment with propiconazole and mancozeb in oil than after a treatment with propiconazole in oil alone. Earlier work indicated that the effect of each fungicide is additive (M.  Guzmán and R.A. Romero, Costa Rica, 1996, unpublished results). The frequency of fungicide-resistant strains in the pathogen population is likely to remain the same after a mixture with mancozeb is applied. The way in which a mixture helps in the prevention and management of resistance to DMI fungicides is thought to be by reducing the size of the population of the pathogen. This results in fewer individuals submitted to selective pressure and less sexual recombination during a season, thus retarding the flow of resistance genes within the population (Van den Bosch et al., 2014). Although a DMI application-free period is advocated as an anti-resistance strategy, the value of this recommendation has yet to be demonstrated in Costa Rica (Jiménez et  al., 1996; Guzmán et  al., 1997; Guzmán, 2007), where

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high populations of the fungus have resistance. After DMI application-free periods of up to 6 months for 2 years, there has been no change in the sensitivity of the pathogen population. This could be because resistance to DMI fungicides does not affect the fitness of P. fijiensis. Certainly, the ability of resistant strains to produce conidia in culture is not affected (Jiménez et al., 1997). However, a DMI application-free period is believed to be of great value in delaying the onset of resistance in those areas where DMI resistance is not present or where the pathogen population is composed of only a few resistant individuals. Environmental impact of chemical control and its mitigation

D.H. Marín, L. Pérez-Vicente and M. Guzmán The control of black leaf streak is becoming ­increasingly more difficult with the number of chemical applications/year needed to protect banana plantations increasing over time. In ­ ­Costa Rica at the beginning of the 1990s, fungicides were applied about 30 times/year. Now, 50 or more applications/year are needed. In Cameroon, 12–14 treatments of fungicides were ­applied/year and now 40–50 applications are required (de Lapeyre de Bellaire et  al., 2010). These changes have occurred because forecasting systems are being abandoned as resistance to systemic fungicide has increased, with control more and more relying on the use of protectant fungicides that must be applied weekly or more often (Marín et al., 2003; de Lapeyre de Bellaire et al., 2010). As a consequence, there has been a significant increase in the cost of disease control and an increase in negative environmental effects. In strategies using protectants alone, 30– 40 kg a.i./ha/year are applied compared with 2–4 kg a.i./ha/year in strategies using systemic fungicides combined with a forecasting system (de Lapeyre de Bellaire et al., 2010). The environmental impact of the chemical control of black leaf streak is related mainly to the use of protectant fungicides known as ethylene bisdithiocarbamates (EBDCs, primarily mancozeb) and chlorothalonil; the former for its risk to human health and the latter because of its risk to aquatic organisms (Caux et  al., 1996).

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Since the 1970s, EBDCs including mancozeb have undergone extensive regulatory review in many countries, primarily because of their frequency of use and worldwide importance to agriculture (IARC, 2001; Gullino et al., 2010). Investigations on the risk posed by mancozeb have focused almost exclusively on its degradation product ethylenethiourea (ETU), which is considered more toxic to humans than mancozeb itself. In soil and agricultural waters, ETU is ­rather rapidly degraded to ethylene urea with its half-life being 1–7 days under field conditions (Cruickshank and Jarrow, 1973; Ross and ­Crosby, 1973). In a banana plantation in south-­eastern Mexico with a history of a long-term application of mancozeb, the ETU in the sediment was near the detection limit (0.01 mg/kg) and did not show any accumulation. However, manganese and zinc in the sediment at all sampling sites exceeded the threshold values for aquatic life. In contrast to the low concentration of ETU in sediment, its concentration in drainage and runnel water (5.9–13.8 μg/l) greatly exceeded the European Union (EU) threshold value for drinking water (0.1 μg/l). However, the threshold value for aquatic life was not exceeded. This study concluded that longterm mancozeb application does lead to a severe accumulation of manganese in sediments and of ETU in surface water (Melgar et al., 2008). In another Mexican study undertaken near banana plantations that had been sprayed with mancozeb for 10 years, high levels of manganese, but low levels of ETU, were detected in soil. High levels of ETU, but low levels of manganese were found in surface and sub-surface waters (Geissen et  al., 2010). In a recent study in Costa Rica, van Wendel de Joode et al. (2016) found high concentrations of manganese in drinking water near banana plantations where mancozeb had been applied from the air. However, as manganese occurred naturally in groundwater in the area, they concluded that aerial spraying was not fully to blame. Urinary ETU concentrations, a biomarker used to assess human mancozeb exposure, were found by van Wendel de Joode et al. (2014) to be high in pregnant women living within 5 km of a banana plantation aerially sprayed with mancozeb. Levels decreased the further these women lived from the plantation. These findings suggested to the authors that the elevated concentrations of urinary ETU are partially due to exposure to mancozeb applied by aircraft (van Wendel de Joode et al., 2016).

Chaves et  al. (2007) investigated the environmental fate of chlorothalonil in a banana plantation of Costa Rica and concluded that it was degraded very rapidly in the soil under tropical conditions. Approximately 45% of the initial concentration had dissipated 24 h after application. Half-lives in the soil and on the surface of the banana leaves were 2.2 and 3.9 days, respectively. However, residues could still be detected after 85 days. Difenoconazole, a systemic fungicide belonging the group of triazols that are applied aerially and by motorized hand sprayer in plantain plantations, has been detected in water samples in Costa Rica (Polidoro and Morra, 2016). The banana industry is concerned about the environmental and health consequences of fungicide applications at the high levels required to control black leaf streak. It spends millions of dollars annually on research to protect the environment and improve fruit quality. Integrated pest management (IPM) strategies are widely practised to limit the use of crop protection chemicals (Marín et  al., 2000). In 1992, the Costa Rican banana industry established the ­ ­Banana Environmental Commission, which is an inter-institutional group composed of representatives from the private and public sectors, including universities. The organization is dedicated to developing and encouraging sustainable banana production. As part of its responsibilities, it undertakes an annual field audit of all banana farms in Costa Rica to ensure an adherence to environmental and social commitments. The agricultural aviation industry adopted electronic navigation systems so that workers on the ground, who used to guide the planes with flags, were no longer exposed to fungicide spray (Marín et al., 2000; Martinez et al., 2011). Later, ‘block’ spraying programmes further reduced the exposure of workers to aerial sprays. Today these systems have been widely adopted in Latin America, the French Antilles and Philippines. ‘Intelligent flowmeters’ (IntelliFlow®), which automatically control and accurately adjust the volume of fungicide mixtures sprayed over the fields, and open/close automatic control valves (SprayOff®) to reduce the spraying of non-target areas have also been introduced. In the near future, other technologies, such as variable-rate application based on disease severity, might be implemented (Martinez et al., 2011). Pesticide drift has become another important public concern. Drift mitigation measures



Fungal Diseases of the Foliage

depend on good spraying practices. No single measure can eliminate product drift completely and each must be regarded as complementary to each other. One measure is the use of windbreaks to prevent fungicides reaching water sources (Ucar and Hall, 2001). Buffer zones (Plate 2.14) can reduce drift by up to a 90%. The width of buffer zones varies from 10 m to 26 m. Where field studies indicate that drift can be greater, 30 m buffer zones planted with native species have been recommended (Valenciano et  al., 2007). Cover crops, such as Xanthosomas spp., and other species are planted in secondary and tertiary drains to intercept fungicides and avoid the contamination of run-off water (Plate 2.14). Cover crops also protect the soil from erosion and encourage soil microbes. The use of ground spray equipment based on electrostatic technology was implemented in the highland production areas of the Philippines in the 2000s as aerial application of fungicides could not be undertaken. This technology, which

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avoids spray drift, was also adopted in other countries where banana plantations were close to populated areas. The use of spray planes and helicopters has been banned in the French Antilles to reduce the environmental impact of black leaf streak management. Here, spraying is carried out by operator teams with knapsack sprayers and ground equipment pulled with tractors or trucks. In Costa Rica, Martinez and Guzmán (2010) reduced disease severity by 30% compared with conventional equipment using an electrostatic knapsack sprayer.

Biological control

M. Guzmán, D.H. Marín and L. Pérez-Vicente introduction. 

Biological control of disease is based on the use of live microorganisms or their metabolites to destroy, totally or partially, plant

Plate 2.14.  Planting of cover plants (Xanthosoma sp.) along the borders of drains inside plantations (left), and reforestation of buffer zones along roads (upper right) and riverbanks (lower right) in Costa Rica helps to prevent fungicide drift (photos: S. Laprade and M. Guzmán, CORBANA).

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pathogens or directly protect plants against their attack at the infection site, before or after the infection has occurred (Agrios, 2004). Biological control can be carried out through the introduction of specific biocontrol agents or through the management of the environment to favour native biocontrol organisms (Arauz, 1998). This type of control has been successful with some soilborne pathogens, but very few successful cases are reported with foliar diseases, such as black leaf streak (Guzmán, 2012). The main reason for this lies in the characteristics of the phylloplane, which is a hostile environment for the antagonistic organisms since they are exposed not only to strong changes on temperature, humidity and UV radiation, but also to the low and heterogeneous availability of nutrients, such as carbohydrates and proteins (Blakeman and Fokkema, 1982; Ceballos et al., 2012). Nevertheless, it is in the leaf surface where the antagonistic microorganisms that may control agents for foliar diseases can be found and selected. Biological control of black leaf streak has been extremely challenging. This is because of the polycyclic nature of the disease, the fact that plants/suckers in all stages of growth are found in plantations and because highly susceptible leaves unfurl every 6–12 days. The banana leaf cuticle is also hydrophobic and suspensions of microorganisms in water droplets tend to run off, resulting in a failure of control. The availability of highly effective fungicides means that the development of biological control measures against P. fijiensis has received little attention and financial support. However, the detection of populations of the pathogen less sensitive or resistant to systemic fungicides and the increasing demand for environmentally safe control measures has increased the interest in finding alternatives for the control of black leaf streak (Marín et al., 2003). bacteria as biocontrol agents.  Bacteria are the main group of microorganisms that have been tested as possible biocontrol agents. Estimated sizes of cultivable bacterial populations in banana and plantain leaves range from 1.25 × 103 to 9.64 × 105 colony-forming units (CFU)/g of fresh leaf (Ceballos et al., 2012) or 2.24 × 102 to 2.72 × 108 CFU/cm2 of fresh leaf, increasing with the age of the leaf (Alfaro, 2013). In pioneering work in Costa Rica, Jiménez et al. (1987)

isolated 225 epiphytic bacteria from banana leaves and evaluated them for activity against P. fijiensis. Only 12 isolates showed efficacy in vitro and just one isolate (Pseudomonas sp.) provided better control than chlorothalonil under glasshouse conditions. However, because species of Pseudomonas selected for biological control have been difficult to establish in the phylloplane, the search for a biological control agent has focused on aerial endospore-forming bacteria, such as Bacillus and Paenibacillus spp. Species in these genera have a greater potential to colonize the foliar surface, form resistance structures, which aid in surviving adverse environmental conditions, and excrete substances with fungicide activity, such as surfactins, iturins and fengycins (Marín et al., 2003; McSpaden-Gardener, 2004; Stein, 2005; Ongena and Jacques, 2008; Ceballos et al., 2012). In the late 1990s, research efforts began on the selection of bacteria from the genera Bacillus and Serratia capable of secreting cell wall hydrolytic enzymes, such as chitinases and glucanases, and with the ability to survive in the phylloplane (Marín et  al., 2003; Guzmán, 2012). González et al. (1996a) isolated 120 chitinolytic bacteria from banana leaves in areas with high and low black leaf streak disease severity in Costa Rica. They found the highest frequency of isolates in areas where disease severity was low. Two isolates of Serratia marcescens showed good antagonistic activity against P. fijiensis under laboratory conditions. These isolates also controlled the ­disease slightly better than propiconazole in a glasshouse trial (González et  al., 1996a, b). However, Miranda (1996) did not observe effective disease control in a field trial using the same ­isolates and three strains of Bacillus. In addition, Guzmán et al. (2005a) obtained very little control of the disease with a commercial formulation of Bacillus subtilis in the field in Costa Rica. An acceptable level of control, comparable to commercial protectant fungicides, has been ­obtained in times of low disease pressure with a commercial formulation of B. subtilis applied in a mixture with low rates of mancozeb. It was thought that the competition between B. subtilis and other microorganisms in the phylloplane was reduced, which improved the inhibitory effect of the bacterium on the leaf surface ­ (J.F. Rodriguez and D.H. Marín, Costa Rica, 2010, personal communication).



Fungal Diseases of the Foliage

Recent studies have focused on isolating endospore-forming bacteria with antagonistic effects on P. fijiensis from the leaf surface of Musa plants (Ceballos et al., 2012; Carr 2013). B. subtilis and B. amyloliquefaciens produced metabolites that altered the cell wall structure of ascospores and mycelia of P. fijiensis (Plate 2.15) and strongly inhibited the pathogen under in vitro conditions. Gutierrez-Román et  al. (2015) evaluated the antifungal activity of individual purified ­chitinolytic enzymes of the strain CFFSUR-B2

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of S. marcescens and observed a reduction on the germ tube growth of ascospores. However, none inhibited germination, even when applied in combination with prodigiosin, a pigment produced by various bacteria but first characterized from S. marcescens, which is reported to have antifungal, immunosuppressive and anti-proliferative activity (Khanafari et al., 2006). A similar effect to that caused by benzimidazoles on ascospore germination was observed. However, none of the purified components produced a greater inhibitory

Plate 2.15.  Isolates of chitinolytic fungi from banana leaves growing in a chitin-rich media – the arrow shows a chitin degradation halo formed by one fungus, which is evidence that chitin is being utilized as a carbon source (upper left). Deleterious effect of chitinases and other secondary metabolites produced by one antagonist bacterium on the germination of Pseudocercospora fijiensis ascospores on water agar (upper right); efficacy of filtered fermentation broth of a chitinolytic fungus on black leaf streak control under greenhouse conditions – the areas of leaf within the black lines were inoculated, but only the square at centre was applied with the ferment of the fungus (photos: C. Carr, F. Alfaro and M. Guzmán, CORBANA).

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effect than the living cells, which completely ­inhibit ascospore germination. Similar results were obtained with chitinases extracted from Streptomyces galilaeus CFFSUR-B12, which failed to inhibit ascospore germination, but inhibited germ tube elongation and mycelial growth. The crude supernatant from cultures and living cells of S. galilaeus were the most effective in inhibiting P. fijiensis (Moreno-Castillo et al., 2016). as biocontrol agents.  Several fungi with chitinolytic properties (Plate 2.15) have been isolated from banana leaves, with some of them having the capacity to produce secondary metabolites with high fungicidal activity against P. fijiensis under in vitro and glasshouse conditions (Carr, 2009). Arzate-Vega et  al. (2006) evaluated the antagonistic effect of 25 strains of Trichoderma spp. isolated from soil in banana plantations of Mexico and two of these also showed a strong antagonistic effect against P. fijiensis in the laboratory and glasshouse. Abiala et  al. (2010) demonstrated the antagonistic effects of culture filtrates of Trichoderma asperellum and T. longibrachiatum on the mycelial growth of P. fijiensis under in vitro conditions. Twenty-nine isolates of Trichoderma were also evaluated under field conditions, with four out of the 29 significantly reducing disease severity. Isolate 2047 identified as T. atroviride was as effective as the fungicide azoxystrobin. Germination of conidia of isolate 2047 was not inhibited in vitro when exposed for 3 h to chlorotalonil, flutriafol, copper oxychloride and azoxystrobin. This was an indication that this biocontrol agent could be applied with fungicides (Sagratzki-Cavero et  al., 2015). Studies with T. harzianum DGA01, B. amyloliquefaciens DG14 also confirmed the inhibition of these pathogens (Alvindia, 2012). In other work on an organic banana farm in Ecuador, a Trichoderma-based product showed an acceptable efficacy against P. fijiensis as well as having a bio-stimulant effect on the cultivar ‘Williams’ (AAA, Cavendish subgroup) during two production cycles (Castro et al., 2015). Oye Anda et al. (2015), working under laboratory conditions, observed a reduction in black leaf streak symptoms caused by the arbuscular mycorrhizal fungus Rhizophagus irregularis MUCL 41833. They suggested that arbuscular mycorrhizal fungi may decrease, at least at the early stage of infection, symptoms of P. fijiensis

fungi

possibly via the induction of a systemic resistance pathway. However, the impact of the pre-colonization of plantlets at the early stage of growth and their effect on black leaf streak in the field has yet to be determined. foliar substrates. 

An interesting and promising way to promote the biological suppression of P. fijiensis has been the application of ‘foliar substrates’. This strategy enriches and modifies the phylloplane to stimulate the establishment and development of selected specific biological control agents or native microorganisms that increase competition and antagonism against the pathogen. Cow’s milk and sugarcane molasses increased the in vitro growth of the biological control agent S. marcescesens and also improved its activity in the glasshouse. However, this stimulatory effect was not observed in field (Ruiz-­ Silvera et  al., 1997). Salazar et  al. (2006) observed a significant increase in the population of chitinolytic and glucanolytic bacteria on banana leaves with the application of a mineral base solution, barley flour and urea. Other substances and compounds that have shown good results as foliar substrates are chitin, glucan urea and calcium nitrate. Combinations of these with milk and molasses in alternation with fungicides made it possible to reduce the number of fungicides applications for black leaf streak control under field conditions in Costa Rica by 40% (Arango, 2002). Similar results were reported by Patiño et al. (2007) in Colombia. They achieved a 40–46% reduction in sprays by alternating fungicides with applications of colloidal chitin, barley flour as a source of glucan, urea and a mineral solution, which promoted naturally occurring chitinolytic and glucanolytic bacterial populations.

microbial ferments, compost teas and biofertilizers.  The foliar application of microbial fermentation products, compost teas, vermicompost leachate and biofertilizers has been tried in attempts to find a sustainable method to achieve black leaf streak control. A mixture of fermentation leachates and the product of the bokashi fermentation of banana waste controlled P. fijiensis on plantain in Colombia (Marin et  al., 2008). Soto and Soto (2009) also demonstrated the effect of liquid biofertilizers enriched with certain microorganisms and determined that



Fungal Diseases of the Foliage

these fermentation products not only are good sources of nutrients, but they also provide and increase the population of antagonistic microorganisms on the leaf surface. Meulemeesters et  al. (2016) reported that a new biofungicide ­obtained as a fermentation product from a soil bacterium controlled the disease similarly to ­ mancozeb when evaluated in a small-plot trial in Costa Rica. plant extracts.  Plant extracts as well as essential and vegetable oils have also been studied as potential sources of natural fungicides for use against black leaf streak. Viveros-Folleco and Castaño-Zapata (2006) evaluated in vitro the activity of extracts from 30 plant species and they observed high inhibitory effects with the extracts of Cinnamomun zeylanicun (cinnamon) and Azadirachta indica (neem), which were similar to those obtained with fungicides. Marín et al. (2008) also evaluated the effect of plant extracts from Swinglea glutinosa, Salvia officinalis, Carica papaya and Azadirachta indica with promising results. The methanolic extract of Topobea discolor (Melastomatacea) and Phyllantus sp. (Phyllanthaceae) at 1000 mg/l completely inhibited germination of ascospores, similar to benomyl at 1 mg/l. Alkaloids, steroids, triterpenes, phenols, flavonoids and tannins, which act against P. fijiensis, were found in plant extracts (Mosquera et al., 2009). Extracts of Heliotropium indicum, Lippia origanoides and Ricinus communis alone and in different combinations were evaluated and compared with benomyl in a small-plot trial. They were found to be antifungal (Vargas-Hernández et al., 2009). Riveros and Arciniegas (2004) evaluated the in vitro effects of 20 plant extracts on ascospore germination and mycelial growth of P.  fijiensis. Extracts from Commelina difusa, Mormodica charantia, Pavonia sp., Plenax sp., Piper hispidum, Piper peltatum, Sida rhombifolia and Sysygium aromaticum had the highest inhibitory effects, with one from the latter species ­being the most active. A natural fungicide (trade name NECO), extracted from the essential oil of fresh leaves of Ocimum gratissimum and sprayed in mixture with mineral oil, significantly reduced the disease after each treatment. Kassi et al. (2014) believed that the product could be integrated into a disease control programme. Oil from betel, garlic

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and capsicum were not successful in killing dormant mycelium, but caused a significant ­delay in development of P. fijiensis (Alvindia, 2012). In contrast, castor, oil palm and soybean oils affected ascospore germination (tube shortening and deformation), whether they were incorporated into or distributed over the surface of the medium, with castor oil being the most active. Under field conditions, castor oil and mineral oil had a better effect on black leaf streak control when applied without fungicide. When mixed with bitertanol, the effect of soybean and oil palm oil was more evident (Madriz et  al., 2008). Martínez and Guzmán (2006) evaluated the efficacy of wax and three formulations of oil palm oil in mixtures with fungicides as alternatives to mineral oil under field conditions. The wax did not improve the efficacy of fungicides, but the oil formulations worked well as fungicide adjuvants. However, with consecutive applications, mineral oil achieved better control of the disease due to its fungistatic effect. The authors believed that organic oils showed potential for use in disease control, but more research is needed in order to optimize their effects (Madriz et al., 2008). To date, biological control has not been used to control black leaf streak effectively on a large scale. However, work on biological control methods has increased considerably over the past 15 years. Because of environmental and consumer concerns, large agrochemical companies are now investing in research and there is increasing interest in incorporating biological control in integrated disease management strategies. This trend indicates that it may not be long before new and more efficient antagonists will be discovered and perhaps utilized. Further studies may include the selection and evaluation of endophytic microorganisms isolated from banana roots to induce systemic resistance against black leaf streak (Guzmán, 2012). This could be a promising area of research, as has been recently demonstrated by Marcano et  al. (2016). More research and investment in the formulation technology of biopesticides for tropical conditions are needed. In addition, a better understanding of the means of their implementation is required. There is no doubt that biological control will become an important tool for disease management in organic and conventional banana production in the future.

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Introduction of resistant cultivars

M. Guzmán, R.A. Romero and L. Pérez-Vicente Cultivars resistant or less susceptible to black leaf streak than local clones have been deployed in areas where the disease has been causing ­significant yield reductions to crops grown by subsistence farmers. In Central America, ABB cooking-­banana cultivars, such as ‘Chato’ (Bluggoe subgroup), ‘Pelipita’ and ‘Saba’, have been tested as substitutes for the local susceptible plantain (Jarret et  al., 1985; Pasberg-Gauhl, 1989). In West Africa, the International Institute of Tropical Agriculture (IITA) micropropagated AAA clones from East Africa and ABB cooking-banana cultivars were distributed to ­local farmers whose own traditional plantain cultivars were severely attacked by black leaf streak (Vuylsteke et al., 1993b). However, due to differences in palatability, these new introductions were not fully accepted. The uptake of certain black leaf streak-resistant plantain substitutes bred at IITA was also resisted because fruit characteristics differ from traditional cultivars (Ferris et al., 1996). The development of disease-resistant hybrids with fruit that suit local palates and cooking qualities that match traditionally grown cultivars remains a major challenge to banana breeders. In Cuba, the government has had more freedom to initiate change in the face of yield declines caused by black leaf streak since 1990. Large areas are now planted with new black leaf streak-resistant hybrids developed by the Fundación Hondureña de Investigación Agrícola (FHIA). The Dominican Republic is another country where hybrids resistant to black leaf streak were accepted on local markets. ‘FHIA-20’ and ‘FHIA-21’(AAAB bred French plantain-­like hybrids) were intensively evaluated in the island and found to have yields superior to local cultivars (Rengifo and Galvan, 2005a, b; Ventura and Jiménez, 2005). In 2015, 20% of the plantain cultivated area in the Dominican Republic, which was estimated to be 61,688 ha, was under ‘FHIA-20’ and 2% under ‘FHIA-21’. ­ ‘FHIA-20’ is grown more because of its higher yields and greater resistance to black leaf streak (D. Rengifo, Dominican Republic, 2016, personal communication). In Jamaica, FHIA hybrids, such

as ‘FHIA-17’, ‘FHIA-20’, ‘FHIA-21 and ‘FHIA25’, have been introduced and accepted by local growers and consumers (J. Conie, Jamaica, 2013, personal communication). A number of FHIA hybrids have also been adopted by growers in Ghana (Dzomeku et al., 2007). After the detection and spread of black leaf streak in Brazil, the Brazilian Agricultural Research Corporation (EMBRAPA) recommended that farmers should grow the resistant cultivars ‘Caipira’ (AAA, Ibota subgroup), ‘Thap Maeo’ (AAB), ‘BRS Conquista’ (a mutation of ‘Thap Maeo’) and some FHIA hybrids. EMBRAPA has more recently released a number of resistant AAAB hybrids from its own breeding programme. These are the Pacovan-types ‘Japira’, ‘Vitória’, ‘Pacovan Ken’ and ‘BRS Preciosa’, and the Prata-­types ‘BRS Garantida’ and ‘Caprichosa’. The Prata-type hybrid ‘BRS Platina’ and the Pome-type hybrids ‘BRS Tropical’ and ‘BRS Princesa’ (all AAAB) are also recommended, but considered as partially resistant (de Oliveira e Silva et  al., 2016; F. Haddad, Brazil, 2016, ­personal communication). In some areas, black leaf streak-resistant clones have gradually been replacing more susceptible types through sheer necessity. Watson (1993) reported that the spectrum of cultivars grown on atolls in the western and central Pacific changed over the 20 years from 1973 to 1993. It was noted that the resistant clones ‘Saba’ (ABB) and ‘Mysore’ (AAB) were much more widely grown in the Federated States of Micronesia and that ‘Saba’ had spread to the Marshall Islands. In the south, ‘Bluggoe’ (ABB) and ‘Rokua Mairana’ (‘Kalapua’-like ABB clone) had replaced other more susceptible cooking types on Tokelau and the northern Cook Islands. ‘Mysore’ was also more widespread. Resistant cultivars, such as ‘Mysore’ and the bred hybrid ‘T8’ (AAAA), have also been used in Australia to reduce inoculum levels and thus buffer the advances made by black leaf streak in the Torres Strait and Cape York Peninsular region of Queensland. The development of hybrids resistant to black leaf streak continues as a major priority of banana breeding programmes, because resource-­ poor people cannot afford chemicals for disease control, and fungicides that are now used by those who can afford them may not be effective in the future. Small-scale, low-technology plantations and subsistence banana plots essential



Fungal Diseases of the Foliage

for food security are seriously affected by the disease. The problem is accentuated by the fact that black leaf streak is pathogenic on plantain cultivars previously resistant to Sigatoka disease. Resistant cultivars offer the only viable solution. Disease management on organic farms

M. Guzmán and L. Pérez-Vicente introduction. 

Since the 1980s, organic banana production has increased exponentially in Latin America and the Caribbean (Wielemaker, 2008; Pocasangre et al., 2015). By 2010, organic bananas were harvested from more than 25,000 ha and comprised approximately 7% of exports. The cultivation of organically grown bananas continues to expand in zones in the dry tropics and subtropics of some South American and the Caribbean countries that have less conducive environmental conditions for the development of black leaf streak. Successful production areas lie in parts of Ecuador, the Dominican Republic (Mao, Cibao Central), Peru (Piura Valley) and Colombia (Las Guajiras) (Pocasangre et al., 2015).

control by disease avoidance. 

The organic production of bananas in the wet tropics has been extremely challenging, because of high disease pressure in an environment much suited to P. fijiensis and the lack of suitable, efficient technologies for disease control. However, production is possible in tropical areas where the pathogen is largely or entirely excluded by unfavourable ­climatic conditions, such as in areas of low rainfall, low relative humidity and unfavourable

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temperatures. Most organic bananas are produced where the rainfall is less than 900 mm/ year and there are long dry periods (Pocasangre et al., 2015). This type of control method is called evasion or avoidance of the pathogen (Agrios, 2004). In Ecuador, the biggest banana exporting country in the world, black leaf streak has become increasingly aggressive. As a result, more fungicide applications are needed for disease control and this has significantly increased production costs (Jiménez et al., 2007). Because of environmental concerns regarding heavy pesticide use and soil degradation, many banana growers in areas with low rainfall have moved towards organic banana farming. Today, over 5000 ha of bananas have been certified as organic in Ecuador, whereas many more are in the process of transition towards organic production (Jiménez et al., 2007). Organic bananas for the export market are also produced in the Dominican Republic mostly by small-scale farmers in Azua, Valverde and Montecristi provinces. More than 12,000 ha are now under organic cultivation, with another 4000 ha in the transition phase (D. Rengifo, Dominican Republic, 2016, personal communication). The production of organic bananas in Peru is unique in that it has developed on sandy soils in the arid tropics, mostly in the northern department of Piura (Plate 2.16). There are currently more than 6000 ha planted with Cavendish cultivars, mainly of ‘Valery’. More recently, the Cavendish cultivar ‘Williams’ was introduced because of its suitability for the particular conditions of the region. Neither black leaf streak nor

Plate 2.16.  Organic banana plantation established on sandy soil in Piura, Peru (left). Note the under-­ canopy irrigation system and the absence of leaf spot disease (right) (photos: E. Montero, Peru).

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Sigatoka leaf spot disease has been detected, because the climatic conditions do not allow their establishment. Temperatures are 14–25°C with an average relative humidity of 69.2% and an annual rainfall of less than 150 mm concentrated mostly between February, March and April (E. Montero, Peru, 2016, personal communication). cultural practices. 

An important component of black leaf streak control in organic plantations is the removal of affected leaves and leaf tissue, for reasons previously explained. The frequency and intensity of sanitation operations can vary during the year according to disease pressure and the amount of diseased tissue. An important procedure is the removal of hands at bunch shooting (de-handing), the number eliminated depending on the photosynthetic ability of the plant to fill the remaining fruit. This practice allows uniform bunch development and the harvest of a quality product, as regards fruit size, weight and green life. During periods of high ­incidence of black leaf streak, more rigorous de-handing can accelerate fruit development, shorten the time to harvest and reduce the effects of the disease on fruit ripening. The amount of de-handing intensity is adapted to local conditions of the banana farm. Fruit from heavily diseased plants have 2 weeks more physiological age than fruit from disease-free plants or those with a low disease severity and should be harvested earlier. Fruit tagging at bunch shooting is customary for monitoring bunch age. However, despite these practices, production records indicate that intense de-leafing can have reduced impact on fruit quality and green life (F. Wielemaker, Costa Rica, 2016, personal communication).

nutrition.  The use of biofertilizers, foliar and/or soil applied, is a common practice in organic plantations. Different systems of composting, such as vermi-compost and lixiviates of compost (compost teas), as well as the so-called ‘biols’ (products of anaerobic and aerobic fermentations of weeds, manure and plant parts using efficient microorganisms), are utilized to supply the nutritional needs of plants. These products have not only been shown to provide nutrition to plants and improve disease control (Alvarez, 2004; Pérez-­ Vicente, 2006), but they also encourage actinomycetes, bacteria and other organisms in the microflora of the soil, and reduce nematode populations (Jiménez et al., 2007; Soto and Soto, 2009).

mineral oil and biofungicides. 

Mineral oils are very important tools in the management of black leaf streak in organic plantations, but their physical and chemical properties must comply with the specifications required for organic production (Wielemaker, 2008). Copper-based fungicides, such as Bordeaux mixture, are also utilized by some farmers, but they are not approved by most organic organizations. In the near future, they will not be permitted to be used in plantations exporting organic fruit to Europe (Pocasangre et al., 2015). In the Dominican Republic, sanitation practices are integrated with the application of biological fungicides, mineral oils, ‘biols’ and leachates that are allowed in organic banana production. Céspedes (2008) described ten organic fungicides used for black leaf streak control in the Dominican Republic based in plant extracts, organic acids, tocopherols, hydrogen peroxide and others substances. Unfortunately, claims for the efficacy of organic biofungicides to control black leaf streak are based mostly on testimonial observations rather than on rigorous experimentation (Pérez-Vicente, 2006; Pocasangre et al., 2015). Commercial formulations of Bacillus subtilis, B. pumilus and tea tree oil have more recently been incorporated into control programmes. Organic farmers used to apply 8–12 treatments/year, but more recently have increased this to 12–16 applications because of an increased disease pressure that was probably caused by improper disease management, more favourable weather conditions for the pathogen and a possible change in the genetics of the causative agent (Pérez-Duvergé et al., 2011). More intensive control measures are implemented in localized areas where black leaf streak is more aggressive. These measures reduce inoculum levels, but the management of the disease becomes more difficult if for some reason they are not undertaken (Pérez-Duvergé et al., 2011).

environmental management.  The same practices recommended for environmental management for black leaf streak control in conventional ­production systems are also suitable for organic plantations. Reduction of relative humidity in the field with an efficient drainage s­ ystem, weed ­control and an appropriate plant ­density are important considerations. Where i­rrigation ­ is required, under-canopy or drip ­ irrigation



Fungal Diseases of the Foliage

s­ ystems are preferred to an overhead system, to avoid wetting the leaves and providing free water on the leaf surface, which is necessary for spore production, release, dissemination and germination. Field observations have shown that plants for local market or family consumption growing under shade trees in agroforestry systems have a lower disease severity than those fully exposed to the sun (Martínez-Garníca, 2004). This shadow effect on the development of the disease has been observed in different cultivars of banana and in different environments (Martínez-­Garníca, 2004; Cordeiro et  al., 2011; Siles et  al., 2013). There are several possible explanations for this phenomenon. The main one is related to the ­activity of toxins produced by P. fijiensis involved in pathogenesis (Lepoivre and Acuña, 1989; Upadhyay et al., 1990a, b). This activity, which is light-dependent, affects the chloroplast and ­disturbs the proton electromechanical gradient across the plasma membrane, increasing electrolyte leakage and inducing an oxidative burst in Musa cells (Harelimana et al., 1997; Lepoivre et al., 2003; Busogoro et al., 2004a, b). Another observed effect is that the shadow acts directly on the establishment of the pathogen on leaf tissue by reducing the formation of dew, which is an important factor for the infection process (Cordeiro et  al., 2011). In agroforestry systems, shade promotes competition by epiphytic microorganism in the phylloplane. Carr (2013) observed that the bacterial populations on leaf surfaces of ‘Grande Naine’ (AAA, Cavendish subgroup) and ‘False Horn’ (AAB, Plantain subgroup) were always higher in the shade compared with those exposed to sun. It is believed that shadow provides an environment with a higher relative humidity and better protection from UV radiation that is more suitable for bacterial populations.

Sigatoka Leaf Spot D.R. Jones Introduction Distribution Until the discovery and spread of black leaf streak disease, Sigatoka leaf spot or yellow Sigatoka, as

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it is now often called, was the most important foliar disease of banana. It was first recorded in Java (Zimmermann, 1902) and later as a problem in the Sigatoka valley on the island of Viti Levu, Fiji (Philpott and Knowles, 1913; Massee, 1914), the location giving its name to the disease. Herbarium specimens indicated that Sigatoka leaf spot was present in Sri Lanka in 1919 and the Philippines in 1921. The disease was first found in Australia in 1924. In the 1930s, Sigatoka leaf spot was reported throughout the Central American–Caribbean region. It was also recorded in Surinam, Guyana and Colombia in South America, Tanzania and Uganda in East Africa, China and West Malaysia. Reports from West Africa, India and Brazil began in the 1940s and the disease was recorded in most other banana-growing countries for the first time in the 1950s and 1960s (Stover, 1962). Sigatoka leaf spot’s rapid global dissemination in the 1930s from original areas of distribution in the 1920s led to speculation that spores of the causal fungus were carried by air currents between continents (Stover, 1962). However, long-distance spread was more likely to have occurred by the uncontrolled movement of propagating material and/or diseased banana leaves used to wrap produce. It is highly likely that the disease may have been present at a low level in many countries for some time before it came to the attention of local plant pathologists. Today, Sigatoka leaf spot is regarded as having a worldwide distribution, although it has not been recorded in the Canary Islands, Egypt or Israel (Meredith, 1970; CMI, 1981). The incidence of Sigatoka leaf spot diminishes in most areas invaded by black leaf streak. The pathogen appears to be much less competitive than P. fijiensis in tropical lowlands, where most banana plants grow. However, despite being at a major disadvantage, the disease can still be found at low levels in certain niches. It was identified in a germplasm collection in coastal Nigeria on ‘SH-3362’, an AA diploid bred by FHIA in Honduras that is susceptible to Sigatoka leaf spot, but has resistance to black leaf streak (C. Pasberg-Gauhl and F. Gauhl, 1994, Nigeria, personal communication). Sigatoka leaf spot would also seem to be less competitive than eumusae leaf spot. In West Malaysia, where eumusae leaf spot is dominant and where black leaf streak is also found, it was detected in ­Selangor in 2008 on ‘Pisang Mas’ (AA, Sucrier

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s­ ubgroup) (K. Grice, Australia, 2016, personal communication). ‘Pisang Mas’ is another cultivar that is susceptible to Sigatoka leaf spot, but has partial resistance to black leaf streak. In northern Thailand, Sigatoka leaf spot is also still found in areas where eumusae leaf spot is dominant (Kaewjan et al., 2012). The situation is different in India, where eumusae leaf spot is also dominant, with no Sigatoka leaf spot detected amongst 99 leaf spot specimens collected (Devi and Thangavelu, 2014). Sigatoka leaf spot is more adapted to cooler environments than black leaf streak and is dominant at higher altitudes in tropical countries where both diseases occur. It is also the only leaf spot found in cooler banana-growing countries, such as South Africa (Surridge et al., 2003a), that are probably climatically unsuitable for black leaf streak. Economic importance and cost of control On a growing banana plant, the most efficient leaves for photosynthesis are the second to fifth, counting down the profile (Robinson, 1996). Lower leaves are progressively less efficient as they are ageing and later senescing. Leaves are not static and, on a vigorous plant of a cultivar in the Cavendish subgroup (AAA) growing under optimal environmental conditions with one new leaf appearing every 7–8 days, the most efficient leaf area is renewed monthly. On a less vigorous plant growing under suboptimal environmental conditions, this renewal time can be extended to several months. Although a banana plant has the internal ability to partially compensate for lost photosynthetic assimilation, due to leaf area destruction, it is important that leaves 2–5 remain as free of excessive shade, severe leaf tearing and disease as possible; otherwise assimilation potential is greatly reduced (Robinson, 1996). Sigatoka leaf spot is very destructive if left uncontrolled. Lesions caused by the disease can coalesce, which leads to the premature death of large areas of leaf tissue and a reduction in vegetative growth in the tropics, as measured by the rate of leaf emergence, rate of increase in plant height and height of the plant at the time of shooting (Leach, 1946). This is because the effects of the disease are not sufficiently great on leaves 2–5 to cause a severe deficit of the photosynthetic assimilates used to power plant growth.

­ nfortunately, the effect of Sigatoka leaf spot on U fruit development is considerable. After shooting, leaf production ceases and the plant is unable to replace those leaves damaged by the disease. Plants of ‘Williams’ (AAA, Cavendish subgroup) with fewer than five viable leaves at harvest – a viable leaf being defined as one with more than 30% healthy tissue – produce lighter bunches (Ramsey et  al., 1990). The greater the damage and the earlier it occurs after shooting, the greater are the effects on yield. On ‘Williams’ plants with between zero and two viable leaves at harvest, yields are reduced on average by around 25–29% (Ramsey et al., 1990). If Sigatoka leaf spot is left uncontrolled, it is not uncommon for plants to have four or fewer leaves at shooting and none shortly afterwards (Meredith, 1970). Yield losses would be expected to be highest in these cases. Sigatoka leaf spot also disturbs the physiology of fruit, resulting in premature ripening (Meredith, 1970; Stover, 1972; Wardlaw, 1972). Premature ripening can occur in the field if plants are severely diseased, or in transit to markets if moderately affected. Some field ripening of bunches occurs on ‘Williams’ plants with fewer than 11 viable leaves at harvest, which reduces marketable yields, and on all bunches on plants with fewer than four viable leaves at harvest, which results in total marketable yield loss (Ramsey et al., 1990). When fruit ripens in transit, consignments are devalued because of uneven ripening and increased risk of postharvest problems. In Australia, badly affected shipments can be condemned because of the risk of infection by fruit fly, which is an interstate quarantine concern. Pulp colour is also affected (Chillet et al., 2014). Sigatoka leaf spot caused widespread disruption to the export trade when first introduced into the Latin American–Caribbean region. In Mexico, production fell from 525,000 t in 1937, the year after Sigatoka leaf spot was first recorded, to 240,000 t in 1941. Exports from the state of Tabasco ceased entirely. In Honduras, production declined to less than one-third of the level prior to the introduction of the disease. Sigatoka leaf spot caused crop losses of 25–50% in Guadeloupe in 1937. In Ecuador, out of 62 million bunches produced in 1954, only 19 million were fit for export because of uncontrolled disease on small farms. In subsequent years, the damage was reduced by the timely application of chemicals. However, 15–17 fungicide applications were required every year to control Sigatoka



Fungal Diseases of the Foliage

leaf spot, which considerably increased the cost of production (Meredith, 1970). The cost of Sigatoka leaf spot control in the North Queensland growing area of Australia has been estimated to be 14% of total production costs (Jones, 1991). Because of its seriousness, developing a commercial cultivar with Sigatoka leaf spot resistance became a priority in banana-breeding programmes. However, this endeavour was not successful in that no bred hybrid was grown commercially as a replacement for the existing export cultivars. Although Sigatoka leaf spot is still important in certain locations, such as Australia and parts of Brazil, black leaf streak is now attracting more of the banana breeder’s attention because it is the major leaf spot problem. Symptoms

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the spot later shrinks and appears sunken and the halo turns a darker brown. The sunken area becomes grey and the halo darker brown, forming a well idefined ring around the mature spot (Plate 2.19), which remains distinct even after the leaf tissue has died (Plate 2.20). On leaves of water suckers and young plants, individual spots tend to be larger and more spherical. Mature spots are normally 4–12 mm in length. Symptom development from specks through streaks to spots has been divided into various stages by different authors (Leach, 1946; Klein, 1960; Brun, 1963). Stover (1972) and Stover and Simmonds (1987) compared these stages. Leaf tissue surrounding spots turns yellow (Plates 2.17–2.20). This is initially more pronounced on the leaf margin side of the spot (Plate 2.17). If the infection density is high, large areas of leaf tissue around spots turn yellow and eventually become necrotic (Plates 2.19 and 2.20). Where

Many descriptions of the development of symptoms have been published (Meredith, 1970). The earliest symptom is a light green, narrow speck about 1 mm in length on the upper surface of the leaf. The speck develops into a streak several millimetres long and 1 mm or less in width, running parallel to the leaf veins. The streak elongates, expands laterally to become elliptical in shape and turns rusty red (Plate 2.17). A water-soaked halo forms around the lesion when the leaf is turgid. This is best seen in the early morning when dew is present on the leaf. This infiltrated tissue quickly turns brown and a young spot is formed (Plate 2.18). The dark brown centre of

Plate 2.17.  Lesions of Sigatoka leaf spot on the upper surface of a leaf of ‘Williams’ (AAA, Cavendish subgroup) in Queensland, Australia. The yellowing leaf tissue visible around the developing spots is more extensive towards the leaf margin (photo: D.R. Jones, QDPI).

Plate 2.18.  Symptoms of Sigatoka leaf spot on the upper surface of a leaf of ‘Williams’ (AAA, Cavendish subgroup) in Queensland, Australia. The mature spots have sunken, straw-coloured centres with dark brown borders. Regular lines of sporodochia are visible within the centres of the spots (photo: D.R. Jones, QDPI).

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Plate 2.19.  Advanced symptoms of Sigatoka leaf spot on the upper surface of a leaf of ‘Williams’ (AAA, Cavendish subgroup) growing in Queensland, Australia. Individual light-coloured spots with dark borders are discernible within an expanse of dead leaf tissue. The necrotic area is itself surrounded by a dark border and yellowing leaf tissue (photo: T. Cooke, QDAF).

mass infection occurs, areas of necrotic leaf become whitish grey within a dark border and the outlines of individual lesions are not well defined. Counting down from the first fully opened leaf, earliest symptoms are first seen on the third or fourth leaf of susceptible plants in an active state of growth, but sometimes appear on the second leaf if conditions are very favourable for infection. Mature spot symptoms are found on older leaves. The overall effect is that the severity of the disease increases in a descending order down the plant. On resistant plants, symptoms may appear only on the very oldest leaves or not at all. Plants with bunches appear more diseased because clean, new leaves are not being produced to displace old ones with symptoms. Symptoms of Sigatoka leaf spot are very similar to those of black leaf streak and eumusae leaf spot.

Causal agent The fungus Mycosphaerella musicola (sexual morph) and Pseudocercospora musae (asexual morph) was the cause of Sigatoka leaf spot under the dual-name nomenclature system. With the introduction of the one-name-for-one-fungus nomenclature system, Pseudocercospora musae is the new species name.

Plate 2.20.  Range of symptoms of Sigatoka leaf spot on the upper leaf surface of a leaf of a cultivar in the Cavendish subgroup (AAA) growing in Queensland, Australia. Some individual lesions are associated with little or no yellowing. Large areas of leaf tissue around other spots are yellowing and becoming necrotic. Mature spots with light centres are visible within large necrotic areas (photo: D.R. Jones, QDPI).

Conidiophores can be formed, if conditions are suitable, at the first brown spot stage. The sporodochia (mass of tightly aligned conidiophores on a dark stroma) develop in the substomatal air chamber and the conidiophores grow through the stoma pore in a tuft-like fashion. As more and more conidiophores emerge, the sporodochia become erumpent, the guard cells ­becoming disrupted, and the adjacent epidermis is pushed back. Sporodochia can easily be seen using a hand-lens (Plate 2.18) and are produced in spots on both sides of the leaf. Conidiophores are pale to very pale olivaceous brown, paler towards the apex, straight or slightly curved, only very rarely branched, without septa, not shouldered or geniculated, narrow towards the apex and without conidial scars. They measure 5–25 μm and are mainly bottle-­shaped, with rounded or nearly truncate apices (Figs 2.12 and 2.13). Conidia are borne terminally and singly on the conidiophore. They are pale to very pale olivaceous brown, smooth, straight or variously curved, occasionally undulate and almost perfectly cylindrical to obclavate–cylindrical (Fig.  2.13). The apex is obtuse or subobtuse and the basal cell is shortly attenuate and has no thickened ­basal hilum. Conidia are usually 2–5-septate or more and usually measure 10–80 μm × 2–6 μm (Meredith, 1970; Mulder and Holliday, 1974b). Spermogonia, which also originate in the substomatal air chambers, develop before the



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10 μm

Fig. 2.12.  Sporodochium of Pseudocercospora musae in vertical section, showing bottle-shaped conidiophores borne terminally on stromatal hyphae (from Meredith, 1970).

10 µm

Fig. 2.13.  Conidia and conidiophores of Pseudocercospora musae (from Mulder and Holliday, 1974b).

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mature spot stage. They are more abundant on the lower surface of the leaf. Microscopically, they appear as small, black, immersed flaskshaped structures, sometimes arising from the stomatic base of old conidial fructifications, and measure 46–77 μm × 37–63 μm. Spermatia are very minute, oblong hyaline cells, which form chains inside the spermogonium and ooze from the ostiole at its apex. They measure 2–5 μm × 0.8–1.4 μm (Meredith and Lawrence, 1970b). Pseudothecia are often present on mature spots and are more numerous on the upper leaf surface. They are dark brown or black, erumpent, with a short protruding ostiole, and have a dark, well-defined wall. Diameters vary from 36.8 μm to 72.0 μm. Inside the pseudothecia, oblong, clavate asci, which measure 28.8–36.0 μm × 8–10 μm, are found. Ascospores inside the asci are 1-septate, hyaline, obtuse–ellipsoidal, with the upper cell slightly broader than the lower. Their dimensions are 14.4–18.0 μm × 3–4 μm. On most agar media, colonies first become visible 4–6 days after inoculation with ascospores or conidia. Growth is slow, but the centre of the colony soon becomes raised, almost hemispherical in side view and unevenly folded. The surface of the colony sectors into distinct patches comprising aerial hyphae differing in texture and colour (Plate 2.21). Colours of colonies include white, light to dark grey, pink and dark green. Black stromatic masses with no aerial hyphae may occur. The submerged mycelium is very compact and dark green to almost black. The optimum temperature for growth is about 25°C. Colonies remain small and diameters of up to only 3 cm after 60 days’ growth have been reported

on potato dextrose agar (Meredith, 1970). Conidia may be produced on colonies and a profusely sporulating culture (Calpouzos, 1954) has been used to provide inoculum for infection experiments (Goos and Tschirch, 1962, 1963). ­Mycelial fragments have been used as inoculum when colonies do not produce or lose the ability to produce conidia (Jones, 1995). Overnight incubation of diseased leaves with early spot stage symptoms at 100% RH and at 25°C should lead to the production of abundant conidia and conidiophores for identification. Conidia closely resemble those produced by P. fijiensis, the cause of black leaf streak disease. However, they are, on average, shorter than those of P. fijiensis and lack a thickened basal hilum. The absence of scars on the conidiophores, which are usually bottle-shaped, also distinguishes P. musae from P. fijiensis. P. musae and P. fijiensis can also be distinguished in leaf tissue and culture by molecular techniques (Johanson and Jeger, 1993; Johanson et al., 1994; Henderson et al., 2003, 2006). More recently, Arzanlou et al. (2007a) developed a test to distinguish P. musae, P. fijiensis and P. eumusae. A genetic analysis of isolates of P. musae from around the world revealed that moderate levels of diversity exist in the Africa, Latin American– Caribbean and Australian regions, with the greatest diversity occurring in Indonesia. All regions had genetically different populations that varied from each other. The make-up of the African population indicated that it was unlikely to have arisen from anywhere other than the Indonesian region. Isolates from the African and Latin American–Caribbean regions were the most similar, with the latter population probably arising from the former. The results also supported the theories that P. musae originated in Southeast Asia and was most likely disseminated between regions with infected planting material/diseased leaves rather than by airborne ascospores (Hayden et al., 2003).

Disease cycle and epidemiology

Plate 2.21.  Colonies of Pseudocercospora musae after 3 months of culture on potato dextrose agar at 25°C (photo: D.R. Jones, QDPI).

The disease develops as described above under ‘Symptoms’, but the period between spore germination and the formation of mature spots ­depends on environmental conditions, the resistance or



Fungal Diseases of the Foliage

susceptibility of the cultivar and the intensity of infection (Meredith, 1970). Infection is believed to occur usually as a new leaf emerges from the pseudostem and unfurls (Stover and Fulton, 1966). If the cultivar is susceptible, the period from infection to mature spot formation can be short. If the cultivar has resistance, mature spots take longer to develop. As a consequence, mature spots are visible on younger leaves of cultivars that are susceptible. Some cultivars appear to be immune, but further studies are needed to determine whether penetration occurs or not. Conidiophores form in the presence of a film of dew or rainwater (Stahel, 1937b). Conidia are dislodged by rainwater to spread the disease to other leaves on the same plant or to leaves on nearby plants if wind-driven. Water drops laden with conidia can fall on unfurling heart leaves and run down the upright cylinder, depositing spores. As the leaf matures, spots develop in lines or other patterns which can be explained by infection of the leaf at different stages of unfurling (Stahel, 1937b; Stover and Fulton, 1966; Meredith, 1970). Other studies have shown that conidia can also be airborne (Burt and Rutter, 1997). Cultures of P. musae derived from a single conidium or ascospore are sterile. The fungus is a heterothallic species and two compatible individuals must be mated before asci are formed. Spermatia, produced in the spermogonia, move in a film of moisture to protopseudothecia of a compatible mating type. There is indirect evidence that one to three hyphae that emerge from protopseudothecia and protrude through the stomata act as trichogynes. Spermatia of P. musae are believed to be functional because spotted leaves that are not wetted by dew or rain, and thus do not have a film of water to facilitate mating, do not produce pseudothecia and ascospores (Stover, 1963). As spots mature, fertilized protopseudothecia develop into pseudothecia containing asci. Ascospores are forcibly ejected from pseudothecia during wet weather or after a heavy dew and can be carried in wind currents. The greatest number of ascospores is discharged after rain that follows a prolonged dry period. Discharge is also enhanced by alternative wetting and drying (Price, 1960). Ascospores are believed to be responsible for the long-distance spread of the disease and form characteristic apical spotting

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symptoms on leaves after disease development. Stover (1980a) reported that leaves on the ground subjected to intermittent rainfall, dew and partial drying in the daytime continued to discharge ascospores in declining amounts for up to 4 weeks. Ascospores have survived for as long as 8 weeks in the shade on leaf tissue above the ground (Stover, 1971a). Ascospores can germinate within 2–3 h of deposition provided that a film of water is present and temperatures are favourable (Brun, 1963). Conidia may take a little longer (Stover, 1965). The optimum temperature for germination is in the range 25–29°C for conidia (Meredith, 1970) and 25–26°C for ascospores (Brun, 1963; Stover, 1965). Germ tubes elongate only in moisture and collapse in hot, dry conditions. Growth can resume later in the presence of rainwater or dew. The optimum temperature for the growth of ascospore germ tubes is 25°C, which is 2°C less than for P. fijiensis (Porras and Pérez, 1997). Germ tubes of P. musae ascospores also grow better at cooler temperatures than P. fijiensis. This physiological difference between the two pathogens may explain the dominance of P. musae in upland plantations (Porras and Pérez, 1997). After a period of epiphyllic growth, which varies from 48–72 h for ascospore germ tubes (Brun, 1963) to 4–6 days for conidial germ tubes, an appressorium is produced above the pore of a stoma and the fungus penetrates by means of a fine infection hypha. Infection through the lower surface occurs more than through the upper surface. Later, hyphae grow out of stomata and can extend 2–3 mm over the leaf surface before forming new appressoria and re-entering the leaf. Temperatures above 21°C favour the disease, with the formation of conidia being optimal at about 25–28°C (Pont, 1960a; Stover, 1965). Disease development is accelerated if infection densities are high; and high light intensity also seems important, as shading can prevent symptom expression (Meredith, 1970). In general, conditions that favour plant growth also favour disease development. The time interval between spore germination and the appearance of the first yellow streak symptoms (incubation period) varies on susceptible plants. It can be as short as 11 days under ideal environmental conditions for disease development (Klein, 1960) or as long as 105 days under unfavourable conditions (Simmonds, 1939).

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Similarly, the time taken for yellow streaks to change into brown spots (transition period) is variable and ranges from 2 to 103 days. Some streaks never develop into spots (Meredith, 1970). In outdoor artificial inoculation experiments in Suriname, Stahel (1937b) found that the first symptoms appeared after 15–17 days, rusty-red streaks after 22–24 days, infiltration of tissue around streaks with water followed by necrosis after 26–27 days and the browning and death of the whole affected area after 30 days. In inoculation studies in misting cabinets in the glasshouse, Goos and Tschirch (1963) noted that conidia germinated after 24–48 h and stomata were penetrated after 4–6 days. The development of disease was favoured by nightly exposure to mist followed by 6–8 h exposure to low humidity during the day. Under this regime of alternating high and low humidity, leaf spots usually developed within 28–35 days. Jones (1995) inoculated young plants derived from tissue culture with mycelial fragments of P. musae and observed that, at 25–26°C, faint chlorotic spot symptoms appeared after 12–16 days and mature lesions after 30 days. Drops of guttation water were frequently seen on water-soaked lesions at the streak-to-spot transition stage, as had been described by Stahel (1937b). Several scales for measuring disease intensity (amount of damaged tissue on individual plants) have been proposed by different authors (Leach, 1946; Guyot and Cuillé, 1958; Brun, 1963; Kranz, 1965; Stover and Dickson, 1970; Stover, 1971b). Methods have been based on allocating a disease rating for each leaf based on the percentage area of leaf damaged by Sigatoka leaf spot. These leaf ratings can then be used to determine disease levels on the plant. The intensity of disease on certain non-­ flowering plants selected at random in plantations has been used to estimate disease prevalence (amount of spotting in a population of plants). Results of such surveys are useful for monitoring changes in disease incidence in plantations over time. However, most systems proposed are time-consuming and cumbersome and have not proved suitable for commercial use. As disease intensity increases, Sigatoka leaf spot lesions first appear on younger and younger leaves. Stover and Dickson (1970) showed that the age of the youngest leaf with ten or more spots correlates well with disease intensity. This

leaf they called the ‘youngest leaf spotted’ (YLS). An estimation of the age of the YLS is obtained by counting down the plant from the first fully unfurled leaf until the YLS is reached. The higher the number, the older the leaf. Disease prevalence is obtained by averaging YLS data from a number of randomly selected medium-sized, non-flowering plants in a plantation. Changes in the average YLS between surveys give an indication of changes in disease prevalence. The average YLS in commercial plantations with regular spray programmes was found by Stover and Dickson (1970) to be 10–11, while the average YLS in unsprayed plots was 3–4.

Host reaction Reaction of Musa germplasm There have been numerous published reports and reviews of the reaction of Musa species and banana cultivars to P. musae in the field (e.g. Parham, 1935; Cheesman and Wardlaw, 1937; Brun, 1962; Simmonds, 1966; Vakili, 1968; Meredith, 1970; Laville, 1983; Tezenas du Montcel, 1990; Daniells and Bryde, 1999; Daniells et al., 1996). In some instances, results have not been the same. Misidentified and mislabelled clones undoubtedly account for some differences. Another possibility is that populations of the pathogen in different locations may differ in their ability to cause disease. However, there is no proof yet that pathogenic variants of P. musae exist, although the appearance of a more virulent form has been suggested to explain the more susceptible reaction of some cultivars in Brazil. Trials to evaluate germplasm for reaction to Sigatoka leaf spot have been undertaken in ­different locations with different environmental conditions. In addition, different methods have been used to measure disease severity and calculate host response. These range from ratings of reaction based on a limited number of observations to a detailed analysis of a variety of ­parameters (Vakili, 1968). Even under the same environmental conditions and using the same evaluation method, readings between plants of the same clone may differ considerably and different interpretations of reaction may be possible. Because of this variability, the layout of screening trials is extremely important, so that



Fungal Diseases of the Foliage

meaningful results can be obtained after statistical analysis. In nature, it is probable that there is an almost continuous gradation of response of Musa germplasm to Sigatoka leaf spot from extremely susceptible to resistant. Therefore, even with statistically validated results, classification into artificial reaction categories is an arbitrary decision. In the following paragraphs, clones that are described as highly resistant to P. musae are those that do not develop symptoms. Clones that have symptoms are divided into partially resistant and susceptible categories. These ­definitions may vary from those adopted by the authors of some papers cited. Screening trials undertaken with wild species in the Eumusa section of Musa have shown that M. schizocarpa, M. balbisiana and M. acuminata ssp. malaccensis, microcarpa, siamea and truncata are on the whole highly resistant to Sigatoka leaf spot (Cheesman and Wardlaw, 1937; Vakili, 1968; Daniells et  al., 1996). Of all the accessions of these wild types tested by Vakili (1968), only one of six accessions of M. acuminata ssp. siamea was seen with symptoms. The reaction of M. acuminata ssp. banksii and ssp. errans is more variable. Vakili (1968) recorded 14 accessions of M. acuminata ssp. banksii as susceptible, seven as partially resistant and five as highly resistant. He also found that one accession of M. acuminata ssp. errans was susceptible, one partially resistant and eight highly resistant. Daniells et al. (1996) screened 16 accessions of M. acuminata ssp. banksii for reaction to Sigatoka leaf spot and found eight to be susceptible, five resistant and three highly resistant. From the above, susceptibility to Sigatoka leaf spot in wild banana populations involved in the evolution of the Eumusa series of banana (see Fig. 1.2) would seem to be the exception rather than the rule. However, Carreel (1995) suggested that M. acuminata ssp. banksii and/ or errans played a major role in this evolution ­because genetic components of one or other or both of these subspecies are found in almost all banana cultivars of the Eumusa series. Therefore, the source of susceptibility to Sigatoka leaf spot in edible banana may have been inherited from susceptible forms of M. acuminata ssp. banksii/errans. All wild Musa in the Australimusa section tested in the field (M. jackeyi, M. textilis, M. maclayi ssp. ailuluai, M. maclayi ssp. maclayi var. maclayi,

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M. peekelii ssp. peekelii and M. peekelii ssp. angustigemma) have been classified as highly resistant (Cheesman and Wardlaw, 1937; Daniells et al., 1996; Daniells and Bryde, 1999). It is not surprising, therefore, that all Fe’i cultivars in the Australimusa series of edible banana so far tested have also been highly resistant (Jones, 1995; Daniells et  al., 1996). Musa ornata in the Rhodochlamys section is also highly resistant (Cheesman and Wardlaw, 1937). Cultivars in the AA genomic group of the Eumusa series of edible banana vary in their ­response to the disease. Vakili (1968) undertook the evaluation of 180 accessions in Honduras in 1963–1964 and found that 45 were highly resistant, 25 partially resistant and 110 susceptible. His results also showed that resistance was present in a higher proportion of accessions originating from Southeast Asia than from the New Guinea–Solomon Island area. Of 82 accessions collected in Papua New Guinea and rated for reaction to P. musae in Australia by Daniells et al. (1996), only three remained free of Sigatoka leaf spot symptoms, indicating high resistance. Twenty-nine fell into the partially resistant category and 50 were rated as susceptible. The AA diploid cultivars ‘Sucrier’ and ‘­Inarnibal’ are generally regarded as susceptible to Sigatoka leaf spot and ‘Pisang Lilin’, ‘Pisang Tongat’, ‘Paka’ and ‘Tuu Gia’ as highly resistant (Cheesman and Wardlaw, 1937; Brun, 1962; Simmonds, 1966; Vakili, 1968; Laville, 1983, Daniells and Bryde, 1999). Vakili (1968) evaluated 16 accessions of ‘Pisang Jari Buaya’, a cultivar that has been used in breeding programmes as a source of nematode resistance, and found 14 to be susceptible and two partially resistant. An accession of ‘Pisang Jari Buaya’ from Papua New Guinea was also found to be partially resistant (Daniells et al., 1996). In the AAA genomic group, ‘Gros Michel’, ‘Lakatan’, ‘Pisang Susu’ and cultivars in the Cavendish subgroup are susceptible (Cheesman and Wardlaw, 1937; Simmonds, 1966; Vakili, 1968; Laville, 1983, Daniells and Bryde, 1999). ‘Red’ and ‘Green Red’ have been rated as susceptible (Laville, 1983; Daniells and Bryde, 1999) and as having degrees of partial resistance (Simmonds, 1933; Simmonds, 1966; Vakili, 1968). Daniells et  al. (1996) classified ‘Mata Kun’ (syn. ‘Red’) as susceptible, but it was less susceptible than a Cavendish cultivar, which was rated at the same trial.

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Cultivars in the Lujugira–Mutika subgroup have been rated as partially resistant (Simmonds, 1966) and ‘Yangambi Km 5’ (Ibota Bota subgroup) as highly resistant (Laville, 1983; Daniells and Bryde, 1999). Kluai Khai Bonng from Thailand is also highly resistant (Daniells and Bryde, 1999) ‘Ney Poovan’ (AB) has been reported as highly resistant by Simmonds (1966) and ‘Safet Velchi’ (syn. ‘Ney Poovan’) was rated as highly resistant by Laville (1983). Reports of the susceptibility of some accessions of this clone (Brun, 1962; Laville, 1983) may be erroneous. Clones in the Plantain subgroup in the AAB genomic group are generally all resistant to P.  musae when tested at or near sea level (Simmonds, 1966; Laville, 1983). However, a ‘Horn’ plantain accession was rated as susceptible close to sea level by Daniells and Bryde (1999). Plantain has been found to be susceptible to Sigatoka leaf spot at elevations above 500 m in Puerto Rico and Colombia (Stover and Simmonds, 1987) and in upland areas of Cameroon (Mouliom-­ Pefoura and Mourichon, 1990). Most workers consider ‘Silk’ (AAB) to be partially resistant (Simmonds, 1933; Brun, 1962; Simmonds, 1966; Laville, 1983) though Daniells and Bryde (1999) rated it as susceptible. Cultivars in the Pome subgroup (AAB) are susceptible (Stover and Simmonds, 1987; Daniells and Bryde, 1999). Cultivars in the Mai’a Maoli– Popoulu subgroup (AAB) have been rated as susceptible (Laville, 1983; Daniells and Bryde, 1999), but resistance may be present in some clones (Parham, 1935). The reaction of ‘­Mysore’ (AAB) has been recorded as varying from high resistance (Parham, 1935; Simmonds, 1966) to high partial resistance (Cheesman and Wardlaw, 1937; Brun, 1962; Laville, 1983; Daniells and Bryde, 1999). Reports of the susceptibility in some clones may be erroneous (Brun, 1962; Laville, 1983). The AAB cultivars ‘Pisang Raja’ (Simmonds, 1966) and ‘Pisang Kelat’ (Daniells and Bryde, 1999) have partial resistance. The situation is less confused for cultivars in the ABB genomic group. ‘Bluggoe’, ‘Pisang Awak’, ‘Kluai Teparot’ and others are known to be either highly resistant or resistant (Simmonds, 1966; Stover and Simmonds, 1987; Daniells and Bryde, 1999). There are probably none that can be classified as susceptible. The high resistance of cultivars in the ABB genomic group has been attributed to their high

‘B’ genome content. As mentioned previously, M. balbisiana, the source of the ‘B’ genome, is highly resistant to P. musae. The general rule is that the greater the ‘B’ component in the genome of the cultivar, the greater the resistance of that cultivar to Sigatoka leaf spot (Meredith, 1970). However, some M. acuminata-derived diploids and at least one triploid are also highly resistant. The AA clones ‘Pisang Lilin’ and ‘Paka’ have been used as pollen sources in breeding for resistance to Sigatoka leaf spot (Stover and Simmonds, 1987). Resistance to the disease ­ ­resides in both M. acuminata and M. balbisiana. All five AAT cultivars tested to date have proven to be highly resistant (Daniells et al., 1996; Daniells and Bryde, 1999) Some hybrids developed in breeding programmes were tested in the field by Daniells and Bryde (1999). They showed that the AAAA tetraploids ‘I.C.2’, ‘Bodles Altafort’ and ‘2390-2’ bred primarily for resistance to Fusarium wilt in the early years of the Jamaican programme, although rated as susceptible, had more resistance to Sigatoka leaf spot than the very susceptible ‘Gros Michel’, which they were bred to replace. ‘T8’, ‘T6’, ‘T12’ and ‘Calypso’, which were AAAA tetraploids bred later in Jamaica, were found to be highly resistant. Two improved AA breeding diploids from the Honduran programme, SH-3142 and SH-3362, were evaluated as very susceptible. The AAAB tetraploid ‘Goldfinger’/ ‘FHIA-01’from the same programme was determined to be susceptible. The results of growth cabinet evaluation techniques based on the inoculation of juvenile plants derived from tissue culture with mycelial fragments (Jones, 1995) have been shown to be similar to results obtained in the field, but exceptions have been noted, suggesting that the technique has its limitations (Daniells et  al., 1996; Daniells and Bryde, 1999). Host–pathogen interactions Black leaf streak attacks many cultivars resistant to Sigatoka leaf spot. Although most cultivars that are resistant to black leaf streak are also resistant to Sigatoka leaf spot, some hybrids bred at the FHIA in Honduras are more susceptible to Sigatoka leaf spot than to black leaf streak. This indicates that the two pathogens may activate different resistance mechanisms in some



Fungal Diseases of the Foliage

instances. The resistance that plantains possess at sea level that is lost above 500 m may also indicate that the gene or genes for resistance in this subgroup active against P. musae may be temperature sensitive. An analysis of differential gene expression has been undertaken by comparing the effects of inoculation on highly resistant M. acuminata ssp. burmanicca (AAw, accession ‘Calcutta 4’) and very susceptible ‘Grande Naine’ (AAA, Cavendish subgroup). Gene transcripts displaying a significant change in expression between the two responses included chitinases, thaumatin, glutathione transferases, a putative avrRpt-2-induced AIG2 protein, a Germin/oxalate oxidase, a regulator of pathogen resistance responses of RPS2 and RPM1 genes and an IAA-amino acid hydrolase. Induced ­ proteins were also isolated for further analysis (Miller et al., 2016).

Control Chemical control Meredith (1970) and Stover (1972, 1990) have reviewed the history of the chemical control of Sigatoka leaf spot. Knowles (1914) in Fiji first advocated the use of Bordeaux mixture to control Sigatoka leaf spot. After the arrival of the disease in the Caribbean and Central America in the early 1930s, large pipeline systems were installed in commercial plantations to deliver Bordeaux, which was applied as a high-volume spray at intervals of 2–5 weeks (Wardlaw, 1941). The costs were high, but control was achieved and the industry saved from possible collapse. Bordeaux worked by controlling conidial production in the leaf spot and conidial infection, but was less effective in preventing infection of the unfurling cigar or heart leaf by ascospores (Leach, 1946). In ­Jamaica, the control strategy was to reduce the number of leaf spots before the rainy season, when ascospores would be released. Spraying with Bordeaux continued until 1957. During this period, zineb and copper oxychloride were found to give good control, but no exceptional substitutes for Bordeaux were found. In the 1950s, the banana industry in the French Antilles was threatened because the hilly terrain made the application of high-volume

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sprays of Bordeaux impracticable. Experiments were conducted with zineb and copper oxychloride in petroleum oil, applied as a low-volume spray. This proved effective and even oil alone applied as a mist gave good control (Guyot and Cuillé, 1954a, b). Oil phytotoxicity caused some initial problems, but improved oils were developed, which led to their widespread use. Oils and oil-based sprays were applied from the ground by various mist blowers or from the air by means of a helicopter or fixed-wing plane at the rate of ­between about 12 l/ha and 15 l/ha, according to disease severity and risks of phytotoxicity at different times of the year. In some years, it was necessary to spray every 10–12 days as new leaves appeared, whereas one application every 3–4 weeks maintained control in other years. Klein (1960) discovered that the development of Sigatoka leaf spot lesions was greatly retarded when oil was applied at the young yellow streak stage and that counts of these streaks on young leaves could be used to forecast when to spray. This method enabled oil to be applied strategically rather than regularly, with associated cost savings. Soon after the adoption of oil spraying, the use of aqueous suspensions of dithiocarbamate fungicides, such as mancozeb, was also shown to control Sigatoka leaf spot. However, it soon became apparent that protectant fungicides were much more effective when applied as oil-inwater emulsions. Benomyl, the first systemic fungicide of the benzimidazole group to be used on banana, was sent to Honduras for trial in 1967. By 1972, it was in widespread use. With the development of systemic fungicides, control was even more effective, as the chemicals could migrate in treated leaves and also from treated to untreated leaves. The penetrant fungicide tridemorph was used soon after benomyl, but was found to be less ­effective. A Sigatoka leaf spot forecasting system based on climatic data (temperature and evaporation), as well as observations of disease symptoms on young leaves, was developed in the French Antilles (Ganry, 1986). A forecasting system based on observations of the most advanced stage of disease development on the first five leaves on a plant has also been published (Ganry et al., 2008). The methods, which rely on the therapeutic action of systemic fungicides, enabled the application of sprays only when necessary.

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Pseudocercospora musae developed resistance to fungicides in the benzimidazole group in the French Antilles in the 1970s and this led to the temporary use of oil alone. Later, propiconizole, a systemic fungicide in the triazole group of demethylation inhibitors (DMIs) was integrated into the forecasting system (Bureau and Ganry, 1987). Flusilazole, another effective fungicide in the triazole group, and imazalil, in the imidazole group of DMI systemic fungicides, have also been used to control Sigatoka leaf spot. In northern Australia in the late 1980s, growers were advised to use mancozeb plus white oil every 14 days in the wet season and every 21–28 days during the dry season, but to use propiconizole in water-miscible oil during periods of high disease pressure. By using propiconizole only when the disease could not be held in check by protectant fungicides in oil, it was thought that the possibility of resistance developing to this valuable systemic fungicide would be lessened. In the Windward Islands, systemic/penetrant fungicides with different modes of action, such as benomyl, propiconizole or flusilazole and tridemorph, were alternated to reduce the risk of the development of resistance. The stages of disease development in young leaves of plants growing in different microclimates were monitored weekly to predict the most appropriate time to spray the crop from aircraft. Plants in locations where Sigatoka leaf spot built up more rapidly were also sprayed from the ground in between aerial applications. This management strategy developed from a forecasting system proposed by Cronshaw (1982). Today, the same fungicides that are used to control black leaf streak continue to be used to control Sigatoka leaf spot. In Australia, resistance to strobilurins is found in all growing areas and also serious shifts in the sensitivity of triazoles have been detected. Growers are urged to de-leaf, and spray protectant fungicides as much as possible with systemics restricted to periods in the summer when the conditions are warm and wet. When systemics are utilized, it is advised that fungicides in different activity groups be rotated to minimize the chances of a build-up of resistance. The maximum number of times any one fungicide group can be used each year is also advised, with strobilurins limited to twice a year and only then if applied with a fungicide from a

different activity group (ABGC, 2015). A protocol for evaluating pathogen isolates for shifts in sensitivity to fungicides has been described (de Lapeyre de Bellaire and Risède, 2008). The efficacy of mancozeb with oil, chlorothalonil, oil in combination with the systemic fungicides propiconazole, tebuconazole, difenoconazole, epoxiconazole and pyrimethanil, and chlorothanil in combinations with the same systemics was evaluated over two cropping ­ ­cycles in northern Queensland, Australia. The results suggested that all the systemics with oil were more effective, or equally effective, than when applied with chlorothalonil and also chlorothalonil alone. Difenoconazole and epoxiconazole with oil followed by propiconazole with oil were the most effective treatments. Pyrimethanil and tebuconazole with chlorothalonil were the least effective (Samuelian et al., 2016). Cultural methods The removal and destruction of badly spotted leaves are recommended as a means of reducing inoculum levels. In Brazil, the excision of necrotic leaves 1 month before harvest resulted in a reduction in premature fruit ripening after harvest (Castellon et  al., 2013). Heavily diseased leaves can also be buried within the plantation or piled on top of one another to prevent the effective discharge of ascospores. Disease severity has been found to be higher on plants growing in nutrient-deficient soils (Freitas et  al., 2016). Banana plants deficient in potassium, nitrogen, phosphorus, sulfur or magnesium had a greater incidence of Sigatoka leaf spot in Brazil than plants supplied with full nutrients and plants deficient in calcium or boron (Freitas et al., 2015). However, in Australia, combinations of calcium and boron levels were also found to influence disease severity (Fitzgerald et al., 2003). Cultural control measures have been discussed more fully for black leaf streak disease. Biological control Meredith (1970) first reported that epiphyllic mycelia of leaf surface-inhabiting fungi could inhibit the germination of spores of P. musae. Bacillus subtilis isolated from the banana rhizosphere has shown promise as a biocontrol agent against P. musae in China (Gang et al., 2010).



Fungal Diseases of the Foliage

Trichoderma harzianum and T. virens isolated from diseased leaves and fruit in northern Queensland were found to be antagonistic against the banana leaf pathogens P. musae, Neocordana musae and Corynespora torulosa (syn. Deightoniella torulosa) in vitro. Several products used by the ­local banana industry to increase production, including molasses, ‘Fishoil’ and ‘Seasol’, were tested as a food source for the isolates. The optimal food substrate was found to be molasses at a concentration of 5%, which, when used in combination with a di-1-p-menthene spreader-sticker, enhanced the survivability of Trichoderma populations under natural conditions. The isolates were sensitive to mancozeb, but oil only marginally suppressed growth in vitro. The author believed that they showed potential to manage leaf spot diseases as a part of an integrated disease approach (Samuelian, 2016). No biological control methods have yet been developed that are used commercially.

Eumusae Leaf Spot D.R. Jones, J. Carlier and X. Mourichon Introduction Specimens of an unknown leaf spot pathogen, which were neither P. fijiensis nor P. musae, were first collected at Onne in Nigeria in 1989 and 1990. The unusual fungus was not immediately investigated, but cultures were retained for future studies (X. Mourichon, France, 2000, personal communication). As the distribution of Sigatoka leaf spot and black leaf streak in South and Southeast Asia was puzzling, with personal observations indicating that the former was not being displaced rapidly by the latter as had occurred in Africa and the Americas, a number of specimens of leaf spot were collected from the region between 1992 and 1995 in an attempt to shed some light on the problem (Jones, 2003). These specimens, collected in southern India, Sri Lanka, Malaysia, Thailand and southern Vietnam, were sent to the Centre de coopération internationale en recherche agronomique pour le développement (CIRAD) in Montpellier, France, for identification.

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Pseudocercospora musae, the cause of Sigatoka leaf spot and a banana pathogen reported in all four countries (Meredith, 1970), was not identified from any of the samples. Some specimens from Johore in West Malaysia and from Vietnam were found to be P. fijiensis and some could not be identified, but all others collected were of an unknown banana pathogen, which was identical to the one found earlier in Nigeria. The same fungus was later identified as the causal agent of leaf spots collected on the islands of Mauritius and Réunion in the Indian Ocean in 1997 and 2000, respectively (Carlier et al., 1999b; Jones, 2003). More recently, the disease has been recognized in Sumatra in Indonesia (Sahlan and Soemargono, 2011). Evidence suggests that eumusae leaf spot has most probably been mistaken for Sigatoka leaf spot and perhaps black leaf streak on visual symptoms in the past. More specimens of leaf spot need to be collected in Asia and analysed to determine its true distribution. However, the disease now appears to be the dominant leaf spot in India, where it has recently been collected from a number of different geographical areas (Devi and Thangavelu, 2014). It has also been confirmed from northern Thailand, where Sigatoka leaf spot was also found (Kaewjan et  al., 2012). In addition, the disease appears to be widespread in plantain-growing areas of Nigeria, with 11 of 95 leaf spot isolates collected identified as the agent causing eumusae leaf spot and the remainder as P. fijiensis (Zandjanakou-Tachin et  al., 2009). Given today’s known distribution, eumusae leaf spot is likely to be present on banana in Asia from India in the west to Vietnam in the east and Indonesia in the south. Its presence on Mauritius, Réunion and Nigeria may be the result of the movement of banana planting material from India.

Symptoms The development of symptoms of eumusae leaf spot is similar to the development of those of Sigatoka leaf spot and black leaf streak. The first obvious symptom seen on inoculated ‘Grande Naine’ (AAA, Cavendish subgroup) has been described as a small yellow speck (Carlier et  al., 1999b). This later broadens and lengthens into a light yellow streak which expands into a large

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light brown spot and darkens. The spot later develops a dark brown border as the centre turns grey. When infection density is low, individual spots are ovoid when mature and surrounded by a chlorotic halo (Plate 2.22 and 2.23). When the infection density is high, developing lesions coalesce and large areas of surrounding leaf tissue become necrotic and are surrounded by a yellow margin (Plate 2.24). Grey spots of individual lesions and more irregular shapes caused by coalescing lesions are visible in the necrotic areas (Plates 2.22 and 2.24). Yellowing of leaf tissues does not appear to be as extensive as that caused by black leaf streak (see Plate 2.8) and Sigatoka leaf spot (Plate 2.20). Causal agent Pseudocercospora eumusae is the name of the fungal species causing eumusae leaf spot. When first microscopically examined, the perfect stage was identified as a Mycospharella and the imperfect stage as a Septoria (Infomusa, 1995). The pathogen was later described and named Mycosphaerella eumusae (asexual morph: Septoria eumusae) with the disease given the name Septoria leaf spot (Carlier et  al., 1999b). However, the asexual morph was subsequently found to be a Pseudocercospora and given the name P. eumusae (Crous and Mourichon, 2002). This led to a proposal to name the disease eumusae leaf spot (Crous and Mourichon, 2002). The initial

Plate 2.22.  Symptoms of eumusae leaf spot on ‘Grand Nain’ (AAA, Cavendish subgroup) near Phicit in northern Thailand. Oval-shaped spots with grey centres are developing from dark streaks and coalescing in places to form irregularly shaped areas of necrotic leaf tissue (photo: D.R. Jones, INIBAP).

Plate 2.23.  Mature oval-shaped spots of eumusae leaf spot with grey sunken centres and dark brown borders are evident on a young leaf of ‘Embul’ (AAB, ‘Mysore subgroup’) in Sri Lanka (photo: D.R. Jones, INIBAP).

Plate 2.24.  High intensity of infection of eumusae leaf spot on a leaf of ‘Anamala’ (AAA, syn. ‘Gros Michel’) growing in Sri Lanka. Developing lesions have coalesced to form large areas of necrotic leaf tissue. Less leaf tissue has yellowed around lesions than has been seen around lesions of black leaf streak and Sigatoka leaf spot. Lesions of eumusae leaf spot can also be seen on the midrib (photo: D.R. Jones, INIBAP).



Fungal Diseases of the Foliage

error in identification arose because sporodochia, which are substomatal and subepidermal, can exude conidia and give the impression they are pycnidia embedded in the leaf tissue. However, as more stromatal tissue is formed, conidiophores become erumpent and the sporodochium breaks through the epidermis becoming acervular-like (Crous and Mourichon, 2002). Reproductive structures (Fig. 2.14) are more prevalent in lesions on the upper leaf surface. Conidiophores are found in dense fascicles that arise from a brown stroma that can be up to 70 μm wide. They are subcylindrical, smooth, hyaline or pale brown below with 0–3 septa. They can be branched or unbranched below with dimensions of 10–25 × 3–5 μm. Conidia are solitary, subhyaline to pale olivaceous, straight or curved with 3–8 septa and dimensions of 18–65 × 2–3 μm, but mostly in the range 30-50 x 2.5-3 μm (Crous and Mourichon, 2002). The hilum is inconspicuous, which distinguishes P. eumusae from P. fijiensis, which produces a conidium with a slightly thickened hilum. Spermagonia are mainly hypophyllous, subepidermal, substomatal, dark brown and up to 75 μm in diameter. Spermatia are hyaline, rod-shaped and measure 3–6 × 1–2 μm. Pseudothecia are globose, black, subepidermal becoming slightly erumpent. They can be up to 80 μm in diameter with an ostiole 10–15 μm wide. Asci are obovoid, straight or slightly incurved and measure 30–50 × 9–15 μm. Eight ascospores are found in each ascus. Each has dimensions of 11–16.5 × 3–4 μm with one ­ ­septa. The ascospore is widest in the middle of the apical cell. The basal cell is slightly longer than the apical cell, with tapering more pronounced towards the lower end (Crous and Mourichon, 2002). Cultures of the pathogen have been initiated from single ascospores ejected from the pseudothecia. All had black stroma-like structures and appeared similar to cultures of P. fijiensis. Pseudothecia were observed 1 week after isolation. A number of micropropagated plants of ‘Grande Naine’ (AAA, Cavendish subgroup) were artificially inoculated with conidia produced by single ascospore isolates of the fungus growing in vitro (Carlier et al., 1999b). Phylogenetic analysis based on sequences of the ITS of ribosomal DNA from P. eumusae,

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P.  musae, P. fijiensis and Mycosphaerella musae confirmed that they are all different species (Carlier et al., 1999b). Genetic diversity analyses carried out using the sequence data of rDNA-ITS region and by RAPD indicated the presence of 8–12 haplotypes among P. eumusae isolates in India and also the existence of 98–100% homology between P. musae, P. fijiensis and P. eumusae (Devi and Thangavelu, 2014). Of the three, P. musae has the largest genome size with P. eumusae the smallest (Chang et al., 2016). All are believed to have been derived from a common ancestor that has been estimated to have appeared about 147–113 million years ago (Chang et al., 2016). ­However, despite a common origin, the three species have different repertoires of candidate effector proteins, suggesting that they likely evolved different strategies for overcoming the host’s defence mechanisms (Chang et al., 2016). Contrary to the timing of their recorded appearance on the global scene as major pathogens on banana, it has been estimated that P. fijiensis evolved first from the common ancestor soon after it appeared and that the progenitor of P.  musae and P. eumusae evolved later about 40–31 million years ago. Finally, both P. musae and P. eumusae may have evolved from their common progenitor 23–17 million years ago (Chang et al., 2016). The factors which determined that P. musae was the first to spread worldwide from its centre of origin are unknown, but may be related to a chance intercontinental movement of infected planting material. Information useful in distinguishing P. eumusae from P. fijiensis and P. musae by morphological differences in their asexual morphs is provided by Crous and Mourichon (2002) and Zapater et  al. (2008b). A molecular method is described by Arzanlou et al. (2007a). Arzanlou et al. (2008) found from the DNA sequence data of isolates from different countries that P. eumusae would appear to be heterogeneous and that further studies would be required to determine if the phylogenetic variation also correlates with differences in ­ morphology. The current distribution of P. eumusae and information on the occurrence of the other two Sigatoka complex pathogens in areas where it occurs suggest that it can, like P. fijiensis, outcompete P. musae. However, its competiveness

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a

b

c

d

e

f

Fig. 2.14.  Pseudocercospora eumusae (syn. Mycosphaerella eumusae). Key: a = asci showing apical chamber; b = ascospores; c = germinating ascospores on leaf surface; d = conidia and conidiophores in vivo; e = spermatophores and spermatia; f = conidia and conidiophores in vitro; scale bar = 10 μm; reproduced from Crous and Mourichon (2002) with permission from Sydowia.



Fungal Diseases of the Foliage

against P. fijiensis is undetermined. The scientific literature suggests that both are found in Nigeria, West Malaysia and Sumatra, Indonesia. In Nigeria, P. fijiensis was identified from leaf spots much more frequently than P. eumusae, but the reverse appears to be the case in West Malaysia. The situation is not clear in Sumatra. Time will tell which of the two, if either, will become the more prevalent in mixed populations. Disease cycle and epidemiology No research has yet been undertaken on this disease in the laboratory or field to determine optimum germination and growth temperatures and main means of dispersal and infection. Host reaction Pseudocercopora eumusae was initially isolated from specimens collected from cultivars in the Sucrier, Gros Michel and Cavendish subgroups plus ‘Pisang Kapas’ (AAB) growing in South and Southeast Asia. The source of isolates in Nigeria was unidentified AAB clones, which were probably cultivars in the Plantain subgroup. Cultures of the pathogen from Mauritius and Réunion were isolated from ‘Grande Naine’ (AAA, Cavendish subgroup). Symptoms have been recognized in southern Thailand on ‘Kluai Lep Mu Nang’ (AA, ‘Pisang Lilin’) and in Sri Lanka on ‘Embul’ (AAA, Mysore subgroup). The response of a number of banana accessions to P. eumusae has been studied in Mauritius (Sulliman et al., 2012). The results indicate that ‘Williams’ (AAA, Cavendish subgroup) is highly susceptible, ‘Petite Naine’ (AAA, Cavendish subgroup), ‘French Clair’ (AAB, Plantain subgroup), ‘GCTCV-119’ (somaclonal variant AAA derived from ‘Giant Cavendish’) and ‘FHIA-18’ (bred AAAB Pome-type hybrid) are susceptible, ‘Pisang Mas’ (AA, Sucrier subgroup), ‘Pisang Ceylan’ (AAB, Mysore subgroup) and ‘FHIA-21’ (bred AAAB Plantain-type hybrid) are resistant and ‘Yangami Km 5’ (AAA, Ibota subgroup) is highly resistant. Much more work is necessary before firm conclusions can be reached. For instance, ‘Mysore’ has been observed to be quite susceptible in India (J. Carlier, 2017, France, personal communication) and also in Sri Lanka (Carlier et al., 2000b).

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Despite P. eumusae being recognized as present and a significant pathogen on banana in ­India in the 1990s, much research has since been published by workers in this country on a leaf spot disease believed to have been caused by P.  musae. However, of 99 specimens of banana leaf spot recently collected in diverse locations in India, all were found to be P. eumusae (Devi and Thangavelu, 2014). It seems highly likely that much of the research undertaken in India in the past was on this pathogen and not P. musae. Evidence for this comes from published information on clonal reactions in the field in India that are not typical of those caused by P. musae. In Nigeria, detached leaf sections of ‘Dwarf Valery’ (AAA, Cavendish subgroup), ‘Agbagba’ (AAB, Plantain subgroup) and the wild seeded diploid M. acuminata ssp. burmannica (AAw, accession ‘Calcutta 4’) were inoculated with mycelial suspensions of each of 11 isolates of P. eumusae and each of 85 isolates of P. fijiensis collected from various locations in banana-growing ­areas. On average, disease severity on ‘Dwarf Valery’ was higher on the plants inoculated with P. eumusae than those with P. fijiensis. While severity averages for P.  eumusae were also higher on ‘­Agbagba’ and ‘Calcutta 4’, differences were not statistically significant. When levels of disease severity caused by the 11 isolates of P. eumusae on the three clones were analysed, two of the isolates were classified as ‘moderately aggressive’, seven were classified as ‘aggressive’ and two as ‘highly ­aggressive’. None were characterized as ‘least aggressive’, as were some P. fijiensis isolates (Zandjanakou-­Tachin et  al., 2013). Transcripts involved in the expression of the resistant response of ‘Manoranjitham’ (AAA) to P. eumusae have also been investigated by comparisons with the susceptible response of ‘Grande Naine’ (AAA, Cavendish subgroup) (Saravanakumar et al., 2016).

Control It is highly likely that eumusae leaf spot can be controlled by the same chemicals that control Sigatoka leaf spot and black leaf streak. Indeed, these fungicides may be being used in countries where the disease occurs. In Thailand, a trial compared the efficacy of weekly sprays of mancozeb alone and in combinations with propiconazole,

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carbendazim and difenoconazole. Mancozeb in combination with the systemic fungicides gave better results than mancozeb alone, with the mancozeb and difenoconazole treatment giving the best result (Kaewjan et al., 2012). In India, water extracts of 33 plant species were tested for their efficacy against P. eumusae, with the most effective, which prevented spore germination, being from Cassia senna, zimmu (an interspecific hybrid of Allium cepa × Allium sativum) and Rhincanthus nasutus. Zimmu was also effective against fungal mycelium. The water extract at 50% concentration reduced disease severity on unsprayed, susceptible ‘Grande Naine’ (AAA, Cavendish subgroup) by 55%. ­Lipid compounds may be responsible for the inhibition (Thangavelu et al., 2013).

Other Leaf Spots Black Cross Leaf Spot D.R. Jones Introduction Black cross leaf spot is found in Australasia– Oceania (American Samoa, Australia, Fiji, New Caledonia, Niue, Papua New Guinea, Samoa, Tonga, Vanuatu) and Asia (Indonesia, Philippines) (Booth and Shaw, 1961; Meredith, 1969; Dingley et  al., 1981; Hyde, 1992). The disease is  not usually regarded as serious. However, in ­Samoa it can be a problem on ‘Fa’i Misiluki’ (AAB, Mysore subgroup) when it is grown in shady areas under trees or coconuts. Under such conditions, black cross leaf spot symptoms can ­cover entire leaves (Gerlach, 1988).

Symptoms Mature symptoms of the disease are found on older leaves and take the form of large, jetblack, four-pointed, diamond-shaped stars or crosses (Plate 2.25). The longer axis of the star runs parallel to leaf veins for up to 6 cm and the shorter arm or arms are at right angles and can extend for about 3 cm. The symptoms are

Plate 2.25.  Symptoms of black cross leaf spot on the underside of a leaf of Musa acuminata spp. banksia (AAw) in the Oro province of Papua New Guinea. Symptoms of Cordana leaf spot are also evident. (photo: D.R. Jones, QDPI).

clearly visible on the underside of the leaf, but are less pronounced on the upper surface, showing only a series of black dots (Booth and Shaw, 1961). In Samoa, a yellow diamond-­shaped spot, interspersed with dark brown lines, has been observed on upper leaf surfaces (Gerlach, 1988). Black cross leaf spot can serve as points of entry for Neocordana spp. (Plate 2.25), which can cause considerable damage on older leaves. ‘Pisang Kepok’ (ABB, Saba subgroup) is severely affected in this way in the wet lowlands of Papua, Indonesia (Plate 2.26) (J. Daniells, Australia, 2016, personal communication). The band pattern of symptoms on the leaf shown in Plate 2.27 indicates that asexual spores may have been carried in water running down the leaf when it was ­unfurling.

Causal agent Black cross leaf spot is caused by the fungus Phyllachora musicola, the black crosses or stars being the distinctive mature stroma of the fungus. On the upper leaf surface, the fungus produces oval to globose perithecia 200–240 μm in diameter, which are immersed in the stroma with the ostiole visible only as a minute pore. Up to 40 perithecia can be found in each stroma. Asci develop from the base of the perithecia among abundant paraphyses. Asci are cylindrical or clavate in shape with the widest part near the centre of the ascus; they measure 115–190 × 16–20 μm



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Plate 2.26.  Leaves of ‘Pisang Kepok’ (ABB, Saba subgroup) growing near Besum in Papua province, Indonesia have been destroyed as a result of Cordana leaf spot gaining entry to wounds caused by black leaf cross (photo: J. Daniells, QDAF).

and contain eight ascospores. The apex is truncate, with a non-amyloid apical ring. Ascospores, which are arranged biserrately in the ascus and surrounded by a deliquescing m ­ ucilaginous sheath, measure 35–52 × 6.5–10 μm, are hyaline, smooth, usually aseptate and obovate to clavate in shape (Booth and Shaw, 1961; Hyde, 1992). The fungus has not been grown in culture. An asexual state was not observed by Booth and Shaw (1961) or Hyde (1992), but Gerlach (1988) has implicated the genus Scolecobasidium,

which he found covering the stroma on the lower side of leaves as a velvety layer. However, Scolecobasidium humicola Barron & Busch has been recorded as growing over black cross fructifications in Papua New Guinea (Shaw, 1984).

Disease cycle and epidemiology White masses of spore exudates occur along the arms of the stroma in humid conditions (Booth

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Plate 2.27.  Symptoms of black cross leaf spot on the underside of a leaf of ‘Pisang Kepok’ (ABB, Saba subgroup) east of Jayapura in Papua province, Indonesia. The broad band of symptoms suggests that spores were carried in water running down the leaf when it was unfurling and erect (photo: J. Daniells, QDAF).

and Shaw, 1961). During wet weather, it is presumed that ascospores are carried in water drops and rain splash to spread the disease along and between leaves. It is also likely that ascospores can become airborne when forcibly ejected from perithecia to disseminate the disease more widely. Lines of black cross leaf spot symptoms have been seen on leaves, indicating that spores were carried in moisture running down the inside of a funnel leaf (Plate 2.27).

Host reaction Meredith (1969) claimed that Knowles (1916) working in Fiji was the first to recognize black cross leaf spot, because his description of the symptoms closely matched the disease attributed to P. musicola. Knowles (1916) stated that the black cross leaf spot was not found on ‘Dwarf Cavendish’ (AAA, Cavendish subgroup), ‘Gros

Michel’ (AAA) and other cultivars grown by ­Europeans, but did occur on ‘Mysore’ (AAB), ‘Blue Java’ (ABB, syn. ‘Ney Mannan’) and native cooking-banana cultivars. Meredith (1969) reported black cross leaf spot on members of the Mai’a Maoli–Popoulu subgroup (AAB), but not on ‘Robusta’ (AAA, Cavendish subgroup) in Fiji. Later, Firman (1972) took the opportunity to rate accessions growing at the Koronivia Research Station in Fiji for reaction to black cross leaf spot. Counting down non-flowering plants, symptoms were seen first on the third, fourth or fifth leaves, with the highest number of crosses being recorded on the fifth or sixth leaf. His findings were similar to the previous reports, which indicated that black cross leaf spot was a disease of cultivars in the AAB and ABB genomic groups. Black cross leaf spot was observed on a wide range of genotypes (AA, AAA, AAB, AAS, ABB and AAAB) in the Papua New Guinea Biological Foundation Banana Collection at Laloki near Port Moresby in 1988. Although the disease was present on 75% of accessions, which included those identified as belonging to the Cavendish subgroup and clones ‘Sucrier’ (AA), ‘Red’ (AAA), ‘Gros Michel’ (AAA) and ‘Pisang Awak’ (ABB), it was not significant on most cultivars. Very severe symptoms were only seen on seven local cultivars, which had been classified as ­belonging to the AAA, AAB, AAS and AAAB ­genotypes (D.R. Jones, Australia, 1988, personal observation). As well as Papua New Guinea, Cavendish cultivars, which were not seen with symptoms in Fiji (Knowles, 1916; Firman, 1972), were affected in the wet lowlands of ­Papua, Indonesia (J. Daniells, Australia, 2016, personal communication). Other records include ‘Lady Finger’ (AAB, Pome subgroup) and ‘Pacific Plantain’ (AAB, Mai’a Maoli-Popoulu subgroup) in the Torres Strait region of Australia (Jones and Daniells, 1991) and ‘Saba’ (ABB) in the Philippines (Meredith, 1969). Symptoms of black cross leaf spot have been seen on M. balbisiana and M. acuminata ssp. banksii in Papua New Guinea (Plate 2.25) and Queensland, Australia (D.R. Jones, Australia, 1988 and 1989, personal observations; Jones and Daniells, 1991). The presence of the disease on many local cultivars in Papua New Guinea and Fiji may be because their genomes are mostly derived from one or both of these two species (Carreel, 1995). No other wild Musa



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species in Papua New Guinea have been seen with symptoms and introduced M. textilis also seems to be unaffected (Booth and Shaw, 1961; D.R. Jones, Australia, 1988 and 1989, personal observation). Control Commercial cultivars of the AAA genotypes, although not immune, would seem to be fairly resistant to the disease in the Pacific. Control is not practised on other cultivars, but planting ‘Fa’i Misiluki’ in open, sunny places rather than in shade is recommended in Samoa (Gerlach, 1988).

Cordana Leaf Spot D.R. Jones Introduction Cordana leaf spot is usually of minor importance, but on occasions it can cause defoliation. Stover (1972) reported that it could be a serious problem in Central America on cultivars in the Plantain subgroup (AAB) during and following periods of wet weather. Epidemics have also occurred on ‘Williams’ (AAA, Cavendish subgroup) in the southernmost area of commercial banana production in New South Wales, Australia (Allen and Dettman, 1990). Most damage occurs when the pathogen gains entry to leaf tissue weakened because of age, adverse environmental conditions, nutritional deficiencies, wounds or through lesions caused by other disease organisms. Cordana leaf spot has a worldwide distribution.

Plate 2.28.  Symptoms of Cordana leaf spot on a leaf of Williams (AAA, Cavendish subgroup) growing in Queensland, Australia. The pathogen has invaded the leaf through lesions caused by Sigatoka leaf spot (photo: QDPI).

Invasion frequently occurs at leaf margins, where the lamina is more vulnerable to tearing and where the effects of senescence and nutritional deficiency problems are first felt. Symptoms are also often seen around lesions caused by other pathogens, such as Pseudocercospora ­fijiensis (see Plate 2.4), P. musae (Plate 2.28), P. eumusae (Plate 2.29) and Phyllochora musicola (Plate 2.26).

Symptoms Causal agents Large, pale brown, oval to fusiform-shaped necrotic lesions, often with light grey centres and concentric ring patterns, characterize Cordana leaf spot. A dark brown border surrounded by a bright yellow halo separates a lesion from normal leaf tissue (Plates 2.28 and 2.29). Often, lesions coalesce to form large areas of dead leaf tissue (Plate 2.29).

The fungus causing this distinctive leaf spot disease was first described as Scolecotrichum musae by Zimmerman (1902) from a specimen collected in Java, Indonesia. It was later transferred to Cordana by von Höhnel (1923). The species was widely distributed. A new species, Cordana johnstonii, which causes a disease very similar in

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A 10 μm

Plate 2.29.  Symptoms of Cordana leaf spot on ‘Novaria’, a radiation induced mutant of ‘Grand Nain (AAA, Cavendish subgroup), growing commercially in Perak State, West Malaysia. The pathogen has invaded through lesions caused by eumusae leaf spot (photo: D.R. Jones, INIBAP).

appearance to the one caused by C. musae, was described by Ellis (1971a) on banana from Papua in Indonesia. It was found to be present in the Cameron Highlands of West Malaysia, Philippines and Tonga. When herbarium specimens of Cordana leaf spot collected in New South Wales in Australia since the 1930s were examined, all were found to be of C. johnstonii. This species was also present on Lord Howe Island off the coast of New South Wales and on Norfolk Island between Australia and New Zealand. Further studies revealed that C. musae was the cause of most Cordana leaf spot in Queensland, which lies to the north of New South Wales. Cordana musae was also present at Darwin in the Northern Territory. In an area of Queensland close to the border with New South Wales, there was a zone where the distribution of both species overlapped (Priest, 1990). Priest (1990) reported that leaf spots caused by C. johnstonii were generally smaller, more regular in outline and more tapered than the larger elliptical to oval spots caused by C. musae. It was estimated that individual leaf lesions of C. johnstonii measure up to 3 cm × 1 cm, while those of C. musae are up to 7 cm × 2 cm. Both fungi had erect, straight to flexuous, septate, denticulate conidiophores, which arose singly in small groups. However, they differed in colour and size. Priest (1990) described C. musae as having pale brown conidiophores that are often nodose, up to 150 μm long and 4–6 μm in diameter. The brown conidiophores of C. johnstonii

B

Fig. 2.15.  Conidia of Neocordana musae (above) and N. johnstonii (below), showing shape and size differences (from Priest, 1990).

were often twisted in the basal part, up to 300 μm long and 5–9 μm in diameter. The conidiophores of both fungi are paler towards the apex (Priest, 1990). Cordana musae and C. johnstonii have conidia that are two-celled, smooth, pale brown to almost hyaline, with a visible thickened hylum. The conidia of C. musae are obclavate to pyriform and measure 14–18 × 8–10 μm on average, while those of C. johnstonii are broadly ellipsoidal to subglobose and measure 19–26 × 14–16 μm on average. Morphologically, the species can most easily be differentiated on conidial size and shape, C. johnstonii having longer and much wider conidia than C. musae (Fig. 2.15) (Priest, 1990).



Fungal Diseases of the Foliage

The two species responsible for Cordana leaf spot symptoms were accommodated in a new genus Neocordana as N. musae and N. johnstonii by Hernández-Restrepo et  al. (2015). These authors described the conidiophores of N. musae as brown, smooth and with dimensions of 46–118.5 x 5–6.5 μm and conidia with dimensions 14.5–19 x 8–11.5 μm. Sexual morphs are unknown in Neocordana. Hernández-Restrepo et  al. (2015) also described another closely related species, found on banana in Mauritius and Mexico, which was given the name Neocordana musicola. Conidiophores of N. musicola were up to 125 × 4–5.4 μm with pale brown conidia, oblong to obovoid, 1–septate with dimensions of 14.5–20 × 6.5–9.5 μm. Neocordana musicola could be differentiated from the other two species in the genus by their narrower conidia (Plate 2.30) (Hernández-Restrepo et al., 2015). Colonies of N. musae, N. johnstonii and N. musicola were effuse, grey to brown, hairy (Hernández-Restrepo et al., 2015). Crous et al. (2016) identified another Neocordana sp. causing a foliar disease on banana in La Réunion. Neocordana musarum resembled N. musae and N. musicola, but it had wider conidia. Based on a megablast search of the NCBIs nucleotide database using the ITS sequence, the highest similarities were with Neocordana musae and Neocordana musicola. Yet another Neocordana species found on banana in Morocco has been described by Crous and Groenewald (2017). Neocordana musigena had conidia with dimensions of 15–18 × 7–9 μm, which were most similar to those of N. musicola,

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but it was phylogenetically distinct from this species, Based on a megablast search using the ITS sequence of the ex-type culture, the best matches were to N. musae, N. musarum and N. musicola. Based on a megablast search using the rpb1 sequence of the ex-type culture, the strain was identical to N. musarum while the actA sequence was 99 % identical to N. musarum. To date, five Neocordana spp. have been identified that cause very similar lesions on banana. Disease cycle and epidemiology Conidia of N. musae are formed at night during rainy periods or when dew is present and violently discharged from the conidiophores just after dawn when there is a sharp decline in humidity (Meredith, 1962a, b). They germinate in a film of moisture on the leaf surface and within 8 h have formed appressoria. Each appressorium produces an infection peg, which penetrates an epidermal cell. One or more fungal cells are then produced, which almost completely fill the host cell. Epidermal cells that have been invaded colour and die as the infection peg swells. The fungus penetrates both living and dead tissue in the same manner. The rate of progress of the disease in living leaves seems to depend on the health of the tissue. Neocordana is a weak pathogen of banana and is primarily a wound invader (Stahel, 1934). The severe outbreaks of Cordana leaf spot that occur near Macksville in New South Wales are in localities with daily temperature ranges of

Plate 2.30.  Comparison of conidia of Neocordana musae (above) and N. musicola (below); scale bar = 10 μm. Images reproduced from Hernández-Restrepo et al., (2015) with permission from Magnolia Press.

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16–28°C in summer and 10–20°C in winter. This is relatively cold for banana cultivation and plants are extremely stressed during the winter months. In this area, N. johnstonii sporulates profusely on the adaxial surface of leaf spots in otherwise green leaves and, to a lesser extent, on spots in leaf litter, during cool, misty weather. Conidia germinate, produce germ tubes and appressoria and penetrate the leaf epidermis, as reported for N. musae by Stahel (1934). The time course for germination and infection was 6–48 h, depending on the temperature, and symptoms appeared after 4–10 days (Allen and Dettman, 1990). Laboratory experiments with leaf lesions detached from living leaves and dead leaf litter showed that new conidia are produced 1 day after wetting and, under moist conditions, this is continued at a constant rate for another 5 days at least. The optimum temperature for production of conidia was 22°C. Using detached banana leaves, it was found that most appressoria were produced at 22–25°C, but some were formed at temperatures as low as 10–13°C (Allen and Dettman, 1990).

mission to collect Musa germplasm in Papua New Guinea, 86% of cultivars collected were observed to have symptoms of the disease. Cordana leaf spot was also seen on a Fe’i banana cultivar (Jones, 1988).

Control Cordana leaf spot is usually of no great economic importance and does not warrant control. However, it can invade leaf tissue through lesions caused by other pathogens and this can lead to extensive necrosis. Control of the pathogens that allow Neocordana spp. entry into banana leaf tissue should control Cordana leaf spot. When the disease has become a problem, fungicides that control Sigatoka leaf spot and black leaf streak, such as mancozeb, propiconazole and pyremethanil, have also been found to be useful. Some work has been reported using biological control agents and plant extracts.

Deightoniella Leaf Spot Host reaction Cordana leaf spot has been reported on M. acuminata ssp. banksii, M. acuminata ssp. banksii × M. schizocarpa, M. balbisiana, Musa boman, M. maclayi, M. schizocarpa and Ensete glaucum in Papua New Guinea (Jones, 1988; Sharrock and Jones, 1989). Cordana leaf spot occurs on a wide range of cultivated banana clones. In addition to cultivars in the Cavendish and Plantain subgroups, typical symptoms have also been seen on ‘Sucrier’ (AA), ‘Inarnibal’ (AA), ‘Gros Michel’ (AAA), ‘Red’ and ‘Green Red’ (AAA), ‘Mysore’ (AAB), ‘Pome’ (AAB), ‘Silk’ (AAB), ‘Pisang Raja’ (AAB), ‘Bluggoe’ (ABB), ‘Pisang Awak’ (ABB), ‘Kluai Teparot’ (ABB) and ‘Saba’ (ABB) in Southeast Asia (Jones and Daniells, 1988; Jones, 1993b, 1994b). A disease survey of the Papua New Guinea Biological Foundation’s Banana Collection at Laloki near Port Moresby in 1988 revealed that 81% of the 238 accessions inspected were affected by Cordana leaf spot. However, only four accessions were very severely diseased. During a

D.R. Jones, E.O. Lomerio, M. Tessera and A.J. Quimio Introduction The fungus that causes Deightoniella leaf spot is a saprophytic colonizer of dead Musa leaves and flowers. It is also a weak parasite of the older foliage of banana and has been reported on young leaves of Musa seedlings (Stover, 1972) and young plants derived from tissue culture (L. Perez-Vicente, Cuba, 2017, personal communication). The leaf spot is more prevalent if plants are growing under poor conditions and humidity is high. Senescing or injured leaves are more prone to the disease. Deightoniella leaf spot, or black leaf spot as it has also been called (Koné et al., 2008), is more of a problem on abacá than on banana. When abacá was grown in Central America, as much as 14% of the total leaf area of ‘Bungolanon’, the most widely planted cultivar, was destroyed (Stover, 1972). However, this was not thought to affect plant growth (Lopez and Loegering, 1953).



Fungal Diseases of the Foliage

The disease has also been recorded on both young and old enset plants and is common in the Sidamo and North Omo regions of Ethiopia (Quimio and Tessera, 1996).

Symptoms Lesions first appear as small, tan to black necrotic spots between 1 and 2 mm in diameter. These increase in size, becoming oval in shape with a black border. Mature spots can be 25 mm or more in diameter and may unite to form bands of necrotic tissue along leaf edges. Deightoniella leaf spot can easily be confused with Cordana leaf spot, but there is a dark, smoky colouring over a tan background in Deightoniella spots (Stover, 1972). On banana, lesions are more prevalent along the edges of the leaf blade and on the older, lower leaves. Symptoms on young plants derived from tissue culture are shown in Plate 2.31. In Georgia, USA, most lesions were less than 10 mm in diameter and tan to black in colour. Larger oval lesions more than 20 mm across with black borders and yellow halos were also observed (Koné et al., 2008). Inoculation experiments on 3-month-old ‘Grande Naine’ plants in Cuba using mycelial fragments showed that symptoms appeared as small yellow spots on the leaf lamina. These then became brownish and finally black, becoming oval in shape with a black border. Lesions, which first appeared 14 days after

Plate 2.31.  Large lesions caused by Corynespora torulosa on leaves of young ‘Grand Nain’ (AAA, Cavendish subgroup) plants derived from tissue culture growing in a nursery in Guayaquil, Ecuador (photo: L. Pérez-Vincente, INISAV)

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inoculation, increased in size enlarging in both length and width. Blotches of various sizes were observed on inoculated leaves, which sometimes covered the whole length of the leaf border (Leiva-Mora et al., 2013). The causal agent can also cause a speckle-­ like spotting of the petiole and a pin-spotting disease of preharvest fruit (see Chapter 4). Deightoniella leaf spot of abacá first manifests itself as small pinpoints on the leaf lamina, which are yellowish at first, then brownish and finally black. Lesions increase in size, becoming greater in length than in width. Finally, blotches of various sizes form, which sometimes run the whole length of the leaf. Well-defined black bands, 1–2 mm in width, and bright yellow haloes surround lesions. Fully developed lesions become brown and dry, but the original small black spots remain distinct. As with banana, a speckle-­ like spotting can be seen on petioles. The causal agent also attacks the pseudostem of abacá. Symptoms on the leaves of enset are shown in Plate 2.32. Areas of dead tissue are evident in places where many lesions have coalesced. Causal agent Corynespora torulosa (Crous et  al., 2013) (syn. Deightoniella torulosa) causes Deightoniella leaf spot. Conidiophores of this fungus arise singly or in small groups. They are brown and swollen at the apex, with up to six successive proliferations. Conidia are produced singly as expanded ends of the conidiophores and their successive

Plate 2.32.  Lesions caused by Corynespora torulosa on a leaf of enset (M. Tessera and A.J. Quimio, IAR).

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Fig. 2.16.  Conidia and conidiophores of Corynespora torulosa (syn. Deightoniella torulosa) (from Ellis, 1957a). Conidia are usually 35–70 × 13.25 μm.

proliferations. They are 35–70 × 13.25 μm, straight or slightly curved, obpyriform to obclavate, subhyaline to olive in colour with three to 13 pseudosepta (Fig. 2.16). Deightoniella torulosa has been described by Ellis (1957a), Meredith (1961a, b) and Subramanian (1968). Information on the change of name from Deightoniella torulosa to Corynespora torulosa can be found in Crous et  al. (2013), which also describes Neodeightoniella phragmitiola.

Disease cycle and epidemiology Corynespora torulosa is present in dead banana leaves and inoculum is produced during periods of rain or dew (Meredith, 1961a). Spores are violently discharged, when the humidity declines, and become airborne (Meredith, 1961b). It has been reported that spores of C. torulosa are present in the air throughout the year in Jamaica (Meredith, 1961c). However, they are not transmitted long distances and viability is lost within 4 days at humidities less than 95% (Meredith, 1961a, b). Spores land on plants and germinate in water. Studies with abacá show that germ

tubes produce appressoria and leaves are directly infected through the epidermis. As the wall of the epidermis is pierced, the epidermal cell is immediately killed and the contents are transformed into a reddish granular substance. The infection tube swells up and fills the infected cell. Corynespora torulosa is commonly found associated with symptoms of black leaf streak diseases in Côte d’Ivoire. Here, it was observed that conidia germination began in less than 1 h and reached 95–100 % within 4 h. Penetration of stomata and pathogenic activity began the third to the fifth day after inoculation. The progress of the disease varied according to the banana cultivar and the amount of conidia. Experimentation showed that 105 spores/ml induced symptoms. It was concluded that C. torulosa alone could induce symptoms and was involved in rapid necrosis of infected leaves under natural conditions (Koné et al., 2007).

Host reaction The problem is mainly reported from cultivars of the Cavendish subgroup (AAA) growing under



Fungal Diseases of the Foliage

plantation conditions. Plantains are also affected (Perez et  al., 1989). ‘Sucrier’ (AA), ‘Grande Naine’ (AAA) and ‘Orishele’ (AAB, Plantain subgroup) were reported as susceptible after ­inoculation with conidia (Koné et al., 2007). In Georgia, USA, 16 of 34 banana cultivars were recorded as affected. ‘Dwarf Namwa’ (ABB, Pisang Awak subgroup) and ‘Dwarf Nino’ (AA, Sucrier subgroup) were infected by inoculation (Koné et al., 2008). The abaca cultivar ‘Bongalonan’ is more susceptible than ‘Libuton’, ‘Maguindanao’, ‘Tangongon’ and ‘Putian’. Most enset clones have been reported as susceptible.

Control No control measures are usually required on banana. However, the removal of dead and diseased leaves will reduce the amount of inoculum in the field. In vitro work in Australia has found that Trichoderma harzianum and T. virens are ­antagonistic against C. torulosa. The optimal food substrate for the Trichoderma species was found to be molasses at a concentration of 5%, which when used in combination with a di-1-p-­ menthene spreader-sticker, enhanced the survivability of Trichoderma populations under natural conditions. This formulation suppressed C. torulosa development under glasshouse conditions (Samuelian, 2016). It is recommended that only resistant abacá cultivars should be planted in areas where the disease causes problems. Other control measures include reducing planting densities, harvesting plants before rotting becomes severe and cutting down and burning seriously affected plants (Ela and San Juan, 1954).

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it only affects banana where Bermuda grass (Cynodon dactylon) is present beneath the banana canopy (Stover, 1972). Meredith (1963a) first reported the disease in Jamaica. Symptoms have been observed on cultivars in the Lujugira–­ Mutika subgroup in Uganda (Tushemereirwe et al., 1993). It was described as being most severe on the youngest leaves and becoming progressively less severe on the older leaves. However, it rarely affected more than 3% of the leaf area. Eyespot is common on enset seedlings and young transplants in the Sidamo and North Omo regions of Ethiopia (Tessera and Quimio, 1994). Older enset plants are not seriously affected. On banana, the first symptoms are minute, slightly sunken, reddish spots with a pale green or faintly yellow border. The spots become oval or lens-shaped in the direction of the leaf veins and the centre turns dark brown. The centre of the spot later dries to a bleached white or grey colour, with a narrow, well defined, dark brown margin and pale green or yellowish-green halo (Plate 2.33). Spots, which may be as large as 16 × 8 mm on leaves, also occur on the midrib and petiole (Stover, 1972). On enset, spots usually coalesce to form large blighted and dead ­areas (Plate 2.34). Severe blighting may be seen on the unfurling leaf and the first and second ­expanded leaves of succulent, rapidly growing plants. The causal agent of eyespot on banana has been identified as Drechslera gigantea. It is a pathogen that causes a zonate eyespot symptom on grasses (Drechsler, 1928, 1929). Sporulation

Eyespot D.R. Jones, M. Tessera and A.J. Quimio Eyespot, also known as Drechslera leaf spot, is a minor disease of banana suckers less than 2 m tall and occurs during periods of wet weather and heavy dew. In Central America and Jamaica,

Plate 2.33.  Symptoms of eyespot caused by Drechslera gigantea on the leaf of an East African highland banana cultivar (AAA, Lujugira-Mutika supgroup) near Kampala, Uganda (photo: D.R. Jones, INIBAP).

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p

elliptical lesions with pale brown to whitish-grey centres and light brown margins coalesced to form necrotic areas. When young plants of ‘Caipira’, ‘Prata Anã’ (AAB, Pome subgroup) and ‘Pacovã’ (AAB, Plantain subgroup) were inoculated with conidia, symptoms developed after 10 days only on ‘Caipira’. Bipolaris sacchari is an important sugarcane pathogen and is known to attack other monocotyledons. It was reported previously as a component of crown rot found on thiabendizole-treated fruit of ‘Robusta’ (AAA, Cavendish subgroup) from the Windward Islands (Wallbridge and Pinegar, 1975).

Malayan Leaf Spot D.R. Jones Plate 2.34.  Necrotic area on the first expanding leaf of enset caused by Drechslera gigantea (photo: M. Tessara and A.J. Quimio, IAR).

­ccurs on eyespot lesions of Bermuda grass o when the grass is wet. Conidia are forcibly expelled from the conidiophore as the humidity decreases and can be picked up by a Hirst spore trap between 8.00 and 14.00 h (especially after rain). Moisture on the leaf surface favours germination and infection. When conidia were placed on the heart leaves of banana suckers, reddish-brown spots appeared after 24 h at 26.7°C. After 5 days, these spots measured 4–5 mm, but no fructifications formed (Meredith, 1963a). The fungus causing leaf spot on enset has been shown to be a Drechslera species. It has been isolated and found to attack leaves of both enset and the banana cultivar ‘Dwarf Cavendish’ (AAA, Cavendish subgroup). Most cultivars of enset are susceptible. Early removal of diseased leaves has been recommended to minimize spread. The causal agent of another eyespot leaf disease of banana has been identified visually, but not by molecular methods, as Bipolaris sacchari (Silva et al., 2008). The pathogen caused a severe leaf spot on young plants of ‘Caipira’ (AAA, syn. ‘Yangambi Km 5’) in a plantation in the state of Minas Gervais in Brazil. Lesions with concentric zones, dark brown centres and dark brown margins surrounded by yellow haloes were seen on young leaves. On older leaves,

Introduction Malayan leaf spot, known also as diamond leaf spot, was first observed in Fiji by Knowles (1916) and was later described from leaves collected at altitude in the Cameron Highlands of West Malaysia (Ellis, 1957b). The disease has also been reported in Tonga and Samoa (CMI, 1990b) and is present in the highlands of Papua New Guinea, where some local cultivars are very susceptible (P. Kokoa, Papua New Guinea, 1988, personal communication; D.R. Jones, Australia, 1989, personal observation), and on Cavendish growing at altitude on the island of Mindanao in the Philippines (Y. Israeli, Israel, 2016, personal communication). The European and Mediterranean Plant Protection Organization (EPPO) database also lists American Samoa and Brunei. Malayan leaf spot has been reported as severe in Fiji during the cool season, particularly in the upper Waidina valley, where it can replace black leaf streak as the most significant problem (Firman, 1971).

Symptoms In Fiji, symptoms on the upper surface of the leaf appear as diamond-shaped, greyish-white spots with dimensions of 2–4 × 3–5.5 mm, the longer axis being parallel to the leaf veins. These



Fungal Diseases of the Foliage

spots, which sometimes have brown centres, are surrounded by a black border about 0.5 mm wide. On the undersurface, the lesion can be covered with a dense, velvety, brown mass (Knowles, 1916). From specimens collected in the highlands of West Malaysia, ellipsoid and round spots were described as well defined, white, grey or pale brown. The dimensions of the ellipsoid spots were recorded as 2–4 × 3–12 mm and those of the round spots as 2–5 mm in diameter. The spots were often very pale on the upper surface and darker on the lower, with dark purple borders (Ellis, 1957b). In the highlands of Papua New Guinea, round and ellipsoid-shaped lesions of various sizes, with well-defined dark brown borders and grey centres, are seen on the leaf lamina and midrib (Plate 2.35). Very few could be described as having a true diamond shape. When infection density is high, surrounding tissue yellows slightly and large necrotic areas develop. The dark brown borders and grey centres of lesions can still be distinguished in dead leaf tissue (Plate 2.35). First symptoms were seen by the author on the second and third fully expanded leaves on ‘Mala’, a highly susceptible local AA cultivar.

Causal agent The fungus Haplobasidion musae causes Malayan leaf spot (Ellis, 1957b; Ellis and Holliday, 1976).

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Microscopical examination of the velvety mass on the lesion on the undersurface of the leaf shows many conidiophores (Fig. 2.17). These arise singly or in groups of two to six at the ends of hyphae or as lateral branches. They emerge through the epidermal wall and cuticle and can be straight or flexuous. Condiophores are pale brown in colour, 0–3-septate, 50–110 × 4–6 μm in size, with an apical cell swollen at the end into a subglobose apical proliferation 9–12 μm in diameter. Spherical sporogenous cells, which are 4–8 μm in diameter and are at first hyaline and smooth and later brown and verrucose, are formed on the surface of each successive apical vesicle. Conidia are borne either singly or in simple or branched chains of two to five. Conidia are spherical, brown, verrucose and 4–6 μm in diameter (Ellis, 1957b).

Disease cycle and epidemiology In Fiji, shade and cool temperatures (monthly means below 23.8°C) favour Malayan leaf spot. The disease is only serious in areas with low sunshine hours, long periods of high humidity and high rainfall (> 2500 mm/year). Leaves seem more susceptible when plants are near to flowering. Plants flowering in the cool season suffer most (Firman, 1971). In West Malaysia, Malayan leaf spot is found at between 1372 and 1525 m (Firman, 1971) and in Papua New Guinea at well over 1000 m. In Samoa, Firman (1971) noticed that incidence increases with altitude. In Tonga, which has a cooler climate than Samoa, Malayan leaf spot occurs on banana growing at sea level. Not much is known about the disease cycle, but infection would seem to occur on young leaves soon after emergence (Ellis and Holliday, 1976).

Host reaction

Plate 2.35.  Symptoms of Malayan leaf spot on a leaf and midrib of ‘Mala’ (AA) in the Southern Highlands province of Papua New Guinea (photo: D.R. Jones, QDPI).

No studies have been undertaken on cultivar reaction, but clones in the Cavendish subgroup (AAA) have been reported as susceptible. Some cultivars, such as ‘Mala’ (AA), growing at altitude in Papua New Guinea, seem to be extremely susceptible.

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Fig. 2.17.  Conidia and conidiophores of Haplobasidion musae (× 500) (from Ellis, 1957b).

Control Leaves sprayed with oil seem more prone to Malayan leaf spot in Fiji and it has been suggested that this is because oil inhibits photosynthesis and transpiration, similar to when plants are shaded. Maneb applied in water delayed the appearance of the disease and symptoms were never severe (Firman, 1971).

Pestalotiopsis Leaf Spot D.R. Jones This disease was first described as naturally occurring on 2–6-month-old seedlings of M. acuminata ssp. banksii, M. acuminata ssp. zebrina and M. balbisiana in Honduras (Vakili, 1963). Pestalotiopsis palmarum, which has since been found to be a dominant endophyte on banana (Brown et al., 1998), was identified as the causal agent,

which formed brownish-yellow, circular spots between leaf veins. Spots developed until they reached vascular tissue, which then appeared as linear extensions of the lesion. Dimensions of 6-week-old spots were 1–4 × 0.3–0.9 cm. The fungus penetrated the epidermis of the Musa seedlings directly by means of appressoria and infection pegs. Growth was intracellular. Inoculum was believed to have originated from sporulating spots on Manaca palm (Orbignya cohune) used as shade. Sporulation did not occur on Musa even after 8 months (Vakili, 1963). Spotting attributed to Pestalotiopsis leprogena has occurred on wounded leaves of M. balbisiana and ‘Bluggoe’ (ABB, Bluggoe subgroup) in Jamaica (Meredith, 1963b) and Central America (Stover, 1972). Grey or fawn spots develop around abrasions or tears, which are usually found at the leaf margin. Delicate concentric zonation and acervuli of the pathogen are present on the upper leaf surface. Acervuli, which are sparse on the lower surface, increase in size and number if the leaf section is incubated for



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24 h. The centre of the spot is surrounded by a narrow, dark brown band and a conspicuous, bright orange or yellow halo. Crescentic areas of necrosis develop on one or both sides of the tear and decay may extend to the midrib. Conidia on M. balbisiana measure 17.5–24.5 × 6.0–7.5 μm and are four- to five-celled, spindle-shaped and slightly constricted at the septa. The basal cell is hyaline, long and cone-shaped with a short hyaline pedicel. The apical cell is also hyaline and cone-shaped and bears three knobless and widely spread setae (Meredith, 1963b; Stover, 1972). Pestalotiopsis menezesiana has been identified as causing a zonate leaf spot on plantain growing near Nanning in Guangxi Province in southern China. The fungus infected leaves at wound sites and initially formed water-soaked symptoms. These developed into brown-black lesions, which later turned into elliptical or irregularly shaped grey lesions with golden-­yellow margins. Acervuli arranged in characteristic concentric circles were frequently observed. ­Conidia, which measured 20–27.5 × 6.3–7.5 μm, were clavate-fusiform, straight or slightly curved, five-celled, slightly constricted at the septa. Colonies grew at 10–34°C with an optimum at 25°C. Both plantain and what are likely to have been Cavendish cultivars were infected by inoculation (Huang et al., 2007). Other Pestalotiopsis spp. recorded on banana are P. disseminata (Guba, 1961), P. versicolor (Prakash and Singh, 1975) and P. chethallensis (Sohi and Prakash, 1978). Current taxonomic knowledge on species in the genus Pestalotiopsis was reviewed by Maharachchikumbura et al. (2011). The authors noted that Pestalotiopsis spp. were taxonomically poorly understood and included endophytes, saprobes, parasymbionts, plant pathogens and animal pathogens. They stated that nomenclature of the genus was confusing and, because host ranges of some species could be extensive, host-based names in databases may be synonymous. It was concluded that isolates needed to be re-examined and sequence data obtained in order for a robust taxonomic system for the genus to be developed. Vakili (1963) considered P. palmarum and P. leprogena to be the same species. Future research into Pestalotiopsis may resolve this and other taxonomic issues.

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Phaeoseptoria Leaf Spot D.R. Jones Introduction Phaeoseptoria leaf spot was first described from Trivandrum, Kerala State, India, where it was the cause of a severe blight of banana leaves (Raghunath, 1963). The disease was not highlighted by Stover (1972), but Punithalingam (1983) reported that several samples of the disease had been received at the International Mycological Institute since the 1950s from different countries. It appears to be widely distributed with the host and probably occurs in most countries where banana is cultivated (Arzanlou and Crous, 2006). Phaeoseptoria leaf spot has been reported in Australasia–Oceania (Australia: Queensland), Asia (India, East Malaysia: Sabah), Africa (Cameroon, Ghana, Kenya, Mauritius, Tanzania: Zanzibar, Uganda) and the Latin American–­ Caribbean region (Colombia, Guyana, Honduras, Trinidad) (Punithalingam, 1983; Jones, 1991; B. Ritchie, UK, 1999, personal communication; Arzanlou and Crous, 2006).

Symptoms Individual lesions mature into ellipsoidal, sometimes ovoid spots. Raghunath (1963) gave the dimensions as 15 × 7 mm, while Punithalingam (1983) described them as 10–20 mm wide. Mature lesions are porcelain white (Raghunath, 1963) or straw yellow (Punithalingam, 1983) in the centre, with dark brown borders and yellow haloes. Often, infection density is high and spots coalesce as they enlarge to form irregular necrotic areas with white to straw-coloured centres. Young leaves are unaffected. Raghunath (1963), who published a photograph of mature lesions, thought that the symptoms of Phaeoseptoria leaf spot were reminiscent of Sigatoka leaf spot. Arzanlou and Crous (2006) described lesions as amphigenous, but more prominent on the upper leaf surface, irregular, elongated, pale brown to grey, surrounded by a dark brown margin and up to 6 mm wide. A photograph taken of Phaeoseptoria leaf spot some time after it was collected from a

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Plate 2.36.  Symptoms of Phaeoseptoria leaf spot on a banana leaf collected from a plant growing on a small plot in South Weipa, North Queensland, in April 1988. Scale bar at upper left = 10 mm (photo: S. Singh, DAFF).

diseased plant in northern Queensland in Australia shows developing lesions to be ovoid and coloured dark brown. As development proceeds, the centre of the lesion becomes straw-coloured leaving a dark brown border. Mature lesions, which are oval-shaped with grey centres and dark borders (Plate 2.36), resemble those of eumusae leaf spot. On another photograph (not shown), lesions appear to be coalescing in places, forming larger areas of necrotic leaf tissue. As no inoculation studies appear to have been undertaken to prove Koch’s postulates, it is not absolutely certain that the leaf spot described above is the symptom of the proposed causal agent.

Although Mycosphaerella was identified as a possible perfect stage by Raghunath (1963), this was not reported by Punithalingam (1976, 1983). A Leptosphaeria sp. was also believed to have been seen in association with one specimen of Ph. musae (G. Kinsey, UK, 1999, personal communication). However, Phaeosphaeriopsis was identified as the genus of the sexual morph by Arzanlou and Crous (2006). Pycnidia, which are immersed in leaf tissue and become erumpent, are found within the spots (Fig. 2.18). They are yellowish brown to dark brown, subglobose and 85–145 μm in diameter. Conidiogenous cells lining the pycnidial cavity are hyaline to pale brown, simple and flaskshaped to doliiform and proliferate percurrently (Fig. 2.18). They give rise to conidia that are generally pale brown, obclavate, apex subacutely rounded, base obconically subtruncate, smooth, 1–4-septate and (17–)20–30(–35) × (2–)3–4 μm in size (Fig 2.18). The ascomata are amphigenous, occurring separately, or on older lesions in association with other leaf spot pathogens. Ascomata are brown, up to 130 μm in diameter with a wall consisting of 2–3 layers of brown cells which is polygonal in cross-section. Asci are bitunicate, subcylindrical, stipitate, fasciculate, 8-spored, 38–60 × 7–8 μm. Pseudoparaphyses, which intermingle among asci, are hyaline, branched, septate and 1–2 μm in diameter. Ascospores are fusoid-ellipsoidal with obtuse ends, widest in the cell above the primary septum, 3-septate, brown, verruculose, guttulate, (15–)17–19(–21) × (3.5–)4 μm. They germinate predominantly from their polar ends with germinating spores distorting, becoming up to 8 μm wide. (Punithalingam, 1983; Arzanlou and Crous, 2006). On oat agar, colonies are floccose and ­greyish sepia with abundant mycelium (Punithalingam, 1976). On 2% malt extract agar, colonies are isabelline in colour with a sepia outer zone and sepia in reverse. After 1 month at 25°C, they reach a diameter of 30 mm and are generally flat, spreading, with sparse to moderate aerial mycelium and smooth, regular margins (Arzanlou and Crous, 2006),

Causal agent Disease cycle and epidemiology The fungus Phaeosphaeriopsis musae (syn. Phaeoseptoria musae) is reported as the cause of Phaeoseptoria leaf spot.

Little is known about the epidemiology of Phaeoseptoria leaf spot. It is assumed that conidia are



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C 10 μm

B 50 μm

A

10 μm

Fig. 2.18.  Vertical section of pycnidium (A), part of the pycnidial wall with conidiogenous cells and conidia (B) and conidia (C) of Phaeoseptoria musae (from Punithalingam, 1976).

dispersed by rain splash and ascospores are forcibly discharged during periods of wet weather. The pathogen is commonly isolated from leaf spots of banana, where it is assumed to be a weak pathogen or secondary invader. It has also been found in lesions associated with Pseudocercospora fijiensis, P. musae, P. eumusae, Sphaerulina musae and Neocordana musae (Arzanlou and Crous, 2006).

as highly susceptible and ‘Poovan’ (AAB, Mysore subgroup) as being affected. The cultivars ‘Annan’ (AB, Kunnan subgroup) and ‘Palayankodan’ (AAB, Mysore subgroup) were also reported as susceptible. No cultivar growing in the area was believed to be immune (Raghunath, 1963).

Host reaction

Control

In Kerala State in southern India, ‘Nendran’ (AAB, Plantain subgroup) has been described

No information on control is available. It is probably not warranted.

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Pyricularia Leaf Spot M. Tessera, A.J. Quimio and D.R. Jones This disease has been described on young banana plants derived from tissue culture. Pyricularia grisea, the causal agent of rice blast, which had previously been identified as the cause of a pitting disease of banana fruit, was believed to be the pathogen involved (Trindade et al., 2002). In September 2010, leaf symptoms were observed on young tissue cultured banana plantlets of ‘Dwarf Cavendish’ (AAA, Cavendish subgroup) at two nurseries in North Queensland, Australia. Symptoms included severe spotting on all leaves, death of some leaves and, in severe cases, death of the whole plant (Male et al., 2011). Lesions started as small, light tan-coloured spots 1 mm in diameter. These developed into circular or elliptical tan-coloured spots 3–4 mm in diameter with water-soaked or yellow margins (Plate 2.37). Mature lesions were brown in colour, 8–10 mm in diameter, with a slightly darker margin and zonate appearance (Plate 2.38). Adjacent lesions coalesced to produce large areas of necrotic tissue (Male et al., 2011). Similar fungal colonies were consistently recovered by Male et al. (2011) from symptomatic (a)

leaf tissue and identified on the basis of conidial morphology as belonging to Pyricularia sp. (Ellis, 1971a, b). Conidiophores were pale brown, darker towards the base, up to 120 μm long and 4 μm wide, sometimes branched, straight to flexuous. Conidiogenous cells were polyblastic, integrated, terminal, sympodial, geniculate, with denticles up to 1.5 μm long separated by a septum. Conidia were ovoid to obpyriform, 19–22 × 6.5– 8 μm, subhyaline to pale brown, smooth, thinwalled, 2-septate, hilum protuberant. The described fungus was found to be close to P. grisea. However, Male et al. (2011) identified the causal agent as Pyricularia angulata. They noted that Hashioka (1971) had observed morphological differences between three isolates of Pyricularia and reassigned P. grisea occurring on Musaceae to its own species, P. angulata. They also reported that Kim et al. (1987) found P. angulata to be pathogenic only on banana plants whereas P. grisea isolated from Digitaria sanguinalis was not. Kim et al. (1987) called the disease ‘banana blast’ and reported that it was frequently encountered in greenhouses of Cheju province in South Korea from 1985 to 1986 and incidence was severe. They reported symptoms of blast on banana leaves to be circular to oval, dark brown spots with yellow margins, which measured (b)

Plate 2.37.  Symptoms caused by Pyricularia angulata lesions on leaves of young plants of of ‘Dwarf Cavendish’ (AAA, Cavendish subgroup) derived from tissue culture: (a) early symptoms; (b) close-up of mature lesions. Images reproduced from Male et al. (2011) with permission from Springer.



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(a)

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(b)

Plate 2.38.  Pyricularia angulata: (a) denticulate conidiophores; (b) conidiophore bearing conidia; scale bars = 10 μm. Images reproduced from Male et al. (2011) with permission from Springer.

1–10 mm in diameter. The symptoms were scattered not only on leaves and fruits of banana, but also on petioles, leaf sheaths, bunch stalks and crowns. The optimum temperature range for mycelial growth of the fungus on PDA was 26–28°C. Molecular research had demonstrated that differences occur between P. angulata, P. grisea and 39 other Pyricularia species and allied genera (Bussaban et  al., 2005). This suggested to Male et  al. (2011) that the original identifications from Hoette (1936) and Meredith (1963) of P. grisea on banana fruit were most likely P. angulata. ‘Banana blast’ was found on young banana plantlets derived from tissue culture in nurseries during rainy season in Eastern India in 2014 to 2015. Taxonomical identification as well as DNA sequence analysis of the internal transcribed spacer region of the causal fungus isolated from affected leaves confirmed the pathogen to be Pyricularia angulata. The same species was found causing pitting disease on fruit in local plantations (Ganesan et al., 2017). It seems very likely that P. angulata is the cause of Pyricularia leaf spot of banana and pitting disease of bananas worldwide. P. angulata is a common inhabitant of hanging banana leaf trash, which acts as a course of inoculum. Pyricularia leaf spot disease is severe on young enset suckers and can lead to premature

b Plate 2.39.  Symptoms of Pyricularia leaf spot on the leaf and midrib of enset in Ethiopia (photo: M. Tessara and A.J. Quimio, IAR).

leaf death. It is common in the Sidamo and North Omo enset-growing areas of Ethiopia (Tessera and Quimio, 1994). Circular, oblong and spindle-shaped lesions with dark borders are found on leaves, midribs (Plate 2.39), petioles and leaf sheaths (Plate 2.40). These can coalesce to cause large areas of necrosis. A Pyricularia sp. isolated from lesions was shown

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c Plate 2.40.  Symptoms of Pyricularia leaf spot on the petioles and leaf sheaths of enset in Ethiopia (photo: M. Tessara and A.J. Quimio, IAR).

to be pathogenic on enset, but not on ‘Dwarf Cavendish’ (AAA, Cavendish subgroup) (Quimio and Tessera, 1996). Most enset cultivars appear to be susceptible, but older plants are not affected. The removal of affected leaves was advocated to minimize the spread of the disease in the sucker production field.

Speckles, Freckle And Rust D.R. Jones Cladosporium Leaf Speckle Introduction Cladosporium leaf speckle is usually found on the older leaves of banana plants growing in humid environments. Although it has been regarded as a minor problem, it can be serious on certain cultivars in certain locations. In West Africa, badly diseased leaves tend to dry out and fall prematurely (Frossard, 1963) and thus lower the photosynthetic ability of the plant. However, no serious attempts have been made to calculate

yield losses. Stover (1972) reported that in West Africa, injury is important only after the first crop following planting. Cladosporium leaf speckle has been reported in Australasia–Oceania (Australia, Papua New Guinea, Samoa, Solomon Islands), Asia (Bangladesh, Hong Kong, Indonesia, Malaysia, Nepal, Sri Lanka, Thailand, Vietnam), Africa (Burundi, Cameroon, Côte d’Ivoire, Democratic Republic of the Congo, Egypt, Ethiopia, Ghana, Guinea, Rwanda, Sierra Leone, South Africa, ­Sudan, Togo, Uganda, Zimbabwe) and the Latin American–­ Caribbean region (Cuba, Ecuador, Honduras, ­Jamaica) (Frossard, 1963; CMI, 1988; David, 1988; Sebasigari and Stover, 1988; Jones, 1993b, 1994b; D.R. Jones, England, 1999, personal observation; Surridge et al., 2003b). A Cladosporium leaf speckle disease has also been reported on enset in Ethiopia. It is said to be a minor problem of old plants in the Awasa Zuria, Sidamo and Welayita regions (Tessera and Quimio, 1994).

Symptoms In Côte d’Ivoire in West Africa, Frossard (1963) described the evolution of symptoms of Cladosporium leaf speckle on ‘Poyo’ (AAA, Cavendish subgroup) in five stages. The first stage was characterized by the appearance, 3–4 weeks after the youngest leaf unfurled, of small dashes resembling dark brown pencil lines measuring 1–5 × 0.5 mm. These lines ran parallel to the leaf veins. During the second stage, the lines elongated and enlarged to streaks measuring 5–0 × 1–2 mm. Lines and violet-brown spots were seen in the third stage, with the size of lesions increasing to 3 × 1.5 cm in the fourth stage. In the fifth stage, lesions coalesced and turned yellow-orange, later becoming a violet-black colour. Symptoms of the disease in Cameroon in West Africa are shown in Plate 2.41. Symptoms on East African highland cultivars growing in West Africa have been recognized as different to those found on clones of the same subgroup in East Africa (Pasberg-Gauhl and Gauhl, 2000). Leaf lesions in West Africa were noted to be diffuse with affected leaves remaining green for a long time. In East Africa, the lesions were more distinct in that they resembled



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spots and affected leaves turned yellow or orange and then brown more rapidly than in West Africa. (F. Gauhl, Austria, 2016, personal communication). These differences could be related to the causal agent involved. Symptoms of the disease in East Africa are shown in Plate 2.42. In South Africa, symptoms initially appear as pale green flecks that elongate into brown streaks

Plate 2.41.  Symptoms of Cladosporium leaf speckle on ‘Essong’ (AAB, Plantain subgroup) in Cameroon. Dark brown lesions of black leaf streak in various stages of development, the more mature ones surrounded by yellowing tissue, are also present (photo: C. Pasberg Gauhl and F. Gauhl, IITA).

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of about 2 cm in length or longer. Leaf streaks frequently turn orange in colour, with sparse grey-green blotching becoming evident on the upper surface of older leaves. Lesions eventually become dark brown, coalesce, and occupy large areas of the photosynthetic leaf surface (Surridge et al., 2003b). From their description, these symptoms would appear to be similar to those shown in Plate 2.42. Stover (1972) in Central America reported that the first visible symptoms consist of diffuse greyish-brown blotching on the upper surface of the oldest leaves (streaks symptoms are not mentioned). Later on, the spots may become orange-­ yellow and then brown and necrotic. Cladosporium leaf speckle symptoms were seen by the author in Malaysia and Thailand in 1990s. Early symptoms were small spots, which elongated into grey and later brown streaks. Coalescing lesions later changed colour to dark brown or violet-black before the leaf tissue was killed (Plate 2.43). Whole leaves could eventually turn necrotic. Lesions were also observed on leaf midribs. On enset, the disease affects the middle and lower leaves (Plate 2.44). When spotting is severe, large areas of dead tissue can be seen along leaf margins. Causal agents The pathogen causing Cladosporium leaf speckle of banana was initially identified as Cladosporium

Plate 2.42.  Symptoms of Cladosporium leaf speckle on an East African highland banana cultivar (AAA, Lujugira–Mutika subgroup) in the Bushenyi district of Uganda (photo: D.R. Jones, INIBAP).

Plate 2.43.  Symptoms of Cladosporium leaf speckle on ‘Kluai Khai’ (AA, Sucrier subgroup) in a smallholding near Kamphaeng Phet, Thailand. High infection densities cause diseased leaf tissue to eventually turn necrotic (photo: D.R. Jones, INIBAP).

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Plate 2.44.  Symptoms of Cladosporium leaf spot on an enset leaf in Ethiopia (photo: M. Tessara and A.J. Quimio, IAR).

musae (Martyn, 1945) and later as Periconiella sapientumicola (Siboe, 1994). More recently, DNA sequence data derived from the ITS and LSU gene regions of isolates show that that the fungus is part of a large group of hyphomycetes in the Chaetothyriales with dematiaceous blastoconidia in acropetal chains. The genus Metulocladosporiella gen. nov. has been proposed to accommodate the pathogen (Crous et al., 2006). Specimens of the Cladosporium leaf speckle fungus from Honduras and Cameroon were genetically analysed by Crous et  al. (2006) and determined to be Me. musae. In addition, specimens they examined from Jamaica and Mexico were found to be the same species. Another species of Metulocladosporiella has been recognized on banana in eastern and southern Africa and given the name Me. musicola (Crous et  al., 2006).This fungus has been identified by DNA analysis on specimens from Kenya, Mozambique, South Africa, Uganda, Zimbabwe and also Queensland (Y.P. Tan, Australia, 2017, personal communication). One of the cultures of an isolate from South Africa analysed by Surridge et al. (2003b) and identified as Me. musae was reported by Crous et al. (2006) to be Me. musicola. Symptoms caused by both Me. musae and Me. musicola are reported as similar (Crous et al., 2006) and it is likely that both have been recorded on visual evidence as Cladosporium leaf speckle caused by Cladosporium musae in the past. More work is now needed to accurately determine the distribution of these two pathogens. The evidence available today suggests that Me. musae is widespread and present in

Africa, the Americas, Asia and Oceania while Me. musicola is present in East Africa, South ­Africa and ­Australia. Mason (in Martyn, 1945) described Me. musae has having long conidiophores, 60–500 × 3.5– 6 μm, and aseptate conidia, 6–22 × 2.5–4 μm. Crous et  al. (2006) described the macronematous conidiophores rising from superficial hyphae as erect, solitary to loosely aggregated and 45–500(–600) μm long. They were medium to dark brown in their lower halves and paler towards their apices. Primary branches were 15–30 × 3.5–5 μm, giving rise to secondary branches or conidiogenous cells. Ellipsoid-ovoid, fusiform or subcylindrical conidia were in branched acropetal chains with dimensions (6–) 8–11(–16) × (3–)4(–5) μm (Fig. 2.19). Colonies growing on potato dextrose agar have smooth, regular margins and sparse aerial mycelium. Their surfaces are pale mouse-grey to mousegrey due to profuse sporulation. The margins of the submerged mycelium are mouse-grey with the reverse greenish-black (Crous et al., 2006). Crous et al. (2006) described the macronematous conidiophores of Me. musicola as arising from superficial hyphae, erect, solitary to loosely aggregated and 80–600(–700) μm long. They were also medium to dark brown in their lower halves and paler towards their apices. Apices were branched, with the branched part composed of usually fairly compact, closely arranged subcylindrical branchlets. Primary branches were 15–85 × 3.5–6 μm, giving rise to 1–3 secondary branches or to conidiogenous cells. Ellipsoid-ovoid, fusiform or subcylindrical conidia were in branched acropetal chains with dimensions of (9–)11–13(–16) × (3.5–)4(–5) μm (Fig. 2.20). Colonies on potato dextrose agar also have smooth, regular margins and sparse aerial mycelium. Centres are darker than the margins because of grey–white aerial mycelium. The colony surface on potato dextrose agar is pale mouse-grey to dirty white–grey at the centre, with margins leaden-black to olivaceous–grey (Crous et al., 2006). Metulocladosporiella musicola can be distinguished from Me. musae by its conidiophores, which are more frequently branched in their apical region with more secondary branches and conidiogenous cells. Its conidia are also longer and wider. Metulocladosporiella musae is more prone to form micronematous conidiophores in culture than Me. musicola. In addition, the scars



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Fig. 2.19.  Micro- and macro-nematous conidiophores and conidia of Metulocladosporiella musae; scale bar = 10 mm; reproduced from Crous et al. (2006) with permission from Elsevier.

on its conidia are barely thickened and only somewhat refractive, while those of Me. musicola are more prominent. Another four Metulocladosporiella spp. infecting banana in Southeast Asia have been recently identified (Marin-Felix et  al., 2018). Metulocladosporiella chiangmaiensis was found in Chiang Mai province, Thailand, Me. malaysiana in Malaysia, Me. musigena in Chiang Mai province,

Thailand and Me. samutensis in Samut Songkhram province, Thailand. Metulocladosporiella samutensis can easily be distinguished from other species of Metulocladosporiella by the production of conidiogenous cells directly from the apex, or loosely arranged primary branches, being almost totally absent of secondary branches. Full descriptions of thse pathogens can be found in Martin-Felix et al. (2018).

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Fig. 2.20.  Conidiophores and conidia of Metulocladosporiella musicola; scale bar = 10 mm; reproduced from Crous et al. (2006) with permission from Elsevier.

The pathogen affecting enset in Ethiopia has been identified as a Cladosporium species. Crous et al. (2006) reported that Me. musae had been found on Ensete gillesii, which is native to West-Central Africa. Disease cycle and epidemiology Conidia, which are carried by air currents, germinate in moisture on leaf tissue. The development

of the disease is favoured by high humidity. Brun (1954) and Frossard (1963) observed that disease peaks in Guinea and Côte d’Ivoire coincided with the beginning and end of the wet season. Frossard (1963) found that cultures obtained from isolates in Côte d’Ivoire grew best at around 25°C with good development between 20°C and 30°C. Growth was weak at 32°C and non-existent at 35°C. Stover (1972) indicated that populations of banana plants of 2000 or more/ hectare in Côte d’Ivoire favour Cladosporium



Fungal Diseases of the Foliage

leaf speckle infection by maintaining high humidity within the plantation. Koné et al. (2006) in Côte d’Ivoire reported that germination of conidia of the Cladosporium leaf speckle fungus was quicker than that of conidia of Pseudocercospora fijiensis. Observations revealed that stomatal penetration and the time to the formation of conidia in the lesion were also quicker. Given the present knowledge on the distribution of Metulocladospriella spp., it is likely that Brun (1954), Frossard (1963) and Kone et  al., (2006) were undertaking research with Me. musae. In Uganda, Cladosporium leaf speckle, which is likely to have been caused by Me. musicola, has been found at altitudes of up to 1850 m. It was described as being the most damaging leaf pathogen above 1600 m (Tushemereirwe et al., 1993). Holderness et  al. (1999) reported that the disease occurred together with Sigatoka leaf spot at higher altitudes where the mean minimum temperature was less than 15°C, conditions unsuitable for black leaf streak. In lower-­altitude zones, Cladosporium leaf speckle was reported to cause slightly more damage than Sigatoka leaf spots. In Kenya, Cladosporium leaf speckle is more frequent in highland areas (Kung’U et al., 1992). A fungus identified as Cladosporium musae in Malaysia has been found to grow in culture between a temperature range of 18°C to 30°C with an optimum of 22°C for mycelial growth and 26°C for sporulation. Conidial germination was optimal at 26°C at a relative humidity of 99–100% (Sahlan, 2003). After 14 days on potato dextrose agar under near-visible ultraviolet light at 25°C, colonies of Me. musae attained a diameter of 37–50 mm while colonies of Me. musicola were 20–30 mm in diameter (Crous et al., 2006). It is possible that Me. musae and Me. musicola have different growth temperatures ranges, with Me. musicola more suited to cooler environments. Host reaction The disease was reported as significant on plant crops of ‘Petite Naine’, ‘Grande Naine’, ‘Poyo’ and ‘Lacatan’ (AAA, Cavendish subgroup) grown on commercial, export plantations in West ­Africa (Frossard, 1963). At the beginning of the rainy season, ‘Gros Michel’ (AAA) was less

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s­ usceptible in the banana collection of the IFAC Station d’Azaguié, Côte d’Ivoire, and cultivars in the AB, AAB and ABB genomic groups were not regarded as susceptible. Only on AAB plantains ‘Plantain Vert’ and ‘Njock Kon’ and ‘Popoulou Popoulou’ (AAB, Mai’a Maoli-Popoulu subgroup) were some symptoms observed. In addition, a Cladosporium sp. was found on leaves of the local plantain cultivar ‘Corne’. ‘Figue Sucrée (AA, Sucrier subgroup) was observed to be resistant. No symptoms were seen on ‘Figue Rose’ (AAA, Red subgroup) and the wild species M. acuminata, M. balbisiana and M. textilis (Frossard, 1963). Some lesions were seen on ‘I.C.2’ (AAAA, bred Jamaican hybrid). Koné et al. (2006), working at the same research station as Frossard (1963), observed no development of symptoms on ‘Orishele’ (AAB, Plantain subgroup), ‘Corne 1’ (AAB, Plantain subgroup) and ‘Grande Naine’ (AAA, Cavendish subgroup) after the appearance of first-stage dashes. These first symptoms of Cladosporium leaf speckle developed on leaves 1–3, counting down from the first fully unfurled leaf at the top of the plant. In Cameroon in April/May 1996, Cladosporium symptoms were observed on East African highland banana cultivars, but not on ‘False Horn’ plantain (AAB, Plantain subgroup) or ‘Valery’ (AAA, Cavendish subgroup) in the Centre Regional Bananier et Plantain at Njombe at 80 m above sea level. In contrast, at the same time, symptoms were seen on ‘French’ and ‘False Horn’ cultivars (AAB, Plantain subgroup) at an elevation of about 650 m at IITA’s M’Balmayo Station in Cameroon (see Plate 2.41) (Pasberg-­ Gauhl and Gauhl, 2000). In the humid tropical lowlands of Nigeria, the youngest leaves with first symptoms and mature symptoms counting down from the first fully unfurled leaf on 19 East African highland cultivars were recorded during an epidemic of the disease in September 1993. Average first-­ symptom leaf readings ranged from 2.2 to 5.5 and the average mature-symptom leaf readings from 4.2 to over 7.8 on the East African highland cultivars, while no symptoms at all were seen on controls ‘Valery’ (AAA, Cavendish subgroup) and ‘Agbagba’ (AAB, Plantain subgroup) (Pasberg-Gauhl and Gauhl, 2000). The authors noted that although black leaf streak, which was also present, established itself more quickly, the amount of leaf area affected by both diseases

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was approximately the same and resulted in a significant reduction in the number of leaves. Under unfavourable circumstances for symptom development, leaves of ‘Valery’ banana and ‘Agbagba’ plantain may have died before advanced, easily visible symptoms of the disease developed. The West African location of the work reported above suggests that the pathogen was likely to have been Me musae. East African highland cultivars (AAA, Lujugira–Mutika subgroup) are affected by Cladosporium leaf speckle in East Africa (Plate 2.42) (Sebasigari and Stover, 1988). In Uganda, it was reported as the second most important leaf disease on East African highland cultivars after black leaf streak and a cause for concern (Holderness et al., 1999). Plantain and sweet banana cultivars have been reported as not affected by ‘streaking’ symptoms caused by Cladosporium leaf speckle, though it is unclear whether they are resistant or not (Tushemereirwe et al., 1993). However, the work of Surridge et al., (2003b) indicate that Cavendish cultivars were hosts in South Africa. In this region, Me. musicola is found (Crous et al., 2006). ‘Kluai Khai’ (AA, Sucrier subgroup) is thought to be the most susceptible clone in Thailand. Damage to leaves is substantial between August and November during the rainy season (Plate 2.43) (Jones, 1994b). ‘Pisang Mas’ (AA, Sucrier subgroup) is also reported as being particularly susceptible in Perak State in Malaysia (Sahlan, 2003). In addition, Cladosporium leaf speckle symptoms have been seen by the author on ‘Orito’ (AA, Sucrier subgroup) in Ecuador. The species of Metulocladosporiella at these locations was not known. ‘Pisang Berangan’ (AAA, Lakatan subgroup) is severely affected in Perak State in Malaysia, where symptoms were observed on young as well as old leaves (Jones 1993b; Sahlan, 2003). ‘Pisang Nangka’ (AAA) has also been seen with symptoms of Cladosporium leaf speckle in Malaysia (Jones, 1993b). Again, the species of Metulocladosporiella involved was not known. Differences in symptoms reaction may relate to the distribution of the species of pathogen in a particular region or country and this may be related to temperature, which in turn may be determined by altitude and latitude. From today’s evidence, the pathogen present in the lowlands of West Africa is likely to be Me. musae. The reason

‘Valery’ was not affected in Nigeria if Cavendish cultivars are severely attacked in Côte d’Ivoire may be related to experimental conditions, such as density of plantings, which would affect ­under-canopy humidity. There may also be local variations in inoculum pressure. More work needs to be undertaken to determine the temperature ranges for growth of the pathogens in culture followed by inoculation studies on various banana clones, such as those in the Sucrier, Plantain, Cavendish and Lujugira–­ Mutika subgroups, in order to clarify the Musa hosts of the six Metulocladosporiella species identified to date. Most enset cultivars in Ethiopia are susceptible to a Cladosporium leaf speckle. The identity of the pathogen is not known.

Control Fungicides used to control Sigatoka leaf spot and black leaf streak are believed to control Cladosporium leaf speckle on banana. In Thailand, the disease was controlled by applying benomyl every 2 weeks during the rainy season at a rate of 0.5 g/l. The early removal of diseased enset leaves is recommended in Ethiopia.

Acknowledgements Cornelia and Friedhelm Gauhl are thanked for useful advice concerning the reasons for the variability in symptom expression on certain cultivars in some countries. The information provided by M. Tessera and A.J. Quimio on Cladosporium leaf speckle affecting enset in Ethiopia is gratefully acknowledged.

Mycosphaerella Leaf Speckle Introduction Mycosphaerella leaf speckle is usually a minor disease of banana leaves. It is found worldwide (Pont, 1960b; Stover, 1969), but has been reported as a problem only in subtropical areas where it is a common cause of leaf death. The



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second most prevalent disease on banana in South Africa was reported to be Mycosphaerella leaf speckle, which in some cases caused severe symptoms resulting in leaf death (Surridge et al., 2003a). The disease has also been recorded on abacá (Anunciado et al., 1977).

Symptoms The first symptoms, which are rarely seen above the fifth fully opened leaf, are water-soaked patches, which exude moisture droplets. These become visible during rain or early in the morning when dew is present. In the absence of moisture, affected areas are first noticeable on the lower leaf surface as irregular light brown or tan-coloured blotches. These may show on the upper surface as smoky patches. Blotches on the lower surface darken in colour and eventually become dark purple to black, irregularly shaped, speckled areas, which are visible on both leaf surfaces. At this stage of disease development, leaf tissue in and around the speckled areas is yellow (Plates 2.45 and 2.46). Later, necrotic tissue within the speckled areas coalesces and bleaches with age to become grey on the lower surface and straw-­coloured on the upper as it dries out. The extensive death of leaf tissue is not often seen above the eighth leaf of an actively growing plant. However, when leaf production ceases with bunch emergence, extensive defoliation can occur before the fruit is ready for harvest (Pont, 1960b).

Causal agent Mycosphaerella leaf speckle has been reported to be caused by Mycosphaerella musae. The perithecia, which measure 45–99 × 34–81 μm, are highly scattered, globose, immersed in leaf tissue and black in colour. Asci, which have dimensions of 24–44 × 8–12 μm are eight-spored and obclavate in shape. Ascospores measure 9–16 × 2–3 μm and are hyaline, obtuse to cylindrical and two-celled, with one cell broader than the other. No conidial stage was observed in vivo on banana leaves. On potato dextrose agar, colonies consist of a thin layer of compact, light grey aerial mycelium

Plate 2.45.  Symptoms of Mycosphaerella leaf speckle on the upper leaf surface of a cultivar in the Cavendish subgroup (AAA) in Vinh Phu province, Vietnam (photo: D.R. Jones, INIBAP).

on a hard, black, sclerotium-like hump. The outline of colonies becomes more irregular with age and droplets of dark brown or black fluid may exude from parts of its elevated surface. The surface mycelium later turns pink. No fruiting bodies have been detected on potato dextrose agar. The optimum temperature for growth of mycelium is 26°C, but the fungus grows well between 20° and 30°C (Pont, 1960b). Stover (1994) reported that conidia of what he called ‘Cercospora non-virulentum’ were associated with cultures derived from ascospores of M. musae. These conidia had dimensions of 55– 200 × 2.6–3.2 μm (average 127 × 2.9 μm) and were usually verrucose, with a basal scar. Conidiophores were 24–46 μm long (average 35 μm) with one septum. Arzanlou et al. (2008) believed that the conidia were of the Stenella-type, but found that Stover’s description did not match any of the three Stenella species they found associated with Mycosphearella leaf speckle-like

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Plate 2.46.  Close-up of symptoms of Mycosphaerella leaf speckle on the upper leaf surface of a cultivar in the Cavendish subgroup (AAA) in southern Queensland, Australia (photo: QDPI).

symptoms. Arzanlou et al. (2008) suggested that isolations need to be made from new collections of leaf speckle in order to resolve its causal agent or agents.

Other Mycosphaerella-like fungi from banana Numerous fungi with Mycosphaerella-like sexual morphs have been recorded on banana. These include the well-known Sigatoka leaf spot pathogens Pseudocercospora fijiensis, P. musae and P.  eumusae featured at the beginning of this chapter and also the leaf speckle pathogen Mycosphaerella musae profiled here. Other recently described fungi with a Mycosphaerella-like sexual stage from banana are Dissoconium musae, Mycosphaerella mozambica, Pseudocercospora assamensis, P. indonesiana, P. longispora, Stenella musae, S.  musicola and S. queenslandica. Cercospora apii,

Mycosphaerella citri, M. communis, M. lateralis, M. thailandica and Passalora loranthi have also been found on other hosts besides banana (Crous et al., 2003; Arzanlou et al., 2008). The author collected the type specimen of what has now been identified as Pseudocercospora longispora from ‘Pisang Mas’ (AA, Sucrier subgroup) growing in a plantation near Melaka in West Malaysia in 1988. Symptoms on photographs taken at the time closely resembled those of the important Sigatoka leaf spots caused by P. fijiensis, P. musae and P. eumusae. This suggests other serious leaf spots in the Pseudocercospora genus may exist as yet undetected in Southeast Asia and could emerge in the future. Little information is available on the symptoms caused by the other Mycosphaerella-like fungi from banana other than the Stenella species, which were isolated from Mycosphaerella leaf speckle-like lesions. Mycosphaerella mozambique from Mozambique was reported to cause leaf spots, which were irregular to subcircular, 1–7 mm in diameter, grey to pale brown on the adaxial surface and grey on abaxial surface, with dark brown margins (Arzanlou et al., 2008). Some of the fungi listed above have recently been reassigned to new genera based on the results of genetic sequencing (Videira et al., 2017). Dissoconium musae is now Uwebraunia musae; an isolate of Mycosphaerella mozambica from Australia is Amycosphaerella keniensis and one from Mozambique is Rachiosphaerella mozambica; Stenella musae is Zasmidium musae; Stenella musicola is Zasmidiun musicola; and Stenella queenslandica is Zasmidium queenslandicum. Other species described by Videira et  al. (2017) from banana are Pseudocercospora marksii and a Paramycosphaerella sp.

Disease cycle and epidemiology Perithecia develop in necrotic leaf tissue as it dries out. They are numerous on the lower leaf surface, but sparse on the upper surface. Ascospores discharge from perithecia when leaves become wet. Germ tubes grow out of the ends of each cell of the ascospores, often simultaneously. The germ tubes give rise to an extensive, branched, epiphytic mycelium. After about 5–6 weeks, lobed appressoria are formed on or near stomatal



Fungal Diseases of the Foliage

pores on the undersides of leaves and guard cells turn light brown. After penetration of the stomata, coarse intercellular hyphae up to 3 μm thick, with finger-like lateral branches, grow through the leaf tissue, killing cells of the spongy parenchyma. The success in control when protectant fungicides are sprayed on the lower surfaces of leaves 4–6 indicates that infection does not take place on young leaves. High humidity favours disease development. Symptoms can appear within 45 days if inoculated plants are held in a saturated atmosphere during the night. Without saturation, the incubation period is 80–102 days. Speckle is ­always more prevalent in sheltered locations, ­hollows or wet spots in a plantation. Disease development is more rapid in senescing or injured leaves. Advanced stages of the disease are usually seen only on older leaves. ­Temperatures below 20°C retard the development of disease, but cooler weather also slows plant growth. As a consequence, speckle is more obvious in winter in subtropical areas, because leaf production does not keep pace with symptom expression, as it does in summer (Pont, 1960b). Stover (1994) has shown that, in tropical America, M. musae is found in association with early streak lesions of both Sigatoka leaf spot and black leaf streak.

Host reaction In Australia, cultivars ‘Mons Mari’ and ‘Williams’ (AAA, Cavendish subgroup), ‘Lady Finger’ (AAB, Pome subgroup), ‘Sugar’ (AAB, Silk subgroup), ‘Ducasse’ (ABB, Pisang Awak subgroup) and ‘T8’ (syn. ‘61–882–1’, AAAA bred Jamaican tetraploid) are susceptible. The disease is also found on the wild banana M. acuminata ssp. banksii (Pont, 1960b) and M. textilis has also been reported as a host (Crous et al., 2003).

Control In the past, copper oxychloride and the dithiocarbomate fungicide zineb gave good control when applied to the underside of leaves (Stover, 1972). Today, fungicides that are applied to control Sigatoka leaf spot disease (mancozeb,

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propiconazole) give adequate control of Mycosphaerella leaf speckle. Removing leaves killed by Mycosphaerella leaf speckle can reduce inoculum levels. Since 2000, Mycosphaerella leaf speckle has become more severe in South Africa and is now the main leaf disease of banana. Necrosis was usually observed only on the older leaves of bunched plants, but associated chlorosis was seen on younger leaves in southern KwaZulu-­ Natal in 2002. Control strategies in South Africa are similar to those used to control Sigatoka leaf spot disease and include removing leaves and applying fungicides (Viljoen et al., 2003).

Taiwan Leaf Speckle Introduction This leaf speckle, named Taiwan leaf speckle in this publication to distinguish it from other speckle diseases of banana, is a minor problem in Southeast Asia, but it has been described as significant on commercially grown cultivars in the Cavendish subgroup (AAA) in Taiwan since 1981 (Hwang and Chen, 1986; Hwang, 1991). Symptoms of Taiwan leaf speckle have also been recognized in Australia, Malaysia, Thailand and Vietnam. Cultivars in the Pisang Awak subgroup are commonly infected in areas where the pathogen occurs. Indeed, the characteristic symptoms of the disease have been associated with the cultivar ‘Ducasse’ (ABB, Pisang Awak subgroup) in North Queensland in Australia (Shivas et al., 2011).

Symptoms On ‘Ducasse’ (ABB, Pisang Awak subgroup) in Australia, symptoms are described as visible on both sides of the leaf and initially appearing as minute linear streaks with dimensions up to 3 × 0.2 mm with up to 30 discrete streaks/cm2. These streaks were black on the upper surface and greyish black on the lower surface, eventually coalescing to form irregular necrotic lesions over the entire leaf surface (Shivas et al., 2011). Plate 2.47 shows dark brown to black streaks in various stages of elongation and

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Plate 2.47.  Leaf symptoms caused by Rami­ chloridium ducassei on ‘Kluai Namwa Khom’ (ABB, Pisang Awak subgroup) in North Queensland, Australia (photo: D.R. Jones, INIBAP).

Plate 2.48.  Dark brown to black streak symptoms of Taiwan leaf speckle on a leaf and midrib of ‘Chuõí Bom’ (AAA, Lakatan subgroup) in Vinh Phu province, Vietnam (photo: D.R. Jones, INIBAP).

development on ‘Kluai Namwa Khom’ (ABB, Pisang Awak subgroup) in North Queensland, Australia. Where there are concentraions of mature streaks, the leaf looks necrotic surrounded by a yellow halo. Symptoms believed to be those of Taiwan leaf speckle on ‘Chuõí Bom’ (AA Lakatan subgroup) in Vietnam are shown in Plate 2.48. Coalescing black streaks are visible on the lamina and also the midrib as in Plate 2.47.

Causal agent The fungal pathogen was identified as Acrodontium simplex in Taiwan (Hwang and Chen, 1986), but the disease is now described as being caused by Ramichloridium ducassei (Shivas et al., 2011; Kirschner and Piepenbring, 2014).

Erect, branched conidiophores (75–)100– 140(–162) μm long arise in fascicles of 7–20 from the stromata on the abaxial side of the leaf. Basal cells are golden to medium brown with upper cells predominantly brown becoming paler at the apex. The main axis with 4–10 septa branches is 3–4 μm wide at the base narrowing to 2 μm towards the tip with the uppermost top 1–1.5 μm. Branches, which arise from nodes at the top of a cell, are found at irregular intervals and have 1–2 cells. Conidiogenous cells at the end of branches are slightly subulate, straight, pale yellow-brown, smooth, (14–)19–31(–40) × 1–2 μm. Conidia are pale brown in colour, solitary, obovoid, oblong or short-cylindrical, smooth with a broadly rounded apex and narrowing base. Their dimensions are (3–)3.5–5.5(–8) × 2–2.5 μm with a darkened, slightly thickened basal hilum (Fig. 2.21) (Kirschner and Piepenbring, 2014). Colonies on potato dextrose agar are slow growing, obtaining a radius of 5–6 mm after 15 days at 25°C. They are dark, velvety, raised and radially furrowed. At the centre they are white with a slight reddish tinge. Colony margins are distinct and irregular with diffuse pale grey spots. The reverse side is a dark greyish brown with a white margin 1 mm broad (Kirschner and Piepenbring, 2014). The optimum temperature for growth in culture is 25°C. Conidia produced in culture have been used to inoculate plants and typical symptoms have been obtained (Hwang and Chen, 1986).

Disease cycle and epidemiology In the south of Taiwan, disease development is greatly affected by climatic conditions. The incubation period is about 35 days in the hot, rainy summer and about 60 days in the cool, dry winter (Hwang, 1991). Leaf speckle builds up to epidemic levels on unsprayed Cavendish cultivars after bunch emergence.

Host reaction Cultivars of the Cavendish subgroup (AAA) are affected in Taiwan and Vietnam. Musa balbisiana



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c

a

b

d

e

Fig. 2.21.  Ramichloridium ducassei from Taiwan. Key: a–c = morphology in situ, d–e = morphology in culture on corn meal agar; a = stroma and base of conidiophore as seen in a transverse leaf section; b = apices of conidiophores; c = conidia; d = conidiophores; e = conidia; scale bars: a = 20 μm, b–e = 10 μm; reproduced from Kirschner and Piepenbring, (2014) with permission from Elsevier.

and cultivars in the AAB genomic group are also reported as susceptible in Taiwan (Hwang, 1991). ‘Pisang Awak’, ‘Kluai Namwa’, ‘Kluai Namwa Khom’ and ‘Ducasse’ (ABB, Pisang Awak subgroup) seem particularly susceptible. Symptoms have also been seen by the author in Southeast Asia on ‘Chuõí Bom’ (AAA, Lakatan subgroup), ‘Pisang Abu Keling’ (ABB, Bluggoe subgroup), ‘Pisang Abu Siam’ (ABB, syn. ‘Kluai Teparot’) and ‘Pisang Abu Nipah’ (ABB, Saba subgroup).

speckle built up to epiphytotic levels during the period of fruit maturation (Hwang and Chen, 1986). Fungicidal sprays containing dithiocarbonates in oil provide effective control (Hwang, 1991). In general, fungicides that are applied in Taiwan today to control black leaf streak and freckle control Taiwan leaf speckle (C.-P. Chao, Taiwan, 2017, personal communication).

Tropical Leaf Speckle Introduction

Control Only in Taiwan has this disease been a significant problem. After the initiation of a forecasting system to control black leaf streak, banana plants were not sprayed with fungicide after bunch emergence. Under these conditions, Taiwan leaf

Tropical leaf speckle is a collective name given to two types of symptom that have been observed on banana leaves in hot, humid tropical environments around the world (Stahel, 1937b; Martyn, 1945; Pont, 1960b; Stover, 1972). Some authors contended that the disease was

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caused by two different fungal pathogens and this was supported by taxonomic studies undertaken by Ellis (1967, 1976). However, de Hoog (1977) was of the opinion, after examining specimens of both types from many locations, that only one fungus was involved. More recently, a modern taxonomic study has shown that two different species and perhaps a third may cause the disease known as tropical leaf speckle. The disease is not regarded as serious and no effect on yield or growth has been documented. Tropical leaf speckle has been described in many countries in the Asian–Pacific, African and Latin American–Caribbean regions and probably has a worldwide distribution. It can be found in humid environments close to rivers and on hills where banana is grown amongst forest trees (L. Pérez-­ Vincente, Cuba, 2016, personal communication). It is also very common in South America where banana is grown as a component of an intercropping system (L. Pocasangre, Costa Rica, 2016, personal communication).

the lower leaf surface. With a hand-lens, these appear as densely aggregated, minute black specks. Individual speckled blotches are smaller than those characteristic of the first symptom, but often merge into extensive discoloured areas. These blotches are less distinct on the upper surface. Plate 2.50 is believed to illustrate this second type of symptom. The conidiophores are clearly visible on the lower surface as a dense, almost velvety coating. These symptoms were seen on midribs (Plate 2.50) and peduncles (Plate 2.51) by the author in the Philippines. Sometimes, in commercial plantations of ‘Umalag’ (AAA, Cavendish subgroup) in the Philippines,

Symptoms In northern Australia, tropical leaf speckle has been reported on young foliage, even as high on the plant as the third fully expanded leaf, where symptoms of Sigatoka leaf spot or Mycosphaerella leaf speckle were not yet evident. However, no extensive necrosis or breakdown of leaf tissue, which could lead to premature defoliation, has been associated with the disease. Lesions are not discrete, but diffuse. Pont (1960b) described two types of leaf symptom. The first appears as roughly circular, chlorotic blotches up to 4 cm in diameter on the upper surface of the leaf and as tan-coloured blotches underneath. Thickly distributed, dark brown or black pinpoint-sized specks are said to be clearly visible on the upper leaf surface. Densely packed, bristle-like conidiophores of the causal fungus were observed on the lower surface, even with the naked eye if the leaf were wrapped around a finger and held against the light. Blotches could merge to form extensive tan-coloured areas on leaves. Plate 2.49 is believed to illustrate this type of symptom. The second type of symptom appears as irregular circular, dark grey to black patches on

Plate 2.49.  Symptoms of tropical leaf speckle on the underside of a leaf of a cultivar in the Cavendish subgroup (AAA) in Australia (photo: Cooke et al., 2009).

Plate 2.50.  Grey to black patch symptoms of tropical leaf speckle on the underside of a leaf lamina and midrib of ‘Umalag’ (AAA, Cavendish cultivar) in a commercial plantation on Mindanao Island in the Philippines (photo: D.R. Jones, INIBAP).



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The pathogen now known as Z. musigenum is believed to be associated with the first type of symptom described above and Z. biverticillatum with the second. However, more work needs to be undertaken before the two causal agents that seem to be responsible for tropical speckle can definitely be associated with the symptoms described above. The main difference in the morphology of the two main species causing tropical leaf speckle of banana is that the conidiophores of Z. musigenum are mainly unbranched while those of Z. biverticillatum are predominantly branched (Kirshner and Piepenbring, 2014). Kirschner and Piepenbring (2014) provide a preliminary key for identifying all species of Ramichloridium found on banana, based on the morphology of conidiophores and conidia found on in situ specimens and in vitro colonies. Zasmidium musigenum (syn. Ramichloridium musae)

Plate 2.51.  Symptoms of tropical leaf speckle on the peduncle of ‘Umalag’ (AAA, Cavendish cultivar) in a commercial plantation on Mindanao Island in the Philippines (photo: D.R. Jones, INIBAP).

the ends of fingers are affected, which renders fruit unacceptable for export. Both types of symptom can occur together on the same leaf in northern Australia (Pont, 1960b).

Causal agents Two species of fungus, Veronaea musae (syn. Chloridium musae) and Periconiella musae (syn. Ramichloridium musae), which were distinguished mainly on conidiophore morphology by Ellis (1967, 1971b, 1976), were initially identified as the causal agents. However, a revision of Ramichloridium and allied genera by Arzanlou et al. (2007b) resulted in V. musae being renamed R. musae and Pe. musae being renamed R. biverticillatum. A recent further revision of some species in the Mycosphaerellacae by Videira et  al. (2017) has changed R. musae to Zasmidium musigenum and R. biverticillatum to Z. biverticillatum.

Zasmidium musigenum (as R. musae) described from banana in Panama had erect, unbranched conidiophores (30–)92–190(–220) μm long that arose singly from external mycelium. They were medium brown in colour, becoming paler and narrower towards the apex, with septa 9–30 μm apart. Conidiogenous cells, which were (9–)13–23(–25) μm long, were terminal and intercalary, cylindrical or slightly subulate, straight, pale brown, smooth. Conidia were solitary, cylindrical or oblong, pale brown, smooth with dimensions of (3–)3.5–5 × (1.5–)2(–3) μm. The basal hilum was darkened and slightly thickened (Fig. 2.22) (Kirshner and Piepenbring, 2014). Colonies on malt extract agar have been reported to reach a diameter of 27 mm after 14 days at 24°C. The colony was mostly submerged with some floccose to lanose aerial mycelium in an olivaceous grey centre, The colony margin was entire, smooth, sharp and coloured pale pinkish olivaceous. The reverse side of the colony was pale orange (Arzanlou et al., 2007b). Zasmidium biverticillatum (syn. Ramichloridium biverticillatum) Zasmidium biverticillatum (as R. biverticillatum), also described from banana in Panama,

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d

c

a b

was described as having erect conidiophores (26–)54–186(–245) μm long arising singly from the external mycelium. They were medium brown in colour, becoming paler towards the apex, with septa 5–28 μm apart. Two to six conidiogenous cells arose at the apex in alternate or opposite lateral positions, giving a branched appearance. However, the conidiophore could also be unbranched. The conidiogenous cell, which was slightly subulate and curved downwards, pale brown, smooth to finely verruculose, measured (9–)11–18(–20) × 2–3 μm. Conidia were solitary, cylindrical or oblong with a broadly rounded apex and narrowing base, pale brown, smooth with dimensions of (5–)6–7(–9) × 2–2.5(–3) μm. The basal hilum was darkened

Fig. 2.22.  Zasmidium musigeum (syn. Ramichloridium musae) from Panama – morphology in situ. Key: a = conidiophores; b = ­conidiophore bases; c = apices of condiophores, the left one with a terminal and an intercalary conidiogenous cell, the right ones with terminal conidiogenous cells arising from a percurrent extension; d = ­Conidia; scale bars a = 20 μm; b-d = 10 μm; reproduced from Kirschner and Piepenbring (2014) with permission from Elsevier.

and slightly thickened (Fig. 2. 23) (Kirschner and Piepenbring, 2014). On malt extract agar, colonies were slow-growing, reaching a diameter of 6 mm after 14 days at 24°C. The surface mycelium was velvety to hairy and coloured dark olivaceous grey. Colonies had an entire margin (Arzanlou et  al., 2007b). Pont (1960b) reported that the fungus now known as Z. biverticillatum grew more slowly than the fungus now known as Z. musigenum on potato dextrose agar, but had similar colony characteristics. The optimum growth temperature for the former was recorded as 27–28°C and for the latter 26°C. Molecular and morphological analysis showed that most specimens of tropical leaf



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b g

e

c

d

f

a Fig. 2.23.  Zasmidium biverticillatum (syn. Ramichloridium biverticillatum). Key: a–b = specimen from Panama – morphology in situ; a = conidiophore bases and apices of branched and unbranched ­conidiophores; b = conidia; c–g = specimen from Taiwan in situ (c–e) and in culture on corn meal agar (f–g); c = conidiophore; d = conidiogenous cells in lateral branches; f = simple and branched conidiophores; g = conidia; scale bars a, b, d, e, g = 10 μm, c = 100 μm, f = 20 μm; reproduced from Kirschner and Piepenbring (2014) with permission from Elsevier.

speckle on Cavendish cultivars from northern Queensland in Australia were associated with Z. biverticillatum. Zasmidium musae-banksii (syn. Ramichloridium australiense) Another Zasmedium sp. has been found affecting banana in Australia (Arzanlou et  al., 2007b; Videira et  al., 2017). However, the symptoms caused by Zasmidium musae-banksii (as R. australiense), which was identified on M. acuminata spp. banksia growing wild in North Queensland, have not been documented. As only one specimen exists, this species is probably not as common as the other three in the same genus that cause tropical leaf speckle and Taiwan leaf speckle. Zasmidium musae-banksii has conidiophores up to 400 μm tall that arise vertically and can be clearly differentiated from creeping aerial hyphae. Intercalary cells are 8–40 × 2–5 μm, subhyaline, later becoming pale brown and warted in the lower part. Subtending hyphae are thick-walled and warted. Conidiogenous cells are terminal

and 10–18 μm long. Conidia are solitary, aseptate, thin-walled, smooth, subhyaline, subcylindrical to obclavate, (10–)12–15(–23) × 2.5–3 μm, with a truncate base and a slightly darkened and thickened hilum (Arzanlou et al., 2007b). In culture, colonies on malt extract agar reached a diameter of 8 mm after 14 days at 24°C. The mycelium was flat and olivaceous grey, becoming granular with gelatinous droplets developing at the margin with ageing. The under surface was pale olivaceous grey. The margin was entire and smooth (Arzanlou et al., 2007b).

Disease cycle and epidemiology Conidia germinate on lower leaf surfaces after 24 h and produce fine hyphae. Eventually a loose network of epiphytic hyphae forms on the leaf surface. The hyphae, which are 1–2 μm thick, have side branches that develop over about every sixth stoma. These side-branches, called stomatopodia, are club-shaped and a

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darker brown than the other hyphae. An infection tube from the stomatopodia enters the leaf through the stoma. Infection hyphae grow across the air space beneath the stoma to the palisade tissue and side-branches enter cells, which then die. However, the fungus spreads no further and necrosis is confined to the tissue adjacent to invaded stoma. This gives the lesion its speckled appearance. Conidiophores arise from epiphytic mycelium. The base of the conidiophore is attached to the cuticle. The causal fungi are weak parasites and in Australia symptoms are only seen in unsprayed plantations located in rainforest clearings in high-rainfall areas. In Central America, the disease is only noticeable on lower leaves during the high-rainfall months. In Papua New Guinea, it is present on banana plants grown in shady, moist locations.

Host reaction Not much is known about cultivar response. Tropical speckle has been seen by the author on cultivars in the AA, AAA, AAB, ABB genomes in Southeast Asia, including members of the Cavendish subgroup. The disease has also been observed by the author on M. schizocarpa and M. acuminata ssp. banksii in Papua New Guinea and Pont (1960a) saw symptoms on M. acuminata ssp. banksii in Queensland, Australia (Pont, 1960b).

Control The disease is of little economic significance, except when it occasionally affects fruit, and control is not usually warranted. Fungicide sprays used to control leaf spot diseases of banana reduce the incidence of tropical speckle.

Freckle Introduction Freckle is the name given to leaf- and fruit-­spotting diseases of banana common in South and East Asia (Bangladesh, Bhutan, Brunei, China,

Hong  Kong, India, Indonesia, Malaysia, Nepal, Pakistan, Philippines, Sri Lanka, Taiwan, Thailand, Vietnam) and Australasia–Oceania (American Samoa, Australia, Cook Islands, Fiji, Hawaii, New Caledonia, New Zealand, Nuie, Papua New Guinea, Samoa, Solomon Islands, Tonga) (Dingley et al., 1981; Zhou and Xie, 1990; CMI, 1990a; Infomusa, 1994; Jones, 1994b). Records from Africa (Congo, Zambia) and the Caribbean ­ (­ Dominican Republic, Jamaica and St Lucia) need confirmation because of confusion that existed in the taxonomy of the fungus causing the disease and because typical symptoms have not been seen by the author in these regions. Freckle is regarded as more serious than black leaf streak on ‘Pei-Chiao’ (AAA, Cavendish subgroup) in Taiwan (Tsai et al., 1993) and it is emerging as a significant disease on plantations of ‘Grand Nain’ (AAA, Cavendish subgroup) in the Philippines, where fruit can be severely blemished, thus affecting its marketability. Together with Cladosporium leaf speckle, freckle also causes a serious disease problem on leaves of ‘Pisang Berangan’ (AAA, Lakatan subgroup) grown on a plantation scale in West Malaysia. On banana grown in smallholdings, freckle is not usually considered a major problem, as blemished fruit is acceptable at local markets. However, heavy infections do lead to the premature death of older leaves of some cultivars. Freckle has also been recorded on leaves and fruit of abaca in the Philippines, though it is not regarded as a serious disease (Anunciado et al., 1977).

Symptoms Two types of leaf spotting have been described, which occur mainly on the upper surfaces of older banana leaves. One consists of very small, dark brown to black spots less than 1 mm in diameter. These give the leaf a sooty appearance (Plates 2.52 and 2.53). Sometimes, spots may cluster in lines and appear as streaks running from the midrib to the leaf edge along veins (Plate 2.52) or diagonally and horizontally along the leaf lamina (Plate 2.53 and 2.54). These streaks indicate where spore-­ ladened moisture has run down leaf veins on horizontal unfurled leaves or down the lamina of an erect and uinfurling leaf. If there has



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Plate 2.52.  Symptoms of freckle on a leaf of a cultivar in the Cavendish subgroup (AAA) in Vinh Phu province, Vietnam. Pycnidia have developed in lines along veins. The leaf is yellowing and turning necrotic where concentrations of pycnidia are high (photo: D.R. Jones, INIBAP).

Plate 2.54.  Necrosis of a leaf of a cultivar in the Cavendish subgroup growing in the Philippines caused by freckle. Note that the midrib has buckled and the leaf lamina is hanging down. Bands of pycnidia running along the leaf indicate that conidia were deposited when water droplets ran down the inside of the leaf when it was erect and unfurling The pathogen here is likely to be Phyllosticta cavendishii (photo: Y. Israeli, JVBES).

Plate 2.53.  Symptoms of freckle on a leaf of ‘Pisang Rastali’ (ABB, Silk subgroup) in Kelantan state, West Malaysia. Bands of pycnidia running along the leaf indicate that infection occurred after conidia-laden water droplets ran down the inside of the leaf when it was erect and unfurling. Later, when the leaf was horizontal, conidia were distributed down leaf veins (photo: D.R. Jones, INIBAP).

been a high ­infection density, diseased areas of leaf become necrotic and the lamina collapses (Plate 2.54). Large individual dark brown to black spots up to 4 mm in diameter characterize the other type of spotting (Plate 2.55). These spots may have a grey or fawn centre and can aggregate to form large blackened areas or streaks. These symptom differences were observed in Hawaii by Meredith (1968). More work needs to be undertaken on determining if the various freckle pathogens have symptomatic differences on ­ ­certain Musa genotypes.

Plate 2.55.  Symptoms of freckle on a leaf of ‘Mondolpin’ (ABB, Bluggoe subgroup) on Badu Island in the Torres Strait region of Australia. Note the larger appearance of the spots, which are also present on the leaf midrib, than those shown in Plates 2.51, 2.52 and 2.53. Surveys in the area indicate that the pathogen is most likely to be Phyllosticta maculata (photo: D.R. Jones, QDPI).

Because the pycnidia are prominent under a raised epidermis, diseased leaves and fruit feel rough to the touch. Severely affected leaves yellow, wither and die prematurely in both cases

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(Meredith, 1968). Spots also form on petioles, midribs (Plate 2.55) and transition leaves. The longevity of freckle-affected leaves has been estimated to be half that of healthy leaves in Taiwan (Chuang, 1984). A description of symptoms on fruit is provided in Chapter 4. Causal agents The fungus Guignardia musae (asexual morph: Phyllosticta musarum) was thought to cause freckle. However, more than one pathotype or pathogen species was suspected to be involved from observations that suggested a differential reaction of some banana cultivars in certain countries. In Taiwan and Hawaii, Cavendish cultivars were reported as susceptible and ABB clones as resistant, which was the reverse of what was found in some locations in Australasia and the South Pacific by the author (Jones, 1994c). Wulandari et  al. (2010) examined specimens of freckle and identified three fungal ­species, largely on the basis of differences in ascospore morphology, which they believed caused the disease. These were named G. musae from a specimen collected in Indonesia, G. stevensii from a specimen collected in Hawaii and G. musicola from a specimen collected in Thailand. The authors advised that genetic sequence analyses of specimens from diverse locations where the disease occurred were needed to fully resolve the issue. Wong et al. (2012) reported that G. stevensii described by Wulandari et al. (2010) was associated with a speckle disease in Hawaii and that G. musicola was a nomen confusum in that its description contained elements of two different species. Genetic sequence analyses in conjunction with morphotaxonomic studies were undertaken by Wong et  al. (2012), who identified three distinct fungal species associated with freckle symptoms on cultivated banana in Asia and Oceania. These were named Phyllosticta maculata, P. cavendishii and P. musarum from their asexual stages and are described below. An assay test was developed to identify the three freckle pathogens (Wong et al., 2013b). Phyllosticta maculata Specimens of this species were found in Queensland and the Northern Territory in Australia,

Sarawak in Malaysia, Sulawasi in Indonesia, Papua New Guinea and American Samoa. This species was collected mainly from ABB clones, but also from the AAB cultivars ‘Mysore’ (Mysore subgroup) and ‘Pisang Tindok’ (Plantain subgroup) (Wong et al., 2012). Additional locations later reported for P. maculata were Fiji, Samoa, Palua and Leyte in the Philippines (Wong et al., 2013b). Fruiting bodies of P. maculata are described as black, solitary or aggregated, on patches of orange-brown lesions. They are erumpent and subepidermal, globose to subglobose, with a central apical ostiole. Ascomata have a diameter of 84–187 μm, with the diameter of the ostiole 12–15 μm. Asci are bitunicate, eight-spored, clavate to pyriform or narrowly ovoid and have dimensions of 47–85 × 18–31 μm. Hyaline, aseptate ascospores are oblong or ovoid, with obtuse ends and dimensions of (17–)19–23(–24) × (8–)9–11(–13) μm (Wong et al., 2012). Pycnidia of P. maculata have a diameter of 84–137 μm with an ostiole 9–12 μm in diameter. Solitary, hyaline, aseptate conidia are oblong or obovoid to subclavate and have dimensions of (15–)16–19(–21) × (9–)10–12(–13) μm. ­Conidia are surrounded by a mucilaginous sheath 2–4 (–6) μm thick with a straight or curved apical mucilaginous appendage (12–)15–26(–37) μm long. Spermagonia are 78–115 μm in diameter. Hyaline, aseptate spermatia are cylindrical with acutely rounded ends, becoming dumb-bell shaped with age, and have dimensions of (10–)11–13 (–14) × (1–)2 μm (Wong et al., 2012). After 1 month on oatmeal agar at 25°C ­under near-ultraviolet light, very slow-growing colonies of P. maculata lack aerial mycelium and appear stromatic, coral-shaped, undulating, extending, superficial with hazel-brown and violaceous black colours interspersed with rosy-buff sectors (Wong et al., 2012).

Phyllosticta cavendishii Phyllosticta cavendishii was found in specimens from Taiwan, Hawaii, Sumatra in Indonesia, Vietnam, East Timor, Sarawak in Malaysia, Tonga and Micronesia. It was collected mainly from Cavendish cultivars, but also from AAB clones, such as ‘Pisang Keling’ (Mysore subgroup) (Wong et  al., 2012). Phyllosticta cavendishii was discovered in the Northern Territory



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of Australia in 2013 and was the subject of an AU$26 million eradication programme which led to the destruction of all banana plants in the affected region. From old records, it seems likely that P. cavendishii had been present in a few locations in northern Australia before 2013. Outbreaks of a freckle-like disease on Cavendish cultivars are recorded in Western Australia at Kununurra in 1979 and Kalumburu in 2001 (Simon McKirdy, Australia, 2001, personal communication). Fruiting bodies of P. cavendishii are described as black, solitary or aggregated on brownish-­ black or orange-brown lesions. They are erumpent and subepidermal, globose to subglobose with a central, apical ostiole. Ascomata have a diameter of 78–129 μm, with an ostiole 9–13 μm in diameter. Asci are bitunicate, eight-spored, clavate or pyriform to narrowly ovoid with dimensions of 45–77 × 19–27 μm. Hyaline, aseptate ascospores are oblong with obtuse ends and have dimensions of (12–)14–17(–18) × (7–)8– 9(–10) μm (Wong et al., 2012). Pycnidia of P. cavendishii have a diameter of 78–137 μm with a central round ostiole 10–14 μm in diameter. Solitary, hyaline aseptate conidia are oblong or ellipsoid with dimensions of (12–) 13–16(–17) × 8–9 (–10) μm; they are surrounded by a mucilaginous sheath 1–3 μm thick and have a straight to curved apical mucilaginous appendage (8–)11–16(–20) μm long. Spermagonia are 67–127 μm in diameter. Hyaline, asepatate dumb-bell shaped spermatia have dimensions of 6–7(–8) × (1–)2 μm (Wong et al., 2012). After 1 month on oatmeal agar under near-ultraviolet light at 25°C, very slow-growing colonies of P. cavendishii lack aerial mycelium and appear stromatic, coral-like, undulating, superficial, salmon-pink to rosy-buff, with shades of hazel-brown at the edge or interspersed within the colony (Wong et al., 2012). Phyllosticta musarum Phyllosticta musarum was collected from a Hill banana (‘Virupakshi’ or ‘Sirumalai’ in the AAB, Pome subgroup) in Tamil Nadu in India and ‘Kluai Namwa’ (ABB, Pisang Awak subgroup) in Thailand (Wong et  al., 2012). It has also been reported in China (Wu et al., 2014). Fruiting bodies of P. musarum are described as black, solitary or aggregated in tight clusters within a black stroma. They are erumpent and

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subepidermal, globose to subglobose, with a central apical ostiole. Ascomata have a diameter of 102–118 μm with a central ostiole 9–13 μm in diameter. Bitunicate, eight-spored asci have a clavate or pyriformin shape and dimensions of 51–78 × 20– 30 μm. Hyaline, aseptate ascospores are oblong with obtuse ends and measure (14–)16–18(–21) × 7–8(–9) μm. They have prominent flame-like mucilaginous caps present at both ends which are 3–4 μm in diameter and up to 7 μm long (Wong et al., 2012). Pycnidia have a diameter of 69–118 μm with a central ostiole 8–14 μm in diameter. Solitary, hyaline, aseptate conidia are oblong, obovoid or ellipsoid in shape and have dimensions of (12–)13–16(–20) × (7–)9–10(–11) μm. They are surrounded by a mucilaginous sheath 1–3 μm thick, and have a straight to curved mucilaginous apical appendage which is (12–)14–18 (–20) μm long. Hyaline, asepatete spermagonia have diameters of 54–111 μm. Spermatia are dumb-bell shaped and measure (6–)7–8(–9) × (1–)2 μm (Wong et al., 2012). Other freckle reports Wickee et al. (2013) stated that if freckle-infected banana tissues are surface sterilized and plated on agar, Phyllosticta capitalensis, which is a common endophyte of banana and many other plants, invariably grows out. Furthermore, if these isolates, which could be weak pathogens, are used in pathogenicity testing they may infect banana and cause a ‘symptom’. Sun et al. (2016) reported that the pathogenicity of P. capitalensis isolated from irregular spots with dark grey centres with dark brown edges surrounded by chlorotic haloes on ‘Williams’ (AAA, Cavendish subgroup) in China was proven by Koch’s postulates. Although it seems likely on the evidence that P. capitalensis did cause the described symptoms, these do not appear to be typical symptoms of freckle. Wu et  al. (2014) described Phyllosticta musaechinensis, which was weakly pathogenic on banana leaves in China and caused a yellow discoloration. The authors did not record if affected leaves felt rough to the touch, but a photograph showed protruding pycnidia. The size of its pycnidia, the size of its conidia, the thickness of the mucilaginous sheath around the conidia and the length of the sheath’s appendage distinguished it from other Phyllosticta spp. on banana.

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Disease cycle and epidemiology Ascospores are discharged from perithecia and conidia are exuded from pycnidia en masse as white, gelatinous tendrils during wet weather or heavy dews. The importance of ascospores in the epidemiology of the disease is unclear, but conidia play a significant role. Meredith (1968) studied what in hindsight was likely to have been P. cavendishii in Hawaii. He found conidia adhered to each other because of the mucilaginous sheath that surrounds each conidium. However, water readily separated them and dissemination occured when drops ran across the leaf or fruit. On immature fruit of ‘Dwarf Cavendish’ (AAA, Cavendish subgroup), germination began as early as 2–3 h at 24°C and, after 12 h, a lateral swelling proliferated into an irregular, hyaline appressorium. After 18–30 h, the appressorium was distinct, being light or dark grey with a thickened wall and a septum where it joined the conidium. Most appressoria were formed in depressions between epidermal cells. Penetration was thought to occur after 24–72 h at 24°C, when single epidermal cells became reddish brown. After 96 h, more than 60% of appressoria were associated with discoloured host cells. This response was more rapid and extensive in areas where appressorium density was high. Fine penetration hyphae entered the cell from the underside of the appressoria. These swelled to a diameter of 3–5 mm on entering the cell lumen. The surrounding tissue was subsequently invaded by intercellular and intracellular hyphae. Lesions were superficial and tissue below five cell layers was rarely invaded. Pycnidia could develop as early as 3 weeks after inoculation (Meredith, 1968). The infection of banana leaves of ‘Brazilian’ (AAA, Cavendish subgroup) in China by what is also likely to be P. cavendishii was similar (Pu et al., 2008). Germination of conidia began 2–3 h after inoculation in a film of water at 25°C. More than 50% and 90% of conidia had germinated at 12 h and 72 h, respectively. Although Meredith (1968) described appressoria arising directly from the conidia, Pu et al. (2008) reported many appressoria forming at the end of an elongated germ tube. The appressoria were also more regularly shaped than those seen by Meredith (1968). One or occasionally two germ tubes developed from each conidium. When there were two, only one

developed a mature appressorium. Appressoria, which were initiated from 6 h after inoculation, were dark brown in colour. Appressorium formation had reached more than 50% after 24 h and 80% after 48 h. A reddish-brown necrosis of host cells appeared under appressoria, most of which were formed in the grooves between adjacent epidermis cells. Penetration was directly through the epidermal cuticle layer and not through ­stomata (Pu et al., 2008). The infection process of P. maculata has been studied using an electron microscope (Wong et  al., 2013a). Germination of conidia commenced within 3 h of inoculation. Germ tubes or hyphae were long and slender, and developed secondary lateral branches at various points on the leaf surface. Most grew in random directions, but some grew along the junction line of two epidermal cells. Appressoria were formed 18 h after inoculation and usually at the tips of germ tubes. They were variable in shape, but those that were obvoid and ellipsoid with an unlobed outline were more common than those that were digitated and lobed ones. Falciform types were rarely observed. Hyphae passed by or grew over open or closed stomata. Penetration was directly through the host cuticle. On fruit, germ tubes were mostly short and swollen, forming sessile appressoria. Each conidium usually produced one slender germ tube, but occasionally two were observed (Wong et al., 2013a). In Taiwan, the incubation period varies ­between about 60 days in the dry, cool season to 20 days in the warm, wet season. The susceptibility of leaves increases as they age (Chuang, 1984). Pycnidia develop in lesions of all sizes, some as small as five dead cells. Streaks of pycnidia are found where water-drops loaded with conidia have run across the leaf surface. Fruit becomes diseased when water runs off infected leaves on to bunches. Often, infection occurs near the ­pycnidium, where the conidia are formed, which leads to an increase in intensity of the disease. On leaves, continuous day-to-day infection results in an overlap of lesions, which results in extensive areas of dead tissue (Meredith, 1968). Host reaction In the South Asia and Southeast Asia region, freckle symptoms have been seen on clones in



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the Cavendish (AAA), Lakatan (AAA), Pome (AAB), Silk (AAB), Plantain (AAB), Mysore (AAB), Bluggoe (ABB), Pisang Awak (ABB) and Saba subgroups (ABB). ‘Pelipita’ (ABB) and ‘­Kluai Teparot’ (ABB) are also susceptible. Observations indicate that clones in the Sucrier subgroup (AA), Gros Michel subgroup (AAA), Red subgroup (AAA), ‘Pisang Nangka’ (AAA) and Pisang Raja subgroup (AAB) may be less seriously attacked. Freckle symptoms have not yet been recorded on the AA cultivars ‘Pisang Lilin’, ‘­Inarnibal’ and ‘Pisang Jari Buaya’ (Jones and Daniells, 1988; Jones, 1993b, 1994b). In Papua New Guinea, freckle, which is probably caused by P. maculata, is present on M. acuminata ssp. banksii and M. schizocarpa. Symptoms are especially severe on hybrids between the two wild species. No symptoms have been seen on M. balbisiana in Papua New Guinea, Taiwan, Hawaii and Southeast Asia. In Australia, P. maculata was found on ‘Lady Finger’ (AAB, Pome subgroup) and ‘Bluggoe’ (ABB, Bluggoe subgroup). In Hawaii, where P. cavendishii is now known to exist, clones in the Cavendish subgroup (AAA) and Plantain subgroup (AAB) were recorded as susceptible, as were ‘Pome’ (AAB), ‘Silk’ (AAB), ‘Sucrier’ (AA), ‘Gros Michel’ (AAA), ‘Red’ and ‘Green Red’ (AAA). ‘Bluggoe’ (ABB), ‘Monthan’ (ABB) and ‘Blue Java’ (ABB, syn. ‘Ney Mannan’) were not seen with symptoms. Cultivars in the AAB Mai’a Maoli–Popoulu subgroup were thought to be resistant (Meredith, 1968). The situation in Taiwan, where P.  cavendishii also occurs, was ­similar (Hwang et al., 1984; Tsai et al., 1993). In the Northern Territory of Australia, P. cavendishii was found on Cavendish cultivars, ‘Lady Finger’ (AAB, Pome subgroup), ‘Sugar’, (AAB, Silk subgroup), ‘Ducasse’ (ABB, Pisang Awak subgroup) and ‘Bluggoe’ (ABB, Bluggoe subgroup). More research needs to be undertaken on finding the range of cultivars affected by each pathogen.

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In Taiwan, control measures include the removal of lower, diseased leaves to reduce inoculum levels and bagging bunches to help prevent fruit infection. Mancozeb has been shown to be more effective than benomyl in controlling freckle, but mineral oil was proved to be ineffective (Chuang, 1983). Propiconazole is now the preferred fungicide. In the Philippines, propiconazole and flusilazole, which control black leaf streak, also control freckle.

Rust Introduction Rust is usually a minor disease of banana. It has been described in Australasia–Oceania (Australia, Fiji, Papua New Guinea, Samoa, Wallis Islands), Asia (Malaysia, Philippines) and Africa (Congo, Nigeria) (Wardlaw, 1961; Mulder and Holliday, 1971; Dingley et  al., 1981; Shaw, 1984). Epiphytotics have occurred on commercial plantations of cultivars in the Cavendish subgroup (AAA) in the Philippines and Malaysia. In Fiji, Cavendish cultivars are quite susceptible to rust when oil-sprayed (Firman, 1972). Symptoms Uredosori appear as small brown linear lesions, mainly on the lower surfaces of older leaves. These elongate, broaden and can coalesce, causing leaf yellowing (Plate 2.56) and necrosis if

Control Freckle symptoms are present on fruit in local markets in Southeast Asia, Papua New Guinea and the South Pacific, but are tolerated by most consumers. Control becomes important when fruit is exported to more discerning buyers.

Plate 2.56.  Symptoms of rust on the underside of a leaf of ‘Williams’ (AAA, Cavendish subgroup) in North Queensland, Australia (photo: D.R. Jones, QDPI).

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the density of infection is great. Powdery, light brown masses of uredospores can often be seen covering developing lesions. Uredosori can act as points of entry for secondary infections caused by Cordana musae.

Causal agents Uromyces musae and Uredo musae are the causal agents of banana rust. Uromyces musae has powdery brown to black, elongate, erumpent sori. The uredosorus is generally hypophyllous, round and pulvinate. Uredospores are globose to subglobose or ellipsoidal, light brown, finely echinulate and 20–28 × 17–24 μm in size with walls 2.5 μm thick. The telia are hypophyllous, scattered or aggregated, oblong or ellipsoidal, 0.5–1 mm in length or up to 3 mm when joined in series. Teliospores are subglobose, ovoid or oblong, brown with dimensions of 23–35 × 17–25 μm on persistent pedicels about 60 μm long (Mulder and Holliday, 1971). Authenticated specimens of this rust fungus are only known from Africa (Firman, 1976). Uredo musae has uredia that are more crowded. Uredospores have a thinner cell wall (1.5 μm) than those of Uromyces musae and more pronounced echinulations (Mulder and Holliday, 1971). All collections of rust on banana on Pacific Islands have been of this fungus (Firman, 1976). The future of the taxonomy of the rusts has been discussed by Marin-Felix et al. (2017). It is possible that one or both of these species may one day be accommodated in Puccinia.

Disease cycle and epidemiology Little has been reported on the life cycle of rust fungi affecting banana. The existence of a host other than banana has not been determined. Wind-blown uredospores carry the disease from banana to banana. These germinate in moisture and invade leaf tissue. Host reaction Many cultivars seem to be susceptible to rust. In Fiji, where rust is common, symptoms have been seen on representatives of the AA, AAA, AAAA, AAB and ABB genomes, including ‘Dwarf Cavendish’, ‘Veimama’ and ‘Poyo’ (AAA, Cavendish subgroup), ‘Gros Michel’ (AAA) and ‘Blue Java’ (ABB, syn. ‘Ney Mannan’). About half the cultivars examined in a screening trial at Koronivia Research Station were diseased, but in most cases symptoms were slight and very few leaves were affected. Rust appeared on two unaffected cultivars in the Cavendish subgroup after spraying with oil (Firman, 1972). Rust has been seen on M. acuminata ssp. banksii in Samoa (Dingley et al., 1981). Control No control is generally warranted. However, s­evere outbreaks that have occurred in some ­commercial Cavendish plantations appear to be related to the continual use of either oil or benomyl alone to control leaf spot diseases (Gerlach, 1988).

References Aba, S.C., Baiyeri, P.K. and Tenkouano, A. (2011) Impact of poultry manure on growth behavior, black Sigatoka disease and yield attributes of two plantain (Musa spp. AAB) genotypes. Tropicultura 29, 20–27. Abadie, C., El Hadrami A., Fouré E. and Carlier, J. (2003) Efficiency and durability of partial resistance components of bananas against black leaf streak disease. In: Jacome, L., Lepoivre, P., Marín, D., Ortiz, R., Romero, R. and Escalant, J.V. (eds) Mycosphaerella Leaf Spot Diseases of Bananas: Present Status and Outlook. Proceedings of the Workshop on Mycosphaerella Leaf Spot Diseases, San José, Costa Rica, 20–23 May 2002. INIBAP, Montpellier, France. pp. 161–168. Abadie, C., Zapater, M-F., Pignolet, L., Carlier, J. and Mourichon, X. (2008) Artificial inoculation on plants and banana leaf pieces with Mycosphaerella spp. responsible to Sigatoka leaf spot diseases. Fruits 63(5), 319–323.



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3 

Fungal Diseases of the Root, Corm and Pseudostem

Fusarium Wilt R.C. Ploetz Introduction Economic impact Fusarium wilt, which is also known as Panama disease, is one of the most destructive diseases of banana (Stover, 1962; Ploetz and Pegg, 1999; Ploetz et al., 2015b). Its influence in the early export trades, the re-emergence of the disease as a factor in Asian and African production, and its importance on diverse banana cultivars in different production sectors emphasize its global significance. Fusarium wilt also affects abacá, but damage on this crop is usually restricted to a reduction in fibre quality (Waite, 1954). It was the most serious disease of abacá in the Western Hemisphere between 1900 and 1960. Despite the disease’s wide impact, its history is frequently associated with only two of the many subgroups of banana that exist. Before the mid20th century, Fusarium wilt was most closely associated with the expansion and demise of export plantations of ‘Gros Michel’ (AAA) (McKenney, 1910; Brandes, 1919; Wardlaw, 1961; Stover, 1962; Ploetz, 2005). Beginning in 1890 in Costa Rica and Panama, virtually every export-producing area in the Americas and Africa was eventually affected. As the trades attempted to keep pace with

the disease’s spread, more areas were brought into, and subsequently lost to, production. Major financial losses accrued, and significant political turmoil and economic hardship developed throughout the so-called ‘banana republics’ (Henry, 1904; Jenkins, 2000; Marquardt, 2001; Koeppel, 2008). Using figures from Stover (1962), Ploetz (2005) calculated that US$2 billion were lost during the ‘Gros Michel’ era (equal to about US$2.4 billion in 2016). To survive, the export industries changed in the mid-20th century to cultivars in the Cavendish subgroup that resisted strains of the pathogen that had ruined ‘Gros Michel’ production. Recently, Cavendish cultivars have been devastated in parts of Asia and Africa by an emerging race of the causal agent, known as tropical race 4 (TR4) (Butler, 2013; García-­Bastidas et al., 2014; Ordóñez et al., 2016; Ploetz, 2015a, b). In Cavendish plantations in Indonesia, Taiwan and Malaysia, respective losses of US$121 million, US$253 million and US$14.1 million/year were estimated (Aquino et  al., 2013). An economic model predicted that TR4-induced losses would eventually reach AU$138 million/year (i.e. 28% of the industry’s total value in 2011) if the pathogen spread to all growing areas in Australia (Cook et al., 2015) Although these are sobering figures, monetary losses caused by TR4 are probably higher in China and the Philippines. By 2006, 40,000 ha had been affected in China (a four-fold increase from 2002) (Li et al., 2013a). Reminiscent of the

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‘Gros Michel’ story, TR4 is spreading to new areas in China and producers are running out of arable land that is not infested. Production has declined in the eastern production areas (Fujian, Hainan and Guangdong), where large tracts have cycled out of banana production, while production has increased in Guangxi and Yunnan in the last decade (A. Drenth, Australia, 2016, personal communication). However, the latter provinces are also affected and production is expected to drop significantly as the disease spreads and consolidates. To date, it is estimated that TR4 has affected over 100,000 ha in China (A. Drenth, Australia, 2016, personal communication). In the Philippines, only producers who have access to new (non-infested) production areas or practise rigorous sanitation measures remain in production (G. Kema, The Netherlands, 2016, personal communication). It is probable that TR4 has already caused greater losses than were experienced during the ‘Gros Michel’ era, and that these losses will continue to increase as the disease spreads. History and distribution In 1874, Bancroft (1876) was the first to describe symptoms of Fusarium wilt of banana and the presence of a fungus in the xylem of affected plants. He was also the first to recognize the contagious nature of the disease and suggest that clean planting material should be used to establish new plantations (Pegg et  al., 1996). Unfortunately, many decades after Bancroft’s (1876) recommendation, infected rhizomes are still used to establish new plantings (Blomme et al., 2014). In other cases, carelessness or bad advice have been key reasons for the spread and impact of this disease (Ploetz, 2015b). Although much could still be done to interdict Fusarium wilt, progress is impeded by human nature and the fact that ‘we do not learn from history’ (Hegel, 1837). When the disease is not taken seriously, when improper measures are used for its management, and when strict sanitation and quarantine procedures are not followed to establish and maintain plantations of susceptible cultivars, rapid spread and serious damage are usually certain (Buddenhagen, 2009; Ploetz, 2015b). Simmonds (1966) proposed that the Fusarium wilt pathogen arose wherever significant

banana production occurred. Although he suggested that susceptible clones could generate their own pathogen, the available evidence indicates that most populations of the pathogen coevolved with banana in Southeast Asia, only to be moved to other locations as this important plant was disseminated (Stover, 1962; Vakili, 1965; Ploetz and Pegg, 1997; Ploetz, 2007). Soon after Bancroft (1876) reported Fusarium wilt in Australia, the disease was described in additional locations outside Southeast Asia (McKenney, 1910; Brandes, 1919; Wardlaw, 1961; Stover, 1962; Ploetz and Pegg, 1999; Ploetz, 2005) (Table 3.1). In many cases, the causal fungus moved from country to country on susceptible banana cultivars. Stover (1962) speculated that it was widely distributed in the western tropics on ‘Silk’ (AAB), which was used as a shade crop for cacao production. However, it was on the widely planted ‘Gros Michel’ that most of the first reports in the West were made (26 of 28 areas listed by Stover (1962)). A similar scenario was evident in West Africa, where the disease was first reported in export production areas on ‘Gros Michel’. However, in East Africa, where insignificant export production occurred, first reports were often on an important cooking banana, ‘Bluggoe’ (ABB) (Wallace, 1952; Jameson, 1953; Stover, 1962). Currently, few banana-production areas remain free of Fusarium wilt. As of the mid-1980s, Stover and Simmonds (1987) indicated that Fusarium wilt was not known in/on South Pacific islands including Papua New Guinea, most of the island of Borneo, Somaliland and countries boarding the Mediterranean. Since their book was published, the disease has continued to spread, and Somalia remains the only country on this list that is apparently free of the disease (Parham, 1935; Russo et al., 1985; Shivas et al., 1996; Shivas and Philemon, 1996; Trujilo, 1971; Davis et al., 2004; Ammar, 2007; García-Bastidas et al., 2014; Ordóñez et al., 2016). General histories of the disease’s spread are found in Stover (1962) and Ploetz and Pegg (1999). Symptoms Fusarium wilt causes a reddish to dark brown discoloration of the xylem. The first internal symptoms occur in feeder roots, which are the



Table 3.1.  First reports, BCGs and references for Fusarium wilt in different banana-producing countries. Country

Africa

Burundi Cameroon Comoros Islands Congo (Republic of the) Congo (Democratic Republic of the) Egypt Ethiopia Ghana Guinea Kenya

Madagascar Malawi Mauritius Mozambique Nigeria Portugal Rwanda Sierra Leone South Africa Spain Tanzania Uganda

Area/region

First report

VCG(s)

Reference(s)

1980 1955 ? 1959 before 1980

0124, 0125, 01212 ? 0128 ? 0124, 0125, 01212

Sebasigari and Stover (1988); Gatsinzi and Sebasigari (1989) Stover (1962) Ploetz and Correll (1988) Stover (1962) Gatsinzi and Sebasigari (1989); Ploetz (1990)

2007 1967 1937 1939 1952

? ? ? ? 0124, 0124-0125, 0124-0125-012801220, 01212 ? 0124, 0125, 01214 ? ? 01213-01216 01210-01215 0120 0124, 0124-0125, 0125 ? 0120, 01215 0120-01215 0124, 0125, 01212 0124, 0124-0125, 0125, 01212

Ammar (2007) CMI (1977) Symond (1939) Heim (1943) Kenya Department of Agriculture (1954); Kung’u and Jeffries (2001)

1961 1969 1930 1970 2015 1924 Madeira Islands 1979 1980 1924 1940 Canary Islands 1924 1951 1953

CMI (1977) CMI (1977); Ploetz et al. (1992) Orian (1932) CMI (1977) Butler (2013) CMI (1954); Ploetz and Pegg (1999) Hernandez et al. (1993) Ploetz and Pegg (1999) Ashby (1926) Ploetz (1990) Ashby (1924); Hernandez et al. (1993) Wallace (1952); Rutherford (2001) Jameson (1953); Kangire et al. (2001)

Fungal Diseases of the Root, Corm and Pseudostem

Continent

Continued 209

Continent

Country

Americas and Caribbean

VCG(s)

Reference(s)

Belize Brazil Cayman Islands Colombia Costa Rica Cuba

1913 1931 1938 1929 1890 1908

Brandes (1919) Deslandes (1937); Ploetz and Pegg (1999) Edwards (1938); Ploetz and Pegg (1999) Anonymous (1929) McKenney (1910); Koenig et al. (1997) Smith (1910); Ploetz and Pegg (1999)

Dominica Dominican Republic Ecuador El Salvador France

1936 1926 1929 1956 1937 1932 1910 1930 1938 1916 1903 1932 1919 1890 1959 1906 1907 1986

? 0120, 0120-01215, 0124 01210 ? 0120, 0120-01215 0120, 0124, 0124-0125, 0125, 01210 ? ? 0120 ? 0120 ? ? ? 0124, 0124-0125-0128 0120, 0124, 0126 0120, 0124, 0125 0124, 0124-0125-0128 0124, 0124-0125-0128 ? ? ? ? 0120, 0124, 0124-0125, 0125, 01210 0124 01215 0124-0125, 0128, 01220

Guadeloupe Martinique

Florida Puerto Rico

Asia

Venezuela Bangladesh

1910 1930 ?

Dominica Agricultural Department (1937) Ciferri (1927) Anonymous (1929); Magdama and Jimenez-Gasco (2015) Stover and Waite (1960) Wardlaw (1937); Ploetz (1992) Kervegant (1932) Prescott (1918) Martyn (1930) Wardlaw (1938); Ploetz and Pegg (1999) Prescott (1918); Ploetz and Correll (1988) Ashby (1913); Koenig et al. (1997) Dampf (1932); Ploetz and Pegg (1999) Brandes (1919); Ploetz and Pegg (1999) McKenney (1910) CMI (1977) Essed (1910) Rorer (1911) Ploetz and Correll (1988); Ploetz and Pegg (1999) Fawcett (1911); García-Rodríguez (2016) Chardon and Toro (1934); Guedez and Rodriguez (2004) Molina et al. (2011)

Chapter 3

First report

Guatemala Guyana Haiti Honduras Jamaica Mexico Nicaragua Panama Peru Suriname Trinidad USA

Area/region

210

Table 3.1. Continued.

Continent

Country

VCG(s)

Reference(s)

Cambodia

?

Molina et al. (2011)

China

?

0123, 0124-0125, 01218, 01221, 01222 0120-01215, 0123, 0124-01222, 0126, 01213-01216, 01218, 01220, 01221 0124-0125, 01220 0120-01215, 0121, 0126, 01213-01215, 01218, 01219

Sarawak,

Western (Peninsular)

2016 2004 2016 2016 ?

1925

Myanmar Oman Pakistan Philippines

1925 1963, 2015 1920

Sri Lanka Taiwan

1930 1967?

Thailand

1925

Vietnam

? 2014

01213 01213-01216 01213-01216 01213-01216 0120-01215, 0121, 0123, 0124-0125, 01223, 01224 0120-01215, 0123, 01213-01216, 01217, 01218, 01222 ? 01213-01216 01213-01216 0122, 0123, 0124-0125, 0126, 01213-01216 0124-0125, 01217 0120, 0121, 0123, 01213-01216 0123, 0124-0125, 01218, 01221, 01222 0124-0125, 0128 01213-01216

Li et al. (2013a)

Basu (1911); Molina et al. (2011) Shivas et al. (1996); Ploetz and Pegg (1999); Wibowo et al. (2011)

Y. Israeli, Israel, 2016, personal communication Garcia-Bastidas et al. (2014); Ploetz et al. (2015a) Chittarath et al. (2017) Ordóñez et al. (2016) CMI (1954); Ploetz (unpublished)

Reinking (1934) Pegg et al., (1994); Ploetz (1994) Reinking (1934) Butler (2013) CMI (1977); Ordóñez et al. (2016) Reinking (1934); Ploetz and Pegg (1999)

Fungal Diseases of the Root, Corm and Pseudostem

Israel Jordan Laos Lebanon Malaysia

1911 Bali, Java, Kalimantan, Papua (Irian Jaya), Sumatra, Sulawesi



First report

India Indonesia

Area/region

Park (1930); Molina et al. (2011) Su et al. (1977); Ploetz and Pegg (1999) Ploetz et al., 1997; Reinking (1934) Vinh et al. (2001) Hung et al. (2017) 211

Continued

212

Table 3.1. Continued. Country

Area/region

First report

VCG(s)

Reference(s)

Australasia, Oceania

Australia

New South Wales, Northern Territory, Queensland, Western Australia Pohnpei Yap

1874

0120, 0124, 0125, 0128, 0129, 01211, 01213-16, 01220

Bancroft (1876); Gerlach et al. (1999)

1999 1999 1935? 1971

0126 0123 ?

L.J. Smith (unpublished) Smith et al. (2002) Parham (1935) Trujilo (1971)

0126 0128 ? ?

Shivas and Philemon (1996); Davis et al. (1999) Davis et al. (2004) Russo et al. (1985) Higgins (1904)

Federated States of Micronesia Fiji Northern Mariana Islands Papua New Guinea Tonga USA USA

Rota, Tinian, Saipan Vava’u Guam (?) Hawaii

1996 2002 1904

Chapter 3

Continent



Fungal Diseases of the Root, Corm and Pseudostem

initial sites of infection. These symptoms progress to the rhizome and are most pronounced where the stele joins the cortex (Stover, 1962). In susceptible cultivars, the pseudostem is eventually colonized, and faint brown streaks and flecks are evident on the outsides of leaf sheaths (Plate 3.1) (Ploetz, 1992; Stover, 1972). The first external symptoms of Fusarium wilt in banana are a yellowing of the oldest leaves (Plate 3.2) or a longitudinal splitting of the lower portion of the outer leaf sheaths on the pseudostem (Plate 3.3) (Wardlaw, 1961; Stover, 1962, 1972). This is followed by a wilt and collapse of leaves at the petiole base (Plate 3.4) (Stover, 1972). In some cases, these leaves remain green (Stover, 1972). As additional leaves are affected, a bunched appearance develops at the top of the pseudostem. Younger and younger leaves develop symptoms (Plate 3.5), buckle and skirt the pseudostem, and the entire plant ultimately collapses. If a bunch has emerged, it is usually stunted and produces small fruit. At this stage, pronounced vascular

213

discoloration is usually evident in the pseudostem (Plate 3.6). Ultimately, as the disease spreads, few plants if any remain unaffected in plantations of susceptible cultivars (Plate 3.7). Symptoms of bacterial wilts resemble those caused by Fusarium wilt. However, external symptoms do not develop on Fusarium wilt-affected plants that are less than about 4 months old, which can occur on plants affected by bacterial wilts (Stover, 1972). In addition, bacterial wilts can cause a brown discoloration of internal portions of fruit and terminal portions of the infructescence can become necrotic when infected by insect-transmitted strains of the pathogens. In abacá, the first noticeable symptoms of Fusarium wilt are the inward curling of the leaf blade at or near the tip of the lower leaves, which gradually droop, wilt and turn from pale yellow to yellowish brown. Newly emerged leaves are reduced in size and also wilt. A reddish-violet discoloration is noticeable in the vascular system when infected rhizomes are cut open. The fibre from decorticated pseudostems of diseased abacá is also discoloured and of a lower quality (Stover, 1972).

Causal agent Fusarium wilt of banana is caused by Fusarium oxysporum f. sp. cubense, a forma specialis (f. sp.) of the F. oxysporum species complex (Michielse and Rep, 2009; O’Donnell et  al., 2009). The complex contains non-pathogenic saprobes as well as plant and animal pathogens (diPietro et al., 2003; Michielse and Rep, 2009). The plant pathogens usually exhibit considerable host specificity, and single formae speciales affect a single or limited set of host plants. Host range

Plate 3.1.  First symptoms in the banana pseudostem after it is invaded by Fusarium oxysporum f. sp. cubense are faint brown streaks and flecks evident on the outsides of leaf sheaths (photo: R.C. Ploetz, UF).

Fusarium oxysporum f. sp. cubense causes disease on Musa acuminata, M. balbisiana, M. schizocarpa, M. textilis and their hybrids (Waite, 1954; Ploetz and Pegg, 1999). Diverse weed hosts of the pathogen have been identified (Waite and Dunlap, 1953; Pittaway et  al., 1999; Hennessy et  al., 2005), but they do not develop symptoms when they are infected (see below). Additional plants

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Chapter 3

Plate 3.2.  A cultivar in the Cavendish subgroup (AAA) with early symptoms of Fusarium wilt caused by Fusarium oxysporum f. sp. cubense subtropical race 4 near Nelspruit in South Africa (photo: D.R. Jones, INIBAP).

that were reported by Waman et  al. (2013) should not be considered hosts, as they were tested with detached leaf assays. Heliconia caribaea, H. crassa, H. collinsiana, H. latispatha H. mariae, H. rostrata and H. vellerig­ era were reported as hosts in tropical America when these plants were members of the Musaceae (Waite, 1961, 1963); they are now in the Heliconiaceae (Barrett et  al., 2014). Isolates that affected these species are no longer considered as Fusarium oxysporum f. sp. cubense, as they

were either weakly pathogenic or avirulent on ‘Gros Michel’ and ‘Bluggoe’; although some isolates affected seedlings of M. balbisiana, none are available for further study (Waite, 1961, 1963). Bentley et  al. (1998) reported the pathogen on Heliconia chartacea in Australia, but they did not indicate whether their isolates were pathogenic on banana. Additional tests are needed to assess the pathogenicity of isolates from the Americas and Australia on different cultivars of banana and Heliconia spp. Until better information is



Fungal Diseases of the Root, Corm and Pseudostem

Plate 3.3.  Splitting of the lower portion of the pseudostem can be an early indication of Fusarium wilt (photo: D.R. Jones, INIBAP).

available, pathogenic isolates from the latter hosts might best be considered a distinct taxon, perhaps ‘F. oxysporum f. sp. heliconiae’. Race Race designations facilitate comparisons among different populations of F. oxysporum f. sp. cubense, but imprecisely classify pathogenic variation in this pathogen because of the unclear genetic bases of resistance and susceptibility in the host and pathogenicity in the fungus (Ploetz, 2006a, 2015b). Better understanding is needed for these important traits. Races 1, 2, 3 and 4 of F. oxysporum f. sp. cubense have been recognized (Stover and Buddenhagen, 1986; Stover and Simmonds, 1987; Stover, 1990; Ploetz et al., 2015b). However, as explained above, F. oxysporum pathogens of heliconia, which were known formerly as race 3, are no longer considered as F. oxysporum f. sp. cubense. Isolates in race 1 were responsible for the ‘Gros Michel’ epidemics and also affected

215

Plate 3.4.  The yellowing and wilting of older leaves, which buckle and collapse at the pseudostem, caused by Fusarium oxysporum f. sp. cubense tropical race 4 on a Cavendish cultivar (AAA) in Hainan, China (photo: R.C. Ploetz, UF).

‘Maqueño’ (AAB, Mai’a Maoli–Popoulu subgroup), ‘Silk’ (AAB), ‘Pome’ (AAB) and ‘Pisang Awak’ (ABB), whereas race 2 affects mainly ABB cooking bananas, such as ‘Bluggoe’. Race 4 has been split into subtropical race 4 (SR4), which affects ‘Cavendish’ and race 1 and race 2 suscepts in the subtropics, and tropical race 4 (TR4), which affects many of the same cultivars as SR4 in the tropics as well as the subtropics (Ploetz, 2006b, 2015b; Ploetz et al., 2015a). Other races may exist. For example, isolates from East Africa and Florida (USA) damage both race 1 and race 2 suscepts, but not Cavendish cultivars (Stover, 1990; Ploetz, 1992, 1993; Kung’u and Jeffries, 2001). Vegetative compatibility group Somatic compatibility has been used to identify genetically isolated populations of the pathogen (Correll and Leslie, 1987; Ploetz and Correll, 1988; Brake et al., 1990; Hernandez et al., 1993; Moore et al., 1993, Rutherford et al., 1998). Usually, genetic

216

Chapter 3

Plate 3.5.  ‘Rasthali’ (AAB, Silk subgroup) dying of Fusarium wilt near Bangalore in India. The young plant in the foreground is also affected. ‘Rasthali’ is very susceptible to race 1 of Fusarium oxysporum f. sp. cubense (photo: D.R. Jones, INIBAP).

complementation (heterokaryon formation) between nitrate-non-utilizing auxotrophic (nit) mutants is used to identify compatible isolates (Correll et al., 1987; Leslie, 1993). When NitM and nit1 and/ or nit3 mutants of different isolates complement each other for nitrate utilization, as evidenced by the production of wild-type mycelium at the interface between ‘tester’ mutants, they are vegetatively compatible and are members of the same vegetative compatibility group (VCG). The distribution of and the relationship among the various VCGs provides

much information on the dissemination and evolution of this pathogen. Some VCGs form complexes, which are defined as groups of VCGs in which cross-compatibility occurs. So-called bridging isolates are able to complement isolates in more than one VCG (Ploetz, 1990; Ploetz et al., 1997). Isolates in these cross-compatible VCGs are closely related (Koenig et al., 1997; Fourie et al., 2011), and the following have been reported in F. oxysporum f. sp. cubense: 0120-01215, 0124-0125-0128-01220, and 01213-01216.



Fungal Diseases of the Root, Corm and Pseudostem

Plate 3.6.  Dark brown discoloration of the vascular tissues in a banana pseudostem caused by Fusarium oxysporum f. sp. cubense (photo: QDPI).

217

Over 20 VCGs of F. oxysporum f. sp. cubense have been reported (Ploetz and Pegg, 1999) and, with the exception of the erroneous VCG 0127, voucher specimens of isolates from VCGs 0120 to 01224 are deposited in international culture collections, including CBS (Utrecht, The Netherlands), FRC (University Park, Pennsylvania, USA), and NRRL (Peoria, Illinois, USA). The pathogen has an unusually large number of VCGs for a forma specialis of F. oxysporum, which may be a result of the diverse banana host and the presumed old age of this pathosystem. Before outbreaks of TR4 were recognized in the tropics, Cavendish cultivars were affected in subtropical areas with cold winters (i.e. Australia, Canary Islands and South Africa) (see Plate 3.2) (Ploetz, 1990; Ploetz and Pegg, 1999). In general, banana is sensitive to low temperatures and photosynthesis (carbon assimilation) is one of the physiological processes that is severely impacted under these conditions (Damasco et  al., 1997). When studying the effect of SR4 on ‘Williams’, Moore et al. (1993) reported that photosynthesis was dramatically reduced by low temperatures. They concluded that cold-stressed plants of this Cavendish cultivar were predisposed to disease development under these conditions. Isolates in VCGs 0120, 0129 and 01211 are responsible for SR4 damage.

Plate 3.7.  Plantation of ‘Lady Finger’ (AAB, Pome subgroup) in northern New South Wales in Australia devastated by Fusarium wilt (photo: D.R. Jones, INIBAP).

218

Chapter 3

Prior to the 1990s, Cavendish cultivars were also affected in the tropics, but in concert with other predisposing factors. For example, plantations were affected in soils in Guadeloupe that were acidified by a volcanic eruption, and in waterlogged soils in Jamaica (Ploetz, 1990, 2006a). Isolates in VCGs 0120, 0124-5 and 01220, which would not normally affect cultivars in the Cavendish subgroup in the tropics, have been responsible for this damage (Ploetz, 1990; Aguilar et  al., 2000; Ploetz and Pegg, 1999; Thangavelu and Mustaffa, 2010a). Elsewhere, another VCG, VCG 0122, was associated in the Philippines with ‘localized, chronic outbreaks of disease that do not spread throughout an area as in Taiwan’ (Stover, 1990). Finally, VCG 0121 has been recovered from affected Cavendish in Indonesia, Taiwan and western Malaysia (Ploetz and Correll, 1988; Aguayo et  al., 2017). Although it is genetically similar to VCGs that are found to attack Cavendish in the tropics (Fourie et al., 2009; Ordóñez et al., 2016), VCG 0121 is relatively uncommon and little is known of the epidemiology of disease it causes on banana. All of the previous occurrences of F. oxy­ sporum f. sp. cubense on Cavendish cultivars contrast with the current TR4 epidemic. Rather than the diverse populations of F. oxysporum f. sp. cubense that are listed above, a unique and uniform pathogen, VCG 01213-01216, is associated with the current TR4 epidemic (Ploetz, 2015b; Ordóñez et al., 2016). Importantly, isolates of VCG 01213-01216 affect Cavendish cultivars in the absence of disease-predisposing factors; serious damage develops and spreads in good soils in the tropics (Ploetz, 2015b). Pathogenic variation in F. oxysporum f. sp. cubense may exist outside the two main phylogenetic clades of the pathogen (see below). Phylogenetically distinct strains of F. oxysporum that do not fall into recognized VCGs have been recovered from wilted banana plants (Bentley et  al., 1998; García-Rodríguez, 2016; Ploetz, 2016, unpublished data; N. Ordóñez, The Netherlands, 2016, personal communication; M. Dita, Brazil, 2017, unpublished data). Significantly, some of these have been shown to cause a vascular wilt on banana (N. Ordóñez, The Netherlands, 2016, personal communication). Thus, as was suggested by Simmonds (1966), some strains of this pathogen might arise in association with banana cultivation, perhaps via the horizontal transfer

of pathogenicity determinants as evidenced in other Fusarium wilt pathogens (Vlaardingerbroek et al., 2016). Understanding how this process occurs in F. oxysporum f. sp. cubense will be key to understanding the evolution of pathogenicity in F. oxysporum (Shahi et al., 2016). Tropical race 4 (VCG 01213-01216) Fusarium wilt occurs throughout the banana belt, but specific populations of the pathogen have somewhat restricted distributions (Table 3.1). A brief history of the TR4 epidemic is provided here to clarify where and how it developed. In 1989, a previously undescribed VCG, designated VCG 01213, was identified in samples the author received from Taiwan (sent by Tsai­ athology young Chuang, Department of Plant P and Entomology, National Taiwan University, Taipei). Although it is now known to distinguish a clonal population of F. oxysporum f. sp. cubense that is responsible for the TR4 epidemic (Ordóñez et  al., 2016), little importance was attached to VCG 01213 at the time. VCG 01213 may have been present in other areas in 1989, but the above diagnosis was the first documented occurrence of TR4. Cavendish cultivars are not preferred in much of Southeast Asia and most banana production in the region had, until recent times, been of other clones. Prior to the 1990s, monoculture production of Cavendish types in the region was most prevalent in the Philippines (Mindanao) and Taiwan (Valmayor, 1990; Hwang and Ko, 2004). Cavendish production in Taiwan began in the early 20th century and by the mid-1960s a large (50,000 ha) export trade had developed (Hwang and Ko, 2004). This peak in production in Taiwan coincided with the first recorded outbreak of Fusarium wilt on Cavendish in 1967 (Su et al., 1977). It occurred in a single field in the main banana-growing region in Chiatung (Su et  al., 1986), but other circumstances surrounding the outbreak are unclear. (At the time of writing, an anecdotal report that germplasm had been i­ntroduced from Sumatra (G. Kema, The Netherlands, 2017, personal communication) could not be confirmed.) Nonetheless, a rapid increase in the incidence, severity and distribution of the disease occurred soon after the first detection and 0.5 million plants/year were affected on the island by 1976. Su et al. (1986) described the panic that developed as the disease



Fungal Diseases of the Root, Corm and Pseudostem

spread, as well as the futile attempts that were made to stem its impact. Beginning in 1992, VCG 01213 was detected in Indonesia (Java and Sumatra) and Western Malaysia on cultivars other than Cavendish (Pegg et al., 1994; Ploetz, 1994). In addition, increasingly severe outbreaks were soon reported in these areas in newly established Cavendish plantations. For the first time, large monocultures of Cavendish were grown in the centre of origin of F. oxysporum f. sp. cubense. Although Ploetz (1994) observed that it was ‘too early to attach much significance to these reports’, TR4 soon devastated Cavendish production in a very wide area (Ploetz, 2006b), and monoculture production of Cavendish became increasingly difficult, if not impossible, in wider and wider areas (Buddenhagen, 2009). To date, TR4 has been reported in Australia (Northern Territory and Queensland), China (Fujian, Guangdong, Guangxi, Hainan and Yunnan), India (Bihar), Indonesia (Bali, Halmahera, Kalimantan, Java, Papua Province, Sulawesi and Sumatra), Jordan, Laos, Lebanon, Malaysia (Peninsular and Sarawak), Myanmar, Mozambique, Oman,

219

Pakistan, Philippines (Mindanao), Taiwan and ­Vietnam (Fig. 3.1; Table 3.1) (Molina et al., 2010; Butler 2013; García-Bastidas et  al., 2014; Freshplaza 2015; Ploetz et  al., 2015b; Ordóñez et  al., 2016; Chitarrath et al., 2017; Hung et al., 2017; Zheng et al., (2018), Thangavelu (2018). A clonal population of the pathogen is responsible for the TR4 outbreaks that have been examined in Australia, China, Indonesia, Jordan, Lebanon, Malaysia, Pakistan, the Philippines and Taiwan (Ordóñez et al., 2016). The origin(s) of this strain is (are) not known, but resistance to TR4 in M. acuminata ssp. malaccensis (Peraza-­Echeverria et al., 2008) suggests that it could have coevolved with this banana ancestor (Ploetz and Pegg, 1997; Ploetz, 2007) on the Malaysian peninsula (Perrier et al., 2011). Although it ‘is probable’ (Buddenhagen, 2009) that infected materials from Taiwan established the pathogen in mainland China, the means by which TR4 spread to other areas are unclear. Morphology In culture, members of the F. oxysporum species complex are fast growing (radial growth

Fig. 3.1.  The TR4 pathogen is found in Australia (Northern Territory and Queensland), China (Fujian, Guangdong, Guangxi, Hainan and Yunnan), India (Bihar). (Bali, Halmahera, Kalimantan, Java, Papua Province, Sulawesi and Sumatra), Jordan, Laos, Lebanon, Malaysia (Peninsular and Sarawak), ­Mozambique, Oman, Pakistan, Philippines (Mindanao), Taiwan and Vietnam.

220

Chapter 3

4–7 mm/day on potato dextrose agar at 24°C), with sparse to abundant aerial mycelium and white, pink, salmon or purple pigmentation (Gerlach and Nirenberg, 1982; Nelson et  al., 1983). Micro- and macroconidia are produced on branched and unbranched monophialides (Nelson et  al., 1983). Microconidia, 5–16 × 2.4–3.5 μm, are one- or two-celled, oval- to kidney-shaped, and are borne in false heads. Macroconidia, 27–55 × 3.3–5.5 μm, are fourto eight-celled, sickle-shaped, thin-walled and delicate, with foot-shaped basal and attenuated apical cells (Gerlach and Nirenberg, 1982). Terminal and intercalary chlamydospores are usually globose, 7–11 μm, and formed singly or in pairs in hyphae or conidia. Atypically, chlamydospores are not produced by isolates of F. oxysporum f. sp. cubense in VCG 01214 (Ploetz and Pegg, 1999). When formed, sporodochia are tan to orange, and sclerotia are blue and submerged. The characteristic, sporodochial wild-type isolate of the species mutates easily in culture either to mycelial types that produce cottony, aerial mycelium, or to slimy, pionnotal types that produce depressed aerial mycelium and macroconidia in pionnotes. The latter attribute lends colonies a wet, yellow-to-orange appearance. These are one-way mutations, in that the mutants never revert back to the wild type. Some strains of F.  ­oxysporum produce strong odours in culture (Domsch et al., 1980), and these have been used to classify isolates of F. oxysporum f. sp. cubense (Brandes, 1919; Moore et al., 1991). Genetic characteristics Fusarium oxysporum f. sp. cubense has no known sexual stage. Since gametic disequilibrium was non-random among pairs of 34 of 36 restriction fragment length polymorphism (RFLP) alleles and because strains with identical genotypes occurred in multiple locations, Koenig et al. (1997) concluded that F. oxysporum f. sp. cubense reproduced in a clonal manner. However, Taylor et al. (1999) re-analysed the above RFLP data and noted little resolution and only one internal branch in one of the largest clades identified by Koenig et al. (1997), which was consistent with recombination. By examining 25 unique genotypes in the clade, they found no statistical support for strict clonal reproduction. Thus, parasexual or

sexual recombination may occur in at least one population. Fourie et  al. (2009) indicated that isolates they examined possessed either the MAT1-1 or MAT1-2 allele, which are responsible for sexual reproduction in ascomycetes (Arie et  al., 2000; Yun et al., 2000). When they paired isolates with different MAT alleles from the same evolutionary clade, protoperithecium-like structures were formed, but no ascospores. These isolates represent potentially heterothallic strains that may have lost, or have rare, fertile mating partners (Turgeon, 1998; Kerényi et al., 2004). As stated earlier, vegetative compatibility in F. oxysporum has been determined with different nitrate metabolism (nit) mutants (Correll et  al., 1987; Leslie, 1993), though other auxotrophic mutants have also been used (Puhalla and Spieth, 1983), such as those for sulfate (sul ­ ­mutants) (Hawthorne and ReesGeorge, 1996). When nit complementation is used to score vegetative compatibility, some isolates are self-­ incompatible, in that their nit mutants do not complement each other, as well as those for other isolates. They are uncommon and are typically discarded in vegetative compatibility analyses. Another category of isolates is self-compatible (produces complementary nit mutants), but is not compatible with previously described VCGs. Genetic markers have shown that some of these isolates are related to previously described VCGs, despite their apparent incompatibility via nit analyses (see below). Finally, incompatible, selfcompatible isolates may be members of unrecognized VCGs. As mentioned above, unaccounted variation in this pathogen probably occurs in Southeast Asia and other areas. With diverse markers, genetic variation has been recognized in F. oxysporum f. sp. cubense (Ploetz and Correll, 1988; Boehm et  al., 1994; Bentley et  al., 1995, 1998; Koenig et  al., 1997; O’Donnell et  al., 1998; Ploetz and Pegg, 1999; Fourie et  al., 2011; Ordóñez et  al., 2016). Two major clonal lineages are evident and most of the known VCGs in each are present in Southeast Asia (Ploetz and Correll, 1988; Koenig et  al., 1997; Bentley et al., 1998; O’Donnell et al,. 1998; Fourie et al., 2011). To date, only VCGs 0129 and 01211 (Australia), 01210 (Caribbean), 01212 (East Africa) and 01214 (Malawi) have not also been found in Asia. However, strains in these exceptional VCGs are phylogenetically related to



Fungal Diseases of the Root, Corm and Pseudostem

other strains in the above lineages, indicating descent from these populations of the pathogen. Isolates that are not related to the above populations have also been recovered from plants affected by Fusarium wilt (Bentley et al., 1998; García-Rodríguez, 2016; Ploetz, USA, 2016, unpublished data; N. Ordóñez, The Netherlands, 2016, personal communication; M. Dita, Brazil, 2017, unpublished data). Notably, some of the isolates that are not from the Southeast Asian centre of origin are also pathogenic on banana (M. Dita, Brazil, 2017, unpublished data; N. Ordóñez, The Netherlands, 2016, personal communication). Thus, the latter results may support the hypothesis of Simmonds (1966) that this pathogen also arose outside Southeast Asia. Clearly, better understandings are needed for the evolution of pathogenicity to banana in these fungi. The Fusarium genome can be divided into two regions (Ma et al., 2010, 2013; Rep and Kistler, 2010). The core region performs essential, ‘house-keeping’ functions in these fungi, whereas the accessory regions, which are located on lineage-specific chromosomes, encode host-­specific pathogenicity factors. The latter chromosomes are dispensable, in that strains can survive if they are lost (Andam et  al., 2015). In general, the core and accessory regions of the Fusarium genome are vertically transmitted (from parent to progeny). However, in rare cases horizontal (lateral) transmission can occur, in which genetic material is transferred and stably integrated into the genome of another fungus. Horizontal gene transfer has been demonstrated among distantly related organisms and is recognized as an important mechanism by which ecological and pathological competence arises in microorganisms (Andam et al., 2015). Horizontal transfer of core and accessory elements has been demonstrated in F. oxysporum (Vlaardingerbroek et al., 2016), and the transfer of pathogenicity genes has been reported in (Laurence et al., 2015; van Dam et al., 2016), or suggested by the polyphyletic nature of (O’Donnell et al., 1998; Baayen et al., 2000), many of the formae speciales in F. oxysporum. Accessory regions in TR4 strain II5 (CBS 102025 and NRRL 36114) contain genes for secreted cysteine-rich proteins, which are potential effectors, and gene clusters for candidate secondary metabolite biosynthesis (L.-J. Ma, USA,

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2016, personal communication). Notable among the accessory genes in plant-pathogenic members of F. oxysporum are those for SIX (secreted in xylem) proteins, some of which are effectors (have a role in disease development) (Rep et al., 2004; Rep, 2005; Rep and Kistler, 2010). For example, in the F. oxysporum f. sp. lycopersici–tomato system, four of the 14 known SIX genes are required for full pathogenicity (Gawehns et  al., 2015). SIX genes have also been detected in F. oxysporum f. sp. cubense. SIX1 is present in all isolates that have been examined, and three homologs of SIX1 are found in TR4, SIX1a, b and c (L.-J. Ma, USA, 2016, personal communication; M. Rep, The Netherlands, 2016, personal communication); SIX1a may have a virulence function on Cavendish cultivars (M. Rep, The Netherlands, 2016, personal communication). SIX8 is present multiple times in the subtelomeric regions of the Fol genome and is also found as multiple copies in strains of TR4 and SR4 (L.-J. Ma, USA, 2016, personal communication). Fraser-Smith et  al. (2014) identified two SIX8 homologs, SIX8a and SIX8b, in over 500 isolates of F. oxysporum f. sp. cubense. Using a PCR and sequencing approach, variation in SIX8 allowed race 4 to be differentiated from race 1 and 2 isolates, and tropical and subtropical race 4 isolates to be distinguished from one another (FraserSmith et al., 2014). Disease physiology Fusarium wilt of banana is a typical vascular wilt disease. Fusarium oxysporum f. sp. cubense is a hemibiotroph that initially establishes a biotrophic relationship with the susceptible banana plant. Although the pathogen does not induce a host response initially and disease ­ symptoms are not produced, it eventually transitions to necrotrophic behaviour in which host tissue is killed. Fusarium oxysporum f. sp. cubense infects roots of both susceptible and resistant banana cultivars, but infection of vascularized portions of the rhizome is most pronounced in susceptible genotypes (Beckman, 1987, 1990). Tyloses, gums and gels are produced in xylem lumena in response to infection, but in resistant cultivars these host products are produced earlier and far more rapidly than in susceptible cultivars. Systemic infection of the pseudostem

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of susceptible cultivars proceeds in advance of these host responses. Fusaric acid, a phytotoxin that is produced by F. oxysporum f. sp. cubense and other members of the F. oxysporum species complex, has been indicated as a cause of the leaf chlorosis symptom (Dong et al., 2012, 2014). Chloroplast damage, reduced photochemical efficiency of photosystem II (FV/Fmax), and reduced photosynthesis (net CO2 assimilation) were associated with the progression of leaf yellowing in artificially inoculated plants of ‘Gros Michel’. Fusaric acid, but not the pathogen, was detected in symptomatic leaves and chlorosis was induced after it was injected into leaf lamina. Transpiration was reduced in diseased leaves because of stomatal closure and reduced hydraulic conductivity detected in diseased stems was associated with the development of the above symptoms (Dong et al., 2012, 2014). Structural and biochemical changes in affected plants were consistent with plant senescence (Dong et  al., 2014), which would be expected during the necrotic phase of disease development. Disease cycle and epidemiology Wardlaw (1935) contended that Fusarium wilt was a ‘conditional infection’ of badly cultivated or nutrient-deficient, exhausted soils. He concluded that the Central American banana industry had exploited ‘the native fertility of virgin soil’, and that once that was exhausted, disease set in. In general, Wardlaw’s (1935) hypothesis has been repeatedly disproven, first during the race 1 epidemics on ‘Gros Michel’ and more recently with the spread of TR4. Devastating losses can develop on susceptible cultivars after the disease initially raises little concern. For example, the first, isolated outbreaks of TR4 in China and the Philippines were not taken seriously, but eventually developed into uncontrollable problems (Buddenhagen, 2009). Nonetheless, there can be great variation in the incidence and severity of this disease over time in a given production area (Stover, 1962). Disease development can be significantly impeded in disease-suppressive soils. These situations are contrasted with soils in which disease predisposition occurs. For example, the ‘disease-conducive’ pockets of soil that Peng et  al.

(1999) studied in Carnavon, Australia were areas in which water logging and/or drought predisposed Cavendish cultivars to damage by normally avirulent populations of VCG 0124-0125. Disease-free areas in these fields were not suppressive, but lacked disease-predisposing conditions (Pegg et al., 1995; Ploetz and Pegg, 1999). Disease-suppressive soils have been classified by the length of time that a susceptible cultivar can be produced in the presence of a virulent pathogen (Stover, 1962; Alvarez et  al., 1981; Chuang, 1988; Stover, 1990). Volk (as reported by Stover, 1962) classified disease-conducive ‘non-resistant’ soils (later called ‘short-life’ soils) as those in which plantations were abandoned 5–10 years after they were first planted. In contrast, plantations in ‘resistant’ soils (later called ‘long-life’ soils) remained in production for more than 20 years. Stotzky et al. (1961) associated disease suppression with chemical and physical factors and found the closest association between suppression and soils in which montmorillonoid clay was present. Later, Stover (1990) recognized suppressiveness in several different areas and listed the attributes of some of these soils. He concluded that the mechanisms for suppressiveness were unknown, but were probably biologically based. Fusarium oxysporum f. sp. cubense is disseminated in several ways, but infected rhizomes are most efficient (Stover, 1962). After the infectious nature of this disease was demonstrated by Brandes (1919), the export industries established new plantations with rhizomes from disease-free fields (Stover, 1962). Those that exhibited vascular discoloration were discarded, and in many cases suckers were also washed and treated with fungicides or biocides (Stover, 1962). However, even with these precautions it was still not possible to establish pathogen-free banana plantations. Surface waters passing over or through infested areas carry propagules of the pathogen and their use for irrigation has been responsible for the pathogen’s rapid dissemination along river basins. Furthermore, F. oxysporum f. sp. cubense is moved in soil and on contaminated tools, farm equipment, clothes and footwear (Rishbeth, 1955; Wardlaw, 1961; Stover, 1962). Recently, Kema (2016, unpublished data) demonstrated that TR4 could be moved on muddy shoes that



Fungal Diseases of the Root, Corm and Pseudostem

were worn in affected banana plantations in ­China and the Philippines. Meldrum et al. (2013a) also detected TR4 on the exoskeletons of the ­banana weevil, Cosmopolites sordidus, which they suggested could be a possible vector or disease-­ predisposing agent. Stover (1962) indicated that banana-free rotations were ineffective measures for managing the disease, and that ‘Gros Michel’ could not be produced in previously affected plantations because of the pathogen’s long survival; 20 years is typical (Stover, 1962), and 40 years has been reported (Simmonds, 1966; Buddenhagen, 2009). Stover (1962) suggested that chlamydospores of the pathogen in decayed banana tissue were responsible for its durability in infested soil. Although chlamydospores enable the patho­ gen to survive inhospitable conditions, it seems doubtful that they would be responsible for its decades-long persistence in the absence of a ­living banana host; extended survival as a non-­ pathogenic parasite of weeds may be more plausible (Ploetz, 2015a). Asymptomatic colonization of the roots of weeds by plant-pathogenic members of the F. oxysporum species complex is common (Waite and Dunlap, 1953; Pittaway et al., 1999; Rekah et al., 2001; Hennessy et al., 2005; Postic et al,. 2012; Altinok, 2013). For example, in banana plantations in Australia, F. oxysporum f. sp. cubense isolates in VCGs 0120 (SR4) and 01213/ 01216 (TR4) were recovered from surface-­ disinfested, asymptomatic roots of, respectively, Amaranthus sp. and Paspalum sp., and Chloris infla­ ta, Cyanthilium cinereum, Euphorbia heterophylla and Tridax procumberens (Pittaway et  al., 1999; Hennessy et al., 2005). Better understandings are needed for the persistence of this pathogen in the absence of its banana host as they may assist disease management efforts and provide insight into the disease’s epidemiology. The survival of the pathogen in banana leaf trash was recognized during the ‘Gros Michel’ era (Rishbeth, 1955; Wardlaw, 1961). Considerable leaf trash and other banana tissues are associated with export shipments of banana fruit. In the early 2000s, the Australian government published a risk assessment for the importation of Cavendish fruit from the Philippines that considered these banana tissues as vehicles in which non-endemic strains of F. oxysporum f. sp. cubense could be moved to Australia (DAFF Australia,

223

2004). It indicated that the fungus could move as both symptomless infections of the vasculature of fruit crowns, and in pieces of infected leaf trash that would be associated with fruit shipments. Large quantities of Cavendish fruit are exported from Mindanao to the Middle East. For example, from 2008 to 2012, Jordan imported 418,000 18 kg boxes/year and substantial quantities of fruit were also imported by other banana-­ producing nations in the Middle East, including Oman and Pakistan (Republic of the Philippines, 2012). Thus, there would have been ample opportunities for the introduction of TR4 to the Middle East if this were a viable means of dissemination. Whether, and the extent to which, fruit crowns and leaf trash are reservoirs for TR4 should be examined experimentally. Once F. oxysporum f. sp. cubense moves into a production area, it is very difficult to exclude from non-infested plantations. Although mixed plantings typically develop moderate losses (Karangwa et  al., 2016), monocultures of susceptible cultivars are usually decimated (Stover, 1962).

Control Rapid identification techniques Molecular assays that confirm the presence of a given pathogen are valuable tools in the diagnostic process. However, early observations of symptoms caused by the pathogen and collections of samples from suspect plants are key first steps in ensuring that new disease outbreaks are recognized and interdicted in a timely fashion (Dita Rodriguez et  al., 2013). This is especially true for highly destructive pathogens that can impact international trade, such as the TR4 strain. Delimiting new disease foci and ensuring that the movement of such pathogens is limited or does not occur requires rapid and decisive responses that are based on reliable information. Ideally, molecular tools that are used for detection should be vetted to ensure that false-positive or false-negative diagnoses are not made. The impact and rapid spread of TR4 has resulted in the development of several different methods to identify this pathogen. In general, TR4 strain-specific DNA of the pathogen is sought that is then detected with different techniques. However, specificity has been an issue. For example,

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a sequence characterized amplified region (SCAR) marker developed in Taiwan for ‘race 4’ detected numerous VCGs in addition to VCG 01213-01216, regardless of the technique that utilized the marker (Lin et al., 2009, 2013; Peng et al., 2014; Aguayo et al., 2017). Another assay, based on a putative virulence factor developed by Li et  al. (2013c), identified VCGs 01213-01216 and 0121 (Aguayo et  al., 2017), as did a SCAR marker developed by Li et al. (2012) and used by Yang et al. (2015) in a Realtime qPCR assay. Aguayo et al. (2017) considered that VCGs 01213-01216 and 0121 both represented TR4, even though VCG 0121 is uncommon in areas in which 01213-01216 has devastated Cavendish. Despite its genetic similarity to VCG 0121301216, VCG 0121 does not seem to possess the epidemiological competence and aggression that makes strains in VCG 01213-01216 so destructive; if it did, VCG 0121, rather than VCG 01213-01216, might be expected to predominate, or at least be more prevalent, in Southeast Asia. However, VCG 0121 has been found in only a handful of locations in Indonesia, Taiwan and Western Malaysia. Better understandings of the pathogenic fitness of VCG 01213-01216 and other populations of this pathogen are needed, as they may enable improved means for managing this disease. A PCR diagnostic measure, which is based on a specific portion of the intergenic spacer (IGS) region of ribosomal DNA, has been used to diagnose VCG 01213-01216 (Dita et al., 2010). It was screened against a wide array of F. ox­ ysporum f. sp. cubense and other Fusarium isolates and a commercial diagnostic kit has been developed for its use (Clear Detections, The Netherlands). High copy numbers of this housekeeping locus exist in the pathogen genome, enabling the detection of low levels of the pathogen. The diagnostic test of Dita et al. (2010) has been used successfully by others (Maryani et  al., 2015; Ploetz et al., 2015a; O’Neill et al., 2016; Ordóñez et al., 2016; Aguayo et al., 2017). However, it detected a non-pathogenic endophyte of banana in Ecuador (Magdama and Jimenez-Gasco, 2015), and falsely detected the pathogen in Mareeba, Queensland, Australia (Deloitte Australia, 2015). A different set of primers for the IGS region than those used by Dita et  al. (2010) were used in a real-time fluorescence loop-mediated isothermal

amplification (LAMP) assay developed by Zhang et al. (2013). The diagnosis of plant pathogens with DNA regions, such as the IGS, that are not linked to pathogenicity has been criticized, as it could result in the erroneous detection of non-­pathogens that possess the same DNA sequence (Aguayo et al., 2017). Clearly, a TR4 strain-specific region that has a role in the pathogenicity of this clone could enable a more specific, albeit less sensitive, assay (copy numbers of such genes are typically low). As mentioned above, the SIX (secreted in xylem) effectors have been investigated for this purpose (Fraser-Smith et al., 2014). The recent outbreak of Fusarium wilt on Cavendish cultivars in North Queensland provides a useful case study in how a new incursion by TR4 could be handled. O’Neill et  al. (2016) confirmed the presence of TR4 by both the IGS diagnostic test of Dita et al. (2010) and traditional VCG analysis. Soon after this detection, affected plants and asymptomatic plants in the immediate vicinity were treated with glyphosphate. Plants were not removed and care was taken not to move any infested material off the property. Recently, with the fiscal support of the banana industry, the entire farm was purchased and taken out of banana production. Most growers in the region now take sanitary precautions to prevent the movement of propagules of Fusarium wilt with disinfectant dips for footwear, car and truck tyres at the entrances/exits to properties, facilities for removing dirt from vehicles, etc. (Australian and Queensland Governments, 2016a). Nonetheless, TR4 was recently detected on another property in Queensland. Clearly, even when the best measures are used TR4 can spread. In other areas, varying degrees of management and success are evident. For example, approaches that are used to manage the disease on Mindanao Island in the Philippines vary widely in their rigour and attention to detail. Predictably, the disease spreads most rapidly on farms in which lax measures are used to restrict traffic within plantations, and damage is worse on farms in which strict foot and tool disinfestation is not practised and where new outbreaks are not isolated promptly with fencing. On the best farms, affected plants are killed with herbicide but are not removed, to avoid disseminating the



Fungal Diseases of the Root, Corm and Pseudostem

pathogen. In general, commercial production continues only on well managed farms. Cultural methods Soil sterilization has been attempted in plantations in Mindanao in the Philippines. Sawdust and rice hulls have been burnt on and around sites where diseased plants have been found. ­Although this measure does not eliminate the pathogen (Plate 3.8), its reduction of the pathogen in soil is thought to be valuable ­ (L.  Trueggelmann, Philippines, 2017, personal communication). Treatment of irrigation water with 2–3 ppm chlorine has also been shown to kill the pathogen (L. Trueggelmann, Philippines, 2017, personal communication). Pathogen exclusion is essential in pathogen-­ free regions. Fusarium oxysporum f. sp. cubense cannot be eradicated from an area once it is

225

i­nfested, and exclusion of the pathogen from non-infested plantations can be very difficult once it moves into a region. Effective quarantine measures are necessary, and regional awareness and contingency programmes, such as those created in the Western Hemisphere, should be considered in all threatened areas (Ploetz et al., 2015a). Although numerous publications indicate that different methods can be used to manage this disease (Thangavelu and Mustaffa, 2012), there are actually few effective measures. Susceptible banana cultivars can usually be grown only if pathogen-free propagation materials are used in pathogen-free soil. Tissue culture-­derived plantlets are the most reliable source of clean material. Even though they are more susceptible to Fusarium wilt than traditional banana seed pieces (Smith et al., 1998), they should be used to propagate this crop whenever possible.

Plate 3.8.  A Cavendish plant with symptoms of Fusarium wilt at upper left indicates the failure of rice hull burning to control Fusarium oxysporum f. sp. cubense tropical race 4 in Mindanao, Philippines (photo: R.C. Ploetz, UF).

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In subsistence agriculture or other situations in which their expense is an issue, tissue-culture plantlets have been used to start nurseries for the production of pathogen-free conventional seed pieces (Lule et al., 2013). Soil health, and soil structure, permeability, mineral content and microbial attributes are recognized as key components of ‘healthy soils’ (Turner and Rosales, 2005; Pattison and Lindsay, 2006). Plantation productivity and the response to some pathogens (e.g. nematodes) is enhanced in healthy soils. Pattison et al. (2014) reported that the severity of Fusarium wilt on ‘Pisang Awak’ was reduced and that yield was increased when pinto peanut (Arachis pintoi) was grown as a ground cover. Pinto peanut reduced water stress, which contributed to disease development, in the banana plants. Although differences between plots with and without ground cover were not great and ‘Pisang Awak’ was not tested in TR4-infested land, Pattison et al. (2014) concluded that a holistic combination of cultural measures and disease tolerance might be useful when combatting TR4. Chemicals and biocontrol agents The perennial nature of banana and the need for long-term efficacy are seldom considered when management strategies are developed for this disease (Ploetz, 2015a, b). Much of the literature on this topic reports results from short-term in vitro or greenhouse studies with no indication that the results would be useful in the field. For example, even though many biological control studies have been conducted, none of the published results indicate that cost-effective management in the field is possible via this approach (Ploetz, 2015a). Likewise, limited or questionable efficacy has been associated with chemical, cultural and physical measures (Ploetz, 2015a). Chemical disinfestants have been examined for cleaning tools, machinery and use in footbaths (Nel et al., 2007; Meldrum et al., 2013b). At the shortest time interval that they tested (30 s), a product that contained quaternary ammonium, Sporekill® (International Chemicals), was more effective than Farmcleanse® (Castrol) in inhibiting microconidium germination of TR4 (Meldrum et  al., 2013b); another product, Domestos® (Unilever), was generally as effective as Sporekill®, but it contained sodium hypochlorite

and was corrosive. Sporekill® remained effective after 6 months of exposure to heat and sunlight, which indicated its potential as a footbath treatment. Nel et al. (2007) also observed significant reductions in microconidium germination of SR4 when different quaternary ammonium products were tested, albeit with much longer exposure times (5 min). Clearly, exposure time should be considered when evaluating these products, as it would be unreasonable to expect labourers to spend much time disinfesting tools or their shoes. Disease-resistant clones In F. oxysporum f. sp. cubense-infested soils, resistant cultivars have been the only consistently effective tool for managing this disease. Cultivars that resist different races of the pathogen exist (Buddenhagen, 1990), but may not be profitable in a given situation. Xu et  al. (2011) analysed the economics of producing race 1- and TR4-­ resistant cultivars in China. They reported that market preferences and the race that was present in infested fields determined which cultivars would be profitable. In TR4-infested soils in which lower rents were charged, Xu et al. (2011) recommended replacing susceptible cultivars with other crops or resistant cultivars. There is a critical need for resistant bananas that meet local and export market standards (Ploetz et al., 2015b). The susceptibility of the major cultivars to different races is well understood (Ploetz and Pegg, 1999), but few are resistant to TR4 (Walduck and Daly, 2007; Ploetz, 2015a). Dessert-banana cultivars (Cavendish (AAA), ‘Pisang Mas’ (AA), ‘Pisang Berangan’ (AAA), ‘Mysore’ (AAB), ‘Silk’ (AAB) and ‘Pome’ (AAB)) and cooking-banana cultivars (‘Mai’a Maoli’ (AAB), ‘Bluggoe’ (ABB) and ‘Pisang Awak’ (ABB)) are susceptible, but there are notable exceptions. Recent results indicate that two other subgroups that are important staple crops in Africa may have useful resistance to TR4. Although the reactions of plantain hybrids from the IITA and FHIA breeding programmes (respectively, ‘Bita-3’ and ‘FHIA21’) suggested that the plantains were vulnerable (Ploetz and Evans, 2015), plantain cultivars have generally done well in screening trials in China, Indonesia and Mindanao (Huang et al., 2005; Molina et al., 2012). Likewise, Molina et al. (2014) indicated that East African highland



Fungal Diseases of the Root, Corm and Pseudostem

banana cultivars (AAA, Lujugira-Mutika subgroup) were also resistant to TR4, based on a study in Mindanao and another in China. These are encouraging results, but few trials in only a few locations have been documented. Additional studies, which could be reported in peer-reviewed journals, are needed over multiple production cycles. Banana improvement programmes, many of which were established to improve Fusarium wilt resistance in the crop, have been established throughout the banana-growing world (Buddenhagen, 1993; Ortiz, 2013). These programmes face enormous challenges. Primitive diploids that have been used as parents usually have very poor agronomic and fruit traits, and introgression of disease resistances that they possess into advanced lines can take several generations. The polyploid nature of the crop, genetic abnormalities that exist in many parental lines, the need for end products to be parthenocarpic and sterile, the low fertility of cultivars that need improvement, and the expense of evaluating large hybrid populations in field settings are additional hurdles that impede progress (Ortiz and Swennen, 2014). Resistance to TR4 is found in bred hybrids, especially those developed by the programme at the Fundación Hondureña de Investigación Agrícola (FHIA) in Honduras. The FHIA hybrids resist Fusarium wilt, as well as black leaf streak and the burrowing nematode, and are important in Africa (Nowakunda and Tushemereirwe, 2004; Dzomeku et al., 2007; Gaidashova et al., 2008; Njuguna et al., 2008; Uazire et al., 2008) and Cuba (Alvarez, 1997; Alvarez and Rosales, 2008). Unfortunately, they do not meet the high standards that are needed for an export banana. For example, the high-yielding dessert clones ‘FHIA-01’ (AAAB, ‘Goldfinger’) and ‘FHIA-02’ (AAA, ‘Mona Lisa’) had lower pulp-to-peel ratios, were not as sweet, and had lower overall consumer acceptance than the Cavendish cultivars ‘Grand Nain’ and ‘Williams’ (Dadzie, 1998). New banana clones are needed that resist TR4 and other races of F. oxysporum f. sp. cubense (Rowe and Rosales, 1999; Ortiz and Swennen, 2014). Recently, hybrids of Cavendish (formerly thought to be sterile) were produced at FHIA with the objective of producing new dessert clones (Aguilar Morán, 2013). Whether useful clones could be produced via this approach remains to

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be seen. Since prior results with ‘Gros Michel’ indicate that disease resistance and desirable fruit characteristics are lost or changed during meiosis, creating a Cavendish-like clone via conventional breeding with a Cavendish parent, which also has resistance to TR4, will be a daunting task. Ideally, new bananas, which do not necessarily look or taste like Cavendish, need to be accepted by consumers. However, with improved understandings of the origins and genetic backgrounds of Cavendish and other important clones (Perrier et al., 2011), it may be possible to reconstitute Cavendish cultivars that are productive and resist TR4 and other problems while retaining the appearance and sensory attributes that consumers associate with these fruit. Cavendish production has a long history in Taiwan (Hwang and Ko, 2004). In the 1980s, S.C. Hwang began evaluating somaclonal variants of ‘Giant Cavendish’ for response to Fusarium wilt. By recurrently selecting tissue-cultured lines in heavily infested fields, he developed variants with improved resistance to this disease. An array of the Giant Cavendish Tissue Culture Variants (GCTCVs) have been produced, several of which are now widely tested and used in Asia (Hwang and Ko, 2004). Cavendish somaclones are also being developed elsewhere. For example, ‘Guijiao No 9’ is a ‘Dwarf Cavendish’ somaclone developed in Guangxi, China (Wei et al., 2016). In general, the Cavendish somaclones are not completely resistant and can usually be grown for no more than two cycles in TR4-infested sites. Poor finger and hand architecture of fruit produced by these lines further complicates their use by the export trades, although the latter traits are not as important for local markets, as in China, or for a preferred market that Taiwanese fruit have in Japan. Nonetheless, the Cavendish somaclones are currently the best TR4-resistant alternatives for standard TR4-susceptible Cavendish cultivars. After decades of research on managing Fusarium wilt of banana, effective alternatives to the use of resistant cultivars have not been found (Ploetz, 2015a, b). In order to manage this disease biologically, chemically or culturally, better understanding would be needed of the factors that are responsible for disease development and how they might be manipulated for disease control. Although the complex

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interactions that occur between F. oxysporum f. sp. cubense and banana are only beginning to be understood (Li et al., 2013b; Bai et al., 2014; Wang et al., 2015), virulent races of this hemibiotrophic pathogen induce apoptosis (programmed cell death) in susceptible cultivars during the necrotrophic phase of host infection (Paul et  al., 2011). Transgenic banana plants with anti-apoptosis genes have enhanced resistance to this disease (Paul et  al., 2011). ­Recently, Magambo et  al. (2016) reported transgenic, anti-apoptotic lines of ‘Sukali Ndizi’ with enhanced resistance to race 1 in glasshouse trials in Uganda, and J. Dale (Australia, 2016, personal communication) observed TR4 resistance in anti-apoptotic lines in the field in the Northern Territory of Australia. Genetic transformation of banana has become commonplace and disease resistance is one of the most sought-after traits (Khanna et al., 2007; Remy et al., 2013; Ortiz and Swennen, 2014). Genetic transformation to create resistant genotypes is particularly important when improvement targets, such as the Cavendish banana, are difficult to improve via conventional breeding. Although a range of transgenes have been tested against race 1 (Subramaniam et al., 2006; Paul et al., 2011; Ghag et al., 2012; Magambo et al., 2016) and TR4 (Yip et al., 2011; Mahdavi et  al., 2012; Hu et  al. 2013), results from field trials have only recently been reported (Dale et al., 2017). As for other disease-control measures, long-term efficacy in the field should be demonstrated for these banana clones before they can be deemed useful. However, even if durable resistance is evident, other agronomic and organoleptic criteria must be met before transgenic bananas would be used as replacements for Cavendish or other important cultivars. Most importantly, consumers must accept these products as safe food. TR4 threatens export and smallholder production of Cavendish in tropical America. Preparation for its arrival in the region has begun. For example, stakeholders have been informed about TR4 (Pocasangre et al., 2011) and responses have been formulated for its possible arrival in the Western Hemisphere (Dita Rodriguez et al., 2013). Similar contingency plans are needed in Africa, the Indian subcontinent and other areas that maybe affected in the future.

Armillaria Corm Rot D.R. Jones Armillaria corm rot of banana is uncommon. It has been recorded in Australia (Blake, 1963), Kenya (Department of Agriculture, 1964; Kung’u, 1995), Malawi (Spurling and Spurling, 1975), Uganda (Hansford, 1945), South Africa (Brodrick, 1971) and Florida, USA (Rhoads, 1942). It is usually associated with new plantations that have been established on recently cleared land. In Australia, the disease was worst where hardwoods were cleared and was rare on bush land (Magee and Eastwood, 1935). The disease has also been called dry rot or stump rot (Stover, 1972). Armillaria mellea and Desarmillaria tabescens (syn. Armillaria tabescens) in the order Agaricales are the causes of the disease (Blake, 1963; Department of Agriculture, 1964). These pathogens are woodland inhabitants and colonize tree stumps or large roots left behind after clearing. Armillaria mellea forms a network of rhizomorphs, which are compacted masses of hyphal strands covered by a melanized outer cortex, under the bark of trees. Thick, creamy white fanlike sheets of mycelium are also found. Desarmil­ laria tabescens does not usually form melanized rhizomorphs in nature (Mihail et al., 2002), but unmelanized rhizomorphs can be formed in culture (Koch et al., 2017). The fungi penetrate the banana’s roots and corm at or below ground level. When the corm is fully invaded, leaves turn yellow and die from the base upwards. If only part of the corm is invaded, only the leaves on the affected side might die. The plant is easily pushed over or breaks off at ground level in the advanced stage of the disease. Rhizomorphs and white fungal strands can be seen permeating the dry, brown corm. Sometimes, fruiting bodies of the fungi are found at the base of the plant (Stover, 1972). Desarmillaria species lack an annulus, a remnant of a partial veil that remains after the pileus expands, at maturity, whereas Armillaria species have a robust annulus at maturity (Koch et al., 2017). Armil­ laria mellea has been described by Pegler and Gibson (1972). To avoid the disease, all tree stumps and large roots should be burnt or cleared from the



Fungal Diseases of the Root, Corm and Pseudostem

plantation site before planting (Blake, 1963). If corm rot is a problem, diseased corms should be dug up together with surrounding plants, cut into pieces and burnt. Replanting should take place some distance away from the position of the affected plant. Control measures adopted in South Africa after the first outbreaks included the removal of infected plant material followed by disinfection of infested and surrounding soil with 0.5 kg methyl bromide/10 m2 applied under plastic cover for 24 h (Brodrick, 1971).

Cylindrocladium Root Rot D.R. Jones Introduction In the late 1980s and 1990s, fungi in the Cylin­ drocladium genus were reported together with endoparasitic nematodes as components of a soilborne pathogenic complex causing root and corm rots in banana plants in the French West Indies (Loridat, 1989), Costa Rica (Semer et al., l987), Côte d’Ivoire (Kobenan, 1991) and Cameroon (Castaing et al., 1996). The intensity of necrosis caused by each component of this disease complex was found to vary with respect to climate and soil characteristics. Both Radolphus si­ milis and the Cylindrocladium spp. individually caused a typical necrotic response, but symptoms were enhanced when the two pathogens interacted.

Symptoms Symptoms consisted of brownish to blackish lesions on the underground corm and root system. These necrotic lesions resulted in a decrease in nutrient uptake, root breakage and plant toppling (Risède and Simoneau, 2004). The decline in the vigour of root systems led to less yields and a consequent drop in production (Loridat, 1989). Tests revealed that necrotic lesions, the first of which were prominent 4 days after inoculation, started as flecks 1–5 mm long that gradually enlarged in length, width and depth. They eventually developed a well-defined oblong

229

shape. Individual lesions could coalesce to rot large areas of root. Symptoms were found on primary, secondary and lateral roots (Risède and Simoneau, 2004).

Causal agents By traditional taxonomic analysis, sequence analysis of the internal transcribed rDNA spacer (ITS) region and RFLP of the rDNA spacer region (IGS RFLP), two major Cylindrocladium taxa were found in association with the necrotic lesions on banana (Risède and Simoneau, 2001). One taxon grouped Cylindrocladium gracile-like isolates that showed atypical Cy. gracile morphology (large, cylindrical, monoseptate condia and clavate vesicles, but close phenetic clustering with typical Cy. gracile reference isolates). The other taxon was shown to be Cy. spathiphyl­ li. In addition, three other rare taxa were discovered that were distinct from the other two. The most common were Cy. scoparium-like, which were found in Cameroon, Martinique and Guadaloupe. One isolate was Cy. floridanum-like and another was typical of Cy. gracile. Both of these isolates were from Cameroon. The authors had previously described Cy. pteridis as the causal agent of the disease in Cameroon, but later decided that these isolates more appropriately belonged to Cy. gracilis-like pathogen. Although the Calonectria sexual morph of Cy. spathiphylii could be obtained by crossing banana isolates, no sexual morph could be obtained from attempts to cross Cy. gracilis-like banana isolates. The authors’ next steps were to determine the relative pathogenic contribution each taxon made to the root-rot lesion problem on banana and the genetic variation of each taxon shown to be pathogenic on banana (Risède and Simoneau, 2004). In this endeavour, 104 isolates from Cameroon, Costa Rica and the West Indies were compared. The results of pathogenicity tests on ‘Grande Naine’ showed that isolates of Cy. spathiphylli were highly pathogenic while Cy.  gracile-like isolates were moderately pathogenic. The isolates that were Cy. scoparium-like, Cy. floridanum-like and Cy. gracile were only very weakly pathogenic. In Costa Rica, the fungus component of the root rot complex was originally named Cy. musae

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(Semer et  al., 1987), but this name was never validly published. Risède and Simoneau, (2004) showed that isolates from Costa Rica were Cy. spathiphylli. Crous et  al. (2006) classified a number of Cylindrocladium isolates on the basis of their morphology and DNA sequence data from their β-tubulin and histone gene H3 regions. They identified the Cy. gracile-like isolates of Risede and Simoneau (2001, 2004) as Cy. flexuosum, which they found to be more morphologically variable than expected. In addition to Martinique and Guadeloupe, this species was reported on banana from St Lucia. Under the one name for one fungus nomenclature, Calonectria spathiphylli, the sexual morph in the old dual name nomenclature, is now the species name for Cy. spathiphylli and Calonectria clavata is the species name for Cy. flexuosum (Crous et al., 2006; Marin-Felix et al., 2017). Calonectria spathiphylli was described by Crous (1993, 2002). Macroconidiophores consisted of a stripe, a pencillate arrangement of fertile branches, a stipe extension and a terminal vesicle. The stipe was septate, hyaline and smooth with dimensions of 50–160 × 5–6 μm. The stipe extensions were septate straight to flexuous, 170–326 μm long, 3–4 μm wide at the apical septum and terminated in a globose or ellipsoid to obpyriform 8–15 μm wide vesicle. The conidiogenous apparatus was 60–150 μm long, 40–90 μm wide with 18–40 × 4–6 μm singleor two-celled primary branches. Secondary branches were single-celled and measured 18– 30 × 4–6 μm. Conidia were straight, cylindrical and rounded at the ends with dimensions of 45–120 × 6–7 μm. Perithecia were orange to red in colour, subglobose to ovoid in shape and 380–655 μm high with a diameter of 340–650 μm. Asci were clavate, two- to eight-spored and measured 120–230 × 7–25 μm. Ascospores, which aggregated in the upper third of the ascus, were hyaline, guttulate, fusoid with rounded ends, one- to three-septate with dimensions of 22–65 × 3–7 μm (see Fig. 3.2). Calonectria clavata from banana was described by Crous et al. (2006). Macroconidiophores consisted of a stipe, a penicillate arrangement of fertile branches, a stipe extension and a terminal vesicle. The stipe was septate, pale brown at base, hyaline, smooth, septate and 60–260 × 5–7 μm. The stipe extension was septate, straight to

­ exuous, 120–450 μm long, 3–4 μm wide at the fl apical septum, terminating in a narrowly clavate vesicle, which was 4–5 μm in diameter. The conidiogenous apparatus was 70–120 μm long, 25–60 μm wide. Its primary branches were aseptate or uniseptate with dimensions of 30–65 × 4–6 μm. Secondary branches were aseptate or one-septate with dimensions of 30–50 × 3–6 μm. Tertiary and quaternary branches were aseptate and measured 15–30 × 3–5 μm. Each terminal branch produced one to four phialides. The phialides were elongated doliiform to reniform, hyaline, aseptate and measured 10–20 × 4–5 μm. The apex had minute periclinal thickening and inconspicuous collarette. Conidia were cylindrical, rounded at both ends, straight with dimensions of (55–)68–75(–95) × (5–)6(–7) μm (average = 70 × 6 μm), one-septate (but up to 5-septate at germination) and lacked a visible abscission scar. They were held in parallel cylindrical clusters by colourless slime. The sexual morph was not obtained in culture. Disease cycle and epidemiology The main species of Calonectria attacking banana in Martinique was found to be Ca. clavata. This fungus produces tan to brownish colonies with irregular margins on 1% malt extract agar. Numerous chlamydospores arranged in chains and brown microsclerotia are produced. Investigations also showed that the fungus could be trapped in soil with tissue-cultured banana plants. Microsclerotia have been detected in naturally infected roots in the field and also in artificially inoculated banana roots. They have been shown to be capable of infecting roots and may be the main survival structure of the fungus in soil. In Cameroon, Ca. spathiphylli is the most common species (J.M. Risède, Guadeloupe, 1999, personal communication). The soil type was found to be an important factor determining the distribution and density of this fungal soilborne pathogen. Calonectria was absent in clay soils rich in smectite, while it prevailed over other soilborne pathogenic fungi in perhydrated andosols on volcanic ash (Delvaux and Guyot, 1989; Loridat, 1989; Risède, 1994). Protocols for the isolation of Calonectria from banana roots and soil samples have been described (Risède, 2008).



Fungal Diseases of the Root, Corm and Pseudostem

231

A

C

B D

Fig. 3.2.  Calonectria spathiphylli (syn. Cylindocladium spathiphylli). Key: A = conidiophore, vesicles and conidia; B = asci; C = chlamydospores; D = ascospores; scale bar = 10 μm (Crous, 1993).

Host reaction Many different banana cultivars were found to be affected in the important Cavendish subgroup

beside ‘Poyo’ and ‘Grande Naine’ (Loridat and Ganry, 1991). Risède and Simoneau (2004) compared the progress of symptom development caused by Ca.

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spathiphylli and Ca. clavata (as Cy. gracile-like isolate) on ‘Grande Naine’ and five other banana cultivars. All banana cultivars inoculated with the isolates showed symptoms after 4 days though with different levels of severity. As root lesions expanded on a particular cultivar, Ca. spathiphylli always caused more damage at any one moment in time than Ca. clavata. The most susceptible cultivar to both isolates was ‘IDN110’ (AA) with ‘Pisang Madu’ (AA) the most resistant. ‘Grande Naine’ (AAA) and ‘Kunnan’ (AB) were intermediate in their response. Semer et al. (1987) reported chemical control of Ca. spathiphylli (as Cy. musae) by benomyl and prochloraz in greenhouse studies. Some antagonistic fluorescent pseudomonads isolated from the rhizosphere of banana plants in Martinique have been shown to inhibit Calonectria in vitro. A strain of Pseudomonas putida was found to totally block fungal growth (Sutra et  al., 2000). Pre-inoculation of banana roots with arbuscular mycorrhizal fungi in glasshouse experiments significantly reduced the effect of Ca. spathiphylli infections (Declerck et al., 2002). Silicon applied to banana reduced root necrosis caused by Ca. spathiphylli by 50% when measured 14 days after application. Growth reduction was also alleviated. These results suggested to the authors that silicon could have a positive effect on the resistance of banana to Ca. spathip­ hylli and provide an environmentally friendly alternative to pesticides (Vermeire et al., 2011).

Rosellinia Root and Corm Rot D.R. Jones Rosellinia bunodes was found by Smith (1929) attacking the root system of banana in Jamaica. However, the existence of another species of Rosellinia in the disease complex was suspected. Rosellinia bunodes, which causes a black root rot disease of many plant species, but mainly of tropical and subtropical woody hosts, is widespread in tropical America and has been found in Central Africa, South Asia and Southeast Asia (Sivanesan and Holliday, 1972). Rosellinia bunodes is a relatively weak pathogen that survives as a saprophyte in many soils.

It affects plants that are debilitated by nutrient imbalances. The fungus usually enters roots as secondary invader after insect or nematode attack. Good soil management has been seen as the key to the problem. Soils that are acidic and have high humidity and/or organic matter levels have been associated with the disease. Pruning and good drainage reduce humidity levels under the canopy. Liming increases soil pH and the removal of organic material from the soil surface helps in control. A Rosellinia sp. was also implicated in a ‘stinking root’ disease of banana in coastal areas of southern Brazil in 1937 by R.D. Gonçalves (Wardlaw, 1961). As in Jamaica, other pathogens were associated with the problem. Fusari­ um spp. were also isolated and unusual sclerotial formations were seen in affected plantations. Rosellinia pepo has been identified as causing ‘star-like wound’ disease of ‘Dominico-­ Hartón’ (AAB, Plantain subgroup) and other clones in Colombia (Belalcazar, 1991). Symptoms on banana include an internal reddish ­necrosis of the corm, necrosis of leaf margins, lower yields and the eventual toppling of plants following the decay of the root system. White, star or fan-shaped patterns of mycelium, which can be seen underneath the outer cortex of the corm and roots, helps distinguish this disease from Fusarium wilt. Rosellinia pepo is a soil inhabitant and is found in organic litter and woody debris. It attacks many plant species, including avocado, breadfruit, cocoa and coffee, but is thought to have a narrower host range than R. bunodes and a more restricted distribution (Booth and Holliday, 1972). In Colombia, it is common on plantain grown as an intercrop with coffee or cocoa and its incidence is greatest where plant residues are highest, such as on land recently cleared of trees. Cultivars in the AAA genomic group are not as susceptible as plantain (­Belalcazar, 1991). The biology and control of R. bunodes and R. pepo have been reviewed (ten Hoppen and Krause, 2006). Because of poor results from field tests utilizing fungicides and biocontrol agents, the authors believed that cultural methods, such as those mentioned above plus the isolation and burning of affected plants, are still the best means of control.



Fungal Diseases of the Root, Corm and Pseudostem

Rosellinia bunodes and R. pepo have been described by Sivanesan and Holliday (1972) and Booth and Holliday (1972), respectively.

Damping-off of Musa Seedlings D.R. Jones A rot that attacks the lower leaves, pseudostem and roots where they join the corm has been reported to affect young, succulent seedlings of Musa species. The plants were frequently killed and fell over, giving the appearance of damping-off (Stover, 1972). A necrosis of the tip of the lower leaf, often followed by the death of the  leaf, was the first symptom of the disease. The rot then progressed to the pseudostem and caused a dark brown discoloration of the leaf sheaths. The rot extended to the inner sheaths and the seedling collapsed and died. Sometimes the discoloration of the leaf sheaths began at or below ground level and moved down to where the leaf sheaths were attached to the developing corm and up to the crown. On occasion, the roots were affected. The disease could also ­affect the tips and base of the coleoptile of seedlings only a few centimetres long. These ­became brown as they emerged from the soil. When conditions were not favourable for disease development, only the older leaves died and one or two leaf sheaths became discoloured. The seedling continued to grow, but was retarded. In some instances, the disease remained below ground and the plant was stunted as a result of root and rhizome injury associated with the necrosis of adjacent leaf sheaths. Infection was greatest during periods of cool, wet weather. As seedlings became older and hardier, only the outer leaf sheaths were affected (Stover, 1963). This damping-off disease was caused by Corynespora torulosa (Crous et  al., 2013) (syn. Deightoniella torulsa) (Stover, 1963), which also affects banana, abacá and enset leaves, abacá pseudostems and banana fruit. Seedlings of the wild species M. acuminata ssp. bank­ si, M. balbisiana, M. schizocarpa and M. textilis were reported as affected, as were seedlings arising from crosses between M. acuminata and

233

edible banana cultivars (Stover, 1972). The problem was controlled by thinning the stands of seedlings, reducing humidity levels and maintaining conditions that allowed the foliage of seedlings to dry rapidly after watering. Banana trash and banana plants with Deightoniella leaf spot were kept well away from seedlings. The disease was of little economic importance and occurred only in the nurseries of banana breeding programmes where hybrid ­ ­seedlings were grown in large quantities.

Deightoniella Pseudostem Rot of Abacá E.O. Lomerio Introduction This disease was first described on the island of Luzon in the Phillipines in 1927 and was reported on the island of Mindanao in 1931 (Agati et  al., 1934). It has been serious in plantations of the cultivar ‘Bongolanon’ and has been reported as causing severe damage to seedlings, but is generally regarded as of minor economic importance. The quality of abacá fibre is lowered because the disease causes an increase in the percentage of dark strands. Severely diseased pseudostems become useless for stripping. In the Philippines, the disease is also known as trunk rot and red-sheath rot.

Symptoms The disease is more severe on the pseudostem of abacá than on the leaves. Lesions about the size of a pinhead form first on the outer leaf sheaths. These have dark brown centres, surrounded by a lighter area, which is delimited by a brown marginal line. As lesions increase in size, they coalesce and become depressed in the middle. Enlargement is more rapid along the axis of the pseudostem so that, in advanced stages, a dark brown to black oval–oblong blemish is produced. Green-brown superficial mycelium of the fungus can be seen in the centre. The disease also progresses inwards, affecting one leaf sheath after another until the stem is girdled

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and sufficiently weakened for it to bend over under the weight of the foliage. Affected leaf sheaths become desiccated and curl backwards and downwards (Ela and San Juan, 1954; Wardlaw, 1961; Stover, 1972).

high-density plantings and proper spacing reduces incidence. Seriously affected plants should be cut down and burned. Losses are minimized if mature plants are harvested before the rot becomes severe (Ela and San Juan, 1954; Stover, 1972).

Causal agent Pseudostem rot of abacá is caused by Corynespo­ ra torulosa (Crous et al., 2013) (syn. Deightoniella torulosa). This fungus has been described under Deightoniella leaf spot of banana, abacá and enset. It also causes a damping-off of Musa seedlings and affects banana fruit.

Disease cycle and epidemiology In the Philippines, the disease is more severe in the rainy season. Cornespora torulosa is a common saprophyte on dead and dying abacá leaves. Spores of the fungus are also found in association with leaf spots and diseased pseudostems. The fungus sporulates at night under conditions of 100% relative humidity and conidia are violently discharged the next morning as the humidity falls and temperatures rise (Meredith, 1961). Germination takes place in water or at 95% relative humidity and above. Wounding the leaf sheath favours infection. Wounds on the pseudostem occur naturally when leaves buckle with age, when sheaths are loosened as older leaves fall and when collapsed leaves are blown by the wind. If wounds are inoculated, the first symptoms appear 15 days after inoculation (Arce, 1954, reported by Stover, 1972). Host reaction All cultivars become infected, but some show field resistance. ‘Bongolanon’ is the most susceptible cultivar (Stover, 1972). Control Serious damage is usually associated with poor cultural practices involving spacing, pruning and weed control. The disease spreads rapidly in

Fungal Root Rot D.R. Jones and R.H. Stover Many genera of fungi have been isolated from rots in the roots of banana. However, there has usually been no proof that the fungi found have been pathogenic. From 1964 to 1966, studies were made of the fungi present in lesions on banana roots and of their pathogenic potential (Laville, 1964; Blake, 1966; Mallessard, 1966; Stover, 1966). Root rot consists of the rotting and dieback of the tips of the main root and the blackening, necrosis and death of the smaller lateral feeding roots. It is common even in the absence of nematode injury (Stover, 1972). Fungi consistently isolated from banana root rots have included Fusarium solani, F. ox­ ysporum and Rhizoctonia sp. However, there has been no proof that F. solani and F. oxysporum (other than f. sp. cubense) are important pathogens by themselves. It is likely that they extend cortical necrosis in nematode lesions, which may reach the stele. In Australia, root lesions caused by the burrowing nematode were found to be more extensive when F. oxysporum was present (Blake, 1966). In Central America, where F. ox­ ysporum is rare in root lesions, F. solani is believed to enlarge lesions caused by nematodes. Rhizoctonia is capable of invading young, intact roots and causing necrosis, but damage is greater if the roots are wounded (Mallesard, 1966; Stover, 1966). Some isolates of Rhizocto­ nia have been shown to be highly pathogenic on Musa seedlings (Stover, 1966), but not all isolates are pathogenic on banana roots. Stover (1972) reported that all isolates from banana roots in Central America were binucleate Rhizoctonia species (BNR) and therefore not the same species as the multi-nucleate R. solani, which is common on other crops. The binucleate members of the Rhizotonia species complex



Fungal Diseases of the Root, Corm and Pseudostem

are designated anastomosis groups (AG) A to U. Multinucleate isolates of R. solani are designated AG1 to AG13. During a root disease survey of banana ­cultivars in Georgia (USA), root lesions and root rot were observed on ‘FHIA-01’ (AAAB, ‘Gold ­Finger’ – bred hybrid), ‘Kandarian’ (ABB, syn. ‘Kandrian’/‘PNG 148’), and ‘Manzano’ (AAB, Silk subgroup). Root lesions were dark brown to black and irregular in shape, with partial or entire roots affected. Lateral roots and outer layers of roots arising from interior layers of the corm of infected plants were blackened and rotted. Isolates were identified as a Ceratobasidium sp. AG-F on the basis of morphological and molecular characteristics plus anastomosis group typing (Yin et al., 2011). AG-G, which is known as Ceratobasidium sp. (Rhizoctonia fragariae), has been described in Cuba as the cause of a root necrosis that develops in a stem and central leaf rot in tissue culture-derived plants of ‘Pisang Awak’ (ABB), ‘FHIA-21’ (AAAB – bred hybrid) and ‘FHIA-23’ (AAAA – bred hybrid) growing in nurseries on a substrate of sugarcane filter press and 10% of zeolite. A mild yellowish discoloration appeared on the margins of the lower leaves that later turned to a characteristic deep yelloworange-reddish colour. The affected plants showed between 10% and 30% of necrotic roots with different levels of severity. Internally, the infection spread from the roots to the developing corm, which became completely necrotic. Growth was slowed and affected plants plants died (Perez et al., 2009). It is possible that under the right conditions many strains of the Rhizoctonia species complex can infect any plant root, but some may be specialized and account for most of the infections on a particular host. To determine which strain or strains predominate on banana, survey work with molecular tools needs to be undertaken (J. Woodhall, USA, 2017, personal communication). Phythium and Phytophthora are isolated infrequently from banana root lesions and are thought not to play an important role in banana root rot (Stover, 1972). However, a Pythium sp. has been implicated in a damping-off disease of abacá seedlings (Roldan, 1933). A species of Zythia has been found to be pathogenic on banana roots in Côte d’Ivoire (Kobenan et al., 1997). Root rots are common when conditions are unsuitable for the optimum growth of the banana

235

plant, such as when soils are waterlogged, compacted or nematode-infested or have a 40% or more clay content (Stover, 1972). In a healthy root system, root rots only affect a very small proportion of the roots. Large, extensive and deep root systems develop in well drained, deep, friable loams. Root rot is less likely to occur if nematodes are controlled and if soil structure and moisture content permit plants to grow under optimum conditions.

Marasmiellus Pseudostem and Root Rot D.R. Jones and E.O. Lomerio Introduction This disease affects banana and abacá and has been reported from Australasia–Oceania (Australia, Papua New Guinea, Fiji, Hawaii), Asia (Philippines, Sri Lanka, East Malaysia), Africa (Mauritius, Uganda, Sierra Leone, Côte d’Ivoire, Ghana) and the Latin America–Caribbean region (Jamaica, Trinidad, Windward Islands, Brazil). The causal agent is a saprophyte of decaying vegetation, including leaf and pseudostem trash. It attacks banana and abacá that have been weakened by drought or poor cultural conditions and is rare in well managed plantations. The disease is reported to be more common on sandy or gravelly areas, but is also reported in inadequately drained soils (Wardlaw, 1961; Stover, 1972).

Symptoms The outer leaf sheaths and leaves of both banana and abacá become dry. Growth is stunted and new leaves are either slow to emerge or fail to emerge. Invaded leaf sheaths turn brown or grey and either stick to underlying layers or rot and become easy to remove. Layers, patches and strands (rhizomorphs) of white fungal mycelium are often visible on and between the dead leaf sheaths. The mycelium is conspicuous on water-soaked, brown lesions, which form on underlying leaf sheath layers (Plates

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3.9 and 3.10). The rot spreads to the inner leaf sheaths, diminishing in size as it advances (Plate 3.11). The disease penetrates to the centre of the pseudostem in severe cases. The dry, diseased outer leaf sheaths at the base of the pseudostem become moist and can be torn off easily. There is usually a characteristic odour of mushrooms. Fungal fruiting bodies (basidiocarps) develop on the pseudostem in wet weather (Plate 3.10) (Ramos, 1941; Wardlaw, 1961; Stover, 1972). Work in Jaffna in Sri Lanka showed that banana plants (which were 2–3 months old when infected) wilted and unfurling leaves became smaller. A necrotic margin was seen in unfurled leaves and the plant eventually died. Pseudostems failed to develop further in plants infected at a medium stage of growth, resulting in either no bunch formation or bunches of reduced size. When plants were infected at an advanced

Plate 3.9.  Symptoms of Marasmiellus pseudostem rot on a cultivar in the Cavendish subgroup (AAA) in Queensland, Australia. The dead outer leaf sheaths have been stripped away to reveal white mycelium on brown lesions (photo: QDPI).

stage of growth or when bunches were emerging, the development of bunches could be affected, with plants toppling even in light winds (­Thiruchchelvan et al., 2013a). On Manus Island in Papua New Guinea, few plants had discoloured corms and true stems with most symptoms in the leaf sheaths. In some cases, fruiting bodies were evident on the outside of the pseudostem and white mycelia were present on brown lesions when the dried outer leaf sheaths were stripped away (J. Daniells, Australia, 2017, personal communication).

Causal agent The disease is caused by Marasmiellus inoderma, a member of the order Agaricales. Its mushroom-

Plate 3.10.  Fruiting bodies of Marasmiellus inoderma visible in the picture on the top right of the pseudostem of an infected abacá plant. The leaf sheaths have been stripped away from the side of the pseudostem facing the camera to reveal white patches of mycelium (photo: E.O. Lomerio, FIDA).



Fungal Diseases of the Root, Corm and Pseudostem

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and kill the roots. Young roots first become covered with white mycelium and develop a blackened tip. Later roots shrivel and the cortex turns brown and soft. Eventually the vascular cylinder darkens and decays. The tissues of the corm are not invaded. Plants that have had their root system partially destroyed can easily be pushed over. In abacá plantations, the fungus has been reported to be present in old pseudostem stumps (Wardlaw, 1961). The fruiting bodies of M. inoderma develop abundantly at soil level, but are also produced on the surface of the pseudostem (Plate 3.10). The basidiospores are formed in the gills of the fruiting body and disseminated by the wind (Wardlaw, 1961). The high annual rainfall on Manus Island coupled with minimal management of the plants (rampant weed growth plus no desuckering and detrashing) were believed to have created favourable conditions for disease development (J. Daniells, Australia, 2017, personal communication). In Jaffna in Sri Lanka, the disease was more evident in the wet season (Thiruchchelvan et al., 2013b).

Plate 3.11.  Interior of the pseudostem of a banana affected by Marasmiellus pseudostem rot showing internal brown discolorations (photo: Cooke et al., 2009).

like fructifications are at first brownish or salmon-­ yellow in colour on the upper surface, later turning pale. The cap, which measures 5–15 mm in diameter, frequently turns upwards, exposing widely separated gills. A white glabrous stalk, which is about 7–9 mm in length and flattened oval in cross-­section, arises from a sclerotic body and is attached to the cap off-centre. The spores, which are hyaline, oval and papillate at the point of attachment to the basidium, measure 7–8.5 × 5–6 μm. The stalk and cap shrivel readily, but recover quickly when placed in water (Wardlaw, 1961).

Disease cycle and epidemiology The plant is usually infected at soil level. As well as the pseudostem, the fungus may also invade

Host reaction The fungus is capable of attacking a wide range of banana and abacá cultivars. Some affected cultivars in Jaffna, Sri Lanka, which includes those in the Kathali subgroup (AA) and the Monthan subgroup (ABB), have been named by Thiruchchelvan et al. (2013a).

Control Cultural practices need improving if the disease is present. Affected plants, particularly stumps, should be removed and trash eliminated from the surroundings. Weeds also need to be kept under control. Plants should not be overcrowded, as high humidity favours infection. Mancozeb and chlorothalanil were the most effective fungicides in inhibiting growth of Marasmiellus cultures (Thiruchchelvan et al., 2013b).

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Pseudostem Heart Rot D.R. Jones and E.O. Lomerio Introduction Pseudostem heart rot of banana is usually a minor disease that is associated with plant injury. However, it was considered to be serious on young plantations of ‘Gros Michel’ (AAA) following severe storms and floods when this cultivar was grown extensively in Central America and the West Indies. After ‘Gros Michel’ was replaced in the export trades by cultivars in the Cavendish subgroup (AAA), the disease was rarely seen (Stover, 1972). Nevertheless, heart rot was reported as a problem on Cavendish cultivars causing economical losses in Egypt in 2006 (Shalaby et al., 2006). Heart rot also affects abacá, occurring mainly in weakened plants growing in dense stands in damp locations under conditions of high humidity. The disease has been observed to spread from centres of infection (Wardlaw, 1961). Other names for pseudostem heart rot are heart-leaf disease, Fusarium stalk rot and stalk heart rot.

the pseudostem become twisted and abortive. This obstructs the upward movement of following leaves, which are damaged and decay. Sometimes, median rather than central sheaths are decayed, resulting in a ring of rot in the pseudostem. In these cases, the plant can recover and the decay is sloughed off during growth. Rots may develop in scattered pockets of infection throughout the pseudostem. However, these usually remain free of secondary rots. The corm is not invaded (Wardlaw, 1961; Stover, 1972). If plants are young and actively growing, central sheaths can recover from heart rot. In these cases, the growing point has either not been damaged or growth through the decayed area has been rapid. However, the lamina of emerged leaves may be ragged, with missing ­portions (Waite, 1956). Plants damaged by windstorms, floods and pruning operations ­frequently recover (Stover, 1972). Symptoms of the disease in abacá are similar. Plants can have their central cylinder completely rotted. Leaves become yellowed, with dry margins and stiff petioles (Plate 3.12).

Symptoms The external symptoms of pseudostem heart rot of banana are the dying and rotting of the inner leaves of the crown. These leaves can become nearly fully unrolled or remain furled as they emerge. Diseased leaves are bright yellow or brown and may collapse at the petiole base. In severe cases, all leaves collapse and the pseudostem is killed. Usually, no more than one pseudostem in a mat may be affected, but several mats may be affected in one area (Wardlaw, 1961). The initial internal symptom is a blackening and decay of the furled heart leaf within the pseudostem. The rot may be firm and odourless at this stage. The decay begins in the upper part of the pseudostem and may extend down to the base, while surrounding leaf sheaths remain unaffected. The central tissue breaks down to a foul-smelling liquid, as a result of a secondary bacterial infection, and healthy sheaths in contact with the diseased core often become streaked brown or purple. Diseased leaves that develop in

Plate 3.12.  Yellowing and decaying leaves of abacá affected by pseudostem heart rot in the Philippines (photo: E.O. Lomerio, FIDA).



Fungal Diseases of the Root, Corm and Pseudostem

Causal agent Pseudostem heart rot was reported to be initiated by Fusarium moniliforme (Stover, 1972). However, because in the past this name has been used for at least six, and probably more, reproductively isolated mating populations (i.e. biological species), and possibly more phylogenetic species, Seifert et  al. (2003) have advised that this name should no longer be used. The pathogen will need to be re-examined using modern taxonomic techniques before any firm conclusions can be reached as to the correct name for the causal agent of pseudostem heart rot. Until then, it is recommended by Seifert et al. (2003) that pathogens be named F. verticillioides (syn. F. moniliforme). Bacteria follow the initial infection and cause secondary infections. There is no evidence that bacteria alone can cause the disease. Other Fusarium spp. and Cephalosporium spp. have later been found in the decay.

Disease cycle and epidemiology The pathogen sporulates profusely on diseased sheaths that have not been affected by secondary rots. Microconidia are mainly produced in false heads by most isolates, but sometimes chains are formed. The fungus has no chlamydospores. Banana plants could not be infected by pouring spore suspensions into crowns between leaf sheaths or by the application of spores or soil. Infections have only occurred when the fungus has been applied to wounded furled leaves inside the pseudostem (Waite, 1956). This indicates that banana tissues have to be injured for infection to take place (Stover, 1972). Ocfemia and Mendiola (1932) and Ramos (1933) reported that water-soaked, light to dark brown lesions could be induced in abacá when Fusarium moniliforme var. subglutinans was applied to unrolled heart leaves kept under moist conditions. Again, the identity of this pathogen is not precisely known given that the organism has been reclassified into at least 11 Fusarium species, including F. musae (Y.P. Tan, Australia, 2017, personal communication). The effects of the pathogen were most severe when the tissues were wounded. Abacá affected by bunchy top, nematode attacks on roots and

239

weevil borer infestations in the corm are more prone to pseudostem heart rot than healthy plants. Ramos (1933) showed that abacá could be predisposed to infection by pruning the root system. In plants affected by bunchy top, the decay is said to begin on the thin, chlorotic margins of the youngest leaves. This extends to the midrib and then to the petiole. During wet weather, the decay then progresses down to the heart.

Host reaction As mentioned above, ‘Gros Michel’ seems more prone to pseudostem heart rot than Cavendish cultivars. However, Cavendish cultivars were reported as susceptible in Egypt with ‘Williams’ being the worst affected (Shalaby et  al., 2006). The disease has also been recorded on ‘Latundan’ (AAB, Silk subgroup) in the Philippines (Lee, 1921, and Serrano, 1925, reported by Wardlaw, 1961). Pseudostem heart rot is rare on cultivars in the AAB and ABB genomic groups in Central America (Stover, 1972).

Control The disease is rare in banana plantations that are well maintained. Badly diseased plants should be eliminated (Stover, 1972). In abacá plantations, other disease problems need to be kept under control and plants should be well aerated.

Ceratocystis Corm Rot D.R. Jones Ceratocystis corm rot, which has been reported on banana in Australia, is caused by Thielaviopsis paradoxa (syn. Ceratocystis paradoxa). Extensive brown or dark water-soaked areas develop in corms after planting and cause a breakdown of the tissue. A dirty-white mould is generally visible in cavities in the affected areas. The pathogen is a soil inhabitant, especially in old pineapple land, and enters the corm through wounds. It is regarded as a minor problem and is controlled by planting healthy corms (Persley, 1993). The

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morphology of spores and spore-bearing bodies is shown in Fig. 3.3.

Sclerotium Corm and Pseudostem Rot D.R. Jones In 1987 a new banana disease, called pseudostem rot, was observed on 3–5-month-old ‘Robusta’ (AAA, Cavendish subgroup) plants. A fungus was observed inside the affected leaf sheaths. The pathogen was isolated in pure culture and identified as Athelia rolfsii (syn. Sclero­ tium rolfsii). The pathogen produced leaf spot diseases (by basidiospores) on 16 tested host

plants from various families (Mohan and Lakshmanan, 1989). During a disease survey in different banana-­ growing regions of India in March 2005, a wiltlike disease was observed in ‘Rasthali’ (AAB, Silk subgroup) in the Tirukattupalli area of Thanjavur district in Tamil Nadu, India. The important symptoms observed were yellowing of leaves from base to apex and extensive pseudostem sheath rot with profuse mycelial growth on the pseudostem. The colour of the rotted portion was yellowish-red to reddish-brown with numerous brown sclerotia present. Splitting of the basal portion of the pseudostem sheath was observed and bottom leaves had desiccated. The infected plant emanated a mushroom odour. The inner tissues of the corm were spongy and colonized extensively with white mycelium. At higher altitudes of 2000–3000 m,

B

E

C

G F

A

D 10μ 100μ Fig. 3.3.  Spores and spore-bearing structures of Thielaviopsis paradoxa (syn. Ceratocystis paradoxa). Key: A = perithecium; B = ascospores; C = chlamydospores; D = perithecial appendages: E = surface of perithecial wall; F = condiophores; G = conidia (Morgan-Jones, 1967).



Fungal Diseases of the Root, Corm and Pseudostem

the cortical portion of the corm was completely converted into a mass of fibre, which was similar in texture to coconut fibre, with profuse growth of fungal mycelium. In some plants, the infection extended up to half of the medulla region. Emerging peeper suckers were necrotic and colonized extensively with white mycelium. In some cases the mycelial growth was seen on the growing tip of the young bud and resulted in reddish-brown necrosis. The symptoms were only observed in plants more than 7 months old (Thangavelu and Mustaffa, 2010b). A fungus was isolated from diseased tissues and cultured on potato dextrose agar. Sclerotia were observed in cultures. The placement of single sclerotium in the pseudostem sheath of tissue-­ cultured plants wounded with pinpricks caused rotting of tissues in 3 days. The same fungus was re-isolated and identified as Athelia rolfsii (syn. Sclerotium rolfsii) (Thangavelu and Mustaffa, 2010b). During a subsequent survey conducted in different parts of banana-growing states of India in 2005, the disease was observed in ‘Rasthali’ and other commercial cultivars, such as ‘Virupakshi’ (AAB, Pome subgroup) and ‘Karpuravalli’ (ABB, Pisang Awak subgroup) in Tamil Nadu, ‘Ney Poovan’ (AB) in Karnataka and ‘Boothibale’ (ABB, Monthan subgroup) in Kerala, In most of the areas surveyed, the disease was observed in only a few plants, but in West Godavari district, Andhra Pradesh and Pudupatti village in Madurai district, Tamil Nadu, the disease incidence in ‘Rhastali’ was more than 50%. The disease is widespread in India (Thangavelu and Mustaffa, 2010b). In 1992, a disease that was observed for the first time at Areka Experimental Station in Welayita, Ethiopia, killed young enset seedlings and new transplants. Roots and leaf sheaths at soil level became rotten and the growth of plants was stunted (Plate 3.13). Severely attacked plants died and enset plots were devastated (Plate 3.14). The causal agent was identified as being in the genus Sclerotium, which gained entry to the enset seedlings and transplants through damaged roots and corms. The pathogen was found to survive in disintegrating root and corm tissue present in the soil. Sclerotial bodies and mycelia isolated from diseased specimens were demonstrated to be pathogenic and initiated root rots and wilt when inoculated into

241

Plate 3.13.  Symptoms of sclerotium root and corm rot of enset in Ethiopia. Leaves are dying on the stunted plant in the foreground (photo: M. Tessera and A.J. Quimio, IAR).

the crown of young, healthy enset seedlings of different cultivars. The disease was controlled effectively by dipping corms in a fungicide suspension (Quimio and Tessera, 1996).

Cephalosporium Infloresence Spot of Enset M. Tessera and A.J. Quimio This minor disease, which causes extensive necrosis of flower bracts and necrotic spots on leaf sheaths of mature plants, was recorded for the first time at Areka Experimental Station, Welayita, Ethiopia, in 1991 (Tessera and Quimio, 1994). The causal agent was believed to be a species of Cepha­ losporium. This fungus was isolated from diseased tissue and then injected into the stems of enset seedlings, where it was found to initiate a necrotic reaction (Quimio and Tessera, 1996).

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Plate 3.14.  Plot of enset at Areka Experimental Station in Ethiopia devastated by sclerotium root and corm rot (photo: M. Tessera and A.J. Quimio, IAR).

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4 

Fungal Diseases of Banana Fruit

Preharvest Diseases D.R. Jones and R.H. Stover

problems with preharvest fruit are covered in the chapters on virus diseases, non-infectious disor­ ders and genetic abnormalities.

Overview

Anthracnose Fruit Rot

Preharvest diseases of banana fruit caused by fungi are important because they produce un­ sightly blemishes, which are unacceptable to consumers, on the fruit peel. Although eating quality is seldom affected, because the pulp is only occasionally invaded, the fruit is rejected ­because its value is significantly lowered. From time to time in the past, severe outbreaks of ­various preharvest diseases have occurred, which have resulted in substantial losses. These diseases were controlled by implementing good sanitation practices in plantations, covering fruit with a poly­ ethylene bag (see Plate 1.6) and spraying fruit with fungicide when necessary. Although many of the problems reported here, such as brown spot and pitting disease, have declined in signifi­ cance, others, such as freckle and speckle, have increased. In the past, fungicides were applied to fruit on commercial plantations to control diseas­ es when necessary. Today, because of health con­ cerns, ­options for chemical control are limited. Bacterial diseases that are insect transmit­ ted can also affect preharvest fruit. In contrast to fungal problems, these invariably affect the pulp and make the fruit inedible. Bacterial diseases of fruit are described in the next chapter. Other

Anthracnose is a common and widespread dis­ ease of postharvest fruit, but in the Asia–Pacific region it can also affect immature fruit on the plant. It has been reported that up to 16% of un­ harvested bunches have been diseased in the ­Philippines (Wardlaw, 1961). Infections begin as small, black circular specks on the flowers and peel at the distal ends of the hands. Salmon-­pink-­ coloured spore masses of the fungal pathogen ap­ pear as the lesions increase in size and coalesce to form large sunken, blackened areas. In severe cas­ es, fingers can be covered in lesions. Affected fruit ripens prematurely, rots and eventually shrivels (Plate 4.1). The disease may spread to crowns and attack other fingers via pedicels (Plate 4.1). The fruit of abacá has also been reported to be affected (Ocfemia, 1924; Agati, 1925). Anthracnose fruit rot is caused by Colletotrichum musae. This fungus is described in the sec­ tion on postharvest fruit. High humidity, high temperatures and the presence of bruises pro­ mote infection, which probably occurs mainly at the remains of the perianth (Wardlaw, 1961). The conidia of C. musae are abundant in banana plantations because the fungus is a common ­inhabitant of decaying banana leaves and old

© D.R. Jones 2019. Handbook of Diseases of Banana, Abacá and Enset (D.R. Jones)

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Brown Spot Introduction Brown spot was a common blemish in some are­ as of the Americas on fruit developing during warm, rainy weather. Incidence varied greatly, but the most severe outbreaks occurred in Mexi­ co, Guatemala and Honduras. In the latter two countries, entire bunches were rejected and up to 20% of hands lost due to the disease. It also caused losses in Costa Rica, Panama, Colombia, Ecuador, Suriname and the Caribbean Islands. Although it was first described only in 1965, it may have been present earlier, because the path­ ogen is a common saprophyte on hanging ba­ nana-leaf trash and on leaves of dead and dying weeds (Stover, 1972). Brown spot is not an im­ portant disease today.

Symptoms

Plate 4.1.  Symptoms of anthracnose fruit rot on fingers of ‘Alukehel’ or ‘Ash Plantain’ (ABB, syn. ‘Ney Mannan’) in Sri Lanka. Note the disease spreading from the crown into a finger via the pedicel (photo: D.R. Jones, INIBAP).

fruit. They are spread by wind and may also be carried by insects that frequent banana flowers. Germination occurs readily in water, but conidia quickly die in the absence of moisture. The dis­ ease is favoured, therefore, by conditions of high humidity and rainfall. In inoculation experi­ ments, the disease was initiated when the fungus was applied to slight wounds in the peel. Very immature fruit, with flowers still attached, were more readily attacked than older fruit (Agati, 1922). Field inoculations carried out in the West Indies were unsuccessful (Wardlaw, 1931). Thirty different cultivars showed susceptibili­ ty in the Philippines. Cavendish cultivars were the most severely attacked, with cooking-banana types being less susceptible (Wardlaw, 1961). However, immature bunches of ‘Blue Java’ (ABB, syn. ‘Ney Mannan’) have been completely destroyed in Fiji (Parham, 1938). In Sri Lanka, ‘Alukehel’ or ‘Ash Plantain’ (ABB, syn. ‘Ney Mannan’) is the main cultivar affected (Plate 4.1) (Park, 1933).

Symptoms of brown spot occur on fingers, crowns and the peduncle. Spots are pale to dark brown with an irregular margin, surrounded by a halo of water-soaked tissue (Plate 4.2). The size of the spots, which are centred over stoma­ ta, varies, but they average 5–6 mm in diameter. Brown spot can be distinguished from Deight­ oniella speckle because brown spot lesions are much larger than those caused by Deightoniella torulosa. They differ from those of pitting disease because they are not sunken, as are lesions caused by Pyricularia angulata. Also, they do not increase in size or number during ripening, as do those of pitting disease. Brown spot occurs on fruit that is 50 days old or older. The pathogen does not sporulate in the fruit spots.

Causal agent Cercospora hayi has been reported as the cause of brown spot. In their treatment of the genus Cercospora, Crous and Braun (2003) considered C. hayi to be a synonym of the older name, Cercospora apii, which is the cause of celery leaf blight and is known to have a wide host range. Based on a comparison of DNA sequence data with the ex-type strain of C. apii (Groenewald



Fungal Diseases of Banana Fruit

257

tissue. Mycelium of the fungus survives in dried leaf tissue for 15 weeks and spores remain viable for at least 5 weeks when exposed to fluctuating temperatures and humidity levels (Stover, 1972). Conidia form within 16 h at 23–26°C in a satu­ rated atmosphere and are released in air cur­ rents with velocities as low as 2.4 km/h. Spore release peaks in the afternoon between 14.00 and 16.00 h at a time that usually coincides with the highest daily wind speeds. Conidia ger­ minate within 3 h on green banana peel. Infec­ tion mainly occurs on fruit less than 21 days old, but spots do not appear until the fruit is ap­ proaching maturity, at about 90 days (Kaiser and Lukezic, 1966). In Honduras, conidia were present through­ out the year, but numbers were highest in July, September and October, when most rain fell. Brown spot incidence was highest during the rainy season, from July to January (Stover, 1972).

Host reaction

Plate 4.2.  Symptoms of brown spot in Honduras. Note the irregular outline of the spots and water-soaked haloes (photo: J. Rivera, FHIA).

et al., 2006), this synonymy appears to be correct. However, in a later publication, the morphologi­ cal similarity of C. hayi and C. musigena was not­ ed, the latter being found in necrotic leaf margins of banana in Thailand. Because sequences from cultures of C. apii-like fungi isolated from banana clustered in three different clades (Groenewald et al., 2013), it was concluded that the name C.  hayi was still unresolved (Nguanhom et al., 2015). The causal fungus is readily isolated from brown spot lesions, but does not produce spores on most media. However, it does sporulate on pro­ pylene oxide-sterilized banana leaf tissue after 7 days. Conidia are hyaline, measure 90–150 × 3–4 μm, contain five to 15 cells and have truncate bases and acute tips (Kaiser and Lukezic, 1965).

Fruit of accessions of Musa acuminata ssp. malaccensis (AAw) and ssp. microcarpa (AAw) have been seen with brown spot, but not fruit of Musa balbisiana (BBw). Fruit of all banana cultivars was susceptible, but the clones in the Cavendish subgroup (AAA) seemed to be more susceptible than ‘Gros Michel’. Dessert-banana cultivars in the AB and AAB genomic groups were also af­ fected, but cultivars in the Plantain subgroup (AAB) were not seriously spotted (Kaiser and Lukezic, 1966).

Control When brown spot was serious, it was controlled in the same way as pitting disease.

Cigar-end Rot Introduction

Disease cycle and epidemiology The pathogen attacks a number of plant species and is also found in dead banana and weed leaf

Cigar-end rot is a disease of banana fruit found throughout the tropics (Hawkesworth and Holl­ iday, 1970). It is of importance in Africa (Baker et al., 2008). A loss of 10,000 bunches was

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r­ eported in Cameroon in 1958 (Stover, 1972). Very high incidences have also been reported in Kenya (Mwangi, 2011). Outside the tropics, the disease has been reported from the Middle East, the Canary Islands and Morocco (Wardlaw, 1961; Stover, 1972; Slabaugh, 1994b; Boubak­ er et al., 2008; Simon et al., 2012). Cigar-end rot is caused by two different fungi.

Symptoms A black necrosis spreads from the perianth into the tip of immature fingers. The pulp usually un­ dergoes a dry rot. The corrugated necrotic tissue becomes covered with the mycelia and spores of the causal fungi and resembles the greyish ash of a cigar-end (Plate 4.3). Diseased tissue is de­ limited from healthy tissue by a black band and a line of chlorosis (Wardlaw, 1931). The rot spreads slowly and seldom affects more than the first 2 cm of the finger tip (Plate 4.4). However, if fruit is attacked soon after emergence, the entire finger sometimes becomes rotten. When Musicillium theobromae is the causal agent, the affected tissue is characteristically dry and fibrous and the spores are grey and powdery. In Australia, this type of cigar-end rot is often associated with ‘choke throat’ (Simmonds, 1966). When the disease is caused by Trachysphaera fructigena, the surface of the lesion becomes cov­ ered with white spores, which later turn pink or brown as they mature, giving the fingertip a greyish, ashen appearance. Internally, the pulp may undergo a dry rot and become mummified (Brun, 1970). A wet rot can occur when second­ ary organisms are present (Dadzie and Orchard, 1997).

Plate 4.4.  Symptoms of a cigar-end rot on fruit of ‘Yangambi Km 5’ (AAA, syn. ‘Ibota Bota’) in Nigeria. Trachysphaera frucigena was one of several fungi isolated from affected fruit. (photo: C. Pasberg-­ Gauhl and F. Gauhl, IITA).

Causal agents Cigar-end rot is caused by either Musicillium theobromae (syn. Verticillium theobromae) (Zare et al., 2007) or Trachysphaera fructigena. However, M. theobromae is more widespread than T. fructigena, which is only found in tropical West and Central Africa plus Madagascar (Holliday, 1970). In these locations, both fungi can cause the dis­ ease, but T. fructigena is the main agent. Musicillium theobromea

Plate 4.3.  Symptoms of cigar-end rot in Australia caused by Musicillium theobromae (photo: Cooke et al., 2009).

Conidiophores of M. theobromae, which measure 150–400 × 4–6 μm, are solitary or in small groups on the surface of the peel. Conidia are borne at the ends of tapering phialides and are aggregated into rounded, mucilaginous, translu­ cent heads. Conidia are 4–9 × 2–3.5 μm and phi­ alides 15–30 μm (Meredith, 1961b) (Fig. 4.1).



Fungal Diseases of Banana Fruit

259

(B) 10 μ

(A)

Fig. 4.1.  Conidiophore (A) and conidia (B) of Musicillium theobromae (Hawksworth and Holliday, 1970).

Trachysphaera fructigena Conidiophores of T. fructigena are simple or bear a terminal vesicle, to which a whorl of pedicellate conidia is attached (Fig. 4.2). The conidia are spherical, strongly echinulate, with an average diameter of 35 μm. They are borne on pedicels, which vary in length up to 30 μm. Oogonia char­ acteristic of the Peronosporales are produced in diseased tissue. They are small, averaging 40 × 24 μm, thick-walled and characterized by the pres­ ence of irregular, sac-like outgrowths (Fig.  4.2). The antheridia are amphigynous, completely sur­ rounding the stalk of the oogonium. Pure cultures can easily be established from conidia. Trachysphaera fructigena was first described on cacao and coffee in West Africa (­Tabor and Bunting, 1923).

Disease cycle and epidemiology In Cameroon, cigar-end rot was found to be at its worst during the rainy season, virtually

­ isappearing in the dry season (Brun, 1954). d Later, a direct correlation was shown between rainfall and incidence of the disease on Caven­ dish cultivars (Beugnon et al., 1970). The prob­ lem was more serious at higher elevations and around the borders of plantations next to native vegetation (Brun, 1954). In Egypt, cigar-end rot was rare in the winter months, reaching a peak in May and June (El-Helaly et al., 1955). In gen­ eral, poorly maintained plantations had the highest levels of disease. In Kenya, infection commonly started in the youngest fruit shaded by the still-attached bracts of the male bud (Mwangi, 2011). Musicillium theobromae is a common colo­ nizer of banana-leaf trash and flowers. Conidia are disseminated in air currents and infect dying flower parts. The optimum growth of M. theobromae in culture is 25°C. Spores of T. fructigena are not believed to re­ main viable for long under field conditions (Mar­ amba and Clerk, 1974). Tezenas du Montcel and Laville (1977) reported that the fungus grew

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10

50 μm

Fig. 4.2.  Conidia, oogonia and antheridia of Trachysphaera fructigena (from Tabor and Bunting, 1923).

best in culture at 24°C, but that cigar-end rot was accentuated when moderate temperatures of around 20°C were followed by higher temper­ atures of above 27°C. The growth and sporula­ tion of three isolates of T. fructigena obtained from various locations in Cameroon in vitro were investigated by Pefoura et al. (2007). The mini­ mum, optimum and maximum temperatures, respectively, were reported to be around 15°C, 21°C and 30°C for growth, and 18°C, 22°C and 27°C for sporulation. In inoculation experi­ ments, advanced symptoms appeared on imma­ ture, wounded fruit after incubation at 18–24°C for 5–7 days (Meredith, 1960). Unlike cigar-end rot caused by M. theobromae, the disease caused by T. fructigena can lead to premature ripening of fruit. This pathogen

also continues to attack and rot fruit after har­ vest. It can invade freshly cut crown surfaces and through wounds in the peel caused by im­ proper handling. Inoculum for postharvest in­ fections originates from fruit harvested with the disease and may be present in de-handing and de-latexing tanks. In the UK, bananas from Ja­ maica that were stored in ripening rooms previ­ ously occupied by diseased fruit from Cameroon became infected (Meredith, 1960). Host reaction Cigar-end rot affects fruit of ‘Gros Michel’ (AAA) and cultivars in the Cavendish subgroup (AAA). Plantains are also susceptible to the disease



Fungal Diseases of Banana Fruit

(­Pefoura et al., 2004). Dadzie and Orchard (1997) believed that it was essentially a disease of plan­ tain, but it is also found in banana and cooking banana.

Control Field sanitation, such as the removal of dead, hanging leaves from plants, will reduce inocu­ lum levels of M. theobromae. The early removal of dead flowers may eliminate the source of in­ fection before it reaches the finger. The use of polyethylene bags on bunches is also advocated as a control measure (Beugnon et al., 1970). Any trash accumulating in the bags should be removed. Perfoura et al. (2004) found that bag­ ging plantain bunches with oil palm leaves sig­ nificantly reduced incidence of T. fructigena. Packing stations and ripening rooms need to be kept clean to minimize the chances of post­ harvest infection by T. fructigena. Fingers affect­ ed by cigar-end rot should be cut from hands in the field to avoid contamination of water in de-­ handing and de-latexing tanks. It may be necessary to apply fungicide when cigar-end rot is prevalent. Brun (1970) found benzimidazole to be ineffective in controlling cigar-­end rot, but this has been used for control in Morocco where resistance to M. theobromae was detected (Boubaker et al., 2008). Tezenas du Montcel (1981) tested different fungicides and found that metalaxyl gave the best control.

261

An internal dry rot of the pulp sometimes occurs later, probably as a result of secondary invasion. The lesion, which frequently has an irregular out­ line, forms more on one side of the finger than on the other (Plate 4.5). Under conditions of high humidity, the surface of the affected tip is often covered with fungal mycelia. The disease does not have the ashy grey appearance associated with cigar-end rot and rarely affects more than the apical third of the finger. It is often associated with an injury, such as sunburn. Fingers in all stages of development can be affected. Corynespora torulosa (Crous et al., 2013) (syn. Deightoniella torulosa), which is a fungus responsible for a leaf spot disease of banana (see Chapter 2) and a pin-spotting disease of banana fruit (see next section in this chapter), is usually isolated from banana fruit with black-tip disease (Meredith, 1961a). However, Musicillium theobromae and Fusarium verticillioides (syn. Fusarium moniliforme) (Seifert et al., 2003) have also have been isolated from black-tip lesions (Meredith, 1965). Both are pathogenic on banana. Musicillium theobromae is the cause of cigar-end rot and F. verticllioides causes a minor, dark brown rot of

Tip-end Rot A number of tip-end rots have been reported on unripe banana fruit. They are not economically significant, but have caused some problems on occasions. Black-tip A black-tip disease was first reported from Bermu­ da by Ogilvie (1927, 1928) and was subsequently found in the Philippines (Reyes, 1934), Trinidad, Guinea, Somalia, Papua New Guinea (Wardlaw, 1961) and Jamaica (Stover, 1972). The disease is confined to the peel in the early stages. The skin turns black and has a dry c­ rumpling consistency.

Plate 4.5.  Symptoms of black-tip on a finger of ‘CEMSA ¾’ (AAB) in Cienfuegos province Cuba. Note that the end of the finger below the infection is ripening (photo: L. Pérez-Vincente, INISAV).

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the pulp of unripe fruit in Israel, known as black heart disease, which can also induce premature ripening (Chorin and Rotem, 1961). Both are also implicated in crown rot disease of ripening fruit. All the fungi implicated are common sapro­ phytes of dead flower parts of banana. Although attempts to reproduce symptoms of black-tip in the field have failed, symptoms were induced in fingers of ‘Lacatan’ (AAA, Cavendish subgroup) by placing C. torulosa in incisions in the corky tissue at the tip of the fingers and incubating the fruit in a saturated atmosphere for 7 days at 28– 30°C. Spraying suspensions of conidia of C. torulosa on to young fruit and then sealing the bunch in a polyethylene bag also occasionally induced black tip (Meredith, 1961d). Perez et al. (1990) found that the optimum temperatures for mycelial growth and spore ­production of C. torulosa were 27°C and 30°C, respectively. The formation of spores was inhib­ ited below 25°C and above 30°C. Spore produc­ tion was optimum on media at pH 6–7. In Cuba, the disease is very common on ‘CEMSA¾’ and other cultivars that retain their flowers. This is because the fungus colonizes decaying flowers during the rainy season and from there invades the tips of the fruit. Fungicides do not totally solve the problem. Very good control was obtained in Cuba when bunches were de­ flowered (L. Pérez-Vincente, Cuba, 2017, personal ­communication). Dothiorella tip-end rot Dothiorella tip-end rot has been reported from Israel (Reichert and Hellinger, 1938; Chorin and Rotem, 1961), Guinea, Côte d’Ivoire (Roger and Mallamaire, 1937, in Reichert and Hellinger, 1938) and Australia (Wardlaw, 1961). The dis­ ease progresses more rapidly in the peel than in the pulp. Blackening is preceded by a narrow, brown, watery margin, which forms a sharp line of demarcation between healthy and diseased tissues. In Australia, the whole fruit may be af­ fected, with the distal end becoming tapered as the diseased tissues shrink, dry and become mummified (Wardlaw, 1961). Dothiorella tip-end rot is now attributed to Neofusicoccum ribis (formerly Dothiorella gregaria) (Crous et al., 2006). Pycnidia are usually produced

in the superficial layers of the peel and burst through the epidermis during development. They may occur singly or together in groups, but with little stromatic material. The black, thick-walled, globose pycnidia have a papillate ostiole and meas­ ure 141–321 × 141–242 μm. The fusoid or ovoid conidia are thin-walled and measure 13–21 × 4–7 μm. The conidia are forcibly ejected en masse and each pycnidium becomes surmounted by a little mass of white powder, which is a character­ istic of the disease. Insects and wind may spread the conidia. In Israel, Dothiorella tip-end rot increases in the second and third year after planting and then decreases (Rotem and Chorin, 1961). Fruit and plants in plots irrigated by sprinklers are more prone to this disease than on plants in surface-­ irrigated plots. Conditions that favour the main­ tenance of high humidity levels, such as are prevalent in Israel in the winter, favour the prob­ lem. Disease levels were reduced in Israel by spraying with zineb and cuprous oxide. Control was also achieved by removing flowers immedi­ ately after formation. However, the practice of breaking off flowers 20–30 days after formation gave no control (Chorin and Rotem, 1961). Neofuicoccum ribis also causes a black rot of orange fruit in Israel, and an isolate of the fungus from oranges caused decay on banana fruit in inocula­ tion experiments (Reichert and Hellinger, 1938). In Jamaica, a similar disease to Dothiorella tip-end rot is reported on ‘Dwarf Cavendish’ (AAA, Cavendish subgroup) at an elevation of around 800 m, especially in damp localities. The large abaxial tepal or sometimes the smaller abaxial tepal are believed to be infected first and then finger tips are invaded. The withered flower parts remain attached to diseased fruit and are a characteristic feature. Often the disease does not progress more than a few millimetres, but decay can sometimes spread 6–8 cm along the finger after 20–30 days, usually advancing more quick­ ly on one side than on the other. In the early stag­ es, the decayed pulp is reddish brown and firm. Later it turns black and is hollowed out by in­ sects. Occasionally, the whole finger is affected and is reduced to a dry, sunken, mummified state. Neoscytalidium dimidiatum (Crous et al., 2006) (formerly Hendersonula toruloidea – Punithalingam and Waterston, 1970) was provisionally adopt­ ed as the causal agent by Meredith (1963a). However, Meredith (1963b) also suggested that



Fungal Diseases of Banana Fruit

this fungus may be the same pathogen as was described by Reichert and Hellinger (1938) as Dothiorella gregaria (now Neofusicoccum ribis). Pyc­ nidia are formed singly or together in groups on the blackened peel and withered flowers. The pyc­ nidium is immersed, globose and black and meas­ ures 110–340 × 100–260 μm, with a papillate ostiole. Most conidia are non-septate, with hya­ line granular contents, and measure 14–24 × 4–8 μm. They are produced at the ends of short, thin-walled cells, which form the innermost layer of the pycnidium wall. They are tightly packed and exude from the ostiole in a mucilaginous sheath. Conidia germinate in 1–2 h at 27°C in a film of water, becoming 1–3-septate in 12 h. The tips of fingers have been artificially in­ oculated by inserting mycelium from cultures in wounds at the junction of the perianth and fin­ ger tip in detached fingers. Symptoms appeared after 6 days on fruit of both ‘Dwarf Cavendish’ ­ ecay and ‘Lacatan’ (AAA, Cavendish subgroup). D was slow at 10°C, but fingers could be c­ ompletely decayed within 12 days at 27°C (Meredith, 1963a). ‘Lacatan’ fruit is not affected in locations where fruit of ‘Dwarf Cavendish’ is diseased. This may be because ‘Dwarf Cavendish’ has a more persistent perianth than taller members of the Cavendish subgroup (Simmonds, 1959). A corky abscission layer, which may preclude further fungal invasion, forms early in ‘­Lacatan’ fruit at the junction of the finger tip and peri­ anth (Meredith, 1963a).

263

Botrytis tip-end rot, caused by Botrytis cinerea, has been found in Israel and was reported as fairly common during the wet winter months. It discolours the peel, which is covered by a fairly dense, brown mycelium, and rapidly rots the pulp (Reichert and Hellinger, 1932). Fusarium and Lasiodiplodia species have also been isolated from tip-end rots (Wardlaw, 1961; C. Pasberg-Gauhl and F. Gauhl, Nigeria, 1995, personal communication). Glomerella cingulata, which has been found in association with brown blotches on green fruit in Queens­ land, Australia (Simmonds, 1966), has also been isolated from tip-end rots in Nigeria. This fungus, together with T. fructigena, was isolated from the tip-end rots illustrated in Plate 4.6 (C. Pasberg-Gauhl and F. Gauhl, Nigeria, 1995, personal communication).

Other tip-end rots Sclerotinia fruit rot is an uncommon, low-­ temperature disease and has been reported from Israel (Reichert and Hellinger, 1930), Bermuda (Waterston, 1947) and Costa Rica (Laguna and Salazar, 1984). A rot, caused by Sclerotinia sclerotiorum, begins at the flower end and spreads down the finger. The normally white pulp tissue becomes reddish-black and then black and is ­finally broken down and hollowed out (Wardlaw, 1961). The entire finger becomes rotten and shrivelled, and abundant sclerotia are found both internally and on the surface of the peel. In Israel, the disease only occurred in the winter. The prob­ lem was controlled by the removal of affected fruit and the application of copper sprays.

Plate 4.6.  Severe symptoms of a tip-end rot on fruit of ‘Fougamou’ (ABB, Pisang Awak subgroup) in Nigeria. The small necrotic fruit on the bunch on the right were most probably infected at an early stage of development (photo: C. Pasberg-Gauhl and F. Gauhl, IITA).

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Deightoniella Fruit Speckle Introduction Speckle, pin-spotting or ‘swamp spot’, as it is known in Jamaica, is a common blemish of ­banana peel. Most fruit has a low incidence of speckle, but it is not regarded as a serious prob­ lem as symptoms usually become much less obvious after ripening, when they are masked by the yellow colour of the peel. However, speckle can cause export fruit to be downgrad­ ed when severe. High levels of the disease can  occur following prolonged higher-than-­ normal rainfall and when a high humidity builds up inside the polyethylene bunch covers because of impeded drainage or lack of suffi­ cient perforations.

Symptoms Symptoms consist of minute, reddish-brown to black spots with small green haloes (Plate 4.7). The spots are usually less than 2 mm in diame­ ter, but some can be up to 4 mm if the disease is severe. The distribution of speckle within a bunch may be uneven. Stover (1972) reported that it was more severe on the outer face of in­ ner-whorl fingers and towards the tips of fin­ gers in the outer whorls. Symptoms can increase as the fruit ages, but resistance to in­ fection also increases with age. Meredith

Plate 4.7.  Symptoms of Deightoniella speckle caused by Corynespora torulosa on a cluster of fingers of ‘Robusta’ (AAA, Cavendish subgroup) in St Vincent, Windward Islands (photo: D.R. Jones, SVBGA).

(1961a) found that 10–30-day-old fingers were more easily infected than 70–100-dayold fingers. Speckling does not increase and spots do not enlarge after harvest. Speckle can be distinguished from spots caused by flower thrips because the latter are raised and bumpy to the touch.

Causal agent Deightoniella speckle is caused by Corynespora torulosa (Crous et al., 2013) (syn. Deightoniella torulosa), which also causes a leaf spot of ba­ nana, abacá and enset, and a pseudostem rot of abacá. Corynespora torulosa can also incite blacktip disease on preharvest banana fruit. The fun­ gus is a weak pathogen of old or injured banana leaves and is commonly found on dead banana leaves. It has been described by Ellis (1957), Meredith (1961a, d), Subramanian (1968) and in Chapter 2 of this volume.

Disease cycle and epidemiology Inoculum is generated during periods of rain or dew on dead banana leaves (Meredith, 1961a). Conidia are violently discharged as the humidi­ ty falls, and they become airborne (Meredith, 1961d). They are not disseminated over long distances and lose their viability within 4 days at a humidity of less than 95% (Meredith, 1961a, c). Conidia germinate in moisture and form appressoria after about 6 h (Meredith, 1961a). Penetration can occur within 12–20 h, as indicated by a reddish-­brown discoloration in the invaded epidermal cell (Meredith, 1961a). Adjacent cells then become discoloured as the fungus spreads intercellularly. Usually, no more than 15 cells are affected in width and depth. Speckles become visible to the naked eye on green banana peel within 72 h (Meredith, 1961a). The fungus does not sporulate in lesions on fruit.

Host reaction Stover (1972) has reported that all cultivars of banana are affected by speckle.



Fungal Diseases of Banana Fruit

Control The disease is more prevalent in badly main­ tained and poorly drained plantations that have abundant leaf trash. High-density planting, in­ adequate weed control and standing water on the ground all help to maintain a moist atmosphere, which encourages speckle. In well-maintained and properly drained plantations, where fruit is covered with a perforated polyethylene bag, speckle is rarely a problem. However, the bag should have sufficient perforations to prevent the build-up of a high humidity level around the bunch. Also, if the bag is knotted at the bottom to stop the polyethylene rubbing on fruit in the wind, the knot should not impede drainage. As leaves decay more rapidly on the ground than hanging on the plant, inoculum levels in the air can be reduced by removing dead leaves (Meredith, 1961a, c). Spraying the young fruit with dithiocarbamate fungicides, as was prac­ tised for the control of pitting disease, also re­ duces the incidence of speckle.

Causal agents of fruit speckle in Central America Research into speckle resumed in the late 1990s because the problem had increased, particularly in the wet season in some areas in Central ­America (Plate 4.8). In Costa Rica, up to 70% of fruit from one farm had been rejected for export.

265

It was discovered that, although symptoms were very similar to Deightoniella speckle, Corynospora torulosa was rarely isolated from lesions. Young, unprotected fruit before bagging is vul­ nerable to damage caused by agrochemicals, such as fungicides and leaf fertilizers, which may cause speckle-like symptoms. Physiological reactions of the fruit leading to small necrotic flecks may also be induced by insects feeding or laying eggs and other fungi penetrating the peel. In total, 16 different sporulating fungi have been isolated from ‘speckle’ lesions during inves­ tigations. Ten of these fungi were selected for ­artificial inoculation experiments. These were a  Cephalosporium sp., Colletotrichum musae, a Cylindrocarpon sp., Corynespora torulosa, Fusarium dimerum, Fusarium chlamydosporum var. chlamydosporum (syn. Fusarium fusarioides), Fusarium incarnatum (syn. F. pallidoroseum and F. semitectum), Fusarium tricinctum, Clonostachys rosea (syn. Penicillium roseum) and a Trichoderma sp. Typical speckle symptoms could be reproduced with D. torulosa, the Cephalosporium sp., Colletotrichum musae, Fusarium chlamydosporum var. chlamydosporum, F. incarnatum and F. tricinctum. Very faint speckling, similar to some symptoms seen in the field, formed after inoculation with the Cylindrocarpon sp., Clonostachys rosea and Trichoderma sp. Speckle is less common on farms where banana plants are treated with fungicide than on farms where banana plants are untreat­ ed. (C. Pasberg-Gauhl, Costa Rica, 1999, personal communication). Different farm management practices for the control of fruit speckle, such as careful bag­ ging practices and early deflowering as well as how to keep humidity levels low, both within the tree bag and in the plantation, were discussed by Pasberg-Gauhl (2002). Causal agents of fruit speckle in North Queensland, Australia

Plate 4.8.  Symptoms of ‘speckle’ on fruit of ‘Grand Nain’ (AAA, Cavendish subgroup) in Costa Rica. Spots closely resemble those of Deightoniella speckle, but are caused by a complex of different fungi (photo: C. Pasberg-Gauhl and F. Gauhl, CB).

The symptoms of fruit speckle of banana in North Queensland were reported by Vawdrey et  al. (2010) as minute reddish-brown to black spots (0.5–1 mm in diameter) often with an oilsoaked or water-soaked margin. Eleven species of fungi were recovered from speckle lesions, but only Colletotrichum musae, Fusarium oxysporum and F. incarnatum (syn. F. pallidoroseum and

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F. semitectum) reproduced speckle-like symptoms on young fruit following inoculation. Studies on fruit speckle epidemiology showed that spraying young fruit with a 10% sap solution before inoc­ ulation with Fusarium spp. caused a threefold increase in the number of speckle lesions, but had much less of an effect on the incidence of speckle following inoculation with C. musae. The presence of flower thrips had little effect on the incidence of symptoms on fruit inoculated with C. musae, but caused a tenfold increase in the incidence of speckle on fruit inoculated with ­ Fusarium spp. Fruit was also shown to be less susceptible to fruit speckle as it matured (Vawdrey et al., 2010). In a field planting of ‘Lady Finger’ (AAB, Pome subgroup), bunches injected at bunch emergence with the fungicide azoxystrobin (0.15 g a.i./l) and sprayed fortnightly with azox­ ystrobin (0.25 g a.i./l) significantly reduced the number of speckle lesions/cm2 compared with bunches injected and sprayed fortnightly with insecticides. This strongly indicated that fruit speckle was caused by fungi and not insects. The fungicides propineb, azoxystrobin, trifloxystrob­ in, copper oxide, mancozeb and chlorothalonil effectively reduced the disease in in vitro experi­ ments, compared with the inoculated control (Vawdrey et al., 2010).

Symptoms Spotting begins to appear as fruit approaches ma­ turity. The first symptom on green peel is a slight­ ly raised, inconspicuous yellow spot 3–5 mm in diameter. The cells colonized by the pathogen do not expand and a longitudinal crack, which is surrounded by a yellow halo, develops as the sur­ rounding tissue grows. The crack increases in length beyond the halo and widens at the centre. The tissue exposed by the crack and the yellow halo becomes necrotic, collapses and turns black. The spot then appears as a black, sunken, dia­ mond-shaped lesion with dimensions of 1.0–3.5 × 0.5–1.5 cm (Plates 4.9 and 4.10). Small spots rarely extend below the peel, but large spots occa­ sionally expose the pulp. The size and number of spots can increase after harvest and inconspicu­ ous spots can become unsightly blemishes as fruit is transported and ripened (Berg, 1968). During the early and late stages of spot develop­ ment, the symptoms are distinctive. However, the middle stage can be confused with pitting disease.

Diamond Fruit Spot Introduction Diamond spot was common in Honduras and Guatemala following the expansion of the cul­ tivation of clones in the Cavendish subgroup (AAA) in the 1960s. It was also a problem in Mexico and frequently encountered on unsprayed bunches on the Pacific coasts of Nicaragua, ­Costa Rica and Panama and in the Philippines. It was not common on the Atlantic coasts of Costa Rica and Panama or in Ecuador, Colom­ bia, Suriname and the Caribbean (Stover, 1972). The disease caused serious losses of fruit in Honduras and Guatemala during the rainy sea­ son from July to January. In these countries, up to 10% of hands were discarded. However, dia­ mond spot is no longer important.

Plate 4.9.  Close-up of a diamond spot lesion on fruit in Honduras (photo: J. Rivera, FHIA).



Fungal Diseases of Banana Fruit

267

Disease cycle and epidemiology

Plate 4.10.  Symptoms of diamond spot on fruit of ‘Agbagba’ (AAB, Plantain subgroup) in Nigeria (photo: C. Pasberg-Gauhl and F. Gauhl, IITA).

Causal agents Diamond spot is caused by Fusarium solani, Fusarium incarnatum (syn. F. pallidoroseum) and possibly other fungi invading lesions initiated by Cercospora hayi (Stover, 1972). Originally, it was thought that F. incarnatum, possibly in con­ junction with other fungi common on the ba­ nana peel, was the cause of the disease (Berg, 1968). The strain of the pathogen that initiates di­ amond spot may be different from the one that causes brown spot. Cercospora hayi was consist­ ently isolated from diamond spot lesions to­ gether with Fusarium spp. When near-mature bananas were inoculated with C. hayi and incu­ bated in a saturated atmosphere, yellow spots up to several millimetres in diameter appear 2–3 weeks later. Certain isolates of F. solani and F. incarnatum, if inoculated at the same time as C. hayi, could invade and extend these initial le­ sions into typical diamond spots. The exact se­ quence of invasion and the stage of fruit maturity when this can take place in the field have not been determined. Fusarium spp. alone cannot penetrate un­ injured banana peel, though they can invade artificial wounds and cause various types of le­ sions. The failure of fruit, in many instances, to produce typical diamond spot symptoms after inoculation with spores of C. hayi and Fusarium spp. is an indication that highly specific condi­ tions may be necessary for disease development. Sporulation does not occur in diamond spot ­lesions.

Cercospora hayi and Fusarium spp. are common inhabitants of dead and decaying banana trash and produce abundant spores under moist con­ ditions. Fusarium incarnatum also sporulates on the dead pistils of banana flowers. Air currents carry the spores, which are common compo­ nents of the air flora of banana plantations, to fruit (Kaiser and Lukezic, 1966; Lukezic and ­Kaiser, 1966; Berg, 1968). Diamond spot was only prevalent following prolonged periods of rain.

Host reaction ‘Gros Michel’ (AAA) was less seriously affected than cultivars in the Cavendish subgroup (AAA) (Stover, 1972).

Control Measures used to control pitting disease also control diamond spot.

Freckle Freckle disease of banana is caused by different fungal species in the genus Phyllosticta. As well as leaves, symptoms of freckle are seen on ba­ nana and abacá fruit. As early as 2–4 weeks af­ ter bunch emergence, a few widely scattered spots or, in some cases, dense aggregates, in the form of streaks or circles, can appear on banana fingers. On green fruit, individual spots first ap­ pear as minute, reddish-brown flecks, surround­ ed by a halo of dark green water-soaked tissue. Large areas of the peduncle and fruit surface sometimes become black, due to dense aggrega­ tions of spots (Plates 4.11 and 4.12). Bracts can also be affected. The severity of the disease in­ creases as the fruit matures. Freckle is particu­ larly severe where fruit is in contact with diseased leaves. During ripening, the centre of individual spots darkens and each spot is sur­ rounded by a halo of green tissue up to 3 mm in diameter. Although this discoloration detracts from the appearance of the fruit, eating qualities are not affected (Meredith, 1968).

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in Japan, the blemishes caused by freckle are un­ acceptable. The disease is, therefore, a problem for export industries based in Taiwan and the Philippines. Information on the causal agents, leaf symp­ toms, epidemiology, host reaction and control of freckle is presented in Chapter 2.

Peduncle Rot

Plate 4.11.  The peduncle and fruit of ‘Mondolpin’ (AAB, Bluggoe subgroup) affected by freckle on Badu Island in the Torres Strait region of Queensland, Australia. Phyllosticta maculata is the likely pathogen (photo: D.R. Jones, QDPI).

Plate 4.12.  Freckle symptoms on a hand of Cavendish fruit in Mindanao, Philippines. The pathogen is likely to be Phyllosticta cavendishii (photo: Y. Israeli, JVBES).

Fruit spots are caused by concentrations of pycnidia. Consumers who buy fruit from local markets in South and Southeast Asia tolerate these symptoms. However, for discerning buyers

In commercial plantations and in many small­ holders’ plots, the male flower bud is broken off the flower stalk after the last female hand has emerged, because this may result in a slight in­ crease in bunch weight and eliminates a habi­ tat for thrips. This practice also reduces the incidence of Xanthomonas, Moko and blood ­ bacterial wilt diseases caused by the insect trans­ mission of their causal agents to male flower bract scars in areas where these pathogens are endemic. However, under certain conditions, fungi can enter the wound and cause a rot (Plate 4.13). Usually, the rot only extends a few centi­ metres and is checked in the vicinity of the false hands (small transition hands between com­ pletely female and male flowers). However, dur­ ing long periods of high humidity, the rot occasionally extends further and prevents the development of lower hands. Sometimes, rots originate in the old male flower bud if it is not removed (Stover, 1972). In northern Australia, fungi isolated from the interface between necrotic and living tissue of affected peduncles include Fusarium oxysporum, F. incarnatum (syn. F. pallidoroseum), F. solani and Musicillium theobromae. In Central America, Lasiodiplodia theobromae is frequently isolated from rotten peduncles and sometimes a bacterial soft rot is present (Stover, 1972). The main agents responsible for peduncle rot in the Philippines are Colletotrichum musae, L. theobromae and Fusarium spp. (Quimio, 1986). Control measures are not usually warrant­ ed. However, in commercial banana planta­ tions, it is common practice to leave one finger of a small apical hand attached to the peduncle when removing the false and small apical hands. This is believed to prevent any peduncle rot from reaching the remaining hands by keeping the vascular tissue in the peduncle ac­ tive, thus enabling the peduncle tissue to resist ­infection.



Fungal Diseases of Banana Fruit

Plate 4.13.  Rot affecting the broken end of the peduncle of ‘Grand Nain’ (AAA Cavendish subgroup) in a commercial plantation in Ecuador. (photo: D.R. Jones, INIBAP).

269

sprays of oil-based fungicides. It was thought at the time by Stover (1972) to be the most important of the fruit spotting diseases in Central America. Pitting disease was a serious problem in Mexico, Taiwan and the Philippines, where it is known as pitmark, in the 1970s. It was also im­ portant seasonally in Ecuador and Colombia, but rare in Suriname and the Caribbean. In Cen­ tral America, the disease was more severe on the Pacific coast than on the Atlantic coast. It has not been reported as a problem in most of Africa, although hands exported from Cameroon in late 1970 had a 14% incidence. It was also thought to be common in Madagascar (Stover, 1972). The disease causes an unsightly blemish, which was unacceptable to consumers of export fruit. In the 1960s, up to 50% of fruit arriving at some packing stations in Central America were dis­ eased, causing significant losses due to the rejec­ tion. Also, because the disease could develop while fruit was in transit and during ripening, consign­ ments with less than 2% incidence at the packing station were known to have a 60% incidence at markets. However, these high incidences only oc­ curred following long periods of heavy rainfall and on plantations where control measures were inadequate (Stover, 1972). Since 1980, the dis­ ease has declined considerably in significance. Symptoms

Pitting Introduction Wastage caused by pitting disease was first de­ scribed by Tomkins (1931) in England on fruit of ‘Dwarf Cavendish’ (AAA, Cavendish subgroup) imported from Brazil. Pitting disease was subse­ quently investigated in Central America (John­ ston, 1932), where it was a serious problem on ‘Gros Michel’ (AAA) and became known there as ‘Johnston spot’, and in Brazil (Wardlaw and McGuire, 1932). In the 1930s, it was also record­ ed in Australia, as ‘black pit’. Here the ­incidence was reported as usually slight, but ­occasionally severe. Pitting disease disappeared in Central America from 1936 onwards, when plantations were routinely treated with high-volume sprays of Bordeaux mixture to control Sigatoka leaf spot. However, it reappeared around 1958– 1960 with the conversion to low-volume aerial

Pitting disease gets its name from the round, sunken pits that appear on fruit as it approaches maturity or after harvest (Plate 4.14). The pits average 4–6 mm in diameter and are surround­ ed by a reddish-brown zone, which in turn is surrounded by a greenish, narrow, water-soaked halo. Although the pit centres sometimes split, the damage does not extend to the pulp. Under favourable conditions during transit and ripen­ ing, the number of pits on fruit may increase fivefold or more. The pathogen does not sporu­ late in lesions on fruit (Stover, 1972). Kim et al. (1987) reported symptoms on fruit to be circular to oval, reddish-brown or dark brown depressed spots, which measured up to 20 mm in diameter. Fruit on the side of the bunch facing away from the pseudostem is more severely affected than fruit facing towards the pseudostem. The larger proximal hands have more pits than the smaller distal hands. Small pits can also occur on pedicels and crowns, which can increase

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t­ ransition leaf and bracts. Conidia are carried to banana fruit by air currents. Although these spores are found in the air throughout the year in Central America, they are much more com­ mon during periods of high rainfall. In the field, conidia germinate and form appressoria on green fruit within 4–8 h in a saturated atmos­ phere. The optimum temperature for infection is 24–26°C. Although pits were produced on young fruit 2–3 weeks after inoculation in the laboratory, symptoms are rarely seen in the field until 70 days after bunch emergence. This is be­ cause the fungus remains dormant until the fruit is nearing maturity and then spreads through the tissue intercellularly (Meredith, 1963b). In Central America, the disease was only severe from 6 weeks after the beginning of the rainy season until the beginning of the dry sea­ son (Stover, 1972). Host reaction

Plate 4.14.  Symptom of pitting disease in Honduras (photo: J. Rivera, FHIA).

­nger drop. After periods of abundant rainfall, fi larger, shallower pits appear on spathaceous bracts (transition leaves) and on young watersuckers. The pathogen can sporulate in these tissues as they decay (Stover, 1972; Slabaugh, 1994c). Causal agent Pitting disease was initially believed to be caused by Pyricularia grisea, the sexual stage Magnaporthe grisea being rarely seen. It was thought that grass hosts of P. grisea, such as Digitaria sanguinalis, were sources of inoculum for banana. However, the causal agent of pitting has been shown to be Pyricularia angulata (see Pyricularia leaf spot in Chapter 2). This species has only been detected on banana.

Fruit of Cavendish types (AAA) was more severe­ ly pitted than the fruit of ‘Gros Michel’ (AAA) or cultivars in the Plantain subgroup (AAB) (Stover, 1972). ‘Silk’ (AAB) was almost always affected (Wardlaw, 1961). Control Collapsed and dying banana leaves, the transi­ tion leaf and bracts need to be removed from plants at regular intervals during the rainy sea­ son to reduce inoculum levels. Dithiocarbamates were more efficacious for control than benomyl and other fungicides in Nicaragua (Guyon, 1970). Two spray applications were found to be better than one and mist spraying was not ­effective. Dusting the polyethylene bag with a fungicide has also been shown to be effective (Slabaugh, 1994c). Today, disease incidence is slight and chemical control measures are seldom necessary (Stover and Simmonds, 1987).

Sooty Mould and Sooty Blotch Disease cycle and epidemiology Pyricularia angulata is a common inhabitant of hanging banana-leaf trash, including the

Sooty mould is the name given to the superficial peel blemish caused by various saprophytic ­fungi growing on honeydew from banana aphids



Fungal Diseases of Banana Fruit

and mealybugs. It is usually only a problem on bunches covered with polyethylene bags that are not insecticide-impregnated. Sooty blotch is dif­ ferent in that the growth of the causal agents is not dependent on honeydew from phloem-feeding insects. Sooty mould is seasonal in appearance, be­ ing most common during cool, rainy weather. The condition gets its name from the black, soot-like mycelium of the causal fungi on the fingers, pedicels, crowns and peduncles. Discol­ oration is usually greatest near the pedicels and between the fingers, as the insects producing the honeydew here are shaded and protected (Plate 4.15).

271

When rubbed, some of the fungal mycelium is removed, but a black or dark brown stain re­ mains. Fungi associated with sooty mould in Latin America have been identified as Chaetothyrina musarum, Leptoxyphium fumago (syn. Fumago vagans), Cladosporium atriellum, Microxyphium sp. and Penicillium citrinum (Stover, 1972, 1975). However, Cladosporium cladosporioides may be the main agent (Stover, 1975). Fruit affected by sooty mould is not marketable. Sooty mould is controlled by applying insecticide sprays to the bunch or using polyethylene bags impregnated with insecticide to protect fruit. Biocontrol agents may also be useful in insect control. Ants that farm aphids and mealy bugs should be con­ trolled in plantations. In Australia, sooty blotch, caused by the fungus Chaetothyrina musarum, is a light brown, superficial blemish (Plate 4.16). Rubbing the fruit makes no difference to the symptoms as it does with sooty mould. It is considered a minor disease, most likely to occur in cool, wet weather. Control is not warranted (Jones et al., 1993).

Postharvest Diseases D.R. Jones and I.F. Muirhead Plate 4.15.  Symptoms of sooty mould on the crown and pedicels of fruit of a Cavendish cultivar on a farm in Guadeloupe following a pesticide reduction programme. The black fungus is growing on the honeydew of mealybugs and aphids. (photo. L. Pérez-­ Vicente, INISAV).

Plate 4.16.  Symptoms of sooty blotch in Australia (photo: QDPI).

Overview After harvest, the banana fruit is subject to a range of fungal diseases with potential to cause significant damage. These diseases and their im­ portance in international trade were described by Wardlaw (1961), Meredith (1971), Stover (1972), Slabaugh and Grove (1982), Stover and Simmonds (1987), Jeger et al. (1995) and Thompson and Burden (1995). These authors traced the history of postharvest disease, begin­ ning at the time when whole bunches ripened and rotted during long sea voyages from Central America to Europe. They revealed how losses were reduced by improved management of rip­ ening through better temperature control and reduced transit times, and how crown rot emerged as a problem when bananas were exported in boxes rather than on the bunch in the early 1960s. They also described the use and effective­ ness of fungicides to control postharvest diseas­ es. However, despite all the progress, postharvest

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diseases of banana are still a significant problem. It was estimated they were responsible for 20% of harvest losses in Sri Lanka in 1997 (Anthony et al., 2004). With some exceptions, fungicides are gener­ ally effective in controlling postharvest diseases when other parts of the fruit-handling system are well managed. However, in recent times, there has been a worldwide trend away from the routine use of pesticides, particularly after harvest (Eckert, 1990). This trend has been due to consumers be­ coming aware of the damage that some agricul­ tural chemicals cause to the environment and to concerns that chemical residues on produce could lead to health problems. At the same time, the consumer is demanding higher-­quality fruit with fewer blemishes caused by pests and diseases. Today, except for organically grown banan­ as, most banana fruit that is exported is treated with fungicide to reduce the incidence of post­ harvest disease. Achieving postharvest disease control in bananas in international trade with­ out using fungicides will require an appreciation and detailed knowledge of all the factors affect­ ing both ripening and fungal development. One of the keys to minimizing losses from postharvest disease is to make use of the natural resistance of green banana fruit by preventing the initiation of ripening. Another is to control the postharvest environment both before and af­ ter ripening begins. The close association be­ tween the activity of pathogenic fungi and the physiological state of the banana fruit is high­ lighted in this chapter. The ripening process in bananas is initiated by ethylene gas, which is produced naturally by fruit tissues. Most postharvest disease problems of banana occur during ripening. The interna­ tional trade in Cavendish bananas makes effec­ tive use of temperature control to prevent fruit ripening during transit. Many ships transporting bananas also have controlled atmospheric condi­ tions to further reduce the incidence of ripening. However, banana is a staple food crop in many tropical and subtropical countries where refriger­ ated road and rail trucks and sophisticated stor­ age facilities are not available. Fruit is also more likely to be damaged in these countries as a result of rough handling during harvest and transit to urban markets. Postharvest diseases are a major cause of wastage under these conditions. Al­ though it has been shown that it is possible to

hold fruit for long periods without refrigeration in simple controlled atmospheres generated with­ in polyethylene bags (Scott et al., 1971), diseases are one of the factors limiting the success of this technology. Most scientific investigative work on post­ harvest diseases of bananas has been undertaken on fruit from Cavendish cultivars, the mainstay of the international export trade. However, the principles underlying the control of postharvest diseases apply equally to bananas from other cul­ tivars that may be marketed in the country of production. Non-infectious problems associated with postharvest fruit are described in Chapter 8.

Crown Rot Introduction Crown rot is a disease of the pad of tissue severed when the hand of fruit is cut from the bunch. Crown rot rose to prominence in international trade after Fusarium wilt in Central and South America led to the replacement of ‘Gros Michel’ (AAA) with the wilt-resistant cultivar ‘Valery’ (AAA, Cavendish subgroup) in the late 1950s and early 1960s (Slabaugh and Grove, 1982). Unfor­ tunately, fruit of ‘Valery’ was more susceptible to scarring and bruising than fruit of ‘Gros Michel’, which had been exported as whole bunches. As a result, the trade moved to de-handing and marketing hands or clusters of fruit in boxes, a procedure that also allowed greater efficiencies in packing and handling. However, the process of cutting the hand from the bunch, dividing it into clusters and pulling off individual rejected fingers exposed wounded crown tissue to infec­ tion by fungi and the development of crown rot. Crown rot is still a major problem that caus­ es losses, as is evidenced by the large number of articles on the subject published in scientific journals in recent years.

Symptoms Crown rot is a firm, dark brown or black rot, which spreads through the crown and may pen­ etrate into the pedicels of individual fingers (Plate 4.17). A layer of fluffy white, grey or pink



Fungal Diseases of Banana Fruit

Plate 4.17.  Fruit of a Cavendish cultivar affected by crown rot. The decay has spread from the crown to the pedicels and fingers. Note the anthracnose spots that develop during ripening on the fingers (photo: D.R Jones, QDPI).

fungal mycelium may cover the cut surface of the crown (Plate 4.18). The mycelium and the rot itself spoil the fresh, clean appearance of the ripening fruit. Individual fingers may fall from the weakened crown if the rot penetrates deeply. In severe cases, the decay reaches the pulp itself and the entire fruit is lost. Disease development stimulates ripening (Slabaugh and Grove, 1982), which is another cause of wastage in interna­ tional trade.

Causal agents Crown rot is caused by a fungal complex. Fungi commonly isolated from diseased tissue have been Musicillium theobromae (syn. Verticillium thoebromae – see Zare et al., 2007), Colletotrichum musae, Thielaviopsis paradoxa (syn. Ceratocystis paradoxa – see de Beer et al., 2014), Thielaviopsis musarum (Pereira de Melo et al., 2016), Lasiodiplodia theobromae (syn. Botryodiplodia theobromae), Nigrospora sphaerica (most likely N. musae – see under ‘squirter’, this chapter), Cladosporium sp., Acremonium sp., Penicillium sp., and Aspergillus sp., as well as many Fusarium spp., including F. incarnatum (syn. F. semitectum and F. pallidoroseum), F. verticillioides, F. sporotrichioides, F. oxysporum and F. solani (Lassois et al., 2010). Many banana-attacking isolates of F. verticilloides have been recognized as a distinct spe­ cies named F. musae. Although morphologically very close to F. verticilloides, which can attack a range of hosts, F. musae is more pathogenic on

273

Plate 4.18.  A white fungal mycelium is visible on the surface of a crown affected by crown rot (photo: D.R. Jones, QDPI).

banana, does not produce the mycotoxin fumoni­ sin and has been isolated only from banana fruit (Van Hove et al., 2011). It is also of interest as a human pathogen (Triest and Hendrickx, 2016). The fungi associated with the disease in boxed bananas exported from the Windward ­Islands to the UK have been studied thoroughly. Johanson and Blazquez (1992) showed that the main pathogens were species of Fusarium, principally F. incarnatum (syn. F. pallidoroseum), and Colletotrichum musae. A series of other fungi known to cause crown rot were also isolated, ­including Lasiodiplodia theobromae, Musicillium ­theobromae and species of Nigrospora. This was consistent with the earlier studies in the Wind­ ward Islands by Wallbridge and Pinegar (1975), Griffee and Burden (1976) and Wallbridge (1981). Knight (1982) considered F. oxysporum, F.  verticilliodes and F. graminearum, which were frequently isolated from crowns, to be the prima­ ry pathogens, whereas L. theobromae, M. theobromae and N. sphaerica (musae?) were considered to be less important. Later, Finlay and Brown (1993) carefully examined the relative impor­ tance of the different pathogens in the Wind­ ward Islands and believed C. musae to be by far the most aggressive species. However, isolation studies undertaken in India suggest that L. theobromae was the major pathogen responsible for crown rot and that C. musae and Fusarium spp. were the minor pathogens. Cross-inoculation experiments, conducted using five different viru­ lent isolates of L. theobromae on five different commercial cultivars of banana, demonstrated that generally the isolates were more pathogenic on the cultivar from which they were isolated

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(Thangavelu et al., 2007). Lasiodiplodia theobromae isolated from various banana cultivars in ­India has shown high genetic variability (Ganesan et al., 2012). Fusarium incarnatum (syn. F. pallidoroseum) and a Verticillium sp. (which may have been Musicillium theobromae) were the most common pathogens isolated from the crowns of fruit cut in North Queensland, Australia (Jones, 1991). Of other fungi isolated infrequently, only an Acremonium sp. caused severe crown rot in inoc­ ulation experiments. Colletotrichum musae was not isolated, but was commonly seen sporulat­ ing on the surface of necrotic pedicels of fruit in the advanced stages of crown rot. Colletotrichum musae and F. incarnatum (syn. F. pallidoroseum) or F. verticillioides (syn. F. moniliforme), acting alone or together, were the dominant crown rot patho­ gens in Mauritius (Lutchmeah and Santchurn, 1991) and Malaysia (Sepiah and Nik Mohd, 1987). Fusarium spp. feature prominently in China (Wang et al., 1991) and Central and South America (Stover, 1972). In Costa Rica, Marin et al. (1996) found F. verticillioides and F. incarnatum were the most pathogenic species. Fungi associated with crown rot disease in organic banana plantations in the Dominican Republic were found to be dominated by Fusarium (47%), followed with Lasiodiplodia (7.4%) and Colletotrichum (1.1%) (Kamel et al., 2016). The fungi associated with crown rot differ in pathogenicity and prevalence in various parts of the banana-growing world. The severity of crown rot and the fungi causing the disease vary during the year and from location to location (Lassois et al., 2010). Fungal populations may differ also between banana plants and even be­ tween crowns (Lukezic and Kaiser, 1966). It is important to know which pathogenic fungi are dominant in any given situation, as the effective­ ness of control treatments may depend upon a correct diagnosis. Disease cycle and epidemiology The fungi causing crown rot are ubiquitous components of the microflora of banana planta­ tions (Stover, 1972; Stover and Simmonds, 1987). They live saprophytically in dead banana leaves, flower bracts, discarded fruit and bunch stems. In the field, airborne spores of fungi, such as

Fusarium spp. and M. theobromae, settle on ba­ nana bunches while water-borne spores, such as the conidia of C. musae, are splashed on to fruit by rain or irrigation water. Spores of some of the crown rot pathogens may remain viable for months in the field under extremes of relative humidity and temperature (Stover, 1972). In Gua­ deloupe, the peak of inoculum of both C.  musae and Fusarium spp. in close proximity to plants occurred between 25 and 40 days after bunch emergence (de Lapeyre de Bellaire and Mour­ ichon, 1997). The flowers and the last bracts were the main sources of inoculum. Colletotrichum musae establishes latent infections, which survive in green banana fruit and probably crown tissue for several months (Muirhead and Deverall, 1981). Thus, at harvest, fruit often carries a heavy load of crown rot inoculum. Cutting the hand from the bunch results in a number of opportunities for crown rot fungi. Firstly, it creates a substantial wound, which is directly accessible to airborne or water-borne fungal spores. Spores present in water used to wash bananas can be drawn into this wounded tissue to a depth of 5–7 mm (Greene and Goos, 1963). Secondly, the de-handing knife itself can drag inoculum from the surface of the crown across the wounded tissue (Finlay et al., 1992). Thirdly, wounding may activate previously estab­ lished latent infections of C. musae (Jones, 1991). Spores that reach the surface of the cut crown germinate in response to the favourable environmental and nutritional stimuli. The pro­ gress of disease from that point depends on sev­ eral factors, principally the amount of inoculum, the physiological state of the fruit, the length of time before ripening commences and the envi­ ronmental conditions before and after ripening (Finlay and Brown, 1993; Chillet and de Lapeyre de Bellaire, 1996b; Lassois et al., 2010). In the Windward Islands, crown rot is a problem all year round, but is more severe during the more humid, wetter and hotter summer months. Kraus and Johanson (2000) reported the highest inci­ dence during the rainy period. In Honduras, crown rot was more prevalent during the sum­ mer and declined in the coldest months (Lukezic et al., 1967). It is thought that heat and moisture favour increased activity of the crown rot fungi in leaf trash and discarded fruit. Several authors (see Johanson and Blazquez, 1992) have mentioned the possibility that ­enzymes



Fungal Diseases of Banana Fruit

produced by saprophytic bacteria may contrib­ ute to tissue breakdown, but this has not been confirmed by detailed research. What is known is that C. musae is able to produce polygalacturo­ nase isozymes capable of effectively macerating green banana tissue (Stanley and Brown, 1994). Phenols have been implicated in the susceptibili­ ty of bananas to crown rot (Ewané et al., 2012). Attempts have also been made to identify genes involved in the response of banana to crown rot (Lassois et al., 2011).

Host reaction Stover (1972) reported that all commercial des­ sert bananas, which are produced on cultivars of the AAA genome, were susceptible and that the problem was rare on fruit from cultivars in the Plantain subgroup (AAB). In the Cavendish subgroup (AAA), ‘Robusta’ and ‘Valery’ were found to be more susceptible to natural crown rot infections than ‘Lacatan’ (Shillingford and Sinclair, 1977). It has also been suggested that fruit of the hybrids ‘FHIA-01’ (AAAB, syn. ‘Goldfinger’) and ‘FHIA-02’ (AAAA, syn. ‘Mona Lisa’) from the Honduran breeding programme is less susceptible to crown rot than Cavendish cultivars (Marin et al., 1996). However, other workers consider that fruit of these two hybrids is more susceptible to crown rot caused by F. incarnatum and C. musae than ‘Grande Naine’ (AAA, Cavendish subgroup) and only ‘FHIA-23’ is more resistant (Perez-Vincente and Hernanan­ dez, 2002). In India, ‘Robusta’ (AAA, Cavendish subgroup) was highly susceptible to all the iso­ lates of L. theobromae tested, with ‘Karpuravalli’ (ABB, Pisang Awak subgroup) being less suscep­ tible (Thangavelu et al., 2007). Fruit of ‘Pisang Berangen’ (AAA) in Malaysia and fruit of ‘Kluai Hom Thong’ (AAA) in Thailand suffer from crown rot (Mohamed et al., 2017; (Jitareerat and Uthairatanakij, 2013). Control Preventing infection The amount of inoculum in the environment is thought to affect levels of crown rot. It is now standard practice on many plantations to ­remove

275

dead banana leaves and flowers from plants and keep areas where fruit is packed free of banana trash. Water used in washing and de-­latexing tanks is also treated with a sanitizer, such as chlorine. This latter procedure recommended for control of crown rot is sound, but in practice may have a limited effect. Slabaugh and Grove (1982) noted that maintaining the desirable concentration of chlorine in wash water did not reduce crown rot, suggesting that sources of in­ oculum other than contaminated water were more important. The importance of field inocu­ lum is highlighted in studies in the Windward Islands, where fruit was cut and packed in the field in an attempt to reduce the physical dam­ age caused during the transport of bunches to boxing plants (Thompson and Burden, 1995). This fruit was not washed and, even though a cellulose pad containing the fungicide thiaben­ dazole was used on the crown to stem the flow of sap, crown rot still caused losses (Johanson and Blazquez, 1992). Finlay et al. (1992) investigated the impor­ tance of different sources of crown rot inoculum in the Windward Islands and the effects of alter­ native handling systems on development of crown rot. In experiments, they reduced the dis­ ease by more than 50% by de-handing (without washing) in a relatively clean packing shed ­instead of the field. Although the benefit was not significant in commercial shipping trials (in which disease levels were naturally low), the work provides a good argument for keeping the packing area free of leaf trash, floral remnants and discarded fruit. Reducing inoculum levels in the field and the packing area should be seen as only the first steps in controlling crown rot. Other methods are generally needed as well. De-handing technique The way in which the hands are removed from the bunch is another factor that affects develop­ ment of crown rot in green tissue. The standard texts recommend a clean cut with a sharp tool and neat trimming. Finlay et al. (1992) showed that rough trimming and breaking crowns from the bunch stalk instead of cutting increased crown rot. The action of breaking crowns left fragments of tissue, which readily dehydrated and senesced, providing conditions favouring the

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establishment of crown rot fungi. The crowns of the broken hands also yellowed more quickly, suggesting premature ripening of the tissue, which also favours fungal development. Smooth­ ly cut large crowns are known in the banana trade to be more resistant to crown rot than ir­ regularly cut shallow crowns. It is always pru­ dent to trim hands leaving as much crown tissue as possible. As well as being more resistant to rot, any rot that may initiate at the cut surface has less chance of spreading to the pedicels and fingers before sale and consumption. Managing the ripening process Bananas harvested green pass through three physiological stages (John and Marchal, 1995):

• • •

the pre-climacteric or pre-ripening phase, during which the fruit’s metabolic activity is relatively low; ripening, which begins with an intense burst of respiration, called the climacteric; and senescence, during which metabolism slows once more.

Fruit grown for international and some do­ mestic markets must be prevented from ripening while in transit. This is achieved by harvesting the fruit at a physiological age to ensure an extended green life and by transporting the fruit under con­ trolled temperature conditions (13–14°C). The minimum temperature for transport is 12°C, as lower temperatures cause the peel to turn brown due to oxidization (Lassois et al., 2010). Sometimes the relative humidity and composition of the stor­ age atmosphere is also controlled. The transit peri­ od may be 1 week or more and fruit must be harvested with sufficient green life to ensure arriv­ al before ripening commences. Applying ethylene in special rooms under controlled conditions then synchronizes ripening. If ripening begins en route, the fruit arrives at its destination ‘mixed ripe’, with hands at different stages of yellowing. Mixed-ripe fruit cannot be marketed successfully, often devel­ ops crown rot and is wasted. In commercial prac­ tice, premature ripening is prevented primarily by harvesting at the appropriate stage of maturity, which is judged by the age of the bunch, the size or ‘grade’ of the fruit and the internal colour of the pulp. Less mature fruit with a longer green life is used for more distant markets.

There is a very close relationship between the stage of ripening and the resistance of the banana to postharvest diseases, including crown rot. During the pre-climacteric phase, the fruit retains much of the resistance of the unharvest­ ed fruit. This resistance is lost once ripening commences and the ripe fruit very rapidly be­ comes fully susceptible. The factors responsible for mediating resistance to C. musae are dis­ cussed under anthracnose and are likely to be similar for other crown-rotting fungi. Incidence of crown rot also depends on how the fruit is managed after harvest. When the period between harvest and artificial ripen­ ing is short and temperatures in transit are con­ trolled, fruit reaches the consumer quickly and crown rot is rarely a problem. This is commonly the case in Australia, where the transit time from farm to market is as short as 1–3 days. In Australia, crown rot is associated with longer transit times, attempts to hold fruit before ripen­ ing to control market supply, or suboptimal envi­ ronmental conditions. In these cases, the resistance of the green fruit is insufficient to pre­ vent disease development and other factors need to be considered. Effects of modified atmosphere The main reason for modifying the atmosphere around green bananas during the pre-climacteric period is to extend their green life and hence prevent the premature initiation of ripening during transit and storage. Ripening is delayed by reducing the oxygen level to 3–7%, increas­ ing carbon dioxide to 10–13% and absorbing ethylene (Wade et al., 1993). The modified at­ mosphere technique, in combination with re­ frigeration, is now in use in international trade. Usually, the atmosphere in the holds of the ships containing the banana cartons is modified using sophisticated equipment. However, the benefits of modified atmosphere storage can be achieved in other ways. Fruit can be packed in polyethyl­ ene bags and a vacuum applied to each bag be­ fore it is sealed and the carton closed. The atmosphere within the bag is modified by the respiratory activity of the fruit. If the fruit is then transported under refrigeration, the system combines the beneficial effects of reduced tem­ perature and modified atmosphere. As well as extending green life, it is claimed that a modified



Fungal Diseases of Banana Fruit

atmosphere also reduces the incidence of crown rot (Slabaugh and Grove, 1982; Stover and Sim­ monds, 1987). Chillet and de Lapeyre de Bellaire (1996a) found that the beneficial effect of a modified atmosphere on crown rot caused by C. musae was more noticeable over a long period (28 days at 13.5°C), as compared with a short storage period. The modified atmosphere, as op­ posed to reduced water loss, controlled the crown rot. Once ripening is initiated, the process is ir­ reversible, shelf-life is limited and storage is im­ practical. The possibility of holding green bananas in a pre-climacteric state without re­ frigeration for 40 days or even longer (Satyan et al., 1992) is potentially important in tropical and subtropical countries where refrigeration is not available. It has been known since at least 1966 that green life can be extended to such ­periods without refrigeration by enclosing pre-­ climacteric bananas in sealed polyethylene bags with an ethylene absorbent (Scott and Roberts, 1966). However, under these conditions, post­ harvest disease is a major constraint and Scott et al. (1971) considered the use of fungicides to be essential. Experimentation on long-term modified at­ mosphere storage reveals a complex pattern of different fungal species causing different amounts of damage, depending on the atmospheric com­ position, the part of the bunch or hand affected and the fungicidal treatment. While some patho­ gens were unaffected or suppressed by modified atmosphere, others, including Alternaria alternata and F. incarnatum (syn. F. pallidoroseum), were stimulated. However, the main pathogens affect­ ing crown tissue were still C. musae and Fusarium species (Wade et al., 1993). Effects of temperature Most fungi causing crown rot come from tropi­ cal environments and are favoured by tropical temperatures. For example, the optimum tem­ perature for growth of C. musae is 28°C. Storage, transport and ripening temperatures are deter­ mined by the requirements of the fruit. As ba­ nanas from Cavendish cultivars are damaged at 12°C or below, the fruit is stored and shipped at 13–14°C. During ripening, the fruit is held at the optimum of 16–18°C, with some adjustments of temperature to control the ripening speed. These

277

temperatures are well below the optimum for the growth of most crown rot fungi. After harvest, fruit should be cooled as soon as possible and certainly within 48 h if transit times are going to be longer than 10 days (Stover, 1972). Cooling soon after harvest maintains green life as well and, to some extent, limits the development of crown rot. The longer that fruit is held at ambi­ ent temperatures before refrigeration, the more chance there is of crown rot developing. The development of crown rot caused by Thielaviopsis paradoxa has been investigated by artificial infection of green mature Cavendish bananas at temperatures between 10°C and 26°C. Results showed the optimal temperature for the development of this particular pathogen was only 18°C, with the fruit pulp being invaded within 6 days. The disease did not develop at 10°C. Only minor mycelial growth was observed on crowns at 22°C and 26°C (Samuelian and Vawdrey, 2014). Effects of a high humidity level Stover and Simmonds (1987) noted that any pack that maintains high humidity and keeps crowns turgid and fresh helps to reduce crown rot, but there appear to have been no specific studies undertaken on the effects of different hu­ midity levels on development of crown rot dur­ ing storage. One would expect the growth of superficial mycelia on the surface of decaying crowns to be encouraged at a high humidity. However, the overriding effect of high humidity levels must be that they prevent the desiccation of the crown, which favours tissue death and fungal colonization. The green life of bananas is also considerably reduced when the humidity is relatively low (30–40%) as a result of ethylene production from the fruit peel (Peacock, 1973). Fungicidal treatments When the world export trade moved from ship­ ping bananas in boxes instead of on the bunch, the cut crown tissue became a major site of fungal infection. However, it was also easy to treat the exposed crowns with a fungicide to protect them during transit and ripening. The major break­ through in chemical control occurred when the first of the highly effective systemic benzimidazole fungicides (thiabendazole and then benomyl)

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­ecame available in the late 1960s and early b 1970s. These systemic fungicides did not pene­ trate far into the fruit, but the effect was sufficient to control crown rot well. Over the past 30 years, many fungicides from other chemical groups have been tested and some, including imazalil and prochloraz, have been approved for use in dif­ ferent countries. In Central America, imazalil is often used for the distant European market and, although more expensive, is regarded as more ef­ fective than the benzimidazoles (Slabaugh and Grove, 1982). Imazalil is also now used on most Windward Islands fruit, with good results. Prochloraz was recommended for use in Australia in the months of glut when bananas were held longer at markets and crown rot tended to be more of a problem (Jones, 1991). This fungicide has recently been found to be the most effective for the control of T. paradoxa in in vitro tests (­Samuelian and Vawdrey, 2014). Before a fungicide can be registered for use on bananas, its efficacy must be proved and au­ thorities satisfied that it will not endanger public health. The regulations of the country where the fruit is to be marketed must also be taken into account. The necessary trials and tests are usually conducted or sponsored by the chemical company promoting its use. Fungicides are applied in many different ways (Thompson and Burden, 1995). The sim­ plest, now in use in the Windward Islands, is to immerse fruit in a solution of fungicide in a pol­ yethylene tub at the farmer’s packing shed. As well as fungicide, the dip solution contains alum to counteract latex staining. The cut surfaces are allowed to drain latex for a maximum of 10 min before immersion. The time between cutting and dipping is regarded as critical, as the risk of crown rot is believed to increase the longer the cut surface is exposed without protection. In commercial packing houses in Central America, treatment is integrated into a mechanized pack­ ing operation. Although dip tanks are common, fungicide solutions can also be sprayed or cas­ caded on to hands or clusters placed crown up on slowly moving trays. In addition, fungicides have been applied to crowns using a brush (Guz­ mán and Villalta, 2008a). There are some practical problems associat­ ed with the use of fungicides. For health reasons, personnel in contact with fungicides, fungicide solutions or fungicide-treated fruit should wear

appropriate protective clothing (Plate 4.19). Such practices are not always adopted in hot, tropical environments, where protective cloth­ ing may be uncomfortable. If the mixture of fun­ gicide is used continually or recycled, its strength declines with time and the passage of fruit. Dirt, sap and spores of fungi tolerant to the fungicide may also build up and reduce efficacy. To main­ tain the fungicide at an effective concentration, it becomes necessary either to use a topping-up procedure or to change the mixture regularly. The correct disposal of the used fungicide mix­ ture is also now seen as an important environ­ mental issue. Some producers in Costa Rica absorb the fungicide on to clay before disposal. In the Windward Islands, the use of charcoal-­ lined disposal pits is advocated. Applying the fungicide in pads, which are placed directly on to the crown of hands, overcomes some of these problems (Johanson and Blazquez, 1992). Un­ fortunately, pads are unsightly and do not ­appeal

Plate 4.19.  Banana worker in protective clothing dipping fruit in a fungicide–alum–water mixture in St Vincent, Windward Islands (photo: D.R. Jones, SVBGA).



Fungal Diseases of Banana Fruit

to consumers. In addition, pads do not work as well on clusters, the form of fruit now commonly exported. The more irregular crown of the clus­ ter means that the pad with the fungicide does not come into contact with all of the cut tissue. If a fungicide used to control crown rot is also used regularly to control leaf spot diseases on the same farms, there is a danger that the population of crown rot fungi in the field may become less sensitive to that fungicide and relat­ ed fungicides (Johanson and Blazquez, 1992). In Guadeloupe, de Lapeyre de Bellaire and Dubois (1997) reported that an average of 23% of iso­ lates of C. musae were tolerant of thiabendazole. Their presence was correlated with the exclusive use of benomyl as an aerial spray to control ­Sigatoka leaf spot in the field. Other control methods With the increasing popularity of organic farm­ ing, there has been an increase in public opposi­ tion to the use of postharvest pesticides of all kinds. Du Pont withdrew benomyl (Benlate®) as a postharvest treatment for fruit and vegetables some time ago. With chemical use on the de­ cline, there is renewed interest in alternative means of control. Recent work has shown that hot water may have the potential to replace chemical fungicides. In tests in Hawaii, a 45°C hot water treatment for 20 min given 15–20 min after de-­handing re­ duced the incidence of crown rot caused by T. paradoxa from 100% to less than 15% (Reyes et al., 1998). Hot-water dips may also be effective against Fusarium and Colletotrichum (LópezCabrera and Marrero-Domínguez, 1998). In the Philippines, C. musae, F. incarnatum, L. theobromae and T. paradoxa are involved in crown rot of fruit of ‘Buñgalan’ (AAA, Cavendish sub­ group). The maximum exposure of fruit to hot water before deleterious effects were recorded was 50°C for 20 min. Hot-water treatment was said to delay ripening and prolong the green life of fruit. It was also seen to improve the overall ap­ pearance of the bananas, reduce weight loss and improve firmness. It was reported to lessen crown rot severity by 50% after 7 days and 33% after 14 days (Alvindia, 2012). However, commercial tri­ als with hot water using naturally infected fruit have not achieved control and ripening delays have also been reported (Lassois et al., 2010).

279

Biological control using microorganisms, such as yeasts or bacteria, is an attractive pros­ pect and much work has been undertaken in this area (Chuang and Yang, 1993; Kraus, 1996; de Costa et al., 1997; Postmaster et al., 1997). While there are significant difficulties in devel­ oping and commercializing biological control agents, there are some factors that would favour this technology for control of crown rot. Firstly, the freshly cut crown provides nutrients and moisture suited to microbial growth. Secondly, the cut crown could be treated within minutes of exposure to most crown rot inocula. Thirdly, most packing systems would allow a controlled dose of microbial agent to be delivered easily and efficiently. Fourthly, the temperature and hu­ midity of the environment while bananas are in transit can be regulated. Fifthly, control is re­ quired only for a limited period, perhaps 1–3 weeks. Finally, the concerns of consumers who influence supermarket chains buying bananas and the size of the international trade might at­ tract commercial investment. Antagonistic mi­ croorganisms would need to be adapted to temperatures of 13–18°C, the range used dur­ ing commercial transit and ripening. However, a much wider temperature range would be needed for storage and handling without temperature control. It is considered by some that biological agents alone will not provide complete control of crown rot and will probably need to be used in conjunction with other measures such as modi­ fied atmosphere packaging (Lassois et al., 2010). Irradiation with ultraviolet light has also been considered for control of crown rot. Un­ fortunately, banana peel is too sensitive to UV for this method to be applicable. Control of postharvest diseases is one of the benefits cited for the postharvest gamma irradiation of fruit. While the doses required for fruit-fly control (75–300 Gy) might be tolerated by the banana fruit (Akamine and Moy, 1983; Burditt, 1994), those required for disease control are 1000 Gy or higher (Moy, 1983) and are likely to be dam­ aging (Thomas, 1986). Mohamed et al. (2017b) reported significant browning of fruit of ‘Pisang Berangan’ (AAA, Lakatan subgroup) following UVC germicidal irradiation except at the lowest dose of 0.01 kj/m2, which reduced crown rot by 46.25%. The opportunities for controlling crown rot by irradiation, therefore, appear limited.

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Natural substances, such as preparations of calcium, plant extracts and organic acids, have been found to have fungicidal activity. However, phytotoxicity has been reported and control is insufficient for commercial operations (see Lassois et al., 2010). Another alternative to using conventional fungicides is to apply edible film-forming poly­ mers or other acceptable food additives, such as organic acids and salts, e.g. potassium sorbate and sodium benzoate. These materials are in­ hibitory to C. musae, one of the major crown rot pathogens, both in vitro and in vivo (Al Zaemey et al., 1993). Although not spectacularly effec­ tive in its own right, this technique might be useful in combination with low levels of fungi­ cide or as part of an integrated control package. Another interesting possibility is treatment with papaya latex, which has given control of crown rot under experimental conditions (Indrakeerthi and Adikiram, 1996). A wax made of sucrose esters with cellulose or fatty acids has also been found to block stomata and inhibit gas exchange. This led to a decrease in oxygen in the fruit with­ out increasing carbon dioxide levels, which cre­ ates an atmosphere for preserving bananas (Banks, 1984). Innovations that improve control over fruit maturity, reduce physical injury and shorten transit times will continue to be investigated and introduced by the banana industries of the world. Improvements in these areas should also reduce the impact of crown rot.

Symptoms Brown spots form on the peel as the fruit ripens to cause a cosmetic blemish, which can deter po­ tential consumers. The spots increase in size and often coalesce to form extensive areas of sunken brown-black tissue (Plate 4.20). Orange or salmon-coloured spore masses develop under ­ the right conditions. Usually, the pulp remains unaffected until the fruit becomes overripe. When green fruit is wounded, black scars are formed. The shape of the scar depends on the type of injury. Injury caused when the ends of fingers impinge on the curved shoulder of fin­ gers on lower hands results in diamond-shaped lesions (Stover, 1972). Lesions are sunken and can have a yellow halo. Sometimes the outer lay­ er of pulp may be invaded. When the fruit is rip­ ened or placed at high temperatures, the lesion increases in size and the fungus invades the pulp. Bananas naturally develop small brown spots on the peel as their sugar content increases on ripening. These ‘sugar spots’ should not be confused with lesions caused by anthracnose.

Causal agent Colletotrichum musae (syn. Gloeosporium musarum) is considered to be the main cause of ­anthracnose.

Anthracnose Introduction Anthracnose is the name given to the disease that appears as sunken brown-black lesions found on the peel of bananas during transport, storage and ripening. Anthracnose arises when dormant infections of the causal fungus in the green peel reactivate as fruit ripens. This results in the gradual necrosis of the yellowing peel from initial brown spots. Anthracnose symp­ toms also appear when green fruit is wounded. Lesions develop to form scars, which spread and become more serious on ripening. Scars detract from the appearance of fruit and lower quality.

Plate 4.20.  Symptoms of anthracnose on fingers of a Cavendish cultivar. The small brown spots that develop during ripening can be seen at the centre of the top finger. Larger black lesions caused following damage to green fruit are evident elsewhere. The white mycelium and salmon-­pinkcoloured masses of conidia of Colletotrichum musae can be seen on the very large lesion on the lower finger (photo: QDPI).



Fungal Diseases of Banana Fruit

Colletotrichum musae is also implicated in crown rot, main stalk rot, stem-end rot and anthrac­ nose rot of preharvest fruit. It is a specialized pathogen of bananas. This fungus produces conidia on conidio­ phores, which arise in acervuli. Acervuli are pri­ marily found on fruit, but can also occur on peduncles, petioles and occasionally leaves. They are usually rounded, elongated, erumpent, up to 400 μm in diameter and are composed of epider­ mal and subepidermal pale brown pseudoparen­ chyma, which becomes subhyaline towards the conidiophore region. Setae are absent. Conidio­ phores, which are formed from the upper pseu­ doparenchyma, are cylindrical, hyaline, septate, branched, subhyaline towards the base and tapered towards the apex. They measure 30 × 3–5 μm,

281

with a single phialidic aperture. Conidia are hya­ line, aseptate, oval to elliptical or cylindrical and often have a flattened base. They measure 11– 17 × 3–6 μm, have an obtuse apex, are variably guttulate and are produced in yellow to salmon-­ coloured masses (Sutton and Waterston, 1970). An acervulus, conidiophores and conidia are ­illustrated in Fig 4.3. Colonies on potato dextrose agar have sparse to abundant white, grey or olive-coloured floccose aerial mycelium. Acervuli, which are dark brown to black, are abundant, irregularly scattered or sometimes confluent and are pro­ duced throughout the culture, particularly in areas devoid of aerial mycelium and around the point of inoculation. Setae are rarely formed. Masses of conidia are salmon-orange and can

20 μm (D)

(C)

(A) 100 μm

(B) Fig. 4.3.  Acervulus (A), conidiophores (B), conidia (C) and appressoria formed from hyphae (left) and germinating conidia (right) (D) of Colletotrichum musae (from Sutton and Waterston, 1970).

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also be formed from separate phialides on the vegetative mycelium. On potato carrot agar, aerial mycelium is sparse and scattered acervuli are not abundant. As the cultures age, the colo­ nies become grey underneath, due to appressori­ um formation. Appressoria are readily formed from vegetative hyphae, where they are navicu­ lar to ovate, becoming irregularly lobed and dark brown, and they measure 6–12 × 5–10 μm (Fig. 4.3). Appressoria formed from germinating conidia are pale brown, subglobose to irregular and up to 8 μm in diameter (Fig. 4.3) (Sutton and Waterston, 1970). The fungus (as Gloeosporium musarum), iso­ lated from fruit of ‘Gros Michel’ (AAA) in Trini­ dad, was reported to produce Glomerella cingulata in culture (Wardlaw, 1934). However, later work indicated that the asexual morph of G. cingulata may have been Colletotrichum gloeosporioides, which can also be isolated from bananas with an­ thracnose (Greene, 1967; Sakinah et al., 2013). Sakinah et al. (2014) found that both C. gloeosporioides and C. musae were associated with ­anthracnose of banana in India, and that C. gloeosporioides was more prevalent than C. musae. Von Arx (1957) noted that C. musae could be ­distinguished from C. gloeosporioides by slightly longer and broader conidia and its faster growth in culture at 24°C (Sutton and Waterston, 1970). Thirteen isolates of the causal agent of an­ thracnose from the fruit of different banana cul­ tivars on sale at a market in Malaysia were compared morphologically and by random am­ plified polymorphic DNA (RAPD) analysis. All were identified as C. musae on morphological characteristics. RAPD banding patterns were also similar, but revealed intraspecific variations within C. musae isolates from different banana cultivars. A dendrogram obtained from the UP­ GMA cluster analysis separated the C. musae iso­ lates from banana into two separate clusters (Zakaria et al., 2009). A similar study of 40 iso­ lates from fruit of different banana cultivars in India showed four different morphological growth types of C. musae in culture. RAPD anal­ ysis of 12 selected isolates revealed a high genet­ ic variability in C. musae isolated from different banana cultivars. A RAPD dendrogram grouped C. musae into three clusters, which may explain differences in morphological characteristics (Sangeetha et al., 2011).

Colletotrichum scovillei (from the C. acutatum complex – see Damm et al., 2012), previously isolated from Capsicum in Indonesia and Thai­ land, has recently been determined to be the cause of anthracnose on fruit of ‘Brazil’ (AAA, Cavendish subgroup) harvested from a planta­ tion in Ledong County, Hainan Province, China. It was reported that 17% of fruit from the plan­ tation was affected (Zhou et al., 2017). Disease cycle and epidemiology Spores of the fungus are produced on senescing banana tissue, including leaves, bracts, discard­ ed fruit and fruit stems. They are dislodged pri­ marily by water. They reach the fruit in the field in rain splash or irrigation water and in the packing shed in recirculated washing water, which accumulates spores from plant debris. The optimum growth and sporulation tem­ perature for C. musae is 27–30°C and there is little growth below 15°C. Conidia adhere to the intact fruit surface (Sela-Buurlage et al., 1991), germinate and form appressoria within 24–48 h. The fungus remains latent until the fruit starts to ripen. Infection also occurs directly through wounds, which stimulate conidial germination and mycelial growth but inhibit appressorial for­ mation (Yang and Chuang, 1996). Anthracnose lesions develop on the main body of the fruit. In commercial trade, the main location is on the angular corners of fruit, which are often scraped, scratched or bruised during packing and general handling. The injuries caused by the tips of the outer whorl of fruit on the inner whorl are also common starting points. These injuries are invaded either by hyphae growing from spores that lodge in the wound or by hyphae growing from latent or quiescent in­ fection that existed before the injury. If the fruit is not injured after harvest, latent infections re­ sume activity during ripening and similar le­ sions form. These lesions develop later in the ripening process and are less damaging and of little concern commercially. The banana–C. musae interaction has been extensively studied under the microscope in ­attempts to understand how latent infection by Colletotrichum species in tropical fruits is con­ trolled. Early work by Simmonds (1941) showed that C. musae could remain inactive in green



Fungal Diseases of Banana Fruit

­ ananas for several months, resuming activity b during ripening. He found that some appressoria produced infection threads and then subcuticu­ lar hyphae before fungal growth ceased. Sim­ monds (1941) believed that these hyphae were the latent structures. Using light microscopy, Muirhead and Deverall (1981) noted that these subcuticular hyphae induced hypersensitive re­ actions in green fruit, visible to the eye as redbrown flecks. The hyphae above cells that had died hypersensitively appeared to play no further role in initiating disease. In contrast, new subcu­ ticular hyphae emerging from appressoria during ripening proceeded to invade the fruit, causing disease. They thus considered the appressoria to be the latent structures. Electron microscopy studies with C. gloeosporioides in avocado fruit showed that the fun­ gus remains latent in the cuticle as an infection peg of variable length (Coates et al., 1993). Similar studies may reveal the same to be true for the ­banana–C. musae interaction. These histopatho­ logical studies are important, because they ­provide a guide to the kinds of physiological mechanisms that control latency. To investigate the physiological processes controlling latency, researchers have followed sev­ eral hypotheses as initially proposed by Simmonds (1963). The first is that a preformed antifungal factor existing in green fruit declines in concen­ tration or effectiveness during ripening, thus al­ lowing the fungus to resume growth. Several such compounds, including 3,4-dihydroxybenza­ ldehyde (Mulvena et al., 1969) and 3,4-dimeth­ oxybenzaldehyde (Abdel-Sattar and Nawwar, 1986), have been identified. Dopamine, the main phenolic substrate responsible for the browning of wounded green banana peel, has also been im­ plicated (Jiang, 1997). However, it remains to be shown that preformed compounds of this kind are present at antifungal concentrations at the sites of latent infection for the duration of the latent peri­ od (Muirhead and Deverall, 1984). The second type of investigation involves antifungal compounds or phytoalexins formed in response to infection. Brown and Swinburne (1981) showed that five inhibitory compounds were produced in necrotic tissue formed when green bananas were challenged by C. musae. One of the major compounds has been identified by Harai et al. (1994) as 2-(4′-hydroxyphenyl)naphthalic anhydride.

283

The host–pathogen interaction is con­ founded by other factors, such as the presence of iron and chelating agents at the fruit surface. Iron inhibits the germination of conidia of C. musae. On the other hand, chelating agents, such as 2,3-dihydroxybenzoic acid (formed from anthranilic acid present in green bananas) and siderophores from epiphytic bacteria, stim­ ulate germination and appressorial formation (­ McCracken and Swinburne, 1979; Harper et al., 1980). Brown and Swinburne (1981) cul­ tured C. musae on media that were either iron-­ replete or iron-deficient and showed that, when conidia from these colonies were used to inocu­ late green fruit, there were variations in necrosis, the formation of antifungal chemical compounds and the speed of disease development during rip­ ening. These interactions are complex and the role of antifungal chemicals in latent infection has not been fully resolved. If antifungal compounds are not involved, stimulation of the fungus by ethylene (Flaishman and Kolattukudy, 1994) or nutrients (Yang and Chuang, 1996) during ripening or changes in the susceptibility of cell walls to enzyme attack (Shil­ lingford and Sinclair, 1980) have been suggested. However, by using an ethylene receptor site inhibi­ tor, ethylene has been shown not to influence the triggering of anthracnose in ripening and wound­ ed fruit (Chillet et al., 2006a). These authors also found that the initiation of wound anthracnose, unlike quiescent anthracnose, was not dependent on peel ripeness. When anthracnose develops in wounds in green fruit, the extent of disease devel­ opment depends, as it does for crown rot, on wheth­ er the fruit is held for a long period before ripening and under what conditions. While the fruit re­ mains green, the lesion is limited, but the longer the pre-ripening period, the larger the lesion. Dis­ ease development accelerates during ripening. Chillet et al. (2006b), working in the French West Indies, found that stressful growing condi­ tions, especially soil flooding, slowed fruit growth, but had no direct effect on fruit susceptibility to C. musae or on green life. They also showed that bananas of the same grade, but of different physi­ ological ages, had markedly different susceptibility to C. musae. Fruit grown in cooler, highland areas was less susceptible to C. musae than fruit of the same physiological age from lowland plantations. A relationship was also found between the ­manganese content of fruit and its susceptibility

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to anthracnose on low-altitude halloysitic and fer­ rallitic soils, where the most variability was ob­ served. Plants producing the most susceptible fruit had high foliar manganese concentrations and low calcium concentrations, and had been grown on acid soils (Chillet et al., 2000). Host reaction Stover (1972) believed that fruit from all dessert cultivars was affected. Of the dessert cultivars, fruit of ‘Gros Michel’ (AAA) is less susceptible to green-fruit wound anthracnose than fruit from cultivars in the Cavendish subgroup (AAA). Fruit of a high caliper grade, which is more sus­ ceptible to wounding than fruit of a lower cali­ per grade, is also more likely to develop wound anthracnose (Stover, 1972). Shillingford and Sinclair (1977) compared the susceptibility of fruit of three Cavendish ­cultivars to postharvest infections. ‘Robusta’ and ‘Valery’ developed more wound anthracnose than did ‘Lacatan’. When inoculated with C. musae, the Jamaican-bred tetraploid ‘Calypso’ (‘65–3405–1’) was resistant to wound anthracnose, ‘Lacatan’ was intermediate in reaction and both ‘Valery and ‘Giant Cavendish’ were equally susceptible. In Malaysia, fruit of ‘Pisang Mas’ (AA, Su­ crier subgroup), ‘Pisang Berangan’ (AAA, Lakatan subgroup), ‘Pisang Awak’ (ABB), ‘Pisang Nangka’ (AAA) and ‘Pisang Rastali’ (AAB, Silk subgroup) have been reported with symptoms of anthrac­ nose (­Sakinah et al., 2014). Anthracnose affects Prata Aña (AAB, Pome subgroup) in Brazil (Maia e ­Silva et al., 2016). Various banana genotypes with resistance to black leaf streak were screened for their reaction to anthracnose in Brazil. ‘Thap Maeo’ (AAB, Mysore subgroup) had the lowest severity while ‘Ambrosia’ (AAAA, Jamaican bred hybrid), ‘PV 42-53’ (AAAB, EMBRAPA bred ­hybrid), ‘FHIA-18’ (AAAB, FHIA bred hybrid) and ‘FHIA-01’ (AAAB, FHIA bred hybrid) were the most susceptible (Pinho et al., 2010). In N ­ igeria, plantain is also reported as susceptible to an­ thracnose (Baiyeri and Nwachukwu, 2005). Control The control of anthracnose depends on the same principles as discussed for crown rot; minimizing

damage to fruit, reducing inoculum levels by us­ ing fresh or clean water to wash fruit, using fun­ gicides to protect fruit and maintaining fruit in a hard, green state until ripening. In the interna­ tional banana trade, injured bananas are reject­ ed during packing operations and subsequent quality checks. Severe cases of wound anthrac­ nose are, therefore, rarely seen. However, scar­ ring is still the main quality defect of green fruit and is a cause for consumer rejection. Anthrac­ nose may cause significant wastage in domestic markets in developing countries, where damage levels are higher and storage temperatures are not controlled. In the semi-arid region in the north of ­Minas Gerais State in Brazil, anthracnose occurs with varying intensities on fruit of ‘Prata Aña’ (AAB, Pome subgroup). Washing the fruit after harvest with neutral detergent and sodium hy­ pochlorite at 2% followed by application of im­ azalil fungicide was determined to be the most efficient control technique (Maia e Silva et al., 2016). In Costa Rica, thiabendazole and imazalil fungicides were used for many years to control postharvest banana diseases with very good re­ sults. However, control problems increased, mak­ ing it necessary to raise the fungicide doses. This led to testing to find alternative fungicides and mixtures of fungicides. With the use of an azoxy­ strobin and thiabendazole mixture it was possible to reduce the fungicide dose by 50% and the treatment cost by approximately 20%, without detriment to control of anthracnose and other postharvest diseases (Guzmán and Villalta, 2008b). In Brazil, the fungicides procloraz and propicona­ zole at a concentration of 100 mg/l and 250 mg/l, respectively, were most efficient for disease con­ trol. Treated fruit remained healthy even after 15-day storage, while the non-treated fruit devel­ oped lesions on 60% of the peel (Sponholz et al., 2004). Attempts have been made to find control measures that do not require the application of manufactured chemicals. In Brazil, bananas sprayed with different concentrations of conidia were immersed in water at different tempera­ tures for different time periods to determine opti­ mum conditions for the control of anthracnose. After 12 days of storage, the disease incidence was only one fruit per bunch treated for 20 min at 45°C, while treatment for shorter time periods



Fungal Diseases of Banana Fruit

were inefficient. The lesioned area was reduced to 15% when fruit was treated for 20 min at 50°C, but to 3% or 0% if treated for 15 or 20 min at 53°C, respectively, compared with 53% in the non-treated fruits (Sponholz et al., 2004). In India, various plant extracts were tested in vitro and in vivo. Fruit of ‘Robusta’ (AAA, Cav­ endish subgroup), ‘Rasthali’ (AAB, Silk sub­ group) and ‘Ney Poovan’ (AB) was treated and kept at a room temperature of 28 ± 2°C and in cool store at 13.5°C. Solanum torvum extract was found to be better than a standard treatment with the fungicide benomyl (0.1%). It was also the best extract tested for increasing the green life and shelf-life of bananas, which was extend­ ed by 16–20 days over the control (Thangavelu et al., 2004). Other workers in India, China, Bra­ zil and elsewhere in recent years have reported success in the control of anthracnose with vari­ ous other plant extracts, antagonistic organisms and essential oils, but there is no information if any have been adopted by commercial growers. In vivo tests using bananas artificially inoc­ ulated with C. musae revealed that coating ba­ nanas with 10% Arabic gum incorporating 1.0% of the biofungicide chitosan was the opti­ mal concentration for controlling anthracnose (80%). It also significantly delayed ripening (Maqbool et al., 2010). Anthracnose was com­ pletely inhibited on bananas artificially inocu­ lated with C. musae and then irradiated with UV-C light (wavelength 254 nm) for 2 h (Bokhari et al., 2013). In the French West Indies, covering bunch­ es with a plastic sleeve had no effect on the quantity of Colletotrichum colonies isolated from flower parts from flowering to harvest. However, the number of anthracnose lesions developing on ripe fruit was considerably less important on sleeved than on unsleeved bunches (de Lapeyre de Bellaire and Mourichon, 1998).

285

clusters of the carton. The spots increase in size during ripening. The problem is initiated by Colletotrichum musae, the cause of anthracnose, but is followed by Fusarium incarnatum, which causes a second­ ary infection. Fungicides used to protect banan­ as against crown rot also control fungal scald.

Stem-end Rot If the pedicel or stem of green banana fruit is in­ jured, a rot similar in appearance to crown rot often develops (Plates 4.21 and 4.22). Stem-end rot is the name given to the decay that advances from the cut end when individual fingers are cut from the bunch and marketed as single fruits, rather than in clusters or hands. The practice

Plate 4.21.  Symptoms of stem-end rot caused by Colletotrichum musae. Salmon-pink-coloured fruiting bodies of the fungus can be seen near the tip of the diseased Cavendish finger (photo: QDPI).

Fungal Scald This disease, which superficially resembles an­ thracnose, can occur in fruit in controlled at­ mosphere packs in transit for more than 14 days. Fungal scald develops where the tips of fruit touch polyethylene liners in the presence of moisture. Reddish-brown sunken spots form on green fruit near the finger tips and on the bottom

Plate 4.22.  Symptoms of stem-end rot caused by Thielaviopsis paradoxa. The white mycelium of this fungus can be seen at the pedicel end of the Cavendish finger (photo: QDPI).

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was once common in Australia and stem-end rot, or ‘black-end’, was a significant problem. When bananas were shipped as whole bunches in the international trade, the pedicel was often creased and bruised when the fingers flexed on the bunch. The ensuing decay, described by Stover (1972), was called Santa Marta stem-end rot and was important because individual fingers often fell from the bunch during transit and han­ dling. Colletotrichum musae (Plate 4.21) was gen­ erally the cause of decay, but other crown rot fungi, such as Thielaviopsis paradoxa (syn. Ceratocystis paradoxa) (Plate 4.22), were also involved. These diseases are not important now, because handling procedures have changed. The princi­ ples and procedures described above for crown rot apply equally to stem-end rot.

Main-stalk Rot Main-stalk rot refers to the decay of the pedun­ cle that occurred when bananas were exported on the bunch. Spreading from the proximal end, the rot had a characteristic sweetish odour and often engulfed the lower hands, causing finger drop (Stover, 1972). The main cause was the fungus Thielaviopsis paradoxa (syn. Ceratocystsis paradoxa), but the fungi Lasiodiplodia theobromae and Colletotrichum musae were also implicated. Although mainly of historical interest, the dis­ ease has recently emerged as a threat to a new method for storing green bananas on the bunch in sealed polyethylene tubes without refrigera­ tion (Satyan et al., 1992). Under modified atmos­ pheres, Fusarium moniliforme var. subglutinans, which is an organism that has since been reclas­ sified into at least 11 Fusarium species including F. musae (P.Y. Tan, Australia, 2017, personal communication), was a major cause of this new stalk rot (Wade et al., 1993).

Botryodiplodia Finger Rot Stover (1972) reported that Botryodiplodia finger rot was one of the most common rots of bananas that were in transit in boxes for more than 14 days. However, it rarely occurs in fruit transport­ ed for 10 days or less. The disease has been re­ ported on fruit from Central and South America,

­ hilippines the Caribbean, India, Taiwan and the P (Slabaugh, 1994a). Botryodiplodia finger rot is a soft rot that ad­ vances from the tip of the fruit below the flower remnants. The pulp is converted into a black mass and the entire fruit can decay. The skin is black and wrinkled, ripens prematurely and be­ comes encrusted with pycnidia. Grey to black woolly mycelium on the surface of the fruit under conditions of high humidity is diagnostic (Stover, 1972). Fully mature fruit is more susceptible to infection and affected clusters ripen earlier. The disease develops faster during ripening and can spread to adjacent fingers (Slabaugh, 1994a). The disease is caused by Lasiodiplodia theobromae (syn. Botryodiplodia theobromae), which is a common inhabitant of decaying banana vege­ tation in the banana plantation. The fungus is easily cultured. Pycnidia are black, flask-shaped with short necks and 250–300 μm in diameter and contain short conidiophores accompanied by paraphyses. Conidia are exuded as single hyaline cells, which mature to two-celled brown, uncon­ stricted, longitudinally striate spores measuring 15 × 25 μm. Wind and water disseminate the co­ nidia. The fruit is infected at the flower end of fin­ gers and through wounds. Control measures used for crown rot are recommended. The wash water should be clean, the fruit should have sufficient green life to reach its destination without ripening, temperature and humidity should be controlled and fungi­ cides may be used (Stover, 1972).

Squirter Squirter, like stem-end rot, is a disease of fingers when they are packed singly. Hands and clusters are not usually affected and the problem is now of little economic importance. It occurred mainly in Queensland and New South Wales in ­Australia, but was reported on fruit in Czecho­ slovakia imported from Guinea (Jechová, 1963). The pathogen enters through the cut pedi­ cel and advances into the fruit as it ripens. The discoloured pulp (Plate 4.23) becomes a liquid mass and may squirt from the end of a fruit if squeezed. External symptoms become visible as a bluish-tan discoloration of the peel as the fruit ripens (Stover, 1972).



Fungal Diseases of Banana Fruit

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in the USDA Fungal database (Farr and Rossman, 2017, reported in Wang et al., 2017). It is unclear if these are saprophytes, endophytes or weak pathogens.

Other Diseases

Plate 4.23.  Internal symptoms of squirter caused by Nigrospora musae (photo: QDPI).

In Australia, the cause was attributed to Nigrospora musae (McLennan and Hoëtte, 1933). This fungus is morphologically almost identical to N. sphaerica. Conidia of N. musae have dimen­ sions of (15–)16–18(–19.5 μm) and N. sphaerica of 16–19(–21) μm (Wang et al., 2017). As such, it was placed in synonymy with N. sphaerica under N. maydis by Jechová (1963) (see Stover, 1972). However, Wang et al. (2017), who studied the phylogenetic relationships between isolates of Nigrospora, using molecular methods, recognized N. musae and N. sphaerica as separate species. Hyphae of N. musae are pale brown, smooth, branched, septate and 2–6 μm in diameter. Co­ nidiophores are aggregated in black sporodochia and are micronematous or semi-­macronematous, flexuous or straight, pale brown, smooth, much branched and 3.5–8 μm in diameter; some ­conidiophores reduced to conidiogenous cells. Conidiogenous cells are aggregated, pale brown, monoblastic, subglobose to ampulliform and measure 6.5–14 × 6–9 μm (average 9.16 ± 1.49 × 7.45 ± 0.74). Hyaline vesicles delimit the conid­ ia from conidiogenous cells. Conidia are sparse, solitary, globose or subglobose, black, shiny and smooth (Wang et al., 2017). Conidia form on dead vegetation, including banana trash. Airborne conidia are common in banana plantations (Meredith, 1961e) and in wash water in packing stations. Like stem-end rot, dipping fruit in a solution of fungicide easily controls the problem (Stover, 1972). Wang et al. (2017) worked with six Nigrospora species isolated from banana leaves in China. These were N. camilliae-sinensis, N. hainanensis, N. lacticolonia, N. pyriformis, N. sphaerica and N. vesicularis. More have been reported from banana

Ring rot has been reported in single bananas from Australia (Jones and Wade, 1995). A dark, water-soaked, 20–40 mm diameter ring devel­ ops on the peel as the fruit yellows during ripen­ ing (Plate 4.24). Internally, a dark, watery rot extends along the centre of the fruit pulp. The symptoms may be confused with squirter (see above). The disease is caused by a phycomycete fungus, which appears to infect from zoospores in the soil or wash tank. Ring rot is more often found after prolonged wet, windy weather. It can be avoided by rejecting fruit from bunches which have been in contact with the ground and by ­using clean wash water.

Plate 4.24.  Symptoms of ring rot caused by an unknown phycomycete in Australia (photo: QDPI).

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Trachysphaera finger rot, described by Stover (1972), is a rare disease of ‘Gros Michel’ fruit from Cameroon. ‘Lacatan’ fruit from Jamaica ripened in rooms in England after fruit from Cameroon have also been affected. The cause, Trachysphaera fructigena, which can also cause the preharvest disease known as cigar-end rot, is an inhabitant of the floral end of African bananas and may invade through wounds in the green peel, causing a dry black rot, which becomes ­fibrous. Banana fruits exhibiting signs of decay were collected from markets in the USA and

I­taly. Fungi isolated from the lesions on the ­banana fruit were Fusarium verticillioides (syn. Fusarium moniliforme), F. subglutinans and F. incarnatum (syn. F. pallidoroseum var. majus and F. semitectum var. majus) (Vesonder et al., 1995). Soft rot on harvested banana fruit caused by Rhizopus stolonifera and R. oryzae have been reported in Korea (Kwon, 2007; Kwon et al., 2012). Infection usually started from cracks in the peel and occurred at harvest time. The disease was not serious on fruit in the pre-climatic stage while still green and hard, but losses were considerable when bananas ripened.

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Tomkins, R.G. (1931) Wastage in banana transport. Tropical Agriculture (Trinidad), 8, 225–264. Triest, D. and Hendrickx, M. (2016) Postharvest disease of banana caused by Fusarium musae: a public health concern? PLoS Pathogens 12(11), e1005940. Van Hove. F., Waalwijk, C., Logrieco, A., Munaut, F. and Moretti, A. (2011) Gibberella musae (Fusarium musae) sp. nov., a recently discovered species from banana is sister to F. verticillioides. Mycologia 103, 570–585. Vawdrey, L.L., Langdon, P. and Westerhuis, D. (2010) Aetiology, effect of chemicals, and influence of fruit exudates, insects and fruit maturity on the incidence of fruit speckle of banana. Australasian Plant Pathology 39(6), 524–529. Vesonder, R.F., Logrieco, A., Bottalico, A., Altomare, C. and Peterson, S.W. (1995) Fusarium species associated with banana fruit rot and their potential toxigenicity. Mycotoxin Research. 11(2), 93–98. von Arx, J.S. (1957) Die arten der gaitung Colletotrichum. Phytopathologische Zeitschrift 29, 413–468. Wade, N.L., Kavanagh, E.E. and Sepiah, M. (1993) Effects of modified atmosphere storage on banana postharvest diseases and control of bunch main-stalk rot. Postharvest Biology and Technology 3, 143–154. Wallbridge, A. (1981) Fungi associated with crown rot disease from the Windward Islands during a two-year survey. Transactions of the British Mycological Society 77, 567–577. Wallbridge, A. and Pinegar, J.A. (1975) Fungi associated with crown rot disease of bananas from St Lucia and the Windward Islands. Transactions of the British Mycological Society 64, 247–254. Wang, B., Liu, C. and Qi, P. (1991) Studies on the Fusarium crown rot of bananas. Acta Phytophylactica Sinica 18, 133–137. Wang, M., Liu, F., Crous, P.W. and Cai, L. (2017) Phylogenetic reassessment of Nigrospora: ubiquitous endophytes, plant and human pathogens. Persoonia 39, 118–142. Wardlaw, C.W. (1931) Banana diseases. 3. Notes on the parasitism of Gloeosporium musarum Cke. and Mass. Tropical Agriculture (Trinidad) 8, 12. Wardlaw, C.W. (1934) Banana diseases. 6. The nature and occurrence of pitting disease and fruit spots. Tropical Agriculture (Trinidad) 11, 8–13. Wardlaw, C.W. (1961) Banana Diseases Including Plantains and Abaca. Longmans, Green, London, 648 pp. Wardlaw, C.W. and McGuire, L.P. (1932) Pitting disease of bananas. Its nature and control. Tropical Agriculture (Trinidad) 9, 193–195. Waterston, J.M. (1947) The fungi of Bermuda. Bulletin of the Department of Agriculture of Bermuda 23, 1–305. Yang, H.R. and Chuang, T.Y. (1996) The nutrient effect on the appressorial behaviour and latent infection of Colletotrichum musae. Plant Protection Bulletin (Taiwan) 3, 247–259. Zakaria, L., Sahak, S., Zakaria, M. and Salleh, B (2009) Characterisation of Colletotrichum species associated with anthracnose of banana. Tropical Life Sciences Research 20, 119–125. Zare, R., Gams W., Starink-Willemse, M., Summerbell, R.C. (2007) Gibellulopsis, a suitable genus for Verticillium nigrescens, and Musicillium, a new genus for V. theobromae. Nova Hedwigia 85, 463–489. Zhou, Y., Huang, J.S., Yang, L.Y. and Wang, G.F. (2017) First report of banana anthracnose. caused by Colletotrichum scovillei. Plant Disease 101, 381.

5 

Diseases Caused by Bacteria and Phytoplasmas

Bacterial Wilt Diseases Xanthomonas Bacterial Wilt G. Blomme and W. Ocimati Introduction Xanthomonas bacterial wilt is currently the most important threat to enset and banana ­production in the East and Central African region. Symptoms were first observed on enset in ­ Ethiopia in the 1930s (Castellani, 1939). However, the bacterial causal organism was not identified and named until 1968 (Yirgou and Bradbury, 1968). It was later found infecting banana plants in Ethiopia in 1974 (Yirgou and Bradbury, 1974). The disease was not identified outside Ethiopia until 2001 when banana in central Uganda (Tushemereirwe et al., 2003) and the eastern Democratic Republic of the Congo (Ndungo et al., 2006) was found to be affected. It has since spread to Rwanda (Reeder et al., 2007), Tanzania (Carter et al., 2010), Kenya (Mbaka et al., 2007; Carter et al., 2010) and Burundi (Carter et al., 2010). Xanthomonas bacterial wilt is now widely distributed in all growing districts in Ethiopia (Yirgou and Bradbury, ­ 1974; Addis et al., 2004), Uganda, Rwanda and ­Burundi and is present in the Kagera region of Tanzania and in western Kenya. In eastern ­Democratic Republic of the Congo, the disease is 296

s­ preading westwards into the Congo basin and southwards towards Katanga province. Xanthomonas bacterial wilt affects all enset genotypes in Ethiopia and edible banana cultivars in the East and Central African region where it occurs (Yirgou and Bradbury, 1974; Eden-Green, 2004; Ssekiwoko et al., 2006b; Addis et al., 2010). There is no yield from infected banana plants (Ssekiwoko et al., 2006b). In the case of enset, varying degrees of tolerance have been observed for a limited set of cultivars. In banana, the fruit prematurely ripens and rots, and/or the whole plant wilts and dies (Smith et al., 2008). Losses on farms can reach up to 100% when control is delayed, especially when ABB genotypes are c­ultivated. This severely compromises food security and livelihoods of affected households (Tushemereirwe et al., 2003, 2004, 2006; Kagezi et al., 2006; Kalyebara et al., 2006, 2007; Karamura et al., 2006; Ssekiwoko et al., 2006b; Blomme et al., 2014). Economic losses have been estimated to be US$200–295 million/year in Uganda (Kalyebara et al., 2006; Abele and Pillay, 2007). The disease spreads quickly by insect transmission and the cultivation of ABB cultivars in central Uganda has often been severely affected by the disease. However, the East African highland banana cultivars (AAA, Lujugira–Mutika subgroup) in market-oriented production systems in southwestern Uganda have had more modest infection levels, often limited to ‘hotspot’ sites. As a consequence, economic losses to ­banana farmers in Uganda have been lower than

© D.R. Jones 2019. Handbook of Diseases of Banana, Abacá and Enset (D.R. Jones)



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initial estimates. A more recent study, which assessed the impact of Xanthomonas bacterial wilt on small-scale farmers’ livelihoods in the Kagera basin of Tanzania, Burundi and Rwanda, reported that 33–75% of banana mats per farm were affected by the disease (Nkuba et al., 2015). The same study reported annual banana production losses valued at US$10.2 million and US$2.95 million in Tanzania and Rwanda, respectively, a reduction in fruit sales of 35% and a doubling of bunch prices. Here, there has been a significant reduction in household food security and ­income. Banana is an important staple food and cash crop in East and Central African countries. It is important for the regional economy with an annual production of about 27 million tonnes of fruit worth US$7.3 billion and forms about 10.3% of the net domestic product (FAO, 2016). Bananas are an indispensable part of life in the region. They provide 3–22% of the total calorie consumption per capita. The average regional daily calorie intake per person from banana over the past five decades has been estimated at 147 kcal. This is 15-fold the global average and sixfold the African average. If unchecked, Xanthomonas bacterial wilt will continue to seriously compromise regional food security and livelihoods associated with banana cultivation and marketing. To cope, most affected households are diversifying into other food crops, such as maize, cassava, taro and sweet potatoes, potentially posing a number of socio-­economic and biological implications that require further investigation (Nkuba et al., 2015; W. Ocimati, Uganda, 2016, personal communication). Severe effects of the disease on the ecosystem health of banana-based agroecosystems have been postulated and studies to quantify these are in progress (W. Ocimati, Uganda, 2016, personal communication). The current impact of the disease to enset farmers seems limited to ‘hotspot’ sites and is in general relatively low. However, enset is harvested 3–5 years after planting and losing only one or two 5-year-old plants is significant to a poor farmer. Symptoms Symptoms on banana plants are typically evident within 2–22 weeks of infection. The severity

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and time to symptom expression and disease progress varies with cultivar, plant growth stage, environmental conditions and point of infection (Ssekiwoko et al., 2006a, b; Addis et al., 2010; Ocimati et al., 2013d). The incubation period has been observed to be generally shorter in East African highland cultivars than in ‘Pisang Awak’ (ABB) (Ocimati et al., 2013d). In general, disease progression is faster in young plants compared with mature plants, and during the wet season compared with the dry season (Mwangi et al., 2007; Tripathi et al., 2008, 2009). The bacterium that causes Xanthomonas bacterial wilt infects banana and enset either through the vegetative parts of the plants (roots, corm, pseudostem comprising leaf sheaths, leaf petioles and leaf lamina) or, in the case of banana, through the inflorescence. Shorter incubation periods have been reported for infections through floral parts compared with those through the vegetative parts in banana (Ocimati et al., 2013c, d). Ocimati et al. (2013d) reported long incubation periods varying between 11 and 110 weeks in attached banana suckers following infections in mother plants. Latent infections are common in banana, with incidences of up to 33% and 53% reported in the East African highland cultivars and ‘Pisang Awak’, respectively. The visible symptoms of floral infections in banana include wilting of male bud bracts (Plate 5.1) followed by drying of the rachis coupled with bacterial exudation (Plate 5.2), often followed by premature ripening of some or all of the fruit (Plate 5.3) and eventually wilting and death of the entire plant (Plate 5.4) (Ssekiwoko et al., 2010; Ocimati et al., 2013b, c, d). Internal cross-sections of the floral stalks show yellow bacterial ooze from the vascular bundles (Plate 5.5) while cross-sections of the fruit show rusty brown stains (Plate 5.6) (Thwaites et al., 1999b; Tushemereirwe et al., 2003; Biruma et al., 2007; Ocimati et al., 2013c, d). Infections through the corm, roots, pseudostem and leaves of banana plants will show a progressive yellowing and wilting of the leaves (Plates 5.7 and 5.8) (Tushemereirwe et al., 2004; Ssekiwoko et al., 2006a; Ocimati et al., 2013b, c, d; Nakato et al., 2014). A cream-­coloured or yellow ooze, typical of many bacterial infections, exudes within a few minutes of cutting tissue (Plate 5.9) and copious quantities may be produced over a period of several hours. Affected pseudostems

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Plate 5.1.  Wilting of male bud bracts of a banana is an initial symptom of Xanthomonas bacterial wilt infection via the inflorescence. The infection can be seen progressing along the rachis towards the fruit (photo: G. Blomme, BI).

Plate 5.3.  Premature ripening and degeneration of fruit of a banana with Xanthomonas bacterial wilt. The disease has progressed from the withered male bud bracts and along the rachis to the fruit (photo: G. Blomme, BI).

Plate 5.2.  Drying and rotting of the rachis of a banana affected by Xanthomonas bacterial wilt. Yellow-coloured bacterial exudations can also be seen on the rachis (photo: G. Blomme, BI)

often wilt and die. Late ­floral symptoms, especially wilting of male bud bracts and drying of the rachis, have been reported in banana for ­infections through the corm, root, pseudostem and leaves of plants ­inoculated at flowering (Ocimati et al., 2013d; Nakato et al., 2014). In most cases, fruits of these plants remain edible though they asymptomatically carry the bacteria. Foliar symptoms of yellowing and wilting of leaves (often associated with loss of turgor as if the leaves had been affected by fire) are often confused with the symptoms of Fusarium wilt, but the excretion of a yellowish bacterial ooze from cut tissues is characteristic of Xanthomonas bacterial wilt (Thwaites et al., 1999b; Tushemereirwe et al., 2003, 2004, 2006). Another difference is that Fusarium wilt causes a far brighter yellow coloration on the leaf lamina while typical wilting due to Xanthomonas bacterial wilt results in a more dull yellow colour, as if the leaf has been burnt by fire (Plate 5.7) (Karamura et al., 2008). In addition, the yellowing and wilting of the leaves in plants affected



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Plate 5.6.  Cross-sections of banana fruit affected by Xanthomonas bacterial wilt shows the pulp stained by bacteria (photo: G. Blomme, BI).

Plate 5.4.  Wilting and drying of leaves of banana plants affected by Xanthomonas bacterial wilt eventually leads to plant death (photo: G. Blomme, BI).

hang down the pseudostem (see Plate 3.4). In  plants affected by Xanthomonas bacterial wilt, the wilting can begin with any leaf and the ­infected leaves tend to snap along the leaf blade (Plate  5.8). The flower and fruit symptoms are similar to those observed for Moko bacterial wilt/bugtok and blood bacterial wilt, but these diseases of ­banana are not known to occur in Africa. Symptoms of Xanthomonas wilt in enset, which have been reported from all growing districts in Ethiopia, are similar to those in banana. Symptomatic leaves lose turgor and wilt (Plate 5.10). Internally, vascular bundles become discoloured, though this symptom is not as conspicuous as the internal discoloration observed in banana. Pockets of yellow or cream-­coloured bacterial matter/slimy secretion may develop within the pseudostem, leaf petioles and midribs (Plate 5.11). Total yield loss most often occurs once the disease takes hold.

Causal agent

Plate 5.5.  Internal cross-section of a banana floral stalk of a banana with Xanthomonas bacterial wilt shows yellow bacterial ooze emanating from the vascular bundles (photo: G. Blomme, BI).

by Fusarium wilt typically progresses from the older to the younger leaves (see Plate 3.2). The wilted leaves may also snap at the petiole and

Yirgou and Bradbury (1968, 1974) described the pathogen as Xanthomonas musacearum sp. n. Young et al. (1978) proposed the name X. campestris pv. musacearum (Xcm) for the pathogen, which was accepted by taxonomic bacteriologists. More recently, Aritua et al. (2008) observed that the bacterium was closely related to X. vasicola species and thus proposed the name X. vasicola pv. musacearum, but this suggested

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Plate 5.7.  Yellowing leaves after tool-mediated infection. Xanthomonas bacterial wilt results in a dull yellow colour as if the leaf has been burnt by fire (photo: G. Blomme, BI).

name change has not yet been approved. Xanthomonas vasicola includes causative agents of several economically important diseases, such as pathovars X. vasicola pv. holcicola pathogenic to sorghum (Sorghum bicolor) and X. vasicola pv. vasculorum pathogenic to sugarcane (Saccharum officinarum) and maize (Zea mays) (Vauterin et al., 1995). Xcm belongs to a group of bacteria that is found only in association with plants or plant materials (Thwaites et al., 1999b). The genus Xanthomonas falls within the gamma-­ proteobacteria that consist of over 420 species and hundreds of pathovars of economically important plant pathogenic bacteria (Vauterin et  al., 1995). Xcm is a motile, Gram-negative, rod-shaped ­bacterium possessing a single polar ­flagellum (Vauterin et  al., 1995). It has the following characteristics: it is oxidase- and ­ tyrosinase-­negative, does not reduce nitrate or hydrolyse starch or gelatin and does not accumulate poly-β-­hydroxybutyric acid (­Yirgou and Bradbury, 1968, 1974). It is non-fluorescent on King’s B medium and produces typical yellow, convex, mucoid colonies on nutrient agar and other media. In media or environments rich in glucose, Xcm produces copious amounts of extracellular polysaccharide, called xanthan ­ gum. While in the plant Xcm infects and

c­olonizes the xylem, d ­amaging tissues and ­depositing ­copious amounts of xanthan gum, which then prevent absorption and translocation of water (Tripathi et al., 2008). Like other xanthomonads, Xcm has been shown to grow more slowly than other bacterial species, such as those in the genera Pseudomonas, Burkholderia and Ralstonia (L. Tripathi, Uganda, 2016, personal communication). This affects its ability to compete with other bacteria outside its host. Initial phylogenetic analysis of partial ­nucleotide sequences of the gyrase B gene and internal transcribed spacer (ITS) region, and ­ genomic amplicon fingerprints using repetitive sequence PCR and fatty acid methyl esters, showed a high homogeneity between all isolates of Xcm in the East and Central African region (Aritua et al., 2007, 2008, 2009; Odipio et al., 2009). However, through genome-wide sequencing, Wasukira et al. (2012) observed a set of single-­­nucleotide polymorphisms among Xcm isolates. They revealed two major sub-lineages of Xcm, suggesting that the outbreaks of Xanthomonas bacterial wilt on Musa spp. in the East and Central African region could have had several introductory events from Ethiopia. Wasukira et al. (2012) specifically observed that the Xcm isolates from eastern Democratic Republic of the



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Plate 5.8.  Collapse of leaves of a banana after the Xanthomonas bacterial wilt pathogen has been introduced on contaminated tools (photo: G. Blomme, BI).

Disease cycle and epidemiology Infection

Plate 5.9.  Bacterial ooze emanating from a cut pseudostem of a banana with Xanthomonas bacterial wilt (photo: G. Blomme, BI).

Congo, Rwanda and Ethiopia clustered together and away from those from ­ Burundi, Kenya, ­Tanzania and Uganda by 86 single nucleotide polymorphisms.

Infection occurs when the bacterium enters the vascular system of the plant through natural or tool-mediated wounds. Banana plant residues, contaminated soils and water, infected plants and traded products, including fruits, leaves and planting materials, have all been confirmed as potential sources of Xcm inoculum (Eden-Green, 2004; Karamura et al., 2008; Nakato et al., 2014). Alternative Xcm hosts are also a potential source of disease inoculum. Screenhouse inoculation of Canna indica, a common weed in the banana-­ growing regions of East and Central Africa, has resulted in characteristic wilting symptoms (Ssekiwoko et al., 2006a; W. Ocimati, Uganda, 2016, personal communication). Xcm has also been isolated from, or observed to cause pathogenic reactions in, non-host crops such as Zea mays

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Plate 5.10.  The leaves of enset affected by Xanthomonas bacterial wilt lose turgor and wilt (Photo G. Blomme, BI).

(Aritua et al., 2008; Karamura, 2012; Rutikanga et al., 2016a), Saccharum spp., Sorghum bicolor and Eleusine coracana (W. Ocimati, Uganda, 2016, personal communication). There are ongoing studies to determine the potential of Canna indica and selected annual and perennial cultivated/ wild crops to act as alternate hosts and perpetuate Xanthomonas bacterial wilt on farms. After infection, the bacteria spread systemically within the initially infected banana plant and often to the other attached stems/suckers in the mat (Ssekiwoko et al., 2010; Ocimati et al., 2013c, 2015). It was initially thought that all the physically interconnected stems in a mat get infected when one stem is infected. However, Xcm has been reported as only partially systemic in banana mats. Only 8–25% of the asymptomatic suckers were reported to be latently infected in controlled experiments in which mother plants were florally inoculated (Ocimati et al., 2015). Similarly, low Xcm incidence (3%) was observed in symptomless suckers sourced from

heavily infected farmers’ fields where plants were mainly infected through contaminated tools (Ocimati et al., 2015). Similar local infections have been reported in plants affected by Moko bacterial wilt and blood bacterial wilt (EdenGreen, 1994). Research on the effect of soil fertility level on plant response to Xcm has been inconclusive. While the application of potassium, calcium and nitrogen to in vitro banana plantlets resulted in significantly reduced susceptibility to Xcm (Atim et al., 2013), the application of nitrogen, phosphorus and potassium in controlled pot experiments did not (Ochola et al., 2014a). The observed differences could be attributed to the interaction of physical and chemical factors in the pot trials, factors that were lacking in the in vitro studies. Field studies are still needed to further elucidate the interaction between soil nutrients and the disease. Higher Xanthomonas bacterial wilt incidence has been reported on farmers’ fields in the



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modes of spread of Xanthomonas bacterial wilt are via insect vectors (Yirgou and Bradbury, 1974; Tinzaara et al., 2006; Shimelash et al., 2008; Addis et al., 2010; Were et al., 2015), contaminated garden tools (Yirgou and Bradbury, 1974; Addis et al., 2010; Ocimati et al., 2013c) and infected planting material (Eden-Green, 2004; Biruma et al., 2007). Infections through soil, though limited, have also been reported (Karamura et al., 2008; Shehabu et al., 2010; Ocimati et al., 2013c). Insect vector transmission

Plate 5.11.  Pockets of yellow or cream-coloured bacterial ooze develop within the leaf sheaths of enset plants with Xanthomonas bacterial wilt (photo: M. Tessara and O.J. Quimio, IAR)

rainy season (Biruma et al., 2007; Blomme et al., 2007). In controlled water stress pot trials, a mild stress of 2 weeks was observed to be beneficial for the plants while extreme stress resulted in a shortened incubation period and exacerbated disease incidence and severity compared with the control treatments under routine watering (Ochola et al., 2014b). These observations suggest that extreme weather conditions could exacerbate the problem, with high soil moisture conditions potentially promoting the multiplication of the pathogens while water stress conditions weaken the ability of the plant to resist the disease. Spread Xcm is easily transmitted by any object/tool or organism that comes in contact with the sap of contaminated plant parts (Brandt et al., 1997). The contribution of these sources of infection to the spread of the disease depends on the survival of the bacterium and its mode of transmission (Eden-Green, 2004; Agrios, 2005). As in other vascular bacterial diseases of banana (EdenGreen, 1994; Buddenhagen, 2006), the main

Insect vectors play an important role in both local and distant spread of Xcm. The most common insects associated with banana inflorescences are stingless bees, honeybees, fruit flies and grass flies (Tinzaara et al., 2006; Shimelash et al., 2008; Rutikanga et al., 2015). Insect transmission occurs mainly, if not exclusively, from male buds of diseased plants to those of healthy plants through freshly exposed wounds left by flowers and bracts that, in many cultivars, are shed daily. Although there is overwhelming circumstantial evidence that insects spread Xanthomonas bacterial wilt of banana, it is still not known if bacteria can be acquired and/or transmitted through contact with intact male flowers. Buddenhagen and Elsasser (1962) argued that the abscission sites rather than the male flowers are the main avenues for both acquisition and transmission of bacteria. If this is correct, then studies of insects that regularly visit the abscission sites may shed some light on the main vector species. A recent study by Were et al. (2015) confirmed the potential role of banana weevils in local disease-spread within a plot or plantation. Xcm was isolated on both the external and internal parts and faecal matter of weevils trapped from infected fields and subsequently deliberately fed on Xcm oozing corms. Infection also occurred in pot trials when these weevils were introduced to potted plants. Transmission via insect vectors largely depends on the population size of a specific vector, their mobility and the size of individuals (Shimelash et al., 2008; Blomme et al., 2014; Rutikanga et al., 2015). For example, fruit flies, which are small and are present in large numbers, spend most of their lives on a few mats and will thus not spread the disease over large areas (Smith

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et al., 2008). Larger insects (e.g. bees and wasps) can visit wider geographical areas and more flowering banana plants daily, thus could potentially spread the disease more widely (Smith et al., 2008). In contrast, banana weevils, though large in size, rarely fly (Gold et al., 1999) or move ­beyond 25 m (Gold and Messiaen, 2000), thus limiting their potential role in disease spread to within a field and neighbouring plots (Were et al., 2015). Insect vector transmission is also influenced by altitude, rainfall and the cultivars grown. Lesser insect vector activity and fewer floral infections have been reported at altitudes above 1700 m compared with lower altitudes (Addis et  al., 2004; Blomme et al., 2005; Shimelash et  al., 2008; Rutikanga et al., 2015). Lower insect activity has also been reported during seasons of intense dry spells and intense rainfall (Rutikanga et al., 2015). Higher insect-mediated floral infections have been reported in banana-growing areas in East Africa and Ethiopia dominated by ‘Pisang Awak’ cultivation (Addis et al., 2004; Ndungu et al., 2004; Blomme et al., 2005; Shimelash et al., 2008). Insects were found to be the main drivers of spread of Xanthomonas bacterial wilt in central Uganda at altitudes of 1050–1400 m, where ‘Pisang Awak’ was predominantly grown during the period 2002–2005 (Tinzaara et al., 2006). Lower disease-spread rates, compared with central Uganda, were observed in the Masisi region of North Kivu province (1700 m in altitude) in eastern Democratic Republic of the Congo and in southwestern Uganda, which are banana-­growing areas both dominated by East African highland banana cultivars. No inflorescence symptoms were observed in the high-altitude (1850 m) Kambata region of southern Ethiopia where ‘Pisang Awak’ is widely grown and de-­budding is not practised. This compared to the sporadic presence of floral infections in the Kaffa region of western Ethiopia (with a slightly lower elevation of 1600 m) where there are also large numbers of ‘Pisang Awak’. A study by Rutikanga et al. (2016b) in Rwanda ruled out any preferential insect attraction to ‘Pisang Awak’ based on nectar volume and sugar concentration. The high incidence in ‘Pisang Awak’ is attributed to non-persistent neuter and male flowers with bracts that fall off leaving fresh wounds prone to colonization by

Xcm (Mudonyi, 2014). Cultivars with persistent flowers/bracts, such as ‘Dwarf Cavendish’ (AAA, Cavendish subgroup), and East African highland banana cultivars belonging to the ‘Nakitembe’ clone set with persistent or semi-persistent bracts, have been reported to escape infection often, even under high disease pressure (Addis et al., 2004; Shimelash et al., 2008; Mudonyi, 2014). However, it is important to stress that these cultivars are not resistant per se, but only escape the disease because of their persistent or semi-­ persistent inflorescences. Enset is harvested before the inflorescence appears. In addition, most, if not all, enset cultivars have persistent bracts and flowers, thus excluding insect-vectored inflorescence transmission in farmers’ fields. Mechanical transmission on farm tools Xanthomonas bacterial wilt has been shown to spread as the result of the use of contaminated farm tools during weeding, de-suckering, transplanting of suckers, leaf pruning and harvesting operations (Addis et al., 2004; Ssekiwoko et al., 2010; Ocimati et al., 2013b, c, d). Contaminated garden tools have been shown to transmit the disease up to 1 week after contact with diseased tissue/bacterial ooze (Buregyeya et al., 2008). Garden tool transmission is the primary mode of disease-spread in high-altitude zones with less insect activity and in intensively managed farms that often de-bud timely to prevent insect-­ mediated infections. Tool transmission by banana traders, who often travel from infected to non-­infected zones in search of banana bunches and leaves for subsequent sale, has also been reported (Nakato et al., 2013). Spread as a result of movement of planting materials Resource poor and subsistence farmers in East and Central Africa depend entirely on suckers obtained from their own or neighbouring fields when planting (Ocimati et al., 2013a). Even if the sucker appears free of disease, it could be infected. Therefore, there is a strong possibility of spreading the disease from farm to farm. Local spread of the disease through planting materials is believed to be common and there is also the potential for long-distance spread.



Diseases Caused by Bacteria and Phytoplasmas

Soilborne inoculum Soilborne infections are generally limited. Xcm penetration through wounds resulting from root damage caused by nematodes and tools has, however, been demonstrated for banana (Shehabu et al., 2010; Ocimati et al., 2013c). Plant death has also been reported in soils contaminated with Xcm ooze (Karamura et al., 2008). Soil inoculum could potentially build up through bacteria that exude from the cut surfaces of diseased plants, such as open wounds resulting from de-suckering of contaminated plants and corm pieces, especially following uprooting of entire mats. This ‘pool’ of bacteria could potentially spread by water or insects and infect plants through wounds resulting from mechanical injuries caused during weeding and animal grazing, or injuries caused by soilborne organisms such as nematodes and insects such as the banana weevil. However, Xcm cannot survive for long in the soil in the absence of living banana tissue (Mwangi et al., 2007). In studies conducted by Mwebaze et al. (2006), the bacterium did not survive beyond 35 days in plant debris left on the ground surface or buried under the ground, suggesting that it lacks a saprophytic phase. Although some bacterial pathogens such as Erwinia amylovora, some pseudomonads and soft rot Erwinia species, are reported to be disseminated by wind-blown rain, there is no evidence yet for the dissemination of Xcm via aerosols. Dispersal by rain splash has also not been reported, though it is likely that this mechanism can move bacterial ooze from exposed surfaces of infected plants to new locations and potentially infect healthy plants through wounds in the roots or corm tissues. Other modes of spread Isolated disease outbreaks in locations far from known sources of infection suggest the involvement of long-distance vectors. Birds (such as hornbills) and fruit bats foraging for food and/or nectar have long been suspected of transmitting the disease (Buregyeya et al., 2008; Smith et al., 2008). Bech-Andersen (1974) implicated starlings in widespread outbreaks of fireblight disease (Erwinia amylovora) in coastal hawthorn hedges in Denmark, but there are few reports of suspected disease transmission through flying birds.

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In a recent study to determine the potential role of birds and fruit bats in the spread of Xanthomonas bacterial wilt, Buregyeya et al. (2014) reported the eastern grey plantain eater, double toothed barbet, sunbird and village weaverbird as the predominant birds visiting male banana flowers and Aidulon helvum, Epomophorus labiatus and Epomaps franquet as the predominant bats. Xcm was isolated from the mouth parts and other external body parts of these species, confirming their potential role in the spread of the disease. Potential disease spread on banana through free-ranging domesticated ruminants (cattle and goats) has also been suggested (Karamura et al., 2008). Wild animals, such as mole-rats, aardvark and porcupine, that feed on corms are thought to be implicated in disease spread in enset gardens (Brandt et al., 1997; Thwaites et al., 1999b). The level of contribution of animals in Xcm spread is, however, not yet well established. The potential of Xcm spread over long distances through marketed banana products, especially bunches and leaves, has also been demonstrated. In a study by Nakato et al. (2013), at the peak of a Xanthomonas bacterial wilt outbreak, Xcm was isolated from 21–63% of banana bunches randomly sampled across different markets in central Uganda and at the borders with neighbouring countries. In a more recent study, lower incidences varying between 0% and 5% were reported in markets in central Uganda where most sampled bunches came from the highlands of western Uganda, where management of the disease is strict and thoroughly monitored (Ocimati et al., 2015). Host reaction To date, no banana and enset cultivars in the East and Central African region have been reported to be resistant when inoculated with Xcm. The levels of susceptibility have been shown to vary from cultivar to cultivar. ‘Genticha’ and ‘Mezye’ have been regarded as having a good level of tolerance to the disease and have been reported to recover after infection. These cultivars have poor postharvest properties and, as such, are not preferred by farmers. In field trials conducted between 1994 and 2000 to determine the response of 103 enset cultivars from the various growing areas in

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­ thiopia and 12 reported as tolerant to XanthoE monas bacterial wilt, all cultivars developed disease symptoms to various intensity levels during the first 45 days after inoculation with Xcm. The percentage of infected plants per cultivar at each disease assessment period after inoculation varied. Several enset clones showed relative ‘tolerance’ to the disease in that only a low percentage of inoculated plants were diseased after 120 days of observation. Unfortunately, the results of some clones were not consistent. For instance, ‘Ado’ collected from Sidama had 77% of plants affected at 120 days after inoculation in one trial and 22% in another. Similarly, ‘Genticha’ from Sidama had 88% of plants affected after 120 days in one trial and 22% in another. In some instances, a few plants in some clones were seen to be affected earlier in the trial, but had resumed normal growth at 90 days after inoculation. These were deemed to have recovered after infection. All plants of enset clones ‘Astara’, ‘Buffare’, ‘Geziwet 2’, ‘Gulumo’ and ‘Kullo’ showed disease symptoms at 30 days after inoculation and could, hence, be used as susceptible checks in future screening trials (Welde-Michael et al., 2008). Differences in infection rates have also been reported among banana cultivars. This variation, as mentioned earlier, has mainly been attributed to differences in susceptibility of banana cultivars to insect-mediated infections (Ocimati et al., 2013d). ABB cultivars, such as ‘Pisang Awak’ and ‘Bluggoe’, have been shown to be very susceptible to insect-mediated infections because of their non-persistent neuter and male bracts and flowers that leave wounds for the entry of the bacteria (Tushemereirwe et al., 2003, 2004; Addis et al., 2004; Blomme et al., 2005; Biruma et al., 2007). ‘Pisang Awak’ was reported to be responsible for the rapid spread of Xcm in central Uganda during the period 2002–2005 (Tinzaara et al., 2006). A 99% incidence was reported in ‘Pisang Awak’ fields exposed to insect-mediated infections in controlled experiments in central Uganda (Nakato et al., 2014). In contrast, some cultivars, although susceptible, can ‘escape’ insect-mediated infections because they have persistent neuter/ male flowers and male-bud bracts (e.g. ‘Mbwazirume’, an East African highland banana cultivar, and ‘Dwarf Cavendish’). Infection ‘escape’ has also been reported for those cultivars, such as the East African highland banana hybrid ‘M9’,

that have semi-persistent neuter/male flowers and male-bud bracts, which are shed after their points of attachment have dried (Tinzaara et al., 2006; Shimelash et al., 2008; Mudonyi, 2014). Planting cultivars of enset and banana that tolerate or escape the disease in mixtures with more susceptible cultivars could potentially slow down the rate of spread of the disease across farms or landscapes. Most farms growing predominantly ABB cultivars are often poorly managed in contrast to farms specializing in growing East African highland bananas. Plots growing ABB types, such as ‘Pisang Awak’, often look abandoned with plants not de-budded. The low levels of management of the ABB farming systems also significantly contribute to the observed high levels of disease incidence. However, all edible banana cultivars in the East and Central African region are easily infected through the use of contaminated garden tools (Yirgou and Bradbury, 1974; Tushemereirwe et al., 2003; Eden-Green, 2004; Ssekiwoko et al., 2006b; Addis et al., 2010; Mudonyi, 2014).

Control In general, bacterial diseases of plants, once established, are difficult to control owing to the lack of an effective chemical or other curative treatment. Early detection and destruction of diseased plants is a key step in preventing disease spread (Karamura et al., 2005). In the case of Xanthomonas bacterial wilt, the situation is complicated as all edible banana cultivars so far studied are susceptible (Ssekiwoko et al., 2006a, b). Because of these difficulties, a disease-control strategy will be more effective where it combines a number of approaches. Disease-control measures are described in more detail below. Detection and diagnosis The rapid detection and correct diagnosis of Xanthomonas bacterial wilt of banana is important for the timely deployment and appropriate choice of control interventions. The ability to find and identify the disease in a field, landscape or country enables control authorities or farmers to decide if they should opt for quarantine exclusion, eradication or control measures. ­Accuracy is important because of the possibility



Diseases Caused by Bacteria and Phytoplasmas

of confusion with symptoms of abiotic stresses, such as nitrogen or water deficiency, and Fusarium wilt. Various diagnostic techniques have been developed ranging from relatively simple and practical microbiological methods (Mwebaze et al., 2006; Mwangi et al., 2007) to serological diagnostic tests (e.g. Nakato et al., 2013) and more sophisticated molecular-based approaches (Adikini et al., 2011; Adriko et al., 2012). visual diagnosis.  Visual diagnosis of symptoms was critical during the initial stages of the epidemic in East and Central Africa as no other means of identification were available at this time. The presence of yellow bacterial ooze in leaf sheaths, real stem and peduncle, and the discoloration of fruit pulp are unique to Xanthomonas bacterial wilt and clear indications of the presence of the disease. microbiological diagnosis. 

Several media have been developed for the isolation, culturing and identification of Xcm. Yellow, mucoid, smooth, dome-shaped and shiny Xcm colonies are distinctively visible after 3–5 days on non-selective media of yeast peptone glucose and agar media (Mwangi et al., 2007) and the semi-selective yeast tryptone sucrose agar (Tripathi et al., 2007). Lighter-yellow colonies are visible after 5–7 days on the semi-selective cellobiose cephalexin agar media (Mwebaze et al., 2006). Yeast dextrose calcium carbonate agar with properties that buffer against acidification (Dowson, 1957) has been used for the long-term storage of Xcm in the laboratory.

serological diagnostic methods. 

ELISA-based Xcm-specific techniques that can handle large numbers of plant samples have been developed and tested (Nakato et al., 2013). In addition, a monoclonal antibody-based lateral flow device was developed for the rapid diagnosis of Xcm in fields and at border points (Hodgetts et al., 2014; Karamura et al., 2016).

polymerase chain reaction (pcr)-based approaches. 

Advanced PCR-based diagnosis with the help of highly Xcm-specific primers (Adikini et al., 2011; Adriko et al., 2012) significantly improved disease diagnosis. This approach allows for high throughput analysis. PCR has also facilitated Xcm genotyping studies. PCR-based diagnosis

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was further simplified through the development and use of DNA kits to capture pathogen and host DNA from infected banana plants in the field. DNA capture kits used included Whatman FTA (Flinders Technology Associates) cards, PhytoPASS kits and two-minute extraction DNA dipsticks (central nitrocellulose membrane upon which lateral flow devices are based) (Ramathani and Beed, 2013). The benefit of these kits is that they are not bulky, can retain DNA at high integrity for several months and can be used with several molecular-based diagnostic techniques in laboratory conditions. Furthermore, they only retain DNA material. As the pathogen is not alive and infective, the material can be moved across country borders without contravening phytosanitary regulations. Quarantine Quarantine forms the first line of defence to prevent the introduction of a disease into a new country or region. Unfortunately, the quarantine infrastructure in East and Central Africa is generally poor. In addition, quarantine laws are often weak or not implemented. Some countries lack the human resources and/or frameworks for implementing quarantine measures. The situation is further complicated by the porous nature of borders between countries, allowing for easy exchange of plant materials and banana bunches. During the initial stages of the Xanthomonas bacterial wilt outbreak, quarantine implementation was also hindered by a lack of surveillance and limited information exchange within and between countries in the region. The more recent development of a lateral flow device for rapid diagnosis of Xcm (Hodgetts et al., 2014; Karamura et al., 2016) could potentially strengthen implementation of quarantine measures, though it comes at a time when the disease has spread to nearly all the banana production zones in the region. Effective control through quarantine is also compromised by the mode of its spread: the movement of insect, bird and bat vectors can hardly be prevented. However, the strengthening of the quarantine laws and infrastructure is still vital to prevent the free flow of diseased planting material, fruit or fresh banana leaves from affected to unaffected areas and thus slowing the rate of spread. There are ongoing studies to map/identify environments

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where banana is vulnerable to Xanthomonas bacterial wilt for the purposes of introducing proactive preventive measures (W. Ocimati, Uganda, 2016, personal communication). De-budding of the male banana ­inflorescence The male flower is the primary infection site for insect-mediated transmission. The early removal of the male bud (immediately after the formation of the last hand) through the use of a forked wooden stick (Plate 5.12) has proved to be very effective in preventing disease incidence. Beneficial side effects of this control practice have been bigger and more evenly filled fruit (Blomme et al., 2005, 2007). The use of a forked wooden stick instead of a knife on consecutive plants, regardless of their disease status, avoids cross-infections. While de-budding is widely used in largescale commercial plantations in Latin America to control Moko bacterial wilt, its adoption has been inconsistent amongst small-scale farming communities in East and Central Africa (Kagezi et al., 2006; Mwangi and Nakato, 2007). Kagezi et al. (2006) and Muhangi et al. (2006) reported that most small-scale farmers remove the male buds only sporadically and often too late to be fully effective in preventing insect vector-mediated transmission. Reasons cited for the observed poor adoption of the technique include lack of labour, use of the male buds to identify infected plants and the traditional belief that early male bud removal would negatively impact on the

Plate 5.12.  The early removal of the male bud (immediately after the formation of the last hand) through the use of a forked wooden stick (photo: G. Blomme, BI).

juice volumes of the cultivars used to make beer (Kagezi et al., 2006). Male buds need to be removed early after the formation of the last hand. However, very early removal may prevent the full development of the lower hands of the bunch or make them curve upwards, negatively affecting bunch size (Biruma et al., 2007). Use of clean garden tools Because of the importance of mechanical transmission of the disease via cutting tools between and within banana plots, farmers are advised to disinfect tools routinely when pruning, de-­ suckering or harvesting from different mats. In East and Central Africa, disinfection by heating tools over a fire (Plate 5.13) or using a solution of household bleach (3.5% sodium hypochlorite (NaOCl)) has been recommended. A solution of 15 ml of household bleach in 0.5 l of water has been reported by FAO as an effective treatment in South Kivu, Democratic Republic of the Congo. Tree branches or wood are not necessarily needed to fuel a small fire within a banana field as crop residues and dried banana leaves produce enough heat to disinfect garden tools (G. Blomme, Uganda, 2015, personal communication). Though shown to be effective, these practices have only been marginally adopted on small-scale farms (Karamura and Johnson, 2010; Blomme et al., 2011). Farmers have reported that flaming contaminated tools in fire is difficult because of constraints on fire-building and bactericidal chemicals are often too expensive and scarce in local markets (Karamura and Johnson, 2010). Nakato and colleagues investigated the antibiotic potential of plants that were growing in banana-growing regions in both Uganda and eastern Democratic Republic of the Congo as alternative garden tool disinfectants over contact periods ranging from 1 min to 24 h. Extracts from Tephrosia, Bidens pilosa, Nicotiana tabacum and Moringa sp. seed eliminated less than 30% of Xcm, while Allium sativum, Carica papaya, Capsicum annum, Solanum lycopersicum and Persea americana eliminated over 90% of the Xcm ­populations. In combination, the plant extracts eliminated between 75% and 100% of the Xcm on the garden tools. This study recommended the use of plant extracts as a practical, affordable and inexpensive means to disinfect garden tools. However, their applicability under small-scale



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Plate 5.13.  Tool disinfection using fire. The tool needs to be held in the fire until the metal is too hot to touch. The fire can be made with dried banana leaves or crop residues (photo: W. Ocimati, BI).

farmer conditions still needs to be tested prior to wide-scale promotion. The required/optimum contact period with the plant extracts for effective elimination of all Xcm bacteria varied between 1 min and 15 min. This work is as yet unpublished (V. Nakato, Uganda, 2016, personal communication). Most farmers work with one machete/knife and may not have the time or patience to apply these time-consuming techniques. Moreover, the 10% Xcm inoculum that survives when the best performing botanicals are singly applied could potentially be sufficient for perpetuating the disease on-farm. Complete mat uprooting and single ­diseased stem removal The uprooting of complete mats has been recommended as part of a control package since the disease arrived in East and Central Africa. However, farmers are not always inclined to remove an entire banana mat when only one stem may be showing disease symptoms (Mwangi and Nakato, 2007). Uprooting of entire mats is cumbersome, arduous and costly. As a consequence, it is often shunned by farmers (Jogo et al., 2013; Ocimati et al., 2015). Nevertheless, its use could

be promoted for eradicating new infections in fields or locations where the disease is reported for the first time. This practice can also be selectively promoted in commercially oriented production systems that can afford the required higher labour costs. Field observations in less intensively managed banana systems, such as in central Uganda and eastern Democratic Republic of the Congo, and research findings suggest that Xcm does not colonize all physically attached lateral shoots of an infected banana plant (Ocimati et al., 2013d, 2015). In work undertaken in 2009–2010, while most East African highland banana cultivars (AAA) and ‘Pisang Awak’(ABB) artificially inoculated through the inflorescence died, relatively few attached lateral shoots showed symptoms (Ocimati et al., 2013d). These findings resulted in the development of the single diseased stem removal (SDSR) control technique whereby, upon symptom expression, the diseased stems are cut off at soil level. The underlying logic of SDSR is that the continued removal of diseased stems in a mat or field will reduce the inoculum level and will bring down disease incidence to an acceptable level (Blomme et al., 2017b). This method aims at control, not eradication. It is quicker and far less labour intensive

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compared with the removal of a complete mat, but SDSR needs to be combined with measures aimed at preventing new infections that can occur through the use of contaminated garden tools or through insect vector transmission. Recovery of moderately infected fields by farmers who were applying this technique has been reported in Uganda (Kubiriba et al., 2012; Jogo et al., 2013; Blomme et al. 2014; Ocimati et al., 2015). Multi-year on-farm assessments of the SDSR technique at more than ten pilot sites in eastern Democratic Republic of the Congo have shown that disease incidence levels can be reduced from a level as high as 90% at the onset of the trials to below 1% after 3–4 months of application (Blomme et al., 2014; ProMusa, 2014; Blomme, 2015; RTB, 2015; Vezina, 2015; Blomme et al., 2017b). Research at these pilot sites has shown that the SDSR technique should be practised at weekly intervals during the first months of intervention; once incidence levels have dropped substantially, this could be reduced to bi-weekly interventions. However, research has also shown that latent infections can occur in infected lateral shoots that are attached to an infected mother plant (Ocimati et al., 2013d, 2015). Symptom appearance in these shoots, which continue to grow vigorously and produce edible bunches, may take up to 24 months (Ocimati et al., 2013d). As symptoms may still appear at later growth stages or during subsequent crop cycles, a continued follow-up, preferably over several years, is recommended when practising SDSR as part of a control package. The need for the application of the technique with other control measures is especially important when dealing with ABB genotypes, such as ‘Pisang Awak’, which are highly susceptible to insect vector transmission or in lowland areas where insect populations and insect-­ mediated transmission levels are high. It is advised to minimize the use of garden tools for de-­suckering or de-leafing operations until very few visibly sick plants with disease symptoms are observed, which corresponds to a 3–4-month period after the start of SDSR application. Chemical destruction of diseased mats using herbicides The use of the herbicides 2,4-D and glyphosate for the destruction of diseased mats was reported to be at least 100-fold more effective than

­ echanical destruction in terms of the labour m and required time (Blomme et al., 2008). ‘Pisang Awak’ plants were observed to snap because of rotting of the pseudostem and corm 3 weeks ­after 1.6 ml of the original concentrate of 2,4-D was injected into the pseudostem. Herbicide treatment also reduces the sprouting of lateral buds (Okurut et al., 2006; Blomme et al., 2008). These studies reported that 2,4-D was more lethal than glyphosate. However, the use of 2,4-D has safety concerns, especially when free-ranging domestic animals are present, as its degradation in or on the soil can take up to 1 month. In contrast, glyphosate is non-persistent. Okurut et al. (2006) thus concluded that glyphosate was the preferred herbicide, as health and environmental safety outweighed the usefulness of 2,4-D. Despite the observed benefit of herbicides, their adoption by farmers in East and Central ­Africa has been poor, possibly due to the inaccessibility of herbicides in rural areas, perceived high cost of herbicides, reluctance to inject an already infected plant, and reluctance to inject symptomatic plants as physically attached asymptomatic plants will also be affected (Blomme et al., 2014). The targeting of market-­oriented and input-­intensive banana farming systems is recommended with this technology. Continuous cutting of re-sprouts Continuous cutting of banana stems and resprouts has been evaluated as an alternative to manual uprooting of complete mats in the eastern Democratic Republic of the Congo. Cut corms took up to 2 years to completely rot while re-­ sprouting (shoot production) ceased by the eighth month after trial initiation (Ntamwira et al., 2016). The injection of 2,4-D into the centre of corms and the addition of a layer of organic manure on the cut corm surface significantly hastened the decay process (Ntamwira et al., 2016). Despite the long time to corm decay, a cost–­ benefit analysis of this method in comparison with complete mat uprooting showed that it has significant benefits. The approach is relatively easy to apply and corms were relatively easy to uproot 7–9 months after trial initiation because of poor anchoring in the soil as a result of an absence of new re-sprouts and a limited number of remaining functional cord roots (Ntamwira et al., 2016). This approach is also environmentally sound, as it prevents the degradation of soils



Diseases Caused by Bacteria and Phytoplasmas

­ ssociated with digging of corms and the in situ a decomposition of roots and corms maintains/ enhances soil fertility. However, for wider adoption, trade-offs between immediate clearing through complete mat roguing for immediate establishment of other crops and the soil fertility and labour benefits associated with continuous cutting will need to be communicated to farmers (Ntamwira et al., 2016). Uprooting entire diseased fields and banana-free fallows Based on the fact that Xcm cannot survive in the soil for long in the absence of living banana tissue (Mwangi et al., 2007), uprooting entire fields coupled with a 6-month banana-free fallow (grass fallow or planting break crops) before replanting with clean banana planting materials was recommended in central Uganda (Turyagyenda et al., 2008). At sites in eastern Democratic ­Republic of the Congo with a high disease pressure, banana-free fallows were only effective after a 12-month break (Sivirihauma et al., 2013; Rutikanga et al., 2016a). This was attributed to secondary reinfections originating in neighbouring fields. A collective effort by farmers across affected landscapes was deemed necessary if the disease was to be eradicated after 6 months of banana-­ free fallow (Rutikanga et al., 2016a). Rutikanga et al. (2016a) reported that such collective action is often hard to implement in areas where extension structures are weak and/or production is not market oriented. In contrast, collective action was successfully used to manage Xanthomonas bacterial wilt in the western province of Rwanda through government enforcement (Rutikanga et al., 2016a). Extension task forces operating in the market-oriented banana production system in southwestern Uganda were also successful (Tinzaara et al., 2013; Rutikanga et al., 2016a).

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conditions and often indexed for viruses. They are thus considered pathogen- and pest-free. propagated plantlets and/or suckers, Macro-­ though not produced under such strict conditions as the tissue culture plantlets, are an acceptable alternative if precautions are taken to ensure freedom from Xanthomonas bacterial wilt. However, access to clean planting material to gap-fill or replant uprooted fields has been a limiting factor in the control of the disease. In banana-producing regions such as Democratic Republic of the Congo, infrastructure for producing tissue-culture plantlets is lacking. This is worsened by the fact that the banana crop in East and Central Africa is produced by resource-poor smallholder households that have no access or capacity to purchase these clean planting materials. Hence, the use of tissue-culture plantlets and other clean seed materials is not common (Ndungo et al., 2008; Ocimati et al., 2013a). Suckers are most often sourced from farmers’ own and neighbouring fields, enhancing disease spread and inoculum build-up (Ndungo et al., 2008; Ocimati et al., 2013c). Farmers’ capacities to use clean seed in the different banana-producing zones or agro-ecologies urgently need to be assessed so that interventions can be tailored to requirements. Use of symptomless suckers sourced from infected fields

As mentioned above, in regions such as eastern Democratic Republic of the Congo, farmer access to clean planting materials is either limited or non-­ existent. Here, farmers rely almost entirely on suckers sourced from their own or neighbouring fields (Ocimati et al., 2013a). Controlled studies had shown a localized spread of Xcm in banana mats (Ocimati et al., 2013d, 2015), indicating that the use of symptomless suckers sourced from diseased fields should be investigated as a possible source of planting material. Initial experiments by Sivirihauma et al. (2013) in eastern Democratic Use of clean planting material Republic of the Congo did not show much success, Clean planting material, in the form of tissue-­ mainly due to new infections, which had entered culture plantlets, macro-propagated plantlets the trial plots from s­ urrounding heavily diseased and/or suckers sourced from clean mother gar- fields. Repeat experiments in which instances of dens, is recommended for the establishment of external infection were limited by tool sterilizanew fields or restoring diseased banana fields tion, fencing trial plots to avoid disease entry via previously uprooted and fallowed. Tissue-culture browsing animals, and de-­budding to minimize mediated spread have shown very low plantlets are produced under sterile laboratory insect-­

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et al., 2013). In a study to determine the role of putative resistance-related host genes (MbNBS, MdNPR1 and PR3), Ssekiwoko et al. (2015) observed a sharp decline in gene expression in the first 6 h and a later recovery. The recovery of gene expression did not stop symptom development in both the resistant and susceptive cultivars. They concluded that the genes studied Use of banana cultivars that escape were not responsible for blocking wilt symptom insect-mediated transmission development in M. balbisiana plants. As Xcm supSome banana genotypes have specific morpho- pressed the key genes in defence pathways durlogical characteristics that enable them to es- ing the early stage of infection, Ssekiwoko et al. cape insect vector-mediated Xcm transmission (2015) thus believed that neither a hypersensi(referred to as ‘wilt-escaping’ cultivars). These tive response, nor systemic acquired resistance genotypes possess physical characteristics/bar- nor induced systemic resistance accounted for riers that prevent the entry of Xcm bacteria via the observed resistance in M. balbisiana. Studies the male inflorescence part. Horn plantains to understand the role of phenolic compounds completely lack the male inflorescence while in M. balbisiana resistance to Xcm have been recothers, such as the ‘Dwarf Cavendish’ (AAA) ommended. If the source of resistance in M. baland some East African highland cultivars, espe- bisiana is identified, the gene(s) can then be cially those of the Nakitembe clone-set, such as incorporated, through genetic engineering ap‘Mbwazirume’, ‘Nakitembe’, and ‘M9’ (bred hy- proaches, into commercial cultivars to impart brid), have persistent or semi-persistent male resistance to the disease. Transgenic technologies have already flowers/bracts that leave no fresh open wounds on the rachis. These attributes prevent insect-­ been explored as a cost-effective alternative solumediated Xcm transmission (Addis et al., 2004; tion for breeding Xanthomonas bacterial wilt-­ Shimelash et al., 2008; Tripathi and Tripathi, resistant materials in Uganda (Tripathi et al., 2010). 2009; Mudonyi, 2014). However, these ‘escape’ Transgenic banana lines with the hypersensitive-­ cultivars can also easily be infected, like any of assisting Hrap and plant ferredoxin-like protein-­ the other edible banana cultivars, through con- producing Pflp genes from sweet pepper (Capsicum annuum), which intensify the hypertaminated garden tools. sensitive response, have been generated using agrobacterium-mediated transfer techniques Development of resistant cultivars and embryogenic cell suspensions (Tripathi et al., The use of resistant cultivars is a cost-effective 2010; Namukwaya et al., 2012; Tripathi et al., method of managing bacterial diseases. Develop- 2014). Transgenic lines that did not show any ment of disease-resistant banana clones through disease symptoms after artificial inoculation unconventional breeding has not been feasible as it der laboratory and screenhouse conditions were requires resistant donor parents. High levels of selected for field evaluations (Namukwaya et al., cell-mediated resistance to Xcm have not been 2012; Tripathi et al., 2014). Promising transidentified in edible banana or enset germplasm genic lines that have proved to be resistant over (Ssekiwoko et al., 2006a; Shimelash et al., 2008; three crop cycles in the field have been identified Tripathi et al., 2009). Additionally, conventional (Tripathi et al. 2014), but await a legal framebreeding of banana is difficult because of the ste- work for release. These transgenes showed normal growth and fruit development, suggesting rility problems and the lengthy breeding cycle. Musa balbisiana, a wild and inedible relative that the over-expression of the Hrap or Pflp gene of banana, has been found to be resistant to Xcm does not alter plant physiology. Transgenic bawhen artificially inoculated (Ssekiwoko et al., nana clones expressing these genes appear to 2006b; Mudonyi, 2014). Slow Xcm multiplica- have significant potential to overcome the Xantion and migration and death of affected plant thomonas bacterial wilt pandemic, which will tissue led to delayed disease development and boost the available set of control options to fight severity and complete plant recovery (Ssekiwoko this epidemic. This technology may also provide ­ isease incidence levels. Between 0% to 4% of d plants in plots established with asymptomatic suckers obtained from fields with an incidence of > 80% were recorded (C. Sivirihauma, Democratic Republic of the Congo, 2016, personal communication).



Diseases Caused by Bacteria and Phytoplasmas

effective control for other bacterial diseases, such as Moko and blood bacterial wilts of banana occurring in Latin America and Asia. Antagonistic bacteria Beneficial microorganisms and their secondary metabolites can directly or indirectly inhibit the growth of other disease-causing microbes through antibiosis, parasitism, competition for niches and nutrients, interference with pathogen signalling and/or priming the host plant’s own defence systems (Cook and Baker, 1983; Wilson and Lindow, 1993; Agrios, 2005; Harish et al., 2008; Reed and Glick, 2013). Bacterial antagonists have been used in many studies to suppress bacterial pathogens. For example, when the biological control agents Pseudomonas fluorescens A506 and Erwinia herbicola C9-1 are applied on the blossoms of pear and apple stigmas at early to mid-bloom, they proliferate and exclude Erwinia amylovora from the infection sites. With two applications, the incidence of fire blight is reduced by about 60% (Johnson et al., 1993; Nuclo et al., 1998). Different bacterial strains, namely Pseudomonas fluorescens, Pseudomonas putida, Bacillus subtilis and Enterobactor aerogenes, have been reported to improve the tolerance of tomato to bacterial wilt of tomato cause by Ralstonia solanacearum (Seleim et al., 2011; Maji and Chakrabartty, 2014). Biological control of Xcm through the use of antagonistic bacteria remains a possible option for controlling the disease. In initial pot experiments in Ethiopia, four bacterial antagonists against Xcm were observed to reduce wilt incidence by 56–75% (Abayneh, 2010). Initial laboratory-based studies on banana in Uganda have shown promising levels of Xcm suppression by some bacterial isolates, namely Burkholderia spp., Herbaspirillum spp. and Enterobacter spp. isolated from the banana tissues from different locations in Uganda (Were, 2016). Combined with other control components, the use of microbial antagonists may prove to be a successful and ecologically safe strategy to reduce the incidence and yield loss due to Xanthomonas bacterial wilt. Use of simulation models in disease management and forecasting Past and current studies have mostly approached the Xanthomonas bacterial wilt problem by

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f­ocusing only on single elements affecting epidemics. Models provide frameworks that allow ideas about the behaviour of a particular system to be conceptualized and communicated (Keeling and Eames, 2005; Groot et al., 2009). Depending on a chosen goal, mathematical modelling can produce valuable theoretical results that can be used to suggest or design epidemic control ­programmes. Mathematical models have been used to study the dynamics of diseases in humans, animals and plants (Fishman et al., 1983; Fishman and Marcus, 1984; Keeling and Eames, 2005; Garner and Beckett, 2005; Garner et al., 2007). Modelling can contribute to better disease control through retrospective analysis of past outbreaks and evaluation of different control strategies, contingency and resource planning, risk assessment and priority setting for target preparedness and surveillance, provision of scientific advice, evaluation of the effectiveness surveillance strategies, underpinning economic impact studies, and provision of realistic scenarios for training exercises and communication (Taylor, 2003; Dubé et al., 2007). The complexity of issues related to the bacterial wilt disease of enset and banana necessitates the use of supporting methodologies and models to inform stakeholders and policy makers, to design alternatives and to explore future disease scenarios. Simulation models through integration of different disciplines look at a problem in a systems dimension and help to explain and/or solve more complex scenarios. Advances in computing power, availability of PC-based modelling and simulation, and efficient computational methodology have made simulation modelling feasible. More recent studies have led to the development of deterministic models to understand the role of the disease transmission parameters and cultural control strategies, more specifically insect vector, tool and mother-to-sucker spread of Xcm, in the reduction of disease threshold and prevalence (Kweyunga, 2011; Nannyonga et al., 2015; Nakakawa et al., 2016). For example, Nakakawa et al. (2016) concluded that the current cultural control practices including de-budding and regular roguing through single stem removal were effective control measures for eradication in the absence of tool-based transmission. While these deterministic mathematical models that are typically based on mean or

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e­xpected value parameters may be useful for ­understanding basic infection dynamics, they are often of limited use as a predictive tool since any one epidemic is unique and unlikely to follow an ‘average’ pattern (Garner et al., 2007). Multi-scale, multi-stakeholder and integrative approaches that look at the problem in a systems dimension are, therefore, still needed to understand all the issues holistically and make accurate predictions and explorations to improve disease management. The increasing sophistication of computers, together with greater recognition of the importance of spatial elements in spread of disease and interest in specific spatially targeted strategies, has led to increased importance of models that incorporate spatial components in epidemiological studies (Garner and Beckett, 2005). There is an ongoing study to elucidate the spatial dimensions of disease dynamics at a regional level so as to identify vulnerable landscapes for proactive preventive measures (W. Ocimati, Uganda, 2016, personal communication). Modelling of the impact of climate on Xanthomonas bacterial wilt in a 300 km2 area in both Burundi and Rwanda, with altitudes ranging from below 900 m to as high as 3000 m, is also in progress (G. Blomme, Ethiopia, 2016, personal communication). Network-based modelling, a relatively new but growing field to study the spread of diseases through contact networks (Keeling and Eames, 2005), is also being explored to understand the epidemiology of Xanthomonas bacterial wilt (G. Blomme, Ethiopia, 2016, personal communication).

Moko Bacterial Wilt and Bugtok S.J. Eden-Green Introduction Moko bacterial wilt takes its name from the ‘Moko’ cultivar (ABB, syn. ‘Bluggoe’), which was virtually eliminated by the disease in Trinidad in the 1890s (Rorer, 1911). Characteristic symptoms were observed in British Guiana (Guyana) some 70 years earlier suggesting that the disease is of considerable antiquity in South America (Wardlaw, 1935; Buddenhagen, 1961).

The commercial significance of Moko bacterial wilt became apparent with the development of large-scale dessert (AAA) banana plantations in Central America after the Second World War. An improved understanding of the disease led to the introduction of effective control measures in the late 1950s and 1960s and serious losses are now rare in intensively managed commercial plantings where continuous surveillance and prompt control interventions can be maintained. Such control measures are much more difficult to sustain under smallholder and domestic plantings, especially where ABB genotypes, such as ‘Bluggoe’, predominate. Under such conditions, significant losses have been reported, for example, in Colombia (Buddenhagen and Elsasser, 1962; Lozano et al., 1969), Peru (French and Sequeira, 1970), Nicaragua (Lehmann-Danziger, 1987), Grenada (Hunt, 1987), Trinidad and Guyana (Phelps, 1987), Mexico (Fucikovski and Santos, 1993), Belize (Black and Delbeke, 1991) and Suriname (Power, 1976). In the New World, Moko bacterial wilt has been reliably reported from Mexico, Guatemala, Belize, Honduras, El Salvador, Nicaragua, Costa Rica, Panama, Colombia, Venezuela, Guyana, Suriname, Trinidad, Grenada, Brazil, Peru, Ecuador (Stover, 1972; Phelps, 1987), Jamaica (EPPO, 2004) and St Vincent (EPPO, 2007). Moko bacterial wilt also occurs in commercial plantations of dessert (AAA) cultivars in the Davao region of Mindanao in the Philippines, where it is believed to have been introduced on propagating material imported from Honduras in 1968 (Rillo, 1979; Buddenhagen, 1994). A  few years earlier, a disease affecting fruit of ‘Saba’ (ABB) and ‘Cardaba’ (ABB) was observed by Roperos (1965) in the same region. Local residents reported that this condition, known locally as bugtok in the southern Philippines and also by the local names tapurok and tibaglon elsewhere in the Philippines, had been present for some time prior to its discovery (Zehr and Davide, 1969). Bugtok was recognized as a disease of fruit, which became discoloured and eventually rotten, resembling bacterial finger-tip rot of ‘Saba’ reported by Stover (1972). However, affected plants developed more extensive vascular symptoms similar to those of Moko bacterial wilt. Bugtok was not recognized in commercially grown dessert-banana cultivars, but in many areas of the southern Philippines the disease



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­ ecame a limiting factor in the cultivation of b cooking banana. The differences in symptoms between Moko bacterial wilt and bugtok derive from the fact that the former occurs on dessert banana in plantations, while the latter is confined to cooking varieties, usually grown in smallholdings. Although caused by the same pathogen, the two diseases are separated geographically, by host genotype, cultivation practices and by the usual modes of transmission. Moko bacterial wilt has recently been reported from peninsula Malaysia (Zulperi and ­Sijam, 2014; Zulperi et al., 2016), but others have reported the presence of blood bacterial wilt (Timin et al., 2014; Teng et al., 2016). The pathogens cause similar symptoms, and although it is unclear which, or whether both, of the pathogens have become established, there is an imminent threat of further spread in other countries in continental Southeast Asia.

Symptoms All parts of the plant may be invaded by bacteria, but the route of infection, host cultivar and the nature of the causal strain determine the ­sequence and severity of symptoms. Infection of dessert (AAA) banana cultivars typically occurs via mature leaves, roots or rhizome. In this ‘bottom-up’ mode of invasion, the first signs of the disease are yellowing and flaccidity of the oldest leaves, which eventually become necrotic and collapse at the base of the petiole. Symptoms spread to the younger leaves, which develop pale green or whitish panels before becoming necrotic. Suckers may wilt ­ (Plate  5.14) with little or no foliar discolouration.Vascular tissues become progressively discoloured (Plate 5.15), initially cream or yellow but later forming dark streaks which may be seen throughout the plant, eventually extending into the daughter suckers. Within a few minutes of cutting, these exude a viscous bacterial ooze, ranging in colour from pale yellow to reddish brown or black. In fruit-bearing plants, discoloured vascular bundles tend to be concentrated within the fruit stem and bases of the younger leaves. Fruit development is arrested and fingers may ripen prematurely or split. Internally, fruits become discoloured and eventually rot.

Plate 5.14.  Collapsed and withered heart leaf of a sucker of ‘Robusta’ (AAA, Cavendish subgroup) caused by Moko bacterial wilt in Grenada. The necrotic margin of the next leaf is also reputed to be a symptom (photo: D.R. Jones, INIBAP).

Early symptoms in the flowers and immature fruits indicate ‘top-down’ infection, by insect transmission to male flower parts or via contaminated tools used for removing male buds (de-­ budding). Symptoms of insect-transmitted disease are very specific to this group of bacterial diseases and are useful distinguishing features from Fusarium wilt. The male flower buds and peduncles become blackened and shrivelled. Within a few days of infection, droplets of cream-coloured or pale yellow bacterial ooze emerge from recently exposed scars of bracts and peduncle cushions (pedicels), which are frequented by stingless bees (Trigona spp.) and other insects that can become contaminated and spread the disease (Buddenhagen and Elsasser, 1962). The bacterium spreads to the fruit and initiates a rot. Fruit clusters may at first appear outwardly normal, but subsequently ripen prematurely (Plate 5.16). By this stage, all fruits typically show internal discoloration

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Plate 5.15.  Brown discoloration in the vascular tissues of the pseudostem of ‘Grand Nain’ (AAA, Cavendish subgroup) caused by Moko bacterial wilt in a commercial plantation in Honduras (photo: D.R. Jones, INIBAP).

and rotting, which can vary in colour (red, brown or black) and texture (firm or gelatinous) (Plates 5.17 and 5.18). The infection continues into the peduncle, causing blackening of the vascular tissues (Plate 5.17). The youngest ‘flag’ leaves, which are subtended from the flower stem, may become discoloured and flaccid, an early diagnostic symptom that is recognized by experienced growers. Eventually the whole mat may become diseased, but bacterial invasion can also be incompletely systemic and may remain confined to the affected pseudostem or fail to spread into all of the daughter suckers. This is typified by bugtok disease of ABB cultivars in the Philippines (Soguilon et al., 1995), but has also been described in ‘Bluggoe’ infected by insect-­ transmitted strains of Moko bacterial wilt in Central America (Stover, 1972; Black and Delbeke, 1991; Albuquerque et al., 2014). Suckers that become infected may wilt and die, or may ­remain symptomless (tolerant) and survive to

Plate 5.16.  Premature ripening and splitting of fruit of ‘Robusta’ (AAA, Cavendish subgroup) affected by Moko bacterial wilt in Grenada (photo: D.R. Jones, INIBAP).

Plate 5.17.  Internal rot of fruit of ‘Grand Nain’ (AAA, Cavendish subgroup) caused by Moko bacterial wilt in a commercial plantation in Honduras. The infection also extends into the peduncle (photo: D.R. Jones, INIBAP).

produce inflorescences, which can be infected by insects. In this way, infected mats may persist indefinitely and act as sources of inoculum for further spread of the disease.



Diseases Caused by Bacteria and Phytoplasmas

Plate 5.18.  Internal rot symptoms of bugtok in the cut fruit of an ABB banana cultivar in the Philippines (photo: S. Eden-Green, NRI).

Causal agent The pathogen causing Moko bacterial wilt belongs to a large and diverse taxonomic group referred to as the Ralstonia solanacearum species complex (RSSC) (Allen et al., 2005), reflecting the polyphyletic and likely geographical origins of strains specializing in different hosts and ­habitats. Ralstonia solanaceraum (Yabuuchi et al., 1995), formerly Pseudomonas solanacearum then Burkholderia solanacearum, is an aerobic, Gram-­ negative rod belonging to the non-fluoresent Pseudomonas rRNA homology group II of ­Palleroni (1984). Rorer (1911) named the causal agent of Moko bacterial wilt as Bacillus musae and noted its similarity to Bacillus solanacearum (now Ralstonia solanacearum). Subsequent investigations showed that banana isolates were pathogenic to solanaceous plants, and the bacterium was reclassified as a member of the species then known as B. solanacearum (Ashby, 1926). However, several decades elapsed before researchers recognized that banana is infected by a distinct subgroup of R. solanacearum, designated as race 2. Independent appearances of new strains, with differing pathological and epidemiological characteristics on cultivated banana, have been noted in regions extending from Costa Rica to Peru, suggesting that these evolved both in wild Heliconia species (Buddenhagen, 1960; Sequeira and Averre, 1961) and in other specialized niches in Central and South America (French and Sequeira, 1970; Buddenhagen, 1986).

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A casamino acid-based indicator medium containing 2,3,5-triphenyltetrazolium chloride at 0.1 g/l (Kelman, 1954) has been widely used to distinguish strains of the Moko bacterial wilt pathogen on the basis of different colony characteristics, as well as allowing for tentative discrimination from contaminating organisms in primary isolations from diseased material. Most strains are readily isolated from infected plants and various selective media, based on Kelman’s medium, have been described to reduce ­contamination (Granada and Sequeira, 1983a; ­Engelbrecht, 1994). Spontaneous avirulent mutants of R. solanacearum arise during repeated subculturing and exhibit non-fluidal, butyrous colonies on Kelman’s medium. This phenomenon, known as phenotype conversion, involves a wide range of phenotypic alterations, including loss of extracellular polysaccharide production and changes in the activity of various exported enzymes. ­Motility by means of polar flagella may not be evident in freshly isolated virulent cultures and is also associated with phenotype conversion (Kelman and Hruschka, 1973; Brumbley and Denny, 1990; Schell, 2000). Traditionally, strains of R. solanacearum have been grouped into five biovars on the basis of carbohydrate catabolism (Hayward, 1964; He et al., 1983) and five races designated by host range (Buddenhagen et al., 1962; He et al., 1983). Strains pathogenic to triploid banana cultivars are designated race 2 and conform to biovar 1. Banana strains are also pathogenic to solanaceous host, such as tomato and potato, and elicit a hypersensitive response when infiltrated into tobacco leaves. This distinguishes them from generalist strains isolated from many solanaceous and non-solanaceous host plants (race 1) and potato pathogens assigned to race 3 (Lozano and Sequeira, 1970; Cellier et al., 2012). Race 1 strains are common in banana-growing areas, even where race 2 is not present (Stover, 1972). While race 1 strains have not been reported to cause bacterial wilt of triploid banana cultivars, they have been implicated in a wilt disease of the wild diploids M. acuminata spp. banksii, M. acuminata spp. zebrina and M. schizocarpa in Honduras (Buddenhagen, 1962) and M. acuminata spp. banksia, M. schizocarpa and M. lolodensis in Australia (Akiew, 1992). In Australia, diseased plants had irregular, soft patches of black rot on the

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­ seudostem, dark brown to black necrotic lep sions in the vascular tissues, and droplets of brown, slimy bacterial ooze on cut surfaces of the pseudostem. Akiew (1992) was also able to infect ‘Sucrier’ (AA), a cultivated diploid. On this host, isolates of race 1 from banana caused yellowing of leaves and stunted growth, whereas race 1 isolates from heliconias caused severe wilt. Ralstonia solanacearum race 1 has also been associated with a bacterial wilt of M. textilis (abacá) in the Philippines (see Abacá bacterial wilt section later in this chapter). Strains of the Moko bacterial wilt pathogen, with distinctive colony characteristics on tetrazolium chloride (TZC) medium, have been described from different ecological niches, reflecting the presumed origins and evolution of the pathogen. Banana (B) strains, persisting in soils and spread mainly through cultivation and planting practices, originated in plantations developed from land previously growing wild Heliconia species, from which isolates pathogenic to banana were obtained (Buddenhagen, 1960). Some isolates from heliconias were indistinguishable in culture from B strains, but caused stunting, ­distortion and slow wilting of young banana plants (D strains). Others produced colonies with dense red formazan pigment from TZC and were pathogenic to heliconias and plantain (AAB), but not dessert-banana cultivars (H strains) or mildly pathogenic only to heliconias (R strains) (French, 1986). Afluidal colony forms (AFV) caused mild wilting (Woods, 1984). The pathogenicity of D, but not R, strains increased after serial passage through banana (Sequeira and Averre, 1961; French and Sequeira, 1970), supporting the view that pathogenicity to banana evolved from naturally infected heliconias. Isolates with a small fluidal round (SFR) colony morphology on TZC medium were first reported from the outbreak of insect-transmitted Moko bacterial wilt on ‘Bluggoe’ (ABB) in Honduras described by Buddenhagen and Elsasser (1962). This strain apparently originated in planting materials introduced from Venezuela (Sequeira, 1998). In contrast to B strains, these produced copious amounts of bacterial ooze from infected inflorescences which was considered to be a likely adaptation to their apparent mode of insect transmission. SFR and other colony variants (SFR-C and A strains) have been described in association with other insect-transmitted

e­ pidemics originating in Colombia and spreading in ‘Chato’ (ABB, Bluggoe subgroup) in the Amazon basin into Peru and Brazil (French and Sequeira, 1970). Strains with SFR characteristics are now widely distributed in South and Central America and the southern Caribbean (Phelps, 1987; ­Lehmann-Danziger, 1987; Albuquerque et al., 2014). The causal agent of the fruit disease bugtok in the Philippines has been shown to be indistinguishable from a B strain of R. solanacearum causing Moko bacterial wilt in Honduras (EdenGreen, 1994a; Thwaites et al., 1999a; Ilagan et al., 2003; Fegan, 2005; Fegan and Prior, 2006 The phylogenetic relationships of banana pathogens within the RSSC have been rationalized into major phylogenetic subdivisions (phylotypes), based on analyses of sequence data from the 16s-23s rDNA interspacial region, and clusters of strains (sequevars) whose endoglucanase gene (egl) partial sequences differ by less than 1% (Remenant et al., 2011; Genin and Denny, 2012). This scheme proposed four distinct phylotypes, largely correlating with the geographical distribution and likely evolutionary origins of different strains: Asian (phylotype I), American (II), African (III), and Indonesian (IV) (Prior and Fegan, 2005a, b; Fegan, 2005; Fegan and Prior, 2005, 2006), consistent with earlier multi-­ location genotype (MLG) clusters (Cook et al.; 1989; Cook and Sequeira, 1994) from whole genome RFLP analyses. Stains of the Moko bacterial wilt and bugtok pathogen clustered into four subgroups within Phylotype II (sequevars IIA-6, IIA-24, IIB-3, IIB-4) whilst Phylotype IV contained the closely related taxa Ralstonia syzygii (causing Sumatra disease of clove) and the blood bacterial wilt bacterium, previously classified as a novel MLG (Cook and Sequeira, 1994). A very low genetic diversity among R. solanacearum strains from banana in the Philippines (Raymundo et al., 1998, 2005) supported the view that a single genotype of the pathogen was ­introduced from Central America. These findings have led to a proposal by Safni et al. (2014) to separate the RSSC into three genospecies based on a polyphasic taxonomic analysis of an extensive set of strains. Under this proposal, bacteria in Phylotypes IIA and IIB (which include all strains of the Moko bacterial wilt and bugtok pathogen) are designated as R.  solanacearum. In addition, bacteria in Phylotype IV are designated as R. syzygii and comprise



Diseases Caused by Bacteria and Phytoplasmas

three subspecies: ssp. syzygii (Sumatra disease of clove pathogen), ssp. celebesensis (blood bacterial wilt pathogen) and ssp. indonesiensis for other broad host range strains of mainly Indonesian origin. A new specific name, Ralstonia pseudosolanacearum, was proposed for Phylotypes I and III. Although the relationships between sequevars, origins, ecological and phenotypic characteristics remain to be fully elucidated, ­banana (B) and heliconia (H) strains clustered with bugtok isolates in sequevar IIB-3, whereas insect-­transmitted SFR and Amazon (A) strains grouped into two different and more distantly related clusters IIB-4 and IIA-6. Two previously unsuspected sequevars were subsequently shown to contain Moko bacterial wilt isolates in Brazil (IIA-41 and IIB-25) and a new cluster was found that was also associated with symptoms characteristic of insect transmission (IIA-53) (Albuquerque et al., 2014). These results suggest that capabilities for insect transmission in banana pathogens have evolved from several independent origins within South and Central America. As yet, little is known about how, from which hosts, and with what frequency insect-transmitted strains that cause Moko and blood bacterial wilts may have adapted to the niche provided by inflorescences of certain types of banana. Interestingly, naturally occurring insect transmission has been observed in diploid banana cultivars infected by strains of R. solanacearum race 1, which is normally considered a pathogen of solanaceous hosts and is not pathogenic to triploid bananas (Vakili and Baldwin, 1966). This suggests that this adaptation might not be an unusual event. Disease cycle and epidemiology In commercial plantings of dessert AAA cultivars, the main routes of survival and spread of the predominantly mechanically transmitted B strains of the Moko bacterial wilt pathogen are through roots following survival and persistence in the soil, in infected banana residues and possibly in association with weed hosts; or from plant to plant through cutting tools during pruning or de-budding operations; and from the introduction of infection via contaminated flood or irrigation water, infected planting materials, on contaminated footwear and possibly via livestock

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(Sequeria, 1958; French and Sequeria, 1970; Stover, 1972). Strains of R. solanacearum from infested soil may infect roots following wounding by soil-inhabiting insects and nematodes (Kelman, 1953) or, under experimental conditions, without mechanical injury (Kelman and Sequeira, 1965). Survival of R. solanacearum in soil is considered to be poor in the absence of amenable host plants (Granada and Sequeira, 1983b). Replanting trials under plantation conditions showed that B strains survived fallow periods of less than 18 months (Sequeira, 1962), reduced to 9 months with dry season tillage (Sequeira, 1958) or 12  months in well drained soils (Lozano et al., 1969). Wardlaw (1972) reported experimental evidence that the bacterium survived less than 6 months in soil maintained at or below 65% water-holding capacity. Infection of weeds may have a role in the Moko bacterial wilt disease cycle, though the persistence of the bacterium in weed hosts has been questioned (Buddenhagen, 1986). Berg (1971) isolated SFR strains from weeds, unrelated to banana, which occurred frequently on Honduras plantations, and B strains were reported to survive in the rhizosphere of certain natural flora (Wardlaw, 1972). Lengthy lists have been published of weed hosts considered important in outbreaks of Moko bacterial wilt in Colombia (Belalcazar et al., 2004; Romo et al., 2012). However, studies in Costa Rica showed that whilst R. solanaceaum was widespread in a range of cultivated and wild hosts in both Moko bacterial wilt-affected and non-infected areas, isolates pathogenic to banana were obtained only from banana or Heliconia spp. (Sequeira and Averre, 1961; Buddenhagen 1960, 1961). Inflorescence infections occur predominantly, if not exclusively, via the male flower parts and are related to the biology and phenology of the male flowers. Banana generally produces a new cluster of male flowers daily and, in many cultivars, clusters of older flowers and bracts are shed (dehisced) with similar regularity. This provides a series of infection courts with newly exposed xylem vessels from which droplets of a ‘nectar-like sap’, attractive to insects, have been observed to exude (Buddenhagen, 2009). Bacteria enter mainly through freshly exposed pedicels (‘cushions’) within about 2 days of abscission, following which inflorescence symptoms develop rapidly.

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SFR stains of Moko bacterial wilt exude from pedicels and bract scars from 15 to 25 days of infection and serve as a source for contamination of stingless bees (Trigona spp.), wasps (Polyoba spp.) and fruit flies (Drosophila spp.) (Buddenhagen and Elsasser, 1962; Buddenhagen and Kelman, 1964). Buddenghagen and Elsasser (1962) observed up to 100 bees and wasps per hour frequenting a single male ‘Bluggoe’ inflorescence, and bacteria with SFR characteristics were recovered from 5% of stingless bees (Trigona corvina) and wasps collected from a patch of diseased plants over a 20-day period. Bagging whole inflorescences to exclude insects, or breaking off the male buds before the first male flower cushions became exposed, prevented infections that otherwise spread rapidly to neighbouring plants. In isolated cases ‘Bluggoe’ plantings up to 8 km apart became infected and acted as new foci of infection. Similar groups of insects have been implicated as vectors of bugtok in the Philippines (Natural, 2011). Insect transmission of both B and SFR strains of Moko bacterial wilt can occur in commercial banana plantations (Stover, 1972) and undetected inflorescence infections can become sources for subsequent spread on cutting tools. However, inflorescence infections are rarely observed, because regular debudding and bagging of flower bunches means that dessert cultivars are likely to escape infection by this route. These features rarely pertain in smallholder plantings, where the preferred cultivars are often ABB types that have dehiscent male flowers and bracts and possibly other characteristics that render them particularly attractive to insects. These factors have contributed to epidemics of insect-transmitted Moko bacterial wilt on banana, which, like blood bacterial wilt and Xanthomonas bacterial wilt, have proved to be difficult to control. Insect-­ borne epidemics spread very rapidly and probably account for the historic accounts of devastating losses in ‘Bluggoe’ banana plants to Moko bacterial wilt in Guyana and Trinidad described by Rorer (1911). Recognition of the mode of insect transmission explained the rapidly spreading epiphytotics of Moko bacterial wilt in Honduras and along the Magdalena Valley in Colombia, where the disease spread over 90 km within 2 years, and also sporadic outbreaks in previously unaffected dessert banana plantations (Buddenhagen and Elsasser, 1962). Similar insect-borne

epidemics of Moko bacterial wilt have been described in Peru (French and Sequeira, 1970) and Nicaragua (Lehmann-Danziger, 1987). Insect transmission most probably accounts for the extensive spread of bugtok throughout central and southern Philippines since the first reports in 1965. Host reaction Following entry via wounds or natural openings, bacteria may initially multiply in the intercellular spaces (apoplast) before colonizing xylem vessels, from where they progressively invade the vascular system and move into surrounding parenchymatous tissues (Kelman, 1953; Vasse et al., 1995). The state of knowledge on mechanisms of pathogenicity and host specificity of the RSSC has been extensively reviewed (Denny, 2006; Genin and Denny, 2012; Ailloud et al., 2015). Resistance and escape All clones of commercial importance are considered susceptible to wound inoculation with B and SFR strains of Moko bacterial wilt, but differences in speed and severity of symptom development have been noted and some banana clones frequently produce apparently healthy suckers from infected mats. Stover (1972) reported artificial inoculation tests in Honduras carried out by injecting ‘B’ and ‘SFR’ strains of R. solanacearum into pseudostems of 345 accessions of banana. About 10% showed some degree of resistance to the SFR strain, with the ABB clone ‘Pelipita’, accessions of Musa balbisiana and the ‘Manang’ accession of M. acuminata ssp. banksii showing high to moderate resistance, and with others, including ‘Sucrier’ (AA), ‘Inarnibal’ (AA), ‘Laknau’ (AAB), ‘Saba’ (ABB) and ‘Pitogo’ (ABB), rated as susceptible. Only M. balbisiana showed resistance to the ‘B’ strain. The majority of 31 diploid (AA) genotypes evaluated in Brazil were susceptible to Moko bacterial wilt, but hybrids designated as ‘F2P2’, ‘1319-01’, ‘1741-01’ and ‘SH-3362’ along with ‘Babi ­Yadefana’, a cultivar from New Guinea, showed some resistance (Silva et al., 2000). A bred tetraploid ‘FHIA-03’ (ABBB) was found to remain healthy after planting in Moko bacterial wilt-infested areas of Grenada (Rowe



Diseases Caused by Bacteria and Phytoplasmas

and Rosales, 1999). However, bacteria injected into ‘resistant’ plants in the Honduran breeding programme could survive for up to 6 months without causing symptoms. Such tolerance to infection may be considered to be an undesirable trait for breeding programmes, because symptomless plants can serve as a hidden source of inoculum in planting materials, but can play an important role in disease management. In bugtok disease, the flowers and fruits of the commonly planted ABB clones such as ‘Saba’ and ‘Cardaba’ are destroyed following insect infection, but the survival and propagation potential of the corm and daughter suckers is usually unaffected. New suckers will grow and produce healthy fruit provided that steps are taken to prevent further insect infection. Other ABB or ABBB clones in the germplasm collection at the Davao National Crops Research and Development Centre in Mindanao reportedly behaved in the same way (Soguilon et al., 1995), indicating that tolerance is linked to the possession of the ‘BB’ genome. As noted above, similar incomplete systemicity has been reported for ABB cultivars affected by Moko bacterial wilt. Cultivars with inflorescence characteristics that are inimical to insect transmission may ­escape infection altogether. The ABB cultivar ‘Pelipita’ was promoted widely in Central ­America as substitute for ‘Bluggoe’ when it was recognized that it escaped infection because of its persistent male bracts and flowers (Stover and Richardson, 1968). ‘Dwarf Cavendish’ types, which tend to retain male flowers and bracts, are less susceptible to inflorescence infection to Moko bacterial wilt than were the ‘Gros Michel’ plantings originally affected in Central America. Most cultivars in the Plantain subgroup (AAB) have male flower parts that are either not shed, are degenerate or are sometimes completely absent (ProMusa, 2016). The non-susceptibility of these types to insect transmission in the field has been recognized to have preventive value against Moko bacterial wilt in South America (Lozano et al., 1969). These features underline the importance of considering different routes of infection when screening for resistance that may prove useful in the field. Mechanical inoculation by injection of bacteria into pseudostem or leaf bases is a convenient technique for screening large numbers of plants (Tripathi et al., 2008), but may be

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­ nnaturally severe, especially if high doses of u inoculum are used, and will reveal nothing about differences in susceptibility to infection via ­inflorescences.

Control When the Xanthomonas bacterial wilt epidemic started in banana in East Africa around 2001, most of the management options were inferred from ‘historic’ studies on Moko bacterial wilt. The situation is now reversed, with lessons for the epidemiology and management of Moko bacterial wilt (at least under smallholder conditions) to be learnt from the body of knowledge on Xanthomonas bacterial wilt that has emerged from East Africa. Control strategies relate to the host cultivar, predominant cultivation practices and the way these interact with the ecology and epidemiology of different strains of the pathogen. In commercial plantations of dessert (AAA) cultivars developed in Central America in the 1950s, pruning, de-suckering and de-budding became routine agronomic practices and this potentiated the spread of B strains of Moko bacterial wilt on cultivation tools and contaminated planting materials. Once these factors were understood, Moko bacterial wilt was successfully brought under control by the early 1960s. It generally remains of minor concern under plantation conditions, though continued and costly vigilance is necessary. In contrast, it can be difficult to gain collective or community-level adoption in smallholder and domestic plantings for timely de-budding and the precautions necessary to prevent spread on cutting tools or implanting materials. Exclusion Moko bacterial wilt is recognized as an important threat to banana cultivation by the Asian and Pacific Plant Protection Commission, the Caribbean Plant Protection Commission, the Pacific Plant Protection Organization and China, and should be excluded from any areas free from of the disease where Musa is grown for local consumption or as a cash crop. Importation of Musa planting material into an area considered to be vulnerable should only be undertaken after a

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thorough assessment of the risks (Diekmann and Putter, 1996). The unregulated movement of banana seed pieces and ornamental heliconia are particular hazardous. Strategies used to ­successfully eradicate R. solanacearum race 2 in ­Australia, following introduction of the disease in heliconia from Hawaii, have been described by Hyde et al. (1992). There are well-documented examples of the introduction of Moko bacterial wilt on infected banana planting material within Central and South America (Buddenhagen, 1961; Lehmann-­ Danziger, 1987) and to the Philippines (Buddenhagen, 1986). Recent spread of Moko bacterial wilt to previously unaffected areas in Ecuador (Delgado et al., 2014), Malaysia (Zulperi and ­Sijam, 2014; Teng et al., 2016), Jamaica (EPPO, 2004) and St Vincent (EPPO, 2007) emphasize the need both for continued vigilance to prevent the introduction of sources of infection and for prompt containment actions to restrict spread when this occurs. Bacteria can also colonize mature fruit (Soguilon et al., 1994b; Hadiwiyono, 2011) so the commercial trade in dessert bananas could provide a pathway for transnational spread of bacteria. An exhaustive analysis concluded that importation of dessert banana fruit from Moko-infested areas in the Philippines posed a significant risk of infection to the ­Australian domestic banana industry, though this could be mitigated by selecting fruit from areas with very low levels of infection (Biosecurity Australia, 2008a). Species of Heliconia may also be hosts of Moko bacterial wilt. Imported heliconias should be quarantined for a lengthy period (see Chapter 13). Eradication Where the pathogen is present, control strategies centre on the prompt detection, isolation and eradication of sources of infection, and adoption of integrated management practices to limit pathogen dispersal, to eradicate sources of infection and to rehabilitate diseased areas. In commercial plantations at risk of Moko bacterial wilt, experienced scouts regularly inspect banana plants for early disease symptoms every 2–4 weeks in advance of pruning operations. Diseased plants together with a ‘buffer zone’ of apparently healthy plants are then destroyed in situ by injections with a systemic herbicide, such as  glyphosate or 2,4-D. To reduce the risk of

t­oppling over, any fruit bunches are cut down, sealed in plastic bags and left to rot on site, with a plastic bag placed over the cut peduncle to exclude insects that might spread the pathogen. Based on early field studies (Sequeira, 1962) and experience under plantation conditions (Stover, 1972), buffer zones of 5 m and 10 m, and fallow periods of 6 months and 12 months, have been recommended for SFR and B strain infections, respectively. This area may either be left fallow or planted with wilt-suppressive crops, such as maize. Where weeds are effectively controlled, the fallow period may be considerably reduced. Lehmann-Danziger (1987) reported that a weed-free period of only 5 months was necessary for the control of Moko bacterial wilt in ­Nicaragua, but a 2-year fallow period was recommended in Grenada, where weed control was less effective (Hunt 1987). Early studies by Sequeira (1958) investigated treatment of infected soil with bactericidal chemicals, but these did not reduce disease incidence in subsequent crops. However, methyl bromide fumigation of soil and diseased residues within the buffer zone has been widely used to reduce replanting time to less than 4 weeks (Stover and Simmonds, 1987). Dazomet and metham sodium have been proposed as more environmentally acceptable substitutes (Castaneda et al., 2002; UNEP, 2003; Martinez and Guzmán, 2011). Plant ‘lixiviates’ (soluble extracts prepared from decomposed banana plant residues), incorporation of French marigold (Tagates patula) and fertilizers have been advocated as alternative soil treatments to reduce R. solanacearum populations and to improve crop nutrition (Alvarez et al., 2015). In the Philippines, rice hulls are burnt over affected mats to reduce soil infection (Natural, 2011; Biosecurity Australia, 2008b). In Belize, systematic surveys of ‘Bluggoe’ and smallholder dessert-banana cultivars, coupled with glyphosate treatment of all banana plants within a 5 m radius, effectively eradicated Moko bacterial wilt and the only subsequent outbreaks arose from possible incursions of the disease over the border from Guatemala. Eradication was achieved with the cooperation of the public and without the need for compensation for healthy plants (mostly ‘Bluggoe’) destroyed (Black and Delbeke, 1991). In the eradication programme in Grenada, compensation was ­given for healthy mats of ‘Bluggoe’



Diseases Caused by Bacteria and Phytoplasmas

­ estroyed, as well as for dessert bananas, as the d fruit was highly valued as a food (Hunt, 1987). Integrated management It has been estimated that, without sufficient precautions against contamination, almost 97% of the dissemination of B strains of Moko bacterial wilt may be due to cultural practices alone (Wardlaw, 1972). Disinfection of digging and cutting tools is, therefore, of great importance in disease control, and sterilization systems suitable for routine field use have been devised (Buddenhagen and Sequeira, 1958; Sequeira, 1958; Stover, 1972). Plantation workers undertaking pruning operations usually carry two knives. One knife is sterilized in a scabbard lined with felt soaked in disinfectant while the other is in use. Tools need to be sterilized between each plant and a dye, such as crystal violet, can be added to the disinfectant in order to monitor the operation. The same sterilizing solution is used to treat the shoes of workers when they leave infested areas. Traditionally, knives have been disinfected with a 10% solution of formaldehyde for 10 s or a 5% solution for 30 s. ­Sodium hypochlorite has become a widely recommended substitute for formalin (Alvarez et al., 2015), but is also toxic, corrosive and relatively unstable. Suggested alternatives include iodophors, such as Vanodine® (Garnica, 1998), creosote oils (Vargas, 2003) and quaternary ammonium products (Martinez and Guzmán, 2011; Philippine National Standard, 2013), such as those marketed in Honduras as Desiquat® and Banaril®. As noted above, the role of weeds in promoting survival and persistence of Moko bacterial wilt remains somewhat controversial, but elimination of potential weed hosts within the buffer zone is considered an important factor in its eradication, as R. solanacearum survives less well in soils where no natural hosts are present. Whilst eradication may be a priority for new outbreaks or where there is a perceived threat of soilborne infection to commercial dessert plantings, it is by no means clear whether this is either necessary or desirable where the main problem is inflorescence infection of ABB types, in which bacterial invasion may be incompletely systemic and infection may not be perennated in daughter suckers. In these cases, the prompt removal

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of only diseased stems may be more appropriate, as recommended for Xanthomonas bacterial wilt on banana, integrated with a rigid programme of breaking off the male bud by hand or by means of a forked stick as soon as the last female fingers have emerged. This is a standard recommendation for all cultivars with dehiscent bracts (Stover, 1972). Timely de-budding, together with appropriate field sanitation measures to reduce the possibilities of mechanical transmission from disease plant residues, have been shown to be highly effective for management of bugtok (Molina, 2006) and are advocated for other banana diseases in Southeast Asia. These practices are considered to have reduced the status of bugtok in the Philippines to that of a minor and sporadic problem (A.B. Molina, Philippines, 2016, personal communication). The introduction of budless mutants described above may provide a long-term solution to the problem of managing insect-transmitted infection, without the need for continuous interventions. With careful attention to ensure that mother plants are free from infection, micropropagation from axillary buds may be particularly appropriate for community or village-level production of large numbers of plants whilst minimizing the risks of infection during plant production (Staver and Lescot, 2015). Other measures that have been proposed to reduce or ameliorate infection include application of table salt (Pava et al., 2003; Talip et al., 2013), microbial antagonists (Ceballos et al., 2014) and resistance inducers (Ramirez et al., 2015). Further carefully controlled field research is needed to investigate and validate these for management of bacterial wilts of banana.

Blood Bacterial Wilt S.J. Eden-Green Introduction The history and biology of blood bacterial wilt in Southeast Asia share striking similarities to those of Moko bacterial wilt in Central and South America (Buddenhagen, 2009). Blood bacterial wilt was originally reported when intensive ­banana cultivation was introduced to two small

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offshore islands in the Selayar archipelago to the south east of the Indonesian island of Sulawesi (formerly Celebes) (Rijks, 1916). In subsequent investigations, Gäumann (1921a, 1923) found the disease to be widespread in cultivated banana in southern Sulawesi and he also observed symptoms in wild (forest) banana species. At some locations, farmers reported historical epidemics (‘waves’ of disease) going back for many years, and had coined the local name penyakit darah on account of the reddish ‘blood-like’ bacterial ooze that emerges from cut vascular tissues. The disease apparently remained confined to Sulawesi until the late 1980s, when an outbreak was identified in West Java (Eden-Green and Sastraatmadja, 1990). Since then it has spread across the Indonesian archipelago from Sumatra in the west to Papua in the east (Davis et al., 2000), a distance of over 4000 km. Blood bacterial wilt has recently been reported from peninsula Malaysia (Timin et al., 2014; Teng et al., 2016; Badrun et al., 2017), but others have reported the presence of Moko bacterial wilt (Zulperi and Sijam, 2014; Zulperi et al., 2016). It is still unclear which, or whether both, of the pathogens, which cause similar symptoms, have become established. However, its appearance on mainland Southeast Asia threatens banana cultivation in nearby countries, such as Thailand and Myanmar. Symptoms Symptoms of blood bacterial wilt resemble those of Moko bacterial wilt and, as with Moko bacterial wilt, the route and mode of infection influence the sequence and severity of symptoms. Thus early symptoms in flowers and immature fruits indicate ‘top-down’ infection, by insect transmission to male flower parts or via contaminated tools used for removing male buds. Yellowing or flaccidity in older leaves, preceding the development of inflorescence symptoms or in pre-­ ­flowering plants, indicates ‘bottom-up’ infection via mature leaves, roots or rhizome (Plate 5.19). The disease affects both dessert and cooking-­ banana cultivars. The latter are more widely grown in Sulawesi, perhaps due to the higher susceptibility of dessert varieties, and the symptoms of blood disease are described from this ABB group. In mature leaves, the disease causes

Plate 5.19.  Withering and collapse of leaves caused by blood bacterial wilt on ‘Pisang Kepok’ (ABB, Saba subgroup) in North Sulawesi, Indonesia. The unaffected male bud indicates a ‘bottom-up’ infection (photo: S. Eden-Green, NRI).

a conspicuous yellowing, which is followed by wilting, necrosis and collapse near the junction with the pseudostem. Younger leaves turn bright yellow, before becoming necrotic and dry, and the emergence of the youngest leaf is arrested. The male flower bud and youngest fruit hands may be blackened and shrivelled (Plate 5.20), though often they appear outwardly unaffected. However, internally the fruit exhibit a reddish-­ brown discoloration and are rotten or dry (Plate  5.21). The name ‘blood disease’ derives from the reddish discoloration visible in the vascular tissues of infected plants. This discoloration extends from affected fruit bunches down into the corm via the peduncle and pseudostem (Plate 5.22). Cut vascular tissue exudes droplets of bacterial ooze, which may vary in colour from cream to reddish brown to black. The vascular discoloration may extend through the mat and into the suckers, though this is not always the



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case and uninfected suckers are sometimes produced that can perpetuate infection (Setyobudi and Hermanto, 2000).

Causal agent

Plate 5.20.  Shrivelling and blackening of the male flower bud and lower hands on a bunch of ‘Pisang Kepok’ (ABB, Saba subgroup) affected by blood ­bacterial wilt in West Java, Indonesia, indicates a ‘topdown’ infection (photo: S. Eden-Green, NRI).

Plate 5.21.  Internal rot of fruit of ‘Pisang Kepok’ (ABB, Saba subgroup) affected by blood bacterial wilt in West Java, Indonesia. Note that no symptoms appear to be visible on the outside of the fruit (photo: S. Eden-Green, NRI).

Plate 5.22.  Brown discoloration of the vascular tissues in the pseudostem of ‘Pisang Kepok’ (ABB, Saba subgroup) affected by blood bacterial wilt in West Java, Indonesia (photo: S. Eden-Green, NRI).

The pathogen causing blood bacterial wilt also belongs to the Ralstonia solanacearum species complex (RSSC) (Allen et al., 2005). Gäumann (1921a) showed the causal agent of blood bacterial wilt was a Gram-negative bacterium, which he named Pseudomonas celebensis. Confusingly, this species was briefly reclassified as Xanthomonas, along with other non-fluorescent pseudomonads (Dowson, 1943). The epithet ‘celebensis’ was then used for another organism and the specific binomial is no longer valid, but has recently been acknowledged in a proposed new taxonomic designation of the pathogen as R. syzygii ssp. celebesensis ssp. nov. (Safni et al., 2014). Little variability has been demonstrated amongst isolates of R. syzygii ssp. celebesensis. All isolates examined by Eden-Green (1994) utilized galactose and glycerol, but not glucose, fructose or sucrose on first isolation. Thus blood bacterial wilt bacteria grow better in culture media if galactose or glycerol are substituted for glucose or sucrose. Inclusion of crystal violet (2.5 μg/ml) and polymyxin B (100 μg/ml), as used for R. syzygii ssp. syzygii (Eden-Green et al., 1992), inhibits many faster-growing contaminating organisms and is useful in selective media for initial isolation. None of the isolates examined by Eden-Green (1994) produced wilt symptoms when inoculated into tomato, eggplant, sweet pepper, groundnut or tobacco, though a hypersensitive reaction was induced in tobacco leaves. In a preliminary test with one isolate, rapid wilting developed after inoculation into ginger (Zingiber officinale) (author’s unpublished results). This contrasted with R. solanacearum strains from Moko bacterial wilt and bugtok ­affected plants, all of which induced wilt symptoms in tomato as well as banana. Despite these phenotypic differences, close similarities were demonstrated between bacteria causing Moko and blood bacterial wilts and Sumatra disease of clove in Indonesia caused by R. syzygii (Roberts et al., 1990) in several serological (Eden-Green et  al., 1988; Robinson, 1993) and molecular

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studies (Thwaites et al., 1999a; Seal et al., 1992, 1993; Taghavi et al., 1996). The phylogenetic relationships of banana pathogens within the RSSC have been rationalized into four major phylogenetic subdivisions (phylotypes I–IV), based on analyses of sequence data from the 16s-23s rDNA interspacial region (Prior and Fegan, 2005a, b; Fegan, 2005; Fegan and Prior, 2005, 2006) and the endoglucanase (egl) gene (Remenant et al., 2011; Genin and Denny, 2012). These phylotypes are consistent with earlier multi-location genotype (MLG) c­ lusters from whole-genome RFLP analyses (Cook et al., 1989), which placed the blood bacterial wilt bacterium in a novel MLG (Cook and Sequeira, 1994), and they largely correlate with the geographical distribution and likely evolutionary origins of different strains. Under this scheme, the blood bacterial wilt pathogen clustered into Phylotype IV together with the closely related taxon R. syzygii (causing Sumatra disease of clove). These findings led Safni et al. (2014) to propose the designation of bacteria in Phylotype IV as R. syzygii, comprising three subspecies: ssp. syzygii (Sumatra disease of clove pathogen), ssp. celebesensis (blood bacterial wilt pathogen) and ssp. indonesiensis for other broad host range strains of mainly Indonesian origin. The lack of genetic diversity among isolates of the blood bacterial wilt pathogen indicates a recent evolutionary history on banana. However, the evolutionary origins of blood bacterial wilt remains an open question. As Buddenhagen (2009) pointed out, the first reports of the disease came from two small islands in the Selayar group following the introduction and establishment in the late 19th century of intensive cultivations of ‘Bluggoe’ for fruit export to mainland Sulawesi. These plantings would have presented numerous infection courts susceptible to infection by insects (and which could possibly have been initiated by migrating bats and birds), setting the scene for the explosive disease epidemics that were first noticed in 1905. However, in the intermediate 20–30 years, the plantings on these islands were apparently healthy, indicating that the disease may have been introduced from other islands either by airborne vectors or in diseased plants. No other similar instances of disease were reported by agricultural officers either from the diverse cultivations of domesticated banana clones or from wild banana species still

widely growing throughout the region. From these observations, and the lack of genetic variability so far observed within R. syzygii ssp. celebesesis, Buddenhagen concluded that the bacterium did not coevolve in banana (Buddenhagen, 2009). Whether or not blood bacterial wilt evolved in Musa, which has a centre of diversity in Indonesia, seems no more nor less likely than whether strains of Moko bacterial wilt evolved in Heliconia, which has its centre of diversity in the New World. Gäumann (1921a) reported observing a cluster of unhealthy-looking wild banana plants on the edge of a forest path in Sulawesi, some of which he showed to be infected with blood bacterial wilt. He also noted that local inhabitants had a long familiarity with the disease and had developed local remedies for dealing with it. From this, he concluded that the disease had probably been present on the main island of Sulawesi before the outbreaks occurred in the Selayar islands. With Sumatra disease of clove, a strong epidemiological association was observed between geographically isolated disease outbreaks and intensive new plantings of clove trees close to natural forest margins in Sumatra and Java (Lomer et al., 1992). The disease has not been found at the centre of origin of clove in the Moluccan Islands, nor has it reliably been reported from Sulawesi. Although natural hosts of R. syzygii ssp. sygygii have yet to be found, closely related myrtaceous species have been shown to be susceptible to experimental infection and to be hosts of insects that transmit the disease. Some of these species form part of the indigenous forest flora of Sumatra and Java. So in this case, it seems reasonable to assume that the clove pathogen may have evolved in closely related indigenous species together with the highly specialized mode of transmission by xylem-­ feeding Homoptera. It is possible that the blood bacterial wilt pathogen evolved to attack banana in a similar manner. Disease cycle and epidemiology Gäumann (1921a) considered it likely that blood bacterial wilt bacteria could survive in soil for at least a year in infested plant residues. However, experimental studies have indicated that



Diseases Caused by Bacteria and Phytoplasmas

R. syzygii ssp. celebesensis does not survive longer than 6 months when added to soils either from pure cultures or in diseased plant residues (Subandiyah, 2011). Tissue cultured plantlets (Subandiyah et al., 2006) and also ‘seedlings’ (presumably suckers) (Hadiwiyono et al., 2007) of ‘Pisang Kepok’ (ABB) have been infected in soils artifically infested with R. syzygii ssp. celebesensis, with and without wounding. Insect-borne epidemics spread very rapidly and probably account for the historic accounts of devastating losses of blood bacterial wilt in Sulawesi and neighbouring islands reported by Gäumann (1921a, 1923). Insect transmission also most probably accounts the rapid spread of blood bacterial wilt throughout the Indonesian archipelago since first reported outside of Sulawesi in 1986 (Supriadi, 2005). Buddenhagen (2009) observed various large wasps, Trigona bees, Oncopsia and flies in contact with ooze discharging from buds and peduncles of affected plants. Insects were seen feeding on fresh cushions; and cushions of bagged plants could be infected with fresh bacterial ooze. Similar groups of insects have been implicated as vectors by other authors (Leiwkabessy, 1999, cited in Supriadi, 2005; Mairawita et al., 2012; Montong et al., 2015). Host reaction All clones of commercial importance are considered susceptible to wound inoculation with the blood bacterial wilt pathogen, but differences in speed and severity of symptom development have been noted and some banana clones frequently produce apparently healthy suckers from infected mats. Gäumann (1921a) observed no resistance to blood bacterial wilt in collections of 100 banana cultivars in Sulawesi. Of 36 Musa cultivars tested by Sudana et al. (1999), cited in Supriadi (2005), ‘Pisang Batu’ (‘Pisang Klutuk’), ‘Pisang Bancan’, ‘Pisang Bunting’, ‘Pisang Bojong’, ‘Pisang DakNangka’, ‘Pisang Kayu’, ‘Pisang Ketip’, ‘Pisang Marga’, ‘Pisang Kepet’, ‘Pisang Muli’, ‘Pisang Papan’, ‘Pisang Rempeneng’, ‘Pisang Susu Ketan’, ‘Pisang Susu Plepeden’, ‘Pisang Telu’ (‘Pisang Telur’) and ‘Pisang Udang’ showed tolerance to blood bacterial wilt. As with Moko bacterial wilt, incomplete systemicity in ABB cultivars is linked to the possession of the ‘BB’

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genome. Cooking-banana cultivars with dehiscent male flowers such as ‘Pisang Kepok’ (ABB) and ‘Pisang Awak’ (ABB) are considered to be particularly susceptible to inflorescence infection by blood bacterial wilt and this has been attributed to the relatively high sugar content of nectar and to possible chemical attractiveness of flowers (Setyobudi and Hermanto, 2000; Buddenhagen, 2009). The male flower parts of cultivars in the Plantain subgroup (AAB) are either not shed, are degenerate or are sometimes completely ­absent (ProMusa, 2016). The non-susceptibility of these types to insect transmission has been recognized to have preventive value against ­ blood bacterial wilt in Indonesia (Setyobudi and Hermanto, 2000). A budless mutant of ‘Pisang Kepok’ (ABB) called ‘Pisang Puju’ that escaped insect infection because of a lack of an infection court was found on Sulawesi in 1991. This clone was collected, propagated and distributed as a goodwill gesture by Chiquita® when it had operations in Indonesia (Buddenhagen, 2009). Other budless mutants of ‘Pisang Kepok’ with different bunch characteristics were collected in Sulawesi in 2005–2007. Two with large bunches of fruit with superior flavours and cooking charcateristics were selected for mass propagation. Budless mutants of the ABB clones similar to ‘Cardaba’ and ‘Bluggoe’ have also been found (Buddenhagen, 2009). In addition, budless mutants of ‘Pisang Loka Bule’ and ‘Pisang Loka Nipah’ (now known as ‘Pisang Unti Sayang’) (Suhartanto et al., 2009) from Sulawesi and neighbouring islands plus ‘Pisang Sepatu Amora’ (now known as ‘Pisang Tanjung’) (Hermanto et al., 2013) from Seram in the Moluccas have been released for growers. Control Control measures are similar to those reported for Xanthomonas and Moko bacterial wilts. A 2-year delay before replanting blood disease-­infested areas was suggested by Gäumann (1923), but little experimental information has been published to show that this is necessary. Survival studies reported above indicate that R. syzygii ssp. celebesensis survives less than 6 months in soils and plant residues and that soilborne infection is not of significant epidemiological

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i­mportance for management of either blood bacterial disease or bugtok. The use of microbial antagonists has been proposed to reduce or ameliorate infection (Marwan et al., 2011; Latupeirissa et al., 2014) but further research is needed to investigate and validate this and other control measures in the field.

Acknowledgements I thank A. ‘Gus’ B. Molina for helpful comments and advice on bacterial wilts in Southeast Asia and Nienke Eernisse for help with Dutch ­translation.

Abacá Bacterial Wilt D.R. Jones and S.J. Eden-Green A disease of abacá, referred to variously as ‘abacá wilt’, ‘vascular disease’ and ‘banana wiltlike disease’, was reported by Palo and Calinisan (1939) to be the most serious disease of abacá in the Davao region of Mindanao in the ­Philippines, where over 1300 ha were found to be ­infected in 1937–1938. Infestations of weevil borer and stem weevil were associated with diseased plants and believed to play a role in infection. Several abacá cultivars were affected by the disease, principally ‘Maguindanao’, ‘Tañgoñgon’, ‘Balindag’ and ‘Bongolanon’. However, the wilting symptom was not observed in ‘Tañgoñgon’. The symptoms of the disease were very similar to those of Moko bacterial wilt except for the appearance of distinctive rusty-brown linear streaks along the leaf veins of infected plants. These streaks could extend from the midrib to the leaf margin. This symptom was followed by a yellowing of the tissues along the veins and ultimately a browning and wilting. In cases of acute infection, affected plants wilted rapidly without showing linear streaks. Other external symptoms were a conspicuous retardation of growth of the central leaf, narrowing of the leaves (most common in the Lauan–Tañgoñgon cultivar) and the appearance of large blackish-brown patches of rotting tissue at the base of the pseudostem. An examination of sections of the rhizome and pseudostem showed that the vascular strands of

affected plants were discoloured, the spread of the infection being easily traced up to the petioles and midribs and out into the leaf veins. No symptoms were reported on flowers and fruit. The disease had previously been considered to be fungal in origin, but Palo and Calinisan (1939) were consistently able to isolate bacteria with characteristics similar to Ralstonia solanacearum from diseased tissue. These isolates induced wilt in abacá plants when introduced into the centre of the pseudostem. As symptoms of bugtok disease, which indicated the arrival of R.  solanacearum race 2 in the Philippines, were not observed until the 1950s (Soguilon et al., 1994a), the 1937–1938 problem may have been caused by R. solanacearum race 1. Zehr (1970) inconsistently isolated R. solanacearum from abacá and banana plants with symptoms of vascular wilt, but the isolates seldom induced severe wilt symptoms when inoculated back into their respective hosts, though they were highly virulent to tomato, potato, pepper and eggplant. Abacá and banana also developed inconsistent and relatively mild disease symptoms after inoculation with isolates from solanaceous hosts. Field observations indicated to the author that bacterial wilts of abacá and banana were relatively rare and that strains of high virulence to these crops probably did not exist in the Philippines. His results indicated that R. solanaceraum race 1 may still have been the main causal agent of abacá bacterial wilt in 1970 and that race 2 at this time was not widespread in abacá growing areas. Rillo (1979) found that virulent isolates of R. solanacearum from diseased ‘Giant Cavendish’ (AAA, Cavendish subgroup) in the southern ­Philippines infected abacá, banana and heliconia. Judging from Rillo’s results, these isolates would have been race 2. However, it is not kown if race 2 was found on abacá in the field at this time. Later, Rillo (1981) differentiated bacterial wilt organisms isolated from abacá and the commercial banana ‘Giant Cavendish’ in terms of host range, colony characteristics, melanin formation, carbon utilization, hypersensitive reaction on tobacco leaves and phage typing. The banana isolates were determined to belong to biovar 1 (typical of race 2) while the abacá isolates where reported to belong to biovars 2 and 3 (typical of races 1 and 3). Rillo (1982) then artificially inoculated a range of test plants with



Diseases Caused by Bacteria and Phytoplasmas

abacá isolates. He found that these infected potato, castor bean, eggplant, tomato and abacá, but not cultivated banana. Three other isolates only infected castor bean, eggplant and tomato, but not abacá or cultivated banana. When seedlings of the same test plants were planted in inoculated soil, only a few castor bean and eggplants became infected. Rillo (1982) believed that the causal organism for abacá bacterial wilt in the field was race 1 of R. solanacearum. Unfortunately, the results of pathogenicity tests undertaken with R. solanacearum isolates from abacá in the Philippines by Rillo (1981, 1982) were inconclusive. In addition, disease symptoms were not pronounced when abacá was reinoculated with abacá isolates. Although the abacá cultivar inoculated was ‘Tañgoñgon’, which does not develop wilt symptoms, the conclusions reached from the work would appear tentative. Nevertheless, the broad host range R. solanacearum race 1 is known to infect some diploid Musa spp. (Buddenhagen, 1962: Akiew, 1992) and it has been widely assumed that the outbreak of abacá bacterial wilt reported in the 1930s was most likely caused by race 1 (Buddenhagen, 1986; Hayward 1986). Abacá bacterial wilt was also reported in Peru (Revilla and Vargas, 1967). Here, the causal agent was determined to be the banana-­attacking strain (race 2) of R. solanacearum. Abacá was found to be resistant to the broad host range race 1 of R. solanacearum. These results suggest that R. solanaceraum race 2 was responsible for abacá bacterial wilt in Peru.

Javanese Vascular Disease D.R. Jones According to Gäumann (1921b), this disease was first observed in 1915 and was subsequently found on almost all cultivated bananas in Java, Indonesia. Affected plants exhibited widespread vascular discoloration, though the disease was not considered to be particularly harmful to its host and 90% of infected plants displayed no external symptoms whatsoever. Where external symptoms were visible, they resembled those of Fusarium wilt and, as many vascular diseases had been ascribed to fungal

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pathogens, Gäumann examined six species of Fusarium, which were isolated from diseased banana plants. None of these fungi was pathogenic to banana, but a bacterium was identified that caused vascular discoloration when introduced into the rhizome, pseudostem or leaf veins. The bacterium was not specific to banana and was also pathogenic on Ravenala, Strelitzia and possibly Heliconia spp. Gäumann named the bacterium Pseudomonas musae, but the name is no longer valid and none of the original isolates survives. The bacterium was described as a Gram-positive, non-spore-forming rod, 0.8–1.2 μm long with one to three polar flagella. The original 1921 study asserted that external symptoms of Javanese vascular disease were caused by secondary infection by Fusarium spp. following primary vascular invasion by P. musae. Gäumann suggested that this was true also of Fusarium wilt and, in so doing, criticized the work of several of his contemporaries. Wardlaw’s (1935) account of the disease detailed the debate that ensued over the role of bacteria in such infections. No subsequent observations of Javanese vascular disease have been made and the original reports have been ascribed to Fusarium wilt.

Rhizome and Pseudostem Bacterial Rots D.R. Jones Overview In the past, different types of bacterial corm and pseudostem rot diseases were distinguished on the part or parts of the plant affected, cultivars affected, the age of plant affected and whether or not suckers became infected (Stover, 1972). Hence there were diseases known as (i) head rot, which seemed limited to the corm and pseudostem at ground level; (ii) rot of the rhizome and pseudostem, which seemed to infect the corm and extend up the pseudostem; and (iii) pseudostem wet rot, which seemed only to affect the pseudostem. The changes in the taxonomy of species that are the causal agents has also confused an already complex problem.

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In the descriptions below, diseases caused by bacterial pathogens attacking the rhizome and/or the pseudostem have been accommodated under the headings used by Stover (1972). This has been thought appropriate because symptom expression may still indicate differences in the pathogens involved. However, associating a pathogen with a particular set of symptoms is difficult. It may only be accomplished if a range of banana and enset genotypes are inoculated, under standardized conditions, with representative genotypes of all known strains of banana and enset soft rot bacteria species in controlled environments and symptom expression development is studied over time. Even then, caution is needed when extrapolating from results obtained with ‘unnatural’ modes of infection, such as wound inoculation of internal tissues with large doses of bacteria, to an ability to cause disease under field conditions. Rhizome rots caused by bacteria were considered to be only a minor problem in the past (Stover, 1972), but recent reports from India and China suggest that they are of growing concern (Lin et al., 2010; Arun et al., 2012). As well as affecting banana in the field, soft rot bacteria have been reported as a major limiting factor in the weaning of micropropagated plantlets (Thangavelu, 2009). Contamination of in vitro cultures has caused big losses to commercial tissue culture laboratories.

Bacterial Rhizome Rot and Tip-Over Introduction A bacterial rot of the banana corm known as rhizome rot and tip-over, but also called head rot and snap-off, was first noted in Honduras in 1948 (Stover, 1959). It was the major cause of poor germination of newly planted rhizomes and severe infection also caused the pseudostem of more mature plants to break. It was reported to be mainly a disease of plants less than 3 years old (Stover, 1972). Stover (1959, 1972) believed that rhizome rot occurred wherever banana was grown in Central America and the Caribbean and was possibly the same disease. He reported that it was most serious on the alluvial soils of Central

America, particularly in areas where rainfall was high. In Honduras, it was frequently observed on ‘Gros Michel’ (AAA) in locations ­flood-­fallowed for 4–6 months. An incidence of 10–15% of mats was common, occasionally rising to 50% in some areas. Losses were greatest in the first 8–20 months after planting. Newly planted rhizomes and plants less than 6 months old were frequently destroyed. Incidence was lower in Cavendish cultivars, but up to 30% of newly planted corms rotted in wet, cool weather. Persistent centres of loss were present in plantations more than 5 years old (Stover, 1972). Stover (1959) also reported high levels of disease in young plants growing in methyl bromide or steam-sterilized soil in the greenhouse. In Jamaica, Shillingford (1974) reported that the pseudostem of Cavendish plants severely diseased by a bacterial soft rhizome rot broke at about soil level just before fruit maturity. Symptoms of a disease of banana that appeared in a new plantation of ‘Basrai’ (AAA, Dwarf Cavendish) in Uttar Pradesh in India in 1970 included damage to the rhizome and the breaking of the pseudostem at soil level (Edward et al., 1973). More recently, a serious collar and rhizome rot disease of banana was observed in the north region of Maharashtra state in India after the rainy season. The infection occurred on a month-old banana plantation in poorly drained soil, but plantations established for 8–10 weeks were also found severely infected (Nagrale et al., 2013). In nothern Karnataka in India, the disease was noticed in all the locations surveyed with incidence ranging from 4% to 65% (Vijayalaxmi et al., 2014). Arun et al. (2012) reported that rhizome rot has been reported from the Indian states of Kerala, Gujarat, Tamil Nadu, Bihar and Karnataka and, apart from affecting banana fields, it  had also been found to be a major limiting ­factor in weaning of micropropagated plantlets (Thangavelu, 2009). The extent of deaths due to soft rot in secondary hardening was reported to be around 2–5% (Thomas et al., 2011). Furthermore, the pathogen was known to contaminate in vitro cultures. A bacterial corm rot has been reported affecting enset in Ethiopia, killing both young and mature plants. Severity of the disease has been said to be high in the lowlands and mid-altitude locations up to 1800 m. Infection is believed to



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occur through damaged roots, the disease then spreading into the corm (Tessera and Quimio, 1994). Symptoms In the field, symptoms occurred at three stages of growth. Firstly, newly planted rhizomes failed to sprout. Secondly, young plants exhibited yellowing and stunted growth. Thirdly, mature plants with severe decay of the rhizome toppled over because of the wind and weight of the fruit (Stover, 1972). Pockets of dark brown or yellow water-­ soaked areas with dark peripheral rings were seen in abundance in the cortex of diseased rhizomes, but occurred throughout the rhizome when in an advanced stage. Here, cavities surrounded by dark, spongy tissue later developed. These cavities could be confused with tunnels caused by burrowing nematodes. Hollow pockets of rot in the outer cortex were colonized by ants (Stover, 1972). In potted plants, the rot often begins in the cortex in association with old leaf sheath remnants attached to the corm from the central growing point. A new rhizome of a young plant usually grows away from the rot in the seed rhizome and remains healthy. However, a dark brown necrosis is sometimes seen in the lamina of older leaves. Stover (1972) believed this may have been the result of a toxin passing into the new plant from the decay in the seed rhizome. Infected young plants became stunted and yellowed as the rot progressed and could be easily pushed over (Stover, 1972). In some cases, the rhizomes of young suckers emerging from an infected parent rhizome became infected, though suckers frequently remained healthy and roots were not attacked. Stover (1972) commented that it was remarkable that a large bunch of fruit could be produced from a rhizome riddled with rot. Plate 5.23 shows a bacterial rot of a rhizome where it joins the psuedostem. External symptoms were seldom seen in affected mature plants, but they were easily blown or pushed over. When the psedostem fell, some infected tissues remained attached to the corm, which split, leaving the rest anchored in the soil by its roots. Such plants are called tip-overs or snap-offs. Tip-overs caused by bacterial rhizome

Plate 5.23.  Bacterial rot of the rhizome seen at the  base of the pseudostem of a Cavendish cultivar in a plantation in central Cuba. This disease was most likely caused by a species of Dickeya (photo: D.R. Jones, INIBAP).

rot can be distinguished from those caused by the burrowing nematode in that, with the latter, the entire stool is pulled out of the ground when the plant falls, exposing the base of the rhizome and roots (Stover, 1972). The symptoms described by Shillingford (1974) were identical to those of Stover (1959, 1972). Rotting of newly planted rhizomes with failure to sprout, stunting and yellowing of young plants and toppling over of mature fruited plants were the major symptoms observed by Thammaiah et al. (2005) in India. Infected young plants became stunted and could easily be pulled up. The mature plants did not show yellowing and stunting, but were easily blown or toppled over and remnants of rhizomes remained attached with the pseudostem. Symptoms were commonly observed during hot summer months and also during the rainy season 3–5 months after planting. Planting infected suckers or

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corm pieces usually results in diseased plants, and their movement can spread inoculum to new ­areas (A.V. Sankar, India, 2017, personal communication). The bacterial corm rot of enset is reported to disintegrate the tissues in the corm, forming a hollow area, which predisposes the plant to toppling by the wind. Wilting of the older leaves is believed to be a distinguishing feature of the disease (Plate 5.24).

Causal agent Tissues in the early stages of decay always yielded bacteria, but no fungal mycelium (Hord and Flippin, 1953). The bacterium isolated from rhizome rot-affected Cavendish banana plants was identified as Pectobacterium carotovorum (syn. Erwinia carotovora) by Stover (1959, 1972). Stover (1959) reproduced the symptoms of rhizome rot

Plate 5.24.  Wilting and collapse of the older leaves of enset caused by bacterial corm rot in Ethiopia (photo: M. Tessera and A.J. Quimio).

when an agar plug containing inoculum was placed inside a banana rhizome. However, cultures of bacteria isolated from banana plants in Jamiaca with identical symptoms and sent to the UK for identification by Shillingord (1974) were found to be closest to Erwinia chrysanthemi. Both bacteria belong to Enterobacteriaceae (class Gammaproteobacteria) and are aerobic, Gram-­ negative rods with peritrichous flagella. A head rot of banana was described in Israel with E. carotovora as the causal agent (Volcani and Zutra, 1967). A survey in Cuba in August 1974 revealed a corm rot in banana caused by E. carotovora ssp. carotovora (Rivera, 1978). Later, a head rot caused by E. carotovora and associated with burrowing nematode was reported in the Philippines (San Juan, 1980). Snehalatharani and Khan (2010) identified bacterial isolates causing a serious tip-over disease of banana in southern India as E. carotovora spp. carotovora, E. chrysanthemi and a bacterium with intermediate characteristics. Erwinia chrysanthemi pv. paradisiaca has also been recorded as the causal agent. Two strains, one more virulent than the other, were believed involved (Nagrale et al., 2013). In Papua New Guinea, a serious rhizome rot caused by E. chrysanthemi, was isolated from diseased tissue and surrounding soil, but not from healthy plants or field soil. Erwinia carotovora ssp. carotovara was also isolated, but pathogenicity tests only indicated a minor role (Tomlinson et al., 1987). It is possible, because aberrant strains will almost certainly be found, that E. carotovora and E. chrysanthemi could have been confused by some authors in the past on the basis of traditional bacteriological tests (S. Eden-Green, UK, 2017, personal communication). Dickey (1979) compared 421 Erwinia isolates. Erwinia chrysanthemi isolates were separated from the other Erwinia species primarily by three physiological characters which were the production of gas from d-glucose, phosphatase production and inability to produce acid from d-trehalose. The 322 isolates of E. chrysanthemi were divided into five infrasubspecific subdivisions based on 12 physiological properties. All 12 isolates from banana (Cavendish cultivars) in Honduras, Panama and Jamaica were accommodated into the IV subdivision. Dickey (1979) seems to have included bacteria isolated by Stover in Honduras and Shillingford in Jamaica in his research.



Diseases Caused by Bacteria and Phytoplasmas

Erwini chrysanthemi is a motile, Gram-­ negative, non-sporing, straight rod with rounded ends and occurs singly or in pairs: it varies from 0.8–3.2 × 0.5–0.8 μm (average 1.8 × 0.6 μm); there are three to 14, but more usually eight to 11, peritrichous flagellae. It has now been placed in the genus Dickeya and subdivided into a number of species (Samson et al., 2005). The bacterium causing a soft rot of the banana rhizome disease which spread up the pseudostem in Guangzhou, China, was identified as a Dickeya sp. by Lin et al. (2010). Four Dickeya species have been isolated from banana. They are D. dieffenbachiae in Cuba, D. paradisiaca in Colombia and Cuba, D. zeae and D. dadantii in China and D. zeae and/or D. dadantii in Côte d’Ivoire, Panama, Jamiaca and Columbia (Samson et al., 2005; Zhang et al., 2014; Liu et al., 2016). The relatedness and clade structure of Dickeya species has been determined by comparison of recA sequences (Parkinson et al., 2009). The Dickeya species that causes the rhizome rot and tip-over diseases reported by Stover (1959, 1972) is as yet unkown. The causal agent of the bacterial corm rot of enset has not been identified (M. Tessra and A.J. Quimio, Ethiopia, 1999, personal communication). Disease cycle and epidemiology The bacterium undoubtedly spreads from infected to healthy rhizomes as a result of seed digging and planting operations, including peeling and heat treatment for nematode control. The causal bacteria can invade the rhizome directly after planting via wounds or decaying leaf sheaths around the central bud. It is likely that the causal agent (or agents) persists in the soil and can be spread by water. The disease is more common on banana and plantain in wet weather, particularly if the ground is poorly drained, cool, and when plants become waterlogged (Stover, 1972; Buddenhagen, 1994). The bacterium is unlikely to be transmitted as a latent pathogen in micropropagated banana (Thomas et al., 2011). Plants derived from tissue cultures have been shown to be susceptible to bacterial rot. During periods of hot weather in the summer in India, excessive heat is known to scorch tender roots, resulting in decay. Soil-borne, soft rot bacteria enter the plant through the wounded roots (A.V. Sankar, India, 2017, personal communication).

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In the Pacific, it is recommended that planting material be collected from only healthy stools. After removing suckers from each plant, knives and other tools should be disinfected by wiping with bleach. Organic matter in the plantation should be well decayed. In Australia, rhizome rot has occurred where banana has been planted into areas with undecomposed organic matter. Plants seen with either banana wilt or rhizome rot symptoms should be removed as soon as possible and burnt. Suckers can be left, as these may be healthy, but they need to be monitored for infection. Plants are more susceptible to rhizome rots when water-stressed. Hot and dry conditions followed by heavy rainfall can lead to outbreaks of infection. Drainage should be improved in plantations where water collects after rain (ACIAR, 2016).

Host reaction Stover (1972) reported that Cavendish cultivars were less susceptible than ‘Gros Michel’ and that, in general, AAB and ABB cultivars were much more resistant than AAA cultivars. This is the reason he gave for rhizome rot not being a problem in plantain plantations. Abacá was also not usually affected. Shillingford (1974) found that the Cavendish cultivars ‘Robusta’ and ‘Valery’ were severely affected, but ‘Lacatan’ was relatively unaffected. In Papua New Guinea, as in Honduras, the disease was most prevalent on experimental plantings of AA diploid cultivars and Cavendish cultivars (Tomlinson et al., 1987).

Control Head rot was believed by Stover (1972) to be only occasionally a severe problem and control measures were only deemed necessary if the risk of infection was high. He advised that if susceptible cultivars are planted in high-rainfall areas, then seed pieces should be carefully selected and inspected. Planting rhizomes during periods of heavy rain and in soil that is not well-drained should be avoided. Loos (1962) reported that disease losses were severe when rhizomes weighing less than 450 g were planted. He also found that peeling

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rhizomes for nematode control did not increase susceptibility to rot bacteria. Working with ‘Gros Michel’, he discovered that rot was negligible when plants were derived from the axillary buds, but severe when plants developed from the centre bud. The removal of the centre bud during paring operations to stimulate axillary buds was also recommended for nematode control. However, this practice increased germination losses of ‘Valery’ (AAA, Cavendish) in some areas and especially in the rainy season. Bacterial attacks  may be effectively controlled by maintaining sufficient moisture around the root zone of young plants derived from tissue culture during the beginning of summer to prevent heat injury to tender roots. Drip irrigation is recommended in the early morning or late evening, but not during the heat of the day. Good drainage should be provided where water stagnates in the field (A.V. Sankar, India, 2017, personal communication). As a rescue measure in India, the infected tissues of affected corms to be used as sources of planting material were scooped out until only healthy white rhizome tissues remained. Care was taken so that no rotten portion remained on the rhizome surface. Such shaved rhizomes were treated with 0.2% methoxy ethyl mercuric chloride solution for 5 min and planted in new pots containing a sterilized mixture of red soil and coir pith in equal proportions. The suckers had little or no roots and the size was reduced to a piece about 2.5 cm in thickness. To promote root formation and plant growth, plants were sprayed twice with mono ammonium phosphate (0.2 %) at weekly intervals. Plants were taken to the field after 45 days of treatment and rated for disease after 4 months. Depending on the cultivar, 82–89% survived to be disease-free (Arun et al., 2012). No effective chemical or biocontrol treatments to reduce incidence have been reported to have been adopted in the field.

Bacterial Soft Rot of Rhizome and Pseudostem Introduction Stover (1972) reported this disease as a minor problem in some areas of Costa Rica, Panama

and Colombia. He described it as a massive, ­ dorous rot of the centre or a portion of the o rhizome. It differed from bacterial rhizome rot described above in that it spread from the rhizome into the pseudostem, causing internal decay and usually vascular discoloration. The progression of this rot destroyed the growing point of the banana stem. External symptoms sometimes resemble those of Fusarium wilt. Bacteria predominated in the rot (Stover, 1972). In Cuba, plants developing from infected rhizomes were slow growing with chlorotic and flaccid leaves, as well as a rotting that spread upwards from the base of the plant to the rest of the pseudostem. These plants sometimes eventually collapsed and died (Rivera and Ezavin, 1980). In India, a soft rot of the banana rhizome that spreads into the pseudostem was described by Nagaraj et al. (2012). The disease, which causes tip-over, was reported as becoming a serious problem in all the banana-growing areas of Kamataka and Andhra Pradesh states, particularly in 2–6-month-old gardens planted with tissue culture plantlets of ‘G-9’ and ‘Robusta’ (AAA, Cavendish subgroup). Disease incidence ranged from 30% to 35% in the districts of Bangalore and Kolar of Karnataka state. Reports of bacterial soft rots of the banana rhizome spreading to the pseudostem have since come from China. It was first noticed in 2009 in Guangzhou city. A survey of three areas of ABB banana production covering 10 ha in Guangzhou revealed that 82% of the fields were affected by a soft rot with similar symptoms at an incidence ranging from 20% to 40% (Lin et al., 2010). The disease was characterized by an odorous soft rot of the centre of the rhizome. The rot progressed up the pseudostem, destroying the growing point and causing internal decay and often accompanied by vascular discoloration. Yellowing and wilting of the leaves were also characteristic symptoms (Lin et al., 2010). The same disease was observed on various banana cultivars of different genotypes in several other cities. Symptoms of the disease included leaf wilting, collapse of pseudostems, and an unusual odour (Zhang et  al., 2014). It is reported that bacterial soft rot is a relatively new banana disease to China and that it is causing significant economic losses.



Diseases Caused by Bacteria and Phytoplasmas

Symptoms The internal rot causes a yellowing and wilting of leaves. If the rot spreads rapidly, the plant dies with its brown, dead leaves remaining erect. The older leaves often fall while green, with the younger centre leaves turning yellow. The emerging heart leaf may be partly necrotic if the rot has extended a considerable distance up the pseudostem. Stover (1972) also observed that the disease usually affected only large, non-fruiting plants and did not invade daughter suckers. In India, the disease also caused rotting of the rhizome and pseudostem, following marginal necrosis or scorching of leaves. Ultimately, affected plants toppled (Nagaraj et al., 2012). In China, a soft rot of banana progresses up the pseudostem, destroying the growing point. This results in internal decay and is often accompanied by vascular discoloration. Yellowing and wilting of the leaves are also characteristic symptoms. An odour of decay was associated with the rot (Lin et al., 2010).

Causal agents A disease characterized by massive soft rot of the rhizome with a typical odour like that caused by Erwinia carotovora was noted in Brazil. The rot progressed up the pseudostem, destroying the growth point, causing internal decay and plant death. External symptoms consisted of yellowing, wilt and collapse of leaves. Pathogenicity tests with tomato plants and potato tubers also suggested that the pathogen was probably of the E. carotovora group (Pereira and Nunes, 1988). In India, nine isolates of bacteria from diseased plants were identified as Erwinia carotovora ssp. carotovora and Erwinia chrysanthemi on the basis of morphological, biochemical, physiological and pathogenicity tests (Nagaraj et al., 2012). Again, traditional biological tests used in the past may have led to some confusion when trying to distinguish E. carotovora and E. chrysanthemi. As mentioned previously, isolates from banana are now designated as Dickeya species. In China, the genome of a strain of Dickeya zeae causing a soft rot of banana that extends up the psedostem has been sequenced. Several of its

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specific properties were compared with those of other D. zeae strains (Zhang et al., 2013). Later, four bacterial isolates representative of a larger group of bacteria recovered from banana plants with soft rot from different regions of China were characterized by Liu et al. (2015). Results showed that the bacterial isolates from banana were Gram-negative, rod-shaped with multiple flagella, and produced a brown pigment on NGM medium (nutrient agar supplemented with 1% glycerol). Physiological and biochemical assays revealed that all four bacteria isolates were D.  zeae and differed from D. dieffenbachiae and D. paradisiaca.

Disease cycle and epidemiology The causal bacteria are probably common in the soil. Spread between plants probably occurs when bacteria are splashed from the soil into the ‘throat’ or funnel made by the leaves, perhaps when the plants are still young. It is thought likely that the disease occurs when water collects in the ‘throat’. Infections of the pseudostem are probably caused by wound invasion. As the disease has been observed to be more severe in the first ratoon crop, spread may occur when suckers are removed from the mother plant. Infection can also occur when the corm is trimmed with contaminated tools before planting (ACIAR, 2016).

Host reaction Yuan et al. (2013) reported a Dickeya sp. affecting banana in Guangdong and Hainan provinces in China. Three isolates of the banana soft rot pathogen were inoculated into 22 species of plants from 15 families. The results indicated that the three isolates had a wide host range. The reaction of seven banana cultivars widely grown in China were also investigated. The results showed that ‘Pisang Mas’ (AA, Sucrier subgroup) had high resistance. ‘Brazil’ (AAA, Cavendish subgroup), ‘Nongke No. 1’ (AAA, Cavendish subgroup) and an ABB cultivar had moderate resistance.  ‘Williams’ (AAA, Cavendish subgroup), ‘Jinfen’ and Guangen’ (ABB, Pisang Awak subgroup) were moderately susceptible.

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Control Most of the control measures reported for bacterial rhizome rot and tip-over also apply to bacterial soft rot of the rhizome and pseudostem. Some biological control work has been attempted (e.g. Nagaraj et al., 2012). Research in smallholder farms in Nicaragua and Costa Rica has shown that banana plants grown in agroforestry systems were characterized by an increase of potential plant-­beneficial bacteria, such as Pseudomonas and Stenotrophomonas, and a decrease of soft rot Erwinia bacteria (Köberl et al., 2015).

Pseudostem Wet Rot Introduction A watery rot of the pseudostem of plantain was initially observed in 1964 in Puerto Tejada, Colombia, a small town near Cali (Llanos, 1967). Pseudostem wet rot then spread into the Cauca valley of Colombia (Llanos, 1967; Fernández and López, 1970), where it caused serious crop losses of 80–90% on nearly 2000 ha of plantain within 2 years of the initial infection being ­observed. Stover (1972) recorded that this disease had been responsible for severe losses of cultivars in the Plantain subgroup (AAB) in Colombia, Venezuela and Ecuador. In the 1970s, the disease caused serious damage in plantains in Cuba, with incidence in some fields of up to 75%. It has also been found in neighbouring Haiti (T. Lescot, Dominican ­Republic, 1999, personal communication). Currently, the disease seriously affects plantations of plantain in El Salvador, Nicaragua, Panama and the Dominican Republic (Dita et al., 2013), where losses up to 50% were informally reported. Dessert banana clones can also be infected, but are more resistant than plantain. The disease is most serious in young, pre-flowering plants. In India, a pseudostem rot disease was detected for the first time on new plantings of ‘Giant Governor’ (AAA, Cavendish subgroup) in West Bengal in 1983. A survey revealed that the disease had been prevalent in the Karimpur area for some time and had spread through infected planting material (Chattopadhyay and Mukherjee, 1986).

Bacterial diseases affecting pseudostems have also been reported in China. Yan et al. (2011) described a rot that began in leaf sheath tissue, but rarely spread along the petiole and leaf veins. Fan et al. (2016) reported a disease that led to complete leaf collapse in serious cases with an incidence of 10–30%. Another bacterial sheath rot of banana was later reported as causing significant economic losses in China by Liu et  al. (2016). Incidences as high as 70–80% were observed in some plantations. Symptoms In Colombia, the pseudostems of plantain were seen with watery, translucent, light yellow spots, later turning reddish and dark. The attack began in the outermost layers of the pseudostem and the plant became bent at the part most severely affected (Fernández and López, 1970). In severe cases the pseudostem is so weakened by rot that the plant doubles over. There are no leaf or fruit symptoms (Stover, 1972). The most characteristic symptoms of the disease on bred tetraploid banana plants were cigar-leaf necrosis, soft rot and unpleasant odour of the pseudostem internal area. Additionally, the infected leaf-sheath tissue showed colouring from light to dark brown. Lower leaves remained in a normal position, without wilting for several weeks. Thereafter, they bent at the petiole level. Vegetative growth ceased in diseased plants (Guzmán and Sandoval, 1996). In 2009, a bacterial disease outbreak occurred in the banana-growing regions of Honghe in Yunnan Province. Typical symptoms included yellow-brown water-soaked spots on the petioles of lower leaves in the early stage, extending gradually along the petiole and becoming dark brown and irregular in shape. Interior pseudostems rotted and turned black-brown, along with necrosis of vascular bundles (Fan et al., 2016). In Yunnan and four other provinces in China, bacteria infecting the middle and bottom parts of the leaf sheath produced black and brown spots. These spots led to folding and bending of the sheath, drooping of leaves, and eventual browning and rotting of the entire leaf sheath (Liu et al., 2016). A similar disease, called bacterial leaf sheath rot, has been reported on enset plants over 3 years



Diseases Caused by Bacteria and Phytoplasmas

Plate 5.25.  Leaf sheath of enset affected by a bacterial wet rot in Ethiopia. The brown-coloured rot has been invaded by secondary infections (photo: M. Tessara and A.J. Quimio)

old. It is said to be common in the Guraghe, Welayita and Sidamo regions of Ethiopia and occurs during the dry time of the year. Yield loss has not been studied, but nothing can be harvested from diseased leaf sheaths. A watery rot starts at the base of the petiole of the outer leaf sheath and then spreads downward (Plate 5.25). The affected leaf becomes yellowish and eventually wilts and dies. The rot spreads inwards until only two or three leaves are left on the diseased plant. The causal agent is an unidentified bacterium. Diseased plants are usually harvested early before the rot penetrates deeply into the leaf sheath layers (M. Tessra and A.J. Quimio, Ethiopia, 1999, personal communications).

Causal agent The cause of the disease in Colombia was found by Fernández (1967) to be an Erwinia sp. Later,

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the pathogen was reported to be a strain of E. carotovora, which was tentatively named var. paradisiaca (Victoria and Barros, 1969). In the next year, the name E. paradisiaca was used by Fernández and López (1970) for the causal agent. In a disease survey in Cuba in 1974, a pseudostem disease of ‘Macho ¾’ (AAB, Plantain subgroup) was reported to be caused by E. chrysanthemi (Rivera, 1978). Up until 1980, the pathogen causing soft rot of the pseudostem of plantain had been designated as either E. carotovora ssp. paradisiaca, E.  carotovora ssp. chrysanthemi and E. chrysanthemi or E. paradisiaca (Dickey and Victoria, 1980). To confuse the situation further, Erwinia carotovora ssp. atroseptica was reported as the causal agent of a pseudostem soft rot disease of ‘Harton’ (AAB, Plantain subgroup) in Venezuela, south of Maracaibo Lake (Cedeño et al., 1990). Once again, traditional bacteriological tests and aberrant strains may have made it difficult to distinguish E. carotovora from E. chrysanthemi in the past. Dickey and Victoria (1980) examined 30 isolates from diseased plantain pseudostems in Colombia and compared them with isolates of closely related Erwinia species and their subspecies. The strains from plantain were phenotypically similar to E. chrysanthemi on the basis of the following characteristics: pectate degradation; production of phosphatase, indole, acetoin, and acid from ethanol; growth at 36–37°C; susceptibility to erythromycin; gas from glucose; utilization of malonate; no growth in 5% NaCl and no production of acid from alpha-methyl-d-glucoside, trehalose, maltose, lactose or palatinose. They could be distinguished from 322 isolates of E. chrysanthemi from 22 other plant hosts by the following phenotypic properties: production of acid from d-arabinose and raffinose; utilization of sodium tartrate; no production of lecithinase or of acid from inulin, mannitol or sorbitol; and inability to liquefy gelatin. It was named E. chysanthemi pv. paradisiaca (Dickey and Victoria,1980). Rivera and Ezavin (1980) and Rivera et al. (1980) found that isolates obtained from plants affected by soft rot of rhizome and pseudostem (described earlier) and plants with pseudostem wet rot differed in pathogenicity and aggressiveness. Isolates from necrotic rhizomes could infect and cause lesions in rhizome cortex and

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pseudostem tissues. However, isolates recovered from pseudostem wet rot lesions were able to affect leaf sheaths, but not the rhizome cortex (reported by Blomme et al., 2017a). In India, the cause of a new pseudostem rot of a Cavendish cultivar was identified as Erwinia chrysanthemi pv. paradisiaca by Chattopadhyay and Mukherjee (1986). The pathogen subsequently became known as Dickeya paradisiaca (Samson et al., 2005). In Guangdong, Guangxi and Hainan provinces in China, the bacterium causing infections of leaf sheaths was identified as Pantoea agglomerans by Yan et al. (2011). Klebsiella variicola was identified as causing the pseudostem disease in Yunnan province, China (Fan et al., 2016). Also in China, a bacterium isolated from diseased pseudostem tissue was identified as a Dickeya sp. (Li et al., 2011). Liu et al. (2016) reported that isolates causing a leaf sheath rot in Guangdong, Guangxi, Yunnan, Fujian and Hainan provinces in China were found to be more similar to Dickeya dadantii and less related to D. paradisiaca and D. zeae based on several phenotype characteristics. Disease cycle and epidemiology The soft-rotting pathogen is presumed to enter the leaf sheaths through wounds left when the petiole is cut. The soft rot advances down the petiole to the leaf sheath to infect the pseudostem and then spreads throughout the pseudostem. Infections also probably arise from the invasion of wounds on the pseudostem. When rotting is severe, the pseudostem is weakened to such an extent that the plant collapses, though leaves and fruit appear unaffected (Victoria and Barros, 1969; Stover, 1972). The causal bacterium or bacteria are common in the soil. Spread between plants may also occur when bacteria are splashed from the soil into the ‘throat’ or funnel made by the leaves, perhaps when the plants are still young. It is thought likely that the disease occurs when water collects in the ‘throat’ (ACIAR, 2016). Under natural conditions, the causal bacterium was found to disappear from the soil surface in 2 weeks, but survived for 4 weeks at a depth of 10 cm (Fernández and López, 1970). These authors believed the disease was spread

chiefly by cutting tools, but also by an insect of the genus Metamasius and rainwater. The Dickeya sp. identified by Li et al. (2011) was reported to survive in stagnant irrigation water and soil, often in association with decaying crop residues. A species-specific polymerase chain reaction (PCR) assay was developed for rapid and sensitive detection of the pathogenic bacterium in diseased plant tissues, soil and irrigation water.

Host reaction In tests in Colombia with nine isolates on various AAB plantain cultivars, ‘Hartón’ proved moderately resistant whereas ‘Maqueño’ and ‘Dominico’ were susceptible. ‘Gros Michel’ (AAA), ‘Manzano’ (AAB, Pome subgroup), ‘Pompo’ (AAB, Mai’a Maoli–Popoulu subgroup), ‘Espermo’ (ABB, Bluggoe subgroup) and ‘Cachaco’ (ABB, Bluggoe subgroup) were rated as moderately susceptible and ‘Grande Naine’ (Cavendish subgroup) moderately resistant (Victoria and Barros, 1969). In Cuba, all the 25 cultivars and species studied (genome types AA, BB, AAA, AAB and ABB) were susceptible to inoculation, though some AAA clones appeared to have some resistance to disease spread (Padrón, 1983). In Costa Rica, a higher incidence of pseudostem soft rot was found in the bred tetraploid ‘FHIA-02’ (4.4% diseased plants) than in ‘FHIA01’ (1.4% diseased plants) (Guzmán and Sandoval, 1996).

Control Control measures recommended by Fernández and López (1970) are the destruction of infected plants and those attacked by a Metamasius sp., which was suspected to be a vector, and sterilization of tools with 10% formalin. The sheath of the pseudostem should not be cut and 15 cm of the petiole should remain when dry leaves are removed. Stover (1972) reported that control measures consist of improving the aeration of plantations, avoiding green or yellow-green leaves when pruning and the frequent sterilization of pruning tools.



Diseases Caused by Bacteria and Phytoplasmas

Heart Rot of Abacá The symptoms of heart rot disease of abacá have been commonly mistaken for those of abacá bacterial wilt. The causal bacterial agent, which has been identified as a Dickeya sp., previously known as Pectobacterium chrysanthemi and Erwinia chrysanthemi, produces a soft rot of the central cylinder of the plant. However, it can also infect the roots and rhizomes. The causal organism is suspected to be related to rhizome rot of banana. Koch’s postulates were satisfied when abacá seedlings were inoculated with the isolated pathogen (Nunez et al., 2012). The pathogen was characterized based on cultural, morphological, physiological and biochemical characteristics and PCR analysis using primers specific to the ­banana rhizome rot pathogen. Virulent and avirulent isolates were obtained. The virulent pathogen produced white, round, elevated, smooth, glistening colonies in nutrient agar and potato dextrose peptone agar. The bacterium is Gram-­ negative, non-spore forming, with short rods measuring 0.6 μm × 1.53 μm; it is catalase positive, facultatively anaerobic and produces acids from glucose, galactose, lactose, sucrose, maltose and dulcitol. Both virulent and non-­virulent isolates gave positive amplifications using Dickeya sp. specific primers and were positively identified as a Dickeya sp. (Nunez et al., 2012).

Bacterial Leaf Blight and Leaf Spot D.R. Jones Leaf Blight A new leaf blight was found affecting banana in Nanning city, China, during a survey between 2003 and 2009 of the bacterial diseases on banana plants. Eight bacterial strains were isolated from affected banana leaves and identified as an intraspecific taxon of  Agrobacterium vitis  based on their 16S rDNA sequence similarities with those of 37 randomly selected bacterial strains registered in GenBank database. The representative strain Ag-1 was virulent on banana leaves and shared similar growth and biochemical

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r­eactions with the reference strain IAM14140 of A. vitis. The strains causing banana leaf blight were denominated as A. vitis pv. musae. The traditional  A. vitis  strains virulent to grapevines were proposed to be revised as  A. vitis  pv.  vitis. This is the first record of a new type of  A. vitis causing banana leaf blight in China (Huang et al., 2013).

Black Leaf Spot The bacterium causing a new bacterial leaf spot disease on dessert banana in Xinping County, Yunnan Province of China, was isolated from the infected plant and identified by cultural and morphological studies, pathogenicity test, physiological and biochemical detection as well as sequences analyses of gyrB, 16S rDNA and rpoB genes. The results showed that the leaf spot disease was caused by Klebsiella pneumoniae ssp. pneumoniae, which naturally occurs in the soil. The pathogen could infect banana leaves, pseudostems and fruit. Black spots appeared on d ­ iseased pseudostems within 3 days after inoculation and they enlarged lengthwise in both directions, causing massive brown necrosis of the internal tissues in 7 days. This is the first report of K.  pneumoniae ssp. pneumoniae causing leaf spot of banana plants in the world (Chen et al., 2016). Klebsiella pneumoniae is also an important bacterial pathogen in humans that is commonly associated with opportunistic and hospital-­ associated infections.

Bacterial Diseases of Fruit D.R. Jones Bacterial Finger-tip Rot Introduction This disease has a widespread distribution, with symptoms observed in Central America, Trinidad, Israel, Taiwan and Australia (Stover, 1972) and the Philippines (Francisco, 2015). It probably occurs wherever ‘Gros Michel’ (AAA) and Cavendish cultivars (AAA) are grown (Stover,

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1972). In Honduras, Stover (1972) reported that around 12% of stems carried an infected finger, with the four smallest apical hands accounting for 85% of total incidence, and that incidence declined during dry weather. In Australia, the disease is known as ‘gumming’ and is most common on November dump fruit, which emerges in  November and is harvested the following ­February–March. About 2–3% of fingers in a bunch can be affected. Losses from the disease are usually not severe and only a few fingers from the youngest hands are affected. Nevertheless, in December 2015, the banana growers and exporters association in the Philippines urged the Department of Agriculture to include the perceived threat and impact of the problem in its priority research area (Francisco, 2015).

Plate 5.26.  Symptoms of ‘gumming’ in fingers of ‘Mons Mari’ (AAA, Cavendish subgroup) in Queesnland, Australia. Initially, parts of the pulp within the finger appear slightly gelatinous and yellow Often, a brown discoloration is evident at the tip of the flower end (upper finger). Later, sections of the pulp can become rusty red in colour and degenerate to a gummy, sap-like constituency (lower finger) (photo: QDPI).

Symptoms Affected fingers are distinguished externally from unaffected fingers because of their distorted shape. They are smaller, straighter and often taper towards the tip, which is a condition known as ‘bottle-neck’. As a consequence, they protrude out of the line of the other fingers (Stover, 1972; Lee et al., 2003). Bacterial finger-tip rot is sometimes known as ‘Mokillo’ on account of the superficial similarity between the internal fruit symptoms of finger-tip rot and Moko bacterial wilt. Often, a brown discoloration is evident at the tip of the flower end (Stover, 1972; Lee et al., 2003). Parts of the pulp within the finger can appear slightly gelatinous and yellow (Lee et al., 2003) (Plate 5.26). Later, sections of the pulp can become rusty red in colour and degenerate to a gummy, sap-like constituency (Plate 5.26). Symptoms never spread further and vascular discoloration within the floral stem, of the type associated with Moko bacterial wilt, does not occur.

Causal agents Internal fruit discoloration has been attributed to a bacterial invasion through the flower end of the finger. Stover (1972) reported that mechanical inoculation of a Pseudomonas species into

the pulp reproduced the symptoms of finger-tip rot within 14 days. Stover (1972) also described a bacterial soft rot of green fruit in Honduras and Nicaragua. This description was largely based on an account of a severe Mokillo infection, published in an earlier edition of Wardlaw’s (1972) work, and may in fact refer to bacterial finger-tip rot. Luna et al. (1988) isolated a non-fluorescent pseudomonad from bananas exhibiting symptoms in Mexico. Isolates produced discoloration on inoculation into the fruit, but not the pseudostem, and did not elicit a hypersensitive reaction in tobacco. The bacterium was oxidase-­ positive and arginine dihydrolase-negative, did not reduce nitrate and accumulated poly-­ βhydroxybutyric acid. Bacterial finger-tip rot is an endemic minor disease of banana in Taiwan (Lee et al., 2003). Isolations from diseased fingers consistently yielded bacterial colonies that were whitish yellow on Luria-Bertani agar and did not produce fluorescent pigment on King’s medium B. They were also Gram-negative and aerobic and grew at 42°C. Rots were caused on onion that were more severe than on potato and carrot slices. The bacterium was subsequently identified as Burkholderia cepacia. An almost complete 16S rDNA sequence of one isolate (strain B9) was determined and compared with



Diseases Caused by Bacteria and Phytoplasmas

available 16S rDNA sequences of members of B. cepacia complex. The sequence was similar (99.87%) to that of B.  cepacia genomovar III LMG 12614 (Lee et al., 2003). Lee et al. (2003) injected bacterial suspensions through the centre of the stigma and the inoculated fingers were enclosed in a plastic bag to maintain high humidity. Typical symptoms appeared in 10–14 days. A small number of control plants inoculated with sterile water also showed symptoms. The authors believed that this may have been the result of finger infestation with epiphytic or endophytic B. cepacia. Young fruits were more susceptible than older fruits. The bacterium was readily re-isolated from diseased fingers and confirmed as B. cepacia (Lee et al., 2003). Subsequent work in Taiwan showed that the banana fruit pathogen had biochemical and physiological characteristics very similar to those of the onion decay pathogen. However, the former, which was named B. cenocepacia, belonged to genomovar III of the B. cepacia ­complex, while the latter, which was named B. cepacia, belonged to genomovar I. RecA PCR– RFLP tests showed the genetic variability of the B. cenocepacia isolates, which contained recA IIIA and IIIB lineages. In addition, PCR and PCR–RFLP assays were developed for the detection and identification of these two pathogens (Lee et al., 2006). To better understand the nature of B. cenocepacia, the genetic diversity of 44 isolates from various banana-growing regions in southern Taiwan was investigated using recA PCR–RFLP, recA nucleotide sequence analysis, and pulsedfield gel electrophoresis assays. All isolates were assigned to genomovar III and consisted of two groups, A and B, which corresponded to recA lineage IIIA and IIIB. The group B strains were separated into B1 and B2 subgroups and the B1 strains were further divided into distinct lineages. The B1 strains were the most frequently detected and occurred in all regions tested (Lee and Chan, 2007). There was no significant difference between strains from each subgroup in their virulence on Cavendish banana fingers. PCR assays were ­further used to determine whether B. cenocepacia contained the cable pilus subunit gene (cblA), IS1356, and B. cepacia epidemic strain marker (BCESM), which are all associated with

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epidemic strains of B. cepacia. The results indicated that cblA and IS1356 were absent, but BCESM was present. The study revealed that banana is a natural reservoir of genetically diversified B. cenocepacia strains (Lee and Chan, 2007). In far north Queensland in Australia, a quality problem was recently encountered ­during the processing of ripe banana for pulp. Affected fingers had reddish brown to blackish centres and pulp breakdown. About 1% of fruit was affected, with incidence more common in the wet season. Pantoea agglomerans and Enterbacter cowanii were frequently recovered from affected fruit. Inoculation experiments showed that both bacteria caused similar symptoms, which were the same as those found at the pulp processing facility. Inoculated fingers were smaller, yellowed prematurely and were out of line with the other fingers in the hand (Pathania et al., 2017).

Disease cycle and epidemiology The precise details of the disease cycle are not known, but insect transmission of the pathogen may be involved. The disease seems to be less common in dry weather, suggesting moisture and high humidity play a role.

Host reaction Fruit of ‘Gros Michel’ (AAA) and cultivars in the Cavendish subgroup (AAA) have been reported as affected (Stover, 1972).

Control In the Philppines, ‘Mokillo’ is controlled through decontamination of cutting tools and farm equipment. Distorted fingers are removed from hands during packing operations and discarded. No treatment was identified by Pathania et al. (2017) in Australia to manage the problem, though it was suggested that insect pest management, proper drainage maintenance and field sanitation during wet and humid weather may reduce incidence.

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Phytoplasma Diseases D.R. Jones Introduction Mollicutes are bacteria without a rigid cell wall. Phytoplasmas are Mollicutes that are bound by a triple-layered membrane. A phytoplasma infection often triggers leaf yellowing, probably due to the presence of phytoplasma cells in phloem, which can affect phloem function and carbohydrate transport, inhibit chlorophyll biosynthesis and trigger chlorophyll breakdown. Phytoplasmas are spread principally by insects of the families Cicadellidae (leafhoppers), Fulgoridae (planthoppers) and Psyllidae (jumping plant lice) (Weintraub and Beanland, 2006), which feed on the phloem of infected plants, ingesting phytoplasmas and transmitting them to the next plant on which they feed. Most modern classifications of phytoplasmas are based on DNA sequences, especially 16S rRNA sequences (Lee et al., 1998). The delineation of groups and subgroups of phytoplasmas are discussed by Zhao and Davis (2016).

China and India The first report of a phytoplasma in banana came from a plant with symptoms of what appeared to be bunchy top disease in China (Li et al., 1999). DNA was extracted and the presence of a phytoplasma was established on the basis of 16S rDNA restriction patterns and a sequence analysis of the 16S rDNA fragment. The RFLP pattern, sequence data and phylogenic tree showed that this phytoplasma belonged to ‘Candidatus Phytoplasma asteris’ in the aster yellows or 16SrI group, which is the largest and most diverse. The second report came from plants in the Aligarh district of Uttar Pradesh in northern India (Singh et al., 2009). According to the authors, the banana was stunted and had symptoms similar to bunchy top disease except the actual bunchy top formation. DNA was extracted from diseased and healthy plants. A phytoplasma was detected in the plants with symptoms, but not in symptomless plants. Based

on RFLP patterns, ­sequence homology and phylogenetic analysis, the phytoplasma, which was initially named ‘banana stunting phytoplasm’, was also identified as being closest to ‘Candidatus Phytoplasma asteris’. ‘Candidatus Phytoplasma asteris’ is naturally transmitted by a wide range of leafhoppers. However, Macrosteles fascifrons (M. quadrilineatus) is reported to be the principal vector (Lee et al., 2004; Weintraub and Beanland, 2006). The banana cultivars affected in the above reports were not identified. Colombia Elephantiasis has been described as a rare problem with localized outbreaks reported from Suriname, Colombia, the Dominican Republic, Costa Rica, Panama and Honduras (Stover, 1972). It was first described from Suriname by Essed (1911), who attributed the cause to Ustilaginoidella oedipigera, which was later identified as a species of Fusarium. However, there was no proof that the fungus was pathogenic and the cause of elephantiasis remains unknown. Fungi are seldom present in affected tissue in Colombia and Central America, but bacteria are common. Inoculation with this pathogen causes some swelling and splitting at the pseudostem base, but fails to produce elephantiasis. In Honduras, the disorder was sometimes present together with ‘yellow mat’ (Prescott, 1917). In 1954, about 24 ha of ‘Gros Michel’ (AAA) in the Dominican Republic became severely affected and were destroyed. The disorder did not reappear after ploughing and replanting. The base of the pseudostem swells and the outer sheaths rupture just above the rhizome (Plate 5.27). Breakage continues inward until the pseudostem falls over, leaving a conical, pineapple-like stump. The breakage is often associated with a narrow band of rot, from which a pathogenic bacterium is obtained. Sometimes leaf symptoms are present, consisting of swollen veins and midrib. Suckers may be normal or deformed and split or are easily broken off close to the ground. In severe cases, the entire mat may be killed. There are no internal symptoms in the rhizome (Stover, 1972). More recently, petioles have been described as remaining rigid, bunching and forming an arrangement similar to that



Diseases Caused by Bacteria and Phytoplasmas

Plate 5.27.  Symptoms of elephantiasis in ‘DominicoHarton’ (AAB, Plantain Subgroup) growing near Armenia, Colombia (photo: L. Pérez-Vicente, INISAV)

of traveller’s palm (Ravenala madagascariensis). In addition, the tips of suckers of affected plants have been reported as necrotic and fruit size reduced (Aliaga et al., 2018). Elephantiasis of ‘Gros Michel’ (AAA) and ‘Dominico Harton’ (AAB, Plantain subgroup) in Colombia has recently been associated with infection by ‘Candidatus Phytoplasma asteris’, ­ which is also found in several tree species in the country (Aliaga et al., 2018). It may also be the cause of elephantiasis in Suriname and Central America. The same type of phtoplasma has been identified in banana in China and India (see above).

Papua New Guinea Cooking-banana cultivars (ABB) with distinct disease symptoms, clearly different from those produced by bunchy top, Fusarium wilt or bacterial wilts, were observed during plant disease

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surveys in Papua New Guinea (PNG) carried out jointly by the PNG National Agricultural Quarantine and Inspection Authority and Australia’s Department of Agriculture, Fisheries and Forestry (Davis et al., 2012). External symptoms of diseased plants consisted of leaf yellowing and later leaf necrosis (Plate 5.28). Affected plants appeared to be dying. Inside pseudostems, small regions of black or brown vascular tissues, usually with wet and necrotic pockets, formed a discontinuous streaking (Plate 5.29) (Davis et al., 2012). In 2008, a sample of an affected plant was taken back to the laboratory for a disease diagnosis (Davis et al., 2012). This sample tested positive for phytoplasma using a nested PCR test in the laboratory of PNG’s Oil Palm Research Association. This finding led to more disease surveys in Madang, Morobe, Milne Bay and East Sepik provinces in 2009 and in the Western and North Solomons provinces in 2010. Samples were taken from 16 yellowing and dying ABB banana plants showing similar external symptoms as described above. In addition, one symptomless sucker from each of two sampled plants were collected for testing. Each sample consisted of approximately 0.5–1.0 g of vascular tissue excised from immediately above and below obvious pockets of necrosis. The samples from the symptomatic plants plus suckers all tested positive for phytoplasma while samples from symptomless ABB plants and ABB plants with leaf yellowing symptoms that did not match those of the disease tested negative. This indicated that phytoplasmas were consistently associated with the distinctive ­ symptoms (Davis et al., 2012). The 16S rRNA gene, 16S-23S spacer region, part of the 23S rRNA gene and the ribosomal protein (rp) S19 (rps19), ribosomal protein L22 (rpl22) and ribosomal protein S3 (rps3) genes of the phytoplasmas in two samples from Madang Province were amplified and sequenced. Both samples contained identical phytoplasmas, which were named ‘Banana wilt associated phytoplasma’ (BWAP) (Davis et al., 2012). Phylogenetic analyses of the 16S rRNA gene of BWAP were conducted using: (i) the ‘­Cocos nucifera’ lethal yellowing phytoplasma (16SrIV group), which had been recently identified in PNG (Kelly et al., 2011), and 44 formally recognized phytoplasma species; and (ii) the rp gene regions

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Plate 5.28.  Banana in the Kalapua subgroup (ABB) showing leaf yellowing and necrosis associated with ‘Banana wilt associated phytoplasma’ at Kjabraka No. 1, East Sepik Province, Papua New Guinea (photo: R. Davies, NAQS).

of 75 phytoplasma species or strains. These analyses showed that the two BWAP isolates from Madang Province had identical 16S rRNA and rp gene sequences to ‘Cocos nucifera’ lethal yellowing phytoplasma. Similarities with 16S rRNA sequences in other phytoplasma s­pecies ranged from 90% to 95.4% and rp gene ­sequence similarities ranged from 70% to 75.4%. This suggested that BWAP was a unique species (Davis et al., 2012). Davis et al. (2012) noted that ‘Cocos nucifera’ lethal yellowing phytoplasma was only found in coconut palms in Madang Province in PNG and yet BWAP had been found in banana

samples from East Sepik, Morobe and Western provinces as well as Madang Province. This meant that banana infected with BWAP may not be acting as a source of inoculum for coconut palms. They speculated that there may be as yet undetected differences between the two phytoplasmas that prevented transmission or differences in the feeding behaviour of insect vectors present. A phytoplasma detected in a sample from Madang Province taken from a banana that had underdeveloped fruit as well as yellowing and streaking belonged to the peanut witches’ broom or 16SrII group. Another phytoplasma in the



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Solomon Islands

Plate 5.29.  Cut pseudostem of a banana cultivar in the Kalpua subgroup (ABB) showing internal symptoms of discontinuous vascular streaking and pockets of necrosis associated with ‘Banana wilt associated phytoplasma’ at Taman, Morobe Province, Papua New Guinea (photo: R. Davies, NAQS).

loofah witches’ broom or 16SrVIII group was found in a sample from Morobe Province taken from a banana with yellows and basal necrosis, but no discontinuous vascular streaking symptom. Undetermined phytoplasmas, which were not cloned and sequenced, were found in samples from Madang, Morobe, Western and North Solomons provinces (Davis et al., 2012). Davis et al. (2012) reported that BWAP and the other phytoplasmas found in banana in PNG were detected only in ABB clones with characteristic leaf symptoms. Cultivars ‘Daru’ and ‘Kalapua’ in Madang Province, ‘Daru’ from East Sepik Province and ‘Kalapua’ from Morobe and Western provinces were positive for BWAP. The phytoplasma in the 16SrII group was detected in ‘Kalapua’ in Madang Province and the one in the 16SrVIII group in ‘­Kalapua’ in Morobe Province. Undetermined ­phytoplasmas were found in ‘Daru’ and ‘Kalapua’ in Madang Province, ‘Kalapua’ in Morobe Province, ‘Kalupua’ and ‘Goya’ in Western Province and ‘Kalapua’ in North Solomons Province.

During a disease survey of the north-west Solomon Islands in 2012, ABB cooking-banana plants were found on Magusaiai Island in the Shortland Islands group with identical symptoms as had been seen caused by phytoplasmas in PNG. Affected banana plants were widespread and had yellowing and/or leaf death with unfilled fruit bunches. Inside the pseudostem, ­discontinuous brown or black vascular streaks were associated with wet, necrotic pockets (­Davis et al., 2015). Samples from three symptomatic plants on Magusaiai Island were tested for phytoplasma by nested PCR. All three samples were found to be positive. The source ABB cultivars were identified by their local names of ‘Vovosi’ and ‘Sauhu’. Because no stands of the same clones of ABB banana were found free of symptoms, control samples were taken from symptomless plants of another ABB clone. These tested negative for phytoplasma (Davis et al., 2015). Two of the three positive isolates, which were obtained from ‘Vovosi’, were cloned and sequenced. Undetermined phytoplasmas found earlier in the North Solomons Province of PNG, which lies very close to the Shortland Islands group, were also cloned and sequenced, as was another undetermined phytoplasma found in the Western Province of PNG. These were all compared with one another and also an isolate of BWAP from Madang Province in PNG using the iPhyClassifier program of Zhao et al. (2009) (Davis et al., 2015). The results of the analysis showed that none matched any known RFLP patterns sufficiently for assignment to any known group. The most similar patterns were in three phytoplasmas belonging to the coconut lethal yellows or 16SrIV group. In another phylogenetic analysis, all phytoplasma isolates formed a clade with the PNG ‘Cocos nucifera’ lethal yellowing phytoplasma also in the 16SrIV group. In addition, the two isolates from Magusaiai Island in the Solomon Islands and the two isolates from the North Solomons Province in PNG had identical sequences and were sisters to BWAP, the undetermined phytoplasma from the Western Province of PNG and the ‘Cocos nucifera’ lethal yellowing phytoplasma (Davis et al., 2015). Further work on the Madang coconut phytoplasma

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provided more evidence for it being identical to BWAP (Lu et al., 2016). ‘Candidatus Phytoplasma cocosnigeriae’, a member of the Nigerian coconut lethal decline or 16SrXXII group, was determined to be a sister to all banana phytoplasmas from PNG and the Solomon Islands and the ‘Cocos nucifera’ lethal yellowing phytoplasma from PNG. ‘Candidatus Phyoplasma cocosnigeraie’ has been shown to be identical to the reference strain of ‘Candidatus Phytoplasma palmicola’ (see Harrison et al., 2014) (Davis et al., 2015). The results suggested to Davis et al. (2015) that phytoplasmas now found in banana plants and coconut trees in the PNG–Shortland Islands region diversified from a recent common ancestor. They noted that the ‘Cocos nucifera’ lethal yellowing phytoplasma in PNG has a partial ­sequence of 168 rRNA gene (756 bp), which is identical to BWAP, and that this relationship may be the result of a host jump from banana to coconut. A phylogenic tree constructed from available evidence suggests that there may be very important differences between the BWAP in Madang Province and the phytoplasmas found on banana plants elsewhere in the region (R. Davis and L. Jones, Australia, 2017, personal communication; P. Kokoa, PNG, 2017, unpublished data).

The BWAP causing an epidemic on banana in the Western Province of PNG was also different from the BWAP in Madang Province. Observations at the outbreak site in the Western Province also indicated that coconuts remained unaffected by any phytoplasma-like disease over a 5-year period. The phytoplasmas found in the Solomon Islands and adjacent North Solomons Province of PNG grouped together. Available evidence suggests that phytoplasma diseases of banana that cause yellowing in the Shortland Islands have not been observed during disease surveys in other areas of the Solomon Islands. An internal quarantine measure to restrict the free movement of banana material from the Shortland Islands to other parts of the Solomon Islands was recommended (Davis et al., 2015)

Acknowledgements D.R. Jones would like to thank S.J. Eden-Green and G. Blomme for their advice on aspects of certain bacterial diseases. The information provided by R. Davis on banana phytoplasmas in Papua New Guinea and the Solomon Islands is also gratefully acknowledged

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6 

Diseases Caused by Viruses

Banana Bunchy Top J.E. Thomas Introduction Bunchy top is widely regarded as the most serious virus disease affecting banana, as it can cause dramatic crops losses (Kumar et al., 2015; Ploetz et al., 2015). The centre of origin of the genus Musa is the South and Southeast Asian– Australasian region (Simmonds, 1962; Perrier et  al., 2009, 2011) and it is probable that the causal agent (banana bunchy top virus, BBTV) also originated somewhere in this broad geographical area (Stainton et  al., 2015). The discovery of a second, related virus in abacá and banana (abacá bunchy top virus, ABTV) (Sharman et  al., 2008), which can cause typical bunchy top symptoms in these hosts, has complicated this situation, though ABTV appears to be less common and to have a more limited distribution than BBTV. The newly recognized ABTV is discussed fully in the next disease description in this chapter. History The first recorded outbreaks of bunchy top disease were on banana in Fiji in 1889, though reports and photographs taken around that time leave little doubt that the disease was present 362

there as early as 1879 (Darnell-Smith, 1924; Magee, 1953). Interest in the disease may have only arisen due to its effects on the new export industry, which commenced production in 1877 and was based on cultivars in the Cavendish subgroup (AAA). Although the outbreaks were first observed in commercial plantations of Cavendish cultivars, the pattern of spread of the disease and the severe symptoms in local cooking-­banana cultivars led Magee (1953) to consider it unlikely that bunchy top originated in Fiji. Cavendish cultivars were not traditionally grown in Fiji and must have been imported at some stage, thus providing the opportunity to also introduce the disease. The industry reached a production peak of 788,000 bunches in 1892, but by 1895 had declined to 147,000 bunches, mainly because of bunchy top. Temporary recoveries in the industry occurred around 1896, possibly due to the planting of ‘Gros Michel’ (AAA), and from 1912 to 1916, to a large extent because of the opening up of new plantations. Surprisingly, a survey in 1937 showed that, although bunchy top incidence was high, with 5–30% of plants in all plantations and gardens affected, no plantings had an incidence of 100% (Magee, 1953). Other early records of the disease include Egypt in 1901 (Fahmy, 1924, in Magee, 1927), the origin of infection unknown, and Australia and Sri Lanka in 1913, both outbreaks probably resulting from the importation of infected planting material from Fiji (Magee, 1953). Bunchy top was first reported in Australia in the Tweed

© D.R. Jones 2019. Handbook of Diseases of Banana, Abacá and Enset (D.R. Jones)



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District of New South Wales (NSW) (Magee, 1927). Rapid expansion of the banana industry took place over the next decade and, with it, the inadvertent spread of the disease. By 1927, bunchy top was present along a 300 km stretch of the east coast of Australia from Grafton in central NSW to Yandina in southern Queensland. An isolated outbreak at Innisfail, a further 1300 km to the north, was detected in 1926, but subsequently eradicated. Production in the expanding Australian banana industry peaked in 1922, but 3 years later had collapsed. In the Tweed and Brunswick districts of NSW, every plantation was affected by bunchy top by 1925, with a disease incidence of 5–90%. In addition, the total area of production had been decreased by 90% (Magee, 1927). A similar situation existed in southern Queensland, where, for example, production in the Currumbin District was reduced by over 95%. C.J.P. Magee noted at the time: ‘It would be difficult for anyone who has not visited these devastated areas to visualize the completeness of the destruction wrought in such a short time by a plant disease’ (Magee, 1927). A severe outbreak of banana bunchy top occurred in Pakistan in the early 1990s (Khalid and Soomro, 1993). Land area under production fell by 55% in 1 year as a direct result of the disease and in some plantations the incidence of bunchy top was observed to be 100% (INIBAP, 1992). The disease has also appeared in Hawaii, causing much damage (Ferreira et  al., 1989). Disease outbreaks in India from 2007 to 2010 caused an annual losses of US$50 million (Selvarajan and Balasubramanian, 2014). In sub-­ Saharan Africa, the disease spreads unabated in many countries, with average incidences in affected areas ranging between 5% and 70%, but in some cases up to 95% (Kumar et al., 2015). The disease was recorded from South Africa for the first time in 2015 (Jooste et  al., 2016) and the next year it was found in Mozambique (FAO/ IPPC, 2016). Bunchy top of abacá was first noticed in the Philippines in 1915 and was causing serious economic damage by 1923 (Wardlaw, 1961). Many production areas were replanted with other crops. Early investigations Initial attempts to attribute a cause to bunchy top implicated a wide variety of agents, including

fungi (Knowles and Jepson, 1912, in Magee, 1927), nematodes (Nowell, 1925 and Fahmy, 1924, cited in Magee, 1927) and bacteria (Darnell-Smith, 1924). Soil and nutritional factors were also considered (Darnell-Smith and Tryon, 1923; Darnell-Smith, 1924), though none of these investigations was conclusive. Although some studies had suggested an association between the banana aphid and the disease (Darnell-Smith, 1924), it was not until the ground-breaking work of Magee (1927) that the viral nature of the disease was established. In the space of 3 years, he established that the causal agent was a virus transmitted by the banana aphid (Pentalonia nigronervosa) and in infected planting material. He also proposed management strategies that still form the basis of Australia’s control programmes today. Geographical distribution Bunchy top of banana occurs in many, though not all, countries in the South and Southeast Asian–Pacific region and in various African countries. Significantly, the banana-exporting countries of the Latin American–Caribbean region are free from the disease, though the aphid vector is present. The countries for which authenticated records of banana bunchy top disease exist are shown in Table 6.1, together with records of the detection of the causal agent. The discovery of a second, related virus in abacá that can cause typical bunchy top symptoms in abacá and banana has complicated distribution records. Although the virus from abacá has only formally been recorded from the Philippines (in abacá) and from Sarawak in Malaysia (in banana) and may have a limited distribution, it cross-reacts in some serological and polymerase chain reaction (PCR) assays with the virus common in ­banana (Sharman et al., 2008) and thus some bunchy top records based on these tests or symptoms alone are not unequivocal. Reports of bunchy top disease in abacá from East Malaysia (Sabah), West Malaysia and Papua New Guinea (Magee, 1953; Wardlaw, 1961) need to be authenticated, as they were associated with atypical symptoms, they were not confirmed by aphid transmission tests and bunchy top of banana was not present.

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Table 6.1.  Distribution records for BBTV. Region

Country

Reference

Africa

Angola Benin Burundi Cameroon Central African Republic

Kumar et al. (2008) Lokossou et al. (2012) Sebasigari and Stover (1988) Oben et al. (2009) Fouré and Lassoudière in Diekmann and Putter (1996) Wardlaw (1961) Busogoro et al. (2009) Magee (1953) Fouré and Manser (1982) Kenyon et al. (1997) FAO/IPPC (2016) Adegbola et al. (2013 Sebasigari and Stover (1988) Jooste et al. (2016) Gondwe et al. (2007) Fouré and Manser (1982) Stover (1972) Thomas and Dietzgen (1991) Buddenhagen (1968) Sulyo and Muharam (1985) Magee (1953) Bananej et al. (2007) Gadd (1926)

Asia

Oceania

Congo (Republic of the) Congo (Democratic Republic of the) Egypt Gabon Malawi Mozambique Nigeria Rwanda South Africa Zambia Bangladesh Cambodia China Hong Kong Indonesia India Iran Japan (Ogasawara-gunto, formerly Bonin Island Japan (Okinawa) Korea Laos

Malaysia (Sarawak) Myanmar Pakistan Philippines Sri Lanka Taiwan Thailand Vietnam Australia Fiji Guam Kiribati (formerly Gilbert Islands) Marianas Islands (Guam) Marianas Islands (Saipan, Tinian and Rota) New Caledonia Samoa Tahiti Tonga Tuvalu (formerly Ellice Islands) USA (American Samoa) USA (Hawaii) Wallis Islands

Kawano and Su (1993) Kiritani (1992) K. Chittarhat, K.S. Crew, A.D.W. Geering, G Kong, J.E. Thomas, Laos and Australia (2015, unpublished results) Su et al. (1993) Furuya and Natsuaki (2006) Soomro et al. (1992) Castillo and Martinez (1961) Bryce (1921); Magee (1953) Sun (1961) Wongsuwan and Chawpongpang (2012) Vakili (1969) Magee (1927) Magee (1927) Beavor (1982) Shanmuganathan (1980) Beavor (1982) Miller et al. (2011) Kagy et al. (2001) Magee (1927) M.Wong, D. Massé and B. Hostachy, Tahiti and Réunion (2017, personal communication) Magee (1927) Campbell (1926) Magee (1927) Dietzgen and Thomas (1991) Simmonds, 1933



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Symptoms The typical symptoms of bunchy top of banana are very distinctive, readily distinguished from those caused by other viruses of banana except ABTV, and were described in detail by Magee (1927). Plants can become infected at any stage of growth and there are some initial differences between the symptoms produced in aphid-infected plants and those grown from infected planting material. In aphid-inoculated plants, symptoms usually appear in the second leaf to emerge after inoculation and consist of a few dark green streaks or dots on the minor veins on the lower portion of the lamina. The streaks form ‘hooks’ as they enter the midrib and are best seen from the underside of the leaf in transmitted light (Plate 6.1). The ‘dot–dash’ symptoms can sometimes also be seen on the petiole (Plate 6.2). The following leaf may display whitish streaks along the secondary veins when it is still rolled. These streaks become dark green as the leaf unfurls. Successive leaves become smaller, both in length and in width of the lamina, and often have chlorotic, upturned margins. The leaves become harsh and brittle and stand more erectly than normal, giving the plant a rosetted and ‘bunchy top’ appearance (Plate 6.3). Suckers from an infected stool can show severe symptoms in the first leaves to emerge

Plate 6.1.  Dark green streak (‘dot and dash’) symptom of bunchy top disease in the minor leaf veins of a cultivar in the Cavendish subgroup (AAA). Note ‘hooking’ as streaks enter the midrib (photo: J.E. Thomas, QDAF).

(Plate 6.4). The leaves can also be rosetted and small, with very chlorotic margins, which tend to turn necrotic (Plate 6.5). Dark green streaks are usually evident in the leaves.

Plate 6.2.  Dark green streaks in the petiole of a cultivar in the Cavendish subgroup (AAA) affected by bunchy top disease (photo: J.E. Thomas, QDAF).

Plate 6.3.  Progressive shortening and bunching of the leaves of a cultivar in the Cavendish subgroup (AAA) affected by bunchy top disease. The symptoms on the lower leaves are caused by black leaf streak disease (photo: J.E. Thomas, QDAF).

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Infected plants rarely produce a bunch after infection and do not fruit in subsequent years. Plants infected late in the growing cycle may fruit once, but the bunch stalk and the fruit will be small and distorted. On plants infected very late, the only symptoms present may be a few dark green streaks on the tips of the flower bracts (Plate 6.6) (Thomas et al., 1994a). Mild strains of the virus, which produce only limited vein clearing and dark green flecks, and symptomless strains have been reported in Cavendish plants from Taiwan (Su et al., 1993). Mild disease symptoms are expressed in some banana cultivars and Musa species. The dark green leaf and petiole streaks, so diagnostic and characteristic of infection of cultivars in the Cavendish subgroup, can be rare or absent (Magee, 1953). Some plants of ‘Veimama’ (AAA, Cavendish subgroup), after initial severe symptoms,

have been observed to recover and to display few if any symptoms. These reports and findings are reported in more detail under ‘Causal agent’ below (Magee, 1948). The symptoms of bunchy top disease on abacá include a reduction in leaf size and lamina area, rosetting of leaves, upcurling and yellowing of leaf margins and stunting of the pseudostem. Chlorotic areas on the leaves sometimes collapse, becoming necrotic, and ‘heart rot’ often occurs in the pseudostem. No fruit is produced and infected plants usually die within 1 or 2 years (Magee, 1953; Wardlaw, 1961). Dark green ‘dots and dashes’ on the minor veins, midribs and petioles occur in only a few abacá cultivars. Magee (1953) concluded that in abacá ‘the green streak symptom occurs so rarely as to lose nearly all its value as an aid in diagnosis’. Symptoms

Plate 6.5.  Intense rosetting of leaves of a sucker of ‘Basrai’ (AAA, Cavendish subgroup) in Pakistan because of bunchy top disease. Note leaves have yellowing margins that are turning necrotic (photo: D.R. Jones, INIBAP).

Plate 6.4.  Suckers arising from the underground corm of a plant that has been removed because it was affected by bunchy top disease. Leaves on two of the four suckers show extensive marginal chlorosis caused by bunchy top disease (photo: J.E. Thomas, QDAF).

Plate 6.6.  Dark green streaks on the tips of the male flower bracts from a banana plant affected by bunchy top disease. (photo: J.E. Thomas, QDAF).



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described for abacá bunchy top in the field are very close to those that develop on abacá inoculated with the bunchy top virus from banana. When enset is experimentally infected with the banana virus, symptoms are similar to those described for abacá (Magee, 1953).

Causal agent Evidence for a viral agent Banana bunchy top virus (BBTV) in the genus Babuvirus, family Nanoviridae, is presumed to be the causal agent of bunchy top of banana, though unequivocal evidence, by reproduction of the disease through inoculation of cloned genomic components using biolistic delivery on gold micro-carriers, Agrobacterium-inoculation (agro-inoculation) (Mware et al., 2016) or purified virions (Thomas and Dietzgen, 1991), is lacking. However, the virions are intimately associated with the disease (Harding et al., 1991; Thomas and Dietzgen, 1991) and have been detected in all symptomatic plants tested (Dietzgen and Thomas, 1991; Thomas, 1991; Thomas and Dietzgen, 1991; Karan et al., 1994). Particle and genome properties The virions of BBTV are icosahedral, ca 18–20 nm in diameter (Plate 6.7), have a coat protein of ca 20,000 kDa, a sedimentation coefficient of ca 46S and a buoyant density of 1.29–1.30 g/ cm3 in caesium sulfate (Wu and Su, 1990c; Dietzgen and Thomas, 1991; Harding et  al., 1991;

100 nm Plate 6.7.  Virions of banana bunchy top virus contrasted with ammonium molybdate (photo: J.E. Thomas, QDAF).

Thomas and Dietzgen, 1991). Purified preparations have an A260/280 of 1.33 (Thomas and ­Dietzgen, 1991). The virus possesses a multi-­ component genome, comprising six circular ssDNA components, each ca 1000–1100 nucleotides long (Wu et  al., 1994; Yeh et  al., 1994; Burns et  al., 1995; Xie and Hu, 1995; Karan et  al., 1997). These components have been found in almost all isolates examined (Karan et  al., 1994, 1997; Stainton et al., 2012). DNA-R encodes the master replication initiation protein (Rep), which directs the replication of itself and the other five integral components (Horser et al., 2001). While the function of DNA-U3 remains unknown, the roles of the other encoded proteins have been determined by functional analysis. DNA-S encodes the coat protein (CP), as shown by N-terminal sequencing and production of antibodies against the cloned and expressed CP (Wanitchakorn et  al., 1997). Green fluorescent protein tagging and confocal microscopy showed that DNA-M and DNA-N are likely to encode the movement protein and the nuclear shuttle protein, respectively. The gene product of DNA-C contains a retinoblastoma protein-­binding LxCxE motif, indicating that it is a cell cycle link (Clink) protein which enhances viral DNA replication (Wanitchakorn et  al., 2000). All components have a similar genome organization with a single open reading frame (ORF), except for some isolates of BBTV that have a second, internal ORF on DNA-R. Replication is thought to occur via a rolling circle mechanism. The components also share a major common region (CR-M), the site of DNA primer binding for dsDNA replicative form synthesis, and a stemloop common region (CR-SL) containing the origin of replication. The CR-SL contains three short repeated sequences (iterons F1, F2 and R1), which are the presumed binding site for the Rep, allowing trans-replication of the other genome components (Vetten et al., 2012). These genome components are present in the plant in vastly different proportions, and these ratios remain relatively constant throughout the course of infection. When compared with DNA-S, present at the lowest relative concentration, these ratios were S1:C2:M6:R44:U359:N692 (Mware, 2016). Two studies have been presented on the suppression of RNA silencing in BBTV infections and the different results obtained by these two groups are interesting. Both have identified MP as the major pathogenicity determinant and

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suppressor of RNA silencing. However, Amin et al. (2011) concluded that Clink was also a suppressor. Niu et al. (2009) concluded that both MP and CP are silencing suppressors acting at different positions in the silencing pathway. Although the experimental systems used by both groups were very similar, they used BBTV isolates from different groups: Asian (Niu et al., 2009) and Pacific (Amin et al., 2011). There is evidence that the two groups have followed separate evolutionary paths (Hu et al., 2007) and it is possible that they have also evolved somewhat different strategies for RNA silencing (Amin et al., 2011). Some BBTV isolates may contain additional self-replicating components (alphasatellites), only distantly related to the master Rep. Similar molecules, thought to originate from nanovirus Reps, are found associated with some begomovirus infections, especially monopartite begomovirus/ betasatellite complexes (Briddon et  al., 2004). Alphasatellites have a size (1000–1100 nt) and structure similar to DNA-R; they code for a Rep protein and have a stem-loop structure, but differ in that the loop but not the stem is conserved, and the putative TATA boxes are located 5′ instead of 3′ to the CR-SL. They also lack the CR-M (Horser et al., 2001). They are phylogenetically distinct and are more closely related to the alphasatellites of members of the genus Nanovirus than to the DNA-R molecules of BBTV (genus Babuvirus) (Briddon et al., 2004). Not all BBTV isolates harbour alphasatellites, and those that do are predominantly from the Asian group of isolates (Bell et  al., 2002; Horser et  al., 2001). Their absence from many isolates provides strong evidence that they are not an integral part of the BBTV genome. The function of the BBTV alphasatellites in not yet known and their presence does not appear to affect the severity of symptoms (Bell et  al., 2002), in contrast to those of begomoviruses and nanoviruses (Briddon et al., 2004; Gronenborn, 2004). Interestingly, Bhadramurthy et al. (2008) reported integration of the complete BBTV CP gene in the genome of four varieties of cardamom (Elettaria cardamomum), with no evidence for transcription, or of the presence of virus particles. Strains of BBTV Two broad groups of isolates have been identified, based on nucleotide sequence differences

between genome components (Karan et  al., 1994). The ‘South Pacific’ group comprises isolates from Australia, the Pacific Islands, Africa and South Asia (including Myanmar), while the ‘Asian’ group comprises isolates from Southeast Asia, southern China, Taiwan and Japan. These differences are present throughout the genome (Stainton et  al., 2015), but no biological differences have been associated with these sequence differences. Despite this general correlation between geographical origin and sequence similarity, extensive recombination and reassortment has been detected between BBTV isolates, including those of the two ‘subgroups’ (Stainton et  al., 2012, 2015). Indeed, interspecies recombination between ABTV and BBTV has been identified (Stainton et al., 2012). Most isolates of BBTV are associated with typical severe disease symptoms. However, as mentioned above, mild and symptomless isolates have been reported from Taiwan (Su et al., 1993, 2003). BBTV has been confirmed in specimens of mild and symptomless infections from Taiwan by ELISA, PCR and sequencing (Su et al., 2003; Fu et  al., 2009; Stainton et  al., 2015) and the isolates can be transmitted by Pentalonia nigronervosa (H.J. Su, Taiwan, 1996, personal communication). Interestingly, the N component appears not to be present in a Taiwanese mild isolate. Magee (1948) noted that certain plants of ‘Veimama’, a Cavendish cultivar originally from Fiji and growing then in NSW, showed a ‘partial recovery’ from bunchy-top symptoms and produced bunches. After an initial flush of typical severe symptoms in three or four leaves, subsequent leaves showed few, if any, dark green flecks. Suckers derived from these partially recovered plants also displayed a flush of typical symptoms followed by partial recovery. The origin of the infection, whether from Australia or Fiji, was uncertain. This partial recovery was noted for some infected plants of ‘Veimama’ only, and in Fiji was noted for one sucker only on a single infected stool from among hundreds of infected stools of ‘Veimama’ observed. Magee was not able to transmit the virus from partially recovered plants and was only able to superinfect them, with difficulty, with high inoculum pressure. This may be an example of a mild strain of BBTV, possibly a nonaphid-­transmitted one, propagated vegetatively,



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reaching only a low titre and conferring a degree of cross-protection, or ‘Veimama’ may not be genetically uniform and individual plants with a degree of resistance may exist. The complete explanation for this phenomenon is unclear. A possible explanation for the above symptom variants may lie in the complement of genome components that they possess. The inability to transmit bunchy top from abacá to banana (Ocfemia and Buhay, 1934) was originally considered to be evidence that two distinct strains of the virus existed. However, as noted by Magee (1953), technical deficiencies in these experiments mean that the results must be viewed with caution. As first voiced by Magee (1953), the many similarities between the bunchy top diseases of banana and abacá, including transmission properties with the vector P. nigronervosa, suggests that both are caused by the same virus. Recent research, however, has shown that abacá and banana can each be infected by both ABTV and BBTV and that symptoms are indistinguishable (Sharman et  al., 2000b, 2008).

A variety of nucleic acid-based assays have been applied to the detection of BBTV in plant tissue and viruliferous aphids, including DNA and RNA probes, labelled either non-radioactively or with 32P (Hafner et  al., 1995; Xie and Hu, 1995). PCR has proved to be about 1000 times more sensitive than ELISA or dot blots with DNA probes (Xie and Hu, 1995). An alternative serological approach is impedance spectroscopy, which is reported to be 100 times more sensitive than ELISA (Majumder et  al., 2013). Substances in banana sap inhibitory to PCR can be circumvented by simple extraction procedures (Thomson and Dietzgen, 1995; Selvarajan et al., 2015) or by immunocapture PCR (IC-PCR) (Sharman et  al., 2000b). PCR products can be specifically detected with a lateral flow device providing increased specificity and sensitivity over gel electrophoresis of amplicons (Wei et al., 2014). Real-time PCR (Chen and Hu, 2013), loop-mediated isothermal amplification (LAMP) (Peng et al., 2012a; Selvarajan et al., 2015) and recombinase polymerase amplification (RPA) (Agdia, 2016) have also been applied in the ­detection of BBTV.

Detection of BBTV Prior to 1990, the only assays available for banana bunchy top were visual assessment of symptoms and aphid transmission to a sensitive banana cultivar. Subsequently, both serological and nucleic acid-based assays have become available. Polyclonal and monoclonal antibodies are now routinely used in ELISA to detect BBTV in field and tissue-culture plants and can detect the virus in single viruliferous aphids (Wu and Su, 1990b; Dietzgen and Thomas, 1991; Thomas and Dietzgen, 1991; Thomas et al., 1995). Triple antibody sandwich ELISA was found to be the optimum serological assay format for routine virus indexing (Geering and Thomas, 1996). All isolates tested from Africa, Australia, Asia and the Pacific region are serologically related (Thomas, 1991). Polyclonal antibodies to BBTV also cross react with ABTV (Sharman et  al., 2008) and cardamon bushy dwarf virus (CBDV) (Mandal, 2010), and some, but not all, monoclonal antibodies cross-react with the abacá virus (Sharman et al., 2008). BBTV is serologically unrelated to the nanovirus faba bean necrotic yellows virus (Katul et al., 1993).

Disease cycle and epidemiology BBTV is transmitted by an aphid vector and in vegetative planting material, but not by mechanical inoculation (Magee, 1927). Distribution and movement within the plant Magee (1927) showed that banana bunchy top was a systemic disease. Following aphid inoculation, symptoms generally do not appear until a further two or more leaves have been produced (Magee, 1927). This period can vary between 19 days in summer and 125 days in winter in the subtropics (Allen, 1978a). The virus can only be recovered by aphids from the first symptom leaf or those formed subsequently (Magee, 1940a). Suckers produced on an infected stool generally develop symptoms before reaching maturity (Magee, 1927). Magee (1939) also concluded that the virus was restricted to the phloem tissue. Microscopic examination revealed hypertrophy and hyperplasia of the phloem tissue and a reduction in the development of the fibrous sclerenchyma

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sheaths surrounding the vascular bundles. The cells surrounding the phloem contained abnormally large numbers of chloroplasts, giving rise to the macroscopic dark green streak symptom. Subsequent investigation, using RNA probes and PCR (Hafner et al., 1995), has demonstrated that BBTV replicates for a short period at the site of aphid inoculation, and then moves down the pseudostem to the basal meristem and finally to the corm, roots and newly formed leaves. Trace levels of virus were eventually detected by PCR in leaves formed prior to inoculation, but replication was not demonstrated. This latter observation is consistent with the inability to transmit the virus by aphids from such leaves (Magee, 1940a) and with the sequential development of single, new leaves from the basal meristem. BBTV has been detected by ELISA and/or PCR in most parts of the plant, including leaf lamina and midrib, pseudostem, corm, meristematic tissues, roots, fruit stalk and fruit rind (Thomas, 1991; Wu and Su, 1992; Hafner et al., 1995; A.D.W. Geering and J.E. Thomas, Australia, 1996, unpublished results). Aphid transmission In Australia, the black banana aphid (P. nigronervosa) (Plate 6.8) had been under suspicion as having a role in the aetiology of banana bunchy top (Darnell-Smith, 1924) and in 1925 it was

Plate 6.8.  Black banana aphid (Pentalonia nigronervosa) (photo: J.E. Thomas, QDAF).

conclusively demonstrated to be the vector (Magee, 1927). A recent taxonomic study has shown by morphological and molecular methods that P. nigronervosa f. sp. caladii should be resurrected as a distinct species, Pentalonia caladii (Foottit et  al., 2010). Although host records overlap, the former species predominantly colonizes Musa spp. while the latter predominantly colonizes other members of the Zingiberales. Both, however, were shown to be vectors of BBTV (Watanabe et al., 2013). The two species have a worldwide distribution, with a host range that includes Musa textilis and other species in the Musaceae. Species in several closely related plant families, including the Araceae (Alocasia sp., Calladium spp., Dieffenbachia spp., Xanthosma sp.), Cannaceae (Canna spp.), Heliconiaceae (Heliconia spp.), Strelitzeaceae (Strelitzia spp.) and Zingiberaceae (Alpinia spp., Costus sp., Hedychium spp.), are also colonized (Wardlaw, 1961; R.N. Allen, Australia, 1996, personal communication). However, the exact geographical distribution and the full normal host range of the two species remain to be determined (Foottit et al., 2010). A degree of host preference has been displayed in past transmission studies and some difficulty experienced transferring aphids between host species. This may well have been due to host preferences of the particular species used. Another related species, Micromyzus (syn. Pentalonia kalimpongensis), is recorded as a vector of CBDV (Mandal et  al., 2013) but appears not to have been tested as a potential vector of BBTV. On banana plants in NSW, aphids are found mostly at the base of the pseudostem at soil level and for several centimetres below the soil surface, beneath the outer leaf sheaths and on newly emerging suckers (R.N. Allen, Australia, 1996, personal communication). Similarly, Hawaiian studies detected aphids more frequently near the base of the plant, especially suckers < 1.5 m tall, and presented a sampling strategy (Robson et al., 2006; Hooks et al., 2011). Aphid numbers decrease during periods of drought (Wardlaw, 1961). Transmission of BBTV, the probable causal agent, is of the circulative, non-propagative type. Most transmission studies have probably been carried out with P. nigronervosa, when the colonies were sourced from banana, though this cannot be certain. The transmission parameters reported from Hawaii (Hu et al., 1996) and Australia



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(Magee, 1927), respectively, are: minimum acquisition access period 4 h/17 h; minimum inoculation access period 15 min/30 min–2 h; ­retention of infectivity after removal from virus source 13 days/20 days. A minimum latent period of at least ‘a few hours’ was recognized by Magee (1940a) and reported as a minimum of 20 h by Anhalt and Almeida (2008). No evidence was found for transmission of BBTV to the parthenogenetic offspring (Magee, 1940a; Hu et al., 1996) or for multiplication of BBTV in the aphid vector (Hafner et  al., 1995, Watanabe and Bressan, 2013). All life stages of the vector are able to transmit BBTV (Magee, 1940a). Transmission efficiency for individual aphids has been reported as ranging from 46% to 67% (Magee, 1927; Wu and Su, 1990a; Hu et al., 1996). The virus was more efficiently acquired and transmitted by nymphs than by adults in the work of Magee (1940a), though the reverse situation was reported by Anhalt and Almeida (2008). Temperature has been shown to have an effect on vector transmission rates. Transmission rates are significantly higher at 25–30°C than at 20°C, and are lower or cease at 10–16°C (Anhalt and Almeida, 2008; Magee, 1940a; Wu and Su, 1990a). Pentalonia nigronervosa and P. caladii have both been shown to transmit BBTV and to have similar tissue tropisms for the virus (Watanabe et al., 2013). Interestingly, the virus transmission properties have more in common with the geminivirids than luteovirids. BBTV was shown to accumulate in the anterior midgut (AMG), which had the highest concentration, haemolymph and principal salivary glands. As the AMG cells have a more prominent secretory than uptake activity, BBTV may not hijack an existing cellular process to traverse the anterior midgut, as suggested for luteovirids, but use an early endosome-­ independent process (Watanabe et  al., 2016). Translocation is rapid within the vector, and the observed direct contact between the anterior midgut and principal salivary gland suggests a possible direct route of transfer, bypassing the haemolymph (Watanabe and Bressan, 2013). In contrast to reports with geminivirids and luteovirids, it appears that the molecular chaperone protein, Buchnera GroEL, from P. nigronervosa does not react with BBTV in vitro or in vivo (Watanabe et al., 2013). Colonies of P. nigronervosa from Australia (where bunchy top occurs)

and from Réunion (where bunchy top does not occur) both transmitted each of six isolates of BBTV with similar efficiency (M.L. Iskra-Caruana, France, 1994, personal communication). Vegetative propagation Bunchy top is efficiently transmitted through conventional planting material, including corms, bits and suckers. All suckers from an infected stool will eventually become infected. Magee (1927) demonstrated 100% transmission of bunchy top through new ‘eyes’ (meristematic growing points), even in a plant that had only been expressing symptoms for 2–3 weeks. Bunchy top is also transmitted in micro-­ propagated banana plants (Drew et  al., 1989; Ramos and Zamora, 1990; Wu and Su, 1991), though not always at rates of 100%. From time to time, apparently virus-free meristems producing apparently virus-free plants can arise from an infected clone (Thomas et al., 1995). In Pakistan in the early 1990s, much of the available planting material of ‘Basrai’ (AAA, Cavendish subgroup) was infected with BBTV. Plantations established from infected suckers and corms were completely unproductive (Plate 6.9). Epidemiology of banana bunchy top The epidemiology of banana bunchy top in Australia is simplified by the presence of a single susceptible host and a single primary vector species (P. nigronervosa) (Magee, 1927). Pentalonia caladii is known to be present in Australia (Foottit et al., 2010), but limited sampling of banana aphids in subtropical Queensland and New South Wales has revealed only P. nigronervosa (M. Webb, K.S. Crew and J.E. Thomas, Australia, 2015, unpublished results). Spread over long distances is by infected planting material and it is by this means that new plantings in isolated areas usually become infected. Dissemination over short distances from these infection foci is by the ­banana aphid. In studies of actual outbreaks of bunchy top in commercial banana plantations in the subtropics, Allen (1978b, 1987) showed that the average distance of secondary spread of the disease by aphids was only 15.5–17.2 m. Nearly two-thirds of new infections were within 20 m of the nearest source of infection and 99% were

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Plate 6.9.  Plantation of ‘Basrai’ (AAA, Cavendish subgroup) affected by bunchy top disease in the Sindh Province of Pakistan (photo: D.R. Jones, INIBAP).

within 86 m. Allen and Barnier (1977) showed that, if a new plantation was located adjacent to a diseased plantation, the chance of spread of bunchy top into the new plantation within the first 12 months was 88%. This chance was reduced to 27% if the plantations were separated by 50–1000 m and to less than 5% if they were 1000 m apart. On average, the interval between infection of a plant and movement of aphids from this plant to initiate new infections elsewhere (the disease latent period) was equivalent to the time taken for 3.7 new leaves to emerge. The rate of leaf emergence varied seasonally, with a maximum in summer (Allen, 1987). Based on the above studies, a computer program that simulates epidemics of bunchy top was developed (R.N. Allen, Australia, 1994, personal communication) that allows epidemiological factors to be varied and their effect on the progress of the epidemic and disease control to be monitored. In Burundi, the disease incubation period was longer under cooler conditions at higher elevations: 21 days at 780 m versus 84 days at 2090 m, because of slower rates of leaf emergence. At both locations symptoms invariably

appeared in the second newly emerged leaf (Niyongere et al., 2013). In the Philippines, Opina and Milloren (1996) also demonstrated that most new infections were adjacent to or in close proximity to primary sources of infection. Smith et al. (1998) also described strong gradients of infection in a commercial plantation where uncontrolled adjacent external sources of infection were present. Niyongere et  al. (2013) noted that placing new plantings at distances as low as 5–30 m from existing affected plantings resulted in a measurable decrease in disease incidence.

Host reaction General host range In the Musaceae, BBTV is known to infect a range of Musa species, cultivars in the Eumusa and Australimusa series of edible banana and Ensete ventricosum. Susceptible Musa spp. include M. balbisiana (Magee, 1948; Espino et al., 1993), M. acuminata ssp. banksii and M. textilis (Magee,



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1927), M. velutina (Thomas and Dietzgen, 1991), M. coccinea, M. jackeyi, M. ornata and M.  acuminata ssp. zebrina (A.D.W. Geering and J.E. Thomas, Australia, 1998, unpublished results). There is some evidence for hosts outside the Musaceae, though reports have been conflicting. Su et  al. (1993) obtained positive ELISA reactions from BBTV-inoculated Canna indica and Hedychium coronarium; and recovery of the virus to banana, though not reported here, was demonstrated (H.J. Su, Taiwan, 1996, personal communication). Ram and Summanwar (1984) reported Colocasia esculenta as a host of BBTV. Pinili et  al. (2013) also reported C. indica and Co. esculenta and additionally Alpinia zerumbet as hosts. Alpinia purpurata has been recorded as a natural host in Tahiti (M. Wong, D. Massé and B. Horstachy, Tahiti and Reunion, 2017, personal communication). However, Hu et  al. (1996) were unable to demonstrate Co. esculenta or A. purpurata as experimental (E) or natural (N) hosts of BBTV in Hawaii. Geering and Thomas (1997) also found no evidence for the following species as hosts of BBTV in Australia: Strelitzia sp. (N), C. indica (E, N), Canna × generalis (N), Canna × orchiodes (N), H. coronarium (E), Heliconia psittacorum (E), Alpinia caerulea (E, N), Alpinia arundinelliana (E), A. zerumbet (E), Alocasia brisbanensis (E, N), Co. esculenta (E, N). Magee (1927) was unable to infect Strelitzia sp., Ravenala sp., Canna spp. (including C. edulis), Solanum tuberosum and Zea mays. Interestingly, the strain subgroup of BBTV used has not been consistent in these experiments, nor is the exact aphid species used known. These inconsistencies may contribute to these conflicting results. Cultivar susceptibility To date, there are no confirmed reports of immunity to BBTV in any Musa species or cultivar. However, differences in susceptibility and symptom severity between cultivars subject to either experimental or field infection have frequently been noted (Magee, 1948; Jose, 1981; Muharam, 1984; Espino et  al., 1993; Mware, 2016; Mwenebanda et  al., 2007; Niyongere et  al., 2011; Sachter-Smith, 2015). Espino et al. (1993) evaluated a total of 57 banana cultivars for their reaction to bunchy top, both by experimental inoculation and field observations. All cultivars in the AA and AAA

genomic groups were highly susceptible. However, low levels of infection (as assessed by symptom expression) or total absence of symptoms following aphid inoculation were noted in some cultivars containing the B genome. These included ‘Radja’ (AAB, syn. ‘Pisang Raja’ – 12.5% of inoculated plants with symptoms), ‘Bungaoisan’ (AAB, Plantain subgroup – 0% with symptoms), ‘Pelipia’ (ABB, syn. ‘Pelipita’ – 10% with symptoms), ‘Pundol’ (ABB, believed to be synonymous with ‘Pondol’ – 0% with symptoms), ‘Katali’ (ABB, Pisang Awak subgroup – 0% with symptoms), ‘Abuhon’ (ABB – 0% with symptoms) and ‘Turangkog’ (ABB – 0% with symptoms). These cultivars were not back-indexed by aphid transmission to a susceptible banana cultivar or biochemically indexed (e.g. by ELISA) and so the presence of symptomless infection cannot be ruled out. Also, greater numbers of aphids than the 15 used here might have resulted in infection. Cultivars ‘Abuhon’ and ‘Bungaoisan’ were susceptible to BBTV by experimental aphid inoculation with an Australian BBTV isolate (A.D.W. Geering and J.E. Thomas, Australia, 1996, unpublished). A field trial in Burundi using 40 cultivars representing a range of genotypes also revealed a wide range of susceptibility to BBTV, and the tendency for the less readily infected cultivars to contain a B genome component. However, as has been noted elsewhere, ‘Gros Michel’ (AAA) was an exception, and plants showed no symptoms at the end of the 28-month observation period, though they were shown by PCR to be infected (Niyongere et al., 2011). Similarly, at two field trial sites in Cameroon observed over a period of 37 months, a wide range of susceptibility was shown among 16 genotypes. Although most were plantains or cooking-banana cultivars (AAB or AAAB genotypes), three AAA dessert-banana genotypes were included. Once again, ‘Gros Michel’ showed a lower level of susceptibility than many other genotypes and, at the site with the lower infection pressure, was not infected within the duration of the trial (Ngatat et al., 2017). Some genotypes can be difficult to infect, but can be shown to be susceptible using increased numbers of infective aphids (Magee, 1948). Furuya et al. (2012) were unable to infect Musa balbisiana var. liukiuensis by aphid inoculation using 5–15 aphids per plant, and Mware

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(2016) reported no infection of banana cultivars ‘Saba’ (ABB), Musa balbisiana accession ‘Butuhan’ (BB) and Musa acuminata ssp. saimea accession ‘Kluai Khae’ (Phrae) (AA) using 20 aphids per plant. It would be interesting to see if larger numbers of aphids could result in infection. A.D.W. Geering and J.E. Thomas (Australia, 1996, unpublished results) noted that ‘Kluai Teparot’ (ABB) was not infected in three separate experiments using five, 12 and 12 plants, and five to six aphids per plant with one Australian BBTV isolate. However, using a second isolate, one plant in five was infected using 15 aphids per plant, and five of five were infected using 50 aphids per plant. Cultivars within the Cavendish subgroup form the basis of the international banana export trade and are generally highly susceptible to bunchy top. However, it appears that not all cultivars with an AAA genome are similarly susceptible. ‘Gros Michel’ exhibits resistance to the disease under both experimental inoculation and field conditions (Magee, 1948; Vakili, 1969; Niyongere et  al., 2011; Kumar et  al., 2015, Ngatat et al., 2017; A.D.W. Geering and J.E. Thomas, Australia, 1996, unpublished information). Magee (1948) considered that the introduction of this cultivar to Fiji in the early 20th century contributed to partial rehabilitadevastated industry. tion of the bunchy-top-­ Compared with ‘Williams’ (AAA, Cavendish subgroup), the concentration of virions of BBTV in infected plants of ‘Gros Michel’ and the proportion of plants infected by aphid inoculation are lower. Symptoms are also slower to develop and are less severe (A.D.W. Geering and J.E. Thomas, Australia, 1996, unpublished results). These factors may contribute to a reduced rate of aphid transmission and field spread in plantations of ‘Gros Michel’. Overall, it appears that real differences exist in cultivar reaction to BBTV and for the time taken before symptoms are expressed – factors that can combine to significantly affect the rate of spread of the disease.

Control No natural resistance to BBTV is known from edible banana cultivars and landraces, though

some differences in susceptibility and symptom expression have been noted. At present, control relies heavily on cultural practices, including the use of clean planting material. Cultural practices On the basis of his research in the early 1920s, Magee (1927) proposed a range of measures for the control of banana bunchy top. These recommendations involved two major components:

• •

Exclusion of the disease from unaffected and lightly affected areas. Eradication of infected plants from both lightly and heavily affected areas.

Magee considered that the measures would fail if left entirely to the goodwill of the growers (Magee, 1936) and so legislation enforcing these control measures was gradually introduced by the governments of New South Wales (Plant Diseases Act and Regulations commencing 1927) and Queensland (Diseases in Plants Act and Banana Industry Protection Act commencing 1929). The legislation in the two states was similar, the major difference being the onus of detection on the growers in Queensland and on the government-funded inspectors in NSW. These measures, which are listed below, still form the basis of control today:

• • • • • •

Registration of all banana plantations. Establishment of quarantine zones. Restrictions on the movement and use of planting material. Regular inspections of all banana plantations for bunchy top. Prompt destruction of all infected plants. Ongoing education and extension programmes for growers.

When adopted, these measures allowed the complete rehabilitation of the Australian ­banana industry. A vital aspect of eradication was the complete removal of infected stools to prevent the re-emergence of infected suckers. When identified, the infected plant was first sprayed with paraffin to kill any aphids. The plant was then chopped down, the corm and all associated suckers dug out and all plant material chopped into small pieces to prevent regrowth. Nowadays, herbicide and insecticide injections have replaced physical removal and destruction.



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A steady rehabilitation and expansion of the Australian banana industry followed in the late 1920s and early 1930s. However, a period of complacency, due to the success of the control programme, combined with a period of overproduction and resultant neglect of unprofitable plantations from about 1933 to 1935, led to a resurgence of bunchy top (Magee, 1936). An intensification of the control programme and increased vigilance by the disease inspectors brought the situation under control again, where it has remained ever since. Allen (1987) evaluated control measures in NSW by computer simulation and showed that bunchy top could be controlled because of the long interval between first appearance of disease symptoms (about two new leaves) and first transmission to other plants (a further 1.7 new leaves). He recommended that inspection intervals be varied according to the growth rate of the plants, with successive inspections made before the growth of four additional leaves. In Australia in a few cases, total eradication of BBTV from districts has been achieved (e.g. Yarrahappini and Richmond Valley areas of NSW and Innisfail, Queensland), but, even with a large input of resources, the disease is usually just reduced to very low, manageable levels. Few other countries have experienced such successful control of bunchy top. In most cases, this has been due to an inability to enforce an organized control programme across whole districts and the lack of a reliable, ongoing source of clean planting material. In addition to plantations, feral banana plants and plants in home gardens must be taken into consideration if bunchy top is to be successfully controlled. Smith et al. (1998) studied a bunchy top epidemic in a commercial banana plantation in Mindanao, Philippines. They found that, despite regular inspection and roguing of infected plants, a clear gradient of infection was observed from the edge of the plantation towards the centre. This could only be explained by a significant, continuing source of infection, probably from banana plots on smallholdings and feral banana plants, in close proximity to the plantation. Attempts to control bunchy top by controlling the aphid vector with insecticide spray programmes have usually met with limited success, due in part to the concealed niches they

occupy. Robson et al. (2007) reported that imidacloprid was effective at controlling aphids upon contact, but that it did not become systemic in the banana plant. Parasitoids have been identified as potential biocontrol agents for the banana aphid (Völkl et al., 1990), but the practice has not been implemented. Natural resistance There is no known natural immunity to BBTV, but cultivars and species differ widely in symptom expression and susceptibility and many cultivars with the B genome have been noted to be less susceptible (see above). Further investigation and utilization of natural sources of tolerance and reduced susceptibility to the disease are warranted. biopriming.  Pot and field trials with rhizosphere and endophytic bacteria using pre- and post-­ planting applications were reported to induce systemic resistance to BBTV, with significant reductions in both percentage infection and virus titre reported (Kavino, 2007a, b, 2008; Harish et al., 2008, 2009). Levels of defence-related enzymes and pathogenesis-­ related proteins were elevated and in field trials enhanced plant growth and fruit yield were also reported. However, only the positive effects on plant growth, but not on virus resistance, were supported by independent research (R. Selvarajan, India, 2016, personal ­communication). somaclonal variants and mutagenesis. 

Putative BBTV-resistant lines of ‘Lakatan’ (AAA) produced by gamma irradiation (Damasco et al., 2006) and through selection of somaclonal variants (Damasco et al., 2008) have been reported from the Philippines and when described were being assessed for the stability of resistance and mechanism of resistance, respectively. Potentially BBTV-resistant abaca lines created through gamma irradiation have also been reported (Dizon et al., 2012). However, there appear to be no follow-up reports on the effectiveness of these approaches. resistance.  In the absence of known sources of natural immunity to BBTV,

transgenic

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focus has been placed on transgenic resistance and a number of different approaches are being investigated. A strategy called InPAct technology has been developed at the Queensland University of Technology (Dale et  al., 2004) and is based on virus-activated cell death mediated by barnase, a cytotoxic RNase from Bacillus amyloliquefaciens. The plant is transformed with a cassette containing a split ‘suicide gene’ with the 3′ and 5′ portions inverted, which is flanked by introns which in turn are flanked by the BBTV intergenic region containing the viral origin of replication (ori). Upon infection by BBTV, the viral Rep nicks the insert at the two ori sites, releasing, circularizing and allowing replication of the cassette. The transcript is processed, leading to removal of the introns and ori, and translation to produce active barnase with subsequent cell death and inability of the virus to move from the inoculated cell. Some challenges with the use of this suicide gene approach include cryptic intron splice sites, cytotoxicity of Rep and Clink proteins, cassette size limits, inserts that are released but not replicated, and high background barnase expression (Bolton, 2009). An alternative strategy is to express defective Rep proteins in the transformed plants, which should outcompete native virus Rep for binding to the infecting virus DNA and thereby greatly reduce or prevent viral replication. At the University of Hawai‘i, modified Rep transformants have been developed and are in the process of field evaluation. These include mutant, antisense, partial Rep genes and inverted repeat of partial Rep gene (Borth et al., 2011). The RNAi strategy based on Rep gene sequences has been shown to be effective in glasshouse trials in India (Shekhawat et  al., 2012; Elayabalan et  al., 2013). By contrast, using an Australian isolate of BBTV, RNAi hairpin cassettes targeting the DNA-R or DNA-N genes provided no resistance or at best very limited tolerance to the virus. However, about 85% of transformed banana lines containing RNAi cassettes targeting the DNA-M gene remained symptomless for up to 10 months post inoculation and tested negative for the presence of BBTV by PCR. Confined field testing of transgenic bananas with RNAi-based BBTV resistance has been approved for Malawi from 2016–2018 (Mkoko, 2016; Mware et al., 2016).

Abacá Bunchy Top J.E. Thomas Introduction Abacá bunchy top disease can be caused by two distinct viruses. One is banana bunchy top virus (BBTV) described above. The other, which is closely related to BBTV, has only recently been discovered. It produces symptoms in banana and abacá similar to those caused by BBTV (Sharman et al., 2008). It has been recorded only from the Philippines and Malaysia (Sarawak), though no targeted surveys have been reported. History and early investigations Bunchy top symptoms on abacá were first recognized in 1910 in Albay province in the Philippines (Ocfemia, 1926) and now abacá bunchy top disease (ABTD) occurs in all major production areas of the country (Raymundo and Bajet, 2000). It has long caused major losses to abacá production in the Philippines, and was a major factor in the decline of abacá exports (Raymundo, 2000; Raymundo et al., 2001). It was originally concluded that ABTD was caused by BBTV (Magee, 1953) because of the similarity of symptoms between ABTD and banana bunchy top disease (BBTD) (Ocfemia, 1931; Magee, 1953), transmission by a common vector, the banana aphid Pentalonia nigronervosa (Ocfemia, 1927, 1931), and the induction of typical ABTD symptoms in abacá infected with BBTV isolates from banana (Magee, 1927). However, the discovery that the causal agent could be a different virus (Sharman et  al., 2008) means that the pathogen associated with much of the earlier work on these two diseases remains equivocal. To date, little work has been undertaken to determine the relative importance of the newly discovered virus affecting banana and abacá production (Sta. Cruz et al., 2016), though in the few instances where molecular analysis has been applied to BBTD samples from the Philippines, only BBTV predominates. Symptoms The symptoms caused by ABTV and BBTV are indistinguishable. Su et al. (2003) noted severe,



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typical symptoms of bunchy top disease on an infected banana in the Cavendish subgroup (AAA) including stunting, vein clearing, reduced leaf size, bunching of young leaves and dark green streaks on the pseudostem. Abacá plants confirmed to be infected with the virus (Sharman et al., 2008) showed vein clearing flecks and narrow, brittle leaves with chlorotic, upturned margins, as previously described for abacá bunchy top disease (Ocfemia, 1930). Causal agent Abacá bunchy top virus (ABTV), genus Babuvirus, family Nanoviridae, can cause ABTD. ABTV has a multi-component genome, comprising six circular ssDNA components, each between 1013 and 1099 nucleotides (nt) long (Sharman et  al., 2008). Though no functional analyses have been done, the genome appears typical of babuviruses. By sequence similarity to BBTV, it comprises DNA-R encoding the master replication initiation protein (Rep), DNA-S encoding the coat protein (CP), DNA-M and DNA-N encoding the movement protein and the nuclear shuttle protein, respectively, and DNA-C encoding a cell cycle link protein and DNA-U3, of unknown function. All components have a similar genome organization with a single ORF (Sharman et al., 2008) and replication is likely to occur via a rolling circle mechanism. The components also share two common regions: a major common region (CR-M), the presumed site of DNA primer binding for dsDNA replicative form synthesis, and a stem-loop common region (CR-SL) containing the origin of replication. Three short repeated sequences (iterons F1, F2 and R1) are contained within the CR-SL. These are presumed to be the binding site for the Rep, allowing trans-replication of the other genome components. Only two isolates of ABTV, one from banana and one from abacá, have been characterized at the molecular level (Sharman et al., 2008). Across the entire genome, they share ca 81% identity, and ca 59% and 63% identity with cardamom bushy dwarf virus (CBDV) and BBTV, respectively. In contrast to BBTV and CBDV, no satellite molecules have yet been found associated with ABTV. Some isolates of BBTV obtained from abacá have erroneously been given the name ABTV (e.g. FJ787433, FJ787434, FJ787435). It must be

stressed that the identity of the virus (ABTV or BBTV) cannot be assumed on the basis of the host from which it was isolated, or from ELISA tests using antisera which may cross-react. Molecular analysis is required for definitive identification. Epidemiology and disease cycle No information has been reported on the epidemiology of ABTV, but by analogy with BBTV it is presumed to be transmitted in the circulative, non-­propagative manner by its aphid vector Pentalonia nigronervosa (Su et al., 2003) (Plate 6.10), and to be transmitted by vegetative propagation through suckers, but not through true seed. Host reaction The virus has been recorded from abacá and banana. No other hosts are known.

Plate 6.10.  Black banana aphids (Pentalonia nigronervosa), vectors of ABTV, feeding on the basal portion of abacá leaf petioles where they emerge from the pseudostem. (photo: E.O. Lomerio, FIDA).

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Control Again, by analogy with BBTV, the use of clean planting material will likely form the basis of control, as no resistance to ABTV has yet been reported.

Bract Mosaic J.E. Thomas, R. Selvarajan, M.-L. Iskra-Caruana, L.V. Magnaye and D.R. Jones

and M. ornata were observed by D.R. Jones to have symptoms at the Kerala Agricultural University’s Banana Research Station at Kannara, near Trichur. Bract mosaic was confirmed from specimens from India sent to Brisbane, Australia (Infomusa, 1995). ‘Embul’ (AAB, Mysore subgroup), seen by D.R. Jones with bract mosaic symptoms at the Horticultural Research and Development Institute’s field station at Walgolla, Sri Lanka, was also found to be affected by the disease following diagnosis (Infomusa, 1995). Symptoms of bract mosaic were also seen in a germplasm collection and in backyard gardens at Gannoruwa near Kandy (Thomas et al., 1999a).

Introduction Losses Bract mosaic disease has a restricted international distribution. It is economically important in the Philippines, India and Sri Lanka. History Symptoms of bract mosaic disease of banana were first noted on several banana cultivars on the Philippine island of Mindanao in 1979 and thought to be different from all other recognized viruses of banana (Magnaye and Espino, 1990). Many accessions in the Southeast Asian Regional Germplasm Collection near Davao were also seen with symptoms. Later, the disease was found to be widespread throughout the Philippines, particularly in plantings of the ABB cooking-banana cultivars ‘Saba’ and ‘Cardaba’. In Mindanao, cultivars in the Cavendish subgroup (AAA) on commercial plantations were also affected. In 1992, D.R. Jones noticed cultivars in the germplasm collections at the Indian Institute of Horticultural Research near Bangalore and at the Tamil Nadu Agricultural University in Coimbatore in India with symptoms of bract mosaic. The disease was also seen in field plots near Coimbatore and Tiruchchirapalli. Filamentous virus particles were found in specimens from affected plants sent for diagnosis (INIBAP, 1993). In 1995 in Kerala State, bract mosaic symptoms were seen by D.R. Jones on plants of cultivar ‘Nendran’ (AAB, Plantain subgroup) affected by a disease of hitherto unknown aetiology known locally as ‘kokkan’ (Samraj et  al., 1966; Shanmugavelu et al., 1992). Other banana cultivars

Data on the economic impact of bract mosaic on banana are limited. In one study conducted in Mindanao, yield losses of up to 40% were noted in cultivars ‘Cardaba’ and ‘Lakatan’ (AAA) (Kenyon et al., 1996; Thomas and Magnaye, 1996). In commercial Cavendish plantations in Mindanao, a correlation between high incidence of bract mosaic and high fruit rejection rates due to misshapen fingers has been noted (Thomas et al., 1999a). Streaks on fruit can also result in rejection. The failure of fruit to fill on infected banana plants has been noted in India (Thomas, 2015). Here, bract mosaic disease causes yield losses of 40–70%, depending upon the cultivar and season (Cherian et  al., 2002; Selvarajan and Jeyabaskaran, 2006). Based on the disease incidence from surveys and the yield loss assessment studies, losses of US$5.8 million per annum are estimated in cultivar ‘Nendran’ alone (Selvarajan et al., 1997). The infected plants showed significant reduction in height, girth, leaf area, finger weight and girth over healthy plants (Selvarajan and Singh, 1997). Geographical distribution The virus causing bract mosaic disease has been recorded from symptomatic banana from the Philippines, India and Sri Lanka (Thomas, 2015) and recently it has been reported to occur in Ecuador and Colombia on banana (Quito-Avila et  al., 2013). Additional records from Thailand and Vietnam have been from banana plants



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containing the virus, but displaying symptoms more typical of banana mosaic. In Western Samoa, the virus was recovered from plants with banana streak symptoms (Rodoni et  al., 1999) and in Taiwan from symptomless plants (Feng, 2006). In Hawaii, the virus has been reported from Alpinia purpurata, but not yet banana (Wang et al., 2010).

Symptoms Typical symptoms of bract mosaic disease on banana are distinctive, but can be influenced by banana genotype as well as other viruses. Mosaic patterns on bracts are diagnostic and distinct from symptoms caused by all other known viruses of banana (Plates 6.11 and 6.12). Mosaic patterns, stripes and spindle-shaped streaks may also be visible on pseudostem bases when the

Plate 6.11.  Mosaic symptoms of bract mosaic disease in a bract from the male bud of ‘Seenikehel’ (ABB) growing in a back garden in Gannoruwa, Sri Lanka (photo: D.R. Jones, INIBAP).

outer leaf sheaths are removed and can extend up the petiole bases. Infection is often associated with an increase in pseudostem pigment. Sometimes the symptoms are chlorotic on a red background (Plate 6.13) and sometimes reddish, yellow or chlorotic on a green background (Plates 6.14 and 15). Symptoms can darken through red to brown and even black (Plate 6.16). Chlorotic streaks and spindle shapes running parallel to the veins are occasionally seen on leaves (Plate 6.17). ‘Nendran’ in particularly is severely affected by the disease, with leaf sheaths separating from the unusually red-coloured pseudostems of young plants. Leaves also become arranged fan-like on one plane (see Plate 6.12), rather like the leaves of the traveller’s palm (Ravenala madagascariensis) . Suckering is also suppressed and suckers that do emerge are

Plate 6.12.  Mosaic symptoms of bract mosaic disease on emerging bracts of ‘Nendran’ (AAB, Plantain subgroup) in a small plantation near Thrissur in India. Another symptom is the fan-like arrangement of leaves. In India, the bract mosaic disease problem was known as ‘kokkan’ (photo: D.R. Jones, INIBAP).

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Plate 6.13.  Chlorotic, spindle-shaped streak ­symptoms of bract mosaic disease on the abnormally red-pigmented pseudostem of ‘Kippu Kadali’ (AB) growing in a germplasm collection of the Tamil Nadu Agricultural University, Coimbatore, India (photo: D.R. Jones, INIBAP).

Plate 6.14.  Yellow spindle-shaped streak symptoms of bract mosaic disease on the deep green pseudostem of ‘Lambi’ (ABB, Bluggoe subgroup) growing in a germplasm collection of the Tamil Nadu Agricultural University, Coimbatore, India (photo: D.R. Jones, INIBAP).

distorted and deeply pigmented (Plate 6.18) (Infomusa, 1995). In the Philippines, chlorotic streaks may be present on peduncles and a high disease incidence is associated with increased levels of malformed fruit in commercial plantations (Thomas et  al., 1999a). In India, petioles and peduncles of ‘Nendran’ become brittle and fruit is only rarely carried to maturity. If fruit does mature, it is undersized. In cultivar ‘Rasthali’ (AAB, Silk subgroup), the infected fruits exhibit necrotic streaks and patches in the peel (Plate 6.19). Subtle mosaics can be seen on the fruit of other cultivars (Plate 6.20). Leaf symptoms, consisting of spindle-­ shaped lesions and streaks running parallel to the veins, are not always evident, but can occur on young plants that have been recently infected. Symptoms in recently aphid-inoculated plants of ‘Lakatan’ (AAA) include broad chlorotic

patches or streaks, sometimes spindle-shaped, along the major leaf veins, (L.V. Magnaye and L. Herradura, Philippines, 1997, unpublished results). Iskra-Caruana et  al. (2008) observed a transient light mosaic on the petiole of newly aphid-inoculated plants of a Cavendish cultivar that completely disappeared until the development of typical spindle-shaped lesions on the final leaf, and streaks on the bracts. In their studies, banana plants from five different accessions either collected in the field or maintained in the greenhouse and continuously showing symptoms were found to be co-infected with banana mild mosaic virus (BanMMV). However, some plants with typical bract mosaic foliar symptoms and not infected with BanMMV have been ­recorded from Sri Lanka and the Philippines (J.E.  Thomas, K.S. Crew and M. Sharman, ­Australia, 2015, unpublished results).



Diseases Caused by Viruses 381

Plate 6.15.  Red spindle-shaped streak symptoms of bract mosaic disease extending up the petiole bases of ‘Suwandel’ (ABB) near Kandy, Sri Lanka (photo: D.R. Jones, INIBAP).

Plate 6.16.  Dark brown streak symptoms of bract mosaic disease on petiole bases of ‘Nendran’ (AAB, Plantain subgroup) near Thrissur, India (photo: D.R. Jones, INIBAP).

In Western Samoa, India (Tamil Nadu and Maharashtra States) and Vietnam, the causal agent has been isolated from banana plants that were showing symptoms typical of banana mosaic and lacking the characteristic symptoms on the bracts (Rodoni et al., 1997, 1999). Some of these plants were shown to have mixed infections with cucumber mosaic virus (CMV).

Particle and genome properties

Plate 6.17.  Chlorotic spindle-shaped streak symptom of bract mosaic disease on a leaf of ‘Nendran’ (AAB, Plantain subgroup) near Thrissur, Kerala, India (photo: D.R. Jones, INIBAP).

Banana bract mosaic disease (BBrMD) is caused by banana bract mosaic virus (BBrMV). Flexuous virus-like particles, each ca 750 nm × 11 nm, have been detected in infected banana (Muñez, 1992; Bateson and Dale, 1995; Thomas et  al., 1997). Purified virions (Plate 6.21) contain a major coat protein of ca 39 kDa (Bateson and Dale, 1995; Thomas et al., 1997). Virions have

a buoyant density in caesium chloride of 1.29– 1.31 g cm−3 and an A260/280 = 1.17 (Thomas et al., 1997). Nucleotide sequence analysis indicates that BBrMV is a distinct potyvirus. The virion encapsidates a monopartite ssRNA genome of 9711 bp in length (Ha et al., 2008). The virus

Causal agent

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Plate 6.19.  Fruit of ‘Rasthali’ (AAB, Silk subgroup) with necrotic patches caused by banana bract mosaic virus (photo: R. Selvarajan, ICAR-NRCB) Plate 6.18.  Distorted and deeply red-pigmented peeper of ‘Nendran’ (AAB, Plantain subgroup) affected by bract mosaic disease at Kannara near Thrissur, Kerala, India (photo: D.R. Jones, INIBAP).

contains a typical large ORF coding for a polyprotein of 3125 amino acids. It contains nine protease cleavage sites, yielding ten matured functional proteins that have motifs conserved among homologous proteins of other potyviruses (Ha et al., 2008; Balasubramanian and Selvarajan, 2012). In addition, a small ORF termed pipo encoding a 7 kDa protein has also been found in BBrMV isolates. The whole genome of BBrMV-TRY (India) and BBrMV-PHI (Philippines) had 94% nucleotide (nt) sequence identity and 88–98% amino acid (aa) sequence identities (Balasubramanian and Selvarajan, 2012). ­Genetic diversity and recombination analysis in the CP gene of 49 isolates of BBrMV revealed a great variation among them and two of the isolates from India were distinct with 18–21% divergence at the nt level and 12–20% divergence at aa level (Balasubramanian and Selvarajan, 2014a). Sequence identity for the helper component protease (HC-Pro) region of BBrMV isolates

was 92–100% at both the nt and aa level (­Balasubramanian et al., 2014). Isolates of BBrMV from India, Sri Lanka and the Philippines are all closely related serologically (Thomas et al., 1997). Weak serological relationships have been demonstrated between BBrMV and other potyviruses, including sugarcane mosaic virus (abacá mosaic strain), dasheen mosaic virus, maize dwarf mosaic virus, wheat streak mosaic virus and sorghum mosaic virus (Thomas et al., 1997). Although bract mosaic-affected banana plants from India and the Philippines sometimes also contain particles of banana mild mosaic virus (Iskra-Caruana et al., 2008; J.E. Thomas and K.S. Crew, Australia, 2015, unpublished results), BBrMV alone appears to be the causal agent of the disease because its virions have been aphid-­ transmitted to healthy banana test plants in which BBrMD has subsequently developed (Caruana and Galzi, 1998; Iskra-Caruana et al., 2008). Detection of BBrMV Virion concentration in infected plants can be relatively low, and the virions are usually not



Diseases Caused by Viruses 383

Plate 6.20.  Fruit of ‘Karpuravalli’ (ABB, Pisang Awak subgroup) near Coimbatore, Tamil Nadu, India with faint mosaic symptoms of bract mosaic disease (photo: D.R. Jones, INIBAP).

Both serological and nucleic acid-based assays are now available for BBrMV. The virus can be detected by ELISA using polyclonal (Thomas et  al., 1997) and/or monoclonal antibodies (J.E.  Thomas, Australia, 1996, unpublished results). The polyclonal antiserum detects all isolates tested (Thomas et al., 1997), though some individual monoclonal antibodies do not react with all isolates (J.E. Thomas, Australia, 1996, unpublished results). The virus can also be detected by reverse transcription (RT)-PCR in total nucleic acid extracts from infected plants, using either virus-­s pecific or degenerate potyvirus genus primers (Bateson and Dale, 1995; Rodoni et al., 1997, 1999; Thomas et al., 1997; Selvarajan and Balasubramanian, 2013). An immunocapture (IC) step with crude sap extracts to replace nucleic acid extracts has been combined in (multiplex) RT-PCR (Sharman et al., 2000b; Selvarajan and Balasubramanian, 2013; Iskra-Caruana et  al., 2008). IC one-step RT-PCR assay enabled the detection of BBrMV in leaf extracts diluted up to 1 × 10–10 (Iskra-Caruana et al., 2008). Quantitative real-time reverse transcription-PCR (RT-qPCR) was reported for the detection of BBrMV using the SYBR Green and TaqMan® chemistry (Wen et al., 2009; Siljo et al., 2014; Balasubramanian and Selvarajan, 2014b). A  reverse transcription loop-mediated isothermal amplification (RT-LAMP) assay was developed for BBrMV. The detection limit for RT-LAMP was similar to that for real time RT-PCR, and up to 100 times more sensitive than conventional RT-PCR (Siljo and Bhat, 2014). An IC-RT-LAMP has also been developed (Zhang et al., 2016).

Disease cycle and epidemiology Plate 6.21.  Virions of banana bract mosaic virus negatively stained with ammonium molybdate. Scale bar represents 200 nm (photo: J.E. Thomas, QDAF).

readily detected by direct electron microscopy of sap. The concentration of virions in the symptomatic bracts is relatively higher than in leaves, leaf sheaths and midribs showing symptoms (R. Selvarajan, India, 2016, unpublished results).

BBrMV is transmitted by at least three species of aphids: Aphis gossypii, Rhopalosiphum maidis (Magnaye and Espino, 1990) and Pentalonia nigronervosa (Muñez, 1992; Thomas, 2015). Pentalonia nigronervosa transmitted BBrMV after an acquisition access period of 1 min, indicating that transmission is of the non-­ persistent type (Muñez, 1992). Efficiency of transmission was less than 10% (Caruana and Galzi, 1998).

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Attempts to transmit BBrMV by sap inoculation to herbaceous indicator plants have so far been unsuccessful (Magnaye and Espino, 1990; Muñez, 1992; Thomas et  al., 1999a; Zhang et al., 2016). However, occasional sap transmission from banana to banana has been achieved (L.V. Magnaye and L. Herradura, Phillipines, 1998, personal communication). The virus can be transmitted through vegetative planting material, including suckers, bits and corms, and via micropropagated plantlets.

reported to have acquired putative resistance to BBrMV following in vitro mutagenesis by gamma-­ irradiation (Dizon et  al., 2012), though these lines have yet to be field tested or evaluated for other agronomic traits.

Host reaction

Introduction

The virus has been detected in a wide range of naturally infected banana cultivars and genotypes (Musa sp.) and no resistance to the virus has been noted. Abacá (M. textilis) is also a host (Thomas et  al., 1997; Sharman et  al., 2000a). Most of the banana cultivars grown in India are known to be susceptible to BBrMV. Occurrence of BBrMV has been reported in small cardamom, Elettaria cardamomum, in India (Siljo et al., 2012) and flowering ginger, Alpinia purpurata, a popular cut flower and tropical landscape plant in ­Hawaii (Wang et  al., 2010). Bract mosaic symptoms were also noted in 2008 on a male bud of Musa velutina from Andhra Pradesh, ­India (R.  Selvarajan, India, 2008, unpublished ­results).

This disease was first described in New South Wales, Australia, in 1930 (Magee, 1930, 1940b). It has been given a variety of names, including infectious chlorosis, heart rot, virus sheath rot, cucumber mosaic and banana mosaic (Magee, 1930; Stover, 1972; Wardlaw, 1972). Banana mosaic is widespread in distribution and assumed to be found in most areas where banana is grown. It is usually only a nuisance to growers establishing new plantings using corms or suckers. However, the disease can be a problem in plantations that are established using tissue culture-derived planting material (Tsai et al., 1986; Niblett et al., 1994). Different strains of the virus pathogen are known to occur. Common strains have not been reported to cause economically important damage to banana. Mosaic symptoms may be absent or occur on only a few leaves. In contrast, severe or heart rot strains cause significant losses due to necrosis of the pseudostem, which results in plant death (Niblett et al., 1994). The heart rot strain is particularly destructive in banana grown under plastic in Morocco (Bouhida and Lockhart, 1990).

Control The use of indexed, virus-free planting material is the best means of control. Roguing/sanitation programmes have been introduced into commercial production areas of banana in the Philippines (Magnaye, 1994). In India, farmers are reluctant to remove the BrMD-affected plants as they yield marketable bunches in the first cycle of virus infection. ­Application of high levels of fertilizer is reported to compensate the yield loss due to BBrMD in the plant crop of ‘Ney Poovan’ and ‘Robusta’ (Selvarajan et  al., 2009, 2012). However, this management strategy can only be adopted where vectors are not present or are controlled with systemic insecticide sprays. Recently, abacá cultivars ‘Tinawagan Pula’ and ‘Tangongon’ were

Banana Mosaic B.E.L. Lockhart, D.R. Jones and J.E. Thomas

Symptoms A wide range of symptoms may be expressed (Ayo-John et al., 2008) and can depend on the strain of the virus pathogen and the temperature. Common or mild strains induce a diffuse mosaic (Plate 6.22) or line patterns and ringspots of the leaf lamina (Plate 6.23) (­Yot-­Dauthy and Bové, 1966; Lockhart, 1986; ­Niblett et al.,



Diseases Caused by Viruses 385

Plate 6.22.  Chlorotic mosaic symptoms of banana mosaic disease on a leaf of ‘Grand Nain’(AAA, Cavendish subgroup) on the island of St Vincent, Windward Islands (photo: D.R. Jones, SVBGA).

Plate 6.23.  Chlorotic line pattern and ring spot symptoms of banana mosaic disease on a leaf of ‘Pisang Mas’ (AA, Sucrier subgroup) growing in Melaka Sate, West Malaysia (photo: D.R. Jones, QDPI).

1994). ­ Occasionally, leaves can be deformed and curl (Plate 6.24). These symptoms appear sporadically and the majority of leaves may be symptomless. Chlorotic mosaics sometimes appear on fruit (Plate 6.25). Symptoms are generally more severe when temperatures fall below 24°C, which occurs in the winter in the subtropics and at altitude in the tropics (Niblett et al., 1994). Severe strains of the banana mosaic virus produce more pronounced symptoms, which can include necrosis of emerging cigar leaves (Plate 6.26) leading to varying degrees of necrosis in the unfurled leaf lamina (Plate 6.27). Internal tissues of the pseudostem can also become necrotic. Leaf distortion is also more severe (Plate 6.26). Plants with severe strains of the virus may die, especially if infected soon after planting.

Plate 6.24.  Distortion of the leaf lamina of ‘Grand Nain’ (AAA, Cavendish subgroup) caused by cucumber mosaic virus on the island of St Vincent, Windward Islands (photo: D.R. Jones, SVBGA).

Plate 6.25.  Yellow mosaic symptoms of banana mosaic disease on fruit of ‘Robusta’ (AAA, Cavendish subgroup) on the island of St Vincent, Windward Islands (photo D.R. Jones, SVBGA).

Causal agent Cucumber mosaic virus (CMV) causes mosaic disease of banana (Yot-Dauthy and Bové, 1966).

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Plate 6.27.  Necrosis of a leaf of ‘Grand Nain’ (AAA, Cavendish subgroup) caused by cucumber mosaic virus on the island of St Vincent, Windward islands (photo: D.R. Jones, SVBGA).

Plate 6.26.  Necrosis of the cigar leaf and the pronounced deformation of other leaves of ‘Grand Nain’ (AAA, Cavendish subgroup) as the result of an infection with a severe strain of cucumber mosaic virus on the island of St Vincent, Windward Islands (photo: D.R. Jones, SVBGA).

This virus is a member of the genus Cucumovirus, family Bromoviridae, and has isometric ­particles 28–30 nm in diameter (Plate 6.28) and a  single-stranded multipartite RNA genome (Francki et al., 1979; Jacquemond, 2012) with a genome of total size of ca 8.6 kb (King et  al., 2012). Most isolates of CMV have three genomic and one subgenomic RNA component (Francki et al., 1979). A fifth RNA species (RNA 5) (Kaper and Waterworth, 1977) occurs in some virus isolates and has been linked to modulation of symptom expression in some plants including banana (Gafny et al., 1996). Isolates of CMV can be separated into two subgroups based on serological relationships, nucleotide sequences and phylogenetic analysis. These two subgroups are designated subgroup I (or DTL, WT) and subgroup II (or ToRs, S) (Devergne and Cardin, 1973; Piazolla et al., 1979; Owen et  al., 1990; Bald-Blume et  al., 2017).

Plate 6.28.  Virions of cucumber mosaic virus stained with uranyl acetate (photo: B.E.L. Lockhart, UM).

Isolates of CMV from banana predominantly belong to subgroup I (Hu et  al., 1995; Singh et al., 1995; Gafny et al., 1996; Ayo-John et al., 2008), which is the subgroup that includes most CMV isolates from the tropics (Niblett et  al., 1994). However, subgroup II isolates from banana are also recognized (Pares et  al., 1998; Hord et al., 2001).



Diseases Caused by Viruses 387

A variety of methods are available for diagnosing CMV infection in banana germplasm (Thomas, 2015). Diagnosis of CMV infection by symptomatology is the least reliable method of disease identification because of the periodicity of symptom appearance, the effect of temperature on symptom expression and the similarity of symptoms induced by other viruses. Disease diagnosis based on symptomatology can be supported by biological assays using herbaceous indicator plants, such as tobacco, squash and cowpea (Francki et al., 1979). Although indicator plant assays have been largely supplanted by other diagnostic techniques, they may still be useful in situations where other methods are unavailable. One value of this method is that it is able to distinguish the presence of CMV, which is transmissible by mechanical inoculation, from banana bunchy top virus (BBTV) and banana streak viruses (BSVs), which are not transmitted by mechanical inoculation and do not infect herbaceous indicator plants. Highly reliable methods have been developed for detection of CMV by enzyme immunoassays. Polyclonal antibodies capable of detecting a wide range of virus strains (Thomas, 2015) and monoclonal antibodies that can be used to differentiate between virus subgroups (Hasse et  al., 1989; Wu et  al., 1997) are available. Genome (nucleic acid)-based methods of virus identification are highly sensitive and are being used increasingly for diagnosis of CMV infection in Musa. Methods based on nucleic acid hybridization (Gonda and Symons, 1978) have been largely replaced by (immunocapture, multiplex) RT-PCR amplification (Wylie et  al., 1993; Hu et  al., 1995; Singh et  al., 1995; Sharman et  al., 2000b). Real-time PCR (Feng et al., 2006), LAMP (Peng et  al., 2012b) and microarray assays (Deyong et al., 2005) have also been developed.

A. glycines, Acyrthosiphon pisum, Myzus persicae, Rhopalsiphum maidis, R. prunifoliae and Therioaphis trifolii (Jacquemond, 2012). As CMV is also seed transmitted in many of its hosts (Jacquemond, 2012), there is often a reservoir of inoculum present in the rural environment. Common and severe strains of CMV do not differ in their epidemiology. Aphid vectors usually acquire CMV from weeds (e.g. Commelina (see symptoms on Plate 6.29) Stellaria, Bryonia and Solanum spp.) or crop hosts (tomato, melon (Plate 6.30), cucumber, sweet pepper) growing near or in banana fields. Viruliferous, winged aphids (alatae) migrate from these plants to infect banana. Most aphid species do not normally colonize banana, but can transmit CMV during exploratory feeding probes. Although some of these transient aphid species may be relatively inefficient vectors of CMV, their large numbers may ensure the effective transmission of the virus from alternative hosts to banana. Movement to banana is most likely by chance, following the disturbance or destruction of preferred hosts. Disease incidence is higher in new plantings, because land is usually cleared of alternative aphid hosts and weeds carrying CMV may later grow in great profusion in exposed soil (Niblett et  al., 1994). Tissue-cultured banana plants, which are low-lying and succulent, are believed to be particularly vulnerable to infection. The means of spread of CMV contrasts with that of BBTV, which is transmitted in a persistent manner by Pentalonia nigronervosa and the

Epidemiology and disease cycle Disease incidence in banana is determined primarily by the number of infected alternative hosts in the vicinity of the crop and the population dynamics of aphid vectors. CMV is capable of infecting over 1000 plant species and is transmitted in a non-persistent manner by over 60 species of aphid, including Aphis gossypii, A. fabae,

Plate 6.29.  Symptoms of cucumber mosaic virus on Commelina diffusa growing under banana on the island of St Vincent, Windward Islands (photo: D.R. Jones, SVBGA).

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Plate 6.30.  Banana intercropped with melon growing under plastic in Morocco. The aphid Aphis gossypii is known to be a vector for severe strains of cucumber mosaic virus from the melon to the banana plants in this situation (photo: B.E.L. Lockhart, UM).

recently recognized P. caladii, together the most common aphid species to colonize banana (Blackman and Eastop, 1984; Foottit et  al., 2010; Duay et  al., 2014). Although Pentalonia has been reported to be unimportant in the spread of CMV (Stover, 1972; Wardlaw, 1972),

it could be important in new plantations established from tissue culture-­derived plants. On juvenile plants, large populations of P. nigronervosa develop on leaves and petioles (Plate 6.31), where aerial dissemination is more likely than from the pseudostem beneath basal



Diseases Caused by Viruses 389

CMV from alternative host to banana as well as from banana to banana. A better understanding of the epidemiology of CMV in banana requires a better understanding of the biology and ecology of the aphid vector species that colonize and/or visit banana and many other alternative hosts of the virus. In theory, a banana plant with banana mosaic should be systemically infected with CMV. Indeed, the leaves of many suckers arising from diseased plants have mosaic symptoms. However, there are numerous instances when the leaves of suckers are symptomless. Farmers report that these suckers, when propagated, develop into plants that exhibit no mosaic symptoms and produce a normal bunch. CMV-infected ‘Williams’ (AAA, Cavendish subgroup) kept under screenhouse conditions in Brisbane have been observed to give rise to symptomless suckers in which the virus was not detected by RT-PCR (K.S. Crew and J.E. Thomas, Australia, 2015, unpublished results).

Plate 6.31.  Colony of black banana aphids (Pentalonia nigronervosa) on a young sucker of ‘Dominico-Hartón’ (AAB, Plantain subgroup) in Colombia (photo: B.E.L. Lockhart, UM).

leaf sheaths as on more mature plants. This hypothesis provides an alternative explanation for the higher incidence of mosaic in banana plantations established from tissue-cultured plants. A second reason for not discounting the possible role for Pentalonia spp. in the epidemiology of CMV is the fact that these aphids can colonize species in both the Musaceae (predominantly P. nigronervosa) and Zingiberales (predominantly P. caladii) (Foottit et al., 2010). In West Africa, where the incidence of CMV is high (Osei, 1995), large populations of Pentalonia have been observed colonizing aerial shoots of Commelina diffusa (B.E.L. Lockhart, USA, 1998, unpublished results), a major alternative host of CMV. Interestingly, Commelina is in the order Commelinales, a sister order of the Zingiberales in the commelinids clade of monocots, and could reasonably be speculated to be a natural host of P. caladii. These observations suggest that in some situations Pentalonia may be an important vector of

Host reaction CMV has one of the widest host ranges of any plant virus, having been recorded as infecting at least 1000 species in at least 85 monocot and dicot plant families (Roossinck, 2001). There are many records of infection of banana (e.g. Singh et  al., 1995; Chou et  al., 2009; Kouassi et  al., 2010) and abacá (M. textilis) is also recorded as a host (Furuya et al., 2006). Many different banana cultivars have been reported with mosaic symptoms, but in some cases the disease may have been confused with other virus diseases. The cucumber mosaic diseases of plantain recorded by Stover (1972) in Honduras and of ‘Poovan’ (AAB, Mysore subgroup) described by Mohan and Lakshmanan (1988) in Tamil Nadu, India, were both most likely banana streak disease. This problem of diagnosis based on symptoms makes it difficult to compile lists of known host cultivars and their reaction to CMV. It is not known if resistance to CMV occurs within Musa. Stover (1972) indicated that M. balbisiana was the only species of Musa that appeared free of virus symptoms in the field, though seedlings had been observed with a whitish chlorotic mottle, which disappeared as

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the plants matured. Incidence of mosaic has been reported to be greater in cultivars in the Cavendish subgroup (AAA) than in ‘Gros Michel’ (AAA) in Central America. Symptoms on Cavendish cultivars have been recorded as being more severe than on diploids (Stover, 1972).

Control It is important to ensure that corms and suckers used for planting are healthy and not derived from plants with mosaic symptoms. Tissue cultures used for the in vitro multiplication of propagating material should be initiated from shoot-tips excised from plants that have been tested for the presence of CMV. Although CMV has been eliminated from diseased material by heat treatment of corms followed by apical meristem culture (Berg and Bustamante, 1974), it is recommended that this technique be attempted only for exceptionally valuable germplasm, such as an important breeding line or hybrid. Otherwise, it would be prudent to discard the material and use an uninfected source of the same clone for propagation. CMV is difficult to remove from in vitro banana material by meristem tip culture alone, as the virus is present in the meristematic dome of highly proliferating banana meristematic cultures (Helliot et al., 2007). There is some evidence to indicate that CMV may be seed transmitted in banana (Gold, 1972) and so seedlings need to be tested if from a plant whose virus disease status is unknown. Symptoms of streak and bract mosaic in banana are similar to those of mosaic (Lockhart, 1986; Magnaye and Espino, 1990). Mosaic symptoms in the bracts are usually diagnostic for bract mosaic. Also, few if any leaf symptoms are associated with bract mosaic disease. Mosaic and streak are harder to differentiate, because both can cause chlorotic and necrotic leaf symptoms, leaf distortion and internal necrosis of the pseudostem. However, a trained eye can usually distinguish between leaf symptoms of these two diseases, as streak symptoms tend to be more linear and conspicuous (Lockhart, 1986). Symptoms of mosaic can also be misidentified as zinc deficiency (see Chapter 9). Eliminating weed hosts of CMV from plantations and surrounding areas and ensuring

that susceptible crop hosts are not present within or near banana fields are other important elements in the control of mosaic. Weeds in and alongside plantings should be controlled efficiently, especially when the crop is young and more vulnerable to infection. Banana should not be intercropped with crop hosts of CMV. In the earliest studies on the disease in New South Wales in Australia, it was reported that outbreaks occurred in plantations near to where cucurbits, tomatoes or other vegetables were growing (Magee, 1940b). In Taiwan, incidence of mosaic was much less when banana was grown next to rice than when grown next to vegetables. The disease was highest in a field intercropped with cucumber (Tsai et  al., 1986). Banana plantlets being acclimatized in nurseries after removal from tissue culture should also be protected from sources of infection. Banana plants with mosaic should be removed from commercial plantations and spaces should be replanted with healthy material. This is because diseased plants often have a poor yield and fruit can have virus symptoms, which makes it unattractive to discerning consumers. Their removal also eliminates the chance of disease spread from banana to banana. On occasions when disease incidence is exceptionally high, fields may have to be completely replanted. Insecticides applied to banana plants are unlikely to be effective in controlling mosaic, because the major aphid vectors recognized do not colonize banana and virus transmission occurs after only a brief probe. Nevertheless, insecticides have been used in commercial plantations in the past and, together with the removal of diseased and surrounding plants, very good control has been achieved (Adam, 1962). However, the cost effectiveness of these operations is questionable (Jeger et al., 1995) and the roguing of diseased plants alone has resulted in adequate control (Stover, 1972). Because it occurs ubiquitously, CMV is not regarded as a quarantinable pathogen. This view does not take into account that severe or heart-rot isolates of CMV, which are far more damaging than common isolates of the virus, do not occur in all banana-producing areas. It is therefore important to avoid the introduction of severe CMV isolates into new areas where they can cause significant damage (Bouhida and Lockhart, 1990).



Diseases Caused by Viruses 391

Abacá Mosaic J.E. Thomas and L.V. Magnaye Introduction The Philippines supplies around 85% of the total world production of abacá fibre. It ranks ninth in importance as an export crop. Abacá mosaic, together with abacá bunchy top, are serious disease constraints to production (Raymundo, 1998; Lalusin and Villavicencio, 2015). The first record of abacá mosaic was on the island of Mindanao in the Philippines in 1925 (Eloja and Tinsley, 1963). Calinisan (1934) first described the disease near Davao and a few years later it was identified in Cotabato province (Celino, 1940). By the late 1930s, abacá mosaic was the major factor limiting abacá production near Davao, being responsible for heavy losses of fibre and much capital investment. This caused the relocation of the industry from the Mount Apo area to more remote regions (Stover, 1972). By 1955, abacá mosaic affected 47% of the 184,000 ha under abacá cultivation and losses of 25–50% in new plantings were common. Abacá mosaic was later reported from Luzon and certain areas in the Visayas. Today, it is found in most areas where abacá is grown in the Philippines and can occur at high levels of infection and causing significant economic losses (Raymundo et al., 2001, 2002; Sta. Cruz et al., 2016). Early infection renders plants unproductive and worthless. Infection can significantly reduce tensile strength, biomass yield, fibre yield, plant height and stalk diameter (de la Cruz and Raymundo, 2009). However, some fibre production is possible when older plants (1–2 years old) become infected and the irregular distribution of the virus in individual abacá clumps can allow some stalks to be temporarily disease free (Magee, 1957; Stover, 1972).

chlorotic streaks, appear parallel to the minor leaf veins. As these chlorotic areas expand, they may develop rusty-brown borders and can extend from the midrib to the leaf margin. Subsequent leaves can develop extensive broad ­yellow or pale green stripes across the width of lamina (Plate 6.32). With older infections, chlorotic, orange or dark-green streaks may be present on the midribs and petioles, even when lamina symptoms are absent (Ocfemia and ­Celino, 1938; Eloja et  al., 1962; Benigno and Rosaro, 1965; Stover, 1972). Leaves can also become distorted (Plate 6.33). In some earlier reports, it is uncertain whether symptom descriptions pertain to abacá mosaic, as some illustrations are reminiscent of infections caused by cucumber mosaic virus (Calinisan, 1939). Diagnosis of the disease based on symptoms alone must be viewed with caution as similar symptoms can be produced in abacá in the Philippines by both the abacá mosaic pathogen of Eloja and Tinsley (1963) and banana bract mosaic virus (BBrMV) (Thomas et  al., 1997;

Symptoms On inoculated abacá, symptoms become apparent after 7–21 days (Celino and Ocfemia, 1941; Eloja et  al., 1962). Small whitish dots, which later elongate to become spindle-shaped

Plate 6.32.  Chlorotic discoloration symptoms of abacá mosaic disease on an abacá leaf in the Philippines (photo: L.V. Magnaye, BPI).

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Plate 6.34.  Virions of sugarcane mosaic virus abacá strain. Scale bar represents 200 nm (photo: J.E. Thomas, QDAF).

Plate 6.33.  Distortion symptoms of abacá mosaic disease on an abacá leaf in the Philippines (L.V. Magnaye, BPI).

Sharman et  al., 2000a). Abacá plants are also frequently subject to a range of mixed virus infections (Sta. Cruz et al., 2016).

Causal agent Flexuous filamentous virions about 680 nm long were detected in abacá mosaic-affected plants (Plate 6.34) (Eloja and Tinsley, 1963) and considered to be those of the causal agent. On the basis of vector transmission characteristics and host range, Magee (1957) suggested that the causal agent of abacá mosaic was related to sugarcane mosaic virus (SCMV). This was subsequently supported by micro precipitin tests showing a serological relationship with SCMV (Eloja and Tinsley, 1963). By ELISA, relationships were demonstrated with members of the sugarcane mosaic subgroup of potyviruses (including SCMV, maize dwarf mosaic, sorghum mosaic and Johnson grass mosaic viruses) and

also a weak relationship with BBrMV (Thomas et al., 1997; Gambley et al., 2004). Sequence analysis of the coat protein gene and 3′ untranslated region has confirmed that this virus causing abacá mosaic is a distinct strain of Sugarcane mosaic virus (SCMV-Ab) (Gambley et  al., 2004). Serological and nucleic acid-based assays are available for SCMV-Ab. The virus can be detected by plate-trapped antigen ELISA using antisera to SCMV or MDMV (Gambley et al., 2004). SCMV-Ab can also be detected by PCR using degenerate potyvirus genus primers (Thomas et al., 1997) or specific primers (Thomas, 2015). Sharman et al. (2000a) demonstrated that BBrMV, widespread in banana in the Philippines, could also be found in naturally infected abacá, and cause abacá mosaic symptoms. This section will focus on SCMV-Ab only.

Disease cycle and epidemiology SCMV-Ab is transmitted in the non-persistent manner by a number of aphid species, including Aphis gossypii, Rhopalosiphum nymphae (Celino, 1940), A. glycines (Wu et  al., 2004), R. maidis (Celino and Ocfemia, 1941), Schizaphis cyperi (syn. A. rotundiventris), S. graminum (Gavarra and Eloja 1969) and Toxoptera citricidus (Eloja et  al., 1977). Aphis gossypii and R. maidis are the principal natural vectors. The ability to transmit is rapidly lost during a series of transfers of the aphids to new hosts, consistent with



Diseases Caused by Viruses 393

a non-­persistent mode of transmission (Celino, 1940). It appears that the important natural vectors of SCMV-Ab do not colonize abacá, and indeed the banana aphid Pentalonia nigronervosa appears not to be a vector of SCMV-Ab (­Celino, 1940; Ocfemia et al., 1947). There is no transmission through true seed of abacá (Calinisan, 1939), but the virus is transmitted through vegetative planting material and tissue culture. Mechanical transmission of SCMV-Ab is possible, though infected abacá is a poor source of inoculum (Eloja et al., 1962). However, once the virus is transmitted to alternative hosts in the Poaceae, either by mechanical inoculation or by aphid transmission, it is readily sap transmissible, even back to abacá (Eloja et al., 1962; Benigno and Del Rosario, 1965).

1956) and a number of banana cultivars, which include ‘Lakatan’ (AAA), ‘Umalag’ (AAA, Cavendish subgroup), ‘Grand Nain’ (AAA, Cavendish subgroup) and ‘Latundan’ (AAB, Silk subgroup) (L.V. Magnaye, Davao, 1998, unpublished results), are susceptible to SCMV-Ab. Resistance has been noted in Musa balbisiana accession ‘Pacol’, the M. balbisiana × M. textilis hybrid ‘Canton’ and Musa ornata (Bernado and Umali, 1956).

Control

The aphid vectors of SCMV-Ab do not colonize abacá. It has been recommended that maize, a host favoured by the aphid vector R. maidis, and other grasses should be excluded from plantations (Stover, 1972). However, studies by Celebrar et al. (1970) showed that total weed removal resulted in twice the incidence of SCMV-Ab as compared with cover cropping or removal of weeds from Host reaction only around the bases of the abacá plants. Insecticide spraying did not result in significant reducHosts are restricted to monocotyledonous plants. tion in disease incidence. Natural hosts include Musa textilis (abacá), A major source of infection is infected Canna indica (Cannaceae) and Maranta arundinaplanting material. When infected corms are cea (Marantaceae), while the following memplanted, symptoms usually start to appear withbers of the Poaceae are experimental hosts: Digin 2 months, but can take 12–18 months (Eloja itaria sanguinalis, Echinochloa colonum, Imperata et al., 1962; Stover, 1972). The virus appears to cylindrica, Rottboellia exaltata, Setaria palmifolia be unevenly distributed in the corm, and it is and Zea mays (Velasco-Magnaye and Eloja, possible to obtain virus-free suckers from an in1966). fected corm if the plants are severed 3–5 weeks The members of the Poaceae listed above, after emergence (Pacumbaba, 1967). Roguing and also Andropogon halepensis, Coix lachryma-­ of infected plants and the use of virus-free plantJobi, Panicum distachyum and Paspalum conjugaing material are considered essential to contain tum, have been observed in the field to have mothe spread of abacá mosaic (Magee, 1957, 1960; saic symptoms and contain viruses transmissible Eloja et al., 1962). to abacá with A. gossypii or R. maidis. However, the symptoms produced in abacá consist of only a few spindle-shaped chlorotic lesions on the lamina and are much less severe than those Banana Streak produced by typical isolates from abacá (Celino and Martinez, 1956; Velasco-­Magnaye and EloM.L. Iskra-Caruana, J.E. Thomas ja, 1966). It is not known whether these grasses and M. Chabannes contain mild strains of SCMV-Ab or other distinct viruses. No cross-­protection was evident Introduction when abacá plants infected with the viruses from R. exaltata or P. distachyum were super-­ infected with typical SCMV-Ab (Celino and Banana streak is one of the most widespread Martinez, 1956). virus diseases of banana. It is caused by a group In the Musaceae, Ensete glaucum, all com- of related viruses, and its epidemiology and mercial varieties of abacá (Bernado and Umali, detection are complicated by the presence of

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mealybug-transmitted episomal infections and of viral sequences integrated into the Musa genome in both ‘activatable’ and ‘dead’ forms (Lava Kumar et al., 2015). Discovery of banana streak Chlorotic leaf streak symptoms first identified as a cucumber mosaic virus (CMV)-mediated chlorosis disease were observed in banana plantations in Côte d’Ivoire’s Nieky Valley in 1958 (Lassoudière, 1974). In 1964, preliminary research was initiated after the occurrence of serious outbreaks in the region. Two years later, Yot-Dauthy and Bové (1966) named this disease ‘banana streak’ by demonstrating that the symptoms were due to a virus distinct from CMV. Lassoudière (1974) was the first to evaluate the impact of banana streak by performing a symptomatology survey using an agronomic trial on banana ‘Poyo’ (AAA, Cavendish subgroup). He reported that symptoms could affect one leaf but not another on the same plant, and that affected plants could lose and regain symptoms between cycles of the crop. He pointed out the importance of using clean planting material to control the disease and the difficulties in identifying it because of the periodic presence of the symptoms. Finally, he recorded yield losses ranging up to 90% and the production of few (if any) exportable bunches. Similar symptoms, turning into necrosis, and severe outbreaks were next reported from southern Morocco (Lockhart, 1986). They had been observed since 1974 in almost every newly established planting of field-grown ‘Dwarf Cavendish’ (AAA, Cavendish subgroup). In some cases, incidence of banana streak exceeded 50%. Lockhart (1986) confirmed the viral aetiology of the disease by purifying and characterizing bacilliform virus particles from these plants and subsequently reported a high serological diversity among viral isolates from around the world (Lockhart and Olszewski, 1993). Distribution and losses After recognition of the disease, numerous banana streak outbreaks were subsequently recorded in Africa (e.g. Nigeria, Cameroon, Zanzibar, Tanzania, Burundi, Rwanda, Democratic Republic of the Congo), in Central and South

America (Honduras, Colombia, Ecuador, Brazil) and in Australia, Tonga and Western Samoa, affecting diverse banana genotypes and improved interspecific hybrids (Stover, 1988; Thomas et al., 1994b; Gauhl et  al., 1999a, b; Geering et al., 2000; Lockhart and Jones, 1999; Daniells et al., 2001; Thomas, 2015). As banana can be infected and show no symptoms, estimates of incidence of the disease based on symptomatology vary greatly. Only a few quantitative studies exist. In some localities in certain countries the disease appears to be ­serious and in other places it is confined to a few plants and is not regarded as important. In southern Cameroon and southern Nigeria, plantains with banana streak have been found in just over half the villages surveyed, with incidences ranging from 0.5% to 17.0% (Gauhl et  al., 1999a, b). Incidence has also been common and widespread in plantain (AAB) around the Sula Valley in Honduras and the Los Rios province in Ecuador (Lockhart and Jones, 1999). The genotype ‘Mysore’ (AAB), a widely distributed dessert clone from India, is known to be almost totally affected by banana streak (Jones, 1994; Lockhart, 1994a; Thomas et al., 1994b; Daniells et al., 1995). The leaf-streak symptoms so common on ‘Mysore’ were first thought to be the consequence of physiological effects (Wardlaw, 1972), but it was later shown that they were due to virus infection (Lockhart and Olszewski, 1993). In Tamil Nadu State in India, ‘Mysore’ is intensely cultivated and can only be grown for three cycles before replanting, because of the rapid decline in productivity of plantations (Lockhart and Jones, 1999). Epidemics of banana streak have only been reported from Uganda (Karamura et al., 1996; Harper et  al., 2002, 2004, 2005). Despite many outbreak records of banana streak disease worldwide over the past 20 years, convincing evidence of other epidemics is lacking (Lava ­Kumar et al., 2015).

Symptoms Symptom expression varies from inconspicuous chlorotic dots to lethal necrosis, depending on virus isolates, banana cultivars and environmental conditions (Gauhl and Pasberg-Gauhl,



Diseases Caused by Viruses 395

1994; Lockhart, 1994a; Dahal et  al., 1998a; Daniells et  al., 1999). However, the most common symptoms on leaves are discontinuous chlorotic or yellow streaks, which run from the leaf midrib to the margin (Plate 6.35). Streaks later darken, often becoming brown or black (Plates 6.36 and 6.37). Necrosis has also been seen on the leaf midrib and petiole. Yellow to orange blotches on leaves can also be associated with banana streak (Plate 6.38) and splitting of the pseudostem is also observed (Plate 6.39). Foliar symptoms appear sporadically over the course of the year and are erratically distributed on the plant. Indeed, symptomatic leaves may be succeeded by leaves expressing few or no symptoms in the same plant although the virus can be detected at all stages in any leaves. When the leaves with symptoms are shed, the plant

Plate 6.35.  Chlorotic stripe and eye-shaped symptoms of banana streak disease on a leaf of ‘Mysore’ (AAB) in Honduras (photo: D.R. Jones, INIBAP).

Plate 6.36.  Streaks on a leaf of a newly bred AAB hybrid infected with both banana streak GF virus and banana streak OL virus turning from yellow to brown (photo: M.-L. Iskra-Caruana, CIRAD).

may appear healthy and it may be many months before symptoms reappear on new leaves. This periodicity of symptoms and their severity is always associated with environmental or physiological changes and is closely correlated with abiotic stresses such as fluctuations in temperature (Lockhart, 1995; Dahal et al., 1998a; Daniells et al., 2001) or water supply (successive dry and wet long seasons). Various other symptoms associated with banana streak include leaf bases falling away from the pseudostem, streaks on the pseudostem (Plate 6.40), narrow, thicker leaves and constriction of the bunch on emergence (‘choking’) (Lassoudière, 1979). Stunting due to a shortening of the internodes, cigar-leaf necrosis (Plate 6.41), internal necrosis of the pseudostem (Plates 6.42), aberrant bunch emergence (Plate 6.43), reduced bunch size and distortion of fingers can also occur (Gauhl and Pasberg-Gauhl, 1994). Peel splitting (Plate 6.44) and necrotic spot symptoms in the peel (Plate 6.45) have been observed on fruit of ‘Grand Nain’ (AAA, Cavendish subgroup) in Costa Rica and Ecuador. In Australia, mosaic symptoms have been noted on the peel of bred hybrids (Plate 6.46). Also, broad yellow lines in the leaf lamina parallel to the midrib, a purple margin to the leaf lamina, leaf twisting, grooves in the base of the pseudostem and an abnormal arrangement of the leaves similar to the traveller’s palm (Ravenala madagascariensis) have been associated with banana streak in ‘Williams’ (AAA, Cavendish subgroup) (Daniells et al., 1998).

Plate 6.37.  Black stripe symptoms of banana streak disease on a leaf of ‘Mysore’ (AAB) growing in the Torres Strait region of Australia (photo: D.R. Jones, QDPI).

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ENYERU (IBANDA)

Plate 6.38.  Yellow-orange blotchy symptoms of banana streak disease on a leaf of ‘Enyeru’ (AAA, Lujigira-Mutika subgroup) in Uganda (photo: B.E.L. Lockhart, UM).

texts (Yot-Dauthy and Bové, 1966; Stover, 1972; Wardlaw, 1972) are now clearly recognizable as those of banana streak. Cucumber mosaic of plantains, described by Stover (1972), and cucumber mosaic of ‘Poovan’ (AAB, Mysore subgroup), described by Mohan and Lakshmanan (1988), are also most likely banana streak.

Causal agent Particle and genome properties Plate 6.39.  A splitting pseudostem of a banana with banana streak disease (photo: M.-L. Iskra-Caruana, CIRAD).

Three phases of symptom expression have been recognized in commercial Cavendish plantations in Ecuador. The first is the appearance of chlorotic streaks in leaves, the second is the appearance of dark blotches on the pseudostem and midrib and the third is the splitting of the outer leaf sheaths of the pseudostem, leading sometimes to the death of the plant. Plants with severe symptoms produced few (if any) marketable bunches. The impact of banana streak may have been underestimated in the past due to confusion with banana mosaic caused by CMV. For instance, some illustrations of banana mosaic in previous

Banana streak viruses (BSVs) cause banana streak disease. They are a related group of non-­ enveloped plant pararetroviruses belonging to the family Caulimoviridae and genus Badnavirus (King et  al., 2012). They have bacilliform particles of 130–150 nm × 30 nm in size (Plate 6.47) (Lockhart and Olszewski, 1993), containing a non-­covalently closed circular dsDNA genome approximately 7.2–7.8 kb long that uses a virus encoded reverse transcriptase (RT) to replicate. Noteworthy and conversely to retroviruses in the animal kingdom, plant pararetroviruses do not require an integration step into the host genome to replicate. Although no extensive studies of the BSV replication cycle have been undertaken, it is likely to parallel that of the species Cauliflower mosaic virus (CaMV) from the genus Caulimovirus and Rice tungro bacilliform virus (RTBV) from the genus



Diseases Caused by Viruses 397

Plate 6.40.  Black stripe symptoms of banana streak disease on the pseudostem of ‘Mysore’ (AAB) growing in Honduras (photo: D.R. Jones, INIBAP)

Tungrovirus (Hohn, 2013; Hohn and Rothnie, 2013). Briefly, the viral life cycle begins with the release of viral double-stranded DNA (dsDNA) from virions into the nucleus. The gaps on both strands remaining after reverse transcription are repaired and the resulting covalently closed dsDNA associates with histones to form a minichromosome (episome). Pol II-mediated ­ ­transcription of the circular minichromosome ­generates a capped and polyadenylated pgRNA with terminal repeats, which is transported to the cytoplasm for translation of viral proteins and subsequently for reverse transcription. Translation for CaMV of pgRNA is initiated by a shunt mechanism in which ribosomes bypass a long leader sequence containing multiple short ORFs (sORFs) and folding into a stable stem-loop structure (Fütterer et  al., 1993; Pooggin et  al., 2006, 2008). Both features are conserved in plant pararetroviruses (Pooggin et  al., 1999). Several consecutive viral ORFs on polycistronic

Plate 6.41.  Cigar-leaf necrosis symptom of banana streak disease on IITA plantain hybrid ‘TMPx 548-9’ in Nigeria. Brown streak symptoms can also be seen on the petiole to the left of the dead cigar leaf (photo: C. Pasberg-Gauhl and F. Gauhl, IITA)

pgRNA are then translated by reinitiation (CaMV) (Ryabova et al., 2002) or leaky scanning (RTBV) (Fütterer et al., 1997) mechanisms. The reverse transcription takes place in cytoplasmic inclusion bodies and presumably is initiated by coat protein-mediated packaging of pgRNA (Guerra-­ Peraza et al., 2000). The resulting open circular dsDNA can re-enter the nucleus for the next round of transcription or be encapsidated into mature virions for movement within the host plant and for transmission to a new host plant by insect vectors. Harper and Hull (1998) were the first to describe the genome structure of BSV from the banana streak OL virus (BSOLV) isolated from ‘Obino l’Ewai’ banana plants in Nigeria. The genome contains three consecutive ORFs on one strand (King et  al., 2012). ORF1 and ORF2 (virion-­ associated protein (VAP)) encode two small proteins of unknown function of 20.8 kDa

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Plate 6.42.  Internal necrosis of the pseudostem of IITA plantain hybrid ‘TMPx 597-4’ growing in Nigeria caused by banana streak disease (photo: C. Pasberg-Gauhl and F. Gauhl, CB).

and 14.5 kDa, respectively. ORF3 is a large polyprotein of 220 kDa encoding at least four proteins encompassing a putative cell-to-cell movement protein, a coat protein (CP – analogous to retroviral gag), an aspartic protease and a viral replicase (analogous to retroviral pol) consisting of RT and RNase H domains (Hull, 2002). This polyprotein is likely to be cleaved into functional units by the aspartic protease once it has been fully translated. A three-dimensional structural model of the banana streak MY virus (BSMYV) CP has been determined, showing it to be a multi-domain structure possessing homologues of the retroviral capsid and nucleocapsid proteins. Additionally, it contains a surface-exposed, intrinsically disordered protein region at the N terminus (NID), to which several linear epitopes have been mapped (Vo et  al., 2016). Two different CP isoforms (ca 44 kDA and 40 kDa) and the VAP are associated with purified virions. The smaller CP

Plate 6.43.  Emergence of the fruit bunch through the side of the pseudostem of a banana streak disease-affected ‘Grand Nain’ (AAA, Cavendish subgroup) in Costa Rica (photo: S. Daillot, CIRAD).

is thought to be a cleavage product of the larger, and to differ in length of the NID domain (Vo et al., 2016). BSV diversity A high degree of heterogeneity exists among BSV isolates which strongly differ serologically, genomically and in symptomatology (Lockhart and Olszewski, 1993; Geering et al., 2000). Five serologically distinct isolates of BSV from Morocco, Rwanda, Trinidad, Honduras and the Philippines were identified by Lockhart and Olszewski (1993). Geering et al. (2000) underlined nucleic acid differences ranging from 21.8% to 33.6% in the sequence of the RT and RNase H regions when comparing four Australian isolates with one from Nigeria. Later, a large molecular diversity was described for BSV from field-grown plants of the East African highland banana group (AAA, Lujugira-Mutika subgroup) in



Diseases Caused by Viruses 399

Plate 6.44.  Peel splitting symptom of banana streak disease of fingers of ‘Grand Nain’ (AAA, Cavendish subgroup) in Costa Rica (photo: C. Pasberg-Gauhl and F. Gauhl, CB).

Uganda (Harper et  al., 2002, 2004, 2005). These sequences represented a number of distinct species of BSV, according to the International Committee on Taxonomy of Viruses (ICTV) species demarcation criteria (Fauquet et  al., 2005). Phylogenetically, the species were distributed across the three main clades formed by the genus Badnavirus (Fig. 6.1). Iskra-Caruana et al. (2014a) showed that all known BSV species (episomal forms) are restricted to Clades 1 and 3. Clade 1 contains BSV species coming from around the world, whereas Clade 3 contains BSV species exclusively from Uganda (Fig. 6.1). Clade 2 contains endogenous, apparently ‘dead’ BSV sequences (Iskra-Caruana et al., 2014a, b; M. Chabannes, France, 2017, unpublished results). To date, nine BSV species have been formally recognized by the ICTV. These are Banana streak GF virus (BSGFV), Banana streak IM virus (BSIMV), BSMYV, BSOLV, Banana streak UA virus (BSUAV), Banana streak UI virus (BSUIV), Banana

Plate 6.45.  Necrotic spot symptoms of banana streak disease in the peel of a finger of ‘Grand Nain’ (AAA, Cavendish subgroup) in Costa Rica (photo: C. Pasberg-Gauhl and F. Gauhl, CB).

Plate 6.46.  Mosaic symptoms of banana streak disease in a finger of IRFA 909 (AAB). The bred hybrid was in a field trial in northern New South Wales, Australia (photo: J.E. Thomas, QDAF).

streak UL virus (BSULV), Banana streak UM virus (BSUMV) and Banana streak VN virus (BSVNV). Sugarcane bacilliform viruses (SCBVs) are genetically variable and genetically related to BSV (Muller et al., 2011). They occur widely in sugarcane and can infect banana and several

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Plate 6.47.  Bacilliform particles of banana streak virus stained with sodium phosphotungstate (photo: B.E.L. Lockhart, UM).

other monocotyledonous hosts (Bouhida et  al., 1993). Banana streak-like symptoms develop in banana after inoculation with SCBV, but BSV does not infect sugarcane (Lockhart, 1995). Endogenous BSV sequences (eBSV) As previously mentioned, BSVs do not require integration into the banana host genome to replicate and do not code for any integrase. However endogenous BSV sequences (eBSV) have been demonstrated for BSOLV, BSGFV, BSIMV and BSMYV in the B genome. They can produce de novo episomal particles causing the same banana streak infections as their episomal counterparts and can cause disease outbreaks (Iskra-Caruana et al., 2010). This was conclusively demonstrated for three BSV species (BSOLV, BSGFV, BSIMV) (Chabannes et  al., 2013; Gayral et  al., 2008; Lheureux et al., 2003) and is strongly suspected for BSMYV (Geering et  al., 2005). These eBSVs contain at least the full-length functional viral genome and are an integral part of the banana genome (Fig. 6.2). BSV integrations were first implied by LaFleur et al. (1996). Ndowora et al. (1999) described the 5′ part of the BSOLV integrant present in the genome of the plantain ‘Obino l’Ewai’. They suggested, based on in silico analysis, that the full-length viral genome of BSOLV could be released following two homologous recombination events. Later, Lheureux et al. (2003) established that the banana streak disease occurring in newly created interspecific AAB hybrids from

virus-free parents, originated from endogenous sequences of BSOLV (eBSOLV) and the identification of a genetic factor named BEL (BSV expressed locus), both present in the Musa balbisiana genome. This work was pursued and extended to include BSGFV and BSIMV (Chabannes et al., 2013; Gayral et al., 2008. Over the past 10 years, extensive genomic, genetic and cytogenetic studies have revealed the sequence, organization and localization of eBSVs from BSOLV, BSGFV and BSIMV in the genome of the seeded diploid M. balbisiana (BBw, accession ‘Pisang Klutuk Wulung’) (Gayral et  al., 2008; Chabannes et  al., 2013). eBSVs present a complex structure composed of a succession of duplicated and/or inverted viral fragments without any Musa embedded sequences (Fig. 6.2). In the BSV species studied, eBSV length ranges from 13.2 kbp to 23.2 kbp. Each eBSV species is integrated at a distinct locus, eBSGFV and eBSOLV being on Chromosome 1 whereas eBSIMV is located on Chromosome 2. eBSGFV and eBSOLV are di-allelic with one infective and one non-infective allele, whereas eBSIMV is monoallelic with potentially both alleles being infective (Chabannes et al., 2013; Gayral et al., 2008). Infective alleles are probably able to reconstitute an infectious viral genome by homologous recombination (Iskra-Caruana et al., 2010; Chabannes and Iskra-Caruana, 2013). With a knowledge of eBSV structure, molecular markers were developed to distinguish eBSV (integrated form) from BSV (episomal virus) and to distinguish eBSV alleles (infective versus non-infective ­alleles) (Chabannes et  al., 2013; Gayral et  al., 2008) (Fig. 6.3). The markers used to genotype M. balbisiana germplasm diversity have so far not revealed a single BB genotype without infective eBSV (Gayral et  al., 2010; Duroy et  al., 2016). They always harbour at least one infectious allele (eBSOLV or eBSGFV). Finally, two recent conclusive studies suggest that current BSV populations of the three most common species (BSOLV, BSGFV and BSIMV) observed from outbreaks worldwide are mainly, if not exclusively, produced by their eBSV counterpart rather than vector-transmitted epidemics. Indeed, by studying the phylogenetic relationship between BSV and eBSV for each species among selected banana plants representative of the diversity of 60 wild Musa species and genotypes, Gayral et  al. (2010) estimated that



Diseases Caused by Viruses 401

95 97

SCBGAV (FJ824813) BSPEV (DQ674317) BSOLV (AJ002234)

84

SCBGCV (FJ439796)

100

91

Clade 1

BSCAV (HQ593111) BSUAV (HQ593107)

BSV and BSV/eBSV

KTSV (AY180137)

63

BSVNV (AY750155)

75

BSIMV (HQ659760)

70

58

BSGFV (AY493509) BSMYV (AY805074)

51

SCBGDV (FJ439817) ComYMV (X52938) BSUEV (AJ968467)

52

BSUFV (AJ968469)

72

BSUDV (AJ968465) BSUGV (AJ968470)

89

98

Clade 2

BSUHV (AJ968472) BSUCV (AJ968464) CSSV-Agou1 (L14546)

100

CSSV-NJ (AJ608931)

100

CSSV-Wobe12 (AJ781003)

72

78

CSSV-CI152 (JN606110)

Endogenous badnavirus sequences in Musa

CiYMV (AF347695) DBALV (X94576)

55

DBSNV (DQ822073) 97 52

BSUKV (AJ968504) BSULV (HQ593109) BSUMV (HQ593110)

66

BSUJV (AJ968501)

100

BSUIV (HQ593108) 100

SCBIMV (AJ277091)

Clade 3 BSV

SCBMOV (M89923) BCVBV (EU034539) TABV (AF357836)

0

1.

Fig. 6.1.  Maximum likelihood phylogeny of badnavirus sequences based on alignment of a 540-bp fragment of the RT/RNase H viral region (adapted from Kumar et al., 2015). Bootstrap values of 500 replicates are given when > 50%. Taro bacilliform virus (TABV) and bougainvillea spectabilis chlorotic vein-banding virus (BCVBV) are given as outgroups. Viral sequences isolated from Musa are in bold. BSV species where a full-length sequence is available are in italics. BSV species in red have only been found as episomal viruses up until now and those in green are both integrated in banana genomes and found as episomal viruses. The scale bar shows the number of substitutions per base. The GenBank accession numbers of sequences are given in parenthesis. Key: BCVBV = bougainvillea spectabilis chlorotic vein-banding virus; BSCAV = banana streak CA virus; BSGFV = banana streak GF virus; BSIMV = banana streak IM virus; BSMYV = banana streak MY virus; BSOLV = banana streak OL virus; BSUAV = banana streak UA virus; BSUCV = banana streak UC virus; BSUDV = banana streak UD virus; BSUEV = banana streak UE virus; BSUFV = banana streak UF virus; BSUGV = banana streak UG virus; BSUHV = banana streak UH virus; BSUIV = banana streak UI virus; BSUJV = banana streak UJ virus; BSUKV = banana streak UK virus; BSULV = banana streak UL virus; BSUMV = banana streak UM virus; BSVNV = banana streak VN virus; CSSV-Agou1 = cacao swollen shoot virus; ComYMV = commelina yellow mottle virus; CiYMV = citrus yellow mosaic virus; DBALV = dioscorea bacilliform AL virus; DBSNV = dioscorea bacilliform SN virus; KTSV = kalanchoe top-spotting virus; SCBMOV = sugarcane bacilliform MO virus; SCBIMV = Sugarcane bacilliform IM virus; SCBGAV= sugarcane bacilliform Guadeloupe A virus; SCBGCV = sugarcane bacilliform Guadeloupe C virus; SCBGDV = sugarcane bacilliform Guadeloupe D virus; TABV = taro bacilliform virus.

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BSV 1Kb eBSGFV-7 eBSGFV-9 Chromosome 1 eBSOLV-1 eBSOLV-2

eBSIMV Chromosome 2

Fig. 6.2.  Overview of eBSV structures in the Musa balbisiana (BBw, accession ‘Pisang Klutuk Wulung’) (adapted from Chabannes et al., 2013). Banana genomic sequences are in green. The BSV genome is represented in linear view with dark blue, light blue and red boxes indicating ORF 1, ORF2 and ORF 3 of the virus, respectively. The intergenic region (IG) is in black. Arrow boxes indicate the orientation of the fragment in the genome. The name of each eBSV allele is given above and red arrows indicate infective alleles. For eBSIMV, it is not yet ascertained if one or both copies are infective. The chromosome location of each eBSV is also indicated. eBSGFV-7 VM1

VV1

VV2 VV3 VV4 Dif GF

Dif GF

VV6 VM2

eBSGFV-9 VV1 VM1

Dif GF VV6 VV2 VV3 VV4 Dif GF VM2 VV5 Dif GF

eBSIMV F1–F3 Musa-F2

F3–F4 F5 - Musa F4–F5

eBSOLV-1 27–28 Dif OL

16–17

29–30

23–24

25–26

eBSOLV-2 16–17

29–30

27–28 Dif OL

20–21

25–26

Fig. 6.3.  Localization of PCR and dCAPs markers within eBSGFV, eBSOLV and eBSIMV of Musa balbisiana (BBw, accession ‘Pisang Klutuk Wulung’). Banana genomic sequences are in green. BSV genome is represented in linear view. Dark blue, light blue and red boxes indicating ORF1, ORF2 and ORF3 of the virus, respectively, for BSV and eBSV. The intergenic region (IG) is in black. Arrow boxes indicate the orientation of the fragment in the genome. PCR amplification fragments are represented by orange, purple and pink lines corresponding to Musa junction, internal and allelic amplifications fragments respectively. PCR markers were developed by Gayral et al. (2008, 2010) and Chabannes et al. (2013).



Diseases Caused by Viruses 403

eBSGFV and eBSIMV integrated into the banana genome relatively recently (ca 640,000 years ago). They also found very little variation between sequences from either integrated or episomal origin. This was confirmed and extended to eBSOLV by Chabannes et al. (2013), who found more than 99% identity at the nucleic acid level between episomal genomes and eBSV for BSOLV, BSGFV and BSIMV in Musa balbisiana (BBw, accession ‘Pisang Klutuk Wulung’), suggesting a major contribution of eBSVs to the current viral populations. Detection of BSV Serological detection of all BSV species remains complicated because of the wide serological diversity among BSV species (Lockhart and Olszewski, 1993). Commercialized polyclonal antisera have been mainly raised against the four BSV species (BSOLV, BSGFV, BSIMV and BSMYV) and various SCBV isolates (Ndowora and Lockhart, 2000; Agindotan et al., 2003) and these recognize viruses from Clade 1 and some species from Clade 3 (M. Chabannes and M.-L. Caruana, France, 2016, unpublished results). They are polyvalent (i.e. they can detect a number of BSV species, though BSMYV is serologically most distinct) and ELISA test kits are available from AGDIA and DSMZ companies. Molecular diagnostics remain a good alternative to overcome such wide serological diversity. Unfortunately direct PCR using total DNA extracts from plants can lead to false positives, because it does not distinguish true BSV infection from PCR amplification of eBSV in the B genome. To overcome this problem, IC-PCR based on immunocapture of viral particles associated with a DNase step and specific PCR amplification (Le Provost et al., 2006; Thomas, 2015) was developed. Alternative techniques are also reported (Thomas, 2015) but IC-PCR remains the most reliable technique to diagnose episomal BSV.

Host reaction Like most badnaviruses, BSVs have a restricted natural host range. Banana streak disease naturally affects cultivated landraces of banana, different species of Musa and natural and synthetic

hybrids found in germplasm collections. No data exists so far to determine if BSV occurs naturally in Musa and Ensete species growing in the wild. Almost all genotypes, cultivars and subgroups of banana appear susceptible to BSVs, but often the particular virus species present has not been identified (Lassoudière, 1974; Sebasigari and Stover, 1988; Dabek and Waller, 1990; Daniells et al., 1999; Dahal et al., 2000; Harper et al., 2002; James et al., 2011). Of note, all interspecific hybrids with one B genome are at risk due to the presence of eBSV. For instance, BSVs have been found in most of the newly created interspecific hybrids emerging from breeding programmes in Guadeloupe (Centre de cooperation international en recherche agronomique pour le développement (CIRAD)), Honduras (Fundación Hondureña de Investigación Agrícola (FHIA)), Nigeria (International Institute of Tropical Agriculture (IITA)) and Brazil (Empresa Brasileira de Pesquisa Agropecuária (EMBRAPA)). Very few data are available on Musa resistance to BSV infection. The major defence mechanism implicated in plant recovery from viral infection is based on RNA silencing. This mechanism generates viral short interfering RNAs (siRNAs) that can repress viral genes both post-­ transcriptionally through RNA cleavage (PTGS) (Mourrain et  al., 2000) and transcriptionally through DNA cytosine methylation (TGS) (Al-Kaff et al., 1998). The resistance mechanisms triggered by Musa to overcome virus attacks have rarely been studied at cellular level except for banana bunchy top virus (Shekhawat et  al., 2012). Regarding BSVs, Rajeswaran et al. (2014) studied the RNA silencing machinery of Cavendish banana plants persistently infected for 15 years with one of the six following BSV species: BSOLV, BSGFV, BSIMV, BSMYV, banana streak CA virus (BSCAV) and BSVNV. They demonstrated that despite the presence of abundant viral siRNAs of different size classes – mainly 21 nt long, reflecting the establishment of a PTGS mechanism – the virus was still present. Besides, they showed that the viral DNA of BSV was largely free of cytosine methylation despite the existence of 24 nt long siRNA, a marker of transcriptional gene silencing (TGS). Thus, the virus is able to evade siRNA-­ directed DNA methylation in the nucleus by a still

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uncharacterized mechanism and thereby retain the potential for active Pol II transcription. This evasion of silencing likely contributes to the persistence of BSV infection in banana plants. Seeded M. balbisiana (BBw) accessions ‘Pisang Klutuk Wulung’, ‘Pisang Batu’ and ‘Kluai Tani’ do not seem to develop banana streak infection from eBSV, whereas those eBSVs are responsible of the disease appearance among interspecific progenies (Lheureux et al., 2003; F. Bakry, France, 2016, personal communication). Lheureux (2002) also demonstrated that ‘Pisang Klutuk Wulung’ never developed banana streak infection after mealybug vector-­ mediated inoculation with BSOLV, BSGFV and BSMYV, concluding that these seeded BB diploids were resistant to multiplication of these three viruses. Preliminary data from Northern blot and deep sequencing analysis of total siRNA in ‘Pisang Klutuk Wulung’ indicated that eBSV are likely regulated at the transcriptional level in banana (Chabannes and Iskra-Caruana, 2013; P.O. Duroy, N. Laboureau, J. Seguin, R. Rajendran, M. Pooggin, M.-L. Iskra Caruana and M. Chabannes, France, 2017, unpublished information). Interestingly, some cultivars are susceptible to BSV despite having two copies of the B genome (ABB genotypes) (Sebasigari and Stover, 1988; K.S. Crew, L.A. McMichael and J.E. Thomas, Australia, 2015, unpublished results). BSIMV has been detected in ‘Pisang Awak’ and ‘Chuṍi Ngu Tien’ and BSOLV in ‘Chuṍi Ngop Diu Duc’, all using IC-PCR with a DNase treatment to eliminate genomic DNA, and bacilliform virions were also observed by electron microscopy in the latter cultivar (K.S. Crew, L.A. McMichael and J.E. Thomas, Australia, 2015, unpublished results).

Disease cycle and epidemiology BSV transmission BSVs as badnaviruses are commonly horizontally transmitted by mealybugs whether the origin of the viral particles is exogenous or endogenous (eBSVs) (Fig. 6.4). Endogenous infections are the result of spontaneous activation of infective eBSV allowing the release of a viral genome (Fig. 6.4). Several genomic and abiotic

stresses have been shown to be involved in the activation of eBSVs. Interspecific crossing context, micropropagation by in vitro culture and strong soil moisture and/or temperature changes play a major role in the occurrence of BSV infections in newly created interspecific and natural hybrids harbouring one copy of the B genome (Harper et  al., 1999; Ndowora et  al., 1999; ­Dallot et al., 2001; Lheureux et al., 2003; Côte et al., 2010). No clear relationships between BSV species and mealybug species have been observed. Experimental mealybug-mediated vector transmission has been demonstrated using Planococcus citri (the citrus mealybug) (Plate 6.48), P. ficus (vine mealybug), Saccharicoccus sacchari (sugarcane mealybug) and Dysmicoccus brevipes (pineapple mealybug) (Kubiriba et al., 2001; Meyer et al., 2008). The mode of transfer is semi-persistent, as reported for Commelina yellow mottle virus (CoYMV), the type member of the badnavirus group. In Nigeria, P. citri has not been reported, but P. musae has been identified in fields where the incidence of banana streak is high (Dahal et al., 1998b). In Ecuador, P. comstocki is the most common mealybug found on banana, but transmission of BSVs by this species has not yet been demonstrated. The diversity of mealybugs involved in BSV transmission is mostly driven by geographical localization rather than biological affinities. Field observations in many countries suggest that natural dissemination of BSVs by mealybug vectors is slow (Daniells et al., 2001), limited in occurrence and does not play a major role in disease epidemi­ ology (Lockhart, 1995; Iskra-­Caruana et  al., 2014b). However, in Ecuador, an efficient collaboration between ants and mealybugs ­ has been observed where ants manage to move mealybugs from plant to plant, thus likely spreading the disease faster than usual (D.R. Jones, Australia, 1996, personal communication). Nevertheless, the spread of BSVs is mainly due to vegetative propagation of ­infected source plants. BSVs are readily transmitted to the suckers of infected plants (Daniells et al., 2001) and through tissue culture (Dallot et al., 2001). Attempts to transmit BSVs by mechanical inoculation have been unsuccessful (Lockhart, 1986) and it is thus



Diseases Caused by Viruses 405

A : Classic triangular BSV interaction Infected plants

Virus free plants

Outcome

Newly infected plants

AAAA without eBSV

AAB hybrid with eBSV

Vector transmission BB (PKW) with eBSV

No virus

B : Special quartet BSV interaction Virus free plants

Outcome

Newly infected plants

AAAA without eBSV

Stress No virus

AAB hybrid with eBSV

re

Ho c o m olo g ous mb i n a ti o n .. ?

BSV

BBw (‘Pisang Klutuk Wulung’) with eBSV No virus

Fig. 6.4.  Schematic representation showing the two ways of BSV spread; either through a classic triangular interaction (A) or an unusual quartet interaction (B) (Iskra-Caruana et al., 2010). The circles superimposed on the virus-free plants represent the one or two copies, where present, of the B genome. Each copy of the B genome contains the three inducible infectious eBSVs. Interaction A. In the ‘classic triangular interaction’, BSV is mealybug transmitted from an infected plant, which is the primary inoculum source, to virus-free plants, which become infected and serve as secondary inoculum sources. BSV spread is described for three kinds of virus-free plants having different genotypes ­regarding eBSVs: a plant without eBSV (AAAA genome); an interspecific hybrid plant (AAB) harbouring inducible infectious eBSVs (only one copy of B genome) and a wild M. balbisiana diploid plant (‘Pisang Klutuk Wulung’) which is a carrier of infectious eBSVs. Disease development occurs in plants without Continued

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Epidemiology of BSV

Plate 6.48.  A colony of Planoccocus citri, a mealybug vector of banana streak virus, on a banana peduncle (photo: B.E.L. Lockhart, UM).

unlikely to be transmitted on cutting tools or during cultural operations. It is also not soilborne. The natural transmission rate of sugarcane bacilliform viruses from sugarcane growing in close proximity to banana is not known, but is expected to be a rare event.

The cause of banana streak epidemics remains unclear though they have been described since 1990 in Uganda, where BSVs are endemic (Tushemereirwe et  al., 1996). The recent data on BSV phylogeny has allowed the development of a hypothesis to describe this unusual BSV epidemic situation. Based on the BSV phylogenetic separation into two clades, Iskra-Caruana et al. (2014a) proposed a scheme retracing the coevolution of BSV and Musa. Briefly, they suggested that BSVs of Clade 1, which are a reflection of past infections, are responsible for outbreaks worldwide due to eBSV activation mainly associated with human activities (breeding, tissue culture) whereas BSVs of Clade 3 are responsible for recent and local epidemics in Uganda. This epidemic context is probably maintained by the use of infected banana planting material rather than mealybug-mediated transmission. Both could represent a real threat for de novo epidemics, but such a situation has not occurred so far. Surprisingly, as opposed to newly created interspecific hybrids, natural and local banana cultivars harbouring infective eBSV for BSOLV, BSGFV and BSIMV species are usually virus-free and do not contribute as latent reservoirs for the initiation of banana streak epidemics. One explanation is that viral fixation of the three BSV species within the genomes of M. balbisiana seeded diploids long ago has contributed to better fitness of banana diploid populations at that time in a period of a strong viral pressure. Nowadays, as the result of coevolution, all seeded diploids of M. balbisiana are virus-free and

Fig. 6.4. Continued. eBSV and interspecific hybrids harbouring eBSVs. These plants then become virus reservoirs for further horizontal/vector transmission. However, ‘Pisang Klutuk Wulung’ appears resistant to BSV as infection never occurs by vector transmission. The mechanism underlying this resistance is still unknown. Interaction B. In the ‘unusual quartet BSV’ interaction, primary infection is caused by BSV arising from within the plant’s genome (eBSVs). Following stress, which triggers eBSV expression, the interspecific hybrid plants are the only ones that become infected. The mechanism underlying the transformation from integration to free circular viral genome is still unknown, although homologous recombination is strongly suspected. After infection, the hybrid then becomes a reservoir for further horizontal/vector transmission as described in Interaction A. ‘Pisang Klutuk Wulung’ (BBw), which harbours the same infectious integrations as hybrid plants, but at a mono- or di-allelic level, presents a natural resistance to this internal activation. Here again, the mechanism underlying this resistance is still unclear, although virus resistance could occur as in petunia and tobacco (Mette et al., 2002; Noreen et al., 2007) via homology-dependent transcriptional or post-transcriptional gene silencing.



Diseases Caused by Viruses 407

r­esistant to BSV multiplication, whatever the origin of the BSV (i.e. eBSV or external source) in regards to the integrated BSV species present. Conversely, natural hybrids with a B genome in their genotypes are younger and can be infected following spontaneous eBSV activation, but never become reservoirs for BSV spread. A natural regulation of BSV infection could be present in such plants, leading to the maintenance of a viral level under the optimal threshold for mealybug transmission.

Control Collection, conservation and utilization of plant genetic resources are essential components for international crop improvement programmes. However, the movement of germplasm involves a risk of accidentally introducing plant pests along with the host plant. Viruses, because they are often symptomless, pose an important risk. This is particularly true for BSVs which are primarily spread by vegetative propagation and whose symptoms appear sporadically during the course of the year. The most effective disease control strategy is to ensure that source plants used for propagation are virus-free. The episomal form of BSVs can be eliminated through meristem tip culture including thermotherapy or cryopreservation steps (Helliot et al., 2002) or chemotherapy with the use of two anti-retroviral and anti-hepadnavirus molecules (Helliot et al., 2003). To ensure that germplasm is BSV-free, different diagnostic procedures are available (see above). Banana cultivars harbouring infective eBSVs still pose a unique problem, because they can be first identified as BSV-free before suddenly become BSV-infected. Indeed, the micropropagation of such virus-free plants gives rise to a low proportion of BSV infected in vitro plantlets, because the in vitro process can activate the release of viral genome from eBSV in a few banana cells (Dallot et al., 2001; Côte et al., 2010). Interestingly, Côte et  al. (2010) demonstrated that competition during the intensive in vitro multiplication existed between virus-free and infected cells, resulting in a high proportion (up to 85%) of virus-free plantlets, according to the cultivar

studied. Similar eBSV activation was observed for all other macro-propagation processes, including the partial internal fragment (PIF) technique. However, the latter results in a higher proportion of BSV-infected plants compared with in vitro culture process, due to a slow cell competition. It has also been shown that eBSOLV and eBSGFV are differently activated in several cultivars that have similar infective eBSVs (Côte et al., 2010). Therefore, it is not possible to guarantee BSV-free planting material with plants harbouring infective eBSVs. Thomas et al. (2015) proposed a strategy to distribute banana germplasm with eBSV including natural and artificial hybrids, which were up till now unavailable due to the difficulties of assessing the risk of BSV emergence. This strategy incorporates several controls, including the full eBSV allelic genotyping of the plant using specific PCR and dCAPS molecular markers for the three BSV species concerned (BSOLV, BSGFV and BSIMV) (Gayral et  al., 2008; Chabannes et al., 2013) and the elimination of any episomal infections to ensure that only eBSVs are present (Fig. 6.5) Because of the high conservation of these eBSVs in natural, cultivated and synthetic hybrids and the majority of known seeded BB diploids (Duroy et  al., 2016), those tools are powerful, robust and useful for marker-assisted selection (MAS) in breeding programmes. This strategy was recently used in collaborative work between CIRAD and CARBAP (Centre africain de recherches sur bananiers et plantains), which resulted in the production of eBSV-free AAB triploid offspring (i.e. without infective and non-­infective eBSV) in mimicking the conventional plantain breeding programme 4×/2× (Noumbissie et al., 2016). In parallel, CIRAD is developing different strategies (self-pollination or duplication of chromosome sets from haploid lines) where eBSV molecular markers are needed to produce BB diploid genitors devoid of any infective eBSV (Umber et  al., 2016). These results pave the way for the safe and quick reintroduction of M. balbisiana genotypes into breeding programmes by MAS. These markers, which are free for use by the worldwide community, should help to control or avoid the dissemination of synthetic hybrids with eBSV capable of activation.

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Musa

Musa clones without B genome

Musa clones with B genome Non-infectious eBSV allele

INDEXING FOR BSV*

INDEXING FOR BSV

BSV not detected

BSV detected

Therapy + indexing + selection

DISTRIBUTION

Musa clones with B genome infectious eBSV allele

BSV not detected* Informed consent from the importer to receive germplasm. Include disclaimer in the health statement : BSVs genomes are present in the B genome of Musa (banana and plantain). Consequently, almost all accessions containing the B genome may develop BSV infection and may express symptoms during any stage of growth.

BSV detected* Therapy + indexing + selection*

DISTRIBUTION

*Note: Usual procedures to eliminate episomal BSV infections not originating from eBSV will be applied. However viral particles may still occur even after treatments due to activation of eBSV allele as indicated in the disclaimer statement.

Fig. 6.5. Schematic plan of the plant health system developed to reduce the risk of spread of banana streak viruses (BSVs) with Musa germplasm (Thomas et al., 2015). The scheme has been developed to facilitate the international distribution of Musa germplasm by reducing the risks of spreading BSVs. The indexing protocols used must also be able to differentiate between active infections (episomal form) and endogenous BSV (eBSV). Episomal BSVs infecting plants are eliminated from germplasm through virus therapy before distribution. The use of virus therapy procedures is mandatory to eliminate all BSV infections. The ability to distinguish between those clones containing non-infectious or infectious eBSV allows greater confidence to be assigned to the risk of distribution of banana streak viruses. Although the physical indexing and therapy treatments of accessions are the same for all three categories, the schematic plan highlights specific points relevant to accessions containing eBSVs. Three categories of accessions are considered when indexing for BSV: 1. Musa accessions without the B genome. If indexed and found free of BSV, they can be distributed. If BSV is detected, they can undergo virus therapy and if free of BSV upon reindexing, they can be distributed. 2. Musa accessions with the B genome and only non-infectious eBSV. If indexed and found free of BSV, they can be distributed. If BSV is detected, this will be very likely be because of episomal infections not derived from eBSV, and they can undergo virus therapy. If free of BSV upon re-indexation, they can be distributed. 3. Musa accessions with the B genome and infectious eBSV. If indexed and found free of BSV, they can be distributed. If BSV is detected, this can be of either non-eBSV origin or derived from eBSV or both. These accessions undergo virus therapy to eliminate all BSV particles to be sure that episomal infection of noneBSV origin is eliminated prior to distribution. When indexed free of BSV and other known viruses, accessions in categories 1 and 2 are available for distribution. For accessions in category 3, it is proposed that the National Plant Protection Organization (NPPO) of the importing country should be notified of the proposed importation of any eBSV-containing germplasm, and their written acceptance of this importation must be obtained before distribution. The health statement attached to the germplasm will include the following disclaimer: eBSVs are present in the B genome of Musa. This is a warning that accessions containing the B genome may develop BSV infection and may express symptoms during any stage of growth.



Diseases Caused by Viruses 409

Enset Streak M. Tessera, A.J. Quimio and D.R. Jones Enset streak was first noticed at Hagereselem in the Sidamo zone and at the Areka Experimental Station in the North Omo zone of Ethiopia in 1991 (Tessera and Quimio, 1994). The disease was later recorded from the major growing areas (Tessera et al., 1996). Chlorotic and yellow mosaics, streaks and stripes are characteristic leaf symptoms of the disease (Plates 6.49 and 6.50). Severely affected plants also have narrow, distorted leaves and become stunted (Plate 6.51). Chlorotic areas of leaves later turn necrotic (Plate 6.50). Early infection results in a significant reduction in yield. Symptoms are more pronounced in the winter than in the summer (Tessera et al., 1998).

Enset streak is believed to be caused by a badnavirus (Tessera et al., 1996). Virus particles are bacilliform in shape and dimensions are given as 118–125 nm × 29.5–30.0 nm (Tessera et al., 1998). The relationship of this virus with species of BSV in banana has not been determined.

Plate 6.50.  Narrow, distorted leaves of a young enset plant affected by enset streak disease in Ethiopia. Areas of leaf tissue with yellow streaks are turning necrotic (photo: A.J. Quimio, IAR).

Plate 6.49.  Yellow streak symptoms on an enset leaf caused by enset streak disease in Ethiopia (photo: M. Tessara, IAR).

Plate 6.51.  Chlorotic and yellow streaks plus distortion on the leaf of an enset plant in Ethiopia with enset streak disease (photo: A.J. Quimio, IAR).

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­ owever, a badnavirus found in Ensete venticoH sum with stripe symptoms in Kenya was identified as banana streak GF virus (Geering and Thomas, 2002). The major means of dissemination of the disease is through infected corms, or suckers arising from an infected corm. A vector has not yet been identified and there is no information on clonal susceptibility, though various cultivars have been seen affected. The use of healthy corms for propagation is advocated for the control of the problem.

Banana Mild Mosaic J.E. Thomas, B.E.L. Lockhart, M.L. Iskra-Caruana, K.S. Crew and M. Sharman Introduction

mosaic and streak symptoms (Plate 6.52). These symptoms, which are also often seen on young ‘Ducasse’ plants derived from infected tissue cultures, may disappear as the plant matures. Transitory chlorotic leaf streaks were noted on some leaves of infected ‘Daluyao’ (AAB, Plantain subgroup) growing under glasshouse conditions (J.E. Thomas, Australia, 1998, unpublished results). ‘Pisang Seribu’ (AAB) displays silvery streaks on the leaf lamina, especially on the distal part, which are best viewed from the underside of the leaf (Plate 6.53). Plants of ‘Gros Michel’ (AAA) from Latin America have displayed large chlorotic blotches on the leaves, stunting and delayed bunching (B.E.L. Lockhart, USA, 1998, unpublished results). Mixed infections of the virus and banana streak virus (BSV) are common (De Clerck et al., 2017) and in such cases symptoms may be similar to those caused by BSV alone (M.L. Iskra-­ Caruana, B.E.L. Lockhart and J.E. Thomas, 1998, France, USA, Australia, unpublished results).

In the 1990s, flexuous filamentous virus-like particles were frequently observed in Musa germ­ plasm accessions and field specimens (Rivera et  al., 1992; INIBAP, 1993; Lockhart, 1994b; M.L. Iskra-Caruana and J.E. Thomas, France and Australia, 1995, unpublished results; Caruana and Galzi, 1998; Belalcázar et  al., 1998). The economic impact of this virus is believed to be low. Observations in Latin America indicate that yield may be affected in both banana and plantain under some conditions (B.E.L Lockhart, USA, 1998, unpublished results). The virus probably occurs worldwide, wherever banana is grown, and is found extensively in germplasm collections (Thomas, 2015; De Clerck et al., 2017). Records exist from Australia, Africa, Southern and Southeast Asia, India, Central and South America and the Caribbean, in a wide variety of Musa cultivars and genotypes (Reichel et al., 2003; Teycheney et al., 2005a; Selvarajan and Balasubramanian, 2016; De Clerck et al., 2017). Symptoms Symptoms are often mild or absent. ‘Ducasse’ (ABB, ‘Pisang Awak’ subgroup) in the field in Australia has been observed with chlorotic

Plate 6.52.  Chlorotic mosaic and streak symptoms of banana mild mosaic disease on a leaf of ‘Ducasse’ (ABB, Pisang Awak subgroup) in Queensland, Australia (Photo: J.E. Thomas, QDAF).



Diseases Caused by Viruses 411

In mixed infections with cucumber mosaic virus (CMV), the presence of the virus seems to correlate with an additional leaf necrosis symptom (Caruana and Galzi, 1998). Plants with mixed infections of the virus and banana bract mosaic virus (BBrMV) display the spindle-shaped lesions on the leaf lamina typical of BBrMV infection (Iskra-Caruana et al., 2008).

Causal agent Banana mild mosaic virus (BanMMV) is an unassigned species in  the family Betaflexiviridae (King et  al., 2012). ­Virions are flexuous filaments about 580 nm long and 14 nm wide (Plate 6.54) (Thomas et  al., 1999b). The coat protein has an apparent Mr of 30 kDa in SDSPAGE (Caruana and Galzi, 1998). The concentration of virions in infected plants is variable, though generally low, with yields of purified

Plate 6.53.  Silver streak symptoms of banana mild mosaic disease on a leaf of ‘Pisang Seribu’ (AAB) in Florida (photo: B.E.L. Lockhart, UM).

virions of 0.2–1.0 mg/kg t­issue (Gambley and Thomas, 2001; Teycheney et al., 2007). BanMMV has a ca 7.4 kb ssRNA genome containing five ORFs (ORF1/polymerase, 205 kDa; ORF2-4/triple gene block, 25.5, 12.4 and 8.0 kDa; and ORF5/coat protein, 26.8 kDa), 3′ and 5′ untranslated regions and a polyA tail (Gambley and Thomas, 2001). Fairly high (28%) nucleotide sequence variation in conserved regions of the polymerase and coat protein encoding regions has been reported between isolates (Teycheney et al., 2005a), though this is not unusual for this family of viruses (King et al., 2012). Polyclonal antisera have been prepared to isolates from ‘M’bouroukou’ (AAB, Plantain subgroup) (B.E.L. Lockhart, USA, 1998, unpublished results), ‘Pisang Seribu’ (AAB) (Teychenney et al., 2007) and ‘Ducasse’ (C.F Gambley and J.E. Thomas, Australia, 1998, unpublished results). All work well in immunosorbent electron microscopy (ISEM), but only the latter is suitable, albeit with limited sensitivity, for ELISA. All isolates tested with polyclonal antibodies have

Plate 6.54.  Virions of banana mild mosaic virus from ‘Ducasse’ (ABB, Pisang Awak subgroup) in Australia. Scale bar = 200 nm (photo: J.E. Thomas, QDAF).

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been shown to be serologically related. Monoclonal antibodies have been prepared to an isolate from ‘Cardaba’ (ABB) and these have been used in tissue blot, Western blot assays (Caruana et al., 1995) and ELISA (Teycheney et al., 2007). Monoclonal antibodies raised against BanMMV can be used in ELISA, but they are very specific to the immunizing strain and therefore not suitable for routine diagnosis with this highly variable virus (Teycheney et al., 2007). PCR primers have been developed that allow detection of BanMMV in both total nucleic acid extracts from infected plants and in immunocapture RT-PCR using crude sap extracts (Teycheney et al., 2007; De Clerck et al., 2017). Some variation in PCR product size has been noted and this seems to be due to variations in the length of the 3′ untranslated region (M. Sharman and J.E. Thomas, Australia, 1999, unpublished results).

However, under natural conditions in Guadeloupe, plants established from virus-free tissue cultures of cultivars in the Cavendish subgroup (AAA) subsequently became field-infected with CMV and BanMMV, frequently as mixed infections, and the incidence of infection increased during the cropping cycle. This implies that natural transmission of BanMMV was occurring, generally simultaneously with the aphid-­ ­transmitted CMV (Caruana and Galzi, 1998).

Host reaction All Musa genotypes appear to be susceptible to BanMMV, though this observation is based only on natural infections.

Control Disease cycle and epidemiology Vegetative propagation, through both conventional planting material and tissue culture, is the only known means of transmission of BanMMV. Indeed, this virus is one of the most common contaminants of international germplasm collections, probably due to both its wide distribution and the frequently symptomless nature of infection (De Clerk et al., 2017). BanMMV has not been transmitted mechanically with sap extracts or partially purified preparations to banana or a range of herbaceous indicators, nor was it transmitted from banana to banana with the citrus mealybug (Planococcus citri) (M. Sharman and J.E. Thomas, Australia, 2002, unpublished results). The banana aphid, Pentalonia nigronevosa, did not transmit BanMMV from singly infected plants (M. Sharman and J.E. Thomas, Australia, 2002, unpublished results), and transmitted BBrMV, but not BanMMV, from banana infected with both viruses (Caruana et al., 1995). No transmission was observed when tissue culture plants of a Cavendish cultivar were grown in soil taken from the rhizosphere of BanMMV-infected ‘Ducasse’ plants, or when surface sterilized corms of ‘Ducasse’ were planted in sterilized potting mix with Cavendish plants and root contact allowed (M. Sharman and J.E. Thomas, Australia, 2002, unpublished results).

The use of indexed, virus-free vegetative planting material is recommended, though the economic impact of this virus is unknown, either alone or in mixed infections with other viruses. Efficient elimination of BanMMV using chemotherapy in vitro has been reported (Busogoro et al., 2006).

Banana Virus X K.S. Crew and J.E Thomas Banana virus X (BVX) was fortuitously discovered in banana ‘Som’ (ABB, syn. ‘Ney Mannan’) while assessing sequence diversity among banana mild mosaic virus (BanMMV) isolates in the CIRAD Guadeloupe field germplasm collection (Teycheney et al., 2005b). On further examination, five additional infected cultivars in Guadeloupe were identified resulting in a total of seven detections from 41 samples examined. A follow-­ up survey (Péréfarres et al., 2009) recorded BVX from plants in the Cavendish subgroup (AAA) (1/301 infected), ‘French Clair’ (ABB, Plantain subgroup) (3/271 infected) and ‘Figue Pomme’ (AAB, Silk subgroup) (3/193 infected). Surprisingly, the virus was not subsequently identified in a survey of the Guadeloupe collection in 2014



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(P.-Y. Teycheney, Guadeloupe, 2017, personal communication). No symptoms have been associated with infected ‘Som’ or other banana cultivars (Teycheney et  al., 2005b; P.-Y. Teycheney, Guadeloupe, 2017, personal communication). Sequence of 2917 nt at the 3′ end of the genome (excl polyA tail) is available. The genome organization resembles that of members of the family Betaflexiviridae, and this partial sequence includes part of the RdRp, the triple gene block proteins and the coat protein. Phylogenetic analysis and sequence comparisons indicate that the virus is only distantly related to other members of this family and may represent a new genus. By analogy to other members of the family, BVX is expected to have filamentous virions. A nested RT-PCR, incorporating direct binding of virions to the PCR tube, has been developed for the detection of BVX (Teycheney et al., 2007). BVX has only been recorded from naturally infected banana (Teycheney et al., 2005b).

Ampeloviruses K.S. Crew, S. Massart and J.E. Thomas Filamentous virions reminiscent of those of members of the Closteroviridae were observed by electron microscopy in five banana cultivars during routine screening of accessions from an international Musa germplasm collection (Crew et al., 2017). These cultivars originated from Indonesia and Vietnam, and were originally detected in plants growing in post-entry quarantine. Back-tracing confirmed that the original cultures in the in vitro collection, and indeed field plants of a sister line growing in an Indonesian collection contained similar virions. No symptoms have been associated with field or glasshouse-grown plants. Virions are extremely flexuous filaments (Plate 6.55). Two complete genomes have been assembled via high-throughput next-generation sequencing (NGS), and are likely to represent two previously undescribed ampeloviruses. These genomes have an organization consistent with

200 nm Plate 6.55.  Ampelovirus virion (arrow) from a partially purified miniprep negatively contrasted with 1% ammonium molybdite (photo: K.S. Crew, QDAF).

members of the genus Ampelovirus, share 50% nucleotide identity and are both 46% identical to the ampelovirus plum bark necrosis stem pitting associated virus (PBNSPaV), the nearest match on GenBank. The genome of virus 1 is 13,235 nt in length and that of virus 2 is 13,229 nt, both apparently comprising eight ORFs, six of which have so far been confidently assigned. At the amino acid level, individual ORFs (1a, 1b/ RdRp, 3/HSP70, 4/p61, 5/CP, 6) share 31–58% identity with each other, and both share 32–50% identity with PBNSPaV isolates, the nearest BLASTp matches, across the same ORFs. The banana ampeloviruses are serologically related to another tentative ampelovirus, sugarcane mild mosaic virus (SMMV). Antiserum prepared against the latter (Lockhart et al., 1992) gave moderate, light or negligible decoration of virions from several banana accessions in electron microscope serology. Partial sequences from several of these accessions also supported the existence of considerable diversity within this group of ampelovirus isolates. The viruses can be identified to the genus level using electron microscope serology with antiserum to SMMV. Reliable nucleic acid-based diagnostic assays have not yet been developed, due to the uncertainty surrounding the extent of virus diversity. As tentative ampeloviruses, they are assumed to be mealybug-transmitted. Banana is the only known host, though no attempts to transmit the viruses to alternative

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hosts have been reported. Virions have been observed in symptomless plants of cultivars ‘Chuṍi Mit’ (AB) and ‘Chuṍi Tay Tia’ (ABB) from Vietnam,

and ‘Pisang Lidi Bikittinggi’ (AA), ‘Pisang Jarum’ (AA) and ‘Pisang Kole’ (unknown genome) from Indonesia.

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Tessera, M., Lohuis, D. and Peters, D. (1996) A badnavirus in Ensete. In: Bekele, E., Abdulah, A. and Yemane, A. (eds) Proceedings of the 3rd Annual Conference of the Crop Protection Society of Ethiopia. Crop Protection Society of Ethiopia, Addis Ababa, pp. 143–148. Tessera, M., Goldbach, R.W. and Peters, D. (1998) Partial purification of the virus associated with enset chlorotic leaf disease. Pest Management Journal of Ethiopia 2, 106–109. Teycheney, P.-Y., Laboureau, N., Iskra-Caruana, M.-L. and Candresse, T. (2005a) High genetic variability and evidence for plant-to-plant transfer of Banana mild mosaic virus. Journal of General Virology 86, 3179–3187. Teycheney, P.-Y., Marais, A., Svanella-Dumas, L., Dulucq, M.-J. and Candresse, T. (2005b) Molecular characterization of banana virus X (BVX), a novel member of the Flexiviridae family. Archives of Virology 150, 1715–1727. Teycheney, P.-Y., Acina, I., Lockhart, B.E.L. and Candresse, T. (2007) Detection of Banana mild mosaic virus and Banana virus X by polyvalent degenerate oligonucleotide RT-PCR (PDO-RT-PCR). Journal of Virological Methods 142, 41–49. Thomas, J.E. (1991) Virus indexing procedures for banana on Australia. In: Valmayor, V.V., Umali, B.E., Besjosano, C.P. (eds) Banana Diseases in Asia and the Pacific: Proceedings of a Technical Meeting on Diseases affecting Banana and Plantain in Asia and the Pacific, Brisbane, Australia, 15–18 April 1991. INIBAP, Montpellier, France, pp. 144–157. Thomas, J.E. (ed.) (2015) MusaNet Technical Guidelines for the Safe Movement of Musa Germplasm. Bioversity International, Rome. Thomas, J.E. and Dietzgen, R.G. (1991) Purification, characterization and serological detection of virus-like particles associated with banana bunchy top disease in Australia. Journal of General Virology 72, 217–224. Thomas, J.E. and Magnaye, L.V. (1996) Banana Bract Mosaic Disease. Musa Disease Fact Sheet No. 7. International Network for the Improvement of Banana and Plantain, Montpellier, France. Thomas, J.E., Iskra-Carana, M.L. and Jones, D.R. (1994a) Banana Bunchy Top Disease. Musa Disease Fact Sheet No 4. INIBAP, Montpellier, France, 2 pp. Thomas, J. E., McMichael, L. A., Dietzgen, R. G., Searle, C., Matalevea, S. and Osasa, A. (1994b) Banana streak virus in Australia, Western Samoa and Tonga. In: Abstracts of the 4th ISSCT (International ­Society of Sugar Cane Technologists) Pathology Workshop, Brisbane, Australia, 4–9 April 1994, p. 40. Thomas, J.E., Smith, M.K., Kessling, A.F. and Hamill, S.D. (1995) Inconsistent transmission of banana bunchy top virus in micropropagated bananas and its implication for germplasm screening. Australian Journal of Agricultural Research 46, 663–671. Thomas, J.E., Geering, A.D.W., Gambley, C.F., Kessling, A.F. and White, M. (1997) Purification, properties and diagnosis of banana bract mosaic potyvirus and its distinction from abaca mosaic potyvirus. Phytopathology 87, 698–705. Thomas, J.E., Iskra-Caruana, M-.L., Magnaye, L.V. and Jones. D.R. (1999a) Bract mosaic. In: Jones, D.R. (ed.) Diseases of Banana, Abacá and Enset. CABI Publishing, Wallingford, UK. Thomas J.E., Lockhart B.E.L., Iskra-Caruana, M.-L. (1999b) Banana mild mosaic. In: Jones D.R. (ed) Diseases of Banana, Abacá and Enset. CABI, Wallingford, pp 275–278. Thomas, J.E., Iskra Caruana, M.-L., Kumar, L.P., Roux, N., Chabannes, M. et al. (2015) Position paper on a strategy to distribute banana (Musa) germplasm with endogenous banana streak virus genomes. Working Group Meeting of the MusaNet Conservation Thematic Group, Montpellier, France, 5 May 2015. Thomson, D. and Dietzgen, R.G. (1995) Detection of DNA and RNA plant viruses by PCR and RT-PCR using a rapid virus release protocol without tissue homogenisation. Journal of Virological Methods 54, 85–95. Tsai, Y.P., Hwang, M.T., Chen, S.P. and Liu. S.S. (1986) An ecological study of banana mosaic. Plant Protection Bulletin (Taiwan ROC) 28, 383–387. Tushemereirwe, W.K., Karamura, E.B. and Karyeija, R. (1996) Banana streak virus (BSV) and an associated filamentous virus (unidentified) disease complex of Highland bananas in Uganda. InfoMusa 5, 9–12. Umber, M., Pichaut, J. P., Farinas, B., Laboureau, N., Janzac, B. et al. (2016) Marker-assisted breeding of Musa balbisiana genitors devoid of infectious endogenous Banana streak virus sequences. Molecular Breeding 36, Article 74 (11 pp). Vakili, N.G. (1969) Bunchy top disease of bananas in the Central Highlands of South Vietnam. Plant Disease Reporter 53, 634–638. Velasco-Magnaye, L. and Eloja, A.L. (1966) Some physical properties and suscept range of the abaca mosaic virus. Philippine Phytopathology 2, 22–30. Vetten, H.J., Dale, J.L., Grigoras, I., Gronenborn, B., Harding, R. et  al. (2012) Nanoviridae. In: King, A.M.Q., Adams, M.J. Carstens, E.B. and Lefkowitz, E.J. (eds) Virus Taxonomy: Ninth Report of the International Committee on Taxonomy of Viruses. Elsevier, Academic Press, Amsterdam, pp. 395–404.

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7 

Nematode Pathogens D.L. Coyne and S. Kidane

Overview Many different plant parasitic nematodes are found in association with banana, abacá and ­enset, but only a relatively small number cause significant damage. These nematodes often occur in mixed populations, which can create difficulties in assessing the damage caused by each species and thus establishing their relative importance. The effect of one species on the host may be similar to that of another, resulting in a general reduction in crop growth and loss of yield. However, each species can cause specific symptoms and may require different control strategies. Conversely, a number of management options may form a common basis for the control of more than one nematode pathogen.

Burrowing Nematode Introduction The burrowing nematode, first described from banana in Fiji (Cobb, 1893), is generally viewed as one of the most important root parasites of banana in tropical areas (Stover, 1986; Gowen et  al., 2005; Jones, 2009) and the cause of a costly disease affecting commercial plantations growing cultivars in the AAA Cavendish subgroup (Sarah, 1989; Stanton, 1994). It is not found on banana growing at altitude, such as in

the ­highlands of Central and East Africa, or in the higher latitude zones, such as the Mediterranean area, Canary Islands, Madeira, the Cape Province of South Africa and Taiwan (Stover, 1972; Bridge, 1988; Sarah, 1989; Gowen et al., 2005), though it may be present in these regions under greenhouse cultivation. The present geographical distribution of the burrowing nematode is a reflection of historical movements of infected planting material, especially corms of Cavendish cultivars, and of the temperature preference of the pathogen (Price, 2006). It also causes black head rot or tip-over disease of abacá (Anderson and Alaban, 1968; Davide, 1972; Castillo et al., 1974). The burrowing nematode destroys root and corm tissue, which reduces water and nutrient uptake. It also has a deleterious effect on root anchorage, which results in the uprooting or toppling of heavily affected plants, particularly during windstorms and heavy rain once bunches have developed. The damage also reduces plant growth and development. In banana, this may lead to severe reductions in bunch weight and a significant lengthening of the crop cycle (Gowen, 1975; Stanton, 1994; Coyne et al., 2005). Crop losses depend on several factors, including the pathogenicity of local burrowing nematode populations, associated pathogens (including other nematode species), banana cultivar, climatic conditions and soil factors, especially fertility. In commercial plantations of

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­ avendish cultivars in areas of Côte d’Ivoire C where soils are poor, reduced bunch weights and ­toppling have been reported to cause losses of over 75% (Sarah, 1989). In such circumstances and without nematode control, banana plants become virtually unproductive after the first harvest. In the more fertile peat soils of Côte d’Ivoire and in the volcanic soils of Cameroon, cumulative crop losses are generally below 30% (Melin et al., 1976; Sarah, 1989). In South Africa, losses have reached 75% (Jones and ­ Milne, 1982). In Central America (Costa Rica and ­Panama) and South America (Colombia), crop losses estimated by counting uprooted plants fluctuate between 12% and 18%. Losses have been recorded as around 5% in the Sula Valley in Honduras (Pinochet, 1986). The actual economic impact of the burrowing nematode on smallholder cultivation is difficult to estimate. However, severe symptoms have been observed in cultivars in the AAB Plantain subgroup in Côte d’Ivoire growing near Cavendish plantations (Sarah, 1985) and damage has reached 50% in experimental plots (Sarah, 1989; Coyne et al., 2013). In Honduras, Stover (1972) reported that there was considerably more uprooting, which resulted in complete loss of yield, in burrowing nematode-infested plots of ‘Horn’ (AAB, Plantain subgroup) than in control plots. In Cameroon, cumulative losses of 60% were recorded in ‘French Sombre’ (AAB, Plantain subgroup) planted in a field naturally infested with the burrowing nematode (Fogain, 2001). In south-­ western Nigeria, yield losses averaging 29% were recorded for 17 banana, plantain and bred hybrid cultivars grown for two cropping cycles in soil with a burrowing nematode-­ dominated population of nematodes (Dochez et al., 2009). A combined burrowing and spiral nematode infestation reduced yields and caused toppling of ‘Obino l’Ewai’ (AAB, Plantain subgroup) in Nigeria with production losses of up to 90% being reported (Speijer and Fogain, 1999). Although not introduced into the ‘lowlands’ of East and Central Africa until the 1960s, there is compelling evidence that the burrowing nematode has contributed significantly to the decline of the AAA East African highland banana in this region (Speijer and Kajumba, 1996; Speijer et al., 1999; Price, 2006).

Symptoms Infection of banana corm and root tissues by the burrowing nematode results in a reddish-brown necrosis. On corms, this is clearly visible after the surface has been washed free of dirt and lightly peeled (Plates 7.1 and 7.2) with the necrosis usually focused around the points where roots leave the corm. Depending on the level of infection, the size of lesions varies from small spots to large ­areas of necrotic tissue. Damage caused by the banana weevil borer (Cosmopolites sordidus) is ­superficially similar to larger lesions, but extends much further into the corm tissue as tunnels (Plate 7.3). Infected roots have dark patches on the surface, which gradually coalesce as the nematode damage advances (Plate 7.4). The root eventually

Plate 7.1.  Banana corm showing necrotic patches caused by the burrowing nematode (photo: D. Coyne, IITA).

Plate 7.2.  Peeled banana corm showing damage caused by the burrowing nematode (left) with a healthy uninfested corm (right). Lesions extend from the root bases on the infested corm (photo: D. Coyne, IITA).



Nematode Pathogens

withers, blackens and dies. Uninfected roots are pale and firm. If infected roots are cut in half and sliced longitudinally, the symptoms of the burrowing nematode can easily be identified as reddish-brown necrotic patches extending from the surface towards the centre, but do not affect the central stele (Plate 7.5). Symptoms of burrowing nematode can be distinguished from those of Fusarium wilt as the latter are confined to vascular tissue and do not extend to the root surface. As discussed earlier, the main impact of the burrowing nematode is to weaken root systems so that plants easily topple during strong winds. Severe nematode damage will be observed in the

Plate 7.3.  Cross-section of the base of the banana pseudostem showing tunnel damage caused by the banana weevil (Cosmopolites sordidus) (photo: D. Coyne, IITA).

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corm and root tissue of such plants, which usually appear unthrifty with thin pseudostems and small bunches. Leaf cover is also reduced (Roderick et al., 2012a) and pseudostem turgidity can be affected, especially under periods of ­water stress. This leads to pseudostems snapping more easily (Coyne et al., 2013). Causal agent Burrowing nematode is the common name for the species Radopholus similis (Plate 7.6). It normally feeds at the advancing edge of necrotic lesions and can be isolated from the reddish tissue that is found here. Large numbers of the nematode can be obtained by teasing affected tissues in a dish of water or using a simple plate extraction method to quantify population densities (Coyne et al., 2014). More sophisticated methods, such as centrifugal flotation and mist extraction, allow for a more accurate quantitative evaluation (Hooper, 1986). It has been demonstrated that populations of R. similis are biologically diverse in terms of their host preference, reproductive fitness, pathogenicity and/or morphology. They can also differ biochemically and molecularly. Two ‘races’ of the nematode, one attacking banana, but not citrus, and the other attacking banana and citrus were demonstrated by Ducharme and Birchfield (1956). Later, Huettel et al. (1984) controversially proposed Radopholus citrophilus as the name of the

Plate 7.4.  Lesions caused by the burrowing nematode on banana roots (photo: J.L. Sarah, CIRAD).

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Plate 7.5.  Cross-section of banana roots showing damage caused by the burrowing nematode. Lesions extend from the exterior of the root to the central cylinder (photo: J.L. Sarah, CIRAD).

Plate 7.6.  Infestation of Radopholus similis in banana root tissue. All stages of the life cycle of the nematode from egg to adult are present (photo: M. Boisseau, CIRAD).

nematode attacking banana and citrus. The concept that they are indeed two pathotypes of the same species has since been supported by further morphological (Valette et al., 1998) and genomic (Kaplan et al., 1998, 2000; Haegeman et al., 2010) studies. Using different criteria, such as chromosome number, pathogenicity, reproduction rate and host preference, three pathotypes of R. simi­ lis were distinguished from Central America and the Caribbean (Edwards and Wehunt, 1971; Pinochet, 1979, 1988a; Tarté et al., 1981; Rivas and Roman, 1985). More recently, pathogenic diversity was reported to be worldwide and clearly linked to reproductive fitness in plant tissues. Isolates from Uganda, Côte d’Ivoire, Costa Rica and Guinea were shown to have higher multiplication rates than those from Martinique,

Guadeloupe, Sri Lanka and Queensland in ­Australia (Sarah et al., 1993; Fallas and Sarah, 1995a; Fallas et al., 1995; Hahn et al., 1996). Enzymatic phosphoglucose isomerase (PGI) and randomly amplified polymorphic DNA (RAPD) analyses revealed two genomic groups of burrowing nematode that are not related to pathogenicity (Fallas et al., 1996). In Africa, damage to banana by R. similis appears more severe than elsewhere (Pinochet, 1979, 1988a, 1988b; Marin et al., 1999). Populations from Uganda seemed to cause the most damage (Fallas et al., 1995). Diversity in the ­reproductive fitness and virulence of R. similis populations from Uganda on banana was later demonstrated, with some isolates being particularly aggressive (Dochez et al., 2005). Further assessment established that some of these aggressive populations were able to reproduce and damage roots on banana carrying the two widely confirmed sources of genetic resistance against R. similis (Plowright et al., 2013). The distribution of these genomic groups appears to be linked to historical movements of planting material. In the case of Uganda, the aggressive populations appear, on the basis of phylogenetic analysis, to originate from Sri Lanka (Plowright et al., 2013). Disease cycle and epidemiology Penetration occurs preferentially at the root apex, but R. similis is able to invade any portion of the root length. The nematode migrates in and between cells in the root cortex, where it feeds on the cell cytoplasm. This results in collapsed cell walls, cavities and tunnels (Blake, 1961, 1966; Valette et al., 1997). On corms, lesions develop where infected roots are attached and then spread outwards. Necroses can extend to the whole cortex of corms (black-head disease) and roots, but the root stele is usually not damaged, except occasionally on very young roots (Mateille, 1994b; Valette et al., 1997). Radopholus similis is a migratory endoparasite, which completes its life cycle in 20–25 days under optimal conditions. Embryonic development takes 4–10 days and the four juvenile stages are completed in 10–15 days, depending on temperature (Van Weerdt, 1960; Loos, 1962). This species has a pronounced sexual dimorphism, in which males present an atrophied stylet and are



Nematode Pathogens

considered to be non-parasitic. Males also survive longer than females, an attribute that apparently enables them to fertilize females after becoming adults without competing for food (Chabrier et al., 2010). Juveniles and adult females are actively mobile. They may migrate into the soil under adverse conditions and move towards new roots. The temperature range for R. similis development lies between 24°C and 32°C, with optimum reproduction occurring at around 30°C (Loos, 1962; Fallas and Sarah, 1995a). It  does not reproduce below 16–17°C or above 33°C (Fallas and Sarah, 1995a, b; Pinochet et al., 1995). Necrosis of root and corm tissues is accelerated if other organisms, such as fungi and bacteria, are present. Fungi commonly associated with burrowing-nematode lesions are Cylindro­ carpon musae, Acremonium stromaticum, Fusari­ um spp. and Rhizoctonia solani (Laville, 1964; ­Pinochet and Stover, 1980). Fungi of the genus Calonectria have been found to be pathogenic on banana in the French Antilles and Cameroon, causing lesions similar to those of R. similis. In association with the nematode, they can cause severe damage (Loridat, 1989). Studies on the interaction between R. similis and Xanthomonas campestris pv. musacearum, the cause of a bacterial wilt of banana, show that root wounds caused by the nematode act as entry points for the bacterium present in the surrounding soil (Shehabu et al., 2010). Environmental factors and stages of plant development will influence nematode population densities. As a rule, R. similis is less influenced by soil conditions than other species, and this may be due to its strictly endoparasitic habit (Quénéhervé, 1988). Rainfall appears to be the main factor (Melin and Vilardebo, 1973; Jaramillo and Figueroa, 1976; Vilardebo, 1976; Jones and Milne, 1982; Hugon et al., 1984; Sarah et al., 1988; Quénéhervé, 1989a, b); too little or too much water suppresses nematode densities in the roots. Temperature also limits development, with R. similis generally absent in cooler banana-­ growing areas. During the crop cycle, R. similis densities increase gradually until after the emergence of the flower bud (Melin and Vilardebo, 1973; Vilardebo, 1976; Sarah, 1986). The increase is faster in the roots of suckers (Sarah, 1986), especially those that are not pruned (Mateille et al., 1984).

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Host reaction Radopholus similis is able to attack almost all banana cultivars, as well as abacá and other seeded Musa species (Gowen et al., 2005). At least 250 host plants have been listed as susceptible, among which there are many weeds and several crops of economic importance, such as black pepper, coconut, tea, tuber crops, fruit trees and ornamentals (Milne and Keetch, 1976; ­O’Bannon, 1977; Bridge, 1987). Parameters used to measure the reaction of Musa to R. similis include the number of nematodes on each plant, the number of nematodes in known weights of root, the percentage of infected roots and assessments of lesion damage on roots and corms (Wehunt et al., 1978; Pinochet, 1988b; Sarah et al., 1992; Fallas et al., 1995; Fogain, 1996; Speijer and Gold, 1996; Price and McClaren, 1996). Trial designs, sampling strategies and methods of statistical analysis have been reviewed by Price and McClaren (1996). Taking into consideration the above methods, Speijer and De Waele (1997) published a manual containing standardized protocols for the assessment of nematode damage on banana. In an attempt to simplify the measurement of parameters useful for identifying nematode resistance in banana under field conditions, Hartman et al. (2010) used an index that included the percentage of dead roots, the number of large lesions and nematode population density. In pot experiments using plants derived from in vitro propagation, root damage measured 12 weeks after inoculation correlated well with nematode infestation levels measured at 6–8 weeks (Fallas et al., 1995). Marin et al. (2000) developed a standard method for screening for genetic resistance using 200 nematodes in pots, while De Schutter et al. (2001) devised a single-­ root method for evaluating banana germplasm using an inoculum of 50 nematodes per 8 cm root section, with final nematode numbers measured after 12 weeks. This method optimizes the use of inoculum and has been modified to simultaneously assess resistance to a number of nematode species (Coyne and Tenkouano, 2005). A pot evaluation of wild Musa species has shown that Musa acuminata ssp. banksii is quite susceptible (Wehunt et al., 1978). In contrast, most accessions of Musa balbisiana tested have been very resistant (Fogain, 1996) or partially

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resistant (Dochez et al., 2006). Musa acuminata ssp. malaccensis, microcarpa, zebrina and burmannica (accession ‘Calcutta 4’) have been found generally to have moderate to good resistance (Wehunt et al., 1978; Fogain, 1996; Dochez et al., 2006). However, one accession of M. acuminata ssp. microcarpa has been rated as moderately susceptible (Wehunt et al., 1978). Musa textilis is moderately resistant (Price and McClaren, 1996). Consequently, R. similis is not reported as a major problem in abacá crops (Anunciado et al., 1977). Radopholus similis has not been recovered from enset. Diploid cultivars vary in their reaction to R. similis. ‘Pisang Mas’ (AA, Sucrier subgroup) and ‘Pisang Lidi’ (AA, syn. ‘Pisang Lilin’) have been found to have moderate resistance (Wehunt et al., 1978; Fogain, 1996). ‘Pisang Batuau’ (AA), ‘Pisang Oli’ (AA) and many accessions of ‘Pisang Jari Buaya’ (AA) are viewed as highly resistant (Wehunt et al., 1978; Fogain, 1996). However, ‘Pisang Jari Buaya’ has been found to be susceptible to some aggressive populations from ­Uganda (Plowright et al., 2013). ‘Guyod’ (AA) and ‘Tuu Gia’ (AA) are very susceptible (Wehunt et  al., 1978; Fogain, 1996). ‘Safet Velchi’ (AB, syn. ‘Ney Poovan’) appears to be very resistant (Price and McClaren, 1996). In a screening study of 55 banana accessions, it was found that some AA cultivars, from the Pisang Jari Buaya and Pisang Batuau subgroups, had good resistance to R. similis. In addition, 17 diploid accessions were observed to have partial resistance (Quénéhervé et al., 2009). Many cultivars in the Cavendish subgroup (AAA) have been estimated to be moderately susceptible to R. similis. ‘Gros Michel’ and its dwarf mutant ‘Cocos’ are less susceptible (­Wehunt et al., 1978; Price and McClaren, 1996). ‘Yangambi Km 5’ (AAA, Ibota subgroup) has very strong resistance to R. similis (Sarah et al., 1992; Price, 1994b; Fogain, 1996; Price and McLaren, 1996; Fogain and Gowen, 1998) and is often used as the resistant check in experiments. However, it has shown susceptibility to some Uganda populations (Plowright et al., 2013). Cultivars in the AAB Plantain subgroup are, on the whole, very susceptible to R. similis (Price, 1994b; Fogain, 1996; Price and McLaren, 1996, Coyne et al., 2005; Dochez et al., 2009). ‘Popoulou’ (AAB, Mai’a Maoli–Popoulu subgroup) is also very susceptible (Quénéhervé

et al., 2009). This susceptibility may be linked to M. acuminata ssp. banksii, the wild banana, which may have contributed both A genomes to these subgroups (Carreel, 1995; Fogain, 1996). ‘Focanah’ (AAB, Pome subgroup), ‘Figue Pomme Ekonah’ (AAB, Silk subgroup), ‘Pisang Kelat’ (AAB) and ‘Pisang Ceylan’ (AAB, Mysore subgroup) all have good resistance (Fogain, 1996; Price and McClaren, 1996). Not many ABB cultivars have been screened against R. similis. Of those that have, ‘Bluggoe’ and ‘Cardaba’ accessions appear moderately susceptible and ‘Pelipita’ moderately resistant (Price and McLaren, 1996; Dochez et al., 2009). Control In general, the control of burrowing nematode is not consciously practised in most smallholdings. This is largely because of a limited understanding of nematodes as the causal agents of the damage that farmers experience. As a consequence, most control methods discussed are those practised in commercial plantations. Chemicals have traditionally been relied upon to keep the burrowing nematode in check. However, with the withdrawal from use of many nematicides over recent years, because of environmental and human health concerns, there has been more alternative research into identifying suitable ­ ­options. Pre-planting measures Reducing nematode densities in the soil before planting and the use of cleansed or nematode-­ free plant material are of primary importance in the control of R. similis. Eradication of R. similis from the soil is virtually impossible. After the first detection of R. similis in South Africa, some drastic measures, which included roguing, burning, soil fumigation with methyl bromide and fallowing, were introduced without total success (Jones and Milne, 1982). However, by implementing a strict quarantine system on the movement of plant material from areas where R. similis was present in South Africa, the spread of R. similis was contained (Willers et al., 2002). Population densities of R. similis may be reduced by fallowing with non-host plants, of which a number have been identified, including



Nematode Pathogens

some cash crops (Milne and Keetch, 1976; Gowen et al., 2005). Using Panicum maximum (Poaceae) in Queensland, Australia (Colbran, 1964) and Chromolaena odorata (Asteraceae) in West Africa (Sarah, 1989) for 1 year proved successful in reducing populations to non-detectable levels. In Côte d’Ivoire, rotation with pineapple (Ananas comosus) helped to reduce R. similis populations (Sarah, 1989) and rotation with sugarcane (Sac­ charum officinarum) also met with some success in Central America (Loos, 1961). In Panama, an 18-month clean fallow period did not eradicate R. similis (Loos, 1961). An alternative method to fallow is soil cleansing. Loos (1961) reported that R. similis was eliminated after land in Honduras and Panama was flooded for 5–6 months. Flooding has also been used in Suriname (Maas, 1969). In Côte d’Ivoire, 6–7 weeks of complete flooding was as effective as 10–12 months of fallow for reducing nematode populations (Sarah et al., 1983; Mateille et al., 1988). However, this method is rarely practical, as land needs to be level and continued treatment requires a permanent water-­ supply. Chemical fumigation, such as with dichloropropene or methyl bromide, has been quite efficient for soil cleansing, However, this method is now generally not used, because of environmental hazards (WHO, 2006). Nematodes may be introduced into clean soil in new growing areas through infected corms and suckers. Even if visually clean, low, undetectable infections will multiply and spread, ultimately affecting the crop. This risk is overcome by the use of in vitro micropropagated plantlets that are free of nematode and other ­infections. Most banana planting material for commercial production is now supplied as tissue-­ cultured material, which should be the only type used when banana is grown in virgin soil. The uptake and use of plantlets derived from ­tissue culture is also increasing amongst some smallholders, especially those located near weaning nurseries (Dubois et al., 2006, 2013). The use of macropropagation techniques can substitute for tissue-cultured material if undertaken correctly and if safeguards are met. One improvised method has been shown to be very effective in producing and supplying healthy planting material to farmers, especially smallholders (Lefranc et al., 2010). The principle ­relies

435

on the use of corm material, sourced from healthy plants. Roots are removed by trimming and any necrotic areas are cut out by paring. The corm is then disinfected with hot/boiling water before being incubated, often split into two halves, in a  nematode-free medium, such as sawdust. A  wooden frame covered in polythene sheeting helps to maintain high humidity (Plate 7.7). The sprouting plantlets can be removed when of a suitable size and weaned in pots until ready for use. For larger suckers, a simple method of disinfection consists of paring the corm tissue to remove necrotic tissue. However, nematodes located deep within the cortex may escape removal. Storing pared material for 2 weeks may further reduce the nematode population (Quénéhervé and Cadet, 1985b), but such techniques cannot be applied to small suckers, which are quite sensitive and need to be replanted rapidly. Paring, followed by hot-water treatments (52–55°C for 15–20 min), has been a common and effective practice in Latin America and ­Australia (Blake, 1961; Stover, 1972; Pinochet, 1986). In commercial settings this can work well. However, hot-water treatments are cumbersome and require careful monitoring of temperature and immersion times to prevent the death of tissues. A recent modification involving

Plate 7.7.  Macropropagation of banana plantlets under locally constructed units (photo: D. Coyne, IITA).

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improve fertility and root development may increase plant tolerance to nematodes. Such ­ measures include ploughing before planting, incorporation of organic matter in the soil, fertilization and irrigation. In smallholder plots in West Africa where cultivars in the AAB Plantain subgroup are grown, Coyne et al. (2005) demonstrated the beneficial effect of mulching with organic matter to reduce losses to nematodes and improve crop performance. Their conclusion was that any mulch was better than no mulch. Post-planting nematode control has primarily relied on the applications of nematicides to banana plants through granular applications or drip irrigation (Gowen et al., 2005; Jones, 2009). In some locations, such as in the Canary Islands, Martinique and Colombia, emulsifiable compounds are applied as liquid sprays or through irrigation systems, and generally on the basis of two to three applications/year. The optimum application time, dose and frequency of applications are determined by nematicide efficiency, environmental conditions, population dynamics and the pathogenicity of local strains. In some banana-growing countries, nematicides have traditionally been applied on a regular basis with no attempt made to determine if treatments were necessary or not. Ideally, nematode levels should be checked periodically to determine treatment needs. The threshold for ‘triggering’ nematicide application will depend on local parameters, such as climatic and soil conditions, as well as aggressiveness of pathotypes. For this exercise to be worthwhile, the check needs to be based on accurate nematode counts. Nematodes must be extracted from plant material and surrounding soil using proven protocols (Sarah, 1991; Speijer and De Waele, 1997). Chemical control has in the past relied heavily upon the regular and repetitive use of the same nematicide. However, this resulted in Post-planting measures the rapid microbial degradation of the active inIn most cases where contaminated planting ma- gredients and/or the build-up of resistance in the terial has been used to initiate new plantations, nematode population rendering the treatment or where clean planting material has been plant- inefficient (Anderson, 1988; Hugo et al., 2014). ed in infested soil, R. similis will multiply quite Most of the previously relied-upon nematicides rapidly. Yield losses can be reduced through were labelled as being either extremely hazardpropping up or guying pseudostems to prevent ous or highly hazardous. Many have been protoppling. Improved drainage is also an important gressively removed from use (WHO, 2006), factor in reducing nematode damage in high-­ resulting in the search for less hazardous and rainfall regions, such as parts of Central America more environmentally friendly products (Zum (Pinochet, 1986). Similarly, any measures that Felde et al., 2009). the immersion of suckers in boiling water for 30 s (Coyne et al., 2010) simplifies the treatment and has been shown to be effective and accurate, especially for smallholder farmers (Tenkouano et al., 2006; Hauser and Messiga, 2010). Planting material can also be disinfected using chemicals. Dipping plant material in a nematicide has proved effective (Jones and Milne, 1982). Another method consists of immersing the planting material in a nematicide–mud mixture, which adheres to the surface, forming a nematicidal coat, and is known as as ‘pralinage’ (Vilardebo and Robin, 1969). Of increasing interest and development is the use of biologically based products for the treatment of planting material, especially plantlets derived from tissue culture. When plantlet roots are exposed to certain beneficial microorganisms in water suspension or by drenching the growing medium, the microorganisms enter the plants and become endophytic (Sikora et al., 2008). The bio-enhanced plants have been shown to be protected to a greater extent against nematodes in the field than plants that have not been treated, reducing nematode damage and improving yields (Waweru et al., 2014). Tissue-cultured planting material is an ideal candidate for enhancement with beneficial microorganisms. Although free of many pests and diseases, in vitro plantlets are also free of beneficial endophytes. Bio-enhancement returns the natural equilibrium to some degree (Dubois and Coyne, 2006). A range of beneficial endophytes, such as non-pathogenic Fusarium oxysporum, Trichoderma spp. and Bacillus spp., have been identified that offer protection (Sikora et al., 2008). However, there is as yet limited knowledge on the persistence of endophytes and how long they may provide protection.



Nematode Pathogens

437

and arbuscular mycorrhizal fungi are also receiving increasing interest for their additional plant-host protection qualities, as are endophytes (see section above). However, difficulties with mass production, shelf-life, and efficiency in regard to host specificity and quite precise soil or environmental conditions (pH, organic-matter content, composition of microfauna/flora) have hampered their development (Stirling, 1991; Cayrol et al., 1992; Davide, 1994). Formulations based on the parasitic fungus Purpureocillium lilacinus (previously known as Paecilomyces lilacinus) are probably the most widely used against R. similis on banana, with a number of products and formulations commercially available. The fungus parasitizes eggs, juveniles and adults. Results vary depending on conditions, but in general are favourable and economically viable. Species of Bacillus, such as B. firmus and B. subtilus, and strains of the bacterium Pseudomonas fluorescens have been demonstrated to inhibit the invasion of roots of banana by R. similis (Aalten et al., 1998; Mendoza  and Sikora 2009). The obligate nematode parasitic bacteria Pasteuria spp. differ in their host range and pathogenicity to nematodes. Pasteuria penetrans has been found parasitizing R. similis (Wang and Hooks, 2009; Sharmila et al., 2012), but has yet to be developed fully for use against this pest. The plant growth-promoting effects of arbuscular mycorrhizal fungi not only provide potential benefits to banana, but have also been shown to reduce R. similis infection and damage (Elsen et al., 2001, 2008). The total R. similis density was reduced by 60% and root necrosis by 56%, in banana plantlets colonized by arbuscular mycorrhizal fungi under greenhouse conditions (Koffi et al., 2012). However, as with many observations when assessing and applying biological control agents, reactions may be quite specific depending on host cultivar and control agent species or strain (Elsen et  al., 2003). Combinations of microbial control agents have also been assessed on a number of ­occasions. Although results can be variable, and Biological control often depend on l­ocal conditions, combined appliPlant-parasitic nematodes have many natural cations have demonstrated better control of R. si­ enemies and a number have been considered as milis than individual applications (Zum Felde et al., possible biological control agents, including om- 2009). Combined application of F. oxysporum and nivorous and predatory nematodes, nematode-­ B. firmus was more effective in controlling R. si­ trapping fungi, nematode-parasitic fungi and milis on banana than either alone or in combinabacteria. Plant health-promoting rhizobacteria tion with P. lilacinus (Mendoza and Sikora, 2009). Management of R. similis in the French West Indies traditionally relied upon the repeated ­application of carbamate or organophosphate nematicides, but an environmentally sound ­ scheme based upon the use of tissue cultures, fallow and intercropping with non-hosts has since been implemented (Risède et al., 2009). A similar scheme in Hawaii includes the incorporation of crop residue into the soil (HBIA, 2010). Synthetic nematicidal products that were developed taking into consideration a greater need for environmental safety continue to be released for use on banana. One such product is based on the fungicide fluopyram, which is marketed as being environmentally friendly. Another is a nematicide utilizing fosthiazate even though it is an organophosphate. As markets and developments evolve and fluctuate, the current status of such products and their efficacy in relation to local conditions need to be assessed and evaluated locally. Soil sanitation can be achieved through a  cleansing system based on injections of the ­herbicide glyphosate into banana plants before uprooting (Risède et al., 2009). Emphasis is also being increasingly placed on efforts to identify suitable biologically based solutions, such as the use of mycorrhizae, endophytes and bio-pesticides (Sikora et al., 2008; Viaene et al., 2013). Applications of plant extracts, some of which appear to provide good control options, while other data and assessments are less consistent or convincing, have received much attention. Of particular note are neem (Azidirachta indica) formulations, which, provided that they originate from reliable sources, can give very good nematode management on banana (Bartholomew et al., 2014). Products based on sesame oil, blends of essential plant oils that include sesame or garlic, furfuraldehydes and products based on Myrothecium verrucaria have been shown to be highly toxic to nematodes and can provide very promising reduction of R. similis.

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Breeding for resistance Fogain and Gowen (1997) demonstrated in field trials that population levels of R. similis were higher on the root systems of nematicide-treated susceptible cultivars than on an untreated resistant cultivar. Their work showed that genetic resistance can effectively control R. similis. Because of differences in pathogenicity among R. similis populations, as well as the other nematode species able to parasitize and damage banana roots, efforts in breeding banana for broad resistance against all these variants will be extremely difficult (De Waele, 1996). Nevertheless, potentially valuable banana cultivars have been evaluated against local populations of the burrowing nematode (Frison et al., 1997). Techniques for the early screening of germplasm in small pots have been developed (Pinochet, 1988b; Sarah et al., 1992; Fogain, 1996; De Schutter et al., 2001; Coyne and Tenkouano, 2005). Such methods allow susceptible germplasm to be ­rapidly identified and inferior lines eliminated, retaining only the most promising germplasm for final evaluation in more costly field trials (Price and McLaren, 1996). Several clones of ‘Pisang Jari Buaya’ (AA) have long been recognized as an exploitable source of resistance to burrowing nematode (­Pinochet and Rowe, 1978, 1979; Wehunt et al., 1978; ­Pinochet, 1988a; Ortiz and Swennen, 2014). The resistance of ‘Pisang Jari Buaya’ has been incorporated into breeding lines and this has led to the production of hybrids of commercial interest (Rowe and Rosales, 1994; Quénehervé et al., 2009). Screening studies have shown that sources of resistance that may be useful in breeding programmes are present in many genotypes (­Sarah et al., 1992; Price, 1994b; Price and McLaren, 1996; Fogain, 1996; Ortiz and Swennen, 2014). Pot tests in Honduras have shown that the bred hybrid ‘Goldfinger’/‘FHIA-01’ (AAAB) has resistance to R. similis, as have the synthetic AA diploids ‘SH-3142’, ‘SH-3362’, ‘SH-3648’ and ‘SH-3723’ (Viaene et al., 2003). Diploid banana hybrids bred for black leaf streak resistance in Nigeria have resistance to R. similis (Tenkouano et al., 2003). Similarly, three leaf spot-resistant banana hybrids (AAA) bred by ­CIRAD and designated ‘FB918’, ‘FB919’ and ‘FB924’ have also proved resistant to R. similis in Martinique (Quénéhervé et al., 2008).

It has been suggested that clones with large numbers of roots may exhibit a higher tolerance to nematode attack and selection for this character should be a worthwhile breeding objective (Gowen, 1996). Research of resistance mechanisms to R. similis have shown that phenolic compounds, especially some tannins and flavonoids, could be involved, reducing the inroads that nematodes make into banana tissues and their multiplication within these tissues (Mateille, 1994b; ­Valette et al., 1996, 1997). Dochez et al. (2009) found that resistance to R. similis is controlled by two dominant genes with additive and interactive effects, where one recessive genotype in one locus suppresses the dominant allele in the other locus. Resistance to R. similis through genetic improvement has long been hindered by difficulties associated with conventional banana breeding (Menendez and Shepherd, 1975; Pinochet, 1988a). Generating hybrids combining hostplant resistance with desired agronomic and quality traits from the cultivars remains a challenge. Nevertheless, good progress has been made in introgressing resistance to burrowing nematode in elite selections (Quénehervé et al., 2009; Lorenzen et al., 2010). New cellular and molecular banana improvement techniques continuously enable the natural limitations of traditional plant breeding to be circumvented (Ortiz, 2013). The genetic modification of existing cultivars is now presenting a realistic option for nematode management with the successful generation of resistant lines (Roderick et al., 2012b), which have confirmed resistance in the field in Uganda (Tripathi et al., 2015). Flow cytometry protocols, DNA markers, resulting genetic maps and the recent sequencing of the banana genome offer yet greater i­ nsights and help to identify useful genes (Ortiz and Swennen, 2014).

Root-lesion Nematodes Introduction Root-lesion nematodes occur widely, but not universally, on banana throughout the tropics (Bridge et al., 1997; Gowen et al., 2005). Like the



Nematode Pathogens

burrowing nematode, their distribution likely increased through the movement of infected clonal planting material. However, they are not found so commonly in commercial plantations of cultivars in the AAA Cavendish subgroup, where R. similis has traditionally been the most important nematode pest. Root-lesion nematodes have, in particular, been reported in association with the AAB Plantain subgroup (Ogier and Merry, 1970; Pinochet and Stover, 1980; Bridge et al., 1995; Speijer et al., 2001) with some evidence to indicate that plantain is more susceptible to these nematodes than are other banana types (Perez et al., 1986). Root-lesion nematodes have also been recorded on abacá (Davide, 1972). One of the root-lesion nematodes found on banana also attacks enset in ­Ethiopia (Addis et al., 2006).

439

plants become stunted, bunch weight is decreased and the production cycle is extended. Damage leads to a reduction in the size of the root system and toppling of plants. Plant toppling may be more prevalent in poor soils. Reduced plant growth, a diminished leaf cover and toppling can increase the exposure of soils to sunlight. This results in a rise in soil temperatures and a reduction in the organic content. Nutrient leaching and erosion may also occur in soils exposed to direct rainfall (Bridge et al., 1997).

Causal agent

It is possible that root-lesion nematodes are overlooked when they occur in mixed populations with R. similis or are mistaken for that species. The damage in roots and corms is identical to that caused by R. similis. Root-lesion nematodes feed on the cytoplasmic contents of cells in the cortex and migrate between and within cells. This causes the formation of cavities within the root tissue and results in characteristic, dark purple lesions and necrotic patches (Plate 7.8). Symptoms are usually confined to the cortex, while the stele tissue is generally unaffected – a useful diagnostic character when examining necrotic roots. ­Infected

Root-lesion nematodes are species of Pratylen­ chus, which can be confused with R. similis when nematodes are viewed under a dissection microscope (× 50). However, unlike R. similis, Praty­ lenchus males have functional stylets. To the experienced technician the genera can be differentiated by the position of the vulva, which is near to the tail in Pratylenchus spp. and at mid-length of the body for R. similis females. Many reports in the banana literature do not identify Pratylenchus to species level. The most widely reported is P. coffeae, with P. goodeyi recognized as probably the second most significant species. Differences in tail morphology help to separate the two species. Pratylenchus coffeae (Plate 7.9) infects a number of important crops, which include potato, yam, citrus, coffee, ginger (Luc et al., 2005), abacá and some ornamental plants. It is the most important nematode pest of banana in the Pacific

Plate 7.8.  Damage to a banana root caused by Pratylenchus goodeyi, a root-lesion nematode (photo: B. Pembroke, UR).

Plate 7.9.  Pratylenchus coffeae, a root-lesion nematode that attacks banana (photo: B. Pembroke, UR).

Symptoms

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and is significant in parts of Southeast Asia, especially in Thailand on ‘Klaui Namwa’ (ABB, Pisang Awak subgroup). This nematode is also reported as the most damaging on cultivars in the AAA Cavendish subgroup in Honduras. Recent surveys have shown an increased incidence of P. coffeae in West Africa, where it is also regarded as aggressive and displacing R. similis (Brentu et al., 2004; Coyne, 2009; Dubois and Coyne, 2011). In Ghana, it was found to be one of the two most widespread species (Afreh-­ Nuamah and Hemeng, 1995; Schill et al., 1996) and in south-western Nigeria it has been recognized as the most important biotic constraint to plantain production (Speijer et al., 2001). In the Cavendish plantations in South Africa, P. coffeae is reported as endemic and in the Limpopo Province responsible for up to 60% losses (De Villiers et al., 2007). Pratylenchus coffeae is also now being observed more often in East Africa (Coyne, 2009) where it seems to be gaining prominence. In Zanzibar, it has been found in 68% of banana fields (Rajab et al., 1999). Its localized distribution and rising status in other African countries indicate that it may have only recently been introduced (Bridge et al., 1997; Coyne, 2009). However, caution is needed in the interpretation of surveys, as some nematodes identified as P.  coffeae in Ghana (Brentu et al., 2004) have since been identified by molecular characterization as P. speijeri, a morphologically similar but separate species (De Luca et al., 2012). Pratylenchus goodeyi is not so widely distributed as P. coffeae and seems adapted to cooler ­climates. Banana, abacá and enset are economically important hosts (O’Bannon, 1975; Peregrine and Bridge, 1992; Tessera and Quimio, 1994). The nematode is found predominantly in Africa and its general absence from commercial plantations of Cavendish cultivars in lowland areas and its presence only on smallholder banana crops indicate that it may be indigenous to this continent (Price and Bridge, 1995). It can occur in extremely high densities, such as on banana in Tanzania (Speijer and Bosch, 1996) and enset in Ethiopia (Peregrine and Bridge, 1992). In Cameroon, P. goodeyi is the most serious nematode pathogen at elevations above 700 m and in the East African highlands it replaces R. similis as the dominant species above 1400 m altitude (Speijer and Fogain, 1999). The nematode has also been associated with banana losses

in coastal Kenya (Seshu Reddy et al., 2007), where prevailing temperatures tend to be higher than is optimal for this species. In Rwanda, no correlation could be established between incidence of P. goodeyi and cooking banana losses (Gaidashova et al., 2009). Pratylenchus goodeyi is regarded as a major pest in commercial Cavendish plantations in the Canary Islands (De Guiran and Vilardebo, 1962) and has also been recorded in Madeira, Egypt and Crete (Gowen et al., 2005). In Australia, P. goodeyi was as pathogenic as R. similis in commercial Cavendish plantations in the subtropics with the former nematode species being more prevalent in the cooler months and the latter in the warmer months (Pattison et al., 2002). It is also common on banana in Hainan Province, China (Zhang et al., 2015). Large numbers of P. coffeae were extracted from abacá roots in Ecuador. Severe damage was observed on the plants in the form of root necrosis, yellowing of leaves and stunted growth (Bridge, 1976). In Ethiopia, P. goodeyi is the predominant nematode species found on enset across agroecologica1 zones (Bogale et al., 2004; Addis et al., 2006). From a total of 71 enset cultivars sampled, all were infected to varying degrees with the nematode. Disease cycle and epidemiology The optimum temperature for invasion and ­development of P. coffeae is 25–30°C, which is the same as for R. similis and appears similar for P.  speijeri. With P. goodeyi, it is nearer 20°C. The life cycle of P. coffeae is completed in about 4 weeks under optimum conditions. The level of damage caused to banana by P.  coffeae varies geographically (Dubois and Coyne, 2011). In some areas of Uganda, very high densities of the nematode have been observed on old banana stands, which still remain productive, and yet in Tanzania lower densities are associated with a high incidence of plant toppling (Bridge et al., 1997). The variability in reported yield reductions caused by P. coffeae has been attributed to the existence of different pathotypes or strains of the nematode and to misidentification of the pathogen (Duncan et al., 1999; De Luca et al., 2012). The question of



Nematode Pathogens

whether there are biotypes of Pratylenchus spp. with different host preferences is under continued investigation. Population densities are usually expressed on the basis of 100 g of fresh roots and results vary according to the extraction technique used. In plantations where damage is obvious, either as uprooting or on visual inspection of roots, densities greater than 10,000 nematodes/100 g roots may be common. In Cameroon, at altitudes over 900 m, the population densities of P. goodeyi on plantations averaged 15,000 with a maximum of 56,000 nematodes for every 100 g of roots (Bridge et  al., 1995). In Uganda, average densities on East ­African highland banana cultivars in the AAA, Lujugira–Mutika subgroup were 25,000 nematodes/100 g roots at ten farms at altitudes over 1600 m, but only 680 nematodes/100 g roots on a similar number of fields at altitudes of 500 m or lower (Kashaija et al., 1994). However, banana plants on these lower-altitude farms were suffering no less, because roots were supporting densities of 32,000 Helicotylenchus multicinctus (spiral nematode) and 6500 R. si­ milis (burrowing nematode) per 100 g roots. These two species were not present at the higher elevation. This illustrates the complexity of determining the relative importance of nematodes in mixed populations and in different ­environments. Host reaction Pratylenchus coffeae is a significant pest on cultivars in the Cavendish, Plantain and Pisang Awak subgroups. In Africa, P. goodeyi is an important pest on cultivars in the Plantain and ­Lujugira–Mutika subgroups (Bridge et al., 1997; Coyne et al., 2005). The occurrence of Pratylen­ chus speijeri is associated with severe damage to ‘Apantu-pa’ (AAB, Plantain subgroup) (Brentu et al., 2004). Resistance (or decreased susceptibility) to Pratylenchus spp. has been demonstrated in glasshouse experiments. In one study, 12 diploids, including the ‘Long Tavoy 1’, ‘Long Tavoy 2’ and ‘Calcutta 4’ accessions of Musa acuminata ssp. burmannica (AAw), exhibited partial resistance to P.  coffeae (Quénéhervé et al., 2009). ‘Yangambi Km 5’ (AAA, Ibota subgroup), ‘Paka’ (AA),

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Kunnan’ (AB) and ‘Pisang Ceylan’ (AAB, ‘­ Mysore subgroup) have been shown to have ­ ­significant resistance to P. coffeae (Collingborn and Gowen, 1997). ‘Yangambi Km 5’ has also been shown to have some resistance to P. goodeyi in pots (Pinochet et al., 1998) and in the field in Cameroon (Fogain and Gowen, 1998). Potted plants of ‘Tjau Lagada’ (AA), ‘Pisang Bungai’ (AA) and ‘Pisang Mas’ (AA) were found to have lower infection levels than ‘Grande Naine’ (AAA, Cavendish subgroup), the susceptible check, ­after inoculation (Moens et al., 2005). Control Cultural, biological and chemical methods of control that are effective against R. similis are in general effective against Pratylenchus spp. These include planting nematode-free suckers or plantlets derived from tissue culture in land free of nematodes and paring suckers to remove roots and infested areas of the corm followed by a hot-water treatment for 20 min at 53–55°C or immersion in boiling water for 30 s. Nematicides that are currently used for the control of R. similis in commercial banana plantations are equally effective on Pratylenchus spp. Similarly, biologically based management options for R. similis are likely to be suitable for Pratylenchus spp. but have in general been less studied than for R. similis. Host-range information is important for developing a management strategy based on healthy planting material. Several crops and common weeds will support reproduction of P. coffeae (Gowen et al., 2005) and a few alternative hosts of P. goodeyi have been discovered in East Africa (Mbwana, 1992). In glasshouse experiments, early inoculation with arbuscular mycorrhizal fungi appeared to increase the tolerance of ‘Grand Naine’ (AAA) to P. goodeyi by reducing the number of lesions on roots and enhancing plant nutrition (Jaizme-Vega and Pinochet, 1997). In the long term, conventional banana breeding, perhaps coupled with genetic transformation, should contribute towards a partial management of Pratylenchus spp. Sources of resistance are currently being identified in Honduras, Cameroon, Uganda and the Canary Islands (Tenkouano and Swennen, 2004; Lorenzen et al., 2010). Screening of leaf spot-resistant ­banana

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hybrids with AAA genomes in Martinique showed four, designated ‘FB918’, ‘FB919’, ’FB920’ and ‘FB924’, to have resistance to P. coffeae (Quénéhervé et al., 2008). As reported earlier, three of these four hybrids were also resistant to R. similis. However, resistance against R. similis will not necessarily confer resistance against P. coffeae. It is important to establish the specific Pratylenchus species involved in causing damage to banana in order to be able to screen accurately for resistance.

Spiral Nematodes Introduction There are several species known collectively as spiral nematodes. Their name comes from the characteristic manner in which they coil when relaxed or heat-killed. Only one is a significant pathogen on banana and, unlike other spiral nematodes, feeds on the crop as an endoparasite. It occurs almost wherever banana is grown and almost exclusively in combination with other important nematode species (McSorley and ­Parrado, 1986). Because of this, opinions have differed as to its importance as a pathogen of banana However, evidence for its role in causing an important disease, often at the edge of the climatic range for banana, is gradually accumulating. In Israel, the spiral nematode has been shown to cause serious damage (Minz et al., 1960) and in Ghana it is regarded as being of equal significance to P. coffeae (Schill et al., 1996). Spiral nematodes have also been found on abacá (Bridge, 1976) and enset (Addis et al., 2006).

Symptoms Like other migratory endoparasites, the spiral nematode feeds on the cell contents in the root cortex, causing necrotic lesions. However, unlike R. similis and Pratylenchus spp., feeding is often restricted to the outer parenchymal cells of the cortex (Zuckerman and Strich-Harari, 1963; Blake, 1966; Mateille, 1994a). In roots where the spiral nematode is the only parasite, lesions are often superficial (Plate 7.10). However, in

Plate 7.10.  Superficial lesions on banana roots caused by Helicotylenchus multicinctus, a spiral nematode (photo: S.R. Gowen, UR).

s­ evere infections, necrosis may extend to the stele causing root death. Therefore, the spiral nematode can also cause toppling of infected plants.

Causal agent The spiral nematode species important on banana is Helicotylenchus multicinctus, although others may be present, especially H. dihystera. Helicotylenchus multicinctus occurs frequently in roots that are infected with R. similis, Pratylen­ chus spp. or Meloidogyne spp. In extracts from root samples, H. multicinctus can be readily distinguished from these other genera by comparison of the lengths of the stylet (which are longer) and by the shape of the body when killed. Dead specimens are curved in the form of a letter C, whereas other spiral nematodes die in a coiled position. Those of Radopholus similis and Praty­ lenchus spp. are generally straight when at rest. Juveniles of Meloidogyne spp. are straight when at rest and smaller in size than H. multicinctus. Helicotylenchus dihystera, H. multicinctus and another unidentified species have been found on enset, though not frequently (1–5% ­incidence).

Disease cycle and epidemiology Unlike other spiral nematodes, which are ectoparasites, H. multicinctus is entirely endoparasitic. Like the burrowing and root-lesion nematodes, H. multicinctus is likely to have been distributed widely on infected planting material. All stages



Nematode Pathogens

(juveniles and adults) are infective and can be found in roots and adjacent soil. When occurring in mixed populations, numbers of H. multicinctus may be greater than those of R. similis (Kashaija et al., 1994). On commercial plantations of ‘Robusta’ (AAA, Cavendish subgroup) in St Lucia, densities reached 24,000 spiral nematodes/100 g fresh roots, three times greater than that of R. similis (Gowen, 1977b). In Venezuela, where H. multicinctus was found coexisting with Meloidogyne incognita on the roots of a cultivar in the Cavendish subgroup, densities of 35,000 spiral nematodes/ 100 g roots were reported (Crozzoli et al., 1995). Population densities in Côte d’Ivoire averaged up to 53,000/100 g root (Adiko and N’Guessan, 2001) and up to 40,000/100 g root in Burkina Faso (Sawadogo et al., 2001). Host reaction There are few reports on differential susceptibility to this nematode in Musa because it has in the past been considered a less serious pathogen than other nematodes. This is an omission that is beginning to be corrected. However, techniques for mass-culturing H. multicinctus have also proved an obstacle as they are not as well established as for R. similis and P. coffeae and which, to a certain extent, constrains critical experimental work. Ssango et al. (2004) were able to separate the effects of different nematode species on cultivars of the AAA Lujugira–Mutika subgroup in Uganda and demonstrate that H. multicinctus caused damage. Evidence from field trials on ‘­Agbagba’ (AAB, Plantain subgroup) in Nigeria also indicated that H. multicinctus was responsible for much production damage when in mixed populations with other nematodes (Coyne et al., 2013). From surveys in plantain fields in the Dem­ ocratic Republic of the Congo, root necrosis was positively and significantly correlated to population densities of H. multicinctus (Kamira et al., 2013). ­However, compared with non-inoculated plants, H.  multicinctus caused no reduction in bunch weight of ‘Grande Naine’ (AAA, Cavendish subgroup) in microplots in Costa Rica, whereas losses were caused by R. similis, Meloidogyne incognita and P. coffeae (Moens et al., 2006). In West Africa, H. multicinctus is highly prevalent on cultivars in  the AAB Plantain subgroup and regularly

443

­ ssociated with necrotic root systems and topa pled plants (Caveness and Badra, 1980; Adiko and N’Guessan, 2001; Speijer et al., 2001; Brentu et al., 2004). In pot experiments, ‘Poyo’ (AAA, Cavedish subgroup) and ‘Gros Michel’ (AAA) were both found to be equally susceptible to H. multicinctus (Mateille, 1994a). In Costa Rica, most of the 31 Musa cultivars assessed by Moens et al. (2005) supported similar or higher densities of H. mul­ ticinctus as the susceptible check ‘Grande Naine’ (AAA, Cavendish subgroup), while ‘Yangambi Km 5’ (AAA, Ibota subgroup) supported low densities. ‘Tjau Lagada’ (AA) and ‘Pisang Bungai’ (AA) appeared resistant, but this finding needs further confirmation. Of 19 bred hybrids (primary tetraploids and improved diploids) screened in India against H. multicinctus in inoculated pot trials and field studies, ‘H 531’ (‘Mysore’ (AAB) × ‘Pisang Lilin’ (AA)) exhibited resistance. ‘H-02-34’, ‘H-03-05’, ‘H-03-13’, ‘H-04-12’, ‘H03-17’, ‘H-04-24 , NPH-02-01 and H 510 were classed as tolerant (Das et al., 2014a).

Control Chemical, biological and cultural control methods used in the management of R. similis will also mostly apply to H. multicinctus.

Root-knot Nematodes Introduction Root-knot nematodes have been found in association with banana in all producing areas. They have also been identified as infecting abacá (Ocfemia and Calinisan, 1928) and enset (O’Bannon, 1975; Bogale et al., 2004; Addis et al., 2006). In general, root-knot nematodes have not been considered important banana pathogens in the past. However, as with the spiral nematode, their importance may be underestimated due to a limited understanding of their role in disease as they regularly occur in combination with other damaging nematode species. On cultivars in the AAA Cavendish subgroup, the burrowing nematode is usually more successful and tends to dominate in situations where both types of

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nematode are found. Root-knot nematodes are more likely to cause problems in areas where Cavendish cultivars have not been introduced or where the climate is too cold for R. similis. Early reports, such as those from Honduras (Pinochet, 1977), Colombia (Zuniga et al., 1979) and the French Antilles (Kermarrec and Scotto la Massese, 1972), did not place any great significance on root-knot nematodes as important pathogens of banana in the Latin American– Caribbean region. However, they are now viewed as having greater importance (Cofcewicz et al., 2005), such as in Brazil, where root-knot nematodes occurred in 79% of root samples (Lima et al., 2013). In East Africa, root-knot nematodes do not appear to be of great significance (Nsemwa, 1991; Kashaija et al., 1994; Speijer and Kajumba, 1996; Speijer et al., 1999). However, they are recorded as a problem in the northern Cavendish-­ growing districts of South Africa, where they have been implicated, in part, in a condition known as false Panama disorder (Deacon et al., 1985; de Beer et al., 2001). Root-knot nematodes and Fusarium oxy­ sporum (but not f. sp. cubense) are associated with this disorder. Treatment with nematicides can prevent the appearance of symptoms (A. Severn-Ellis, Australia, 1999, personal communication with D. De Waele). They have also been regularly recovered during field surveys in Central Africa. Root-knot nematodes were the second most frequently occurring nematodes in the Democratic Republic of the Congo after H. mul­ ticinctus, being present in 48% of fields in the lowlands of Bas Congo and 61% of fields in the highlands in South Kivu (Kamira et al., 2013). In West Africa they can also be common, occurring in association with H. multicinctus in 90% of fields or more (Caveness and Badra, 1980; Adiko and N’Guessen, 2001; Sawadogo et al., 2001). In North Africa, root-knot nematodes have been recognized as a problem and are believed likely to contribute significantly to production losses (Gowen et al., 2005). In the Philippines, root-knot nematodes are also found on Cavendish cultivars. Large and widespread populations have been detected in commercial growing areas around Davao in Mindanao. The average population density in 82% of plantations examined was 3539 nematodes/ 100  g fresh roots. Large root-knot nematode

densities were also found on the roots of all local cultivars sampled (Davide et al., 1992). In Southeast Asia, root-knot nematodes are widely distributed on local diploid and triploid dessert cultivars, and also on cooking-banana cultivars. In West Malaysia, they were widespread in a commercial Cavendish plantation, presenting extensive root galls and average densities of 2300 individuals/200 ml soil (Razak, 1994). They were also the most predominant species recovered from banana roots (Rahman et al., 2014). Root-knot nematodes are commonly found on local banana cultivars in Thailand (Prachasaisoradej et al., 1994), Malaysia (Razak, 1994) and Indonesia (Hadisoeganda, 1994), where they have largely been regarded as being of minor importance. In North Vietnam, rootknot nematodes along with P. coffeae are the two major nematode species associated with banana (Van den Bergh et al., 2006). Root-knot nematodes are very common throughout the banana-­ growing regions of Australia, but were not shown to cause yield loss (Stanton, 1994). Root-knot nematodes have been described as common and abundant on banana in Mediterranean countries, such as Crete (Vovlas et al., 1994) and Lebanon (Sikora and Schlosser, 1973). Studies on interactions between banana and root-knot nematodes to determine production and yield losses are relatively few compared with other nematode species. In field experiments in the Philippines, Davide and Marasigan (1985) reported a yield loss of 26.4% after ‘­Giant Cavendish’ (AAA, Cavendish subgroup) was inoculated with 1000 juveniles per plant. A 45.4% yield loss was caused by inoculations with 10,000 juveniles and a 57.1% yield loss by 20,000 juveniles. In Costa Rica, bunch weights of ‘Grande Naine’ (AAA, Cavendish subgroup) were reduced by 32% after inoculations of 1000 root-knot nematodes per plant. This loss was greater than that caused by the same numbers of burowing, root-lesion and spiral nematodes (Moens et al., 2006). In North Vietnam, field studies showed that ‘Chuõí Ngu Tien’ (AA) and ‘Grande Naine’ (AAA, Cavendish subgroup) inoculated with 8700 root-knot nematodes per plant suffered yield reductions of 23% and 19%, respectively (Van den Bergh et al., 2006). In a field study in Nigeria using ‘Agbagba’ (AAB, Plantain subgroup), bunch weights in the plant crop were reduced by 50% following an inoculation of 2000 root-knot



Nematode Pathogens

445

Plate 7.11.  Swollen and necrotic banana roots caused by root-knot nematodes (photo: D. Coyne, IITA).

nematodes per plant and by a similar amount in the following two crop cycles (Coyne et al., 2013).

Symptoms On banana, galls and swellings on primary and secondary roots are the most obvious symptoms of root-knot nematode infection (Plate 7.11). Sometimes, the root tips are invaded and there is little or no gall formation, but growth ceases and new roots proliferate just above the infected tissues. Infected plants may have a much lower number of secondary and tertiary roots and root hairs (Claudio and Davide, 1967). Dissection of galls reveals the typical swollen females in various stages of development (Plate 7.12). At maturity, the females are saccate. Eggs are laid within a gelatinous matrix to form an external egg sac or egg mass. In thick, fleshy primary roots the egg masses may be contained within the root, resulting in swollen roots. On banana roots grown under in vitro conditions, protruding egg masses were observed 28 days after inoculation (Coosemans et al., 1994). Different root-knot nematode species may occur in the same gall (­Pinochet, 1977; Cofcewicz et al., 2005). They may also colonize the outer layers of the corm up to 7 cm deep (Quénéhervé and Cadet, 1985a). Above-ground symptoms caused on banana by root-knot nematode in Pakistan included yellowing and narrowing of leaves, stunting, reduced plant growth and less fruit production (Jabeen et al., 1996). Stunted growth has also been attributed to

Plate 7.12.  A swollen banana root caused by root-knot nematodes in longitudinal section. White females are clearly present at the centre of the dark-coloured areas (photo: D. Coyne, IITA).

root-knot nematodes in India (Sudha and Prabhoo, 1983) and Taiwan (Lin and Tsay, 1985). On abacá, galls on roots have been reported to be 3–10 mm in diameter and may run together to form an irregular club-shaped body up to 5 cm long and over 1 cm in thickness. Infected roots become brown and then almost black in colour. The surface of the galls crack with age and become rough to the touch. Leaves turn pale green or yellowish. The youngest leaf is generally the worst affected. Later, leaves become narrower and shorter. Plants appear stunted and leaves tend to bunch. Galls on the primary and secondary roots of enset are associated with root-knot nematodes. Infected plants become stunted and have yellow leaves, which may wilt in the dry season. Young seedlings can be seriously affected.

Causal agents Root-knot nematodes (Meloidogyne spp.) are ubiquitous pathogens with a global distribution

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and infect a wide range of host plants (Perry et al., 2009). Identification to species level is often not undertaken because of a lack of diagnostic expertise or facilities. However, this is changing and the list of species found infecting Musa is ­expanding. The species most commonly recorded on banana are Meloidogyne incognita and M. javanica. Meloidogyne arenaria and other Meloidogyne species are also variously reported. In the Caribbean region, M. arenaria was the most frequently occurring species on banana at 62% of sampled sites in Guadeloupe, French Guiana and Martinique. In the same survey, M. cruciani and M. hispanica were infrequently found (Cofcewicz et al., 2005). Meloi­ dogyne graminicola was recorded causing damage on ‘Tianbao’ (AAA, Cavendish subgroup) in Fujian Province, China (Zhou et al., 2015), while M. enterolobii was additionally recorded causing galling damage on ‘Baxi’ (AAA, Cavendish subgroup) in the same area (Zhou et al., 2016). The Meloidogyne species found on abacá have not been identified, but are reported in combination with R. similis and Helicotylenchus spp. (Bridge, 1976). Root-knot nematodes so far identified from enset roots include M. incognita, M. javanica and M. ethiopica (Peregrine and Bridge, 1992; Tessera and Quimio, 1994; Mandefro and Dagne, 2000).

Disease cycle and epidemiology The life cycle of Meloidogyne spp. on banana is similar to its life cycle on other hosts. The endodermis is penetrated by vermiform infective ­juveniles, which enter the stele and induce the vascular parenchyma or differentiating vascular cells in the central part of the stele to form multi-nucleate giant cells. The formation of these giant cells disturbs or blocks the surrounding xylem vessels (Dos Santos and Sharma, 1978; Sudha and Prabhoo, 1983; Vovlas and Ekanayake, 1985; Jabeen et al., 1996). The nematode becomes sedentary and feeds from these giant cells as it develops into a mature female and reproduces. They have spherical bodies with slender necks. Multiple life cycles can be completed within the same root, depending on the longevity of the root and the severity of the necrosis. The males are vermiform and generally rare.

Root-knot nematodes may require much more time to become established in banana roots than root-lesion nematodes. In Cuba, Meloidogyne spp. needed 24–30 months to establish themselves on ‘Dwarf Cavendish’ (AAA, Cavendish subgroup) (Fernandez and Ortega, 1982). Root-knot nematodes are influenced by rainfall and soil conditions, such as temperature, texture and pH. After establishment, soil moisture and temperature are mainly responsible for fluctuations in populations (McSorley and Parrado, 1981; Fernandez and Ortega, 1982; Mani and Al Hinai, 1996; Youssef and Aboul-Eid, 1996). Regardless of inoculum levels, M. incog­ nita usually reach highest densities in the soil during the rainy season and then decline to reach lowest levels during the dry season. The climate also affects the host. During the dry season, not enough new roots are available for nematodes to infect, resulting in low nematode densities. In Egypt, the highest densities of M. in­ cognita on banana were also positively correlated with the highest soil temperatures (26–30°C) observed at the experimental site (Youssef and Aboul-Eid, 1996). In the Philippines, Davide (1980) reported that the highest population densities of M. incognita were observed in sandy loam soils and at pH 5–5.6. Meloidogyne spp. and R. similis can jointly infect banana. However, root-knot nematodes are usually reduced or completely replaced by R. simi­ lis, as the latter species destroys the roots, which provide the feeding sites for the root-knot nematodes (Santor and Davide, 1992). In West and Central Africa, it is quite common to find Meloid­ ogyne spp. and H. multicinctus in combination, ­often with lower densities of other nematodes. In banana roots, Meloidogyne spp. often ­occur together with fungi capable of colonizing weakened or wounded tissue. In Yemen, Sikora (1980) observed higher levels of root rot in banana plantations where M. incognita and rootrot fungi (Fusarium and Rhizoctonia spp.) were present together in the soil. Synergistic effects of M. incognita and Fusarium oxysporum f. sp. cubense, the cause of Fusarium wilt, on roots of ‘Rasthali’ (AAB, Silk subgroup) have also been reported (Jonathan and Rajendran, 1998). Root-knot nematodes are often dispersed in run-off water and can also be spread with irrigation water and contaminated planting material.



Nematode Pathogens

447

Host reaction

Control

In general, the widely grown banana cultivars tend to be susceptible to root-knot nematode. Often, inconclusive results have been obtained when banana genotypes have been screened for resistance to Meloidogyne spp. on a large scale. In Brazil and India, of numerous Musa genotypes screened against M. incognita and M. javanica, none were found resistant or even moderately resistant (Zem and Lordello, 1981; Patel et al., 1996). In the Philippines, Davide and Marasigan (1985) screened 90 different Musa genotypes for reaction to M. incognita. They reported that the response of cultivars varied considerably, ranging from mild to severe root-gall formation. ‘Viente Cohol’ (AA), ‘Dakdakan’ (AA, syn. ‘Viente Cohol’), ‘Pogpogon’ (AA), ‘Alaswe’ (AAA), ‘Inambak’ (AAA), ‘Pastilan’ (AAA), ‘Sinker’ (AAA), ‘Mai’a Maole’ (AAB, Mai’a Maoli–Popoulu subgroup) and ‘Pa-a Dalaga’ (ABB) showed some resistance to M. incognita with generally only a few nematodes infecting the roots, which had trace to slight gall formation. In Malaysia, the popular cultivars ‘Pisang Mas’ (AA, Sucrier subgroup), ‘Pisang Embun’ (AAA, Gros Michel subgroup), ‘Pisang Nangka’ (AAA), ‘Pisang Berangan’ (AAA, Lakatan subgroup), ‘Pisang Rastali (AAB, Silk subgroup) and ‘Pisang Tandok’ (AAB, Plantain subgroup) were susceptible to root-knot (Razak, 1994). Using 26 Vietnamese banana accessions from the AA, AAA, AAB, ABB and AB genome groups and some wild accessions, no source of resistance was found against a mixture of Meloidogyne spp. (Van den Bergh et al., 2002). Of 31 Musa accessions screened for nematode resistance in Costa Rica, no resistance was observed against M. incognita (Moens et al., 2005). Nor was any source of resistance to M. incognita or M. arenaria found from 55 Musa accessions screened for resistance in Martinique (Quénéhervé et al., 2009). In India, Das et al. (2014b) screened 19 bred hybrids in inoculated pot trials and in the field against M. in­ cognita. ‘H 531’ was found to be resistant and ‘H-02-34’, ‘H-03-05’, ‘H-03-13’, ‘H-04-12’, ‘H-04-24’ and ‘NPH-02-01’ were classified as tolerant. To date, no assessment appears to have been made on abacá and enset.

Chemical, biological and cultural options utilized in the management of the burrowing nematode will also mostly apply to Meloidogyne spp. Meloidogyne spp. can be disseminated with infected planting material. Risks can be minimized by using healthy planting material derived from tissue culture or by removing/peeling the outer tissues of the corm or sucker followed by a hot-water, boiling-water or nematicide treatment before planting (Haddad et al., 1973). In Yemen, heavy banana losses, associated with severe infection by M. incognita, were reduced through the use of Meloidogyne-free propagative stocks (Ibrahim, 1985). Root-knot nematodes have a wide host range and associations with other plant hosts, including numerous weeds, are far more numerous than for the other banana nematode pests. Special attention should be given to the maintenance of weed-free fallow and the selection of cover crops in rotation systems and intercrops. In India, intercropping with Coriandrum sa­ tivum, Sesamum indicum, Crotalaria juncea, Tagetes erecta and Acorus calamus have significantly ­reduced M. incognita on ‘Robusta’ (AAA, Cavendish subgroup) in field trials (Charles, 1995). The same effect on Meloidogyne spp. was obtained in crop rotation trials with Pangola grass, maize and sugarcane in Cuba (Stoyanov, 1971) and with Tagetes patula in South Africa (Milne and Keetch, 1976). Rotation with paddy rice can also drastically reduce root-knot nematode densities, though this was a result of flooding (Sivakumar and Marimuthu, 1986). Fallowing to eradicate root-knot nematodes may, however, be ineffective, as Meloidogyne spp. have been shown in Cuba to persist in soil in the absence of banana for up to 29 months (Stoyanov, 1971). Numerous field experiments have shown the effectiveness of nematicide in the control of root-knot nematodes. Dipping banana corms in a solution of nematicide for 10 min before planting may protect the plants for a few months against nematode infection. Immersion of peeled corms in a 1% solution of sodium hypochlorite (NaOCl) for 5 or 10 min also controlled Meloido­ gyne spp. and was considered by Lordello et al. (1994) as an effective, low-cost and non-toxic pre-planting treatment. By knowing the seasonal fluctuation in nematode population densities,

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an effective nematicide application strategy can be developed (Badra and Caveness, 1983). Control is most effective when treatments are timed to coincide with the build-up of nematode populations that usually occurs at the onset of the rainy season. In Puerto Rico, oxamyl applied to leaf axils of ‘Giant Cavendish’ (AAA, Cavendish subgroup) four times at 30-day intervals during the growing season effectively controlled M. incognita (Robalino et al., 1983). Hoan and Davide (1979) reported that root extracts of 11 plant species showed nematicidal effects when tested against M. incognita in the Philippines. Root extracts from African marigold (Tagetes erecta), ipil-ipil (Leucaena leucocephala), Bermuda grass (Cynodon dactylon) and makahiya (Mimosa pudica) prevented eggs from hatching. The performance of these root extracts was comparable to that of commercially produced chemical nematicides. Results of another study revealed that leaf extracts from kaatoan bangkal (Antho­ cephalus chinensis) and water lily (Eichornia cras­ sipes) and extracts of garlic (Allium sativa) and onion (Allium cepa) bulbs were also effective against M. incognita (Guzman and Davide, 1992). Characterization of the active nematicidal principle showed a phenolic aldehyde from kaatoan bangkal, a carboxylic acid from water lily and a ketone from onion. Culture extracts of 17 species of microorganisms have been evaluated under laboratory and greenhouse conditions in the Philippines for nematicidal activity against M. incognita infesting ‘Giant Cavendish’ (Molina and Davide, 1986). Purified extracts of Penicillium oxalicum, P. anatoli­ cum and Aspergillus niger showed high nematicidal activity. Purpureocillium lilacinus and Penicillium oxalicum have been very successful in controlling Meloidogyne spp. and other nematodes on banana (Davide, 1994). Arbuscular mycorrhizal fungi are also being investigated as biological control agents. Inoculation of microprapagated ‘Grande Naine’ (AAA, Cavendish subgroup) with two isolates of Glomus mosseae suppressed gall formation and build-up of M. incognita in roots under greenhouse conditions. The presence of nematodes had no effect on the colonization of roots by these fungi (Jaizme-Vega et al., 1997). Inoculation of the same banana cultivar with Glomus intraradices did not affect the build-up of M. incognita in the roots, but increased plant

growth by enhancing plant nutrition (Pinochet et al., 1997). Following the commercial production of Pasteuria penetrans for control of Meloidogyne spp. (Smith et al., 2004), the prospects of improving the management of root-knot nematodes through biological control moves closer.

Other Nematodes In addition to the burrowing nematode, root-­ lesion nematodes, spiral nematodes and root-knot nematodes, numerous other nematode species have been recovered from banana. Most are of little consequence and occur in relatively low numbers. Some, however, although not generally viewed as key pathogens, have a localized significance and require mention. Rotylenchulus reniformis, or reniform nematode, has been found in association with banana in all producing areas throughout the world. Documented reports come from South America (Zuniga et al., 1979; Crozzoli et al., 1993), Hawaii (Wang and Hooks, 2009), the Caribbean (Oramas and Roman, 1982), Africa (Fargette and Quénéhervé, 1988; Adiko and N’Guessen, 2001; Kamira et al., 2013; Daneel et al., 2015), Asia (Chau et al., 1997; Rahman et al., 2014) and the Mediterranean (Aboul-Eid and Ameen, 1991). In St Lucia, densities of up to 2500 juvenile and infective immature female nematodes were found in 100 cm3 samples of soil taken from around the fine secondary roots in which mature adult females were permanently lodged (Gowen, 1977b). In West Malaysia, R. reniformis were the most prevalent nematode species recovered from the banana rhizosphere (Rahman et al., 2014). In Guangdong Province in China, R. reniformis was present in 61% of fields and identified as a major nematode parasite of banana in the area (Shaomei et al., 2006). However, although R. reniformis is believed to cause damage to the root system (Edmunds, 1968), little quantitative data on the effect of this nematode species on growth and yield of banana have been reported. Rotylenchulus reniformis penetrates the cortex of banana roots perpendicularly to the stele and establishes a permanent feeding site in the endodermis. Nematode feeding induces the ­fusion of endodermal, pericycle and vascular



Nematode Pathogens

­ arenchymal cells to form a syncytium, with p hypertrophied nuclei and prominent nucleoli (Vovlas and Ekanayake, 1985). These permanent feeding sites are generally located on the secondary roots (Ayala, 1962; Edmunds, 1968). Rotylenchulus reniformis is usually found in association with other pathogenic nematode species. Most nematicides effective against rootknot nematodes, including oxamyl applied to leaves (Gowen, 1977a; Robalino et al., 1983), were also effective against R. reniformis. Information on either cultural or biological control is limited. In India, intercropping with Corian­ drum sativum, Sesamum indicum, Crotalaria ­juncea, Tagetes erecta and Acorus calamus significantly reduced R. reniformis densities on ‘­Robusta’ (AAA, Cavendish subgroup) in field trials (Charles, 1995). The nematode has also been reported to infect the roots of abacá (­Davide, 1972). Hoplolaimus pararobustus, which can be found in relatively high densities on banana (1000–18,000 nematodes/100 g roots) (Guerout, 1974; Hunt, 1977; Price, 1994b), appears to occur only in the subepidermal cortex (Mateille, 1994a). The potential of this nematode to cause damage has been questioned, but, if large populations of this relatively large migratory endoparasite are present, it was believed that they must be having some effect on plant development (Price, 1994a). One of the few studies to have assessed the damage this species causes was conducted on ‘Agbagba’ (AAB, Plantain subgroup) in the field in Nigeria. Following inoculation with 2000 H. pararobustus nematodes/plant, bunch weights were reduced by 53% in the plant crop cycle with significantly more stems snapping than in the controls as a result of reduced water uptake (Coyne et al., 2013). Heterodera oryzicola occurs on banana in southern India and its incidence is probably related to the cropping system, where banana is grown in rotation with paddy rice. Pathogenicity studies suggest that this nematode could cause yield loss (Charles and Venkitesan, 1993). Pratylenchus spp., such as Pratylenchus minutus, and Paratrichodorus spp., such as Par­ atrichodorus minor, have been mentioned in some studies (Daneel et al., 2015), but remain of

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limited and possibly local importance, with little information on their pathogenicity on Musa. Nematode black leaf streak disease of enset was first recorded in 1991 in Ethiopia (Tessera and Quimio, 1994), where it occurs in most growing areas. The disease is caused by Aphelenchoides ensete (originally reported as an Ektaphelenchoides species) (Swart et al., 2000) and can severely damage enset suckers and seedlings. The most characteristic symptom is small black streaks on leaves (Plate 7.13). Streaks sometimes coalesce to form long necrotic stripes. Severe streaking can cause the premature death of leaves (Plate 7.14). The nematode lives and multiplies in leaf tissue (Plate 7.15) and spreads to neighbouring healthy leaves by rain splash or during watering operations. The nematode is carried to new farms on infected plants. Most enset clones seem susceptible. The early removal of infected leaves helps to control the disease and minimize the chance of spread. An Ektaphelenchoides sp. was also recovered from 30% of enset roots of surveyed fields (Addis et al., 2006). During the same survey, other plant-parasitic nematodes, recorded mostly from the root rhizosphere, included Scutellonema paralabiatum, Scutellonema sp., Rotylenchulus sp., Tylencholaimellus sp. and Tylenchorhynchus levit­ erminalis.

a Plate 7.13.  Symptoms of nematode black leaf streak disease on the leaf of a young enset seedling (photo: M. Tessera and A.J. Quimio, IAR).

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Plate 7.15.  Aphelenchoides ensete in leaf tissue of enset (photo: M. Tessera and A.J. Quimio, IAR).

Acknowledgements The authors gratefully acknowledge the contributions of the nematologists who wrote the ­disease descriptions for the first edition of this publication. Their text and images formed the starting point for this update.

Plate 7.14.  Leaf of an enset sucker with severe symptoms of nematode black leaf streak disease caused by Aphelenchoides ensete (photo: M. Tessera and A.J. Quimio, IAR).

• • • •

J.L. Sarah - Burrowing Nematode; S.R. Gowen - Root-lesion Nematodes and Spiral Nematode; D. De Waele - Root-knot Nematodes; D. De Waele, S. R. Gowen, M. Tessera; and A.J. Quimio – Other Nematodes.

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Sarah, J.L. (1989) Banana nematodes and their control in Africa. Nematropica 19, 199–216. Sarah, J.L. (1991) Estimation of nematode infestation in banana. Fruits 46, 643–646. Sarah, J.L., Lassoudière, A. and Guérout, R. (1983) La jachère nue et l’inondation du sol, deux méthodes intéressantes de lutte intégrée contre Radopholus similis dans les sols tourbeux de Côte d’Ivoire. Fruits 38, 35–42. Sarah, J.L., Kéhé, M., Beugnon, M. and Martin, P. (1988) Expérimentation avec l’aldicarbe pour lutter contre Radopholus similis COBB (Nematoda, Pratylenchidae) et Cosmopolites sordidus (GER-MAR) (Coleoptera, Curculionidae) en bananeraie. 2. Expérimentation réalisée en Côte d’Ivoire. Fruits 43, 475–484. Sarah, J.L., Blavignac, F., Sabatini, C. and Boisseau, M. (1992) Une méthode de laboratoire pour le criblage variétal des bananiers vis-à-vis des nématodes. Fruits 45, 35–42. Sarah, J.L., Sabatini, C. and Boisseau, M. (1993) Differences in pathogenicity to banana (Musa sp., cv. Poyo) among isolates of Radopholus similis from different production areas of the world. Nematropica 23, 75–79. Sawadogo, A., Thio, T., Konate, Y.A. and Kiemde, S. (2001) Nematode parasites of banana in western ­Burkina Faso. InfoMusa 10, 28–29. Schill, P., Gold, C.S. and Afreh-Nuamah, K. (1996) Assessment and characterization of constraints in plantain production in Ghana as an example for West Africa. In: Ortiz, R. and Akorada, M.O. (eds) Plantain and Banana: Production and Research in West and Central Africa. Proceedings of a Regional Workshop, Onne, Nigeria, 23–27 September, 1995. IITA, Ibadan, Nigeria, pp. 45–51. Seshu Reddy, K.V., Prasad, J.S., Speijer, P.R., Sikora, R.A. and Coyne, D.L. (2007) Distribution of plant-­ parasitic nematodes on Musa in Kenya. Infomusa 16, 18–23. Shaomei, L., Kelin, X., Chungsheng, L. and Gyan, F. (2006) A study to the reniform nematode disease of dwarf banana in Guangdong Province of China. Journal of South China Agricultural University 1987, 4. Sharmila, R., Kumar, S. and Ramakrishnan, S. (2012) Parasitizing ability of Pasteuria penetrans on phytonematodes. Journal of Biopesticides, 5 (Supplementary), 33–35. Shehabu, M., Addis, T., Mekonen, S., De Waele, D and Blomme, G. (2010) Nematode infection predisposes banana to soil-borne Xanthomonas campestris pv. musacearum transmission. Tree and Forestry ­Science and Biotechnology 4, 63–64. Sikora, R.A. (1980) Observations on Meloidogyne with emphasis on disease complexes and the effect of host plant on morphometrics. In: Proceedings of the 2nd Research Planning Conference on Rootknot Nematodes, Meloidogyne spp., Region VII. Athens, Greece, 1979. North Carolina State University, Raleigh, North Carolina, USA, pp. 93–104. Sikora, R.A. and Schlosser, E. (1973) Nematodes and fungi associated with root systems of bananas in a state of decline in Lebanon. Plant Disease Reporter 57, 615–618. Sikora, R.A., Pocasangre, L., Zum Felde, A., Niere, B., Vu, T.T. and Dababat, A.A. (2008) Mutualistic endophytic fungi and in-planta suppressiveness to plant parasitic nematodes. Biological Control 46, 15–23. Sivakumar, M. and Marimuthu, T. (1986) Population dynamics of phytonematodes associated with betelvine (Piper betle L.), banana and paddy rice with special reference to the crop. Indian Journal of Nematology 16, 277. Smith, K.S., Hewlett, T.E. and Griswold, S. (2004) Pasteuria for nematode control: development of a commercial production process. In: Proceedings of the 2004 Annual International Research Conference on Methyl Bromide Alternatives and Emissions Reductions, Orlando, 31 October – 2 November, 2004. Available at: http://www.mbao.org/2004/Proceedings04/mbrpro04.html Spejier, P.R. and Bosch, C.H. (1996) Susceptibility of Musa cultivars to nematodes in Kagera region, Tanzania. Fruits 51, 317–222. Speijer, P.R. and De Waele, D. (1997) Screening of Musa germplasm for resistance and tolerance to nematodes, INIBAP Technical Guidelines I. INIBAP, Montpellier, France, 48 pp. Speijer, P.R. and Fogain, R. (1999) Musa and Ensete nematode pest status in selected African countries. In: Frison, E., Gold, C., Karamura, E. and Sikora, R.A. (eds) Mobilizing IPM for Sustainable Banana Production in Africa. Proceedings of the International Workshop on Banana IPM. November 1998. Nelspruit, South Africa, INIBAP, Montpellier, France, pp. 99–108. Speijer, P.R. and Gold, C.S. (1996) Musa root health assessment: a technique for the evaluation of Musa germplasm resistance. In: Frison, E.A., Horry, J.P. and De Waele, D. (eds) New Frontiers in Resistance Breeding for Nematode, Fusarium and Sigatoka, Proceedings of the Workshop held in Kuala Lumpur, Malaysia, 2–5 October 1995. INIBAP, Montpellier, France, pp. 62–78. Speijer, P.R. and Kajumba, C. (1996) Yield loss from plant parasitic nematodes in East Africa Highland ­banana (Musa AAA). MusAfrica 10, 26.

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8 

Non-infectious Disorders of Banana E. Lahav, Y. Israeli, D.R. Jones and R.H. Stover

Introduction Infectious agents have not been shown to be the cause of the disorders described in this chapter. All the disorders are believed to result from the plant’s response to special growing conditions, often unfavourable. They are not major problems except in localized areas and then usually for short periods. Problems affecting the plant are described first, followed by disorders that specifically affect the fruit.

Plant Disorders Heart Leaf Unfurling Disorder of Plantain In Central America, the heart leaves of ‘French’ and ‘Horn’ plantain (AAB, Plantain subgroup) occasionally fail to unfurl normally. The first sign of unfurling abnormality is the bending over of the point of the unfurled heart leaf, which appears chlorotic. The first 30–60 cm at the tip then fails to unfurl or only one-half of the blade unfurls. The unfurled portion remains chlorotic and eventually rots. Sometimes only portions of the lamina unfurl, giving the leaves a torn and tattered appearance. Affected leaves appear shorter than normal, because the tip portion has rotted away. After the emergence of several abnormal heart leaves, normal leaf emergence resumes. 462

The cause of this disorder is unknown, but it is believed to be associated with an uneven growth rate as a result of adverse growing conditions, such as periods of drought followed by rain (Stover, 1972).

High Mat This is a common defect in banana growth, especially in plantations several years old (Stover, 1972). The upper portion of the rhizome grows out of the soil, exposing a considerable area of root-bearing tissue. Short roots emerge above ground when the weather is moist. The base of the rhizome is usually only 2.5–5 cm below the soil surface. Such plants are not as well anchored as plants with buried rhizomes and uprooting is more common. According to Wardlaw (1961), compact soils are a contributing factor, as well as pulling trash and soil away from the base of the plant in hoeing operations. Brouhns (1957) believed that lateral bud formation high on the rhizome was a contributing factor. Subra and Guillemot (1961) pointed out the importance of selecting axial suckers rather than lateral or terminal suckers for good anchorage. The axial sucker is the first sucker to emerge from the base of the rhizome on the side opposite to where the mother plant emerged. However, according to Charpentier (1966), all rhizomes normally tend to move upward out of the soil. Hasselo (1957)

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recommended earthing-up by making a mound of earth around the base of the banana mat in a circle of 45 cm radius so that the root-bearing region is covered to a depth of 10–15 cm. On volcanic soils in Cameroon, this reduced uprooting losses to an insignificant level. Charpentier (1966) stated that the tendency for rhizomes to come out of the ground resulted from a natural phototropism of the vegetative parts. This occurred when a rhizome was planted at a normal depth. When rhizomes were planted excessively deep, a second corm formed above the deeply placed corm and there was some foliar deformation. By earthing-up or darkening the base of the pseudostem, a second corm also formed higher up, but there was no foliar deformation. As earthing-up is used to stimulate corm formation for speeding up vegetative reproduction (Barker, 1959; Charpentier, 1966), it would seem unlikely to offer a more than temporary solution to the ‘high mat’ problem. In Central America, the high mat condition is common in some areas and is not related to compacted soils. Also, lateral buds that emerge from well below the soil surface can develop into high mats by the time the plant has flowered. Clones in the Cavendish subgroup (AAA), ‘Gros Michel’ (AAA) and cultivars in the Plantain ­subgroup (AAB) are affected. ‘High mat’ is found quite often in the first cycle of plants derived from tissue culture (Robinson and Galán-Saúco, 2010), but this is due to the early development of buds at the bottom of the developing corm (Plate 8.1) that push the growing corm upwards. This is not a disorder, but a normal development of tissue-cultured plants.

Plate 8.1.  Buds that develop on the bottom of the corm of tissue-cultured plants push the mat upwards (photo: Y. Israeli, JVBES).

Leaf-edge Chlorosis A yellowing of the leaf margins of Cavendish cultivars was described in Central America by Stover (1972). A similar condition occurs on the island of St Vincent in the Caribbean. Chlorosis begins at or near the edges of the leaf and spreads up the lamina towards the midrib. In severe cases, the yellowing can extend almost to the midrib (Plate 8.2). The affected tissue is sometimes invaded by weak pathogens and becomes necrotic. Leaves 3–6 are most commonly affected and the condition is worse in plants bearing bunches. The chlorotic areas of the leaf do not continue to expand in size as the leaf ages. The symptoms are also found in fertile plantations, but their effect on photosynthesis is negligible. The cause of this chlorosis is not known, but it may be associated with a transitional nutritional deficiency brought on by periods of stress. In St Vincent, although the symptoms resemble those caused by calcium deficiency, tests have shown that the affected leaf tissue has sufficient calcium. However, this does not completely rule out calcium deficiency as the cause. An analysis of leaves with symptoms reveals low levels of boron, manganese, phosphorus and sulfur. Stover (1972) reported less nitrogen, manganese and potassium than normal.

Roxana Disease This is a rare disorder reported in 1956 from an abandoned ‘Gros Michel’ planting near Roxana,

Plate 8.2.  Symptoms of leaf-edge chlorosis on ‘Robusta’ (AAA, Cavendish subgroup) in St Vincent, Windward Islands (photo: D.R. Jones, SVBGA).

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Limon Province, Costa Rica (Allen, 1957). About 10% of the plants were affected. Later in the year, another small outbreak occurred at Quebrada Honda about 80 km from Roxana. The problem was also identified in the Sula Valley in Honduras in 1957 (Waite, 1960). The external symptoms are very similar to those of rayadilla disease, caused by zinc deficiency. In some ways, they also resemble certain cases of ‘yellow mat’ attributed to adverse soil conditions, where chlorotic striping and narrowing of the leaves occur along with phloem dysfunction. Leaves are erect and rosetted in appearance. The size of leaves is reduced, with laminae irregularly scalloped and cupped upwards. Irregular, elongated, white or yellowish patches or stripes occur, principally in the inter-veinal areas parallel to the veins. Mottling and striping are most prominent on new leaves. Suckers may remain healthy in appearance until approaching the shooting stage, when leaf numbers are reduced from 12–16 to about six malformed, chlorotic leaves. Petioles and leaf sheaths have reddish-brown necrotic streaks in severely affected plants. Rhizomes from affected plants produce affected and stunted suckers, though severity of symptoms varies greatly. According to Allen (1957), there is phloem disorganization in leaves, with symptoms exactly as described by Magee (1927) for bunchy top. The only other disease in Central America in which the phloem is sometimes affected is yellow mat. The cause of Roxana disease has never been determined, though a virus was suspected because of its systemic nature. However, this is unlikely, as the condition never reappeared when Cavendish cultivars were planted extensively in the same province (Stover, 1972).

Yellow Mat This disorder was first described affecting ‘Gros Michel’ (AAA) by Prescott (1917) and called ‘Colorado disease’, as it occurred commonly in the Colorado District near Tela in Honduras. Later, it was reported as most severe in the Changuinola area of Panama (Dunlap, 1923; Permar, 1925). When these areas were abandoned because of Fusarium wilt, no further records were noted until the Changuinola area

was replanted with wilt-resistant cultivars in the Cavendish subgroup (AAA) in the late 1950s and early 1960s. The disorder reappeared and was called ‘yellow mat’, or mata amarilla in Spanish, because of the pronounced yellowing of the foliage. Yellow mat occurs sporadically, rarely more than 5% of the crop being affected, in localized areas along the Atlantic coast of Central America and in Suriname. Barnes (1962) described a disease with similar symptoms on the Cavendish cultivars ‘Lacatan’ and ‘Giant Cavendish’ in wet areas in Grenada. In Panama, all cultivars of dessert banana and plantain (AAA, AB, AAB) were affected. According to Dunlap (1923), yellow mat was also prevalent in abacá throughout the Changuinola area. In other banana-growing areas of Central America, the condition is rare or absent. The first symptoms are a yellowing and browning of the lower leaves, usually on large plants or plants with fruit. The leaves die and the disorder progresses acropetally until only a few of the youngest leaves remain green. New leaves that emerge after symptoms appear are small and narrow. Leaves of large suckers may or may not be affected at first, depending on the severity of attack. If affected, sucker leaves may be narrow, strap-shaped and chlorotic, with chlorosis sometimes in bands. Often, leaf yellowing is the only symptom present on suckers (Stover, 1972). Early symptoms of leaf yellowing and browning with generally poor plant growth are similar to those present on banana growing on heavy clay and in poorly drained areas. However, suckers usually succumb with yellow mat. Often, young sword suckers turn black and die. The death of suckers helps to distinguish yellow mat disease from symptoms of poor growth as a result of adverse soil texture and drainage. Affected plants are usually stunted, the fruit is small and short-fingered and the bunch may not be vertical. Eventually, all the sword suckers attached to plants showing symptoms die. Water suckers will continue to emerge for a few months before the entire mat is destroyed. In a minority of cases, there may be some recovery and renewal of normal growth. When symptoms first appear, an examination of the plant below soil level shows shallow rooting, extensive necrosis and a generally poor root system. Internal symptoms, when present, consist of vascular



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discoloration in the lower pseudostem, usually paler and less extensive than symptoms caused by Fusarium or bacterial wilts. This discoloration was shown by Reinking (quoted by Dunlap, 1923) to involve the phloem and not the xylem. On ‘Gros Michel’, external symptoms were often similar to those of Fusarium wilt. An examination of the lower pseudostem became a quick microscopic method of distinguishing between cases of Fusarium wilt (which affects the xylem) and yellow mat on the wilt-susceptible ‘Gros Michel’. The presence of phloem discoloration was confirmed in outbreaks in Panama in 1964. The phloem sieve tubes were plugged with a dark amorphous material. However, in some cases, phloem discoloration is slight or absent. Rhizomes taken from mats with well-­ developed symptoms may or may not grow. In some cases, growth is normal for several months and then symptoms appear and the plant dies. In a minority of cases, normal growth and recovery from the disease occur. Root growth from affected rhizomes is usually reduced and roots are weak in comparison with roots from normal rhizomes. The above-ground symptoms are believed to be related to deterioration and rotting of the root system. The precise cause of yellow mat is not known. Because of vascular discoloration, a disease caused by a species of Fusarium was suspected, but Reinking (1926) showed that Fusarium spp. associated with the disorder were not pathogenic. Outbreaks of yellow mat are associated with clay or light clay soils, land with drainage problems, extreme fluctuations in the water ­table and rainfall well above average. There is no evidence of yellow mat spreading like a soilborne disease. Flooding alone does not cause symptoms. It is believed that certain toxic soil factors are involved which affect not only the root system but the rhizome as well, and in some way the physiology of the rhizome storage tissue is disturbed. As a result, rhizomes from affected plants usually cannot produce normal plants. The production of these toxic factors is, in some unknown way, related to soil structure, permeability and drainage. In areas where sporadic outbreaks occur persistently, optimum yields are not obtained. Evidence of high clay content and poor soil structure is frequently associated with yellow mat, indicating that it is related to some

soil condition that adversely affects root growth. However, the possibility of a virus being involved cannot be excluded. Early symptoms of yellow mat resemble those of potassium deficiency. Low oxygen and high carbon dioxide levels are known to impede potassium uptake (Hammond et al., 1955). It is possible that potassium uptake may also be impeded when soil conditions favourable for yellow mat are present, though there is no plant response to potassium or minor elements in yellow mat areas. Following extensive improvements in soil drainage in Panama, incidence of yellow mat declined. Existing drains were deepened and new drains opened across contours to remove water more rapidly during periods of heavy rainfall. As a result, water-tables were lowered and root growth increased (Stover, 1972).

Fruit Disorders Alligator Skin A rare blemish called ‘alligator skin’ is occasionally found on green fruit at harvest. Portions of the peel are reddish brown and the surface becomes hard and raised and cracks into rectangular areas, somewhat resembling the skin of an alligator (Plate 8.3). Usually, the blemish occurs on the inner face of the fingers and only a few hands or fingers on a stem are affected. Sometimes a small amount is seen on the outer ridges

Plate 8.3.  Alligator-skin blemish on a hand of green Cavendish bananas at harvest (photo: R.H. Stover).

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of the fingers. The cause of this defect is unknown, but irritation of the young fruit epidermis is suspected. Light abrasions from leaves or bracts that rub the developing fruit when are probably a contributing factor. Finger-ridge scarring can be caused by abrasions from polyethylene bags when blown against the fruit in high winds (Stover, 1972).

Dark Centre of Ripe Fruit The presence of a black centre (Plate 8.4) in otherwise normally ripened fruit is a common source of complaints from consumers. On biting into a ripe banana, the centre is found to be black and soft. There are no outward symptoms of the internal blemish. Dark centres can be induced by the impact of dropping boxes containing fruit with a colour index of 4–5. Dropping individual fingers on their flower tips six times from a height of 15 cm consistently produced dark centres without bruising. Dark centres have also been induced by applying pressures of

70 g/cm to the flower ends of fingers. As ripening advances, dark centres are less likely to develop and green fruit is not susceptible. The condition is believed to result from rupture of latex ducts when ripening fruit is roughly handled. Phenolic substances are released, which, on reaction with polyphenoloxidase enzyme, turn brown or black. Latex cells are less susceptible to rupture in green fruit, as tannins, which inhibit the browning enzymes, are present (Stover, 1972).

Break Neck In Central America, fruit of cultivars in the Cavendish subgroup sometimes falls from the plant as a result of breakage of the peduncle where it curves inward towards the pseudostem. Breakage usually occurs during or shortly after the time the fingers emerge from the inflorescence. Sometimes the peduncle may break at some point within the pseudostem or at the point of emergence from the pseudostem. The peduncle is very brittle at this stage. There are no symptoms of disease or abnormal growth. Incidence is less than 1% and the disorder does not occur throughout the year. ‘Lacatan’ (AAA, Cavendish subgroup) was more susceptible to break neck than other Cavendish cultivars in Honduras (Stover, 1972). The reasons for the weakness of the peduncle are not known, but water stress possibly contributes. Up to 40% of bunches may be lost in hot dry environments. The effects can be moderated with irrigation, but even when well watered, bunch losses can double if pan evaporation increases from 8 mm to 10 mm/day (Hill, 1990). Break neck is also known in extremely cold conditions when weak pseudostems are unable to support heavy bunches. This occurs especially in ‘May flowering’ or ‘November dump’ fruit grown in the subtropics. Nitrogen excess can also cause break neck.

Hard Lump Plate 8.4.  Symptoms of dark centre in ripening fruit of a cultivar in the Cavendish subgroup (AAA) (photo: R.H. Stover).

Fruit of ‘Rasthali’ (AAB, Silk subgroup) commonly suffers from a disorder called ‘hard mass’ or



Non-infectious Disorders of Banana 467

‘lumps’ in India. These lumps, which are pinkish brown in colour and firmer than the usual soft pulp, taste like immature or unripe fruit. Hard lump is of concern because it impairs the eating qualities of affected ‘Rasthali’ bananas, which are highly prized in India. It has been reported that the occurrence of hard lump is seasonal and only fruit harvested from July to December has the problem. The application of 2,4-dichlorophenoxyacetic acid (2,4-D) to bunches, cut ends of peduncles or hands reduced incidence (Shanmugavelu et al., 1992).

Malformed Fingers and Hands Individual fingers and occasionally entire hands are twisted and excessively curved, destroying the symmetry of the hand (Plate 8.5). The symptoms are similar to those caused by low or high temperatures or by water stress. This problem is well known in fruits packed in cartons in the packing house and therefore most of the information is on the Cavendish subgroup of cultivars. Such hands are discarded when packed, because

Plate 8.5.  Highly malformed hands and wild fingers (photo: Y. Israeli JVBES).

they cannot be arranged normally in the box and may cause damage to other hands. Malformed hands commonly occur in the upper part of the bunch. Usually, the basal or second hands are affected, but sometimes other hands as well. The first basal hand is usually the largest in the bunch with an average of 25 fingers, but can vary from four to 33 fingers. A hand with few fingers growing on an elongated crown is called a ‘flying hand’. Hands with numerous fingers in three crowded layers cause finger malformation. From the third hand downwards, malformations are rare. A common cause of fruit malformation is the prevention of normal finger curvature and growth by the presence of leaves or bracts between the fingers or upon the young developing hands of fruit. Sometimes, fruit malformation is caused by the weight of the polyethylene bag obstructing normal curvature of the fingers. This can occur when a young bunch is covered before the fingers have curved upward. Removal of leaves and bracts that touch the young fruit and a delay in bagging of very young fruit reduce malformation. However, even in the absence of these obstructions to finger curvature, malformed fingers and hands can still be found. High-density plantings resulting in reduced illumination and low temperatures during inflorescence development increase the rate of twisted hands in ‘Grand Nain’ (AAA, Cavendish subgroup). In Central America, 1.5–3% of de-handed fruit at the boxing station is discarded because of malformed fingers. Weight loss might be high, especially in young and vigorous plantations with heavy bunches. Packing clusters can reduce the amount of rejected fruit. A frequent malformation found in the Cavendish sub-group is ‘wild finger’ (also called ‘malformed finger’ or ‘naughty finger’) where a single finger is twisted towards the bunch central axis. A high frequency (50%) of bunches with wild fingers was found in a 4-year-old plantation of ‘Grand Nain’ and ‘Williams’ in lowland Philippines. More wild fingers were found at the sides of the hand than at its centre, and more in the upper one to six hands than the lower hands (Y. Israeli, 2016, Israel, unpublished data). Damage is reduced by removing the ‘flying hand’ and the external fingers of the six upper hands. This operation is undertaken in young bunches before bagging and at the same time as deflowering.

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Plate 8.6.  The fusion of fingers in the proximal hand is one of the most common genetic defects (photo Y. Israeli, JVBES).

The removal of young fingers causes latex contamination, but it dries and drops down during fruit growth. De-fingering facilitates hand-tubing (wrapping individual hands in the bunch) and the removal of 12–20 fingers with a high potential of malformation improves much the remaining fruit. The fusion of fingers in the proximal hands is one of the most common genetic defects, especially in large Cavendish bunches (Plate 8.6). Occasionally up to four fingers may be fused together. About 5% of bunches may have fused fingers and up to 0.5% of the weight of banana clusters can be rejected at the packing station because of this defect. Another disorder that causes the rejection of fingers is ‘nipple tip’ related to stress periods, but different from the ‘blunt tip’ that is typical in highland cultivation at lower temperatures. Some other bunch and fruit abnormalities are ‘long peduncle’, ‘short fruit’, ‘fruit ribbing’ and ‘pale male bud’ (Daniells et al., 1999). Fruit malformations are connected to genetic factors and therefore differ among cultivars of the Cavendish subgroup. Relatively more malformations are found in ‘Grand Nain’ than in ‘Williams’ and much less in ‘Dwarf Cavendish’ or the high cultivars such as ‘Valery’ or ‘Lacatan’.

Maturity Stain Maturity stain, which is known as ‘maturity bronzing’ in Australia, is a peel blemish that

Plate 8.7.  Maturity stain on Cavendish fruit (photo: Y. Israeli, JVBES).

appears only on fruit with an excessive caliper grade. Longitudinal streaks and blotches appear mostly on the shoulder of outer whorl fingers of basal hands (Plate 8.7). This stain should not be confused with the reddish discoloration from red rust thrips or from a chemical injury (Stover, 1972). Maturity stain is stress induced and correlated in Australia and Costa Rica with periods of heavy rains, high temperatures, high humidity and a possible potassium/magnesium imbalance. On the Pacific side of Panama and Costa Rica, the normal rejection level of fruit because of maturity stain has been reported to be 8–10%. However, this rises to 20% during periods of heavy rain (Stover and Simmonds, 1987). Maturity stain is also associated with a late harvest when fruit is of a high caliper grade. A considerable amount of research on maturity stain has been undertaken in Australia. Campbell and Williams (1976) suggested that it was caused by high turgor pressure in the non-­dividing cells of the peel epidermis, which resulted in the breakdown of the middle lamellae, cell rupture and oxidation of the released cell constituents. Anatomical studies on the peel of the developing fruit showed that stretching of the epidermis exceeds its elastic limit and leads to cracks and cell disruption in the peel surface and, subsequently, to formation of lesions characteristic of maturity stain (Williams et al., 1990). Sealed polyethylene bags and the delayed harvest of bunches in these bags increased maturity stain compared with open bunch covers (Johns and Scott, 1989; Daniells et al., 1992). It has been found to be greatest in fruits kept in



Non-infectious Disorders of Banana 469

darkness and was prevented with continuous exposure to white light (Wade et al., 1993). The disorder was found to begin developing on fruit of water-stressed bunches at a much thinner finger diameter than in fruits of unstressed controls (Daniells et al., 1987). It showed a positive association with the amount of copper in the fruit skin and calcium in leaves, and a negative association with the amount of calcium and magnesium in the fruit skin (Campbell and Williams, 1978). The removal of leaves reduced the incidence of the fruit-peel disorder, while bunch trimming increased the incidence (Daniells et al., 1994). However, it was concluded that the ­reduction in maturity stain achieved by leaf removal is of no immediate benefit to the growers. Treatments of partial benefit are better crop ­nutrition and irrigation, and an aerial spray of calcium nitrate (Ca(NO3)2) incorporated with fungicides. Harvesting fruit at a lower grade can also reduce incidence.

Neer Vazhai Neer vazhai is a disorder affecting 1–2% of ‘Nendran’ (AAB, Plantain subgroup) grown in the Tiruchirappalli district of Tamil Nadu State in southern India. Affected plants appear normal until after flowering, when fingers on many of the hands fail to fill (Plate 8.8). Unfilled fingers tend to have persistent female flower parts. When suckers from affected plants develop into mature plants, they in turn produce deformed bunches. The problem may be genetic (Shanmugavelu et al., 1992).

Physiological Finger Drop When bananas were shipped on stems (peduncles), losses were encountered as a result of fingers falling from the bunch (loose fingers). This usually occurred after ripening had begun. In the French Antilles, finger drop or ‘de-grain’ with

Mixed Ripe Fruit Mixed ripe fruit, where some of the fruit ripens before marketing while some stays green, is a more widely spread phenomenon than premature ripening. The time that fruit ripens is related to its physiological age and not its size. Variable age of bunches harvested is a major contributing factor, with the oldest bunches more prone to mixed ripening. Some fruit can ripen while still small if growth has been delayed due to stress conditions (Turner and Rippon, 1973; Rippon, 1975). Recording flowering time on emergence and harvesting at the right physiological age reduce mixed ripening. Polyethylene-sleeved fruit is known to ripen earlier than unbagged fruit. Methods to maximize green life, age-grade control and proper postharvest handling were summarized by Daniells (1991). Marriott (1980) listed some of the factors shortening the green life of fruit and possibly inducing mixed ripening. High levels of Sigatoka leaf spot diseases on plants as bunches develop and mechanical damage to fruit also stimulate earlier ripening. In general, environmental stresses affecting fruit growth, such as drought or chilling, may stimulate mixed and premature ripening.

Plate 8.8.  Neer vazhai disorder in a bunch of ‘Nendran’ (AAB, Plantain subgroup) growing in India (photo: S. Sathiamoorthy, TNAU).

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a physiological cause was a serious seasonal problem as stem fruit ripened (Daudin and Guyot, 1965; Guillemot and Colmet-Daage, 1965). Phillips (1970) reported a similar finger drop problem in the Windward Islands. Finger drop should not be confused with finger loss resulting from mechanical injury to the necks of the finger, which may or may not be accompanied by a fungal infection, or with stemend rot. With physiological finger drop, the fruit detaches itself from the pedicel as ripening progresses. The fibrous system of the pedicel is not involved, since the break takes place at the placental axis of the fruit, usually before the finger is fully ripe. Physiological finger drop is the premature separation of the finger from the pedicel. Fruit with finger drop tends to yellow more quickly, while the pedicel remains green. The skin is dull yellow, with a silky, soft texture. Guillemot and Colmet-Daage (1965) found a close relationship between the pedicel ratio (pedicel length divided by width) and a tendency for finger drop. They believed that finger drop was promoted by sharp fluctuations in the amount of soil nitrogen and recommended that levels should be kept high during the period following the first heavy rain. Application of nitrogen was to be avoided at the end of the dry season. Phillips (1970) found that the pedicel ratio was adversely affected by dense planting (3000 plants/ha or more) and heavy applications of fertilizer. Finger drop has been associated with nitrogen imbalance (Martin-Prevel and Montagut, 1966). It occurs during the hot, wet season in the tropics when there is a low potassium supply and ammonium nitrogen (N-NH4) accumulates. The excess nitrogen delays bunch emergence and produces bunches with widely spaced hands, which are easily damaged in transport. Short-cycle fruit (fruit reaching maturity in a short time from planting) was found to be more susceptible to finger drop than long-cycle fruit (Daudin and Guyot, 1965). However, no anatomical differences were found between shortand long-cycle fruit. It was thought that finger drop was related to seasonal factors and any other factor that reduced the maturation time of the plant crop to less than 350–365 days from planting. Vallade and Rabechault (1963) could find no differences in the anatomy of the pedicel that would explain susceptibility to finger drop.

In Australia, Hicks (1934) attributed a seasonal finger dropping to ripening at high temperatures and humidity. Finger drop was reduced when ripening fruit was removed to lower humidity or green fruit was held at lower temperatures before ripening. Semple and Thompson (1988) reported that temperatures up to 30°C and ethylene increased finger drop, while relative humidity had no effect. Some bred tetraploid cultivars have an inherent weakness in finger attachment to the pedicel at ripening. This defect has made these cultivars unsuitable for the commercial trade (New and Marriott, 1974).

Premature Field Ripening This specific problem occurred in the Changuinola area of Panama and was common from 1955 to 1961 on ‘Gros Michel’ and cultivars in the Cavendish subgroup. There was an irregular development of hands, with the proximal or upper hands on the hanging bunch filling out and ­maturing more rapidly than the apical or lower hands. The full, proximal fingers ripened on the plant and, in advanced stages of ripeness, the fingers fell from the stem while the apical hands were still green. Outbreaks of the condition were sporadic, starting with many bunches followed by a decline to few or none. The worst outbreaks followed a drought, a flood or a night with frost in the autumn. Therefore, it appears that this problem is probably related to a serious check in the growth of the plant during the period when the fruit is filling. Inspections indicated that the root systems were deficient on plants with premature field ripening (Stover, 1972).

Sinkers Fruit normally floats in water when the hands are removed from the peduncle and placed in de-latexing tanks at the packing station. However, sinkers appear during some periods of the year (such as winter in the subtropics) and from some areas. The term ‘sinker’ is applied to fruit that sinks to the bottom of the tank, where, unless quickly removed, it remains and becomes excessively scarred due to abrasion damage (Stover, 1972).



Non-infectious Disorders of Banana 471

Sinkers are most common in fruit with a slow sap flow, from areas where growing conditions have been poor because of soil conditions, or when there is adverse weather, such as excessive rainfall. The appearance of sinkers in Honduras is seasonal and may reach 5% of the fruit (Johnson, 1979). Sinkers have relatively low levels of potassium and can be avoided by applying adequate amount of this nutrient or organic matter. Sinkers do not necessarily ripen sooner than normal fruit.

Split Peel Split peel (Plate 8.9) results from several causes, some of which have not been defined. Split peel of green fruit in the field results from a too rapid filling out of the fingers, usually on the proximal or largest hands. The rapid expansion in pulp volume results from highly favourable growing conditions or failure to harvest fruit when the

correct harvesting caliper grade is reached. Split peel may occur during transit and when ripening. A too-high ripening temperature is a contributing factor (Stover, 1972). Split peel was common on ‘Gros Michel’ ­ripened in November to February in Honduras. Banana growth during this period is retarded, as a result of low temperatures, and fruit maturation time is increased. Dipping the fruit in hormone solution, such as 2,4-D, before ripening reduced the amount of split peel (Freiberg, 1955). Sometimes split peel occurs in transit on fruit that is not over mature. The cause is unknown. Split peel may also occur as a result of sudden frost or drought stress. It also an expression of a rare somaclonal variant of plants of Cavendish cultivars propagated by tissue culture (­Israeli et al., 1991) and is a common postharvest defect in the fruit of the Fusarium wilt-tolerant ‘GCTCV-119’ selection from Taiwan.

Uneven De-greening Uneven de-greening, which becomes evident following ethylene treatment, is a serious ripening disorder of bananas harvested from some selections of cultivars in the Cavendish subgroup (AAA) in subtropical areas. In Taiwan, it affects fruit from plants of ‘Pei-Chiao’, which flower between December and March, the period of lowest temperatures. In mild cases, either affected fruit only partially yellows or yellowing is delayed. In severe cases, the peel remains green (Plate 8.10). The disorder mainly affects the top

Plate 8.9.  Symptoms of split peel on green Cavendish fruit at harvest (photo: E. Lahav, VI, and Y. Israeli, JVBES).

Plate 8.10.  Uneven de-greening of bananas harvested from ‘Pei-Chiao’ (AAA, Cavendish subgroup). Normally de-greened bananas of uniform yellow colour (right) contrast with unevenly de-greened bananas (left) (photo: S.C. Chiang, TBRI).

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three hands. Temperatures below 20°C and genetic characteristics are believed to be the major contributing factors. An integrated strategy, including the elimination of selections of ‘Pei-Chiao’ that are prone to the problem and the use of brown paper bags as bunch covers instead of blue polyethylene, were tested to overcome the problem. Brown paper bags affect the microclimate of the bunch, as well as blocking infrared (IR) radiation and blue-green light, resulting in changes to the configuration of epidermal cells (Chiang et al., 1998; Tang, 1998).

Withered Pedicels In boxed fruit, pedicels sometimes wither during ripening, especially following transit periods in excess of 10 days. The pedicel surface appears wrinkled and shrivelled, and is dull-brown or grey in colour in contrast to the normal greenish-yellow of turgid pedicels. Clusters with shrivelled pedicels are found alongside normal, turgid pedicels. Occasionally, in some locations, up to 50% of the clusters can be affected. This occurs even when the fruit is under high humidity in polyethylene bags. The cause of withered pedicels is not known. Incidence increases the longer the fruit is held at ambient temperatures prior to shipboard storage at 13–15°C. Incidence is also greater in fruit from areas with poor soil or poor drainage or when growing conditions are not ideal (Stover, 1972).

Yellow Pulp The colour and texture of the pulp of green fruit when harvested at the correct grade is normally dull white and firm. Sometimes, the pulp has a

yellow or honey-coloured tint and is soft in the centre of the finger. Such fruit is said to have ‘yellow pulp’. In some cases, a pinkish cast may be present. Colour and texture changes are most pronounced near the centre of the finger, and are usually indicative of approaching ripening or breaking of skin colour prematurely (Stover, 1972). Any factors delaying the filling out of the fruit or increasing the time to harvest maturity can cause yellow pulp. They include loss of foliage, due to Sigatoka leaf spot diseases or defoliation, excessive shade or drought. In Guadeloupe, yellow pulp was detected following prolonged drought by cutting open fingers at the packing station. Fruit with yellow pulp is discarded because it would ripen in transit. In Ecuador, in an effort to avoid fruit ripening in transit, a wing finger is cut on each bunch to ensure that the pulp has the correct colour. In Central America, populations of 1976– 2470 plants/ha on good loam soils can result in a softening of fruit before harvest and yellow pulp. Competition for limited light and space results in fruit requiring abnormally long periods to reach harvesting grade. Such fruit is ‘stale’ at harvest and often has a tendency to ripen in transit (Stover, 1972). In Cameroon and Guinea, up to 70% of fruit from areas where yellow pulp was most severe has been rejected for export on occasions in the past. In these areas, high potassium/magnesium ratios, high soil-calcium levels or deficiencies in magnesium or manganese may have caused yellow pulp (Dumas and Martin-Prevel, 1958; Charpentier and Martin-Prevel, 1968). Yellow pulp was avoided by applying high doses of sulfur to the soil, which improved the balance of cations by reducing excess calcium and increasing the absorption of manganese (Marchal et al., 1972).

References Allen, R.M. (1957) A virus-type disease of Gros Michel bananas in Costa Rica. Turrialba 7, 72–83. Barker, W.G. (1959) A system of maximum multiplication of the banana plant. Tropical Agriculture (Trinidad) 36, 275–284. Barnes, R.F. (1962) Grenada banana disease. In: Report on Banana Investigations 1962. Regional ­Research Centre, University of the West Indies, Trinidad, pp. 25–26. Brouhns, G. (1957) Note sur la croissance du bananier ‘Gros Michel’. Fruits 12, 261–268. Campbell, S.J. and Williams, M.T. (1976) Factors associated with maturity bronzing of banana fruit. Australian Journal for Experimental Agriculture and Animal Husbandry 16, 428–432.



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Campbell, S.J. and Williams, M.T. (1978) Mineral relationships in ‘maturity bronzing’ of banana fruit. Australian Journal for Experimental Agriculture and Animal Husbandry 18, 603–608. Charpentier, J.M. (1966) La remontée du méristème central du bananier. Fruits 21, 103–119. Charpentier, J.M. and Martin-Prevel, P. (1968) Carences et troubles de la nutrition minéral chez le bananier. Guide de diagnostic pratique. Institut Français de Recherches Fruitières Outre-Mer, Paris, 75 pp. Chiang, S.C., Tang, C.Y., Chao, C.P. and Hwang, S.C. (1998) An integrated approach for the prevention of uneven degreening of bananas in Taiwan. Acta Horticulturae 490, 511–518. Daniells, J. (1991) How to avoid mixed ripe problems with bananas. Queensland Fruit and Vegetable News 62(4),12–13. Daniells, J.W., Watson, B.J., O’Farrel, P.J. and Mulder, J.C. (1987) Soil water stress at bunch emergence increase maturity bronzing of banana fruit. Queensland Journal of Agriculture and Animal Sciences 44, 97–100. Daniells, J.W., Lisle, A.T. and O’Farrell, P.J. (1992) Effect of bunch covering methods on maturity bronzing, yield and fruit quality of bananas in North Queensland. Australian Journal of Experimental Agriculture 32, 121–125. Daniells, J.W., Lisle, A.T. and Bryde, N.Y. (1994) Effect of bunch trimming and leaf removal at flowering on maturity bronzing, yield and other aspects of fruit quality of bananas in North Queensland. Australian Journal of Experimental Agriculture 34, 259–265. Daniells J. W., Smith M. K. and Hamill S. D. (1999) Banana Offtypes, an Illustrated Guide. Information Series QI99019. Department of Primary Industries, Queensland. Daudin, J. and Guyot, H. (1965) A study of finger drop affecting the Giant Cavendish banana in the French West Indies. In: The Banana Industry and Research Development in the Caribbean. Caribbean Organization, Puerto Rico, pp. 131–144. Dumas, J. and Martin-Prevel, P. (1958) Contrôle de nutrition des bananeraies en Guinée (premiers résultats). Fruits 13, 375–386. Dunlap, V.C. (1923) Weekly Reports on Banana Investigations. United Fruit Co., Panama Division. Freiberg, S.R. (1955) Effect of growth-regulators on ripening, split peel, reducing sugars, and diastatic activity of bananas. Botanical Gazette 117, 113–119. Guillemot, J. and Colmet-Daage, F. (1965) Factors affecting quality of bananas in the West Indies: finger drop – effects of variation in the nitrogen content of the soil on finger drop. In: Banana Industry and Research Developments in the Caribbean. Caribbean Organization, Puerto Rico, pp. 36–55. Hammond, L.C., Allaway, W.H. and Loomis, W.E. (1955) Effects of oxygen and carbon dioxide levels on absorption of potassium by plants. Plant Physiology 30, 155–161. Hasselo, H.N. (1957) Earthing-up of Gros Michel bananas. Tropical Agriculture (Trinidad) 34, 59–64. Hicks, E.W. (1934) Finger dropping from bunches of Australian Cavendish bananas. Journal of the Council for Scientific and Industrial Research, Australia 7, 3. Hill, T.R. (1990) Scheduling irrigation for bananas with tensiometers. In: Growing Bananas in an Extreme Environment. First National Banana Symposium, 25–27 June 1990. Kununurra, Western Australia, pp 54–55. Israeli, Y., Reuveni, O. and Lahav, E. (1991) Qualitative aspects of somaclonal variations in banana propagated by in vitro techniques. Scientia Horticulturae 48, 71–88. Johns, G.G. and Scott, K.J. (1989) Delayed harvesting of bananas with ‘sealed’ covers on bunches. 2. Effect on fruit yield and quality. Australian Journal of Experimental Agriculture 29, 727–733. Johnson, T.J. (1979) Effects of potassium on buoyancy of banana fruit. Experimental Agriculture 15, 173–176. Magee, C.J.P. (1927) Investigations on the bunchy-top disease of bananas. Bulletin Australian Council for Scientific and Industrial Research 30, 30–64. Marchal, J., Martin-Prevel, P. and Melin, P. (1972) Le soufre et le bananier. Fruits 27, 167–177. Marriott, J. (1980) Banana: physiology and biochemistry of storage and ripening for optimum quality. CRC Critical Reviews in Food Science and Nutrition 13, 41–88. Martin-Prevel, P. and Montagut, G. (1966) Essais sol-plante sur bananier. 8. Dynamique de l’azote dans la croissance et le développement du végétal. Fruits 21, 283–294. New, S. and Marriott, J. (1974) Post-harvest physiology of tetraploid banana fruit: response to storage and ripening. Annals of Applied Biology 78,193–204. Permar, J.H. (1925) Colorado disease in the Farm. 8. Lacatan planting. In: Annual Report United Fruit Company for Period December 27, 1924 to December 5, 1925, Panama Division. Phillips, G.A. (1970) Fruit quality problems of the Windward Islands’ banana industry. PANS 16, 298–303.

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Prescott, S.C. (1917) Diseases of the Banana. Bulletin 2. United Fruit Company Research Department, La Lima, Honduras, 35 pp. Reinking, O.A. (1926) Fusaria inoculation experiments: relationship of various species of fusaria to wilt and Colorado disease of bananas. Phytopathology 16, 371–392. Rippon, L.E. (1975) Knowing bunch age can increase growers’ profits. Banana Bulletin 39(4), 2 Robinson, J.C. and Galán Saúco, V. (2010) Bananas and Plantains. Crop Production Science in Horticulture 19, CAB International, Wallingford, UK, 311 pp. Semple, A.J. and Thompson, A.K. (1988) Influence of the ripening environment on the development of finger drop in bananas. Journal of the Science of Food and Agriculture 46, 139–146. Shanmugavelu, K.G., Aravindakshan, S. and Sathiamoorthy, S. (1992) Banana Taxonomy, Breeding and Production Technology. Metropolitan Book Co., New Delhi, India, 456 pp. Stover, R.H. (1972) Banana, Plantain and Abacá Diseases. Commonwealth Mycological Institute, Kew, UK, 316 pp. Stover, R.H. and Simmonds, N.W. (1987) Bananas. Longman Scientific and Technical, Harlow, UK, 468 pp. Subra, P. and Guillemot, J. (1961) Contribution à l’étude du rhizome et des rejets du bananier. Fruits 16, 19–23. Tang, C.Y. (1998) Response to selection of uneven degreening in the clones of a Cavendish banana cultivar (Musa cv. AAA) in Taiwan. Fruits 53, 355–363. Turner, D.W. and Rippon, L.E. (1973) Effect of bunch covers on fruit growth and maturity in bananas. Tropical Agriculture (Trinidad) 50, 235–240. Vallade, J. and Rabechault, H. (1963) Etude des caractères anatomiques des pédicelles de bananes, en corrélation avec le degrain. Fruits 18, 129–140. Wade, N.L., Kavanagh, E.E. and Tan, S.C. (1993) Sunscald and ultraviolet light injury of banana fruits. Journal of Horticultural Science 68, 409–419. Waite, B.H. (1960) Virus diseases of bananas in Central America. Proceedings of the Caribbean Region, American Society of Horticultural Science 4, 26–30. Wardlaw, C.W. (1961) Banana Diseases. Longman, Green, London, pp. 40–42. Williams, M.H., Vesk, M. and Mullins, M.G. (1990) Development of the banana fruit and occurrence of the maturity bronzing disorder. Annals of Botany 65, 9–19.

9 

Mineral Deficiencies of Banana E. Lahav and Y. Israeli

Introduction For optimal growth and fruit production, banana plants require high amounts of nutrients, which are often only partly supplied by the soil (Table 9.1). Thus, to obtain high yields, large quantities of mineral nutrients have to be replaced in order to maintain soil fertility. This is achieved by applying organic manure and/or, more efficiently, mineral fertilizers, which supply the elements in a concentrated and readily available form. The overall requirement of mineral nutrients can be estimated from an analysis of the whole plant and on an estimation of plant growth. The grower must know the ability of the soil to meet the requirements necessary for good growth and whether supplementary fertilizers are needed. Two approaches have been utilized to help solve this problem. In one approach, field experiments can be established on a range of soil types. As the results of these trials are also dependent upon local conditions of climate, the cultivar grown, the type of irrigation system and the effect of pests and diseases (especially nematodes), their reliable extrapolation is limited. In order to make the results of field experiments more meaningful, plant tissues and soil samples are usually analysed as part of a second approach to determine mineral nutrient levels with the aim of estimating the amount of fertilizer and microelements required to optimize yields (Lacoeuilhe

and Martin-Prevel, 1971; Marchal et al., 1972; Warner and Fox, 1977). Deficiency symptoms are useful in diagnosing mineral nutrient imbalance and are summarized in Table 9.2. However, once visual symptoms are seen, yield loss may have already been experienced.

Nitrogen Deficiency Growth of the banana plant is more sensitive to a lack of nitrogen (N) than of any element. Under sand-culture conditions, nitrogen deficiency more than halves the rate of leaf production, while deficiencies of other elements have only a slight effect (Murray, 1960). For banana, the relationship between total dry-matter production and total nitrogen uptake is close (Lahav and Turner, 1983). Therefore, the first indication of nitrogen deficiency is a reduced rate of growth resulting in a reduction in yield. After the application of nitrogen, there is an immediate increase in growth and fruit weight. Nitrogen is rapidly redistributed from old banana leaves to young ones (van der Vorm and van Diest, 1982). Hence, deficiency symptoms appear quickly and soon all leaves are affected. The leaves are pale green, with the midribs, petioles and leaf sheaths becoming reddish pink (Plate 9.1) (Murray, 1959). The distance between successive leaves is reduced, giving the plant a ‘rosette’ appearance. Nitrogen deficiency

© D.R. Jones 2019. Handbook of Diseases of Banana, Abacá and Enset (D.R. Jones)

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Table 9.1.  Average amount of nutrients (kg/ha) in banana and plantain, based on 2000 mother plants/ha with followers and an average bunch weight of 25 kg bananas (50 t/ha) and 15 kg plantains (30 t/ha); roots not included (from Lahav, 1995). Amount removed by bunches Nutrient N P K Ca Mg S Cl Na Mn Fe Zn B Cu Al Mo

Amount in remaining plants

Proportion removed by fruit (%)

Total

Banana

Plantain

Banana

Plantain

Banana

Plantain

Banana

Plantain

189 29 562 10–26 49 23 75 1.6 0.5 0.9 0.5 0.7 0.2 0.2 –

76 11 243 9 11 9 – – 0.2 0.4 0.1 – 0.04 – –

199 23 660 126 76 50 450 9 12 5 4.2 0.57 0.17 2 0.0013

189 16 945 149 53 19 – – 6.8 3.1 0.7 – 0.1 – –

388 52 1222 136–152 125 73 525 10.6 12.5 5.9 4.7 1.27 0.37 2.2 –

265 27 1188 158 64 28 – – 7 3.5 0.8 – 0.14 – –

49 56 46 8–17 39 32 14 15 4 15 12 55 54 9 –

29 41 20 6 17 32 – – 3 11 12 – 29 – –

Table 9.2.  Summary of mineral deficiency symptoms. Age of leaf

Symptoms on leaf lamina

Additional symptoms

Element

All ages

Uniform light green or yellowing coloration

Pink petioles; stunted growth

Nitrogen

Midribs curving resulting in weeping; drooping leaves

Copper

Young leaves

Yellow to almost white coloration with inter-veinal chlorosis Pale green to yellow coloration including veins Streaks across veins Yellow stripes along veins Marginal chlorosis

Old leaves

‘Sawtooth’ marginal chlorosis Yellow discoloration in mid-blade; midrib and margins remain green Dirty yellow-green Yellow-orange and brown scorching along margins

symptoms are often observed under conditions of poor rooting and when there is weed competition. In general, nitrogen deficiency may result from any factor that reduces growth, such as drought or poor drainage.

Iron Thickening of secondary veins; leaves deformed Leaf lamina incomplete Reddish coloration on lower side of young leaves Thickening veins; necrosis from margins inward; leaves deformed Petiole breaking; dark green-purple colour of young leaves Limit of chlorotic borders not clearly defined; pseudostem disintegrating Leaf bending; rapid leaf desiccation

Sulfur Boron Zinc Calcium Phosphorus Magnesium

Manganese Potassium

The banana plant cannot store nitrogen (Martin-Prevel and Montagut, 1966) and it is easily leached from the soil. Therefore, nitrogen is almost universally in short supply, even for plants grown on the very fertile soils of Central



Mineral Deficiencies of Banana 477

Plate 9.1.  Symptoms of nitrogen deficiency in banana in St Lucia, Windward Islands. Note the pink colour of the petioles and the ‘rosette’ effect due to the short internodes between petioles (photo: E. Lahav, VI).

America (Butler, 1960). In order to ensure availability and to look after the environment, nitrogen should be supplied in frequent small applications and by controlled release. Nitrogen deficiency is easily corrected by a variety of nitrogen fertilizers. The most commonly used is urea (CO(NH2)2); however, it should be noted that its constant application might significantly reduce soil pH. Other fertilizers are ammonium nitrate (NH4NO3), ammonium sulfate ((NH4)2SO4), potassium nitrate (KNO3) and calcium nitrate (Ca(NO3)2). The nitrogen compound used will depend on soil pH, the presence and type of irrigation, and cost (Lahav, 1995). Nitrogen is also a component of compound fertilizers, which also include potassium as 4:0:12 or 10:0:30 and phosphorus as 5:25:0.

Phosphorus Deficiency Deficiency symptoms are rarely seen in the field. This is because phosphorus (P) requirements of the plant are low and there are usually adequate amounts in the soil, as it is not leachable. In addition, banana plants accumulate the phosphorus they require over an extended period of time and a relatively small quantity of the element is taken from the plant with the harvested fruit. Phosphorus is also easily redistributed from old to young leaves (van der Vorm and van Diest, 1982), from leaves to the bunch (Lahav, 1974) and from mother plant to followers (Walmsley and Twyford, 1968). Hence, phosphorus deficiency,

Plate 9.2.  Yellow chlorotic stripes along the secondary veins due to phosphorus deficiency (photo: J. W. Daniells, QDAF).

if observed at all, will be seen early in the development of the plant crop. Deficiency symptoms have been recorded in the field in Dominica (Stover and Simmonds, 1987) and Guadeloupe (Lacoeuilhe and Godefroy, 1971), and in sand culture (Charpentier and Martin-Prevel, 1965). A low supply of phosphorus results in stunted plants and poor root development. Older leaves initially exhibit yellow chlorotic stripes along the secondary veins (Plate 9.2) and marginal chlorosis, in which ­purplish-brown flecks develop. Eventually, a ‘sawtooth’ necrosis appears. The affected leaves curl, the petioles break and the younger leaves have a deep bluish-green colour. Sucker and plant growth is reduced, but bunch weight is affected only when the deficiency is severe and prolonged. It is presumed that the banana plant obtains adequate phosphate from soils through its mycorrhizal association (Lin and Fox, 1987; Fox, 1989). Vesicular–arbuscular mycorrhizal fungi can invade banana roots and improve plant growth in soils that differ considerably in phosphorus content.

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The most commonly used phosphorus fertilizers are superphosphate, monoammonium phosphate (NH4H2PO4), diammonium phosphate ((NH4)2H2PO4) and rock phosphate. These fertilizers can be applied at any time of the year, but for more soluble phosphorus fertilizers, such as phosphoric acid (H3PO4), application, especially in the subtropics, should be confined to the growing season. Phosphoric acid, because of its strong acidity, is not recommended for soils with low pH.

Potassium Deficiency Potassium (K) is a key element in banana nutrition. When deficient, leaves become smaller in size and midribs curve so that the tip of the leaf points towards the base of the plant (Plate 9.3) (Murray, 1959; Martin-Prevel and Charpentier, 1963; Lahav, 1972). However, the predominant effect of potassium is on the longevity of the leaf (Murray, 1960; Lahav, 1972) and the most common symptom of potassium deficiency is the appearance of spreading orange-yellow patches in the oldest leaves, followed by their rapid desiccation (Plate 9.3). Other effects of potassium deficiency are choking, delay in flower initiation and reduced numbers of fingers/hand and numbers of hands/bunch. Fruit size is also seriously affected by a shortage of potassium (Lahav, 1995) with its growth restricted because a smaller leaf area and delayed stomatal activity result in a

reduction in total dry-matter production, carbohydrate translocation and photosynthesis. Protein synthesis is also impaired and the conversion of sugars to starch is restricted. Potassium uptake is highest during the first half of the vegetative growth phase of the banana plant and is much reduced 2–3 weeks after bunch emergence, even when the potassium supply is abundant. As the bunch attracts potassium from other parts of the plant, it is important that the plant has an adequate supply of the element early in its development (Lopez and Espinosa, 1995). Therefore, a temporary potassium deficiency is often seen at flowering. This is different to the situation with cultivars in the plantain subgroup. Here, plants take up more than half of their potassium requirements after flowering (Lahav, 1995). Potassium deficiency is common in the less fertile banana soils and there are numerous reports of positive responses of banana plants to the application of potash (Bhangoo et al., 1962; Stover and Simmonds, 1987). On volcanic soils in Cameroon, Hasselo (1961) described a premature yellowing of ‘Lacatan’ (AAA, Cavendish subgroup), which was corrected by potash application. Similarly, in Jamaica, Tai (1959) stated that potash fertilizer reduced a premature yellowing of banana. In Honduras, banana plants on certain leached, acid soils responded to potash if nitrogen was also applied (Bhangoo et al., 1962), but no response to potash was obtained in fertile alluvial soils (Butler, 1960). Potassium chloride (KCl) is the form of potassium usually applied to banana plantations. However, if soil salinity is a problem, fertilizers containing chlorides should be avoided and the high potassium requirement of the plant satisfied by using potassium sulfate (K2SO4) or potassium nitrate (KNO3), which increases soil pH. Compound fertilizers with nitrogen and potassium present in the ratios as 4:0:12 or 10:0:30 can also be used as a potassium source for banana.

Calcium Deficiency Plate 9.3.  Symptoms of potassium deficiency in banana grown in sand culture in Israel. Note the orange-yellow patches on the leaf, which are beginning to desiccate, and the downward bend of the midrib (photo: E. Lahav, VI).

Symptoms of calcium (Ca) deficiency first appear on the youngest leaves as thickened lateral veins, being especially noticeable near the midrib. Later, the tissue between veins at the leaf margin becomes chlorotic (Plate 9.4). This symptom is



Mineral Deficiencies of Banana 479

Plate 9.4.  Symptoms of leaf-edge chlorosis due to calcium deficiency in banana in Costa Rica (photo: E. Lahav, VI).

generally more common near the tip of the leaf. The affected tissue expands towards the midrib, turns necrotic and gradually gives the leaf a ‘sawtooth’ appearance (Martin-Prevel and Charpentier, 1963). It should be mentioned that the symptoms resemble those of leaf-edge chlorosis (see Plate 8.2). Calcium deficiency symptoms also include ‘spike leaves’. Here, the lamina is ­deformed (Plate 9.5) or almost absent, due to a temporary shortage of calcium caused by a flush of rapid growth (Lahav and Turner, 1983). Symptoms appear mainly on soils poor in calcium, with a low pH, or as a result of large amounts of potassium (antagonist to calcium) applied with inadequate rainfall. The plants recover when roots grow after rains or when ­potassium is leached below root volume. ‘Spike leaves’ can appear in soils rich in calcium as a result of low root activity in the early spring when the sucker regenerated growth and new leaves are emerging.

Plate 9.5.  Deformed lamina as a result of calcium deficiency in banana in New South Wales, Australia (photo: E. Lahav, VI).

Calcium is supplied to banana plantations in the carbonate form as lime or dolomite and also as a component of superphosphate (21% calcium). Dolomite is preferable to lime since it has higher buffering capacity and it also contains magnesium. Carbonates are usually applied to adjust soil pH rather than to increase the supply of calcium as a plant nutrient, though the two go hand in hand. The amount of lime applied depends upon the change in soil pH ­required and the buffering capacity of the soil. A common rate for banana is 1–2 t/ha annually or 3–6 t/ha every 3–5 years.

Magnesium Deficiency Magnesium (Mg) deficiency has been reported from many areas where banana is cultivated. It usually occurs where banana has been grown for 10–20 years without magnesium fertilizer

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(Chalker and Turner, 1969) on soils with low cation exchange capacity or where high amounts of potassium have been given for a number of years (Messing, 1974). A large range of leaf symptoms has been attributed to magnesium deficiency. These include marginal yellowing extending to near the midrib, changes in the arrangement of leaves, purple mottling of the petioles, marginal necrosis and separation of leaf sheaths from the pseudostem. The most common symptom in the field is that leaf margins of older leaves usually remain green, while the area between the margin and the midrib becomes chlorotic (Plate 9.6) (Murray, 1959; Martin-Prevel and Charpentier, 1963; Chalker and Turner, 1969; Turner and Bull, 1970). Imbalances in the magnesium/potassium ratio can also cause problems. The magnesium deficiency disorder called ‘blue’, because of the bluish and brown marbling on the petioles (Plate 9.7), may be accentuated by potassium excess. Charpentier and Martin-Prevel (1968) indicate that a ‘clearing blue’ disorder may be because of potassium deficiency as a result of excess magnesium. However, the magnesium-­ deficient ‘blue’ disorder is more common, because application of potassium fertilizer is more frequent than that of magnesium. In this disorder, the fruit maturation period is increased and there is a tendency to yellow pulp. Analysis reveals that soils are only slightly saturated in bases and calcium addition could improve the potassium– magnesium imbalance. Charpentier and Martin-­ Prevel (1968) concluded that potassium, calcium

Plate 9.6.  Yellow discoloration in the mid-leaf area due to magnesium deficiency. Midrib and margins remain green (photo: E. Lahav, VI).

and magnesium deficiencies involve an imbalance of at least two of the three cations. Soil and tissue analyses are therefore essential to correct the problem. A practical formula that calculates potassium–calcium–magnesium leaf relations is given by Lopez and Espinosa (1995). Although the expression of magnesium deficiency symptoms is not necessarily associated with reduced yields, magnesium is usually applied when symptoms occur. However, the simple addition of magnesium-containing compounds to the soil does not always give the expected rapid response (Turner and Bull, 1970). It may be necessary to replant and incorporate magnesium into nitrogen, phosphorus and potassium mixtures so that small, regular additions of magnesium are given to the banana plants (Messing, 1974). Soil-applied soluble magnesium fertilizers, such as keiserite (magnesium sulfate monohydrate (MgSO4.H2O)), give a more rapid response than the less soluble compounds as dolomite,

Plate 9.7.  Banana in Côte d’Ivoire with symptoms of ‘blue’, the magnesium deficiency disorder which is caused by a potassium/magnesium imbalance (photo: R.H. Stover).



Mineral Deficiencies of Banana 481

but they are also more expensive. Since magnesium deficiency is unlikely to become a sudden problem in the field, the regular application of small amounts of magnesium seems advisable.

Sulfur Deficiency Deficiencies of sulfur (S) have been reported in the field, as well as in sand-culture experiments (Charpentier and Martin-Prevel, 1965; Messing, 1971; Marchal et al., 1972). Sulfur is actively redistributed from old to young leaves (van der Vorm and van Diest, 1982). When there is a shortage, young leaves first turn yellowish white (Plate 9.8). As the deficiency progresses, necrotic patches appear on leaf margins and a slight thickening of the veins occurs, similarly to calcium and boron deficiencies. Sometimes the morphology of the leaf is changed and a bladeless leaf appears, again similarly to calcium and boron deficiencies. Growth is also stunted and the bunch is very small or choked. Temporary sulfur

deficiency is very typical in tissue-culture plants during the first 2 months after planting, since they have no sulfur storage and relatively small root systems. The most rapid uptake of sulfur occurs between the sucker and shooting stages. After shooting, uptake rate is reduced and sulfur needed for fruit growth comes from the leaves and pseudostem (Walmsley and Twyford, 1976). Most sulfur is supplied to banana as ammonium sulfate ((NH4)2SO4), potassium sulfate (K2SO4) or superphosphate (Ca(H2PO4) + CaSO4). In the Windward Islands, 127 kg of sulfur/ha is needed to establish the plant crop (Walmsley and Twyford, 1976). Subsequent losses in fruit removal amount to 23 kg/ha/year, which is about three times the losses in plantain (Lahav, 1995). Nitrogen–phosphorus–potassium (NPK) fertilizer containing 4% sulfur is used to avoid yield losses caused by sulfur deficiency (Messing, 1971). Sulfur-coated urea is also considered a good source (Jaramillo, 1976). Regular applications of sulfur, amounting to 50 kg/ha/year, are recommended to avoid deficiencies (Marchal et al., 1972). If sulfur-containing fertilizers, such as ammonium sulfate, potassium sulfate and superphosphate, are not used, sulfur should be applied separately.

Manganese Deficiency

Plate 9.8.  Yellowish young leaves due to sulfur deficiency in banana in Taiwan (photo: E. Lahav, VI).

Jordine (1962) reported manganese (Mn) deficiency in banana in Jamaica. Later, Charpentier and Martin-Prevel (1965) investigated the effects of deficiency and excess of manganese in sand culture in Côte d’Ivoire. Though found naturally in many African peat soils, manganese deficiency can be artificially induced by adding excessive amounts of lime. The characteristic feature of manganese deficiency is a ‘tooth-comb’ chlorosis in leaves. The fungus Corynespora torulosa is usually found in these chlorotic areas. The chlorosis starts marginally on the second or third youngest leaf. Sometimes, a narrow green edge is left at the leaf margins. Chlorosis then spreads along the main veins, towards the midrib, with interveinal areas remaining, green giving a ‘tooth-comb’ appearance. While normal-sized bunches are produced by deficient plants (at least in the plant crop), the fruit is covered with black spots. Poor

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fruit development is partly associated with the premature desiccation of the leaves. There is evidence that, under salt stress, high levels of manganese are found in the leaf margins. Manganese deficiency can be corrected by foliar sprays of 2% solution or by the ground application of manganese sulfate (MnSO4) (Jordine, 1962). A tentative annual rate of 7–11 kg manganese/ha has been suggested (Twyford and Walmsley, 1968; Marchal and Martin-Prevel, 1971).

Iron Deficiency Iron (Fe) deficiency symptoms associated with lime-induced chlorosis have been reported in the field in Hawaii (Cooil and Shoji, 1953), Jamaica (Charpentier and Martin-Prevel, 1968) and Israel (Ziv, 1954). It is also known in the Caribbean, where banana is grown on soils originating from coral reef mother rock. The most common symptom is interveinal chlorosis. In Israel, this symptom is expressed more intensively in the spring, since the soil pH is higher at that time, the roots are less active and the leaves are starting to develop. Later, the entire lamina becomes chlorotic (Plate 9.9). Leaves may become yellow-­ white, with an iron level as low as 3.4 ppm (­Cooil and Shoji, 1953). In Hawaii, chlorotic plants flowered when still small, producing small bunches or no bunches at all. Four applications of an iron sulfate (FeSO4) spray at weekly intervals controlled the chlorosis for up to 8 weeks. Application of iron salts to the soil doubled the amount of growth compared with non-treated plants. Iron deficiency can be overcome with foliar sprays of 0.5% iron sulfate with surfactant or iron chelated with ethylenediamine-N, Nʹ-bis (2-hydroxyphenyl) acetic acid (C18H20N2O6). However, lime-induced chlorosis is better corrected by soil application of 5 kg of 6% iron chelate/ha/ year, which can be applied directly to the soil or through irrigation water.

Zinc Deficiency The most widely reported minor element deficiency of banana is that of zinc (Zn) (Moity, 1954;

Plate 9.9.  Severe chlorosis in an iron-deficient banana growing on calcareous soil in Western Galilee, Israel (photo: E. Lahav, VI).

Cardenosa-Barriga, 1962; Jordine, 1962). It has often been confused with virus infection (Freiberg, 1966). Zinc deficiency is more common on naturally high-pH soils or on highly limed soils, because zinc ions in the chelate complex can be replaced by calcium ions. Zinc availability may also be restricted in organic and peat soils. In Carnarvon, Western Australia, yields declined from more than 60 t/ha at pH 4.5 to about 30 t/ ha at pH 8.7. This was thought to be associated with the supply of zinc to the plants, and leaf analysis data supported this interpretation (Turner et al., 1989). Symptoms of zinc deficiency can be associated with water logging, poorly drained soils, or compacted soils. While the plant expresses zinc deficiency symptoms, the problem of the soil condition, either drainage or compaction, needs to be addressed for zinc applications to be effective. Zinc is moderately redistributed from old to young leaves (van der Vorm and van Diest, 1982). The characteristic deficiency symptoms



Mineral Deficiencies of Banana 483

appear in young leaves, which become significantly smaller in size and more lanceolate in shape (Jordine, 1962). The emerging leaf also has a high amount of anthocyanin pigmentation on its underside (Plate 9.10), which often disappears as the leaf unfurls. The unfurled leaf has alternating chlorotic and green bands. The fruit is sometimes twisted, short and thin, with a  light green colour. As with sulfur deficiency, young tissue-culture plants also have no zinc storage and therefore show zinc deficiency symptoms immediately after planting. Zinc deficiency symptoms may appear without any apparent reduction in growth or yield, but, if the deficiency persists, plants of the next cycle are stunted (Charpentier and Martin-Prevel, 1965). Rayadilla disease (Cardenosa-Barriga, 1962), now attributed to zinc deficiency, was first reported in 1944 in Colombia on plantains in the Cauca Valley and ‘Gros Michel’ (AAA) in the ­Sevilla area. In the Sevilla area, the condition is also called lengua de vaca (cow’s tongue), because of the narrow strap-shaped leaves. Rayadilla is seasonal in appearance, being most common during dry seasons and tending to disappear during rainy months. It is worst west of the Orihueca and Rio Frio districts, where high pH and salinity problems may occur during the dry season, when plants are irrigated (Colmet-Daage and Gautheyrou, 1968). Plants have a stunted appearance, with erect, narrow leaves and small bunches. The bunch may be horizontal or partially parallel to the ground, rather than hanging perpendicularly. There is a general, uneven chlorosis, with stripes of yellow to white tissue alternating with green in young leaves (Plate 9.11).

This tissue may become necrotic as the leaves mature. The narrow strap-shaped leaves may have a thickening over the veins, resulting in upward curling. These leaves become fragile and brittle. Plants approaching maturity are most affected and suckers may appear normal until they are 150 or 180 cm high, when chlorotic mottling and rosetting symptoms appear. Normal leaves may follow chlorotic, stunted leaves, or the disease symptoms may intensify with each emerging leaf. Because of the symptoms, a virus disease was originally suspected, but attempts at transmission with six species of aphids were unsuccessful (Cardenosa-Barriga, 1962). Application of zinc to chlorotic foliage resulted in the resumption of normal growth and pigmentation. Symptoms of zinc deficiency in Jamaica were very similar to those of rayadilla in Colombia (Jordine, 1962). It also seems highly likely that ‘alkali chlorosis’, reported in Haiti, is a zinc deficiency disorder (Wardlaw, 1938).

Plate 9.10.  Anthocyanin pigmentation on the underside of young banana leaf as a result of zinc deficiency (photo: E. Lahav, VI).

Plate 9.11.  Symptoms of zinc deficiency in mature banana plants. Note the chlorotic stripes along the secondary veins. (Photo: E. Lahav, VI.)

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The zinc requirement of banana is relatively low. On the basis of whole-plant analysis, Twyford and Walmsley (1968) suggested a rate of about 1.0 kg of zinc/ha/year. Field experiments on acid soils in Australia showed that double and triple this amount was required to correct the deficiency, while in Central America and the Philippines up to 20 kg/ha is applied. Since zinc is leached under acid conditions and fixed under alkaline conditions, it is easy to correct its deficiency with a spray of 0.5% zinc sulfate (ZnSO4) (Jordine, 1962). The amount of zinc applied can be reduced to 5 kg/ha if zinc sulfate is incorporated with some sprays to counter black leaf streak disease. Work in Western Australia has shown that foliar applications need to be applied every 3 months to maintain zinc concentration in the leaves above a critical level (Turner et al., 1989). The long-term solution seems to be the regular addition of zinc to the soil. Zinc chelates are also sometimes applied.

Boron Deficiency Boron (B) deficiency has been recorded in the field in Ecuador (Tollenaar, 1969) and in sand culture (Charpentier and Martin-Prevel, 1965; Norton, 1965; Coke and Boland, 1971). Symptoms include reduced leaf area, curling and lamina deformation. However, the most characteristic symptom is chlorotic stripes at right angles to the veins on the underside of the leaf (Plate 9.12). New leaves may have an incomplete lamina, similar to that in sulfur and calcium deficiencies. Thickening of secondary veins and inhibition of root and flower formation have also been reported. All affected roots later darken and die. In Taiwan, a heart rot of plants derived from tissue culture has been attributed to boron and calcium deficiencies. Symptoms were a yellowing and necrosis of leaves, degeneration of roots and the development of a blackish cavity at the apical meristem of the corm before and after planting out in the field. Plants failed to develop symptoms when sprayed with either sodium borate (NaBO3) or calcium sulfate (CaSO4) (Ko et al., 1997). Charpentier and Martin-­Prevel (1968) pointed out that there is no clear distinction between sulfur and boron deficiency symptoms.

Plate 9.12.  Chlorotic stripes at right angles to the line of the leaf veins as a result of boron deficiency (photo: E. Lahav VI).

Soil concentration of boron should be in the range of 0.1–1.0 ppm (Walmsley and Twyford, 1976) and optimal level in the leaves 11–25 ppm. About 12 kg of borax/ha has been suggested as an amount that should overcome boron deficiency (Twyford and Walmsley, 1968). In India, 2 ppm of boron sprayed as boric acid (H3BO3) gave the best results (Srivastava, 1964).

Copper Deficiency The copper (Cu) requirements of banana are small, total copper uptake being about 1% of manganese uptake (Walmsley and Twyford, 1976). Copper is actively absorbed and readily translocated within the plant (van der Vorm and van Diest, 1982). Deficiencies have been described in pot culture (Charpentier and Martin-­ Prevel, 1965) and in the field in Côte d’Ivoire (Moity, 1961). Deficiency symptoms appear on all leaves. They are similar to those of nitrogen deficiency in that a general uniform paleness of the laminae occurs. Petioles are not pink, but the midrib bends, giving the plant an umbrella-like appearance. Yield is greatly reduced. Deficient plants have been reported as being sensitive to fungal and virus attacks (Moity, 1961). In acute cases, the rigidity of the funnel leaf is lost and it bends over the upper leaves during unfurling. It is preferable to correct copper deficiency with a foliar spray, and an application of up to



Mineral Deficiencies of Banana 485

0.5% solution of neutralized copper sulfate (CuSO4) has been advocated (Lahav, 1995). However, soil applications can also be used. Twyford and Walmsley (1968) suggested 1 kg of copper sulfate/ha as an annual rate for banana.

Srivastava (1964) increased the growth of suckers by applying a nutrient solution containing 4 ppm copper as copper sulfate to the soil. Copper oxide (CuO) and copper chelates can also be applied.

References Bhangoo, M.S., Altman, F.G. and Karon, M.L. (1962) Investigations on the Giant Cavendish banana. I. E ­ ffect of nitrogen, phosphorus and potassium on fruit yield in relation to nutrient content of soil and leaf tissue in Honduras. Tropical Agriculture (Trinidad) 39, 189–201. Butler, A.F. (1960) Fertilizer experiments with the Gros Michel banana. Tropical Agriculture (Trinidad) 37, 31–50. Cardenosa-Barriga, R. (1962) La ‘Rayadilla’ del platano en Colombia. Turrialba 12, 118–127. Chalker, F.C. and Turner, D.W. (1969) Magnesium deficiency in bananas. Agricultural Gazette of New South Wales 80, 474–476. Charpentier, J.M. and Martin-Prevel, P. (1965) Culture sur milieu artificiel. Carences atténuées ou temporaires en éléments majeurs, carence en oligo-éléments chez le bananier. Fruits 20, 521–557. Charpentier, J.M. and Martin-Prevel, P. (1968) Carences et troubles de la nutrition minéral chez le bananier. Guide de diagnostic pratique. Institut Français de Recherches Fruitières Outre-Mer, Paris, 75 pp. Coke, L. and Boland, D.E. (1971) Boron nutrition of banana suckers. In: Proceedings of the 2nd ACORBAT Conference, Kingston, Jamaica, 12-16 July 1971, pp. 59–66. Colmet-Daage, F. and Gautheyrou, J.M. (1968) Etude préliminaire des sols de la région bananière de Santa Marta (Colombie). Fruits 23, 21–30. Cooil, B.J. and Shoji, K. (1953) Studies reduce banana chlorosis. Hawaii Farming and Science 1, 1–8. Fox, R.L. (1989) Detecting mineral deficiencies in tropical and temperate crops. In: Plucknett, D.L. and Sprague, H.B. (eds) Westview Tropical Series 7. Westview Press, Boulder, Colorado, pp. 337–353. Freiberg, S.R. (1966) Banana nutrition. In: Childers, N.F. (ed.) Fruit Nutrition. Horticultural Publications, New Brunswick, New Jersey, pp. 77–100. Hasselo, H.N. (1961) Premature yellowing of Lacatan bananas. Tropical Agriculture (Trinidad) 38, 29–34. Jaramillo, R. (1976) Efecto de urea y de urea-azufre en la produccion de banano ‘Giant Cavendish’ en Guapiles, Costa Rica. Turrialba 26, 90–95. Jordine, C.G. (1962) Metal deficiencies in bananas. Nature 194, 1160–1163. Ko, W.H., Chen, S.P., Chao, C.P. and Hwang, S.C. (1997) Etiology and control of heart rot of banana t­issue culture plantlets. Plant Pathology Bulletin (Taiwan) 6, 31–36. Lacoeuilhe, J.J. and Godefroy, J. (1971) Un cas de carence en phosphore en bananeraie. Fruits 26, 659– 662. Lacoeuilhe, J.J. and Martin-Prevel, P. (1971) Culture sur milieu artificiel. 1. Carences en N, P, S chez le bananier, analyse foliaire. 2. Carence en K, Ca, Mg chez le bananier, analyse foliaire. Fruits 26, 161–167, 243–253. Lahav, E. (1972) Effect of different amounts of potassium on the growth of the banana. Tropical Agriculture (Trinidad) 49, 321–335. Lahav, E. (1974) The influence of potassium on the content of macro-elements in the banana sucker. Agrochimica 18, 194–204. Lahav, E. (1995) Banana nutrition. In: Gowen, S. (ed.) Bananas and Plantains. Chapman and Hall, London, pp. 258–315. Lahav, E. and Turner, D.W. (1983) Banana Nutrition. Bulletin 7, International Potash Institute, Worblaufen-­ Bern, Switzerland, 62 pp. Lin, M.L. and Fox, R.L. (1987) External and internal P requirements of mycorrhizal and non-mycorrhizal banana plants. Journal Plant Nutrition 10, 1341–1348. Lopez, A.M. and Espinosa, J.M. (1995) Manual de Nutricion y Fertilizacion del Banano. Instituto de la P ­ otasa y el Fosforo, Quito, Ecuador, 82 pp. Marchal, J. and Martin-Prevel, P. (1971) Les oligo-elements Cu, Fe, Mn, Zn dans le bananier – niveaux foliaires et bilans. Fruits 26, 483–500.

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Marchal, J., Martin-Prevel, P. and Melin, P. (1972) Le soufre et le bananier. Fruits 27, 167–177. Martin-Prevel, P. and Charpentier, J.M. (1963) Culture sur milieu artificiel. Symptomes de carences en six éléments minéraux chez le bananier. Fruits 18, 221–247. Martin-Prevel, P. and Montagut, G. (1966) Essais sol-plante sur bananiers. 8. Dynamique de l’azote dans la croissance et le développement du végétal. 9. Fonctions de divers organes dans l’assimilation de P, K, Ca, Mg. Fruits 21, 283–294, 395–416. Messing, J.H.L. (1971) Response to sulphur in Windward Island soils. In: Proceedings of the 2nd ACORBAT Conference, Kingston, Jamaica., 12–16 July 1971, pp. 51–58. Messing, J.H.L. (1974) Long term changes in potassium, magnesium and calcium content of banana plants and soils in the Windward Islands. Tropical Agriculture (Trinidad) 51, 154–160. Moity, M. (1954) La carence en zinc sur le bananier. Fruits 9, 354. Moity, M. (1961) La carence en cuivre des ‘tourbières du Nieky’ (Côte d’Ivoire). Fruits 16, 399–401. Murray, D.B. (1959) Deficiency symptoms of the major elements in the banana. Tropical Agriculture (Trinidad) 36, 100–107. Murray, D.B. (1960) The effect of deficiencies of the major nutrients on growth and leaf analysis of the banana. Tropical Agriculture (Trinidad) 37, 97–106. Norton, K.R. (1965) Boron deficiency in bananas. Tropical Agriculture (Trinidad) 42, 361–365. Srivastava, R.P. (1964) Effect of microelements Cu, Zn, Mo, B and Mn on the growth characteristics of banana. Science and Culture 30, 352–355. Stover, R.H. and Simmonds, N.W. (1987) Bananas. Longman Scientific and Technical, Harlow, UK, 468 pp. Tai, E.A. (1959) Annual Report, 1956–57, 1957–58. Banana Board Research and Development, Kingston, Jamaica. Tollenaar, D. (1969) Boron deficiency in sugarcane, oil palm and other monocotyledons on volcanic soils of Ecuador. Netherlands Journal of Agricultural Science 17, 81–91. Turner, D.W. and Bull, J.H. (1970) Some fertilizer problems with bananas. Agricultural Gazette of New South Wales 81, 365–367. Turner, D.W., Korawis, C. and Robson, A.D. (1989) Soil analysis and its relationship with leaf analysis and banana yield with special reference to a study of Carnarvon, Western Australia. Fruits 44, 193–203. Twyford, I.T. and Walmsley, D. (1968) The status of some micronutrients in healthy Robusta banana plants. Tropical Agriculture (Trinidad) 45, 307–315. van der Vorm, P.D.J. and van Diest, A. (1982) Redistribution of nutritive elements in a ‘Gros Michel’ banana plant. Netherland Journal of Agricultural Science 30, 286–296. Walmsley, D. and Twyford, I.T. (1968) The uptake of 32P by ‘Robusta’ banana. Tropical Agriculture (Trinidad) 45, 223–228. Walmsley, D. and Twyford, I.T. (1976) The mineral composition of the banana plant. 5, Sulphur, iron, manganese, boron, zinc, copper, sodium and aluminum. Plant and Soil 45, 595–611. Wardlaw, C.W. (1938) Banana diseases. 12. Diseases of the banana in Haiti, with special reference to a condition described as ‘plant failure’. Tropical Agriculture (Trinidad) 15, 276–282. Warner, R.M. and Fox, R.L. (1977) Nitrogen and potassium nutrition of the Giant Cavendish banana in Hawaii. Journal of the American Society for Horticultural Science 102, 739–743. Ziv, D. (1954) Chlorosis of bananas and other plants in the Jordan Valley due to iron deficiency. Hassadeh 35, 190–193 (in Hebrew).

10 

Injuries to Banana Caused by Adverse Climate and Extreme Weather Y. Israeli and E. Lahav

Introduction Banana is a tropical crop that grows and yields best in regions where the temperature, humidity and rainfall are optimal for normal crop development. However, banana is sometimes grown in areas, such as those found in the subtropics, where climates are suboptimal for good growth for certain periods of the year. Here, adverse temperature conditions extend the period it takes to produce a crop and can give rise to fruit quality disorders. In addition, extreme weather events, such as strong winds and hail, cause injuries that lead to yield reductions. Tropical storms with or without excessive rain are common problems for banana growers as well as long periods of drought. Banana cultivation at high elevations (up to altitudes of 2200 m) also exposes plants to specific limiting environmental factors.

Seasonal Wind and Windstorms Banana leaves are extremely sensitive to tearing by wind because of their wide thin lamina and parallel secondary veins. Skutch (1927) stated that leaf tearing is a normal phenomenon in banana. Leaf tearing might have some advantage in reducing boundary layer resistance, increasing the transpiration rate and maintaining the leaf energy balance (Taylor and Sexton, 1972).

Light wind at a velocity of 15 km/h is assumed to have a positive effect on the leaf energy balance (Turner, 1994). However, it was proved that intensive leaf tearing reduces the rate of growth and decreases yield. Ziv (1962) found that bunch weight was reduced by 17% after leaves were torn from the edge of the lamina to the petiole every 30–50 mm and Peretz (1992) measured a 10% yield reduction after tearing leaves into strips 50 mm wide. When leaves were torn into strips 12.5 mm wide, bunch weight decreased by 17.9% (Eckstein et  al., 1996). The negative effect of leaf tearing is explained by a decrease in photosynthesis resulting from a decrease in radiation interception by the torn lamina pieces. In addition to leaf tearing, wind increases leaf water loss, which may result in water stress, stomatal closure (Shmueli, 1953) and reduced photosynthesis. Seasonal winds in Israel can be strong enough to totally shred the lamina (Plate 10.1) or even fold leaves, especially in the upper part of the pseudostem. Wind can also cause fruit damage by rubbing fingers against leaves or bunch covers (Plate 10.2). Wind-carried dust, sand or salt particles may also collide with the fruit, causing bruises. In many banana-growing areas, especially in the subtropics, banana is artificially protected from the wind. In the Canary Islands, banana is commonly grown behind windbreaks made of bricks or sheltered in greenhouses (Galán-Saúco, 1992). Artificial windbreaks (Plate 10.3) were widely used in Israel (Lorch, 1959; Ziv, 1962)

© D.R. Jones 2019. Handbook of Diseases of Banana, Abacá and Enset (D.R. Jones)

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Plate 10.1.  Wind damage to banana in Israel. The leaves on some plants have been shredded until only the midribs remain (photo: Y. Israeli, JVBES).

Plate 10.2.  Abrasion damage to fruit of ‘Robusta’ (AAA, Cavendish subgroup) in St Vincent, Windward Islands. The line scars are caused by the polyethylene bunch cover rubbing on bananas, usually the finger ridges, in the wind (photo: D.R. Jones, SVBGA).

and in New South Wales, Australia (Freeman, 1973). Closely planted trees are often employed to block strong winds: Casuarina is used in Egypt, Cupressus in Israel (Plate 10.4), Erythrina in Martinique and Guadeloupe and pines in Queensland, Australia (Daniells, 1984). A significant

increase in the rate of growth and up to a 25% increase in bunch weight have been recorded in Israel in plants protected from the afternoon breeze in midsummer (40 km/h, 9 h/day) (Ziv, 1962; Peretz, 1992; Eckstein, 1994). On the downside, living windbreaks may shade nearby plants and thus reduce yield (Eckstein, 1994), compete with the banana plants for water and nutrients (Ziv, 1962) or host various pests and diseases (Krambias et  al., 1973). The cost of windbreaks (in terms of productive farmland occupied) and greenhouses (in terms of maintenance) might also be high, but improved fruit quality is a major factor in their adoption. Since the late 1970s, protected banana cultivation has been gradually developed in the subtropics. It began with cultivation in plastic greenhouses in Morocco and in the Canary Islands (Galán-Saúco et al., 1992; Cabrera Cabrera and Galán-Saúco, 2012) (Plate 10.5). In Turkey, more than half of the total area of land under banana was in greenhouses (Gubbuk, 2010). In Israel, where greenhouses are unsuitable because of the summer heat, about 90% of the 2500 ha growing banana were covered by screenhouses



Injuries to Banana Caused by Adverse Climate and Weather

Plate 10.3.  Artificial windbreak for banana in Israel (photo: E. Lahav, VI).

Plate 10.4.  Cupressus trees used as a windbreak for banana in Israel (photo: Y. Israeli, JVBES).

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Plate 10.5.  Plastichouses in the Canary Islands (photo: Y. Israeli, JVBES).

in 2016. When new, screens transmit 85–88% of the photosynthetic active radiation. Light transmission is reduced to about 75% during the dusty summer, but recovers after winter rains (Israeli et al., 2002) (Plate 10.6). Greenhouses or screenhouses protect plants by decreasing wind damage, decreasing excessive solar radiation (especially at midsummer), increasing the relative humidity around the plants, elevating to some extent autumn and spring temperatures and giving partial protection against hail damage. Greenhouses also protect from salt, which affects plants growing close to the coast. Leaves are also untorn, with no shredding, which partially contributes to enhanced photosynthesis. The improved conditions increase the rate of growth, especially of the plant crop, and final plant size. In the longer term, bunch weight increases, fruit quality improves and losses associated with field cultivation are reduced, which results in more marketable fruit. Other advantages are that seasonal changes in the pattern of fruit production are less noticeable and water use is much more efficient. Some variation

has been recorded in regard to long-term density and production cycle data, possibly due to differences in management of the dense foliage and the plant population. Protected cultivation technology has been thoroughly studied (Galán­Saúco et al., 1992; Eckstein et al., 1998; G ­ ubbuk and Pekmezci, 2004; Möller et al., 2010; Cabrera Cabrera and Galán-Saúco, 2012; Tanny et  al., 2012, 2014; Pirkner et al., 2014; Haijun et al., 2015) and Robinson and Galán-­Saúco (2010) summarized in detail the advantages and ­limitations. Tropical storms above 50 km/h, such as hurricanes in South America and the Caribbean and cyclones in Southeast Asia and Australia, can cause total devastation by breaking or uprooting banana plants over large areas (Plate 10.7). Heavy rains and floods can also accompany strong winds. Toppling of banana plants occurs more often in wet soil, especially when roots and corms are damaged by nematodes or weevil borer (Gowen, 1993). Damage recovery after uprooting is slow, because of a delay in the emergence of following suckers (Robinson and



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Plate 10.6.  Screenhouse in the Jordan Valley, Israel (photo: Y. Israeli, JVBES).

Plate 10.7.  Cyclone damage in Queensland, Australia (photo: J.W. Daniells, QDAF).

Galán-Saúco, 2010), and quite often replanting is required. The location of the majority of commercial export industries have been deliberately chosen

so that they lie outside areas where hurricanes and cyclones occur. Thus, commercial banana plantations in the Philippines have been sited on the island of Mindanao, which was considered

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cyclone-free, unlike the island of Luzon, where 19 cyclones are recorded annually (Valmayor, 1990). Recently, however, and possibly as a result of global warming and climate change, typhoons seem to be traversing the Philippines further south than usual. One that struck the main banana area of Mindanao in 2012 destroyed about 10,000 ha. Typhoons in the Philippines have also resulted in the spread of the Fusarium wilt pathogen that attacks Cavendish cultivars in plantation flood waters (Plate 10.8). A method of planting high-density, single-­ cycle plantations was developed in Taiwan in ­order to avoid the typhoon season (Tang and Liu, 1993). Planting occurs about 1-2 months before the season so that only young suckers, which can withstand strong winds without critical damage, are present in the field during the period when a typhoon may strike. In Australia in 2011, the foliage of mature test plants was either partially or completely removed just prior to a devastating cyclone in an attempt to determine if the practice minimized wind resistance and thus reduced damage. The method was shown to be successful and replanting was not needed as is usually the case after severe storms (Lindsay, 2016). Rate of toppling differs among cultivars (Ahmad and Quasem, 1975; Stover and Simmonds, 1987; Daniells and Bryde, 1993, 1995; Tang and Chu, 1993), as was demonstrated by Monnet (1961) (Table 10.1). The main factor involved is stature, but pseudostem circumference is also important. Leaf petioles in tetraploid cultivars are not as strong as in triploids and their leaves break more easily (Stover and Simmonds, 1987). Sarma et al. (1995) reported that the Musa balbisiana selection ‘Bhimkol’ (BBw), which is cultivated in Assam in India, although a tall clone, has a fibrous pseudostem, which makes it resistant to toppling. The relative wind tolerance of ‘Grand Nain’ in contrast to ‘Poyo’ (both AAA, Cavendish subgroup) was demonstrated in Côte d’Ivoire when the two cultivars grew side by side. ‘Grand Nain’, which is shorter than ‘Poyo’ by 50–70 cm and has a stronger pseudostem, suffered significantly less toppling (Y. Israeli, Israel, 2016, personal communication). In St Lucia, the taller Cavendish cultivar ‘Robusta’ suffered 2.5 times more wind damage than the shorter ‘Williams’ (Holder and Gumbs, 1983a). ‘Goldfinger’ (AAAB) in Australia, although

Plate 10.8.  Yellowing plants show the spread of Fusarium wilt in a Cavendish plantation after floods in the Philippines (photo: R. Barnett, Philippines). Table 10.1.  The effect of mature plant height on blow-down losses and yield in six different Cavendish cultivars (AAA) and ‘Gros Michel’ (AAA) during four crop cycles in Guinea (after Monnet, 1961). Cultivar ‘Dwarf Cavendish’ ‘Grand Nain’ ‘Seredou’ ‘Poyo’ ‘Maneah’ ‘Lacatan’ ‘Gros Michel’

Average height (cm)

Blow-down losses (%)

Yield (t/ha)

180

3.3

82.0

210 260 280 290 320 360

2.4 20.1 35.3 57.5 60.2 71.6

89.1 60.9 23.1 40.0 19.6 17.4

taller than ‘Grand Nain’ or ‘Williams’, has a larger and stronger pseudostem. Consequently, it suffers less from strong winds (Daniells et al., 1995). The effect of strong wind on three cultivars of different heights is illustrated in Plate 10.9. It can be concluded that cultivar selection and timing of planting to avoid adverse seasonal weather (Obiefuna, 1986) contribute to decreased wind damage.

Low Temperature The response of the banana to temperature has been thoroughly reviewed (Stover and Simmonds, 1987; Turner, 1994, 1995, 1998; Turner et al., 2007; Robinson and Galán-Saúco, 2010). This



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Plate 10.9.  Effect of plant height on losses from wind. The devastated plot in the centre contains the tall Cavendish cultivar ‘Lacatan’ (360 cm in height). Much less damage has been caused to the smaller Cavendish cultivar ‘Valery’ (290 cm in height) in the plot to the right and to ‘Cocos’, a dwarf mutant of ‘Gros Michel’, in the plot to the left (photo: R.H. Stover).

review will focus on the effect of low winter temperatures, chilling and frost on the banana plant. Minimum temperature for meristem activity and new leaf formation is 9–11°C (Green and Kuhne, 1969; Ganry, 1973), but almost all new leaves emerge only when the ambient temperature is above 14–16°C (Robinson and Galán-Saúco, 2010). Leaf emergence rates are optimal between 27°C and 32°C (Ganry, 1973; Turner and Lahav, 1983; Allen et  al., 1988). The banana grows best between 22°C and 31°C. The optimum temperature range for carbon dioxide (CO2) assimilation by the banana leaf has been found to be 24–36°C, with maximums recorded at 31–32°C (Israeli, 2000), 29.4°C (Thomas et al., 1998) and 32.5°C (Eckstein and Robinson (1995b). The minimum temperature for root growth is about 12°C (Robinson and Alberts, 1989). The optimum calculated by Turner (1995) is 24°C, which is somewhat lower than the optimum of 26.5°C found by Israeli et al. (1979) under controlled conditions. The lowest critical temperature for fruit growth is between 11°C (Green and Kuhne,

1975) and 14.5° (Ganry and Meyer, 1975). Turner and Barkus (1982) showed that relative fruit growth rate increases linearly between 13.3°C and 21.6°C. In Israel, fruit growth continues during winter when daily average temperatures are as low as 13–14°C and leaf emergence has almost totally stopped (Israeli and Lahav, 1986). It seems that, under such conditions, the fruit grows during daytime when temperatures are higher. Leaves do not emerge, because the daily temperature fluctuations in the meristem zone are small. The optimum temperature for fruit growth is 30°C (Hord and Spell, 1962; Ganry and Meyer, 1975), which is close to the optimum leaf CO2 assimilation rate (Israeli, 2000). The relationship between temperature and rate of fruit growth enables harvest dates to be predicted, but the interaction of many additional factors (water availability, nutrient supply, amount of light, strength of winds, cultivar factors, pest and disease incidence) makes the use of such models very difficult (Turner, 1995). The relationship between temperature and other climatic factors and the banana rate of growth enables expected flowering time to be

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predicted (Cottin et al., 1987; Israeli et al., 1988). Fortescue et al. (2011) identified a facultative response of banana to long day photoperiod. Flowering to harvest time is affected significantly by temperature. Under tropical conditions, annual variation of 30 days can be expected (Stover and Simmonds, 1987). Fruit-filling periods increased from 90 to 120 days when average temperatures decreased from 29°C to 23°C (Hord and Spell, 1962). Temperatures below 20°C are quite rare in tropical lowlands, though every 5–8 years a cold spell may reduce temperatures to 11°C and cause chilling damage in the growing areas of Mexico, Honduras or Guatemala (Stover and Simmonds, 1987). Long fruit-filling periods resulting from low temperatures are also related to increased elevation (Daudin, 1955; Sanchez-­ Nieva et al., 1970; Galán-Saúco et al., 1984). In the Philippine highlands at an elevation of 600– 1000 m, where night temperature may decrease as low as 14°C, fruit-filling time is increased by 3–5 weeks compared with the lowlands. As a consequence, the shoot-to-shoot cycle is also longer. Temperatures within the plantation can be increased by eliminating suckers, which reduces shade and allows more sunlight to penetrate. The plant population is smaller, but advantages, such as heavier bunch weight, bigger fruit size and higher fruit sugar levels at ripening compensate. Plantain in Africa is sometimes grown at even higher elevation. At 1066 m, agronomic results are still good, but a decline was noted at an increased planting elevation with an associated decrease in prevailing temperatures (Sivirihauma et al., 2016). Low winter temperatures in the subtropics significantly increase the period from flowering to harvest. In Israel, Cavendish bunches emerging in the spring are harvested after 80 days, while those emerging in the autumn may need as long as 240 days to mature (Oppenheimer, 1960; Israeli and Lahav, 1986). In South Africa, this variation is between 118 and 213 days for ‘Williams’ (AAA, Cavendish subgroup) (Robinson, 1981). The first indication of low-temperature damage, observed at 17/10°C day/night temperatures, is shortening of the distance between petioles of recently opened leaves. This symptom is referred to as ‘choking’. Choking is the result of a disturbance in normal leaf emergence and is caused not only by low temperatures, but also

by other environmental stress conditions. Leaves formed at low temperature may also remain erect and have narrower laminae (Porteres, 1950; Prest and Smith, 1960; Cann, 1964; Turner and Lahav, 1983). Low-temperature induced choking has also been recorded on plantain growing at high elevation in Central Africa (Sivirihauma et al., 2016). One of the most significant symptoms of low winter temperature damage is ‘choke throat’, a condition that is characterized by only partial emergence of the inflorescence from the pseudostem. The basal hands that are trapped in the trunk do not open normally and the whole bunch is positioned almost horizontally (Plate 10.10). ‘Choke throat’ results from the crowding of petioles at the top of the pseudostem and the loss of elasticity of leaf sheaths during winter when growth ceases. In extreme cases, the bunch is completely trapped inside the pseudostem (Plate 10.11) or emerges through the side of the pseudostem (Plate 10.12). Such bunches were differentiated in autumn and are ‘choked’ during the end of the winter or early spring (Oppenheimer, 1960). The condition has been described in many subtropical growing

Plate 10.10.  ‘Choke throat’ – partial emergence of the banana bunch (photo: Y. Israeli, JVBES).



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and ‘Grand Nain’. ‘Williams’ is also affected in South Africa (Robinson and Galán-Saúco, 2010). In Taiwan, ‘Grand Nain’ suffers from ‘choke throat’ while the taller local cultivar ‘Pei-Chiao’ suffers less (Tang and Chu, 1993). ‘May flowering’ in the northern hemisphere (or ‘November dump’ in the southern hemisphere) is another condition associated with low temperatures during floral differentiation (Summerville, 1944; Oppenheimer, 1960; Fahn et al., 1961; Ziv, 1962; Cann, 1964; Blake and Peacock, 1966; Kuhne et al., 1973; Israeli and Blumenfeld, 1985; Robinson, 1993; Turner, 1995; Robinson and Galán-Saúco, 2010). It is interesting to note that some bunch symptoms similar to ‘May flowering’ and ‘November dump’ appear all year round at high altitudes (Sivirihauma et al., 2016). The main symptoms of this disorder are as follows:

• •

Plate 10.11.  Banana bunch trapped inside the pseudostem as a result of rosetting and loss of elasticity of the upper leaf petioles caused by low temperatures (photo: Y. Israeli, JVBES).

regions (Stoler, 1960; Ziv, 1962; Kuhne et  al., 1973; Robinson, 1993) and ‘Dwarf Cavendish’ (AAA, Cavendish subgroup) is especially affected. This, amongst other factors, has led to a decline in the use of ‘Dwarf Cavendish’ in subtropical areas and its replacement by less susceptible Cavendish cultivars, such as ‘Williams’ or ‘Grand Nain’. Winter or early spring flowering is often followed by a high percentage of female flowers with a persistent perianth (Plate 10.13) with the residues of many male flowers persisting on the axis (Daudin, 1953; Israeli et  al., 1980; Israeli and Blumenfeld, 1985). Within the Cavendish subgroup, the sensitivity to low temperatures generally decreases as plant height increases and as the peduncle above the basal hand becomes longer (Kuhne, 1980a; Hill et al., 1992). Although the tendency to choking decreases with height, experience gained in Israel shows that low temperatures can induce choking even in the relatively taller ‘Williams’

•

• • • •

Fewer than five anthers in a flower. Fewer than three loculi in an ovary. Sometimes the loculi are totally missing. Flowers with reduced number of loculi develop small fruit scattered irregularly in the hand. Enlargement of the style base due to unification of several filaments. Sometimes the loculi penetrate into the style base. Such thickening forms a nipple at the tip of the finger (Plate 10.14). Nectary longer than normal. Fingers short and thick with a round cross-section and straight growth. The distal part of the finger is often larger than the proximal part. Variability in finger size within a single hand (Plate 10.15). A reduction in the number of hands to up to half the usual number (Plate 10.16).

In Israel, abnormal flowers appear during a period of 3–4 weeks on bunches emerging in late May or early June. If one assumes that there are 11 leaves hidden in the pseudostem during floral initiation (Summerville, 1944) and at the end of initiation there are still nine leaves left to emerge (Ziv, 1962), then these abnormal flowers would appear to be formed in bunches differentiated in late autumn or winter when temperatures are low. Further evidence for low temperatures being the cause of the condition comes from studying the location of the abnormal flowers on the bunch over the ‘May flowering’ period.

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Plate 10.12.  A banana bunch that has emerged through the side of the pseudostem (photo: Y. Israeli, JVBES).

At the beginning of the period, abnormal flowers are located mainly on the distal hands of emerging bunches while the basal hands are more affected on bunches emerging at the end of the period. As the basal hands complete their differentiation earlier than the distal hands, it is believed that they are not affected at the beginning

of the period, because they are formed before temperatures decrease, whereas the distal hands are formed when conditions are colder. Conversely, at the end of the period, it is the basal hands that are differentiated during the colder weather, whereas the distal hands are formed later, when spring temperatures are increasing



Injuries to Banana Caused by Adverse Climate and Weather

Plate 10.13.  Banana inflorescence that emerged in the winter with persistent perianths (photo: Y. Israeli, JVBES).

Plate 10.14.  Typical malformations of female flowers in May flowering banana bunch (photo: Y. Israeli, JVBES).

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Plate 10.15.  Symptoms of May flowering: plump banana finger tips, variable finger size, mixed male and female flowers on the same hand, and persistent male flowers (photo: Y. Israeli, JVBES).

(Fahn et al., 1961). These symptoms may appear one at a time or simultaneously and their severity depends on the temperature. They are more frequent in ‘Dwarf Cavendish’, but also appear in ‘Williams’ and ‘Grand Nain’. The relationship between temperature at the apical meristem and quality of bunches was demonstrated in the Jordan Valley in Israel by Stoler (1962). Inflorescences differentiated in the spring, when meristem temperatures were 20–24°C, flowered in July–August and produced excellent bunches with many hands and normal fingers. Earlier-differentiated bunches, when temperatures were less than 18°C, contained few hands with abnormal flowers. High temperatures ranging from 26°C to 30°C at the zone of the apical meristem resulted in bunches with few hands, but normal fingers. The average daily air temperature during bunch differentiation is related to the number of hands formed on the bunch (Israeli, 1976). Numbers of hands increase

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Plate 10.16.  May flowering bunch with only three hands of bananas (photo: Y. Israeli, JVBES).

when the bunch is differentiated at temperatures between 18°C and 27°C, but decrease quickly when differentiation occurs at temperatures above 27°C. A special sucker pruning system, aimed at avoiding bunch differentiation during spring or autumn, was developed in Israel (Oppenheimer, 1960; Ziv, 1962; Aubert, 1971; Ticho, 1971). By using this method, the actual rate of abnormal bunches is significantly reduced. Low winter temperature effects are minimized in greenhouses or reduced in screenhouses. An increase in bunch weight is the main reason for greater production in greenhouses in the Canary Islands than in the field (Galán-Saúco et al., 1992), while reduced bunch loss is the main reason for increased production under screenhouses in Israel.

Chilling and Frost Low temperatures in tropical regions caused by cold fronts result mainly in the cessation of growth and a delay in fruit filling. Chilling damage is rare (Stover and Simmonds, 1987). However,

temperatures may drop much lower and chilling or frost damage may occur in subtropical regions. Chilling injury to the banana canopy takes place when temperatures drop to 5–8°C (Oppenheimer, 1960; Ticho, 1971; Kuhne and Green, 1980; Robinson, 1993; Turner, 1994). The effect of chilling temperatures on photosynthesis in warm-climate plants was described in detail by Allen and Ort (2001). Repeated exposure to low night temperatures (Eckstein and Robinson, 1995b) and strong solar radiation the following day (Smillie et al., 1988) results in photoinhibition, and leaf bleaching. The colour of leaves changes to pale green, yellow (Plate 10.17), orange and finally brown (Plate 10.18) because of the destruction of chlorophyll. Leaf necrosis takes place after 3–4 weeks. Chilling injury is a product of time and temperature and can occur after short exposure to temperatures of 2–3°C (Robinson, 1993) or a long exposure to temperatures of 5–8°C. Slight chilling injury might be reversible. A young leaf affected by chilling and showing some chlorophyll destruction may return to normal activity within a few weeks if not exposed to further chilling (Shmueli, 1960). It has been shown that high proportions of desaturated phosphatidyl-glycerol within lipids of the chloroplast membrane of a few plants, which includes banana, indicate a sensitivity to chilling (Roughan, 1985; Kenrick and Bishop, 1986). Some degree of chilling injury is a frequent event in high-elevation banana plantations in the subtropics and tropics, but is usually tolerated. The minimum temperature in Israel during January–February is about 8°C and production is still satisfactory – about 65–70 t/ha/year. However, frost is an event causing significant damage. Normally, banana cannot be grown where the frequency of frosts is high. Banana plants are sensitive to low temperatures because of their anatomical, morphological and physiological characteristics (Shmueli, 1960). They have large leaves, with large cells surrounded by large intercellular cavities. The leaves are highly hydrated, being very sensitive to any change in hydration level. The osmotic pressure of leaf sap is relatively low, while its freezing-point is high. Plants can experience freezing injury when there is a frost (Snyder and de Melo-Abreu, 2005). Frosts fall into two categories. An advection frost is associated with a large-scale incursion of cold



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Plate 10.17.  Severe chilling injury. The banana leaves will gradually dry out (photo: Y. Israeli, JVBES).

Plate 10.18.  Advanced stage of leaf chilling resulting in a loss of chlorophyll. Note necrosis of the banana leaf tissue next to the petiole because of direct exposure to the sky (photo: Y. Israeli, JVBES).

air with a well mixed, windy atmosphere and below-zero temperatures even during daytime. This type of frost is typical at high latitudes and is less common in banana-producing areas. Radiation frost, most common in the subtropics, is associated with cooling as a result of energy loss through radiant exchange during clear, calm and dry nights. In some cases, a combination of day-advection and night-radiation frost conditions may occur. Frost is said to have occurred when the air temperature near the soil surface or in a standard meteorological shelter at 2 m

above the ground drops below zero (Kalma et al., 1992; Snyder and de Melo-Abreu, 2005). The sky-exposed banana leaf, which loses energy and cools due to a loss of long-wave radiation, is the coldest part of the plant during a bright winter night with its temperature close to that near the soil surface on open ground. The water in the banana leaf ’s intercellular cavities starts to freeze when its temperature drops below 0°C (Gilead and Rosenan, 1957; Shmueli, 1960; Israeli and Chudi, 2015). Internal, protected plant parts have higher temperatures. Frost damage occurs when sap freezes. Leaves blacken during the days following the frost and this leads to complete leaf desiccation and necrosis (Plate 10.19). Israeli and Chudi (2015) and Israeli et  al. (2016) studied frost events in the Jordan Valley in Israel over a 54-year period. The amount of frost damage, which was recorded in 19 of those years, was a function of how low the temperature dropped, how long the temperature remained below a critical level during the night and how many consecutive frost-nights occurred during the frost event. Significantly higher damage was recorded when the frost arrived early in the winter following a period of high temperatures and intensive growth. Fruit was particularly sensitive during this time. The observations suggested that some degree of hardening or resistance to

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Plate 10.19.  Severe frost damage to banana in Western Galilee, Israel (photo: E. Lahav, VI).

damage was induced if there was a gradual decrease in temperatures over several weeks before the frost event. A second highly frost-sensitive period occurred in the spring when growth and intensive floral initiation intensity were resumed. At this time, there was no dense mother-plant foliage to shade and protect the followers. A frost at this stage could also have a detrimental effect on the yields obtained during the following season. In some cases, the effects were more severe in the growing season of following year (Israeli et al., 2016). Production was most affected after two consecutive frost years. Banana plantations affected by frost resulting in loss of canopy can still produce marketable fruit if the bunches are protected and have reached the half-full filling stage. Such fruit absorbs mostly water during its last stages of development and green leaves are not critical for bunch filling and maturation (Israeli and Lahav, 1986). However, bunches affected at a less advanced stage will not reach commercial requirements (Smith, 1973; Robinson, 1993). Frost damage on fruit appears as water-­ soaked areas on the fingers (Plate 10.20). In a frost-damaged plantation, the bunch is no longer

Plate 10.20.  Banana fingers with water-soaked areas after frost damage to cell membranes (photo: Y. Israeli, JVBES).

protected from the sun by leaves and symptoms of sunburn may appear on the peduncle and upper hands. Often, the peduncle snaps and the bunch falls to the ground. Plants that do not produce bunches before frost have been estimated to need four to six newly emerged leaves after frost to support normal bunch development (Kuhne and Green, 1980; Robinson, 1993), though ten leaves may be more realistic (Smith, 1973). Bunches that emerge soon after frost will



Injuries to Banana Caused by Adverse Climate and Weather

probably be choked and have abnormal flowers. Such bunches have no commercial value. Sensitivity to chilling and frost varies among cultivars and clones. Cultivars containing the B genome inherited from Musa balbisiana are more resistant than those based solely on the A (Musa acuminata) genome. Liang et al. (1994) tested the effect of chilling temperatures on potted AAA and ABB banana cultivars. The AAA banana clones were significantly injured while no damage was observed on the ABB clones. Peroxidase activity was initially much higher in leaves of ABB plants than in the AAA plants. It remained high in the ABB banana after exposure to chilling, but decreased in the AAA plants. Paclobutrazol (growth retardant) reduced the damage caused by low temperature. ‘FHIA-01’, which is also known as ‘Goldfinger’ (AAAB, bred hybrid), is relatively more frost-resistant than ‘Grand Nain’ (AAA, Cavendish subgroup) (Rowe and Rosales, 1993; Z.C. de Beer, South Africa, 1995, personal communication). Within the Cavendish subgroup, the clones that are relatively low in stature suffer more than the tall ones. However, their rate of growth and development is faster and they recover quickly. This height-­ related difference in the frost sensitivity of Cavendish clones was well demonstrated in Israel at locations where short cultivars, such as ‘Dwarf Cavendish’, grew next to taller ones, such as ‘Williams’ and ‘Grand Nain'. Frost damage can be reduced in several ways. The most important is the selection of a frost-free area for planting. Before planting in an area prone to frost, a topographical/climatic survey should be undertaken (MacHattie and Schnelle, 1974; Gat, 1984) to aid the selection of the best sites (Lomas et al., 1989). Land with heavy clay soils in locations where cold air cannot escape is prone to frost and should not be planted. Methods for predicting frost and passive or active measures to protect against frost have been reviewed for many crops (Bagdonas et al., 1978; Barak and Israeli, 1989; Kalma et  al., 1992; Snyder and de Melo-Abreu, 2005; Snyder et  al., 2005), including banana (Shmueli et  al., 1959; Oppenheimer, 1960). Favourable conditions for the development of radiation frost include initial a low air temperature, a clear sky, low air humidity and no wind. During the night, the soil loses heat through long-wave

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radiation. This radiation heats the plants above, which slows their cooling process. Therefore, keeping the soil surface clean of weeds or trash will help heat transfer (Shmueli et  al., 1959) and reduce frost damage to some extent. High-­ density plantations will be damaged less than low-density plantations because relatively less of their canopy is exposed to the sky. A dense canopy is especially important in protecting the lower young suckers. Pseudostems should not be cut down after harvest if frost risks still exists. The lower parts of vegetative windbreaks should be pruned, so that air can move freely over the soil surface. Frost damage may be actively reduced or prevented by using heaters or solid-fuel fires, overhead sprinklers to provide artificial rain, misters to provide artificial fog or wind machines or helicopters to mix the lower cool air layers with higher warmer layers. All these methods have been tested in banana plantations (Shmueli et  al., 1959; Oppenheimer, 1960; Barak and Israeli, 1989), but none are used commercially, because of low efficacy or economic considerations. The most efficient means for protection against environmental stresses, including chilling and frost, is to grow banana as protected cultivation (Plate 10.21). This technique is widely used in Morocco (Janick and Ait-Oubahau, 1989; Choukr-Allah, 1990), Canary Islands (Galán-Saúco et al., 1992) and southern Turkey (Cevik et  al., 1984) and to a smaller extent in Sardinia, Sicily, Crete and South Korea (Ahn et  al., 1990; Pala and Ovadia, 1990; Colombo et  al., 1993). Greenhouses also give protection against wind and excessive radiation. Fruit quality in greenhouses is much improved and the bananas produced can compete successfully in the market with those from the tropics. However, the cost and profit should be considered in each location. Greenhouses without active heating or without a special cover that minimizes outward long-wave radiation during the cold night and inward ultraviolet (UVA) radiation during the following day have a limited frost protective effect. Screenhouses covered with 10–15% shade crystal net only marginally affect the temperature during frost nights and visible foliage damage seem to be similar to that in uncovered plantations. However, the effect on the next year’s

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Plate 10.21.  Banana plants growing inside of a polyethylene-covered greenhouse used to protect bananas from frost in Tenerife, Canary Islands (photo: Y. Israeli, JVBES).

­ roduction is in favour of screenhouses. This difp ference is attributed to the higher vegetative strength of the screenhouse-grown plants and their better recovery (Israeli et al., 2016). The ready availability of in vitro planting material today makes the re-planting of frostdamaged plantations much easier than in the past,when growers needed to look for suckers or corms in frost-affected plantations.

Hail Stover and Simmonds (1987) stated that hail damage is a rare event in tropical lowlands, referring probably to the export banana producing zones, but hail is very frequent in tropical India where a quarter of the world’s bananas are produced, and in other Asian countries (Malhotra, 2013). Hail is typical of many subtropical growing regions and of certain tropical highlands. Hail can tear banana leaves to narrow shreds, which later turn brown and desiccate (Plate 10.22). Leaf shredding by hail is probably more damaging than leaf tearing by wind. Additional damage is

caused when hail accumulates in the funnel leaf or strikes the upper petioles and other exposed plant parts. Plant tissue in direct contact with ice freezes and desiccates within a few hours. Hail also damages fruit, the points of impact being characterized by sunken scars (Plate 10.23). In extreme cases, fingers can split (Kuhne and Green, 1980). Polyethylene sleeves (especially when more than 0.1 mm in thickness) give fruit partial protection (Kuhne and Green, 1980; Eckstein and Fraser, 1996). Greenhouses or screenhouses may protect the banana when the hail is not too heavy (Robinson and Galá-Saúco, 2010), but may collapse under the weight of the heavy hail. However, there are technical solutions to this problem. The salvage of hail-damaged plantations depends on the severity of the damage. Marketable bunches should be harvested immediately. Plants with no functional leaf area and severely damaged bunches should be cut to let the suckers grow. Undamaged bunches close to harvest should be left to complete their development with a minimal number of functional leaves. For plants that have not flowered, the damaged leaves should be removed in order to accelerate



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Plate 10.22.  Banana leaf shredding as a result of hail (photo: Y. Israeli, JVBES).

as Egypt, Israel, Jordan, Pakistan, Oman, northern India and Western Australia, if an adequate water supply is available. However, exceptionally high temperatures were recorded in some of these areas in 2015 and 2016. In Rajasthan in India, which is an important banana production area, 50.6°C was recorded on 20 May 2016. Banana leaf growth stops at an ambient temperature of 38–40°C (Stover and Simmonds, 1987; Robinson and Galán-Saúco, 2010) and photosynthesis decreases at temperatures above 35°C (Eckstein and Robinson, 1995b; Israeli, 2000). A study of the energy balance of the baPlate 10.23.  Hail damage to uncovered banana nana in tropical lowlands showed that a leaf in a fruit (photo: Y. Israeli, JVBES). horizontal position exposed to direct sunlight may reach 12°C above ambient temperature. new growth and development from reserves Under conditions of reduced transpiration, as a stored in the pseudostem. When four or more consequence of limited water-supply or after arnew leaves emerge after the hailstorm, a comtificial stomata closure using petroleum jelly, mercial bunch may be obtained. If flowering is temperatures rise about 20°C above air temperaexpected before a sufficient number of new ture (Stoutjesdijk, 1970). Taylor and Sexton leaves emerge, the plant should be sacrificed for (1972), working with Heliconia latispatha, rethe sake of a good follower (Kuhne and Green, corded a temperature of 46°C in a leaf exposed 1980; Robinson, 1993; Robinson and Galán-­ to direct sun during the dry season, but a temSaúco, 2010). perature of only 40°C in a leaf with torn lamina. They demonstrated in Strelizia nicolai, another relative of the banana, that a leaf exposed for a Heat short time to 47.5°C would turn necrotic a few days later. The authors concluded that, in plants Banana can usually withstand high tempera- in the Musaceae, tearing of the leaf lamina tures and grow in hot and semi-arid areas, such along the secondary veins is a natural process,

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reducing the boundary-layer resistance, increasing the transpiration rate and helping to decrease excess heat. The last unfurled leaf is the most sensitive to overheating (Plate 10.24). This is because the stomata have not begun to function and transpiration is limited (Eckstein and Robinson, 1995a; Israeli, 2000). The last expanded leaf shows signs of burns at an ambient temperature of 42–43°C even when the water-supply is plentiful (Robinson, 1993). The cigar or funnel leaf is also very sensitive to heat stress. Under high temperatures, it fails to unfurl normally and necrotic spots are observed on the side facing the sun (Plate 10.25). This happens in the plantation, inside the screenhouse and even with potted plantlets in 50% shaded nurseries (Plate 10.26). The cigar leaf is most affected by heat during periods of fast growth when there is insufficient time for hardening. Under these conditions, it is often distorted and sometimes bends horizontally. Plants close to bunch emergence,

Plate 10.24.  Heat stress damage to a young banana leaf in the Jordan Valley, Israel (photo: Y. Israeli, JVBES).

which is a crucial moment in their development (Lassoudiere, 1978), are especially vulnerable to heat stress, which results in thin and degenerated fingers (Plate 10.27). When banana corms were kept for 6 weeks at 32–37°C, chlorotic leaves with deformed laminae or with no laminae emerged (Anonymous, 1976). Similar symptoms have been observed in the field after a heat wave (Stover and Simmonds, 1987). Israeli et al. (1979) grew banana plants in aerated nutrient solutions at mean temperatures of 16.2°C, 25.0°C, 26.2°C and 33.6°C. At the highest temperature treatment, dry matter and potassium content declined in the corm and roots and the root/shoot dry-matter ratio decreased. Ramcharan et al. (1995) grew banana plants in pots for 6–10 weeks under controlled conditions with root zone temperatures of 28°C, 33°C, 38°C and 43°C. As temperatures increased, the dry weight of the roots decreased. In this experiment, leaf width decreased at 38°C and 43°C. At 43°C, roots were brown and suberized and lacked root tips. Lack of root tips may affect the hormone balance of the plant and thus the distribution of photosynthates. Ingram and Ramcharan (1988) investigated the physiological response of banana roots exposed to a range of temperatures between 30°C and 60°C. Damage to cell membranes was measured by electrolyte leakage and was first detected at 45°C. At 48°C, 225 ± 36 min exposure was needed to cause 50% of the maximal electrolyte leakage, while at 57°C 7 ± 4 min were needed. For heat injury, as with chilling damage, there is an interaction between temperature and time of exposure. Lahav et al. (1993) investigated the effect of polyethylene soil covers on soil temperatures in a banana plantation in Israel. During hot afternoons, the covers raised the soil temperature from 30.6°C to 37.9°C. This rise caused partial stomata closure, increased leaf temperatures from 35.7°C to 37°C and decreased photosynthesis. The number of hands per bunch was also reduced and bunch weight decreased by 28%. Above-optimum temperatures at the apical meristem during the time of bunch differentiation resulted in bunches with fewer hands (Stoler, 1962). The number of hands per bunch declined rapidly from 11.7 hands at 26.5°C to 9.4 hands at 30°C. Covering the soil with black or transparent polyethylene increased soil temperature



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Plate 10.25.  Heat effect on the opening of the unfurled banana leaf (photo: Y. Israeli, JVBES).

by 2°C and 6°C, respectively, and resulted in stunted and chlorotic suckers, which survived for only a few weeks (Stapleton and Garza-Lopez, 1988). On the other hand, covering the soil with trash or planting at an extremely high density (up to 4500 plants/ha) reduces soil temperature in very hot regions (Shanmugavelu et al., 1992).

Another aspect to be considered is the effect of heat stress on the fruit. Bananas maturing under high temperatures do not ripen normally and soften quickly, with the pulp having less aroma and taste (Tang and Liu, 1993). High temperatures in ripening rooms delay the normal decrease in chlorophyll concentration and

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Plate 10.26.  Sunburn symptom on a young banana plants in the nursery (photo: Y. Israeli, JVBES).

yellowing (Seymour et  al., 1987), resulting in over-soft, green fruit, called ‘cooked fruit’ (Thompson et al., 1972; Rippon and Trochoulias, 1976). Increasing temperatures in ripening rooms from 20°C to 35°C accelerates the softening processes, ethylene production and the accumulation of sugars, but above 30°C the de-greening process is inhibited. At 40°C, a total cessation of the ripening process was recorded (Yoshioka et  al., 1978). This is related to decreased protein synthesis, decreased acid phosphatase activity and the suppression of isoamyl acetate synthesis (Yoshioka et al., 1980a, b, 1982). High temperatures in the field may interfere with fruit development. Stover and Simmonds (1987) described a heat wave in Panama where daily maximum temperatures of 33–35°C, high radiation and a limited amount of rain caused changes in latex vessels in the peel. Latex globules exploded and cell walls and cells turned reddish brown. This resulted in symptoms of under-­peel discoloration similar to those caused by chilling. Under-peel discoloration caused by heat stress (45°C at bunch emergence) is quite common in Western Australia (Robinson, 1993). However,

Plate 10.27.  Thin fingers and premature ripening resulting from high temperatures during last stages of banana inflorescence development and emergence. The basal hands, usually the largest, were the mostly affected (photo: Y. Israeli, JVBES).

bananas are produced successfully under summer maximum temperatures above 40°C when temperature rise is gradual, water supply is adequate and supporting cultural practices, such as mulching of the soil surface and optimizing the plant population to maintain mutual shading, are implemented.

Solar Radiation The natural habitat of the wild banana is the open spaces in the tropical forest. Under shade, the banana declines rapidly (Simmonds, 1962). However, in many banana-growing areas in tropical lowlands, sunlight is limited to 4–5 h/day because of cloudiness. Nevertheless, very good crops can be obtained under such conditions. Stover (1984) showed that 14–18% of incident



Injuries to Banana Caused by Adverse Climate and Weather

radiation normally passes through the canopy to reach the ground in plantations in Central America. When plant density was increased so that only 10% of the light reached the ground, plants affected by reduced radiation could be found. Israeli et al. (1995) showed that shading slows the rate of growth of plants and time of appearance of the first ratoon. The visible symptoms of heavy shade are dark green leaves (the result of higher concentrations of chlorophyll), thinner leaf lamina, reduced rate of growth, reduced height, ‘rossetting’ of leaves, reduction in the size and number of suckers, reduction in bunch size and small, defective fingers. Generally, screenhouses with 12% shading give positive results, explained by effective wind protection, increased relative humidity, reduced evapotranspiration, the conversion of a part of the incoming radiation to diffused radiation and reduced peak hot summer transmitted radiation due to summer dust accumulation (Israeli et al., 2002; Möller et  al., 2010; Tanny et  al., 2012, 2014; Pirkner et al., 2014) Excessive radiation may also cause problems. Exposed parts of the peduncle and fruits, especially those on the upper hands, can yellow and blacken due to sunburn (Plate 10.28). These organs are prone to overheating because of their relatively small transpiring epidermis surface. The damage is more severe in bunches covered with transparent polyethylene sleeves (Plate 10.29). Bunch covers made from non-­ transparent white polyethylene significantly reduce the effects of strong sunlight, as does paper placed over the upper hands (Rippon and Turner,

Plate 10.28.  Sunburn symptoms on the banana bunch (photo: Y. Israeli, JVBES).

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1970; Kuhne, 1980b; Ke et al., 1981). Sunburn can be prevented by folding the leaf over the bunch. In plantations, most bunches are protected by the leaf canopy, but those on plants next to roadways are vulnerable (Plate 10.30). As it is believed that banana plants gradually become acclimatized to high radiation, severe sunburn may result from the sudden exposure of a protected bunch to direct sunlight. The detrimental effect of solar radiation during daytime following a chilling or frosty night was described earlier in this chapter. Wade et  al. (1993) found that ultraviolet (UVC) radiation (but not UVA) induced bronzing spots on the fruit, but when white light was applied immediately afterwards, its effect was much reduced.

Drought Droughts are major natural disasters causing huge damage and losses of human life. Millions of people died in Africa and Asia because of starvation brought by drought during the last century (Masih et  al., 2014). Unlike floods or tropical storms, which are highly devastating disasters but of short duration, drought is less acute but may last for a number of years and even decades. Agricultural drought may be moderate when rainfall is 20–59% less than the normal average or severe when there is a 60% deficiency (Samra, 2004). Local banana production in the wet tropics and also in drier parts of the world relies on rain. Drought is an important factor limiting banana production (Shou, 1990; Valmayor, 1990; Akomeah et al., 1995). Banana plants grown by smallholders are more drought-affected than those in export plantations, of which more than two-thirds are irrigated even in areas where annual precipitation is high (Carr, 2009). Van Asten et al. (2011) studied the effect of rainfall on production of East Africa highland banana (EAHB) plants in three sites in Uganda and found a linear correlation between bunch weight and the amount of rainfall received during the 12 months before harvest. Rain decline of 100 mm resulted in bunch weight loss of 1.5–3.1 kg. EAHB plants that received less than 1100 mm suffered production losses of 20–65%

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Plate 10.29.  Sunburn symptoms on bagged banana fruit (photo: Y. Israeli, JVBES).

compared with those in wetter areas. These observations demonstrate the potential of damage of even a mild to moderate drought to rain-fed banana. Minimum annual rainfall range of 1200– 2690 mm/year is necessary for optimal production (Robinson and Alberts, 1986). Stover and Simmonds (1987) believed large-scale commercial banana production could be achieved in the tropics without irrigation when the minimum annual rainfall was above 2000 mm and uniformly distributed over the whole year. However,

dry spells of a few months duration occur once or twice a year in some tropical growing areas. In North Queensland, Australia, annual rainfall is 2000–4000 mm, but some months get less than 100 mm. Irrigation during the dry season increases production by 15–20% and improves fruit quality (Daniells, 1984; Behncken, 1990). Irrigation systems are being steadily installed in commercial plantations around the world to maximize production. Drought is frequently accompanied by high temperatures and high evaporative demand,



Injuries to Banana Caused by Adverse Climate and Weather

Plate 10.30.  Symptoms of sunburn injury on the banana peduncle (photo: Y. Israeli, JVBES).

which causes a depletion in valuable water resources. A lack of water has very serious consequences. Measures such as a means to transport water from one region to another, the availability of large-scale water storage facilities and the use of recycled or desalinized water need to be put in place to combat this problem (Molden, 2007; Government of India, 2011; Dai, 2013). Milburn (1993), Turner (1994, 1995, 1998, 2013), Turner et al. (2007, 2009), Carr (2009) and Robinson and Galán-Saúco (2010) have reviewed the physiological and agronomical changes that take place in banana as a result of water deficit. Banana has a high water consumption (Ghavami, 1972; Sarraf and Bovee, 1973; Meyer and Schoch, 1976; Israeli and Nameri, 1987; Turner, 1987; Robinson and Alberts, 1989) and high sensitivity to water stress (Shmueli, 1953; Aubert, 1968; Doorenbos and Kassam, 1979; Robinson and Bower, 1987; Eckstein and Robinson, 1996; Turner and Thomas, 1998; Mahouachi, 2007, 2009). It also grows and produces much better under irrigation (Arscott et al., 1965a, b; Trochoulias, 1973; Krishnan and Shanmugavelu, 1980; Meyer, 1980; Holder and Gumbs, 1983b; Robinson and Alberts, 1986).

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Banana needs an ample supply of water, because it has a large vegetative mass, a high hydration level, broad thin leaves, which absorb much energy and transpire water easily, and a sparse and shallow root system (Shmueli, 1953; Olsson et  al., 1984; Robinson, 1985). Eckstein (1994) showed that the first symptoms of water stress (partial closure of stomata and reduction in carbon fixation) in the field are expressed 4 days after cessation of irrigation. Twelve days without water resulted in a 79% reduction in CO2 assimilation, caused mainly by stomatal closure and partially by reduced photosynthetic system activity. The reduced assimilation results in reduced growth and production. The first symptom in the crop, even when no other symptoms are observed, is reduced finger length. If the fruit is affected during its initial stages of development, several weeks or even months may pass between the period of water stress and the emergence of the affected bunch. The effect of water deficiency is especially noticeable during periods of high temperatures and otherwise optimal growth conditions (­Arscott et al., 1965a; Robinson and Green, 1981). Water stress slows down the rate of fruit growth and at the same time stimulates maturation (­Arscott et al., 1965a; Srikul and Turner, 1995). The fruit will be affected even when a limited ­period of water stress occurs between bunch emergence and harvest (Hegde and Srinivas, 1989; Mahaouachi, 2007). The first visible symptoms of water stress are usually expressed in the leaves as a colour change from dark to pale green and the continuous folding of the lamina halves during the day when temperatures are high. Later, leaves become yellowish green followed by necrosis at the margins (Plate 10.31). The tissue on both sides of the midrib, where the lamina is directly exposed to the sun, also becomes necrotic. These symptoms are more pronounced in leaves sprayed with oil to control leaf spot diseases. Water deficit delays bunch initiation and flowering. During continuous water stress, the petioles are crowded at the top of the pseudostem and the bunch is choked on emergence similar to ‘choke throat’ caused by low temperatures (Kuhne and Green, 1980; Holder and Gumbs, 1983a). Water stress also affects bunch shape, such as non-uniform distances between hands, large differences in finger length between the upper and the lower

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Plate 10.31.  ‘Grand Nain’ (AAA, Cavendish subgroup) after 5 months of drought in the arid Jordan Valley, Israel (photo: Y. Israeli, JVBES).

hands and extreme curving of fingers towards the bunch axis or in the opposite direction to form an open hand (Holder and Gumbs, 1983a) (Plate 10.32). Leaves can also age earlier, followed by leaf folding and desiccation (Wardlaw, 1961a; Kallarackal et  al., 1990). Later, plants become stunted and small with a reduced leaf area. Finally, growth ceases and plants have small, malformed bunches (Wardlaw, 1961a; Stover and Simmonds, 1987; Robinson and Galán-Saúco, 2010). Hill et al. (1992) reported up to 35% unmarketable bunches due to pseudostem breakage and snapping of the peduncle in Carnarvon, Western Australia, from October to March. This is a period of strong winds, high temperature and low humidity, which cause extreme moisture stress. Bunches are also frequently lost in the Windward Islands during the dry season because of the breakage of drought-weakened pseudostems (Plate 10.33). Loss of green leaves results in peduncle and fruit sunburn, early field ripening and split fingers. The B genome of M. balbisiana increases relative resistance to drought (Simmonds, 1962). Thomas et al. (1998) demonstrated that the decrease in leaf stomatal conductance in response to an increase in the leaf to air vapour-pressure difference is lower in cultivars with the B genome. Banana clones with the AAB genome are more resistant than those with the AAA genome, and

Plate 10.32.  Inward curving of banana fingers due to drought (photo: Y. Israeli, JVBES).

cultivars in the ABB group are the most drought-­ resistant (Stover and Simmonds, 1987; Valmayor, 1990). However, conflicting observations on the drought resistance conferred by the B genome



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Plate 10.33.  Breakage of a drought-weakened pseudostem of ‘Robusta’ (AAA, Cavendish subgroup) in St Vincent, Windward Islands. Propping can help to prevent such losses (photo: D.R. Jones, SVBGA).

were reported by Bananuka et  al. (1999). Ekanayake et al. (1995) showed that differences may occur between clones in the same genome-group according to their tendency to close stomata under high vapour-pressure deficit during the afternoon. Enset is considered to be a drought tolerant crop (Simmonds, 1962). Our understanding of the genetic, genomic and proteomic basis of the banana response to water stress is still limited, but much research is being undertaken (Vanhove et al., 2012; Kissel et al., 2015, 2016) in order to support a future genetic improvement strategy to improve drought tolerance in banana. Within the AAA group, the Cavendish subgroup seems to be more drought-sensitive than ‘Gros Michel’ (Stover and Simmonds, 1987). Variations also exist within the Cavendish subgroup. ‘Grand Nain’ experiences water stress, expressed as choking and fruit deformation, in climatic conditions where ‘Williams’ is symptomless. ‘Williams’, on the other hand, is more sensitive to drought than ‘Robusta’, showing a 14% reduction in the number of hands compared with 7% in ‘Robusta’ (Holder and Gumbs, 1983a).

Israeli (2000) studied the effect of withholding irrigation on ‘Dwarf Cavendish’ and ‘Grand Nain’ during a hot, arid summer in the Jordan Valley in Israel. He found that the former cultivar was much more resistant than the latter in both gas exchange and plant growth parameters. Ploidy level may also influence reaction to water stress. Triploids were found to be more susceptible than diploids and tetraploids (Ramadass and Sheriff, 1993). The ecological and physiological adaptations of banana in terms of its reaction to stresses caused by insufficient water were discussed by Turner et al. (2009). Banana would appear to be a typical isohydric plant that maintains close to constant midday leaf water potential even under drought conditions by reducing stomatal conductance as necessary to limit transpiration (for definitions of isohydric and unisohydric plants, see Meseda and Fernandez, 2006; McDowell et al., 2008; Limpus, 2009). Experiments on the response of banana gas exchange to withholding irrigation under field conditions with high evaporative demand has revealed a fast decrease in stomatal conductance during the middle of the day, followed by increase in leaf temperature,

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reduced carbon fixation and a complete cessation of growth (Robinson and Bower, 1987; Eckstein and Robinson, 1996; Thomas and ­ Turner, 1998; Israeli, 2000). Although measurable diurnal and seasonal changes in leaf turgor pressure represent environmental effects on the ­banana leaf water status (Zimmermann et  al., 2010), changes in leaf water potentials between irrigated and non-irrigated suckers are minimal. The banana roots respond to the soil water deficit by sending an abscisic acid signal, which is known to be active in mediating the adaption of plants to stress, to the leaves (Thomas and Turner, 1998; Mahouachi et al., 2014) while the epidermis is directly sensing an increase in the leafto-air vapour-pressure difference. Both signals ­control the stomatal conductance so that the leaf water potential remains almost constant. When stomata close, transpiration is reduced and leaf energy balance is affected. The down-folding of the leaf lamina halves, which is at least partially independent of their water status (Thomas and Turner, 2001), helps to mediate the increase in midday leaf temperature. However, this is ineffective when transpiration is much reduced. Then, stomatal limitation coupled with the direct damage to the photosynthetic apparatus leads to carbon starvation. Turner et al. (2009) pointed to the hypothesis that the extreme sensitivity of banana to soil drying is actually a means of survival: the plant is maintaining its water potential at the expense of productivity. This is not ideal from an agricultural point of view, but an excellent strategy for a plant that has an underground rhizome with many protected growing points that can survive drought and resume growth when rain returns (Plate 10.34). When transpiration is low, it is the root pressure that must maintain the water potential of the plant (Turner et  al., 2009). The function of root pressure in banana is demonstrated in the guttation of leaf sap that may amount to 400 ml/m2 during six night hours (Shapira et al., 2013). In summary, the high sensitivity of the banana to soil and atmospheric water deficit and its ability to maintain its hydrated state under these conditions is an ecological adaptation to its native environment, its morphology and life cycle. An increase in the banana water use ­ ­efficiency, an important agricultural target, depends on genetically changing these adaptations.

Plate 10.34.  Regrowth of banana underground buds and their survival over 6 months during the dry, hot summer of the Jordan Valley, Israel. The other species that survives these harsh conditions is the deep-rooted, drought-tolerant Syrian mesquite (Prosopis farcta) (photo: Y. Israeli, JVBES).

Excess Water Roots need an appropriate supply of oxygen for their normal growth and function. Under poor drainage conditions, the roots develop close to the surface, where some oxygen is still available. In the absence of oxygen, the roots lose their rigidity, present a pale bluish-grey colour and gradually rot (Delvaux, 1995). Aguilar et  al. (2003) studied the oxygen consumption of the banana roots in detail. Respiratory oxygen consumption was at a peak in the root apex and in the nearby root hair zone. They found that the respiration activity of the stele was high and this tissue was the first to experience the effects of oxygen deficiency. Nutrient loading of the stele was one of the earliest root functions to be affected, followed by root tip elongation. Anoxia for more than 6 h killed the root tip (Aguilar et  al., 1998) and, after proper drainage, lateral



Injuries to Banana Caused by Adverse Climate and Weather

roots grew from behind the necrotic root apex and formed a morphological feature that is commonly called ‘chicken feet’. ‘Chicken feet’ is a useful diagnostic criterion if the effect of waterlogging is being evaluated after the actual event. Reduced latex flow from severed latex ducts, which occurs a few days after waterlogging has occurred, is also useful in diagnosis (D. Turner, Australia, 2015, personal communication). A genetic variability in the response to soil oxygen deficiency was identified and the possible future use of this variability was postulated (Aguilar et al., 2008). Excess water is a significant problem in many banana plantations in tropical lowlands. Drainage canals and ditches are prepared before planting in order to avoid waterlogging. Ideally, such drainage systems should allow excess water to be channelled away as early as possible after rain and lower the water table to below 1 m about 24 h later (Stover and Simmonds, 1987). In some areas, drainage pumps are required (Lassoudiere and Martin, 1974). Drainage problems are more severe in clay soils with low hydraulic conductivity. If planting in such soils is

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unavoidable, drainage ditches should be positioned quite close to one another. Growth ceases in plants flooded for 48 h. The yellowing of leaves is the first symptom to become apparent. Longer periods of flood (72– 96 h) result in irreversible damage (Plate 10.35). Flood damage is more severe when water is standing than when water is flowing. This may be related to the lower oxygen content of standing water. The negative effect of excessive soil humidity is aggravated when soil temperatures are high (Ramcharan et al., 1995). Irizarry et al. (1980) showed that lowering the water table from 12 cm to 36 cm increased production in ‘Maricongo’ (AAB, Plantain subgroup) from 5.6 t/ha to 37.8 t/ha and that this was correlated with an increase in root production. Ghavami (1976), working with ‘Valery’ (AAA, Cavendish subgroup), found that damage first became evident when the water table rose above 60 cm and was more significant during the second and third crop cycles, probably because excess water not only caused root necrosis, but also prevented root formation.

Plate 10.35.  Yellowing of leaves of ‘Kluai Namwa’ (ABB, Pisang Awak subgroup) due to floods along the Mekong River in Thailand (photo: D.R. Jones, INIBAP).

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Plate 10.36.  Irrigation of banana growing on embankments in old paddy rice fields near Bangkok, Thailand (photo: D.R. Jones, INIBAP).

Both Ghavami (1976) and Holder and Gumbs (1983c) related significant yield decreases to rises in the water table. Floods associated with tropical storms may cause more damage than the strong winds (Stover and Simmonds, 1987; Tang and Liu, 1993). In addition to the direct damage caused by waterlogging, flood water also weaken the resistance of plants to diseases, such as Fusarium wilt (Stover and Malo, 1972; Valmayor, 1990; Shivas et al., 1995). The construction of ridges is an efficient method to reduce the damage to banana plants in areas of heavy soil and high water table (Avilan et  al., 1982; Daniells, 1984). In Thailand, banana is grown successfully on raised embankments in redundant paddy rice fields (Plate 10.36).

Lightning Strike This is a rare event that has been reported in Jamaica, Suriname (Maas, 1967), Honduras, North Queensland and Hawaii. The damage is

caused by the extreme heat and shock waves generated by the electricity in the lightning bolt. Banana plants collapse at the base or mid-­ pseudostem or at the crown (Plate 10.37). The pseudostems explode because of the high temperature and pressure created by the strike that turns the plant fluid into steam (Nelson, 2008). Other symptoms include the yellowing, browning and dieback of the leaf tip. The midrib also collapses about 20–30 cm from the leaf tip, causing it to hang down. These collapsed yellow leaf tips or yellow ‘flags’ attract attention to otherwise normal green plants in plantations. A purple-­ brown or reddish discoloration is present in the midrib from the point of leaf collapse to the pseudostem. The entire leaf eventually turns brown and collapses. Discoloration advances from the collapsed leaves along the midrib and petiole into the pseudostem and leaf sheaths. The inner leaf sheaths show brown, elongated lesions up to 2.5 cm in length on the outer epidermis. The vascular tissue may turn brown. The pseudostem begins to rot from the top downwards and collapses as a soft rotten mass, but the rhizome is not affected. The condition appears in localized



Injuries to Banana Caused by Adverse Climate and Weather

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Plate 10.38.  Under-peel discoloration of banana fruit caused by chilling temperatures. The latex ducts are discoloured and such fruit ripens to a dull greyish-yellow colour. Compare with the normal, non-chilled fruit (photo: Y. Israeli, JVBES).

more slowly than normal fruit, will have a dull yellow colour and, if the discoloration is severe, fruit may not yellow, but may become dull grey. Also, the pulp may be hardened. Eating quality is affected only when chilling injury is severe. Under-peel discoloration from chilling may occur in the field when temperatures drop below Plate 10.37.  Collapse of the banana canopy due 12.8°C for several hours as a result of cold to lightning injury (photo: S. Nelson, Hawaii). fronts. More commonly, it occurs in transit because of excessive refrigeration or ventilation in areas of 0.5–2 ha and affects up to 50% of the ships and vans. Fruit exposed to winter temperaplants. Symptoms appear rapidly over a period tures while in transit in non-tropical areas may of about a week and then gradually cease after also be chilled. 10–14 days. Plants destroyed by lightning strikes Chilling in transit is avoided by maintaining should be cut down and the ground replanted. strict control over the temperature of inflowing air. Air temperatures should not be below 13.2°C for fruit of cultivars of the Cavendish subgroup Under-peel Discoloration (AAA) and 12.2°C for fruit of ‘Gros Michel’ (AAA). Chilling is most likely to occur on fruit Under-peel discoloration consists of a reddish-­ stored near air delivery ducts. Periods of storage brown streaking in the vascular tissue just be- beyond 14 days can also lead to under-peel discollow the fruit epidermis (Plate 10.38). Symptoms oration even at 13.3°C. Under-peel discoloration are visible when the epidermis is peeled back associated with long storage or transit periods is with a knife. Where under-peel discoloration is probably related to desiccation as well as low present, latex flow is reduced or stopped and the temperature. Von Loesecke (1950), Wardlaw (1961b), peel tends to adhere strongly to sub-­epidermal Palmer (1971), Mattei (1978), Marriott (1980) layers. Under-peel discoloration usually indicates and later Duan et al. (2007) reviewed the physithat the fruit was subjected to chilling tempera- ology of postharvest chilling of banana fruit. tures. Less frequently, it can also be a symptom Ratule et al. (2006) noted changes in the degree of fruit desiccation, ageing at low humidity, with of browning, peel electrolyte leakage, soluble or without high temperatures, and bruising. solids content, weight loss and change in colour Fruit with under-peel discoloration will ripen of fruit of ‘Pisang Berangan’ (AAA, Lakatan

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subgroup) during storage at 5–15°C for 4–16 days and found the last three parameters to be the most quantitative indicators for chilling damage. The use of a colorimeter has enabled changes in peel colour resulting from chilling, either before or after ripening, to be evaluated objectively and quantitatively (Hewage et  al., 1996). Thus chilling damage can be identified even before external symptoms are visible and after very moderate chilling temperatures (11°C), resulting in very light chilling symptoms. Electrical conductivity of the peel is another parameter used to assess damage (Deullin, 1980). Recently a modern automatic physiological monitoring system was suggested for ships that transport export bananas so that warnings could be given if fruit in containers had temperature or reduced greenlife problems. Emphasis was placed on avoiding chilling or heat stress (Jedermann et al., 2014). The fruit of banana genotypes vary in their response to chilling temperatures (Marriott and New, 1975; Broughton and Wu, 1979; Mohammed and Campbell, 1993). Fruit of Cavendish cultivars is more sensitive than ‘Gros Michel’ and cultivars in the AAB and ABB genomic groups (Satyan et al., 1992; Morrelli et al., 2003; Promyou and Ketsa, 2010). Chilling injury is a function of temperature and exposure time. It is less pronounced if the chilling process is gradual, when oxygen is reduced and when relative humidity increases in the ambient atmosphere (Mattei, 1978). Rate of under-peel discoloration may decrease when the fruit is protected by vegetable oil, mineral oil or dimethylpolysiloxane solution. Treating fruit with these chemicals enabled fruit to be stored at 9°C for 24 h with no significant damage (Jones et al., 1978). Heat treatment (38°C for 3 days) prior to storage at 8°C resulted in chilling tolerance in which the enzyme phenylalanine ammonia-lyase was involved, at both transcriptional and translational levels (Chen et al., 2008). Pre-treatments with abscisic acid, jasmonic acid derivative and putrescine helped to maintain partial membrane integrity in banana fruit peel exposed to low temperatures, thus alleviating chilling injury (Duan et al., 2007). Chilling injury is related to a decrease in activity of peroxidases (Toraskar and Modi, 1984), oxidation of phenols, especially dopamine (Abd El-Wahab and Nawwar, 1977), and an increase in the level of α-farnesene, which

precedes the appearance of external chilling symptoms (Wills et  al., 1975). A possible effect of chilling on phase transition in membrane structural lipids (Lyons, 1973) was demonstrated by the identification of saturated lipid acids, typical in sensitive plants, in banana (Roughan, 1985; Kernick and Bishop, 1986). Also, chilling reduces unsaturated lipid acids, especially linolenic acid (Wang and Gemma, 1994), and changes membrane permeability (Gemma et al., 1994).

Global Climate Change The earth has been experiencing global warming and climate change. The change is worldwide, because it covers both sea and land surfaces in the northern and the southern hemispheres. Jarvis et al. (2008) used global climatic models to describe the expected effect on temperatures and rainfall in typical banana production regions for 2050 if current trends continue. An increase in mean annual temperature of 1.5–3.2°C was obtained with increases in annual precipitations of > 100 mm for East Africa, South Asia and Ecuador, and decreases of < 100 mm for Caribbean islands. In addition, the authors used the EcoCrop model (CIAT, 2016) to evaluate the suitability of the expected climate for banana production in different regions compared with the present climate. The analysis revealed a global increase of 6% in suitability, but most of it in regions with low production density. Changes in crop management, varietal changes and migration to different zones of production might be needed, as well as genetic improvement (Ramirez et al., 2011). Van den Bergh et  al. (2012) used the EcoCrop model to evaluate the suitability of nine selected subtropical banana production sites after climate changes predicted for 2020 and 2050. The results indicated no significant change for the Canary Islands, the Mozambique–South Africa border area, northern Morocco and southern coastal China, improved suitability for two sites (South Brazil and southern non-coastal China) and a decrease in suitability for three sites (Formosa Province in Argentina, northernmost India, and Paraguay). Furthermore, the study indicated that subtropical environmental conditions suitable for banana cultivation are widely represented globally and a transfer of



Injuries to Banana Caused by Adverse Climate and Weather

technology to the most promising may increase banana production. A recent detailed assessment of global banana production as a result of climate change was given by Calberto et  al. (2015). This work was more optimistic and suggested that, following a 3°C increase in average temperatures by 2070, land suitable for banana production, which would be mostly in the subtropics and in the highland tropics, will increase by about 50%.

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However, the authors were aware of the main weakness of prediction models, which are based on averages, and recognized that extreme weather events may also occur. These events may sometimes decrease rather than increase suitability (Dai, 2013; Masih et  al., 2014; Sabiiti et  al., 2016; USAID, 2016). Adaptation to the changes relies not only on a coordinated global response, but also on the actions taken at the level of the small-scale farmer (Below et al., 2010).

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11 

Chemical Injury to Banana E. Lahav and Y. Israeli

Introduction Cultural and biological control practices may ­reduce the need to use agricultural chemicals to sustain plant growth and protect crops from pests and diseases (Constantinides and McHugh, 2003; Pattison et  al., 2005; Isaac et  al., 2012; King, 2016). However, commercial banana production requires the input of significant quantities of chemicals to maintain optimum soil fertility, control weeds and combat diseases and pests. Occasionally, these agrochemicals are applied either in excess or improperly and injury to the banana plant or fruit can occur. Banana plants can also be damaged if grown in soil containing minerals in toxic concentrations or in environments where the air is polluted. Damage associated with chemical injury to banana is described in this chapter.

Injury Caused by Agricultural Chemicals

the leaves, ­followed by necrosis. The first symptom, which appears on leaves 2–5, counting down from the first fully opened leaf, is a whitish discoloration or brown band similar in appearance to sunburn. Brown spots may appear parallel to the edge of the leaf, but not at distal or proximal positions. These spots become ­necrotic and the tissue disintegrates. The leaf becomes thin and greyish white, and tends to puncture and fragment (Charpentier and M ­ artin Prevel, 1968). These symptoms usually disappear when the excess is leached from the soil by rain or irrigation. If the symptoms persist, this is most probably the result of lack of drainage or the presence of excessive salts associated with saline soils. Some fertilizers, as ammonium and amides, result in soil acidification and related problems, such as aluminium toxicity (Serrano, 2005). The addition of dolomitic lime (CaMg(CO3)2) or gypsum (CaSO4.2H2O) might interfere with relationships between nutrients and result in a deficiency of potassium or iron.

Fertilizers

Pest and Disease Control Chemicals

Excessive fertilizer application increases salinity, which is expressed by increased osmotic tension of the soil solution and drought stress (in addition to the specific toxic effect of elements, such as sodium and boron). Typical ­salinity symptoms are marginal chlorosis of

Fungicides are used to treat banana foliage in the plantation, young bunches before bagging and harvested fruit in the packing house. When applied under unfavourable conditions or when inappropriate practices are followed, phytotoxic damage may occur. Mineral oil spray, used against

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banana leaf spot diseases since the late 1950s, initially caused significant injury. External symptoms of oil toxicity are leaf burns, usually along the midrib, and brown to black necrotic spots on the lamina (Plate 11.1). This type of damage was much reduced with the change to oils of a narrow distillation range (346–354°C), containing less than 12% aromatic compounds and above 90% unsulfonated residues (Calpouzos et  al., 1961; Calpouzos, 1968). However, sprayed plants without external injury symptoms produced bunches 8% lighter than untreated control plants, demonstrating oil’s symptomless phytotoxicity (Israeli et al., 1993). Tridemorph, a fungicide used to control Sigatoka leaf spot and black leaf streak diseases, may sometimes cause leaf yellowing when used in combination with oil. This yellowing is stronger in mature leaves exposed to the sun (Cronshaw, 1987), in water-stressed plants when radiation is high and after the application of some herbicides. Damage can also be caused to young fruit before bagging when the fingers are still pointing horizontally or downward (the top side being exposed to droplets of fungicide). Foliage damage may also occur when the protectant fungicide chlorothalonil is applied following an oil-systemic mixture application (R. Barnett, Philippines, 2016, personal communication). Damage to fruit of

‘Grand Nain’ (AAA, Cavendish subgroup) as a result of the spray application of different fungicides, oil and water is illustrated in Plates 11.2 and 11.3. Insecticides, such as chlorpyrifos, imidacloprid, bifenthrin or acephate, used for bud injection, a common practice in Australia, Asia and Hawaii where flower thrips and scab moth can attack developing fruit (Broadley et  al., 2004; Nelson et al., 2006; Newley et al., 2008; Sankar, 2014), can damage very young flowers. The timing, point of injection, quantity delivered and concentration must follow the recommendations in order to avoid phytotoxicity. The initial indication of damage is the blackening of the male bud tip. Later, when the hands open, the bud looks twisted and the last four to five hands turn black. In addition to phytotoxic problems, physical damage to the bottom hands may occur if the bud is injected at the wrong point. Recently opened bunches can be sprayed with fungicides, such as thiophanate-methyl or carbendazim, before bagging to control pitting disease or diamond spot. Sometimes ionic copper concentrate is applied to control bacterial soft rot. Insecticides may also be used again at this stage if scab moth, rust thrips or scarring beetle pressure is high (Broadley et  al., 2004). These chemicals are rarely phytotoxic, but the mineral

Plate 11.1.  Damage to a banana leaf caused by a spray application of mineral oil on a hot day (photo: R. Barnett, Philippines).



Chemical Injury to Banana 529

Plate 11.2.  Water-soaked speckling caused by a combination of fungicides, oil and water on fruit of ‘Grande Naine’ (AAA, Cavendish subgroup) in Costa Rica (photo: C. Pasberg-Gauhl and F. Gauhl, CB).

Plate 11.3.  Necrotic speckling caused by a combination of fungicides, oil and water on fruit of ‘Grande Naine’ (AAA, Cavendish subgroup) in Costa Rica (photo: C. Pasberg-Gauhl and F. Gauhl, CB).

oil carrier or poor application techniques, especially too high a concentration/volume or large droplets, may cause damage. Thiophanate-­methyl is still the main fungicide used against pitting disease. Sometimes, when over-application occurs or when the worker fails to agitate the spray tank before application, the peel is damaged (Plates 11.4 and 11.5). Fruit with damaged peel is rejected in the packing house. Carbendazim has resulted in significant phytotoxicity in the Philippines when applied to plants suffering from extended drought stress (F. Odtojan, Philippines, 2016, personal communication). Quaternary ammonium compounds used for disinfecting tools to prevent the spread of Moko bacterial wilt are phytotoxic if applied to fruit.

Plate 11.4.  Damage to young fruit of a Cavendish cultivar caused by spraying with the herbicide thiophanate-methyl (photo: R. Barnett, Philippines).

Plate 11.5.  Chemical burn caused by the herbicide thiophanate-methyl applied as a spray to young fruit of a Cavendish cultivar (photo: Y. Israeli, JVBES).

In the packing house, alum sulfate is applied to fruit to reduce latex flow and fungicides, such as iprodione, imazalil or prochloraz, are commonly applied to the fresh-cut cushion to control the postharvest disease, crown rot. Because there is a risk that fruit may carry excess residues of fungicides, chlorine dioxide (ClO2) or calcium hypochlorite (Ca(ClO2)) has been used to generate the disinfectant chlorine in tank water (Lassoudiere, 2007). However, exceptionally high levels of chlorine or contact with the chlorinated water for more than 30 min can cause a reddish flecking or blackening of the crown and peel. The monitoring of chlorine levels by measuring its concentration in the tank using the ­orthotolidine test and also taking the pH of the ­water every 30–60 min is recommended (L. Truggelmann, Philippines 2016, personal communication).

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Herbicides

and environmental impact should also be considered. The use of herbicides in dessert banana and planThe herbicide paraquat is not translocated tain plantations was reviewed by Lassoudiere and systemically. Yellowing, chlorosis and necrosis Pinon (1971), Feakin (1972), Lassoudiere (1972, only appear where the herbicide has been in 2007), Irizarry (1987), Stover and Simmonds contact with the leaf tissue (Lassoudiere, 1972; (1987), Tezenas du Montcel (1987), Galán-­ Liu and Singmaster, 1990) (Plate 11.6). ThereSaúco (1992), Shanmugavelu et al. (1992) and fore, it is recommended, as with other contact Robinson and Galán-Saúco (2010). Modern herbicides, that it be used only when banana ­approaches that incorporate use of cover crops, suckers are taller than 1 m (Feakin, 1972; Stover dry mulch and manual chopping in order to and Simmonds, 1987; Galán-Saúco, 1992; minimize herbicide have also been suggested ­ Robinson and Galán-Saúco, 2010). Manual (Constantinides and McHugh, 2003; Pattison weed control is recommended when suckers are et al., 2005; Newley et al., 2008; Isaac et al., 2012; smaller. Planting material derived from tissue Weis et al., 2012; King and Lindsay, 2014). Herbi- culture is extremely sensitive to paraquat, as cide phytotoxicity has been reviewed by Lassoudiere well as ­other herbicides, as it is initially low-lying (1972) and Soto (1985). Other ­publications on with dense foliage (Kwa and Ganry, 1990; Marie this subject are by Guillemot (1975), Velez-­Ramos et al., 1993). Drift is the main risk in using paraand Vega-Lopez (1977), Ramadass et al. (1980), quat. This risk may be reduced if special equipLiu et  al. (1985), Cronshaw (1987), Liu and ment, such as a low-pressure spray, special spray Rodriguez-Garcia (1988), Liu and Singmaster nozzle and mechanical shield, is used. Keeping (1990) and Gonzalez and Piedrahita (1994). the spray nozzle close to the ground and the adSymptoms associated with herbicide damage are dition of specific adjuvants may also help. Under summarized in Table 11.1. no circumstances should herbicides be sprayed There is much evidence that weed competi- when conditions are windy (Feakin, 1972; Liu tion affects the rate of growth and yield of banana and Singmaster, 1990). The precision spraying and that mechanical or chemical weed control is of paraquat using a shielded nozzle is the comnecessary for maximum production (Mishra and mon choice for young plantings when manual Das, 1984). Mechanical weed control may some- weeding is not possible. times give better results than chemical control Glyphosate is a contact herbicide with long(Venero, 1980), but in most cases this method is range, systemic activity. After contact with banana more expensive (Irizarry, 1987) or less efficient leaves or the pseudostem, it is absorbed and trans(Isaac et  al., 2012). As a result, herbicides are located to the meristems, which causes interveincommonly used. The most popular contact her- al chlorosis and the deformation of young leaves bicides are paraquat, glyphosate and glufosinate (Plate 11.7). Sometimes the lamina is totally (Stover and Simmonds, 1987; Shanmugavelu absent (Guillemot, 1975; Achard, 1993). The et al., 1992; Achard, 1993; Dave, 1993). Simazine, major danger from glyphosate is its delayed action diuron, oxyfluorfen and ametryne are contact and cessation of growth even when no external herbicides with residual effect (Liu et al., 1985; symptoms are observed. Frequent applications, Irizarry, 1987; Liu and Santiago-Cordova, 1991; which are recommended to reduce thick stands Dave, 1993). Because of human health concerns, of weeds, should be applied as close to the ground most herbicides have either restrictions on their as possible (Liu and Rodriguez-Garcia, 1988). use or are prohibited. When glyphosate was applied to the pseudostem All herbicides, if not properly used, might of banana suckers taller than 1 m in controlled delay growth or cause damage to banana plants. experiments, no phytotoxicity was recorded However, the degree of damage differs greatly (Aguero, 1998; Brenes-Prendas and Aguero-­ according to the compound in use and is affected Alvarado, 2012), but in practice banana suckers by the stage of plant development, cultivar, climate, and plants derived from tissue culture are highly soil type, method of application, agro-techniques sensitive. This is probably because of the difficulbeing applied and presence of other chemicals. ty in controlling the actual spray volume and Phytotoxicity is only one factor to be taken into coverage. Glyphosate is used in high concentraaccount for herbicide selection. Efficiency, cost tion to destroy banana plants affected by Moko

Name of herbicide Paraquat

Commercial product and Pre- or content of a.i. post-emergence Gramoxone® 200 g/l

Dalapon

Post

Rate (g a.i./ha)

Mode of action

Phytotoxicity hazard

200–1000

Contact

Low

Post

Glufosinate Basta® 200 or 600 g/l

Post

300–600

Mainly contact

Ametryne

Pre and post

800–5000

Oxyfluorfen Goal® 240 g/l

Pre and post

500–1200

Contact, systemic residual Contact, Moderate systemic, residual

Diuron

Karmex® 80%

Pre

1000–4800

Residual

High

Simazine

Gesatop® 500 g/l

Pre

500–3000

Residual

Moderate

Gesapax® 500 g/l

a.i., active ingredient.

Post

3000–12,000 Systemic, residual 500–1500 Systemic

Low

Phytotoxicity symptoms

References

Local chlorosis and necrosis on direct contact with leaves or live leaf sheaths Not reported

Lassoudiere and Pinon (1971); Liu and Singmaster (1990); Achard (1993) Irizarry (1987); Shanmugavelu et al. (1992) Guillemot (1975); Achard (1993)

High

Reduced growth rate, typical chlorosis and lamina deformations on young leaves Low, moderate Local yellowing and browning Perkins (1990); Gonzalez on young, in vitroafter direct contact, spreading and Piedrahita (1994); propagated plants towards the cigar leaf of Achard (1993); Y. Israeli young, in vitro-propagated (unpublished) plants Low Temporary chlorosis on young Guillemot (1975); Soto leaves; reduced fruit size (1985); Cronshaw (1987) Leaf necrosis after direct contact; marginal leaf necrosis when affected by gaseous form Yellowing along midrib of older leaves, spreading towards the margins; reduced growth rate

Marginal leaf necrosis

Escorriola et al. (1979); Liu and Singmaster (1990); Y. Israeli (unpublished) Guillemot (1975); Velez-Ramos and Vega-Lopez (1977); Ramadass et al. (1980); Cronshaw, (1987); Shanmugavelu et al. (1992) Soto (1985); Y. Israeli (unpublished)

Chemical Injury to Banana 531

Dowpon® 85% Glyphosate Roundup® 480 g/l



Table 11.1.  Characteristics of herbicides in use and their phytotoxic symptoms on banana.

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Plate 11.6.  Leaf burn on the older leaves of young plants of a Cavendish cultivar caused by a spray application of the herbicide paraquat (photo: Y. Israeli, JVBES).

Plate 11.7.  Interveinal chlorosis and lamina deformation caused by the herbicide glyphosate on a banana sucker growing in Martinique, French Antilles (photo: Y. Israeli, JVBES).

bacterial wilt, bunchy top or Fusarium wilt and also those growing in abandoned plantations (CSFRI, 1983). Repeated applications of the same herbicide results in development of populations of resistant weeds. This has happened with both paraquat and glyphosate, which are widely and commonly used worldwide. A natural response of field workers is to increase herbicide concentration and/or volume applied, As a consequence, an increase in phytotoxic damage is recorded. New herbicides applied alone or in a mixture with glyphosate to control Diffenbachya oerstedii were tested by Brenes-Prendas and Aguero-Alvarado (2012) in Central America with partial success. Isaac et al. (2012) successfully demonstrated an integrated approach to weed management by combining chemical control with mulch and cover crops to overcome the problem of Commelina diffusa. The use of glyphosate was minimized by mixing it with other herbicides. Glufosinate is a partially systemic contact herbicide (Perkins, 1990; Gonzalez and Piedrahita, 1994). Chlorotic and necrotic spots are induced when glufosinate is in direct contact with banana leaves. Young plants derived from tissue culture are especially sensitive and contact with exposed leaves results in chlorosis and necrosis of the funnel leaf several days later (Plate 11.8).



Chemical Injury to Banana 533

Plate 11.8.  Chlorosis and necrosis of young leaves of tissue culture-derived banana plants caused by the herbicide glufosinate in Martinique, French Antilles (photo: Y. Israeli, JVBES).

Oxyfluorfen is both a contact and systemic herbicide with a residual effect in the soil. Because of its high volatility, it may also act in gaseous form. It may be used in combination with glyphosate in order to shorten the time for a visible effect and overcome some resistant species. Brown necrotic spots are formed on banana leaves following direct contact with oxyfluorfen (Plate 11.9) with a marginal necrosis symptom the result of translocation (Plate 11.10) (Escorriola et  al., 1979) Usually the damage is limited, but oxyfluorfen use at high temperatures, when volatility is enhanced, should be avoided. It is also specifically problematic in screenhouses, where air circulation is limited and its gaseous form may reach high concentrations (Plate 11.10). Diuron was one of the most popular residual herbicides (Feakin, 1972; Lassoudiere, 1972; Tosh et  al., 1982; Soto, 1985; Irizarry, 1987; Stover and Simmonds, 1987) and is still in partial use today (Soto, 1985; Shanmugavelu et al., 1992; Dave, 1993; Robinson and Galán-Saúco, 2010). It has been used alone or with contact herbicides. Phytotoxicity symptoms on banana have been reported by Lassoudiere (1972),

Guillemot (1975), Velez-Ramos and Vega-Lopez (1977), Ramadass et  al. (1980), Tosh et  al. (1982) and Shanmugavelu et al. (1992). Initial symptoms are yellowing along the central midrib and later towards the margins of mature leaves (Guillemot, 1975; Ramadass et al., 1980). The bunch is also greatly damaged (Plate 11.11). Phytotoxic effects occur more frequently in plants growing during the dry season in light soils, where the organic matter is low (Lassoudiere, 1972). Lower concentrations of diuron should be used under these conditions. Plantain is more sensitive to diuron than other banana types (Tezenas du Montcel, 1987). Ametryne is one of the oldest herbicides, but is hardly used today. It is both a contact and a pre-emergence herbicide (Liu et  al., 1985; Stover and Simmonds, 1987; Dave, 1993; Robinson and Galán-Saúco, 2010). Ametryne is usually not phytotoxic, but chlorosis might be induced when high concentrations are absorbed by banana roots (Lassoudiere, 1972). The range of efficiency of ametryne is limited and it is usually mixed with other herbicides, such as paraquat, simazine or other triazynes (Soto, 1985).

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Plate 11.9.  Necrosis on a leaf of a young Cavendish plant derived from tissue culture caused by the herbicide oxyfluorfen (photo: Y. Israeli, JVBES).

Plate 11.10.  Marginal leaf scorch caused by the herbicide oxyfluorfen on Cavendish plants growing inside a screenhouse (photo: Y. Israeli, JVBES).

Such combinations might induce phytotoxic symptoms in banana when applied at double-­strength concentrations (Guillemot, 1975). Because of its persistence in soil, ametryne, as with other residual herbicides, may be displaced downhill during heavy rains. Displacement depends on the slope and rain intensity (Liu et al., 1985).

Plate 11.11.  Cavendish bunch showing toxicity symptoms after the excessive application of the herbicide diuron in Western Galilee, Israel (photo: E. Lahav, VI).



Chemical Injury to Banana 535

Simazine is a well known residual herbicide, but it is limited in its use today because of registration limitations. It can be applied alone as a pre-emergent (Lassoudiere and Pinon, 1971; Tosh et  al., 1982; Dave, 1993; Robinson and Galán-Saúco, 2010) or in combination with other herbicides, such as ametryne or paraquat. When used at recommended concentrations, no phytotoxic symptoms are observed; but when high concentrations are used, browning and necrosis appear at leaf margins (Lassoudiere, 1972; Guillemot, 1975; Soto, 1985). Such symptoms were seen in Israel after simazine was applied at 2000 g of active ingredient (a.i.)/ha. Phytotoxicity symptoms are followed by a significant retardation of growth. Stronger phytotoxic symptoms occur in the presence of amitrol. The hormone herbicide 2,4-dichlorophenozyacetic acid (2,4-D) and its derivatives were used in the past as weed killers in banana plantations (Feakin, 1972; Ramadass et  al., 1980; Stover and Simmonds, 1987; Shanmugavelu et  al., 1992). Not just banana leaves but also pseudostems and suckers are known to be extremely sensitive to 2,4-D. Even in gaseous form, 2,4-D will damage banana suckers (Lassoudiere, 1972). The herbicide is a systemic auxin that affects the apical meristem, resulting in deformation of leaves and total death of the whole plant. When applied to banana plants after bunch emergence, 2,4-D interferes with normal fruit growth and may cause finger malformations. It has been replaced in banana plantations by paraquat, glyphosate and other contact herbicides. However, because of the high sensitivity of banana to 2,4-D, it is sometimes used for pruning unwanted suckers (Plate 11.12) (Chundawat and Patel, 1992). It is specifically suited for destroying banana plants in plantations affected by Moko bacterial wilt, as tool-transfer of the inoculum is avoided. Severe damage may result if it is used in confined spaces in screenhouses or glasshouses.

Disinfectants Disinfectants used in modern banana plantations as a part of a bio-security strategy do not normally have direct connection to the plants. However, caution is needed if they are added to

Plate 11.12.  Twisting, breaking and splitting of the leaf sheaths of a Cavendish sucker caused by the herbicide 2,4-D used for de-suckering (photo: E. Lahav, VI).

the irrigation water. Chlorine compounds that produce HClO ions are commonly used to keep the drip irrigation water in the water supply pipes clean of organic contamination. The injection of the chlorine is automatically controlled, but if an excess is erratically injected and reaches the banana root system, it may cause marginal chlorosis and necrosis in the leaves of suckers (Plate 11.13). The damaging effects only occur for a short time as the chlorine is neutralized by the soil organic matter. Chlorine is also used in nurseries to arrest the spread of pathogenic bacteria among potted plantlets derived from tissue culture. When the concentration of HClO ions in the irrigation water exceeds 6 ppm, leaves turn chlorotic. No new damage is evident after the concentration is brought back to the 6 ppm level (Y. Israeli, Israel, 2016, unpublished information).

De-Suckering Agents In young plantations established from plants derived from tissue culture, a relatively large number of suckers are produced. These need to be pruned. Chemicals used for killing suckers, such as 2,4-D and kerosene, may cause damage when applied at too high a concentration (Stover and Simmonds, 1987; Chundawat and Patel, 1992; Robinson and Galán-Saúco, 2010). If too much kerosene paraffin is used, the corm of the mother plant might be injured and the incidence of pathogenic bacteria may increase.

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Plate 11.13.  Toxicity symptoms on a leaf of a Cavendish cultivar caused by an excess of chlorine in water applied in a drip irrigation system (photo: Y. Israeli, JVBES).

Containers used for herbicides should not be used for the storage of de-suckering chemicals. Even small amounts of paraquat or glyphosate residues in kerosene injected into suckers might significantly affect the mother plant and result in the loss of the bunch.

Table 11.2.  Summary of symptoms on banana caused by excess minerals. Symptoms Description of on symptoms Petioles Leaf

Injury Caused by Elements in Toxic Concentrations Symptoms caused by excess of various elements are summarized in Table 11.2.

Sodium and Chlorine The sensitivity of banana to salinity or sodium chloride (NaCl) toxicity depends on environmental conditions, especially soil characteristics and evaporative demand. Problems do occur in India, Pakistan, Philippines, Haiti, Dominican Republic, Ecuador, Brazil, Colombia, Jamaica, Israel, Jordan, Egypt, Morocco and the Canary Islands. Dunlap and McGregor (1932) found that plants grew satisfactorily at 100–500 ppm total soluble

Fruit

Roots

Blue coloration Marginal browning followed by necrosis Irregular chlorosis followed by necrosis Marginal chlorosis followed by necrosis Marginal blackening followed by necrosis Chlorotic stripes Water-soaked lesions followed by necrosis Not filled Weak bunch, widely spaced hands, long peduncles Growth inhibited Club-like root tips

Element Magnesium Salinity (sodium chloride) Magnesium

Sodium, boron, calcium Iron, manganese Arsenic Fluorine

Arsenic Nitrogen

Copper Aluminium



Chemical Injury to Banana 537

salts, whereas plants and fruit were visibly ­affected from 500 to 1000 ppm. Above 1000 ppm, plants were stunted or dead. Saline soils produce marginal chlorosis, leading to necrosis (Plate 11.14), stunted growth and thin, deformed fruit. The adverse effect of salinity was demonstrated in some parts of Israel where banana plants were irrigated with water containing chlorides at 500–600 ppm. When the salty water was replaced by fresh water, the salts were leached from the soil and a significant improvement of growth and production was recorded. Under saline conditions, sucker growth is reduced and fruits do not fill. Plants growing very near to the ocean in areas affected by sea spray also show signs of salt toxicity (Plate 11.15). The effect of salinity on banana is assumed to be because of an increase in osmotic pressure of the soil solution, which results in difficulties in water uptake. The banana plant may be able to stand higher-than-normal salinity levels when evaporative demand is not too high, the water-supply is adequate and the drainage system is functioning well (Ben-Shalom et al., 2014). However, the plant may be adversely affected when evaporative demand increases or the water-­ supply decreases. Salinity is conventionally managed by increasing the amount of irrigation water, thus

leaching the salts from the rhizosphere to the deeper salt layers. However, leached salts can reach groundwater reservoirs, thus contributing to the deterioration of both soil and water quality. In Israel, the removal of excess salt from the water before it reached the field by means of desalination resulted in 56% less water having to be applied. This strategy increased bunch weight by 26–30%, improved fruit quality and decreased the salt load of the groundwater (­Silber et al., 2015). In most cases, it is difficult to distinguish between the effects of general salinity, excess sodium (Na) and excess chlorine (Cl). Shapira et al. (2009) described the symptoms of sodium toxicity as beginning as marginal chlorotic spots and then turning into a band of continuous marginal chlorosis, which became necrotic in older leaves. Eventually about one-third of the leaf could affected be affected. Excess sodium adversely ­ growth and yield and caused flowering to be ­delayed for 35 days. There was also a 31% drop in bunch weight and a 23% decrease in finger weight (Israeli et al., 1986). The specific negative effect of sodium on production of banana has also been demonstrated in the Canary Islands (Palacios et al., 2000). A significant accumulation of sodium (but not chlorine or potassium) was reported in affected leaf margins (Shapira et  al., 2009) ­

Plate 11.14.  Marginal necrosis of a leaf of a Cavendish cultivar caused by salt toxicity in Santa Marta, Colombia (photo: E. Lahav, VI).

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Plate 11.15.  Salt damage to leaves of Cavendish cultivar grown near the ocean on the island of Tenerife, Canary Islands (photo: Y. Israeli, JVBES).

(Plate 11.16). A combination of highly ion-­ selective bundle sheath cells around vascular tissue and high rates of guttation facilitated the disposal of excess sodium from the lamina. In the absence of guttation, the lamina margins were sacrificed by the plant in order to maintain a high photosynthetic potential in the remaining healthy tissue. Thus the distinct anatomy of the marginal vein in the banana plays a major role in the accumulation of sodium in the margins, the latter serving as a ‘dumping site’ for this toxic element. Under saline conditions, sodium levels in the lamina and petiole may change only slightly, Plate 11.16.  Early symptoms of sodium toxicity in a while sodium levels in the roots greatly increase leaf of a Cavendish cultivar (photo: Y. Israeli, JVBES).



Chemical Injury to Banana 539

as a result of selective transport. It has been suggested that sodium is toxic in the lamina at a concentration of 0.5% (Adinarayana et  al., 1986), but the roots seem to be a better indicator for sodium and chlorine levels. Tentative toxicity levels in the roots are 1.0% for sodium and 3.3% for chlorine (Israeli et al., 1986). When the soil exchangeable sodium percentage is high, water permeability is restricted. This condition might be corrected by gypsum or sulfur applications (Richards, 1954). Soils in S ­ anta ­Marta, Colombia, were treated this way and sodium toxicity damage was remarkably ­ reduced. ­Under saline conditions, it is also recommended to fertilize with potassium nitrate (KNO3) or sulfate (K2SO4) instead of potassium chloride (KCl).

Calcium A case of marginal leaf necrosis (Plate 11.17) resulting from calcium (Ca) in excess of 4% dry weight was reported from the Jordan Valley, ­Israel (Israeli, 1981).

Manganese Worldwide, manganese (Mn) excess is considered to be a greater problem than manganese

deficiency. Butler (1960) reported experiments in Jamaica where 20 kg of manganese/ha/year reduced yields by 5%. However, in sand-culture experiments, high levels of manganese have not reduced yield (Charpentier and Martin Prevel, 1965; Turner and Barkus, 1983). Black spots and marginal necrosis on leaves have been associated with high concentrations of manganese in Australia (Lahav and Turner, 1983). Toxic concentrations may be the result of natural high concentrations of available manganese in the soil or from a build-up of the element applied in fungicides, such as mancozeb. However, the tolerance of banana to high concentrations of manganese in the soil solution is high. Manganese toxicity observed in the field may be attributed to indirect effects, such as low calcium, magnesium and zinc availability in acid soils or waterlogging, rather than to a high concentration of manganese. High manganese levels in the soil are corrected by the application of lime.

Iron A black, necrotic marginal scorch on older leaves has been associated with iron (Fe) concentrations of up to 800 ppm in the Canary Islands (Ben-Meir, 1979; Lahav and Turner, 1983)

Plate 11.17.  Marginal necrosis of a leaf on a plant of a Cavendish cultivar – a symptom of calcium toxicity in the Jordan Valley, Israel (photo: Y. Israeli, JVBES).

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(Plate 11.18). Marginal scorching can sometimes be found in fertile plantations, but it is not necessarily an indication of a problem, unless associated with severe necrosis.

Plate 11.18.  Marginal scorch on a leaf of a Cavendish cultivar caused by excess of iron in Côte d’Ivoire (photo: Y. Israeli, JVBES).

Boron Boron (B) toxicity is quite rare. It may be found when recycled water is used for banana irrigation. When recycled water was used for irrigation in Israel for a number of years, the boron concentration in the water taken up by banana rose from 0.5 mg/l to 6.0 mg/l, which led to boron levels in the leaves above 1000 ppm. This reduced pseudostem height, bunch weight and number of bunches per hectare (Ben-Shalom et al., 2014). When boron is by regulation excluded from laundry powder ­detergents, its level in recycled water is much reduced. Another source of excessive leaf boron is an exaggerated boron application in response to a deficiency. A few grams of borax per plant are usually sufficient to correct a deficiency. With higher dosage, and since the boron is not easily leached, the concentration in the root zone may be high and symptoms of toxicity occur (Plate 11.19). Paling of the leaf margins and necrosis of leaves have been reported as in plants with more than 850 ppm of boron in the leaf lamina (Lahav and Turner, 1983). Shapira et al. (2012) studied the transport of boron in the banana lamina. They found that excess boron remains confined to the vascular system and is not translocated to the lamina

Plate 11.19.  Marginal necrosis of leaves of a Cavendish cultivar caused by the application of too much boron (photo: Y. Israeli, JVBES).



Chemical Injury to Banana 541

mesophyll, because of selective transport through the enclosing bundle sheath cell layer. By this mechanism, excess boron is directed towards the leaf margins and is effectively disposed of in the guttation fluid. Impaired guttation resulted in high boron toxicity damage.

Copper The accumulation of copper (Cu) in some acid soils of Central America, as a result of applications of Bordeaux spray for Sigatoka leaf spot control from the mid-1930s to the late 1950s, has affected root growth of cultivars in the Cavendish subgroup (AAA). Copper injury is not evident when the crop is first planted, but appears after the second or third cycle and becomes worse as the plantation ages (Stover, 1972). This toxicity is believed to result from: (i) the uptake of copper by the roots and its release when the roots die, which results in an accumulation of the element in the root zone close to the rhizome; (ii) a gradually increasing soil acidity in the surface soil; and (iii) a ‘high mat’ condition (when the base of the rhizome is only 5–10 cm below the soil surface 3–4 years after planting). Copper levels in the top 5–10 cm of the root zone of affected plants varied from 600 ppm to 2000 ppm and pH from 5 to 6. Root injury was reduced after deep ploughing to dilute copper concentrations in the upper soil layer and the application of lime to raise the pH.

Aluminium Aluminium (Al) is not an essential element for growth, but can cause toxicity problems, especially when the soil pH is below 4.0 (Godefroy et  al., 1978; Lopez and Espinosa, 1995). The main symptom of aluminium toxicity in banana is reduced root growth (Lassoudiere, 2007) and deformed roots (Lopez and Espinosa, 1995). Aluminium uptake by banana roots significantly decreases the banana plant’s transpiration rate, which in turn decreases the uptake of calcium and magnesium. The toxicity of aluminium significantly reduces dry matter, stunts the roots and induces magnesium deficiency (Rufyikiri et al., 2000b, 2001). Aluminium adsorption on root exchange sites is one of the mechanisms that can be responsible for aluminium toxicity in banana. Plants with mycorrhizae exhibit

greater resistance to aluminium toxicity than those that do not have beneficial fungal associations with their roots (Rufyikiri et  al., 2000a, 2003). Matsumoto and Yamaguchi (1990) selected aluminium-tolerant variants from gamma-­ irradiated protocorm-like bodies of ‘Nanicão’ (AAA, Cavendish subgroup) growing in vitro in high-aluminium liquid medium at low pH that simulated natural conditions of aluminium toxicity. One variant showed higher stress tolerance and was transferred to the field for evaluation. However, no further results have been reported.

Arsenic Arsenic (As) toxicity was reported by Fergus (1955) in Australia. Banana leaves had chlorotic stripes and bunch development was poor. An analysis of affected plants showed 0.25–2.00 ppm of arsenic as dry leaf matter, while healthy plants had 0–0.50 ppm. Herbicides were thought to be the source of the excess arsenic. The condition can be overcome by spreading around each plant 50–100 kg of clean soil, which adsorbs arsenic and renders it unavailable to banana.

Fluorine Symptoms of toxicity resulting from excess airborne fluorine (F) have been reported in Taiwan (Su et  al., 1978). Symptoms were described as typical marginal scorching, characterized by water-soaked lesions that tuned necrotic at the leaf margin (Plate 11.20). The content of ­fluorine

Plate 11.20.  Water-soaked leaf spots on banana developing into necrotic lesions as a result of fluorine toxicity in Taiwan (photo: E. Lahav, VI).

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in leaves with symptoms was 117–2119 ppm, as compared with 36–82 ppm in leaves from plants without symptoms. The corresponding values in fruit from affected and unaffected plants were 450 ppm and 105 ppm, respective-

ly. Similar symptoms (without a specific indication of fluorine toxicity) were more recently observed on banana growing in an area near Guangzhou in China with heavy ­industrial air pollution.

References Achard, R. (1993) Weed control in banana plantations in Cameroon. Fruits 48, 101–105. Adinarayana, V.D., Balaguravaiah, Y., Narasimha, Rao, P. and Subba Rao, I.V. (1986) Potassium and sodium disorders in banana. Journal of Potassium Research 2, 102–107. Aguero, R.A. (1998) Recommended use of ranger herbicide (glyphosate) in successive cycles, did not affect the growth and yield of the banana plant. In: Arizaga, L.H. (ed.) Memories XIII Reunion ACROBAT, Guayaquil – Ecuador, Nov. 23–27 1998, pp. 293–305. Ben-Meir, J. (1979) A case of iron excess in Canary Is. Bananas. International Banana Nutrition Newsletter 1, 11–14. Ben-Shalom, Ch., Or, G., Levingart-Aycicegi, A., Yirmiahu, U., Tarchitzki, Ch. and Lahav, E. (2014) Response of banana irrigated with recycled wastewater to boron in the Western Galilee: Part C. Alon Hanotea 68(6), 54–57 [in Hebrew]. Brenes-Prendas, S. and Aguero-Alvarado, R. (2012) Toxicidad de herbicidas promisorios para el control de Dieffenbachia oerstedii en hijos de banana. Agronomia Mesoamericana 23(1), 47–53. Broadley, R., Chay-Prove, P., Rigden, P., Daniells, J., Treverrow, N. et al. (2004) Subtropical Banana Information Kit. Agrilink, your growing guide to better farming. Agrilink Series Manual QI04011. Department of Primary Industries, Queensland Horticulture Institute, Brisbane. Butler, A.F. (1960) Fertilizer experiments with the Gros Michel banana. Tropical Agriculture (Trinidad) 37, 31–50. Calpouzos, L. (1968) Oils. In: Torgason, D.C. (ed.) Fungicides: An Advanced Treatise, Vol. 2. Academic Press, New York, pp. 367–393. Calpouzos, L., Delfel, N.E., Colberg, C. and Theis, T. (1961) Relation of petroleum oil composition to phytotoxicity and sigatoka disease control on banana leaves. Phytopathology 51, 317–321. Charpentier, J.M. and Martin Prevel, P. (1965) Culture sur milieu artificiel. Carences atténuées ou temporaires en éléments majeurs, carence en oligo-elements chez le bananier. Fruits 20, 521–557. Charpentier, J.M. and Martin Prevel, P. (1968) Carences et troubles de la nutrition mineral chez le bananier. Guide de diagnostic pratique. Institut Français de Recherches Fruitières Outre Mer, Paris, 75 pp. Chundawat, B.S. and Patel, N.L. (1992) Studies on chemical desuckering in banana. Indian Journal of Horticulture 49, 218–221. Constantinides, L.N and McHugh, J.J. Jr (eds) (2003) Pest Management Strategic Plan for Banana Production in Hawaii. Workshop summary. Honolulu, Hawaii, 71 pp. Cronshaw, D.K. (1987) The role of calixin in phytotoxic and stress-induced symptoms on banana leaves. In: Proceedings of the 7th ACORBAT Conference, 1985, San José, Costa Rica. Centro Agronomico Tropical de Investigación y Enseñanza (CATIE), Turrialba, Costa Rica, pp. 171–173. CSFRI (1983) Using glyphosate to kill bananas in Panama wilt-affected plantations. Citrus and Subtropical Fruit Research Institute Information Bulletin 131, 9. Dave, B. (1993) Weed control in banana plantations of Martinique. Fruits 48, 95–99. Dunlap, V.C. and McGregor, J.D. (1932) The Relationship Between Soil Alkalinity and Banana Production in St. Catherine District, Jamaica. United Fruit Company Bulletin 45. Escorriola, J.M., Arce, J. and Beltran, A. (1979) Oxyfluorfen: nueva herramienta con gran potencial herbicida para la industria bananera. In: Proceedings of the 4th ACORBAT Conference, 1979, Panama. Union de Paices Exportadores de Banano (UPEB), Panama, pp. 267–268. Feakin, S.D. (1972) Weeds. In: Feakin, S.D. (ed.) Pest Control in Bananas. PANS Manual No. 1. Centre for Overseas Pest Research, Foreign and Commonwealth Office, Overseas Development Administration, London, pp. 5–14.



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Fergus, I.F. (1955) A note on arsenic toxicity in some Queensland soils. Queensland Journal of Agricultural Science 12, 95–100. Galán-Saúco, V. (1992) Control de malas hierbas. In: Los Frutales Tropicales en los Subtropicos II: ­Platano (Banano). Ediciones Mundi-Prensa, Madrid, pp. 87–88. Godefroy, J., Lassoudiere, A., Loissois, P. and Penel, J.P. (1978) Action du chaulage sur les caractéristiques physico-chimiques, et la productivité d’un sol tourbeux en culture bananière. Fruits 33, 77–90. Gonzalez, S. and Piedrahita, W. (1994) Glufosinate: new molecule for the weed control on banana. In: Proceedings of the 10th Conference ACORBAT, 1991, Villahermosa, Mexico. Corporation Bananera Nacional (CORBANA), San José, Costa Rica, pp. 117–122. Guillemot, J. (1975) Tests sur l’efficacité et la phytotoxicité de quelques herbicides en bananeraie. Fruits 30, 75–81. Irizarry, H. (1987) Intensive plantain production in the humid mountains of Puerto Rico. In: Proceedings 3rd Meeting of the International Association for Research on Plantain and Bananas, Abidjan, Ivory Coast, 1985, pp. 55–59. Isaac, W.A.P., Brathwaite, R.A.I. and Ganpat, W.G. (2012) Weed management challenges in fairtrade banana farm systems in the Windward Islands of the Caribbean. In: Alvarez-Fernandez, R.Dr. (ed.) Herbicides – Environmental Impact Studies and Management Approaches. InTech, Rijeka, Croatia, pp. 209–222. Israeli, Y. (1981) A case of calcium excess in Israeli bananas. International Banana Nutrition Newsletter 3, 11–12. Israeli, Y., Lahav, E. and Nameri, N. (1986) The effect of salinity and sodium adsorption ratio in the irrigation water, on growth and productivity of banana under drip irrigation conditions. Fruits 41, 297–301. Israeli, Y., Shabi, E. and Slabaugh, W.R. (1993) Effect of banana spray oil on banana yield in the absence of Sigatoka (Mycosphaerella sp.). Scientia Horticulturae 56, 107–117. King, N. (2016) Banana Best Management Practices – Environmental Guidelines for the Australian ­Banana Industry. Queensland Department of Agriculture and Fisheries, Brisbane, 142 pp. King, N. and Lindsay, S. (2014) Providing the Framework for Practice Change – Building an Australian Banana Industry Environmental Best Management Practices Guide. Queensland Government ­Department of Agriculture, Fisheries and Forestry, Brisbane, 18 pp. Kwa, M. and Ganry, J. (1990) Utilisation agronomique des vitroplants de bananier. Fruits, Numéro special, 107–111. Lahav, E. and Turner, D.W. (1983) Banana Nutrition. Bulletin 7, International Potash Institute, Worblaufen-­Bern, Switzerland, 62 pp. Lassoudiere, A. (1972) Utilisation de herbicides en culture bananiere. Etude bibliographique. Fruits 27, 87–105. Lassoudiere, A. (2007) La Bananier et sa Culture. Quae Editions, Versailles, France, 383 pp. Lassoudiere, A. and Pinon, A. (1971) Indications préliminaires sur des essais de desherbage chimique en bananeraie. Fruits 26, 333–348. Liu, L.C. and Rodriguez-Garcia, J. (1988) Optimum time interval and frequency of glyphosate application for weed control in plantain (Musa sp.). Journal of Agriculture of the University of Puerto Rico 72, 297–300. Liu, L.C. and Santiago-Cordova, M. (1991) Persistence of three herbicides in a drip-irrigated banana field. Journal of Agriculture of the University of Puerto Rico 75, 19–23. Liu, L.C. and Singmaster, J.A. (1990) Herbicide drift control in plantains and taniers. Journal of Agriculture of the University of Puerto Rico 74, 471–475. Liu, L.C., Fernandez-Horta, D. and Santiago-Cordova, M. (1985) Diuron and ametryn runoff from a plantain field. Journal of Agriculture of the University of Puerto Rico 69, 177–183. Lopez, A.M. and Espinosa, J.M. (1995) Manual de Nutricion y Fertilizacion del Banano. Instituto de la Potasa y el Fosforo, Quito, Ecuador, 82 pp. Marie, P., Dave, B. and Cote, F. (1993) Use of banana vitroplants in the West Indies: assets and constraints. Fruits 48, 89–94. Matsumoto, K. and Yamaguchi, H. (1990) Selection of aluminium-tolerant variants from irradiated ­protocormlike bodies in banana. Tropical Agriculture (Trinidad) 67, 229–232. Mishra, A.K. and Das, P.C. (1984) Weed control in banana. South Indian Horticulture 32, 61–64. Nelson, S.C., Ploetz, R.C. and Kepler, A.K. (2006) Musa species (banana and plantain) Musaceae (­banana family). Species Profiles for Pacific Island Agroforestry, 1–33.

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Newley, P., Akehurst, A. and Campbell, B. (2008) Banana Growing Guide – Cavendish Bananas. NSW Department of Primary Industries, Orange, New South Wales, 28 pp. Palacios, M.P., Haman, D.Z., Del-Nero, E., Pardo, A. and Pavon, N. (2000) Banana production irrigated with treated effluent in the Canary Islands. Transactions of the American Society of Agricultural ­Engineers 43(2), 309–314. Pattison, T., Smith, L., Moody, P., Armour, J., Badcock, K.A. et  al. (2005) Banana root and soil health (BRASH) project – Australia. In: Turner, D.W. and Rosales F.E (eds) Banana Root System: towards a better understanding for its productive management. Proceedings of an International Symposium. International Network for the Improvement of Banana and Plantain, INIBAP, Montpellier, France, 149–165. Perkins, G.R. (1990) Basta – a new herbicide for horticulture. In: Proceedings of the 9th Australian Weeds Conference, pp. 544–547. Ramadass, R., Vaithialingam, R., Bhakthavathsalu, C.M. and Veerannan, L. (1980) Control of weeds in banana with the aid of herbicides. In: Proceedings of the National Seminar on Banana Production and Technology, Tamil Nadu, India. Tamil Nadu Agricultural University, Coimbatore, India, pp. 93–97. Richards, L.A. (ed.) (1954) Diagnosis and Improvement of Saline and Alkali Soils, Agriculture Handbook No. 60. USDA, Washington, DC, 160 pp. Robinson, J.C. and Galán-Saúco, V. (2010) Bananas and Plantains, 2nd edn. Crop Production Science in Horticulture 19. CAB International, Wallingford, UK, 311 pp. Rufyikiri, G., Declerck, S., Dufey, J.E. and Delvaux, B. (2000a) Arbuscular mycorrhizal fungi might alleviate aluminium toxity in banana plants. New Phytologist 148, 343–352. Rufyikiri, G., Dufey, J., Nootens, D. and Delvaux, B. (2000b) Effect of aluminium on bananas (Musa spp.) cultivated in acid solutions. I. Plant growth and chemical composition. Fruits 55, 367–379 Rufyikiri, G., Nootens, D., Dufey, J.E. and Delvaux, B. (2001) Effect of aluminium on bananas (Musa spp.) cultivated in acid solutions. II. Water and nutrient uptake. Fruits 56, 3–14. Rufyikiri, G., Genon, J.G., Dufey, J.E. and Delvaux, B. (2003) Competitive adsorption of hydrogen, calcium, potassium, magnesium, and aluminum on banana roots: experimental data and modeling. Journal of Plant Nutrition 26, 351–368. Sankar, A.V. (2014) Banana bunch care and methods to maximize bunch size. Available at: http://­tcbanana. blogspot.co.il/2014/12/banana-bunch-care-development.html Serrano, E. (2005) Banana ban acidification in the Caribbean coast of Costa Rica and its relationship with increased aluminium concentrations. In: Turner, D.W. and Rosales F.E (eds) Banana Root System: towards a better understanding for its productive management. Proceedings of an International Symposium, San Jose, Costa Rica. INIBAP, Montpellier, France, pp. 142–148. Shanmugavelu, K.G., Aravindakshan, K. and Sathiamoorthy, S. (1992) Weeding. In: Banana Taxonomy, Breeding and Production Technology. Metropolitan Book Co., New Delhi, India, pp. 97–100. Shapira, O., Khadka, S., Israeli, Y., Shani, U. and Schwartz, A. (2009) Functional anatomy controls ion distribution in banana leaves: significance of Na+ seclusion at the leaf margins. Plant, Cell and Environment 25, 476–485. Shapira, O., Israeli. Y., Shani, U. and Schwartz, A. (2012) Salt stress aggravates boron toxicity symptoms in banana leaves by impairing guttation. Plant, Cell and Environment 36, 275–287. Silber, A., Israeli, Y., Elingold, I., Levi, M., Levkovitch, I., Russo, D. and Assouline, S. (2015) Irrigation with desalinated water: a step toward increasing water savings and crop yields. Water Resources Research 51, 450–464. Soto, M.B. (1985) Manejo y control de las malas hierbas. In: Cultivo y Comercializacion de Banano. Litografia e Impresa LIL, San José, Costa Rica, pp. 242–265. Stover, R.H. (1972) Banana Plantain and Abaca Diseases. Commonwealth Mycological Institute, Kew, UK, 316 pp. Stover, R.H. and Simmonds, N.W. (1987) Bananas, 3rd edn. Longman, Scientific and Technical, Harlow, UK, 468 pp. Su, H.J., Ko, W.H., Chuang, S.Y., Huang, M.T. and Hwang, S.C. (1978) Etiological Studies on Marginal Scorch of Banana, with Special Reference to Air-polluted Fluoride Associated with the Disease. Banana Research Institute, Special Issue No. 21, Taiwan Banana Research Institute, Pingtung, Taiwan, 20 pp. Tezenas du Montcel, H. (1987) Weeding. In: Plantain Bananas. Macmillan Publishers, Basingstoke, UK, pp. 51–53. Tosh, G.C., Mohanty, D.C. and Nanda, K.C. (1982) Herbicides for the control of weeds in interplanted pineapples and bananas in India. Tropical Pest Management 28, 431–432.



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Turner, D.W. and Barkus, B. (1983) The uptake and distribution of mineral nutrients in the banana in response to supply of K, Mg and Mn. Fertilizer Research 4, 89–99. Velez-Ramos, A. and Vega-Lopez, J.A. (1977) Chemical weed control in plantains (Musa acuminata × Musa balbisiana, AAB). Journal of Agriculture of the University of Puerto Rico 18, 411–416. Venero, R. (1980) Chemical weed control in banana and its influence on yields. Cultivos Tropicales 2, 127–137. Weis, M., Keller, M. and Ayala, V.R. (2012) Herbicide reduction methods. In: Alvarez-Fernandez R.Dr. (ed.) Herbicides – Environmental Impact Studies and Management Approaches. InTech, Rijeka, Croatia, pp. 95–120.

12 

Genetic Abnormalities of Banana1 Y. Israeli and E. Lahav

Introduction Being inter- or intraspecific hybrids, mostly polyploid and almost totally sterile, spontaneous vegetative mutations have much contributed to the diversity of edible banana (Champion, 1967; Stover and Simmonds, 1987). Mutations occur in all cultivars and may change plant stature, leaf morphology, foliage and fruit pigmentation, waxiness, hairiness, bract and male flower persistence and finger length and shape. They can also result in the absence of female flowers or the male part of the inflorescence. Simmonds (1962) estimated that the rate of mutation in banana was 1–2 per 1 million, while Daniells and Smith (1993) thought spontaneous mutation rates were in the order of 1/100,000. Most mutants are agronomically inferior and affected plants are discarded when they are identified. Mutants may be characterized not only by morphological abnormalities, such as extreme dwarfism, abnormal leaf emission and a pronounced mosaic, but also by physiological traits, such as a difficulty in iron uptake. Some are kept in botanical collections and gardens as curiosity, but most are eliminated. Nevertheless, the rare beneficial somatic mutations in the ­edible banana have created the genetic diversification needed for the selection of new cultivars (Heslop-Harrison and Schwarzacher, 2007;

1

De Langhe et al., 2009: Perrier et al., 2011) and helped to create the 1000 cultivars known today (MusaNet, 2016). Since the introduction of the banana in vitro propagation techniques (Ma and Shii, 1972), the rate of off-type or mutated plants has greatly increased and has been reported from many different places in the world (Israeli et  al., 1995; Bairu et al., 2011). Off-type frequency in banana plants derived from tissue culture can vary from an extremely high 91% (Daniells and Smith, 1993) and 25% (Stover, 1987) to as low as 3% (Hwang and Ko, 1987) or 1% (Arias and Valverde, 1987). Variable rates of off-types have also been reported in plantain (Sandoval et al., 1991; Vuylsteke et al., 1991; Krikorian et  al., 1993). Since awareness of the problem has much increased, the appearance of mutated plants is now much reduced. Off-types might differ permanently or temporarily from the source plants. The latter, the result of an epigenetic effect, is characterized by a non-heritable change and is reversible. The permanent off-type, referred to as a somaclonal variant, is heritable and is an expression of pre-­ existing variation in the source plant or is due to the creation of de novo variation via variable genetic mechanisms (Larkin and Scowcroft, 1981). More general information about somaclonal variation, its occurrence and causes may be found in reviews by Scowcroft (1984), Lee and

  This chapter is dedicated to the memory of Dr Oded Reuveni.

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Phillips (1988), Bajaj (1990), Swartz (1991), Côte et al. (1993), Jain (2001) and Strosse et al. (2004). The causes of somaclonal variation in banana are only partially understood (Reuveni and Israeli, 1990; Reuveni et al., 1996; Sahijram et al., 2003; Krishna et al., 2016). The identification of an off-type phenotype as a somaclonal variant requires observations over several ratoon cycles (Scowcroft, 1984). The type and rate of variation are specific to the genotype (Stover, 1987; Smith, 1988; Robinson and Nel, 1989; Israeli et al., 1991; Vuylsteke et al., 1991; Newbury et al., 2000; Matsumoto et al., 2006) and this is true not only for different cultivars, but also for different selections of the same cultivar (Israeli et al., 1996). The micropropagation procedure itself, including number of transplanting cycles, type of explant used and type and concentration of hormones used, may also affect the rate of somaclonal variation (Smith, 1988; Drew and Smith, 1990; Reuveni and Israeli, 1990; Israeli et al., 1995; Reuveni et al., 1996; Bairu et al., 2006; Shirani et al., 2009; Bidabadi et al., 2010). During the period when micropropagation techniques were being developed, the high rate of off-type was a major concern. Improvements in laboratory procedures, such as the selection of stable source plants, limiting the number of transplanting cycles in culture and the number of explants produced from a single primary explant to about 1000, minimizing the use of adventitious buds and limiting the concentration of specific plant hormones in the medium, as well as identification and elimination of off-types during acclimation and nursery stages, resulted in the rate of off-types being reduced to less than 5%. This rate is regarded as commercially acceptable and it has become common practice for tissue culture-derived plants to be used as the planting material of choice by commercial producers worldwide. Small growers in Asia, Africa and Central and South America also have greater access to tissue-cultured banana (Kodim and Zapata-Arias, 2001; Al-Amin et al., 2009; Kahangi, 2010; Nijuguna et  al., 2010; Roy et al., 2010; Lule et al., 2013); Some banana breeders have realized that the variation produced in vitro may also be of use in genetic improvement. Variation rates have been increased by using gamma radiation or chemical mutagens (colchicine, ethyl methanesulfonate) (Jain, 2010; Chen et  al., 2013; Demtsu et  al.,

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2013) and material then screened for required traits. Success in these endeavours has so far been limited. However, the utilization of somaclonal variation resulting from tissue culture procedures has resulted in clones of ‘Pei-Chiao’ (AAA, Cavendish subgroup) being identified and selected that have more field resistance to tropical race 4 of Fusarium oxysporum f. sp. cubense (Hwang and Ko, 2004). These clones, developed in Taiwan, are currently allowing plantation ­banana cultivation to survive in areas where Cavendish cultivars can no longer be grown economically. Some successful genetic improvement of plantain using somaclonal variation has also been reported (Vuylsteke, 1998). The use of molecular tools to identify genetic variation, including traits that cannot be identified through visible phenotypic symptoms, is at present the centre of interest (Ray et al., 2006; Sahijram et  al., 2010; Bairu et  al., 2011). Oh et  al. (2007) have suggested that some specific regions of the Musa genome respond with higher rates of rearrangements and mutations to the stress conditions created during the tissue culturing process.

Variations in Stature Mutations of genes controlling banana stature are common and were originally described by Cheesman in 1933 and Champion in 1952. In the Cavendish subgroup (AAA), Cheesman described tall mutants of ‘Dwarf Cavendish’ and suggested that ‘Grand Nain’ was probably derived from a mutant of ‘Dwarf Cavendish’. Stover and Simmonds (1987) listed nine cultivars with dwarf counterparts. According to Richardson (1961), ‘Gros Michel’ (AAA) has mutated to distinct dwarf variant forms in Jamaica (‘Highgate’), Panama (‘Cocos’) and Honduras (‘Lowgate’). Richardson also reported the reverse mutation of dwarf ‘­Cocos’ to the tall ‘Gros Michel’. He noted that changes from short to tall plants are more frequent than changes from tall to short plants. Over a 3-year period, Richardson (1961) noted that reversions 50–70 tall appeared among 28,000 ‘Cocos’ mats. He believed that this change was controlled by the somatic loss of the dominant dwarf allele rather than by a reverse gene mutation. ‘Tall’ variants are less frequent with micropropagated cultivars ‘Williams’ and ‘Grand Nain’

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(AAA, Cavendish subgroup) than ‘dwarf ’ variants (similar, but not necessarily identical to ‘Dwarf Cavendish’), accounting for about 90% of all off-types (Israeli et al., 1991; Reuveni et al., 1996). The shorter stature selection ‘Nathan’ gives only extra dwarfs as height off-types (Reuveni et al., 1986). It seems that the genetic factor determining stature in the Cavendish subgroup is unstable in both conventional and micropropagated plants, but the pattern and rate of change differs in vitro and in situ and is clonal-dependent. However, under extreme diversification-inducing tissue culture conditions, ‘Grand Nain’ produces all known Cavendish statures from ‘Extra Dwarf ’ to ‘Lacatan’ (Stover, 1987, 1988). Height variation has also been recorded in off-types of the ABB group (Ventura et al., 1988). Matsumoto et al. (2006) studied the possible factors affecting the high frequency of dwarf offtypes during the micropropagation of ‘Nanicão’ and ‘Grand Nain’ (AAA, Cavendish subgroup) and concluded that high levels of cytokinin enhances changes to dwarfism, but the higher rate of multiplication of the shoots of the dwarf mutants in culture itself contributes to their increased proportion in the population during repeated subcultures. The ‘dwarf ’ variant is the most frequent ­micropropagation off-type in the Cavendish subgroup, but does not appear in micropropagated ‘Red’ (AAA). However, the mosaic-like leaf variant is found in both micropropagated Cavendish cultivars and ‘Red’, which indicates that different genomes differ in the type and rate of variants generated in culture (Vuylsteke et  al., 1988; ­Israeli et al., 1995). In the Cavendish subgroup, dwarfs are very similar to ‘Dwarf Cavendish’ and vary significantly from the original cultivar. Their height, leaf size and morphology, distance between petioles (Plate 12.1), bunch length, bunch mass and finger dimensions are smaller than those of the original cultivar. The rate of growth of dwarf plants is fast and they flower earlier than the original cultivars from which they are derived (Israeli and Nameri, 1985; Smith and Drew, 1990). There is no difference in the number of hands or in pseudostem and bunch stalk circumference. Some of the flowers have a persistent perianth and the male flowers and bracts are persistent on the male axis (Plate 12.2). Their fruits differ in size and shape from those of the original cultivars. All these are typical

c­haracteristics of ‘Dwarf Cavendish’. It was demonstrated that dwarf off-types show improved tolerance to photoinhibition induced by low temperature, as compared with the normal ‘­Williams’ (Damasco et  al., 1997). Most dwarfs produced by Cavendish clones resemble each other in their basic characteristics, whether or not they originated from different cultivars, propagation batches or production plots. Tissue culture-derived dwarfs can be detected during their growth period in the nursery, ­before transplanting to the field. Differences in height, distance between petioles, morphology of petioles and leaf size can be distinguished if all plants are grown under uniform conditions. When the height of normal plants from the ground to the base of the youngest petiole is 35–40 cm, dwarfs of ‘Grand Nain’ and ‘Williams’ are about 5 and 10 cm shorter, respectively. Because of the greater height difference, the ‘Williams’ dwarfs can be screened out more efficiently at this stage (Israeli et al., 1991). Efficient detection of dwarfs in the field, based on the above parameters, is achieved during periods of uniform growth. Differences in height, leaf index (length–width ratio) and distance between petioles is pronounced. The leaf index is less than 2.0 in more than 90% of dwarf ‘Williams’ plants and 2.2 or greater among normal ‘Williams’ plants (Israeli et al., 1991). Inflorescence parameters, such as shape and size of fingers and persistence of bracts and male flowers on the axis, are the most reliable means of characterizing dwarfs. Smith and Hamil (1993) showed that, in the Cavendish subgroup, dwarfs can be identified and removed when plantlets are 7 weeks old and 20 cm high. The best morphological characteristics for screening at this stage are length of lamina and petiole and the ratio between them. Reuveni (1990) demonstrated that dwarfs may be identified at the tissue-culture propagation stage by adding gibberellic acid (GA3) to the growth medium. Normal plants respond with intensive growth, while the reaction of dwarfs is very slow. The same response is achieved as a result of spraying GA3 after de-flasking (Damasco et  al., 1996a). Significant differences are found in endogenous GA3 levels between dwarf, normal and giant ‘Grand Nain’ (Sandoval et al., 1995). The level of GA3 was 3.6 times higher in the ­normal ‘Grand Nain’ than in the dwarf off-type



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Plate 12.1.  A ‘dwarf’ variant of Cavendish cultivar (left) that has arisen in tissue culture compared with a normal plant (right). Note the short internodes, the short petioles and the different leaf length–width ratio on the dwarf variant (photo: Y. Israeli, JVBES, and E. Lahav, VI).

and 4.6-fold in giant ‘Grand Nain’ as c­ ompared with the dwarf. The differences in GA20 were 4.6 and 7.3, respectively. Ortiz and Vuylsteke (1995) investigated the inheritance of dwarfism in plantain and concluded that a single recessive gene controls the trait. Banana cultivars and clones have been identified by modern methods of molecular analysis (Howell et al., 1994; Bhat and Jarret, 1995). These methods have also been used for identification of dwarf off-types. Damasco et  al. (1996b) identified a random amplified polymorphic DNA (RAPD) marker specific to the dwarf off-type produced during in vitro propagation of ‘New Guinea

Cavendish’ and ‘Williams’ cultivars. The marker provided a reliable means with which to detect dwarf variants in tissue culture. The detection of dwarf off-types in plantain using the RAPD technique is also reported (Ford-Lloyd et  al., 1993). Later, Damasco et al. (1998) reported the use of the sequence-characterized amplified region (SCAR)based marker for the early detection of dwarf offtypes in micropropagated Cavendish. However, the practical use of these tools, as well as additional molecular tools, in standard routine mass micropropagation seems quite limited. An ‘extra-dwarf ’ variant is an extremely low-stature plant about 1 m tall. The leaves are

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Plate 12.2.  Fruiting ‘dwarf’ variant of ‘Grand Nain’ (AAA, Cavendish subgroup) (left) compared with normal ‘Grand Nain’ (right). The dwarf variant has persistent perianths and bracts plus horizontal hands (photo: Y. Israeli, JVBES, and E. Lahav, VI).

greatly condensed, forming a ‘rosette’, and are very short and broad. The bunch stalk is very short and the bunch becomes choked in the pseudostem on emergence. As a result, the bunch tends to lie horizontally and is sometimes even orientated vertically. The fingers are very short and the fruit has no commercial value. These characteristics are typical of the ‘Extra-­ Dwarf Cavendish’ cultivar described by Stover and Simmonds (1987). The incidence of the extra-dwarf variant is very high in ‘Nathan’ (­Israeli selection of ‘Dwarf Cavendish’). Very few extra-dwarfs arise naturally in the Cavendish cultivars ‘Williams’ and ‘Grand Nain’. ‘Dwarf Parfitt’ (Plate 12.3), an ‘extra-dwarf ’ Cavendish

variant found in Australia, has been shown to be resistant to subtropical race 4 of Fusarium oxy­ sporum f. sp. cubense. Compared with ‘Williams’, it also has a higher photosynthesis rate and a better tolerance to low temperatures (Moore et al., 1993). The ‘extra-dwarf ’ variants, contrary to ­other stature variants, can be detected in tissue ­culture by their compact appearance and the formation of rosette-like leaves. They can be recognized and removed easily at the hardening and nursery stages (Israeli et al., 1991). The ‘J.D. Dwarf ’ is an off-type selection from tissue-cultured ‘Williams’ (Daniells, 2002). It is slightly shorter than ‘Williams’, but more important



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Plate 12.3.  ‘Extra-dwarf’ Cavendish variant ‘Dwarf Parfitt’ in Queensland, Australia. Note that the bunch has ‘choked’ on emergence (photo: QDPI).

is its sturdier pseudostem, which make it possible to carry fruit without support. This selection presents some advantages, but also some disadvantages, as is often the case with somaclonal variants. ‘Giant’ variants often arise when ‘Grand Nain’ is micropropagated, but in small numbers. The plants resemble the tall cultivars of the ­Cavendish subgroup, such as ‘Valery’ (Plate 12.4) or sometime even ‘Lacatan’. Giants can be easily detected in the nursery, as they are approximately 30% taller than normal plants (Plate 12.5) and are clearly visible in the field. It has been suggested that their low frequency is a result of slow in vitro multiplication in contrast to the

high multiplication rate of dwarf mutants (­Matsumoto et al., 2006). Giant variants are also rarely observed in conventionally propagated banana.

Abnormal Foliage The most common genetic foliage abnormality is the ‘mosaic’ or ‘mosaic-like’ variant, which is also called ‘Masada’ in Central America and South Africa. The variably thick, rubbery, narrow leaves, with different degrees of pale green mottling, may resemble banana mosaic disease to

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Plate 12.4.  A ‘giant’ tissue-culture ‘Valery’-type variant stands out in a plantation of normal ‘Grand Nain’ (AAA, Cavendish subgroup) (photo: Y. Israeli, JVBES).

the inexperienced eye. The mottling, which is ­irregular and not restricted by secondary veins, can be seen more clearly when the lamina is lit from behind. The upper leaf surface shows a pattern of bright spots and is covered with depressions and protuberances (Plate 12.6). In addition, a very narrow and irregular lamina, wavy leaf margins and longitudinal bright stripes along the midrib and petiole down to the leaf sheath are sometimes observed. The plants grow relatively slowly and inflorescence emergence is delayed. The number of hands is normal, but the peduncle is thin and the fingers are very small with no commercial value. Reuveni (1990) recorded exceptionally low numbers of

stomata on the mosaic variant leaf. Mosaic variants have appeared so far in all clones of the ­Cavendish subgroup and also in ‘Red’, but usually in small numbers. They can be detected at the nursery stage when tissue-cultured plants reach a height of 25–30 cm, with a leaf length exceeding 20 cm. The symptoms observed on the eighth to tenth leaves are more easily distinguished than those on the older leaves. One case only of a mosaic sucker originating from a normal mother plant was recorded in the Cavendish cultivar ‘Williams’ in conventionally propagated plants (Israeli et  al., 1991). The in vitro environment enhances the occurrence of these off-types.



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Plate 12.5.  A ‘giant’ tissue-culture off-type of ‘Grand Nain’ (AAA, Cavendish subgroup) at the end of the nursery stage (photo: Y. Israeli, JVBES).

An extreme mosaic with more pronounced characteristics is also known (Plate 12.7). It is very droopy in appearance and has very narrow laminae (only a few centimetres wide) and very intense mottling. The extreme mosaic can be ­detected at the hardening stage or immediately after potting. Mosaic plants have been found to be aneuploids (Reuveni et  al., 1986; Sandoval et  al., 1996). It is clear that the in vitro procedure promotes their development. A change in chromosome

number has also been reported for somaclonal variants of other species (Lee and Phillips, 1988). The mosaic-variant plants are similar, but not identical, to each other, which points to a difference in the level of aneuploidy, as confirmed by Sandoval et al. (1996). The extreme mosaic might have a larger number of extra chromosomes than the common mosaic variant. The connection between various off-types and chromosome number in meristematic cells of root tips of ‘Grand Nain’ was investigated by

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Plate 12.6.  Leaf of ‘mosaic’ variant of a Cavendish cultivar covered with irregular bright spots. The leaf surface is also covered with depressions and protuberances and the margins are wavy (photo: Y. Israeli, JVBES, and E. Lahav, VI).

Plate 12.8.  ‘Drooping-leaf’ variant of ‘Grand Nain’ (AAA, Cavendish subgroup) (photo: Y. Israeli, JVBES, and E. Lahav, VI)

Plate 12.7.  ‘Extreme mosaic’ variant of a Cavendish cultivar (photo: Y. Israeli, JVBES, and E. Lahav, VI).

Sandoval et  al. (1996). About 5% of cells in plants propagated conventionally had abnormal chromosome numbers, while 14% of cells were affected in normal in vitro-propagated plants.

Dwarf variants had 22% abnormal cells and ­mosaic variants 35%. A ‘drooping-leaf ’ variant was observed by Smith (1988) and Vuylsteke et  al. (1988) in plantain (AAB), and by Israeli et al. (1995) and Daniells et  al. (1999) in the Cavendish cultivar ‘Grand Nain’ (Plate 12.8). This variation is associated with slow growth, delayed flowering and agronomically inferior bunches. Bunches of the plantain variant were of the ‘False Horn’ type, but extremely small. The origin of this variation is suggested to be due to a high ploidy level. A  variant with upright leaves was found in ­Australia (Daniells et al., 1999) (Plate 12.9). Randomly scattered pale patches, without chlorophyll in the leaf lamina, characterizes the ‘variegated-leaf ’ variant. To the untrained eye, it can sometimes be mistaken for symptoms of cucumber mosaic virus. The variegation is irregular and differs from plant to plant. Sometimes it appears as red colour on a pale background. Variegation is very likely a sectorial chimera. A very



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Plate 12.9.  ‘Erect foliage’ variant of ‘Williams’ (AAA, Cavendish subgroup) (photo J.W. Daniells, QDAF).

pronounced variegated mother plant with both variegated and normal followers is illustrated in Plate 12.10. In some plants, variegation disappears after several months and the newly emerged leaves are symptomless and the followers normal. However, other variegated plants retain the symptoms and can produce variegated fruit (Plate 12.11). This type of variant occurs in both tissue-cultured and conventionally propagated plants of ‘Williams’ and ‘Grand Nain’ in Israel (Israeli et al., 1991). The first detection of variegated leaves depends on the intensity of symptoms. Extremely variegated plants can be detected in tissue-­cultured plants at the test-tube stage or at the very beginning of the hardening stage. Less intense symptoms can be detected at the nursery stage before transplanting to the field. Another type of leaf aberration is the ‘­deformed-lamina’ variant. The leaf blade is deformed or lobed (Plate 12.12) and in some cases the affected part has a bright mosaic. The deformed portion of the leaf is usually close to the petiole. This variant was found in in vitro plants of banana (Israeli et  al., 1991) and p ­lantain

Plate 12.10.  Pronounced variegation in the leaves of ‘Pelipita’ (ABB) growing in Nigeria. Note that the lead follower is not variegated, but the smaller sucker appears affected (photo: D.R. Jones, INIBAP).

(Vuylsteke et  al., 1996). It was only once recorded in conventionally propagated banana in Israel (Israeli et  al., 1991). Affected tissue-­ cultured plants can be detected easily in the nursery ­before planting in the field. In most cases, the deformation disappears gradually in the field, but in few cases it is inherited for many cycles. It is most probably a long-lasting epigenetic ­effect. An ‘iron-deficient’ variant was found in ‘Dwarf Cavendish’ plantation in the Jordan ­Valley, Israel, during the 1970s. This mutant has reduced ability for iron uptake and produces chronically yellow leaves unless iron-chelate is applied (Plate 12.13). The physiology of this abnormal variation has not yet been studied. Interestingly, it never occurred among micropropagated Cavendish. More rare foliage aberrations have been documented in Australia (Daniells et al., 1999). These were a yellow flecking of the lamina, chlorotic/

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necrotic patches, excessive anthocyanin pigmentation, choking, dark/shiny leaf bases, rolling of petiole wings, black streaking of midribs or leaf bases and striping of midribs or leaf bases.

Variations in Pseudostem Pigmentation

Plate 12.11.  ‘Variegated-leaf’ variant of a Cavendish cultivar with variegated leaf and fruit. The white tissue lacks chlorophyll (photo: Y. Israeli, JVBES, and E. Lahav, VI).

The ‘reddish pseudostem’ variant is characterized by a decrease in black pigmentation and the appearance of a reddish colour on the leaf sheaths and petioles (Plate 12.14). The red colour is pronounced, especially in young plants, either in the nursery or in the field. These plants also have bright green leaf sheaths, petioles, midribs and fruit peel. Another typical characteristic of this variant is an elongated funnel leaf, which does not open normally during the hot midsummer months. ‘Reddish pseudostem’ variants are quite rare in both conventional and in vitro plants. They are easily distinguished when they exceed a height of 30 cm in the nursery. Detection is more reliable under conditions of high light ­intensity, as o ­ pposed to partial shade, where normal pseudostems are also pale and diagnosis is more difficult. As a consequence, symptoms are obvious on plants in the field. Variants with a

Plate 12.12.  Leaves of a ‘deformed-lamina’ variant of a Cavendish cultivar (photo: Y. Israeli, JVBES, and E. Lahav, VI).



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Plate 12.13.  ‘Iron-deficient’ variants of ‘Dwarf Cavendish’ (AAA, Cavendish subgroup) (Photo: Y. Israeli, JVBES).

Plate 12.14.  ‘Grand Nain’ (AAA, Cavendish subgroup) with a reddish pseudostem (photo: Y. Israeli, JVBES).

reddish pseudostem produce satisfactory commercial fruit. The ‘black pseudostem’ variant is almost completely black (Plate 12.15). Coloration is also found in the petioles and central veins. The characteristic persists during many cycles (­ Israeli et  al., 1991). The ‘black pseudostem’ variant ­occurs in conventionally propagated plants, as well as in tissue-cultured plants. ‘Pisang ­Klutuk Wulung’ is a black-stemmed Musa balbisiana ­ from Indonesia. Other pseudostem pigmentation variants are greenish (hardly any anthocyanin in the lamina and leaf sheaths), striped to various degrees, black spotted (the fruit is also spotted) and red/green variation, which is common in the Red banana subgroup (AAA) (Israeli et  al., 1991). The ‘Red’ to ‘Green Red’ shift can be explained by the periclinal chimera structure of the mother plants (Stover and Simmonds, 1987). Waxiness is another pseudostem variation. Its inheritance was investigated by Ortiz et  al. (1995), who found that waxiness is controlled by a recessive gene, with a significant

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Plate 12.15.  ‘Grand Nain’ (AAA, Cavendish subgroup) with a black/dark-violet pseudostem. Coloration extends to petioles and midribs (photo: Y. Israeli, JVBES).

dosage effect. All the above-­mentioned off-types have also been found in conventional banana propagation, but at a very low rate. Additional rare pseudostem abnormalities are a slender-to-thick shift, a collapse before bunching, a traveller’s palm appearance and wax stripes (Daniells et al., 1999).

Chimeras Typically for long-term vegetatively propagated plants, banana presents chimeric variation (­Tilney-Basset, 1986; Swartz, 1991; Jain, 2010). The variegated leaves and the black-­violet pseudostem, as well as the deformed leaf variants described above, are clearly chimeras. Within a few cycles, some of the suckers of such plants will revert to the normal phenotype (Israeli et al., 1995). The chimeras are known, but rare, in conventional propagation and occur at a modest incidence and low rate with micropropagation (Israeli et al., 1991).

Plate 12.16.  Green/Red chimera of a bunch of ‘Red’ (AAA, Red subgroup) (photo: J.W. Daniells, QDAF)

In fruit, the condition whereby stripes of abnormally pigmented tissue stand out against the normal green background is most probably a chimera. They often consist of well-defined reddish or brown bands on the peel. Occasionally, part of an entire bunch is affected. Split peel is a similar fruit chimeric variation. The ‘Red’ banana is a well-known chimera. According to Stover and Simmonds (1987), the red peel of the fruit of this cultivar is a chimera consisting of a ‘red skin’ on a ‘green core’. An unusual chimeric bunch with red/green fruit is presented in Plate 12.16. Association between chimerism and somaclonal variation in banana was also suggested by Daniells and Smith (1991). Swarts (1991) and Reuveni et  al. (1993) suggested that the high rate of dwarfs in the Cavendish subgroup may also point towards the involvement of chimera dissociation. Cytochimeras are variants that contain cells with variable ploidy, such as polyploidy, aneuploidy, mixoploidy or other cells variations. It was demonstrated that even conventionally propagated banana sucker meristems contain cells with variable ploidy (Sandoval et  al., 1996). In vitro culture and especially



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­ utation induction (physical or chemical) inm creases the chimerism significantly (Roux et al., 2001) and efficient methods for chimera dissociation are needed. The use of embryogenic (single) cell suspension is one of the possible means to reduce cytochimeras (Roux et  al., 2004; Jain, 2010).

Inflorescence and Fruit Variations Inflorescence and fruit off-types are the most difficult of all to detect, since they can only be identified in the field at a late stage of plant development when their elimination causes economic loss. Most of the variants described below are common to conventional and in vitro propagated ­bananas and are quantitatively unimportant. Variations have been documented in flower, fruit and bunch characteristics (Stover and ­Simmonds, 1987; Côte et  al., 1993; Israeli et  al., 1995; ­Daniells et al., 1999). With a ‘persistent flower’ variant, the perianth of the female flower does not abscise, but remains attached to the ovary. There is no nectar secretion and bracts remain attached to the cushion and are delayed in their withering. The fruits are less curved and take an ‘open-hand’ form. A large part of the male axis is covered with residual male flowers. This mutant (Plate 12.17) is known in both tissue-cultured and conventionally propagated plants (Israeli, 1977). The ‘inflorescence with no female flowers’ variant (Plate 12.18) is caused by interference to the parthenocarpic development of the ovary. It is an off-type known in conventional plantations of ‘Dwarf Cavendish’, ‘Williams’ and ‘Grand Nain’, but is very rarely found in micropropagated plants. The suckers of such plants grow rapidly as a result of lack of competition for nutrients from developing fruit. The ‘French reversion’ variant in plantain is characterized by the appearance of a typical ‘French’ inflorescence in ‘False Horn’ plantain. The plant produces relatively small fruit and ­persistent intermediate flowers with the male bud present at maturity. All these traits are totally different from the original plant. In vitro-propagated plantains produce a high rate of this variation (Pool and Irizarry, 1987). Krikorian et al. (1993) suggested a chimera breakdown mechanism as an explanation for this high variation rate. The

Plate 12.17.  Persistent flowers and bracts of a mutant of ‘Williams’ (AAA, Cavendish subgroup). The hands are horizontal and the peduncle is straight (photo: J.W. Daniells, QDAF).

‘Monganga’-type is another variation in plantain, producing a degenerated ‘False Horn’ inflorescence (Vuylsteke et al., 1996). In the ‘split-finger’ variant, fingers split longitudinally on their external side (see Plate 8.9). Similar fruit has been found in conventionally propagated plants. Fingers in some ratoon bunches can be normal, which points towards a chimera dissociation. The ‘multi-bunching’ variant has multiple pseudostems, peduncles and male flower buds. This condition has been described several times (Cheesman, 1933; Champion, 1952, 1967; ­Nayar et  al., 1958; Gill, 1968; Stover and ­Simmonds, 1987). Davis (1984) observed about 35 different cases of abnormal bunching. In 1991, a variant of ‘Giant Cavendish’ with three bunches was found in a farmer’s field in s­outhern Taiwan and propagated in vitro (Tang, 1995). When plants arising from this tissue c­ ulture were planted in the field, 21.1% were observed to be multi-­ bunching variants. These variants were described as a single bunch with multiple male buds (Plate 12.19), multiple bunches on a ­single peduncle

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Plate 12.18.  Inflorescence of ‘Grand Nain’ (AAA, Cavendish subgroup) with aborted female flowers (photo: Y. Israeli, JVBES).

and multiple bunches on independent peduncles. In the ratoon crop, the number of multi-bunching variants rose to 61.1% and a new type with multiple pseudostems appeared. This variation suggests that the abnormal division of the meristematic tissue occurred at various stages of inflorescence development. The author mentioned that there was no economic advantage in the multi-bunching, since the total fruit weight was smaller than the weight of a single bunch. A similar experiment was conducted in Nigeria with ‘UNN’ plantain (Tenkouano, 2000). Reverse of the double to a single-bunch phenotype was observed and few plants underwent an additional dichotomization event to produce three bunches. It was suggested that multi-­ bunching may be due to random genetic events instead of a stable mutation. A bunch that bends horizontally and has widely spaced hands (Plate 12.20) has been observed on a tissue culture variant of ‘Williams’ (AAA, Cavendish subgroup). Other bunch and

Plate 12.19.  ‘Multi-bunching’ banana variant with multiple male buds arising from a single bunch (photo: E. Lahav, VI).

fruit variants are a long tapering bunch with no male bud (Plate 12.21), waxy fingers and blunter fruit tips. These variations appear mostly in the Plantain subgroup (AAB) and ABB group (Stover and Simmonds, 1987). Daniells et  al. (1999) have noted variants with pointed-tip fingers (which were more susceptible to bacterial finger-tip rot disease), severe fruit scarring, long peduncles, longer fruit, short fingers, incomplete fruit that does not fill, different-coloured male buds (Plate 12.22) and bunches in which the two top hands fail to fill. During recent years, a poor-fruit off-type of ‘Grand Nain’ has been identified in Israel (Plate 12.23). The plant is slow in growth and flowers late with a partially choked bunch, a thin and sometimes twisted peduncle and short ­nipple-tipped fruit. Petioles are choked at the upper part of the pseudostem. This variant initially occurred at a low frequency in many recently planted plantations, but incidence has increased and in an extreme case has reached 7% of the population (Gal Or,



Genetic Abnormalities of Banana

Plate 12.20.  Tissue-culture variant of ‘Williams’ (AAA, Cavendish subgroup) with widely spaced hands and a bunch tending to the horizontal (photo: D.R. Jones, QDPI).

2016, Israel, personal communication). The plantlets were produced by a tissue-culture laboratory working to the best-known standards. However, the problem is still not solved. There is an urgent need to revise the protocols to make sure that plants of this off-type will not be selected as mother plants for shoot-tip propagation.

Concluding Remarks This chapter’s descriptions of genetic abnormalities are derived from the documented information available on the subject. Most have been primarily off-types (natural and in vitro-induced) of ­cultivars in the Cavendish subgroup. This reflects the major economic importance of Cavendish cultivars, which account for almost 50% of the world’s total production of bananas (­Daniells et al., 2013).

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Plate 12.21.  ‘Tapering bunch’ tissue-culture variant of ‘Chinese Cavendish’ (AAA Cavendish subgroup) (photo: D.R. Jones).

Genetic abnormalities exist in cultivars, which are important food sources, in other subgroups. Many of these cultivars have not been routinely multiplied in vitro and it is possible that, when they are, new variants will arise. The off-types listed in this chapter are mainly deleterious. However, some important somaclonal variants have been identified that have greater resistance to disease, tolerate environmental stress conditions or have improved horticultural performance. Other variants, which could be of great economic importance, are awaiting discovery. The use of in vitro micropropagation methods to produce banana planting material faced some initial setbacks. The main reason was the initial high rates of mutations, which were later minimized. Improvements in the quality of tissue-­cultured plants and their ready availability at an attractive cost has resulted in a worldwide increase in their use. This, in turn, increases the need for good-­quality explant sources, which are

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Plate 12.22.  Yellow-coloured male bud of ‘Williams’ (AAA, Cavendish subgroup) (photo: J.W. Daniells, QDAF).

not easy to obtain. An average-sized laboratory that produces 10,000,000 plantlets/year will need to inoculate 10,000 explants in culture (1:1000 propagation rate). About 3000 fieldgrown mother plants that are kept under continuous supervision, including clonal fidelity and horticultural performance, are needed as sources of these explants. Mother plants must also be free of diseases that can be carried in tissue culture. This is achieved by selecting material from regions free of these diseases or by ­disease indexation in quarantine before use. Molecular tests to identify the potential risk of invisible genetic variations need to be further developed in order

Plate 12.23.  ‘Late shooting – poor bunch’ tissue culture variant of ‘Grand Nain’ (AAA, Cavendish subgroup), with short fruit, thin distorted peduncle and choked top leaves (photo: Y. Israeli, JVBES).

for these to become routinely applied. The production of first-rate planting material free of genetic aberrations is the interest of both plantlet propagators and banana producers.

Acknowledgement J.W. Daniells is thanked for his important contribution to this chapter.

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Gill, M.M. (1968) A note on dichotomy of the inflorescence in the plantain (Musa paradisiaca Linn.). Tropical Agriculture (Trinidad) 45, 337–341. Heslop-Harrison, J.S. and Schwarzacher, T. (2007) Domestication, genomics and the future for banana. Annals of Botany 100, 1073–1084. Howell, E.C., Newbury, H.J., Swennen, R.L., Withers, L.A. and Ford-Lloyd, B.V. (1994) The use of RAPD for identifying and classifying Musa germplasm. Genome 37, 328–332. Hwang, S.C. and Ko, W.H. (1987) Somaclonal variation of bananas and screening for resistance to Fusarium wilt. In: Persley, G.J. and De Langhe, E.A. (eds) Banana and Plantain Breeding Strategies, Proceedings of an International Workshop, Cairns, Australia. ACIAR Proceedings No. 21, Australian Centre for International Agricultural Research, Canberra, Australia, pp. 151–156. Hwang, S.C. and Ko, W.H. (2004) Cavendish banana cultivars resistant to fusarium wilt acquired through somaclonal variation in Taiwan. Plant Disease 88(6), 580–588. Israeli, Y. (1977) Deciduous and persistent flowers in the Cavendish banana. MSc thesis, Hebrew University of Jerusalem, Israel, 60 pp. [in Hebrew, with English summary]. Israeli, Y. and Nameri, N. (1985) Off-types of banana plants multiplied in vitro. In: Report on Observations and Experiments in Bananas in the Jordan Valley in the Years 1978–84. Banana Experimental Station, Jordan Valley, Israel, 24, 50–59 [in Hebrew]. Israeli, Y., Reuveni, O. and Lahav, E. (1991) Qualitative aspects of somaclonal variations in banana propagated by in vitro techniques. Scientia Horticulturae 48, 71–88. Israeli, Y., Lahav, E. and Reuveni, O. (1995) In vitro culture of bananas. In: Gowen S. (ed.) Bananas and Plantains. Chapman and Hall, London, pp. 147–178. Israeli, Y., Ben Bassat, D. and Reuveni, O. (1996) Selection of stable banana clones which do not produce dwarf somaclonal variants during in vitro culture. Scientia Horticulturae 67(3–4), 197–205. Jain, S.M. (2001) Tissue culture-derived variation in crop improvement. Euphytica 118, 153–166. Jain, S.M. (2010) In vitro mutagenesis in banana (Musa spp.) improvement. Acta Horticulturae 879, 605–614. Kahangi, E.M. (2010) The potential of tissue culture banana (Musa spp.) technology in Africa and the anticipated limitations and constraints. Acta Horticulturae 879, 281–288. Kodim, A. and Zapata-Arias, F.J. (2001) Low-cost alternatives for the micropropagation of banana. Plant Cell, Tissue and Organ Culture 66, 67–71. Krikorian, A.D., Irizarry, H., Cronauer Mitra, S.S. and Rivera, E. (1993) Clonal fidelity and variation in plantain (Musa AAB) regenerated from vegetative stem and floral axis tips in vitro. Annals of Botany 71, 519–535. Krishna, H., Alizadeh, M., Singh, D., Singh, U., Chauhan, N., Eftekhari, M. and Sadh, R.K. (2016) Somaclonal variations and their applications in horticultural crop improvement. Biotech 6(54), 1–18. Larkin, P.J. and Scowcroft, W.R. (1981) Somaclonal variation: a novel source of genetic variability from cell cultures for improvement. Theoretical Applied Genetics 60, 197–214. Lee, M. and Phillips, R.L. (1988) The chromosomal basis of somaclonal variation. Annual Review of Plant Physiology 39, 413–437. Lule, M., Dubois, T., Coyne, D., Kisitu, D., Kamusiime, H. and Bbemba, J. (2013) Trainer’s Manual. A Training Course for Banana Farmers Interested in Growing Tissue Culture Bananas. International Institute of Tropical Agriculture, Ibadan, Nigeria, 126 pp. Ma, S.S. and Shii, C.T. (1972) In vitro formation of adventitious buds in banana shoot apex following decapitation. Journal of the Chinese Society of Horticultural Science 18, 135–142. Matsumoto, K., Styer Caldes, L. and Yamamoto, Y. (2006) Response of banana dwarf somaclonal variants to benzylaminopurine. Infomusa 15 (1–2), 27–29. Moore, N.Y., Pegg, K.G., Langdon, P.W. and Whiley, A.W. (1993) Current research on Fusarium wilt of banana in Australia. In: Valmayor, R.V., Hwang, S.C., Ploetz, R., Lee, S.W. and Roa, N.V. (eds) Proceedings, International Symposium on Recent Developments in Banana Cultivation Technology, Chiuju, Pingtung, Taiwan, 14–18 December 1992. INIBAP/ASPNET, Los Banos, Laguna, Philippines, pp. 270– 284. MusaNet (2016) Global Strategy for the Conservation and Use of Musa (Banana) Genetic Resources (compiled by B. Laliberte). Bioversity International, Montpellier, France. Nayar, T.G., Seshadri, V.S. and Bakthavathasalu, C.M. (1958) Abnormalities in bananas – III. Indian Journal Agricultural Science 28, 401–402. Newbury, H.J., Howell, E.C., Crouch, J.H. and Ford-Lloyd, B.V. (2000) Natural and culture-induced genetic variation in plantains (Musa spp. AAB group). Australian Journal of Botany 48, 493–500.



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Njuguna, M.M., Wambugu, F.M., Acharya, S.S. and Mackey, M.A. (2010) Socio-economic impact of tissue culture banana (Musa spp.) in Kenya through the whole value chain approach. Acta Horticulturae 879, 77–86. Oh, T.J., Cullis, M.A., Kunert, K., Engelborghs, I., Swennen, R. and Cullis, C.A. (2007) Genomic changes associated with somaclonal variation in banana (Musa spp.). Physiologia Plantarum 129, 766–774. Ortiz, R. and Vuylsteke, D. (1995) Inheritance of dwarfism in plantain (Musa spp. AAB group). Plant Breeding 114, 466–468. Ortiz, R., Vuylsteke, D. and Ogburia, N.M. (1995) Inheritance of pseudostem waxiness in banana and plantain (Musa spp.). Journal of Heredity 86, 297–299. Perrier, X., De Langhe, E., Donohue, M., Lentfer, C., Vrydaghs, L. et al. (2011) Multidisciplinary perspectives on banana (Musa spp.) domestication. PNAS 108(28), 11311–11318. Pool, D.J. and Irizarry, H. (1987) Off-type banana plants observed in a commercial planting of ‘Grand Nain’ propagated using the in vitro culture technique. In: Galindo, J.J. and Jaramillo, R. (eds) Proceedings of the 7th ACORBAT Conference held at San Jose, Costa Rica. CATIE Technical Bull. 121, 99–102. Ray, T., Dutta, I., Saha, P., Das, S. and Roy, S.C. (2006) Genetic stability of three economically important micropropagated banana (Musa spp.) cultivars of lower Indo-Gangetic plains, as assessed by RAPD and ISSR markers. Plant Cell, Tissue and Organ Culture 85, 11–21. Reuveni, O. (1990) Methods for detecting somaclonal variants in ‘Williams’ bananas. In: Jarret, R.L. (ed.) Identification of Genetic Diversity in the Genus Musa. Proceedings of an International Workshop held at Los Banos, Philippines, 5–10 September 1988. INIBAP, Montpellier, France, pp. 108–113. Reuveni, O. and Israeli, Y. (1990) Measures to reduce somaclonal variation in in vitro propagated bananas. Acta Horticulturae 275, 307–313. Reuveni, O., Israeli, Y., Eshdat, Y. and Degani, H. (1986) Genetic Variability of Banana Plants Multiplied via in vitro Techniques. Final report submitted to IBPGR (No. PR 3/11), Agricultural Research Organization, The Volcani Center, Bet-Dagan, Israel. Reuveni, O., Golubowicz, S. and Israeli, Y. (1993) Factors influencing the occurrence of somaclonal variations in micropropagated bananas. Acta Horticulturae 336, 357–364. Reuveni, O., Israeli, Y. and Lahav, E. (1996) Somaclonal variation in banana and plantain (Musa species). In: Bajaj, Y.P.S. (ed.) Somaclonal Variation in Crop Improvement II. Springer, New York, pp. 174–196. Richardson, D.L. (1961) Note on the reversion of the dwarf banana ‘Cocos’ to ‘Gros Michel’. Tropical Agriculture (Trinidad) 38, 35–37. Robinson, J.C. and Nel, D.J. (1989) Mutations in tissue-culture field plantings of banana. Citrus and Subtropical Fruit Research Institute Information Bulletin 200, 7. Roux, N., Dolezel, J., Swennen, R. and Zapata-Arias, F.J. (2001) Effectiveness of three micropropagation techniques to dissociate cytochimeras in Musa spp. Plant Cell,Tissue and Organ Culture 66, 189–197. Roux, N.S., Strosse, H., Toloza, A., Panis, B. and Dolezel, J. (2004) Detecting ploidy level instability of banana embryogenic cell suspension cultures by flow cytometry. In: Jain, S.M. and Swennen, R. (eds) Banana Improvement: Cellular, Molecular Biology, and Induced Mutations. Science Publishers, Enfield, New Hampshire. Roy, O.S., Bantawa, P., Ghosh, S.K., Da Silva, J.A.T., DebGhosh, P. and Mondal, T.K. (2010) Micropropagation and field performance of ‘Malbhog’ (Musa paradisiaca, AAB group): a popular banana cultivar with high keeping quality of North East India. Tree and Forestry Science and Biotechnology, 4(1), 52–58. Sahijram, L., Soneji J.R. and Bollamma, K.T. (2003) Invited review: analyzing somaclonal variation in micropropagated bananas (Musa spp.). In Vitro Cellular and Developmental Biology – Plant 39, 551–556. Sahijram, L., Soneji, J.R. and Rao, M.N. (2010) Molecular and genetic characterization of somaclonal variation in micropropagated bananas (Musa spp.). Genes, Genomes and Genomics 4(1), 9–17. Sandoval, J.A., Tapia, F.A.C., Miller, L. and Villa Lobos, A.B. (1991) Observations about the variability encountered in micropropagated plants of Musa cv. ‘False Horn’ AAB. Fruits 46, 533–539. Sandoval, J., Kerbellec, F., Cote, F. and Doumas, P. (1995) Distribution of endogenous gibberellins in dwarf and giant off-types banana (Musa AAA, cv. ‘Grand Nain’) plants from in vitro propagation. Plant Growth Regulation 17, 219–224. Sandoval, J.A., Côte, F.X. and Escoute, J. (1996) Chromosome number variations in micropropagated trueto-type and off-type banana plants (Musa AAA ‘Grande Naine’ cv.). In Vitro Cellular and Development Biology–Plant 32, 14–17.

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Scowcroft, W.R. (1984) Genetic Variability in Tissue Culture: Impact on Germplasm Conservation and Utilization. Technical Report to the International Board for Plant Genetic Resources (IBPGR), Rome (84/152), 41 pp. Shirani, S., Mahdavi, F. and Maziah,M. (2009) Morphological abnormality among regenerated shoots of banana and plantain (Musa spp.) after in vitro multiplication with TDZ and BAP from excised shoot tips. African Journal of Biotechnology 8(21), 5755–5761. Simmonds, N.W. (1962) The Evolution of the Bananas. Longman, Green, London, 170 pp. Smith, M.K. (1988) A review of factors influencing the genetic stability of micropropagated bananas. Fruits 43, 219–223. Smith, M.K. and Drew, R.A. (1990) Growth and yield characteristics of dwarf off-types recovered from tissue-cultured bananas. Australian Journal of Experimental Agriculture 30, 575–578. Smith, M.K. and Hamill, S.D. (1993) Early detection of dwarf off-types from micropropagated Cavendish bananas. Australian Journal of Experimental Agriculture 33, 639–644. Stover, R.H. (1987) Somaclonal variation in ‘Grand Nain’ and ‘Saba’ bananas in the nursery and field. In: Persley, G.J. and De Langhe, E.A. (eds) Banana and Plantain Breeding Strategies. Proceedings of an International Workshop, Cairns, Australia. ACIAR Proceedings No. 21, Australian Centre for International Agricultural Research, Canberra, Australia, pp. 136–139. Stover, R.H. (1988) Variation and cultivar nomenclature in Musa. AAA group, Cavendish subgroup. Fruits, 43(6), 353–356. Stover, R.H. and Simmonds, N.W. (1987) Bananas. Longman Scientific and Technical, Harlow, UK, 468 pp. Strosse, H., Van den houwe, I. and Panis, B. (2004) Banana cell and tissue culture – review. In: Jain, S.M. and Swennen, R. (eds) Banana Improvement: Cellular, Molecular Biology and Induced Mutations. Science Publishers, Inc., Enfield, New Hampshire. Swartz, H.J. (1991) Post culture behaviour: genetic and epigenetic effects and related problems. In: Debergh, G.J. and Zimmerman, R.H. (eds) Micropropagation: Technology and Application. Kluwer Academic Publishers, Dordrecht, The Netherlands, pp. 95–112. Tang, C.Y. (1995) Variation of bunch types in mericlones of a multi-bunching banana. Infomusa 4(2), 17–18. Tenkouano, A. (2000) Persistence and horticultural value of inflorescence dichotomy in plantain. Hortscience 35, 933–936. Tilney-Basset, R.A.E. (1986) Plant Chimeras. Edward Arnold, London, 200 pp. Ventura, J.de-la-C., Rojas, M.E., Yera, E.C., Lopez, J. and Rodriguez-Nodals, A.A. (1988) Somaclonal variation in micropropagated bananas (Musa spp.). Ciencia y Tecnica en la Agricultura, Viandas Tropicales 11(1), 7–16. Vuylsteke, D. (1998) Field performance of banana micropropagules and somaclones. In: Jain, S.M., Bra, D.S. and Ahloowalia, B.S. (eds) Somaclonal Variation and Induced Mutation in Crop Improvement. Kluwer Academic Publishers, Dordrecht, The Netherlands, 219–231. Vuylsteke, D., Swennen, R.L., Wilson, G.F. and De Langhe, E.A. (1988) Phenotypic variation among in vitro propagated plantain (Musa sp. cultivar ‘AAB’). Scientia Horticulturae 36, 79–88. Vuylsteke, D.R., Swennen, R.L. and De Langhe, E.A. (1991) Somaclonal variation in plantains (Musa spp., AAB group) derived from shoot-tip culture. Fruits 46, 429–439. Vuylsteke, D.R., Swennen, R.L. and De Langhe, E.A. (1996) Field performance of somaclonal variants of plantain (Musa spp., AAB group). Journal, American Society Horticultural Science 121, 42–46.

13 

Quarantine and the Safe Movement of Musa Germplasm D.R. Jones

A trained plant pathologist best serves his country and humanity when he detects and stops a dangerous pest before it can become established. (Carefoot and Sprott, 1967)

Introduction The present distribution of serious diseases of banana is outlined in Table 13.1. Some have a relatively limited distribution while others have a continental or near-global distribution. Factors controlling the rate of spread are numerous. They include the life cycle of the pathogen, the effectiveness of quarantine measures, the presence or absence of insect vectors, climate and geographical features, such as oceans, deserts, forests and high mountain ranges, which may act as natural barriers. The sheaths of wild banana must have been useful to early humans as a source of fibre and the leaves as wrapping materials and temporary thatch. The male buds, male flowers and immature fruit may also have been utilized as vegetables, as they still are in some areas of Southeast Asia today. Early cultivators would have eagerly transplanted suckers of banana with fruit showing the first signs of parthenocarpy and sterility, as this feature would have added to the attractiveness of an already versatile plant. Superior edible types would have been propagated and distributed widely among and between ethnic

groups. In this way, banana cultivars would have spread throughout tropical Asia. Early migrants and traders are thought to be responsible for the later transfer of planting material from Asia to Africa. Relatively recently in historic terms, Polynesians distributed banana throughout the Pacific and Europeans took the plant across the Atlantic to the New World (Simmonds, 1962). Musa is affected by a number of viruses, bacteria, phytoplasmas, fungi and nematodes. Some minor pathogens of banana and abacá have a worldwide distribution, because they have common alternative hosts in most environments suitable for Musa and are not reliant on specific vectors. This allowed for an early and possibly natural dissemination globally. Others originated in or near the centre of origin of banana and abacá or had a limited distribution in an area where they evolved to attack Musa. The mode of dissemination of many of these pathogens and the restrictions on human movement in historical times meant that their initial spread with germplasm exchanged between tribes would have been slow. These pathogens had to wait for the assistance of modern humans with efficient and relatively cheap modes of long-distance transport to enable them to spread more rapidly and for the large-scale cultivation of Musa for their effects to become noticeable. When the detrimental effects of these diseases on yield threatened the economics of commercial banana industries, their causes were investigated seriously in

© D.R. Jones 2019. Handbook of Diseases of Banana, Abacá and Enset (D.R. Jones)

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Table 13.1.  Known distribution of some important or potentially important pathogens of Musa. Pathogen Fungi Pseudocercospora fijiensis Pseudocercospora musae Pseudocercospora eumusae Phyllosticta cavendishii Fusarium oxysporum f. sp. cubense (race 1) Fusarium oxysporum f. sp. cubense (tropical race 4) Bacteria X. campestris p.v. musacearum (X. vasicola p.v.) Ralstonia solanacearum Ralstonia syzgii spp. celebesensis ‘Banana wilt associated phytoplasma’ Viruses Banana bunchy top virus Banana bract mosaic virus Banana streak virus Cucumber mosaic virus Abacá mosaic virus Nematodes Rhadopholus similis Pratylenchus coffeae Pratylenchus goodeyi

Mainland Asia

Pacifica

Latin America

Caribbean

Africa

Black leaf streak Sigatoka leaf spot Eumusae leaf spot

L W W

W W –

W W –

W W –

W W L

Freckle Fusarium wilt

W W

L L

– W

? W

? W

Fusarium wilt

L

L

–

–

L

Xanthomas bacterial wilt

–

–

–

–

L

Moko bacterial wilt Blood disease

? ?

K L

W –

L –

– –

Phytoplasma wilt

–

–

–

–

Bunchy top Bract mosaic Banana streak Banana mosaic Abacá mosaic

W L W W –

W L W W L

– – W W –

– – W W –

L – W W –

Burrowing nematode Root lesion nematode Root lesion nematode

W

W

W

W

W

W

W

W

W

W

–

Lb

–

–

L

Disease

Includes Australia, East Malaysia, Indonesia, Papua New Guinea, the Philippines and Taiwan lying off the Asian mainland. Subtropical Australia. Distribution: L, distribution limited; W, distribution widespread; –, absent; ?, records need confirmation or are questionable.

a b

order to find solutions and return the industries to profitability. For these reasons, the first banana pathologists were employed by either export enterprises or colonial governments. It is important that care is taken to slow or prevent the further dissemination of pathogens of Musa. The greatest risk to commercial production in the Latin American–Caribbean region would be the introduction of populations of Fusarium oxysporum f. sp. cubense that are capable of attacking Cavendish clones in the tropics. If this occurred and a wilt-resistant replacement for Cavendish cultivars were not found, then the

damage already caused in plantations in West Malaysia (Ong, 1996) and elsewhere would be repeated on a much larger scale. Major threats to smallholders and subsistence farmers come from the continued spread of bunchy top, bract mosaic and bacterial wilts. However, it is not only the most serious diseases that may pose problems. A disease that may cause minor damage in one environment may cause significant damage in another. It is prudent, therefore, to try to exclude all exotic pathogens from gaining entry to countries where they do not occur. Even if a disease is present in a



Quarantine and the Safe Movement of Musa Germplasm

country, it is good policy not to be indifferent to its reintroduction, because of possible strain or race differences that could increase problems. The basis of good quarantine policy relies on a knowledge of the diseases present in a country and of exotic diseases that threaten crops in that country. An understanding of the epidemiology of exotic diseases and how they may gain entry is also important. Without this information, it is very difficult to develop regulations and guidelines for plant importations. For this reason, it is crucial that countries have the capability of diagnosing disease problems. Basic taxonomic training of plant pathologists should be encouraged and supported, especially in developing countries. It is hoped that this book will provide sound assistance to those who identify problems of Musa and Ensete. However, diagnosis should not rely on matching the illustrations with symptoms seen in the field. Only a few diseases and disorders are obvious from their effects on the plant. Others may have symptoms that closely

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resemble different problems. The plates and figures in the book, therefore, serve only as a guide. Further diagnostic work is necessary to confirm suspicions about a possible causal agent. In this respect, the numerous references cited in this book are meant to serve as a source of information that may allow a diagnostician to move forward to a positive end result.

Movement of Pathogens Most banana diseases can be spread from one location to another with planting material. Until fairly recently, banana planting material consisted of rhizomes, with or without attached leaf and pseudostem tissue (Plate 13.1). The potential for moving pathogens with rhizomes is high, as adhering soil can harbour root-attacking organisms, such as nematodes, fungi and bacteria. In addition, the rhizome tissue itself could be infected with all three and also viruses. Any

Plate 13.1.  Suckers of ‘Klaui Namwa’ (ABB, Pisang Awak subgroup) on sale in northern Thailand. This planting material comes from an area near the Mekong River, where race 1 of Fusarium oxysporum f. sp. cubense is commonly found on this cultivar. Some suckers would undoubtedly carry the soilborne and rhizome-borne pathogen (photo: D.R. Jones, INIBAP).

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attached leaf tissue and adhering leaf trash could also carry foliar pathogens. Risks can be significantly reduced if the rhizome is washed to remove soil and pared to trim off roots, the outermost tissue layers, discoloured areas and any leaf material. However, risks are not eliminated and there is still the possibility that some pathogens, which may not show obvious symptoms, could be present. It is known that F. oxysporum f. sp. cubense is disseminated with rhizomes and infested soil. Bancroft (1876) first recognized the importance of planting uninfected rhizomes to control the disease, but, as corms can carry the disease without showing obvious symptoms, his recommendations were easier said than done. Nematodes have also been introduced to uninfested land with planting material and accompanying soil. The appearance of the burrowing nematode in some new areas has been associated with shipments of rhizomes of highly susceptible cultivars in the Cavendish subgroup. It has been suggested that the bacterium that causes Moko bacterial wilt may have been introduced to Mindanao in the Philippines on Cavendish rhizomes imported from Central America (Buddenhagen, 1986). Banana bunchy top virus (BBTV) was most likely introduced into Australia with planting material from Fiji (Dale, 1987). In many parts of the banana-growing world, banana leaves are used to wrap food and pack produce, which may be carried or taken long distances by travellers and traders. Although it is unlikely that heavily diseased leaves would be deliberately utilized for this purpose, it is possible that an occasional leaf with an odd lesion may be selected. The spread of black leaf streak in the Americas may have been accelerated by the use of infected banana leaves to cushion transported fruit. Banana leaves are not allowed entry into Australia, because of the risk of introducing foliar pathogens (Jones, 1991). Dried leaf tissue could also harbour foliar and fruit disease pathogens (Stover, 1977). Banana fruit can carry crown rot fungi and other pathogens. Fungi causing crown rot are found everywhere, and quarantine measures to exclude them are not warranted. Bananas can also be affected by serious bacterial diseases and not show obvious external symptoms. Often, the presence of these diseases is only discovered when fingers are peeled or cut to expose the pulp.

Diseased fruit could be moved from one location to another and then discarded in the garden when found to be rotten. This may allow the bacterial pathogen to spread to nearby banana plants. A quarantine regulation restricting the movement of bananas in Indonesia, as well as vegetative parts of banana, may well have delayed the spread of blood bacterial wilt from Sulawesi to West Java until 1987 (Eden-Green, 1994). There is also a risk that leaf trash with viable fruiting bodies of fungal pathogens may accompany bananas packed in cartons. For these reasons, the risks associated with the movement of banana fruit from one banana-growing country to another need to be carefully evaluated. Some pathogens move with seed of crop plants. Seed, of course, is produced in fruit of wild banana species (Plate 13.2). Seed is also found in the fruit of plants following artificial pollination in breeding programmes. Although few, if any, seeds are usually found in fruit of cultivated banana, sometimes edible cultivars, such as ‘Pisang Awak’ (ABB), which retains a high degree of fertility, produce them. There is strong evidence that banana streak viruses are seedborne (Daniells et  al., 1995) and it is possible that cucumber mosaic virus (CMV) may also be seed-borne (Gold, 1972; Stover, 1972). Although not absolutely proven, bacterial agents of Moko, bugtok and blood bacterial wilts, which attack banana fruit, may also move with seed as surface contaminants. Some banana pathogens can affect alternative hosts, which then become sources of infection. Not much is known about the host range of many banana pathogens and so risks have not

Plate 13.2.  Rows of seed inside a finger of Musa acuminata ssp. banksii (AAw) in Papua New Guinea (photo: D.R. Jones, QDPI).



Quarantine and the Safe Movement of Musa Germplasm

been totally defined, but some threats have been identified. Heliconia is known to pose a significant risk, as it has long been recognized as a host of the bacterium that causes Moko bacterial wilt. In 1989–1990, a survey of 20 commercial heliconia farms in Hawaii found five with plants affected by heliconia wilt. Of 17 isolates of Ralsto­ nia solanacearum obtained from diseased heliconia, five were pathogenic on banana. Fortunately for the banana industry in Hawaii, these banana-­ attacking isolates had not spread from Heliconia to banana. The results of the survey suggested that quarantine regulations governing the import of Heliconia into Hawaii and intrastate movement of vegetative cuttings or rhizomes were needed (Ferreira et al., 1991). At the time of the Hawaiian survey, the threat that ornamental heliconia posed to the banana industry in Australia had long been recognized and quarantine regulations were in place that required imported Heliconia to be held in post-entry quarantine before release. The quarantine period was 3 months in a private post-entry quarantine house for plants from areas free of Moko bacterial wilt and 9 months in a government post-entry quarantine facility for those from an area where the disease was known to occur. All needed to be inspected regularly by a government plant pathologist before release. After learning of the detection of heliconia wilt in Hawaii, the Australian quarantine authorities immediately organized a survey of heliconia plants recently imported from Hawaii into Australia. A number of plants were found to be wilting in a private post-entry quarantine house in a banana-growing area. Some were affected with strains of R. solanacearum that were not pathogenic on banana, but two plants were shown to carry R. solanacearum race 2, strain SFR. Fifty-five plants already released from quarantine because of an absence of obvious wilt symptoms during the 3-month quarantine period had been planted outside in the importer’s nursery. Two were found to be diseased and were removed. The nursery was quarantined, unaffected heliconia plants were returned to quarantine, banana plants on the property were destroyed and soil that might have been contaminated was removed and treated with formalin (Jones, 1991; Hyde et al., 1992; Akiew and Hyde, 1993). Luckily, the pathogen was contained and eradicated, due to prompt quarantine action. This incident

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highlights the need for extreme caution if Heliconia is allowed entry to banana-growing countries.

Aspects of Spread of Some Major Banana Health Problems Fusarium wilt Only a few of the major disease problems known today seem to have been disseminated during the first movements of Musa planting material out of Southeast Asia to Africa and elsewhere. The distribution of Musa germplasm during the 20th century seems to be responsible for the long-distance spread of most important banana pathogens from their centres of origin. Fusarium oxysporum f. sp. cubense, the cause of Fusarium wilt, is a notable exception to this rule. Intercontinental spread of the pathogen is considered to have occurred long before the disease was described for the first time in 1876 (Stover, 1962). It is fortuitous that populations of the Fusarium wilt fungus capable of attacking Cavendish clones in the tropics did not disseminate outside Asia at this time. However, the most important Cavendish-attacking strain of F.  oxysporum f. sp. cubense in the tropics (VCG 01213) is now expanding its area of distribution having reached China, Australia, the Middle East and ­Africa in recent years. Its introduction to Africa at the Matanuska commercial export plantation in Monapo in the Nampula province of Mozambique (IPPC, 2013) has been thoroughly investigated, but still no definite conclusions have been reached on how the pathogen gained entry to the continent. Some think propagules may have arrived in soil attached to the boots of plantation workers from Southeast Asia. It is ironic that this first outbreak on the African continent was associated with an aid project that was hoped to provide employment and incomes for local people. A second export banana plantation in Mozambique in the Chiúre district in Cabo Delgado province has now also been found to be affected by the disease. There are grave concerns that the problem will spread to other areas in Mozambique and into neighbouring countries. The great fear is that the pathogen may also spread to the main banana-­ growing regions in the Americas and become established and perhaps disseminated widely before any containment actions can be taken.

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Recently, there have been reports of F. oxy­ sporum f. sp. cubense VCG 01213/16 in Cavendish plantations and plots in Vietnam (Hung et al., 2017) and Laos (Chittarath et al., 2017). These new areas of Cavendish production were established because yields of Cavendish fruit in China have been falling as a result of inroads made by the TR4 strain in that country. Although small plots of Cavendish along the Red River in Vietnam may have succumbed to the disease because of pathogen propagules washed downstream from Yunnan province in China, it seems likely that new commercial plantations became affected because of the movement of contaminated soil with trucks and other farm equipment, contaminated soil on the shoes of workers and also with hardened tissue-cultured planting material from nurseries in China, notably Guangxi province (A. Molina, Philippines, 2017, personal communication). TR4 has now also been reported in the Greater Mekong subregion of Vietnam and in Myanmar (Zheng et al., 2018). Plants in a 2ha Cavendish plantation at Baoo-Pooran in the south of Sindh province of Pakistan were first seen with Fusarium wilt symptoms in 2012. This outbreak was later shown to be caused by TR4 (Ordóñez et al., 2016). The problem has since spread further north to the centre of the Sindh province (H. Laghari, Pakistan, 2017, personal communication). Outbreaks of Fusarium wilt caused by TR4 have also been detected in the Katihar and Purnea districts of Bihar state in India (Thangavelu, 2018). At the time of going to press, there were unconfirmed reports that the pathogen had also been found in the Faizabad district of ­Uttar Pradesh, the Burhanpur district in Madhya Pradesh and at Surat in Gujarat (Kulkarni, 2018). It is likely that TR4 will spread to all banana growing areas in South and Southeast Asia in the coming years. In Australia, VCG 01213 devastated banana production in the Northern Territory following its discovery in 1997. Its spread to new farms was in spite of coordinated attempts to contain the problem. On 3 March 2015, the pathogen was detected on a Cavendish banana farm in the Tully Valley, which is about 140 km south of Cairns in North Queensland. Other outbreaks were detected on the same farm. The Tully Valley is a major banana-growing location in the largest banana-growing area in Australia and the pathogen’s arrival was taken very seriously by the biosecurity authorities, the government

and the growers. In an attempt to control the disease, all plants on the property were subsequently destroyed. The land was quarantined and surrounded by a security fence. Because it was estimated that the pathogen could survive in the soil for at least 40 years, further banana cultivation was stopped and possible dissemination in watercourses was studied. A ground cover was established to reduce the run-off of rainwater. The contaminated land was subsequently purchased by the national association of banana growers with help from the government to compensate the farmer for loss of production. As the disease is spread through soil, mud and infected plant material, it was essential that people, vehicles, machinery and equipment be appropriately decontaminated on entering and exiting the property. A ‘grower’s kit’ was issued by Biosecurity Australia which provided growers with important information to help protect their properties. All growers in the region were encouraged also to take precautions by installing their own disinfectant dips for vehicles and the shoes of people entering their property from elsewhere. A new outbreak on another farm near Tully was detected on 12 July 2017 showing how difficult it is to contain the problem. Again, time will tell if the pathogen can be contained or whether further spread is inevitable. Surveys have indicated that F. oxysporum f. sp. cubense most likely evolved in Indonesia and Malaysia, where diversity is highest. It is in these countries that isolates attacking Cavendish are also endemic (Buddenhagen, 2009). The increasing trend of the monoculture of susceptible cultivars in large plantations has provided ideal conditions for epidemics in this region. VCG 01213 has also been isolated from banana plants in villages in the region where banana is still grown. It is likely that a combination of different strains of the F. oxysporum f. sp. cubense in the local soil plus the practice of growing many different cultivars of varying genotypes together has reduced the impact of VCG 01213. Buddenhagen (2009) reported that wilt was seldom seem in village plots in southern Sumatra and believed it could be explained by villagers abandoning their favourite, highly susceptible cultivars, such as ‘Pisang Berangan’ (AAA), but continuing to grow less susceptible clones. This form of management allows the banana to remain an important smallholder crop in Southeast Asia.



Quarantine and the Safe Movement of Musa Germplasm

Black leaf streak Pseudocercospora fijiensis is an example of a pathogen with a high capability for spread. Initially, the disease caused by P. fijiensis was most likely of minor concern in village plots in the Southeast Asian–Pacific region because of the diversity of Musa genotypes grown, some being susceptible, others having resistance. The first recognition of black leaf streak in Fiji in 1963 was probably because of a combination of a susceptible cultivar being grown in large contiguous plots, thus encouraging an epiphytotic, and the presence of colonial plant pathologists, who realized the symptoms were of something new. The pathogen’s subsequent recognition elsewhere and spread to new areas was dramatic. As a fungal disease of banana foliage, black leaf streak can be carried long distances with leaf trash, fresh leaves used to wrap produce and leaves attached to suckers. Airborne spores are also thought to carry the pathogen considerable distances and to ensure that, once established in a new region, a steady and inexorable spread to surrounding banana-growing districts and countries is inevitable. When black leaf streak was first found on islands in the Torres Strait and on the tip of Cape

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York Peninsula in Australia (Jones and Alcorn, 1982), attempts were made to eradicate the disease and prevent the spread of the fungal pathogen to other parts of the country (Plate 13.3). All plants were destroyed by burying, ploughing or burning. When eradication failed (Jones, 1984), most probably as a result of inoculum blown in the wind from Papua New Guinea during cloudy weather, susceptible cultivars growing in the area where the disease was found were replaced with resistant ones to reduce inoculum levels and the chance of spread south (Jones, 1990). Since then, black leaf streak has been discovered occasionally on susceptible banana plants growing in isolated communities on Cape York Peninsula to the south of the affected zone. These outbreaks have been eradicated as they occur. Because of the vigilance of the quarantine authorities and the natural barrier to spread afforded by the sparsely populated Cape York Peninsula, the commercial banana-growing areas of Australia were spared until the pathogen was found in a plantation in the main growing area near the town of Tully in North Queensland in April 2001 (Jones, 2003: Petersen et al., 2003). The disease was detected on 14 commercial properties and 11 clumps of unmanaged or ‘feral’ plants. Its distribution and intensity of

Plate 13.3.  Quarantine sign at airport on Horn Island in the Torres Strait area of Australia in the late 1980s. In order to prevent the spread of black leaf streak disease, air passengers are warned that it is prohibited under quarantine legislation to carry banana material (photo: D.R. Jones, QDPI).

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symptoms indicated that it had probably arrived about 6–12 months before its discovery. Government funds, prompt action by the local horticultural authorities, the cooperation of banana growers plus the onset of the dry season were all factors that led to its eradication in the region. The success of the campaign was brought about by the de-leafing of 4500 ha of commercial plantations, weekly fungicide sprays for 6 months and the destruction of all non-­ managed plants (Petersen et  al., 2003). This is the one success story in the otherwise relentless spread of the disease to all areas where banana is cultivated. Black leaf streak still remains a major threat to banana cultivation in tropical Queensland. The disease can still be seen on susceptible banana cultivars growing on those Torres Strait islands closest to Papua New Guinea. The disease has not been totally eradicated from Australia and is waiting for an opportunity to stage a future incursion into commercial growing areas to the south. This incursion may be through the activities of humans moving planting material of leaves or the long-distance dissemination of spores during cyclonic disturbances to ‘stepping-­ stone’ clumps of banana on Cape York Peninsula. Continued vigilance is important. Regular disease surveys are the only way to prevent a reoccurrence of black leaf streak in the growing areas of North Queensland or to spot an incursion before it becomes established. The relentless spread of black leaf streak from its centre of origin in the Southeast Asian– New Guinea region to the Americas and Africa has been documented in Chapter 2.

Freckle Freckle disease of banana is now known to be caused by at least three species of Phyllosticta (Wong et  al., 2012). However, it was originally thought that there was only one pathogen. When freckle was discovered in the Torres Strait and Cape York regions in far north Queensland (Jones and Alcorn, 1982), the available information seemed to suggest that it was a major disease of Cavendish cultivars, which are grown almost exclusively in Australia’s commercial growing areas. It was for this reason that eradi-

cation was attempted at the same time that the eradication of black leaf streak was attempted in the same regions. These attempts failed (Jones, 1984). Later visits by the author to Papua New Guinea and the South Pacific showed freckle to be widespread and prevalent on ABB cultivars. No symptoms were seen on Cavendish clones. This had also been the case in the Torres Strait and Cape York regions of Australia. How could this be when Cavendish, but not ABB cultivars, were attacked in Taiwan (Hwang et  al., 1984) and Hawaii (Meredith, 1968)? It was then thought that it was possible that two or more races or species of fungus were causing freckle (Jones, 1999). The species attacking banana in Queensland was later discovered to be P. maculata, which has been found on ABB cultivars, but not Cavendish clones (Wong et al., 2012). Freckle was also found in Australia at Kununurra in northern Western Australia in 1979 seriously affecting the first banana plants to be grown commercially in the area. The planting was of a Cavendish cultivar and, though not known at the time, the pathogen was most likely P. cavendishii (Wong et  al., 2012). This planting was destroyed before any other commercial banana plants were grown. In 2001, the Northern Australia Quarantine Service (NAQS) undertook a banana disease survey in Broome, Kalumburu and Kununurra in northern Western Australia and found a Cavendish cultivar with freckle at Kalumburu. An investigation found that the plants had most likely come from Kununurra in the early 1980s before the destruction of banana plants at the outbreak site of 1979 (McCurdy, 2001). In August 2013, banana freckle was discovered on Cavendish cultivars at a number of properties in the Howard Springs area near Darwin in the Northern Territory of Australia. This was found to be P. cavendishii. In October 2013, a National Management Group with representatives from government and the banana industry agreed to a nationally cost-shared response programme to eradicate P. cavendishii. The eradication response, which was carried out by the Northern Territory government, involved search and surveillance of thousands of properties for the presence of freckle disease. All banana plants within a 1 km radius of affected properties were destroyed and ongoing monitoring continued for 12 months. The Plant Health Act (NT) gave inspectors broad powers to enter, inspect and



Quarantine and the Safe Movement of Musa Germplasm

search properties, require information and seize and treat plants. Section 18 of the Act also allowed the Chief Inspector of Plant Health to declare quarantine areas established to prevent further movement of the disease and cultivation of banana plants. May 2015 to April 2016 was a host-free period for the areas where eradication was being attempted. May 2016 to April 2017 was when sentinel host banana plants were grown in the area of eradication and others allowed to plant only freckle-free plants derived from tissue culture. Further observations from May 2017 onwards will determine whether the eradication has been successful. The above gives the reader some idea as to the serious nature with which P. cavendishii is considered by the Australian government and the local banana industry. Close cooperation between government (with a professional plant health service) and growers has been the key to success in keeping important exotic banana pathogens in check in Australia.

Xanthomonas bacterial wilt Symptoms of Xanthomonas bacterial wilt were first seen on enset in Ethiopia in the 1930s (Castellani, 1939). The causal agent was named in 1968 (Yirgou and Bradbury, 1968) and was later found to affect some of the few mats of banana that grew in the enset-growing areas (Yirgou and Bradbury, 1974). The potential for spread and severe damage to banana crops worldwide was recognized at the time by Yirgou and Bradbury (1974), who warned that great care needed to be taken to see that the bacterial wilt of enset did not ‘escape’ from Ethiopia. However, the long period of time after its discovery on banana and its non-appearance on banana in any another country, plus the belief that it was primarily a major disease of enset, may have made plant pathologists complacent about its potential to cause damage elsewhere. Thus, its appearance in 2001 on banana in the Mukono district in Central Uganda (Tushemereirwe et al., 2004) and at Masisi in the North Kivu province in the eastern Democratic Republic of the Congo (Ndungo et  al., 2004) came as a complete surprise to most. What is more surprising is that the disease has still not established in the main ba-

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nana-growing area of Ethiopia in the Rift Valley near Arba Minch. The Rift Valley here is surrounded by and is quite close to enset-growing areas, which are at a higher elevation (G. Blomme, Ethiopia, 2017, personal communication). The high degree of genetic sequence similarity among a number of isolates of the pathogen collected in different countries on banana and enset indicates a very recent origin of the pathogen in Ethiopia (Wasukira et  al., 2012). Genetic analysis also suggests that there are at least two major sub-lineages of the pathogen that vary slightly from one another.  Isolates from Uganda, Kenya, Tanzania and Burundi are genetically different from isolates collected in Ethiopia, Democratic Republic of the Congo and Rwanda. This indicates that there may have been more than one introduction from Ethiopia to countries in the Great Lakes region. One pathway may have been to Uganda and another to the Democratic Republic of the Congo (Wasukira et al., 2012). The area where the disease first appeared on banana in Uganda is about 570 km from the Ethiopian border and Masisi in the Democratic Republic of the Congo is about 1000 km away. The mode of this long-distance spread is unknown. The most obvious means would have been by the movement of infected enset/banana planting material over many kilometres from the highlands of Ethiopia. The pathogen causing Xanthomonas wilt of enset and banana has been found to be a pathovar of Xanthomonas vasicola and the name X. vasicola pv. musacearum has been suggested (Aritua et al., 2007). Other pathovars of X. vasi­ cola attack sugarcane, maize and sorghum in Africa. Sorghum, which originated in the semi-­ arid regions of sub-Saharan Africa, is a host of X. vasicola pv. holcicola. It has been suggested that this pathogen infecting sorghum in close proximity to enset may have adapted to attack enset (Aritua et al., 2009). In retrospect, it is a great pity that the pathogen could not have been eradicated shortly after its discovery in Uganda and the Democratic Republic of the Congo. However, containment would have been difficult given its dissemination by flying vectors (insects, birds and bats). Spread westward throughout the plantain-growing areas of the Democratic Republic of the Congo and to other surrounding countries seems now to be almost inevitable.

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Bunchy top The spread of some other significant diseases has been less rapid. Bunchy top was recorded in Fiji in the late 19th century, but evidence indicated that it most likely originated elsewhere. It was next seen in Tuvalu (Ellice Islands) in 1926 and then Egypt and Australia in 1927. Early spread was almost certainly because of the movement of infected planting material along colonial trade routes. In subsequent years, it was recognized in other countries in Asia, the Pacific and Africa (see Chapter 6). Its identification at new locations over time has been steady, but generally slow, though in recent years the rate of new country records seems to have accelerated, with banana found infected in an additional 16 countries since 2000. The long-distance dissemination of banana bunchy top virus (BBTV) relies primarily on the movement of infected planting material from one country to another. The unrestricted movement of banana suckers, corms and tissue cultures across international borders is against the quarantine regulations of many banana-growing countries, but undoubtedly occurs from time to time as legitimate channels of entry are circumvented. Propagating material, including meristems used to initiate in vitro cultures, would not normally be taken from plants with obvious bunchy top symptoms, but there is a possibility that germplasm could be selected for propagation after infection, but before symptom expression. Stainton et al. (2015) analysed 171 banana leaf samples from 14 countries and recovered, cloned and sequenced 855 complete BBTV components, including 94 full genomes. Full genomes were determined from eight countries where previously no full genomes had been available (Samoa, Burundi, Republic of the Congo, Democratic Republic of the Congo, Egypt, Indonesia, the Philippines and the USA-Hawaii). Accounting for recombination and genome component reassortment, the authors examined the geographical structuring of global BBTV populations, which revealed that BBTV likely originated in Southeast Asia and that the current global hotspots of BBTV Diversity were Southeast Asia/Far East and India. The research also indicated that the global patterns of geographical structure that are evident within the BBTV phylogeny are not consistent with the virus having been spread

during the prehistoric dissemination of banana across the globe, nor are they consistent with frequent human-mediated transcontinental BBTV movements during the past few decades. They are, however, entirely consistent with more gradual, natural, or human-facilitated movements of the virus (via infected propagules and/or aphids) over the past 300 years from its centres of diversity. The results also suggested frequent shorter-distance BBTV movements. For example, there is evidence indicating that during the past 100 years there must have been multiple BBTV movements between Taiwan, China, the Indian subcontinent and Indonesia. In countries where bunchy top has become established, total eradication has proved elusive and so it is important to prevent introduction by enforcing strict quarantine measures. This is not easy in developing countries with porous borders. Streak Banana streak viruses have been reported as being found in over 50% of Musa germplasm accessions containing the B genome indexed by Biosecurity International’s banana indexing centres (De Clerck et al., 2017). Almost all selections of ‘Mysore’ (AAB), a cultivar believed to have originated in India, where it is known as ‘Poovan’, are infected with banana streak virus (BSV). The distinctive periodic symptoms of BSV were once thought to be a physiological problem and were used in part by the author to identify the clone on germplasm collecting missions. The ‘peculiarities’ of ‘Mysore’ were only relatively recently suspected of being disease symptoms and were not associated with BSV infection until 1993 (Infomusa, 1993). The virus most probably spread with this clone when planting material of the clone was introduced into other countries (Plate 13.4). Symptoms of BSV in ‘Mysore’ are not always visible and effects can be much more subtle than those of BBTV. The virus thus became widely disseminated. Further spread of BSV from ‘Mysore’ to other clones via mealybug vectors may be an uncommon event in many countries, as ‘Mysore’ is often the one of a few clones with BSV symptoms in long-established field germplasm collections. Other germplasm accessions



Quarantine and the Safe Movement of Musa Germplasm

Plate 13.4.  Symptoms of what is likely to have been banana streak Mysore virus in ‘Mysore’ (AAB) in FHIA’s germplasm collection in Honduras in 1994 (photo: D.R. Jones, INIBAP).

with BSV may be those plantain types where the male bud degenerates on maturity. These are commonly seen with BSV symptoms in the field in Central and South America. The great variability of BSV in Uganda suggested that it arose in part because of a series of introductions of infected banana cultivars into Uganda, each with a different complement of infecting viruses (Harper et al., 2004). Five new BSV species have been recognized from a number of variants collected in Uganda and Kenya by James et al. (2011b). The creation of episomal BSV infections from integrated sequences in the banana genome of clones containing the B genome during embryo rescue and tissue culture procedures has led to complications in conventional banana breeding programmes and the safe movement of improved Musa germplasm (see Chapter 6). The bred hybrid ‘FHIA-21’ (AAAB) was seen by the author with symptoms of BSV when on ­trial in Honduras in 1995 (Plate 13.5). During a survey for BSV conducted in Cuba, banana streak GF virus (BSGFV) and banana streak IM ­virus (BSIMV) were detected separately in plants of the introduced hybrid ‘FHIA-25’ (AAAB) and both together in other plants of the same hybrid. Banana streak OL virus (BSOLV) was found in ‘FHIA-18’ (AAAB) and ‘FHIA-21’ (AAAB), and both BSGFV and BSOLV in other ‘FHIA-21’ plants (Javer et  al., 2009). CIRAD-bred AAB hybrids ‘IRFA 909’, ‘IRFA 910’ and ‘IRFA 914’ were also found with BSGFV when undergoing trials in ­Australia (Geering and Thomas, 2002). This problem is still of major concern to banana breeders.

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Plate 13.5.  Symptoms of a banana streak virus in ‘FHIA 21’ (AAAB – bred hybrid) in a trial the Baracoa area of Honduras in 1995 (photo: D.R. Jones, INIBAP).

Principles of the Safe Movement of Musa Germplasm The safe movement of Musa germplasm was ­reviewed by Stover (1977), who outlined the danger of disease and pest transfer on traditional planting material and emphasized the advantages of using tissue-cultured plantlets for the first time. The movement of conventional propagating material of Musa, such as rhizomes and suckers, from one banana-growing country to another should no longer take place. Even when accompanied by certification that the material has been inspected and found free of disease symptoms, there is still a risk, because certain pathogens may be carried without exhibiting symptoms. Only if the importing country has the capability of immediately initiating shoot-tip cultures from the imported material and testing plants derived from these cultures for virus pathogens, which may be carried as latent infections, should this be contemplated. Seed of Musa and Heliconia imported into Australia is submerged in a solution of sodium hypochlorite (1% available chlorine) for 10 min as a precaution against the introduction of the Moko disease pathogen (Jones, 1991). If seeds of species in these genera need to be imported, they should also be sown in post-entry quarantine and plants observed regularly for disease symptoms. The safest way of introducing new germplasm is as tissue culture (Plate 13.6). This eliminates

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Development of a System for the Safe Movement of Musa Germplasm History

Plate 13.6.  Banana plantlets growing in tissue culture (photo: QDPI).

the risks posed by fungi, bacteria and nematode pathogens, as these would ­almost certainly contaminate the culture medium if present. Artificial media used to culture germplasm introductions should not contain antibiotics or colouring, which might inhibit or mask contaminants. The germplasm exchange system currently provided and administered by the International Network for the Improvement of Banana and Plantain (INIBAP) utilizes in vitro cultures (Van den houwe and Jones, 1994). Although the risks are significantly reduced using tissue cultures, they are not eliminated. The possibility of moving virus and phytoplasma pathogens with tissue cultures of Musa still remains. In recent years, efforts have been made to develop and improve tests to detect Musa viruses in germplasm held in collections and in quarantine. In the future, the risks posed by phytoplasmas needs to be analysed and this will entail confirming or finding their vectors. Detection methods will also need to be tested and antibiotics evaluated for their role in elimination of phytoplasmas in tissue culture.

Until relatively recently, the detection of banana viruses relied almost entirely on the inspection of plants for symptoms. Before the mid-1980s, the only threats recognized were from BBTV and CMV. Introduced banana was usually grown for 9–12 months in quarantine and examined at regular intervals for disease symptoms. As it had been reported that some strains of CMV could be carried as latent infections (Stover, 1972), the routine inoculation of cotyledons of indicator plants, such as cowpea (local lesion host) and cucumber (systemic host), with sap from young banana leaves was advocated. Soon after its creation in November 1984, INIBAP established an in vitro banana germplasm collection at the Laboratory of Tropical Crop Husbandry of the Katholieke Universiteit Leuven (KUL), Belgium. The function of this collection was not only to conserve valuable germplasm, but also to actively distribute superior clones to departments of agriculture, regional centres, international institutes and universities undertaking banana improvement programmes. To reflect this aspect of its work, the facility was named the INIBAP Transit Centre (ITC). The storage, multiplication and transfer of tissue-­ cultured material began in 1985 and it quickly became obvious that germplasm health was an important issue that needed to be addressed. One of the major concerns was to ensure that BBTV was not introduced to the western hemisphere through germplasm exchange. Another problem emerging worldwide with the development of ­tissue-culture techniques was how to ensure that material selected as a source of shoot tips for culture initiation and subsequent mass propagation was free of virus disease. It was now even more important than ­before to develop quick, reliable tests that would detect banana viruses (Dale, 1988). Virology laboratories in Australia, France, the Philippines and Taiwan became involved in projects to characterize BBTV and to develop rapid indexing tests that would allow for its early detection in germplasm in quarantine and material selected for in vitro propagation.



Quarantine and the Safe Movement of Musa Germplasm

In 1988, the Food and Agriculture Organization of the United Nations (FAO), the International Bureau of Plant Genetic Resources (IBPGR) and INIBAP organized a meeting of banana virologists and quarantine experts at the University of the Philippines, Los Banos (UPLB) to define protocols for the international transfer of Musa germplasm. The meeting reaffirmed that all Musa movement should be by tissue culture and that the only quarantine risk from in vitro material was the possible spread of virus diseases. The virus diseases identified were bunchy top, banana mosaic, banana streak and bract mosaic, which had recently been recognized in the Philippines as something different from mosaic. The detection of banana virus diseases by visual observation was seen as inadequate, following reports proving that BBTV could be carried in banana tissue cultures without expressing obvious symptoms (Drew et  al., 1989). Unfortunately, although herbaceous indicator and serological tests were available to detect CMV, reliable diagnostic tests for other banana viruses were only just being developed. The spread of BBTV was perceived to be the biggest threat associated with the movement of banana in tissue culture, and protocols advocated at this time for international transfer and quarantine reflected this concern. It was thought important that the quarantine environment should be optimum for the expression of BBTV symptoms and that, as a consequence, plants in quarantine should be grown at 32°C under conditions of high natural light, if necessary supplemented with artificial light. The recommendations of the meeting were published as the FAO/IBPGR Technical Guidelines for the Safe Movement of Musa Germplasm (Frison and Putter, 1989). As a result of the UPLB meeting, an international system of safe germplasm transfer was initiated under the auspices of INIBAP. INIBAP established two virus indexing centres, utilizing the banana virology expertise available at the Queensland Department of Primary Industries (QDPI) in Brisbane, Australia, and the Centre de coopération internationale en recherche agronomique pour le développement (CIRAD) in Montpellier, France. The task of these centres was to quarantine and test accessions held by INIBAP at the ITC so that they could be cleared for international movement. Approximately 20 tubes of proliferating tissue cultures of each

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Musa germplasm accession are held at the ITC. All this material is derived from a single shoot tip. Following recommendations made in the FAO/IBPGR Technical Guidelines for the Safe Move­ ment of Musa Germplasm, a representative sample of five plantlets per accession was dispatched to an INIBAP virus indexing centre. Four were grown in quarantine for 1 year (or longer if the eighth-leaf growth stage had not been reached after 1 year), inspected weekly for virus symptoms and indexed for CMV. The fifth plant was held in reserve in case of loss of a test plant. As an extra precaution against the spread of BBTV, accessions originating from continents where bunchy top was present were sent for testing at both of  INIBAP’s virus indexing centres. When an accession had been cleared, INIBAP could distribute tissue cultures derived from stocks remaining at the ITC. The next step forward came in 1990, when INIBAP organized a meeting of bunchy-top specialists at the laboratories of CIRAD’s Institut de recherche sur les fruits et agrumes (IRFA) in Montpellier, France. The objective of this meeting was to compare the various diagnostic tests for BBTV that had been developed and make recommendations on indexing requirements. An addendum to the FAO/IBPGR Technical Guide­ lines on the Safe Movement of Musa Germplasm was published as a result of the meeting. This addendum detailed agreed changes to the BBTV indexing protocol, which now included the use of BBTV-specific monoclonal antibodies (for use in enzyme-linked immunosorbent assay (ELISA) tests) and complementary DNA (cDNA) probes. Petiole sap was recommended to test for BBTV, as virus concentration was high in phloem tissue, and it was thought that tests should be conducted twice during the quarantine period. It was deemed important that ten new leaves should be produced by each plant in quarantine, the youngest being at least 1 m in length, to allow for the expression of bunchy top symptoms (Iskra-­ Caruana, 1994). The ‘state-of-the-art’ quarantine protocols being used in Australia at this time to prevent the entry of banana pathogens reflect the leading role that virologists and quarantine specialists in this country were playing in determining international standards (Jones, 1991; Thomas, 1991). Over the next 5 years, it became evident that BBTV, although still important, was not the

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main cause for concern. Not once was BBTV found in ITC germplasm indexed at INIBAP virus indexing centres. In fact, accessions that were originally collected because they were ­infected with BBTV also indexed negative, indicating a possible loss of the virus after years of storage and subculturing. Work in Australia had also suggested that BBTV is not always passed on to all plantlets derived from infected tissue cultures (Drew et al., 1992). BSV was the pathogen being detected more and more frequently at INIBAP virus indexing centres in germplasm from many countries and also from banana-breeding programmes. Previously, BSV had only been found in a few African countries, but it now appeared to have a worldwide distribution. Streak symptoms were often slow to develop in plants in quarantine and sometimes they failed to appear during the quarantine period. The most reliable technique for detecting BSV at the time was the examination of leaf sap under the electron microscope for bacilliform particles, and this became mandatory at INIBAP virus indexing centres. Antisera also became available as a result of research into sugarcane bacilliform virus (SCBV) and BSV that was being undertaken at the University of Minnesota by B.E.L. Lockhart. However, there were serious doubts as to whether they could detect all strains of the virus. At this time, as part of its International Musa Testing Programme sponsored by the United Nations Development Programme (UNDP), INIBAP commissioned Dr Lockhart to develop a reliable test that would detect all strains of BSV. Increasingly, through the routine use of the electron microscope to screen ITC and other germplasm, isometric and filamentous particles that could not be identified were being found in banana sap (Jones, 1994). Sometimes symptoms were associated with the presence of these viruses, but often no obvious symptoms could be seen. The significance of the viruses was unknown and work was needed to clarify the situation. Until 1995, the Philippines was the only known location of bract mosaic disease. In 1995, D.R. Jones recognized that virus symptoms in many banana cultivars in southern India were identical to those of banana bract mosaic virus (BBrMV) in the Philippines. It was later confirmed that BBrMV was the probable cause

of kokkan disease of ‘Nendran’ (AAB, Plantain subgroup) and diseases of other cultivars in India. Symptoms of BBrMV were also seen by D.R. Jones on banana in Sri Lanka. This was confirmed from banana specimens sent from Sri Lanka (Thomas et al., 1997). It became apparent that the distribution of this serious pathogen was greater than had been previously thought and that a reliable diagnostic test for BBrMV was urgently needed. INIBAP proposed and later funded research to characterize, investigate and develop diagnostic tests for BBrMV and the unknown viruses seen under the electron microscope. Because INIBAP had been adjusting its indexing protocols and requirements in line with new knowledge on the detection and distribution of Musa viruses, the procedures being advocated by INIBAP in 1994 to screen banana germplasm for virus diseases differed significantly from those outlined in the FAO/IBPGR Technical Guidelines on the Safe Movement of Musa Germplasm (Jones and Iskra-Caruana, 1994). It became obvious that another meeting of banana virus experts was necessary to update the FAO/IBPGR Technical Guidelines of 1989 and its addendum of 1990. The recommendations of the meeting of banana virologists that took place under the auspices of FAO and the International Plant Genetic Resources Institute (IPGRI) in Rome in June 1995 closely followed the protocols for virus detection followed by INIBAP (Jones, 1996). Modifications advocated included the testing of a larger proportion of a tissue-cultured accession held at the ITC, to reduce the chances of the sample being unrepresentative of the health status of the ‘mother stock’, and a shortening of the quarantine period, because of increased confidence in indexing tests. The recommendations, as the new FAO/ IPGRI Technical Guidelines for the Safe Movement of Germplasm Musa spp. (Diekmann and Putter, 1996), were available on request from IPGRI. The guidelines offered the best possible advice at the time of publication for institutions and commercial companies involved in the international transfer of Musa. They also included descriptions of abacá mosaic, banana mosaic, bract mosaic, bunchy top and banana streak diseases and the protocols of tests to detect the virus pathogens.



Quarantine and the Safe Movement of Musa Germplasm

Since the 1995 Rome meeting, it has been recognized that the high rate of BSV infection in bred tetraploids may be related to the de novo synthesis of the virus from BSV genomic sequences integrated into the DNA of Musa. This synthesis seemed to be associated with in vitro culture and possibly other stress factors related to embryo rescue In addition, banana virus X, a new virus disease problem, was reported. It was

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thought at the time that further advances in our knowledge of banana virus problems would undoubtedly continue to be made and that these advances would eventually require further revision to the Technical Guidelines. A new set of guidelines was published in 2015 (Plate 13.7) (Thomas, 2015). Germplasm that has been fully indexed is available for distribution from the ITC.

Plate 13.7.  Front cover of J.E. Thomas (ed.) (2015) Technical Guidelines for the Safe Movement of Musa Germplasm, Bioversity International, Rome (photo: D.R. Jones).

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Virologists concerned with the safe movement of Musa germplasm The ITC, which is operated by Bioversity International under the auspices of FAO, is hosted by the Katholieke Universiteit Leuven, Belgium. Its role, as before, is to facilitate safe exchange of Musa germplasm. With over 1500 accessions, the ITC contains the largest banana in vitro collection in the world (Thomas, 2015). Virology research units having expertise with Musa viruses, and collaborating with Bioversity International for the virus indexing of banana, currently include the following.

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Centre de coopération internationale en recherche agronomique pour le développement (CIRAD), Montpellier, France (Officer-­inCharge: Marie-Line Iskra-Caruana); University of Queensland, Brisbane, Australia (Officer-in-Charge: John Thomas); University of Liege, Gembloux Agro-Biotech, Gembloux, Belgium (Officer-in-Charge: Sébastien Massart), currently Bioversity International’s Virus Indexing and Sanitation Centre (VISC); and International Institute of Tropical Agriculture (IITA), Ibadan, Nigeria (Officer-­in-­ Charge: P. Lava Kumar).

The virologists in charge of the banana virus indexing centres listed above together with the assistance of Murray Sharman (Queensland Department of Agriculture and Fisheries, Australia), Ludivine Lassois and Caroline De Clerck (University of Liege, Gembloux, Belgium), Matthieu Chabannes (CIRAD, Montpellier, France), Pierre-Yves Teycheney (CIRAD, Capesterre-Belle-Eau, Guadeloupe), Ines Van den houwe (ITC, Leuven, Belgium) and Nicolas Roux (Bioversity International, Montpellier, France) developed the current guidelines and recommendations for the a safe movement of Musa germplasm in 2015.

Current Recommendations for the Safe Movement of Musa ­Germplasm The following general recommendations apply for all international Musa germplasm exchanges (Thomas, 2015).

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Germplasm should be obtained from the safest source possible. A pathogen-tested Musa germplasm collection is held in vitro at the ITC from which safe germplasm can be ordered online (http://www.crop-diversity. org/banana). All germplasm moving from one continent to another should transit through the ITC or, if possible, be obtained from the ITC. Other sources of safe germ­ plasm may be available where indexing laboratories have the capacity and expertise to test for the complete range of viruses. All germplasm should ideally be distributed in the form of tissue culture. If this is not possible, full quarantine measures, including containment in secure post-entry quarantine facilities, must be taken until the vegetative material or seed is cultured in vitro. Germplasm should be tested for all viruses known to affect Musa according to the protocols described in these guidelines. However, in some instances tests may be omitted if there is reliable evidence that particular viruses are not present in the country of origin of the germplasm. Indexing procedures and results should be  documented in a germplasm health statement. The movement of germplasm should be carefully planned in consultation with quarantine authorities and the relevant indexing laboratory, and should comply with the regulatory requirements of the importing country. International standards for phytosanitary measures (ISPMs) as published by the Secretariat of the International Plant Protection Convention (IPPC) should be followed during the movement of germplasm. In accordance with IPPC regulations, any material being transferred internationally must be accompanied by a phytosanitary certificate.

The following technical protocols should be followed when preparing vegetative material for virus indexing and distribution (Thomas, 2015).

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Select a sucker from a plant without symptoms of systemic pathogen infection. Trim the sucker to remove soil, roots and any other extraneous material, leaving part of the central corm containing the meristem



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Quarantine and the Safe Movement of Musa Germplasm

and about 10 cm of the pseudostem tissue above it. The overall dimensions of the block of tissue should be about 20 cm high and 10−15 cm in diameter. Air-dry the block for 24 h and wrap in newspaper. Label the material and dispatch in a cardboard box using the fastest transportation method. Plastic should not be used for wrapping. Send the material to the nearest appropriate tissue culture laboratory in the country of origin, or if this is not possible, to a tissue culture laboratory preferably in a non-­ banana-growing area. The meristem tip should be excised, surface disinfected (Hamill et  al., 1993) and cultured aseptically (Strosse et  al., 2004; Van den houwe and Swennen, 2000; Vuylsteke, 1989; Wong, 1986). A single meristem-tip-derived subculture should be further cloned to seven plantlets, of which five should be sent to an indexing facility and two should remain in culture for future multiplication. At the indexing facility, four plants should be established in a potting mix in a vector-free, insect-proof greenhouse under conditions conducive to vigorous plant growth. The fifth plant serves as a back-up. The plants to be indexed should be physically isolated from other banana plants so as to minimize the possibility of cross-contamination. After 3 and 6 months of growth, tissue samples should be taken from the three youngest expanded leaves and indexed for viruses. In addition, electron microscopic observations should be undertaken at both 3 and 6 months to look for the presence of any undescribed viruses. If all tests are negative, the four indexed plants may be released and the cultures derived from the two remaining plants in vitro may be further propagated and distributed in vitro. If found to be infected, the material should be subjected to a virus eradication treatment if available. The plant material must not be released until its virus-free status has been confirmed For the movement of in vitro material, neither charcoal nor antibiotics should be

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added to the culture medium, as these may mask contamination in the culture. In vitro cultures should be shipped in transparent tubes, preferably plastic, and visually inspected for bacteria, fungi and arthropods. Contaminated germplasm should be destroyed. Special care should be taken to protect the material from extremes of temperatures (not below 15°C and not above 35°C) during shipment.

Musa seed and embryos can transmit some viruses. If the germplasm is collected as seed, the following technical protocols should be used before international movement (Thomas, 2015).

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Collect seeds from plants that are free from pests and disease symptoms. Make sure seeds are free of pulp and there is no visual evidence of insects, mites or nematodes. Air-dry and, if necessary, fumigate. Banana seeds are sensitive to desiccation and should be moved immediately to an appropriate tissue culture laboratory in the country of origin, or if not possible, to a tissue culture laboratory preferably in a non-banana-growing area. Seed should be surface-disinfected. The embryos should be isolated and germinated in vitro to establish sterile cultures (Bakry, 2008; Pancholi et  al.,1995; Uma et  al., 2011). Materials derived from embryo culture should be indexed and moved in the same way as material derived from meristem tip culture.

Virus Detection and Therapy The virus detection methods now recommended by Bioversity International’s virus indexing centres are as follows (Thomas, 2015).

Banana bunchy top virus (BBTV) BBTV can be detected by ELISA on samples from field plants, tissue cultures and infective aphids (Thomas and Dietzgen, 1991; Thomas et  al., 1995). Monoclonal and polyclonal antibodies

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likely that meristem tip culture will be effective, as it is for BBTV (Ramos and Zamora, 1990; Wu and Su, 1991; Thomas et al., 1995).

are commercially available (Wu and Su, 1990; Dietzgen and Thomas, 1991. Polymerase chain reaction (PCR) assays are a sensitive alternative for detection (Hu et  al., 1996; Sharman et  al., 2000; Mansoor et  al., 2005) and DNA probes and primers are available for BBTV DNA components 1 to 6 (Burns et  al., 1995; Stainton et  al., 2012). Recently methods based on real time PCR (Chen and Hu, 2013), loop-mediated isothermal amplification (LAMP) (Peng et al., 2012b), electrochemiluminescence PCR (Tang et al., 2007) and impedance spectroscopy (Majumder et al., 2013) have also been described for the detection of BBTV. However, these techniques have not yet been tested against a genetically diverse range of BBTV isolates to determine their wider applicability. Multiplex PCR assays have also been reported (Sharman et al., 2000; Liu et al., 2012). The optimal tissue for indexing is the midrib of the youngest symptomatic leaf, though the virus can also be detected in the leaf lamina, pseudostem, dormant meristems, the bunch stalk, bracts and fruit peel from infected plants. The virus can sometimes be detected in the leaf formed before the youngest symptomatic leaf. Meristem or shoot tip culture (< 1 mm to 2  mm), possibly combined with heat therapy (Ramos and Zamora, 1990; Thomas et al., 1995; Helliot et al., 2001; Lassois et al., 2013), has been successful in achieving a proportion of virus-free plantlets. Testing after treatment (ideally after an 8−12-month period in the greenhouse) is essential in case the virus is reduced to undetectable levels, but not eliminated, during tissue culture.

The virus can be detected by ELISA using antibodies prepared against the abaca strain of sugarcane mosaic virus (Gambley et  al., 2004), or by RT-PCR (A.D.W. Geering and co-workers, Australia, 2014, unpublished information). No information on therapy methods have been published.

Abacá bunchy top virus (ABTV)

Cucumber mosaic virus (CMV)

The virus can be detected by ELISA in field plants. Some commercially available BBTV monoclonal and polyclonal antibodies (from Agdia, Inc., Elkhart, Indiana) are known to cross-react with ABTV, albeit with a reduced sensitivity. Specific PCR assays are a sensitive and reliable alternative for detection (Sharman et  al., 2008). Some PCR primers for BBTV are known to cross-react with ABTV (Su et al., 2003). ABTV is likely to be phloem limited and the optimal tissue for indexing is the midrib of the youngest leaves. There are no reports on attempts to eliminate ABTV from infected germplasm, but it is

A range of techniques can be used to detect CMV, including observation of symptoms, electron microscopy, the use of indicator plants, nucleic acid hybridization, serology and PCR. The latter two are recommended for reliable and sensitive routine detection of CMV. A number of ELISA kits are commercially available and several PCR assays have been described (e.g. Wylie et  al., 1993; Hu et  al., 1995; Sharman et  al., 2000), including multiplex assays (Sharman et al., 2000; Liu et al., 2012). Different methods have been developed and tested for CMV eradication, such as thermotherapy, chemotherapy, meristem culture and cryotherapy.

Banana bract mosaic virus (BBrMV) Leaf symptoms can be erratic and bract symptoms are evident only during flowering. Dead leaf sheaths must be removed to reveal mosaic and streaking on the pseudostem. The virus can be detected in extracts of leaf lamina, midribs and flower bracts by ELISA using BBrMV-specific polyclonal and/or monoclonal antibodies. The virus can also be detected by RT-PCR (Iskra-Caruana et  al., 2008), immunocapture RT-PCR (Sharman et al., 2000) or LAMP assays (Siljo and Bhat, 2014). Multiplex (IC)-PCR ­assays are available (Sharman et al., 2000; Liu et al., 2012). Both meristem culture and heat treatment of shoot cultures have resulted in elimination of BBrMV from infected ‘Señorita’ (AA) (Ramos and Zamora, 1999). Abacá mosaic virus (AMV)



Quarantine and the Safe Movement of Musa Germplasm

A combination of these techniques can improve the eradication rate (Gupta, 1986; Helliot et al., 2002, 2004). The efficiency of the eradication technique depends on the characteristics of the virus strain, the type of tissue treated and the banana genotype. However, meristem culture from heat-treated in vitro plantlet is recommended for best results. Protocols are available in Lassois et al. (2013). Banana streak virus (BSV) The main difficulty in diagnosing BSV is to distinguish active infections (episomal) from integrated viral sequences. Although infectious eBSV are only present in the M. balbisiana genome, all Musa spp. also carry partial or ‘dead’ integrated BSV sequences in their genomes. Thus PCR tests on any banana DNA can result in BSV-positive reactions. Diagnostic PCRs must be designed to accommodate this problem. Serological detection of BSV is complicated by the wide serological diversity among virus isolates, some of which are serologically unrelated (Lockhart and Olszewski 1993). A polyvalent polyclonal antiserum raised against BSV species and Sugarcane bacilli­ form virus species (Ndowora, 1998) is capable of detecting all known isolates by immunosorbent electron microscopy (ISEM) in partially purified extracts, even in asymptomatic leaf tissue (B.E.L. Lockhart, USA, 2014, unpublished information). Plants derived from in vitro material should be tested at least 3 months after plantlets have been acclimatised and planted out. ELISA test kits for BSV are commercially available from Agdia and DSMZ companies. Samples of laminar and midrib tissue from the three youngest expanded leaves, or symptomatic tissue if present, should be tested by ELISA. A partial purified extract is used to observe virus particles under the electron microscope (Geering et al., 2000). Immunocapture PCR (IC-PCR) to detect specific BSV species has been described (Geering et al., 2000; Le Provost et al., 2006). Immunocapture conditions were optimized to minimize non-specific binding of plant genomic DNA to the immunocapture tubes. Alternatively, to avoid contamination by plant genomic DNA, a DNase I treatment is performed. Rolling circle amplification (RCA) is a technique that selectively amplifies circular DNA

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(such as encapsidated BSV genomic DNA) resulting in a specific amplification of DNA from episomal forms of BSV only. James et al. (2011a) developed and used RCA with the addition of BSV-specific primers for the detection of episomal BSV and demonstrated the ability of this method to differentiate between episomal and integrated viral genomic sequences. This method avoids the need for antiserum, but does require knowledge of the virus sequence in the selection of restriction enzymes required for visualization of the virus-specific products. Identification of a particular BSV also requires subsequent PCR or sequencing. LAMP assay is a DNA amplification technique that does not require thermal cycling (Notomi et  al., 2000). LAMP products can be detected using conventional agarose gel electrophoresis or using visual observation to estimate turbidity or colour changes (Mori et  al., 2001). Peng et al. (2012a) used the LAMP assay to detect BSV particles in infected banana plants. However this technique cannot differentiate between integrated and episomal forms and so false positives could be very frequent. Also, primer design is complicated and is likely to be a constraint with the genetic diversity found in germplasm indexing. Based on the sequences and the structure of the different eBSV described in the seeded diploid M. balbisiana (BBw, accession ‘Pisang Klutuk Wulung’), several PCR markers have been developed in order to genotype those integrations in different banana genotypes (Gayral et al., 2008, 2010; Chabannes et al., 2013), including: (i) integration markers targeting eBSV insertion sites; (ii) structure markers targeting internally reorganized eBSV structures; and (iii) allelic markers that allow the distinction between alleles of the same eBSV species. The targeted eBSV, the type of marker developed, the name and the sequence of the primer as well as the size of the PCR product were described in Chabannes et al. (2013). Cryopreservation followed by apical meristem culture has a high rate of success in eliminating episomal BSV infections in ‘Williams’ (AAA, Cavendish subgroup), a banana genotype free of eBSV (Helliot et al., 2002). Two anti-retroviral and anti-hepadnavirus molecules have also been effective in eradicating the episomal form of BSV from banana plants (Helliot et al., 2003). Methods are described in Lassois et al. (2013).

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Banana mild mosaic virus (BanMMV) Polyclonal antisera to BanMMV have been developed (Teycheney et  al., 2007; J.E. Thomas, Australia, 2015, unpublished), though they are not commercially available. These antisera can be used in immunocapture RT PCR or in ISEM. BanMMV displays a high degree of genetic diversity, making detection by PCR particularly challenging. A set of 154 partial RdRp sequences showed a mean pairwise nucleotide sequence divergence of 20.4% (Teycheney et  al., 2005) and similarly, a set of 14 complete coat protein sequences showed a mean of 20.6% divergence (J.E. Thomas, Australia, 2014, unpublished information). A group of partial CP sequences, primarily from Côte d’Ivoire, was particularly variable compared with all other isolates (mean 35%) and also within the group itself (mean 35%). Teycheney et al. (2007) described a nested immunocapture RT PCR assay for the detection of BanMMV, targeting the RdRp region of the flexiviridae genome and based on the primers of Foissac et al. (2005). This assay is not specific for BanMMV, but does not detect banana virus X, another member of the Betaflexiviridae (Teycheney et al., 2007). Alternative primers, targeting the coat protein gene (upstream primer) and the poly-A tail (downstream primer), have been developed (Gambley et  al., 1999; Teycheney et  al., 2005; M. Sharman and J.E. Thomas, Australia, 2014, unpublished information). The published upstream primer sequence of BanCP2 (Teycheney et  al., 2005) contains a probable typographical error at positions 7042−7066 on the BanMMV genome. This primer has from one to three mismatches within the first eight 3′ bases in 23 of 54 available partial CP sequences on GenBank. Upstream primer BanMMV CP2 similarly has mismatches in the first seven bases against 12 of 54 sequences. However, there is good correlation between ISEM results and IC RT PCR results when the latter primer has been used for extensive testing of ­banana germplasm (K.S. Crew and J.E. Thomas, Australia, 2014, unpublished information). The latter primer has been modified to eliminate the majority of mismatches in the 3′ end (J.E. Thomas, Australia, 2014, unpublished information). BanMMV has been eliminated from ‘Pisang Awak’ and ‘Pisang Seribu’ using meristem culture and chemotherapy with a number of chemicals

(of which ribavirin was clearly the most e­ ffective) (Busogoro et  al., 2006). Meristem culture alone has also inadvertently eliminated BanMMV from selections of ‘Ducasse’ (ABB, Pisang Awak subgroup) (S.D. Hamill, Australia, 2014, personal communication).

Banana virus X (BVX) A nested RT PCR has been reported for the detection of BVX (Teycheney et al., 2007). No method for its eradication has been published.

Results of Virus Indexing at the Virus Indexing Centres The results of virus indexing of the ITC’s Musa accessions undertaken at the Virus Indexing Centres from 2007 until 2013 has been published by De Clerck et al. (2017). The first step in the indexing process, called pre-indexing, involved only molecular tests and was designed as a pre-screen for new germplasm lines or existing accessions. The second step, called full indexing, was undertaken on either older existing accessions or newer accessions, which had tested negative during previous pre-indexing. It involved observation in a secure post-entry quarantine facility (Plate 13.8) for virus symptoms, molecular tests and inspections for virus particles by transmission electron microscopy (TEM). The pre-indexing protocol of 434 samples representing 270 accessions revealed a high ­level of virus infection. The results showed 66.9% of samples were infected with BSOLV and/or BanMMV. Mixed infections were common. BSOLV was found in 43.9% of samples and BanMMV in 47.8%. BBrMV was detected in 1.8% and BBTV in 0.8%. CMV was not ­detected. Full indexing of 243 accessions, 68 of which had passed the preliminary pre-indexing tests, revealed 68% were free of virus. BBTV, BBrMV and CMV were not detected in any accession. BanMMV was detected in 46 accessions from Honduras, India, Papua New Guinea, ­Tanzania, Uganda, Vietnam and locations unknown. BSV was found in 44 accessions from Cameroon, ­India, Indonesia, Papua New Guinea, Tanzania,



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Plate 13.8.  Young banana plants established from introduced tissue cultures in a secure post-entry quarantine facility in Brisbane, Australia (photo: D.R. Jones, QDPI).

Vietnam and locations unknown. BSOLV was the most commonly detected BSV species, being found in 32 of the 44 accessions with BSV. B ­ SGFV was found in four accessions and BSIMV in another four. In another four accessions, which had ­tested negative by PCR for the six known BSV species, bacilliform particles indicating BSV were observed. This result highlighted the importance of the non-specific ISEM assay for BSV detection. Mixed infections of BSV and BanMMv were found in 22 accessions. BSV and an unidentified virus in three, and BanMMv and an unidentified virus in one. Higher rates of BSV infection were detected in B genome-containing Musa genotypes (58% for pre-indexing and 40% for full indexing) compared with accessions lacking the B genome

(12% for pre-indexing and 3% for full indexing). No clear difference in the percentage of ITC accessions that tested positive for BSV was observed between Musa genotypes containing the B genome (BB, AB, BBB, ABB, AAB, AAAB, AABB, ABBB). A significant proportion of the samples tested during pre-indexing was infected with at least one virus, demonstrating to the authors (De Clerck et  al., 2017) the utility of this early pre-screening step. The accessions that went through both pre-indexing and full indexing recorded fairly similar virus detection results, showing the reliability of the methods used. The authors concluded that the results suggested a strong complementarity between the two steps and showed the value in sequential testing.

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Index of Musa Species, Banana Subgroups and Banana, Abacá and Enset Cultivars

Page numbers in italics refer to illustrations. Musa species

Musa acuminata 9–10 Musa acuminata ssp. banksii 29–30, 31, 570 bacterial wilts  317, 320 fungal leaf diseases  123, 134, 138, 159, 166, 171 nematodes 433 viruses 372 Musa acuminata ssp. burmannica 56, 63, 78, 84, 87, 131, 434, 441 Musa acuminata ssp. errans  29–30, 123 Musa acuminata ssp. malaccensis  29, 123, 219, 257, 434 Musa acuminata ssp. microcarpa  29, 123, 257, 434 Musa acuminata ssp. siamea  123, 374 Musa acuminata ssp. truncata 123 Musa acuminata ssp. zebrina  29, 30, 317, 373, 434 Musa balbisiana  9–10, 27, 30, 31 bacterial wilts  312, 320 drought tolerance  510 fungal leaf diseases  80, 123, 134, 138, 144, 155, 160–161 nematodes 433 viruses  372, 374, 389–390, 393, 400, 404 wind tolerance  492 Musa boman 138 Musa chiliocarpa 25 Musa coccinea 373 Musa ensete see enset in general index



Musa ingens 2 Musa jackeyi  123, 373 Musa lolodensis  31, 317 Musa maclayi  31, 123, 138 Musa ornata  123, 373 Musa paradisiaca 22 Musa peekelii  31, 123 Musa sapientum 23 Musa schizocarpa  27, 123, 138, 166, 171, 317 Musa textilis see abacá in general index Musa velutina 373

Banana Subgroups

Bluggoe 27 Cavendish  3, 9, 14–17 fruit diseases  256, 257, 270, 275, 284, 341 fungal leaf diseases  131, 134, 140, 143, 155, 160, 166, 171 black leaf streak  52, 78–79, 123 Fusarium wilt  207, 217–219, 227, 572 genetic variants dwarfs 547–551, 549, 550, 551 foliage  552, 554 giants 551, 552, 553 malformed fruit  467, 468 nematodes 434 postharvest chilling  515 pseudostem heart rot  239

593

594

Index of Musa and Ensete Species

Cavendish (Continued ) root and rhizome rots  231–232, 330, 333 viruses 390 weather damage  492, 501, 511 Fe’i 2, 11, 31–32, 32, 123 Gros Michel†  16 Ibota 19 Iholena  21, 23 Kathali 237 Lakatan 19 Laknau 24 Lujugira–Mutika  7, 16, 17, 18, 30, 49, 124, 156, 227 Mai’a Maoli-Popoulu  21, 22–23, 124, 134, 171 Monthan  25–26, 237 Mysore 24 Ney Mannan  28 Pisang Awak  28 Pisang Lilin  13, 14 Plantain 19–22 bacterial diseases  321, 327, 336, 338 fruit diseases  261, 270, 275 fungal leaf diseases  124, 135, 141, 145, 155, 171 black leaf streak  46–47, 48, 49, 79 Fusarium wilt  226 genetic variants  549, 559 nematodes  430, 434, 439, 441, 443 non-infectious disorders  462 Pome  21, 22, 124 Red 19 Saba  26, 28 Silk  23, 24 Sucrier  13, 14, 171

Banana, Abacá and Enset Cultivars

*abaca **enset

1319-01 320 1741-01 320 2390-2 124

Abuhon  26, 373 Ado** 306 Agbagba  20, 21, 131, 156, 267, 443–445, 449 Alaswe 447 Alukehel  26, 256, 256 Amas 14 Ambrosia 284 Anamala 16, 128 Annan 147

Apantu-pa 441 Arthiyakol 10 Ash Plantain  256 Astara** 306

Babi Yadefana  320 Balingdag* 328 Basrai 371 Bhimkol 10 Blue Java  171, 172, 256 Bluggoe  25, 27, 27, 112, 124, 138, 171, 226, 306, 314, 434 Bodles Altafort  124 Bongolanon*  234, 328 Boothibale 241 Brazil 335 BRS Conquista  112 BRS Garantida  112 BRS Platina  112 BRS Preciosa  112 BRS Princesa  112 BRS Tropical  112 Buffare** 306 Bungaoisan 373

Cachaco 338 Caipira  112, 142 Calypso  124, 284 Caprichosa 112 Cardaba  321, 434 Cevvazhai/Sevazhai  20 Chato  112, 318 Chuo͂́i Bom  161 Chuo͂́i Mit  414 Chuo͂́i Ngop Dui Duc  404 Chuo͂́i Ngu Tien  404, 444 Chuo͂́i Tay Tia  414 Cocos 434, 493, 547 Corne 155 Curraré 67–68

Daluyao 410 Daru 345 Dominico 338 Dominico Hartón 343 Ducasse  159, 161, 171, 410 Dwarf Bluggoe  25, 27 Dwarf Cavendish  14–15, 16, 17, 548, 550 bacterial wilts  304, 312, 321 choke  17, 495 drought 511 fruit disorders  263, 468 fungal leaf diseases  79, 172 iron deficiency  555



Dwarf Dwarf Dwarf Dwarf

Index of Musa and Ensete Species

Namwa  141 Nino  141 Parfitt  550, 551 Valery  131

Embul 378 Espermo 338 Extra-Dwarf Cavendish  15, 16, 549–550, 551

F2P2 320 Fa’i Misiluki  132, 135 False Horn  19, 21, 155, 554, 559 FB918  438, 442 FB919  438, 442 FB920 442 FB924  438, 442 FHIA-01 (Goldfinger)  124, 227, 235, 275, 284, 438, 492, 501 FHIA-02 (Mona Lisa)  227, 275, 338 FHIA-03 320 FHIA-17 112 FHIA-18  64, 78, 131, 284, 577 FHIA-20 112 FHIA-21  64, 112, 131, 235, 577 FHIA-23  235, 275 FHIA-25  112, 577 Figue Pomme  412 Figue Pomme Ekonah  434 Figue Rose  155 Figue Sucrée  155 Focanah 434 Fougamou  78, 84 French  19, 21, 22, 22, 462 French Clair  131, 412 French Horn  19, 21

GCTCV-119  131, 471 Genticha**  305, 306 Geziwet** 306 Giant Cavendish  15, 16, 18, 227 Goldfinger (FHIA-01)  124, 227, 235, 275, 284, 438, 492, 501 Grand Nain  166, 393, 467, 468, 492, 495, 501, 511 genetic variants  547–548, 550, 551, 552, 553, 554, 555, 559, 560–561, 560, 562 Grande Naine  15, 16, 17, 79, 84, 87, 141, 155, 232, 338, 441, 444 Green Red  19, 123, 138 Gros Michel  9, 14 bacterial rots  330, 338 fruit disorders  260, 284, 341 non-infectious  470, 471

595

fungal leaf diseases  123, 134, 138, 155, 171, 172 Fusarium wilt  207, 215 genetic variation  547 low temperature damage  515 nematodes  434, 443 phytoplasma 343 pseudostem heart rot  238 viruses  373, 374, 410 Guangen 335 Gulumo** 306 Guyod 434

Hartón 338 Horn  19, 21, 124, 312, 462

I.C.2  124, 155 Inambak 447 Inarnibal  13, 14, 123, 138, 320 IRFA 909  577 IRFA 910  577 IRFA 914  577

Japira 112 J.D. Dwarf   550–551 Jinfen 335

Kalapua  13, 26, 345 Kandarian 235 Karpuravalli  241, 275 Katali 373 Kluai Hom Thong  275 Kluai Khai  14, 156 Kluai Khai Bonng  124 Kluai Khai Thong Ruang  87 Kluai Lep Chang Kut  29 Kluai Lep Mu Nang  15, 131 Kluai Namwa  7, 26, 161, 169 Kluai Namwa Daeng  26 Kluai Namwa Khao  26 Kluai Namwa Khom  26, 161 Kluai Nhoen  12 Kluai Tani  404 Kluai Teparot  12, 26, 29, 124, 138, 171, 374 Kullo** 306 Kunnan  13, 441

Lacatan  155, 263, 275, 284, 333, 466, 468, 492, 493 Lady Finger  134, 159, 171 Lakatan  123, 375, 393 Laknau 320 Latundan  239, 393

596

Index of Musa and Ensete Species

M9  306, 312 Maguinanao* 328 Mala 143 Manoranjitham 131 Manzano  234, 338 Maqueño  215, 338 Mata Kun  123 Mbwazirume  49, 312 Mezye 305 Mons Mari  159 Monthan 171 Mshare 17 Muraru 17 Musakala 16 Mysore  23, 112, 124, 131, 138, 168, 171, 226, 394, 576

Nakabululu 16 Nakitembe  16, 304, 312 Nathan  548, 550 Nendran  147, 378, 379, 469 Ney Poovan (Safet Velchi)  13, 14, 15, 124, 241, 434 Nfuuka 16 Njock Kon  155 Nongke No. 1  335

Orishele 141 Orito 156 Orotava 19

Pa-a Dalaga  447 Pachanandan 21 Pacific Plantain  23, 134 Pacovan  21, 22 Pacovan Ken  112 Paji 19 Paka  13, 63, 78, 123, 441 Palayankodan 147 Pastilan 447 Pata Sina  28 Pei-Chiao  166, 471, 495 Pelipia 373 Pelipita  26, 28, 112, 171, 321, 434 Petite Naine  131, 155 Peyan 26 Pisang Abu Keling  161 Pisang Abu Nipah  161 Pisang Abu Siam  161 Pisang Ambon  19 Pisang Awak  26, 570 anthracnose 284 bacterial wilts  304, 306, 327 fungal leaf diseases  124, 134, 138, 159, 171

fungal root diseases  226, 235 viruses 404 Pisang Batu  404 Pisang Batuau  434 Pisang Barangan  19, 20 Pisang Berangan  7, 20, 156, 166, 226, 275, 284, 447 Pisang Bungai  441, 443 Pisang Ceylan  78, 131, 434, 441 Pisang Embun  447 Pisang Jambe  12 Pisang Jari Buaya  13, 123, 434, 438 Pisang Jarum  414 Pisang Kelat  23, 124, 434 Pisang Keling  168 Pisang Kepok  28, 327 Pisang Klutuk Wulung  404, 557 Pisang Kole  414 Pisang Lidi Bikittinggi  414 Pisang Lilin (Pisang Lidi)  13, 78, 123, 434 Pisang Loka Bule  327 Pisang Loka Nipah (Pisang Unti Sayang)  327 Pisang Mas  78, 115–116, 131, 156, 226, 284, 335, 434, 441, 447 Pisang Masak Hijau  15, 16 Pisang Nangka  19, 156, 171, 284, 447 Pisang Oli  434 Pisang Puju  327 Pisang Raja  23, 25, 124, 138, 171, 373 Pisang Rastali  284, 447 Pisang Sepatu Amora (Pisang Tanjung)  327 Pisang Seribu  25, 26, 410 Pisang Susu  19, 123 Pisang Tandok  447 Pisang Tindok  168 Pisang Tongat  123 Pisang Tongka Langit Alifuru  32 Pitogo  29, 320 Plantain Vert  155 Pogpogon 447 Pome  138, 171, 215, 226 Pompo 338 Poovan  25, 147, 576 Popoulou Popoulou  155, 434 Poyo  155, 172, 443, 492 Prata Anã  21, 22, 284 Pundol 373 PV 42-53  284

Rasthali  24, 216, 466–467 Red  19, 123, 134, 138, 171, 548, 557, 558, 558 Red Dacca  19 Robusta  7, 17, 79, 275, 284, 333, 492, 511

Saba  112, 134, 138, 171, 320, 321, 374 Safet Velchi (Ney Poovan)  13, 14, 15, 124, 434



Index of Musa and Ensete Species

Sakali Ndizi  25, 26 Santa Catarina Prata  22, 22 Sauhu 345 SH-3142  124, 438 SH-3362  124, 320, 438 SH-3648 438 SH-3723 438 Silk  23, 124, 138, 171, 215, 226, 270 Silver Bluggoe  25, 27 Sinker 447 Som 412 Sucrier  13, 123, 134, 138, 141, 156, 171, 318, 320 Sugar 159 Sukali Ndizi  228

T6 124 T8  63, 78, 112, 124, 159 T12 124 Tañgoñgon*  328, 384 Thap Maeo  112, 284 Tinawagan Pula*  384 Tjau Lagada  441, 443 Turangkog 373 Tuu Gia  78, 123, 434

597

Umalag 393 Utin Iap  32 Uzakan  23

Valery  67–68, 113, 155, 272, 275, 284, 333, 468, 493 Veimama  172, 366, 368 Viente Cohol  447 Virupakshi 241 Vitória 112 Vovosi 345

Williams 17 fruit malformations  468 fungal leaf diseases  79, 87, 113, 116, 131, 135, 159 genetic variants  548, 555, 555, 559, 559, 560, 561, 562 rots  239, 335 weather damage  492, 495, 511

Yangambi Km 5  64, 78, 84, 87, 124, 131, 434, 441, 443

General Index

Page numbers in italics refer to illustrations. abacá bunchy top virus (ABTV)  362, 363, 369, 377, 584 abacá mosaic virus (AMV/SCMV-Ab)  392, 392, 584 abacá (Musa textilis)  2, 32–34, 33 abacá mosaic  391–393, 391, 392 bacterial heart rot  339 bacterial wilt  318, 328–329 bract mosaic  384 bunchy top  362, 363, 366–367, 369, 376–378, 377 Deightoniella leaf spot  138, 139, 141 Deightoniella pseudostem rot  233–234 freckle 166 Fusarium wilt  207, 213 Marasmiellus pseudostem and root rot  235–237, 236, 237 Mycosphaerella leaf speckle  159 nematode pests  429, 434, 439, 440 root-knot  445, 446 pseudostem heart rot  238, 238, 239 acetic acid treatment  91 acybenzolar-S-methyl 101–102 aerial application of fungicides  96–97, 106–107 Africa 568 BBTV 364 black leaf streak  43, 47–48, 48–49, 60–62, 63 Fusarium wilt  209 see also East Africa; West Africa; individual countries Agrobacterium vitis (leaf blight)  339 agrochemical-induced damage



de-suckering agents  535–536 disease and pest control agents  527–529, 528, 529 disinfectants 535, 536 fertilizers 527 herbicides 530–535, 532, 533, 534, 535 alligator skin  465–466, 465 Alpinia spp.  373 altitude banana 494 black leaf streak vs Sigatoka leaf spot  73–74 Cladosporium leaf speckle  155 enset 35 aluminium 541 ametryne  531, 533–534 ampeloviruses 413–414, 413 animals, as vectors of Xanthomonas bacterial wilt  305 anthracnose postharvest disease  51, 280–285, 280 preharvest disease  255–256, 256 antimicrobial compounds (phytoprotectants)  86–87, 283 Aphelenchoides ensete (nematode black leaf streak of enset)  449, 449, 450 aphid vectors of abacá bunchy top  376, 377, 377 of abacá mosaic  392–393 of banana bunchy top  370–371, 370 of banana mosaic  387–389, 389, 390 of bract mosaic  383 arbuscular mycorrhizal fungi  110, 232, 437, 448 Armillaria corm rot  228–229 arsenic 541

599

600

General Index

ascospores/asci black cross leaf spot  133 black leaf streak  58, 58, 69–73 Cylindrocladium root rot  230, 231 eumusae leaf spot  129, 130 freckle  168, 169 Mycosphaerella leaf speckle  157 Phaeoseptoria leaf spot  146 Sigatoka leaf spot  120, 121 Asia 568 BBTV 364 black leaf streak  44, 45–46, 60 Fusarium wilt  210–211 see also individual countries Athelia rolfsii (Sclerotium corm and pseudostem rot)  240–241, 241, 242 ATP-binding cassette (ABC) protein family  86 Australia Armillaria corm rot  228 bacterial finger-tip rot  340 black leaf streak  45, 112, 573–574 bunchy top  363, 374–375 Cordana leaf spot  136 freckle  169, 574–575 fruit speckle  265–266 Fusarium wilt  212, 224, 572 nematodes 440 quarantine  571, 572, 573, 587 Sigatoka leaf spot  126 azoxystrobin  101, 266, 284

Bacillus spp.  108–109, 126, 437 bacterial finger-tip rot of fruit  339–341, 340 bacterial leaf blight  339 bacterial leaf spot  339 bacterial rhizome and pseudostem rots  329–330 heart rot of abacá  339 pseudostem wet rot  336–338, 337 rhizome rot and tip-over  330–334, 331, 332 soft rot of rhizome and pseudostem  334–336 bacterial wilts abacá bacterial wilt  328–329 blood bacterial wilt  323–328, 324, 325 compared with Fusarium wilt  213, 298–299 Javanese vascular disease  329 Moko bacterial wilt and bugtok  314–323, 315, 316, 317 Xanthomonas bacterial wilt  296–314, 298–303 bagging of fruit on the plant  7–8, 9, 261 problems caused by  265, 467, 468, 472, 507 postharvest  276, 277 banana blast (Pyricularia leaf spot)  148–150, 148, 149, 150

banana bract mosaic virus (BBrMV)  381–383, 383, 580, 584 see also bract mosaic banana bunchy top virus (BBTV)  362, 367–369, 367, 370–371, 576 detection  369, 579, 583–584 see also bunchy top Banana Environmental Commission  106 banana mild mosaic disease  410–412, 410, 411, 586 banana mosaic  384–390 causal agent  385–387, 386, 578, 584–585 control  390, 584–585 host susceptibility  389–390 symptoms 384–385, 385, 386 on Commelina weeds  387 transmission 387–389, 389 banana streak  393–408 causal agent  396–403, 400 detection  403, 585 endogenous sequences (eBSV)  400, 402, 403, 404, 407, 585 phylogeny 399, 401 control  407–408, 585 diagnosis 403 distribution and spread  394, 576–577 epidemics 406–407 host susceptibility  403–404 and safe movement of germplasm  580, 581, 585 symptoms 394–396, 395, 396 fruit  398, 399 leaves  395, 396, 397, 577 pseudostems  396, 397, 398 transmission 404–406, 405, 406 banana virus X (BVX)  412–413, 581, 586 banana weevils  303, 304, 431 banana wilt associated phytoplasma (BWAP)  343–345, 344, 345, 346 BanMMV (banana mild mosaic virus)  411–412, 411, 586 bats, as vectors  305 BBrMV (banana bract mosaic virus)  381–383, 383, 580, 584 see also bract mosaic disease BBTV (banana bunchy top virus)  362, 367–369, 367 detection  369, 579, 583–584 and safe transfer of germplasm  578–580, 584 transmission 370–371 see also bunchy top beer 17 benomyl  96, 98, 125, 156 benzimidazoles black leaf streak  96, 98, 101–102 Cladosporium leaf speckle  156 fruit damage  529, 529 resistance  100, 126, 279 Sigatoka leaf spot  125 Bermuda grass  141–142



General Index

biofertilizers  110–111, 114 biological control black leaf streak  107–111, 109, 114 crown rot  279 Cylindrocladium root rot  232 Deightoniella leaf spot  141 eumusae leaf spot  132 nematodes  436, 437, 441, 448 Sigatoka leaf spot  126–127 soft rot  336 Xanthomonas wilt  313 biopriming  375, 436 Bipolaris sacchari 142 birds, as vectors  305 bitertanol 101 black banana aphid (Pentalonia nigronervosa)  370–371, 370, 377, 383, 388–389, 389 black cross leaf spot  132–135, 132, 134 black-end (stem-end rot)  285–286, 285 black heart disease  262 black leaf spot (bacterial)  339 black leaf spot (Deightoniella leaf spot)  138–141, 139 black leaf streak (black Sigatoka)  42 causal agent  56–64 diagnosis 59 genetics  59–63, 64, 85–86 growth and development  57–59, 58, 65 mating types  59, 60 pathogenicity  63–64, 84–86 taxonomy 56–57 compared with Sigatoka leaf spot  53, 55, 73–74 control 88–89 biological 107–111, 109, 114 chemical 95–107 costs 51–53 cultural 89–95, 89, 92, 93, 94, 95, 114–115 on organic farms  113–115 resistant cultivars  80–82, 87–88, 112–113 disease cycle  65 disease development time  68, 75 histology 83–84 incubation period  66–67, 75 infection 64–66 spore formation and dispersal  68–73 symptom evolution time  67–68, 69 distribution and spread  42–48, 60–62, 73–74, 573–574 economic impact  48–53 host reaction  74 evaluation methods  74–78 host–pathogen interactions  83–88 resistance/susceptibility  55, 63–64, 78–79, 79, 84, 86–88 results from the field  80–83 symptoms  42, 53–56, 54, 55, 56, 83

601

evolution time  67–68, 69 fruit  52 black leaf streak of enset  449, 449, 450 black-tip 261–262, 261 bleach (hypochlorite)  308, 323, 447, 529, 577 in irrigation water  535, 536 blood bacterial wilt  323–328, 324, 325 ‘blue’ (magnesium deficiency)  480, 480 Bordeaux mixture  95, 114, 125 boron deficiency 484, 484 excess 540–541, 540 boscalid 101 Botryodiplodia finger rot  286 botrytis tip-end rot  263 bract mosaic disease  378–384 causal agent  381–383, 383, 580, 584 control  384, 584 distribution  378, 378–379 economic impact  378 host susceptibility  384 symptoms 379–381 bracts  379 fruit  382, 383 leaves  379, 381 peepers  382 petiole bases  381 pseudostems  380 transmission 383–384 bracts  1, 3 Brazil  22, 50, 112 break neck  466 brewing 17 brown spot  256–257, 257 BSV (banana streak virus)  396–403, 400 detection  403, 585 endogenous sequences (eBSV)  400, 402, 403, 404, 407, 585 phylogeny 399, 401 and safe movement of germplasm  580, 581, 585 see also banana streak disease bugtok 314–315, 317, 318, 321, 323 see also Moko bacterial wilt bulla 37 bunches 4 bunchy top  362–376 on abacá  362, 363, 366–367, 369, 376–378, 377, 584 causal agents  362 ABTV  363, 369, 377, 584 BBTV 367–369, 367, 579, 583–584 control  374–376, 378, 584 disease cycle  369–372, 377 distribution and spread  362–364, 576 economic impact  362–363 host susceptibility  372–374, 375

602

General Index

bunchy top (contiuned ) and pseudostem heart rot  239 and safe transfer of germplasm  578–580, 584 symptoms 365–367, 365, 366, 372, 376–377 Burkholderia spp. (finger-tip rot)  340–341 burrowing nematode (Radopholus similis) 429–438 control 434–438, 435 disease cycle  432–433 economic impact  429–430 and fungal root rots  234, 433 host susceptibility  433–434 pathogen characteristics  431–432, 432 symptoms 430–431, 430, 431, 432 BVX (banana virus X)  412–413, 586 BWAP (banana wilt associated phytoplasma)  343–345, 344, 345, 346

calcium deficiency  463, 478–479, 479 excess 539, 539 Californian worm compost  94 Calonectria spp. (Cylindrocladium root rot)  229–232, 433 Cameroon 63 Candidatus Phytoplasma asteris  342, 343 Candidatus Phytoplasma cocosnigeriae  346 Canna indica  373, 393 carbendazim 529 Caribbean region  7, 13, 568 black leaf streak  43–44, 47, 50, 52–53, 62–63 Fusarium wilt  210 nematodes 446 see also individual countries carotenoids 32 castor oil  111 Central America see Costa Rica; Honduras; Panama Cephalosporium inflorescence spot of enset  241 Ceratobasidium sp.  235 Ceratocystis corm rot  239–240, 240 Cercospora hayi brown spot  256–257, 257 diamond fruit spot  267 Chaetothyrina musarum (sooty blotch)  271, 271 chemical control black leaf streak  95–96 application methods  96–98, 106–107 choice of fungicide  96, 98–102 costs 51–53 environmental impact  105–107 plant extracts  111, 114 resistance  97, 98–100, 101, 104–105 timing of application  102–104 Cladosporium leaf speckle  156 Cordana leaf spot  138 Cylindrocladium root rot  232

damage to leaves/fruit caused by fungicides/insecticides 527–529, 528, 529 herbicides 530–536, 532, 533, 534, 535 eumusae leaf spot  131–132 freckle 171 fruit diseases postharvest  272, 277–279, 284, 529 preharvest  261, 270 Fusarium wilt  226 health and safety issues  106, 278, 278 insecticides  375, 390, 528 Malayan leaf spot  144 Marasmiellus rot  237 Mycosphaerella leaf speckle  159 nematodes burrowing nematode  436, 436–437 root-knot nematodes  447–448 root-lesion nematode  441 Sigatoka leaf spot  125–126 Taiwan leaf speckle  161 ‘chicken feet’  513 chilling injury  498, 499 postharvest 515–516, 515 chimeras 558–559, 558 China bacterial diseases  334, 336, 338 black leaf streak  45–46 Fusarium wilt  207–208, 572 phytoplasma diseases  342 chitinases 108–110 chlorine as a disinfectant  308, 323, 447, 529, 577 in irrigation water  225, 535, 536 chlorothalonil  98, 105, 106, 528 choking/choke throat  494–495, 494, 495, 496 cigar-end rot  257–261, 260 Cladosporium cladosporioides (sooty mould)  271 Cladosporium leaf speckle  150–156, 151, 152 classification 12–29 see also taxonomy climate (effect on crop) see weather damage climate (effect on disease) black leaf streak  64–65, 66, 67, 71, 74, 113–114 and fungicide applications  102–103 cigar-end rot  259–260 Cladosporium leaf speckle  154, 155 Cordana leaf spot  138 Fusarium wilt  217 Malayan leaf spot  143 maturity stain  468 Mycosphaerella leaf speckle  159 nematodes  433, 446 Sigatoka leaf spot  116, 121 Taiwan leaf speckle  160 Xanthomonas wilt  302, 304 climate change  74, 516–517



General Index

clove (Sumatra disease of  )  326 CMV (cucumber mosaic virus)  385–387, 386, 578, 584–585 ‘Cocos nucifera’ lethal yellowing phytoplasma  344, 345–346 Colletotrichum gloeosporioides 282 Colletotrichum musae anthracnose postharvest  51, 280–285, 280 preharvest 255–256, 256 other postharvest diseases  273, 286 Colletotrichum scovillei 282 Colocasia esculenta 373 Colombia  47, 49, 342–343, 483 colony morphology anthracnose 281–282 black leaf streak  58–59, 58 Cladosporium leaf speckle  152, 155 freckle  168, 169 Fusarium wilt  219–220 Mycosphaerella leaf speckle  157 Phaeoseptoria leaf spot  146 Sigatoka leaf spot  120, 120 Taiwan leaf speckle  160 tropical leaf speckle  163, 164, 165 Xanthomonas bacterial wilt  300, 307 Commelina diffusa  387, 389 compost/compost teas  94, 110, 114 conidiophores/conidia anthracnose 281, 281 black leaf streak  57, 68–69, 72–73 Ceratocystis corm rot  240 cigar-end rot  258–259, 259, 260 Cladosporium leaf speckle  152–153, 153, 154 Cordana leaf spot  136–137, 137 Cylindrocladium root rot  230, 231 Deightoniella leaf spot  139–140, 140 eumusae leaf spot  129 freckle  168, 169 Fusarium wilt  220 Malayan leaf spot  143, 144 Mycosphaerella leaf speckle  157 Pestalotiopsis leaf spot  145 Pyricularia leaf spot  148 Sigatoka leaf spot  118, 119, 120, 121 squirter 287 Taiwan leaf speckle  160, 161 tropical leaf speckle  163, 164, 164, 165, 165 cooking bananas  3, 17 copper deficiency 484–485 excess 541 copper-based fungicides  95, 114, 125 Cordana leaf spot  55, 133, 135–138, 135, 136 corms (rhizomes)  3 Armillaria rot  228–229 banana weevil  431

603

burrowing nematode  430, 430, 432 Ceratocystis rot  239–240, 240 Cylindrocladium root rot  229–232, 433 high mat  462–463, 463 injection of fungicides  97 rhizome rot and tip-over  330–334, 331, 332 rhizome rot/head rot/snap-off   330–334, 331, 332 Rosellinia rot  232–233 Sclerotium rot  240–241, 241, 242 soft rot  334–336 Xanthomonas bacterial wilt control  310–311 yellow mat  465 see also Fusarium wilt Corynespora torulosa black-tip  261, 262 damping-off of Musa seedlings  233 Deightoniella leaf spot  138–41, 139 Deightoniella pseudostem rot of abacá  233–234 fruit speckle  264–265, 264 manganese deficiency  481 Costa Rica  49, 52, 74, 106 Côte d’Ivoire  430 cover crops to improve soil health  91–92, 226 to reduce fungicide drift  107, 107 creamy pulp  52, 90 crown rot  272–280 and black leaf streak  51 causal agents  273–274 control  275–280, 529 disease cycle  274–275 host susceptibility  275 symptoms 272–273, 273 Cuba  47, 50, 52, 97, 112 cucumber mosaic virus (CMV)  385–387, 386, 578, 584–585 cultivation abacá 33–34 banana 5–9 enset 34–36 cultural control abacá mosaic  393 Armillaria corm rot  228–229 banana mosaic  390 black leaf streak  89–95, 89, 92, 93, 94, 95, 114–115 bract mosaic  384 bunchy top  374–375 cigar-end rot  261 Deightoniella  141, 233, 234 freckle 171 frost damage  501 fruit diseases peduncle rot  268 speckle 265 tip-end rots  262

604

General Index

cultural control (continued ) Fusarium wilt  225–226, 225 Marasmiellus pseudostem and root rot  237 Moko bacterial wilt  322–323 nematodes  436, 447, 449 pseudostem wet rot  338 rhizome rot and tip-over  333, 334 Rosellinia root and corm rot  232 Sigatoka leaf spot  126 wind damage  492 Xanthomonas bacterial wilt  308–311 culture of fungal colonies see colony morphology Cylindrocladium root rot  229–232, 433 cytochimeras 558–559

dalapon 531 damping-off of Musa seedlings  233 dark centre of fruit  466, 466 de-budding of the male flower  268, 308, 308, 323 de-handing  114, 272, 275–276 de-leafing 89–91, 89, 92 leaves left on the ground  70, 91 see also tools, contaminated de-suckering agents  535–536 deformed-lamina leaves  555, 556 Deightoniella damping-off of Musa seedlings  233 Deightoniella fruit diseases black-tip  261, 262 speckle 264–265, 264 Deightoniella leaf spot  138–141, 139 Deightoniella pseudostem rot of abacá  233–234 Desarmillaria tabescens (corm rot)  228 dessert bananas  3 diamond fruit spot  266–267, 266, 267 diamond leaf spot (Malayan leaf spot)  142–144, 143 Dickeya spp. heart rot of abacá  339 pseudostem wet rot  338 rhizome rot  333 soft rot  335 diethofencarb 98–100 difenoconazole  106, 126, 132 disease development time  68, 75 disinfection of contaminated tools etc.  226, 308–309, 309, 323, 529 contamination of irrigation water  535, 536 of fruit  529 of planting material  435–436, 447, 577 dithiocarbamates  98, 161, 270 diuron  531, 533, 534 domestication of the banana  9–12, 29–32 Dominican Republic  112, 113 Dothiorella tip-end rot  262–263 drainage systems  91, 94, 513 Drechslera leaf spot (eyespot)  141–142, 141, 142

drooping-leaf variant  554, 554 drought  17, 507–512, 510, 511, 512 dwarf variants  15, 547–551, 549, 550, 551

East Africa banana production  2, 17, 30 black leaf streak  48, 48–49 Cladosporium leaf speckle  150–151, 152, 155, 156 Fusarium wilt  209 nematodes  440, 441 Xanthomonas bacterial wilt  296–297, 304, 306, 575 see also Ethiopia; Mozambique; Uganda economics abacá 32–33 banana  2–3, 223 enset 34 impact of disease abacá mosaic  391 black leaf streak  48–53 bract mosaic  378 bunchy top  362–363 burrowing nematode  429–430 Fusarium wilt  207–208, 218–219, 568 Moko bacterial wilt  314 Sigatoka leaf spot  116–117 Xanthomonas bacterial wilt  296–297 organic farming  113 Ecuador  113, 116 elephantiasis 342–343, 343 enset (Ensete ventricosum)  1, 2, 34–37, 34 bacterial corm rot  330–331, 332, 332 bacterial leaf sheath rot  336–337, 337 black leaf streak (fungal)  80 black leaf streak (nematode)  449, 449, 450 bunchy top  367 Cephalosporium inflorescence spot  241 Cladosporium leaf speckle  150, 151, 152, 154 Deightoniella leaf spot  139, 139 drought tolerance  511 enset streak  409–410, 409 eyespot  141, 142, 142 Pyricularia leaf spot  149–150, 149, 150 root-knot nematode  445, 446 root-lesion nematodes  439, 440 Sclerotium rot  241, 241, 242 Xanthomonas bacterial wilt  296, 297, 304, 575 cultivar susceptibility  305–306 symptoms 299, 302, 303 Ensete glaucum  138, 393 Enterbacter cowanii 341 environmental impact of chemical control agents  105–107, 530, 535 erect leaf variant  555 Erwinia spp.



General Index

pseudostem wet rot  337–338 rhizome rot  332–333 soft rot  335 Ethiopia Cladosporium leaf speckle  150, 156 enset 34–37 Xanthomonas bacterial wilt  296, 575 ethylene  8, 283 ethylene bisdithiocarbamates (EBDCs) see mancozeb ethylenethiourea (ETU)  106 eumusae leaf spot causal agent  128–131, 130 compared with Sigatoka leaf spot  115–116 control 131–132 distribution 127 host susceptibility  131 symptoms 127–128, 128 evolution of the banana  9–12, 29–32 export bananas  7–9, 49, 207, 223, 272, 276–277 eyespot 141–142, 141, 142

fallow periods  311, 322, 434–435, 447 fenbuconazole 101 fenpropidin 100 fenpropimorph 100 feral banana plants  375 fermentation products (biocontrol agents)  110–111 fertilizers and black leaf streak control  94, 110, 114 boron 484 calcium 479 copper 484–485 iron 482 magnesium 480–481 manganese 482 nitrogen 477 excess  470, 527 phosphorus 478 potassium 478 sulfur 481 zinc 484 fibre (abacá)  33 Fiji  42, 142, 362 finger rot Botryodiplodia 286 Trachysphaera 288 finger-tip rot  339–341, 340 fingers 3 malformed 467–468, 468 physiological finger drop  469–470 flag leaf (abacá)  33 flooding  492, 512–514, 513 as a control method  435 prevention  514 flowers  1, 3

605

blood bacterial wilt  325, 327 cold damage  495–497, 497, 498 insecticide damage  528 Moko bacterial wilt  315, 319–320, 321, 323 morphologically resistant forms  312, 321, 327 removal of the male flower  268, 308, 308, 323 variant forms  559, 559, 560, 562 Xanthomonas bacterial wilt  297, 298, 298, 299, 308, 312 see also bract mosaic fluorine 541–542, 541 flusilazole 126 flutriafol 97 foliar disorders see leaf disorders foliar substrates  110 followers 4 see also suckers food additives (on harvested fruit)  280, 285 food products banana  3, 17 enset 36–37, 36 forecasting systems for fungicide application  102–104, 125, 126 freckle eradication measures in Australia  169, 574–575 fruit  166, 267–268, 268 leaf  166–171, 167 frost damage  498–502, 500 enset 35 fruit abacá 33 bagging 7–8, 9, 261 postharvest  276, 277 problems caused by  265, 467, 468, 472, 507 enset 35 growth and development  4, 493 orange pulp varieties  31, 32 ripening and transport  8–9, 272, 276–277 as a source of disease  570 fruit disorders agrochemical damage diuron  534 thiophanate-methyl  529 bacterial infections blood bacterial wilt  324, 325 finger-tip rot  339–341, 340 Moko bacterial wilt/bugtok  315–316, 316, 317 Xanthomonas bacterial wilt  298, 299 non-infectious alligator skin  465–466, 465 break neck  466 dark centre  466, 466 finger drop  469–470

606

General Index

fruit disorders (continued ) hard lump  466–467 malformed fingers and hands  467–468, 467, 468 maturity stain  468–469, 468 mixed ripeness  469 neer vazhai  469, 469 premature field ripening  470 sinkers 470–471 split peel  471, 471, 559 uneven de-greening  471–472, 471 withered pedicels  472 yellow pulp  472 postharvest fungal infections  271–272 anthracnose  51, 280–285, 280 Botryodiplodia finger rot  286 cigar-end rot  260 crown rot  51, 272–280, 273, 529 fungal scald  285 main-stalk rot  286 ring rot  287, 287 soft rot  288 squirter 286–287, 287 stem-end rot  285–286, 285 Trachysphaera finger rot  288 preharvest fungal infections  255 anthracnose 255–256, 256 brown spot  256–257, 257 cigar-end rot  257–261, 260 creamy pulp  52, 90 diamond fruit spot  266–267, 266, 267 freckle  166, 267–268, 268 peduncle rot  268, 269 pitting 269–270, 270 premature ripening  48, 52, 90, 116, 298, 470 sooty mould/sooty blotch  270–271, 271 speckle 264–266, 264, 265 tip-end rots  261–263, 261, 263 variant forms  559–561, 560, 561, 562 variegated chimeras  556, 558, 558 viral infections banana mosaic  385, 385 banana streak  398, 399 bract mosaic  380, 382, 383 weather damage cold (postharvest)  515–516, 515 cold (preharvest)  497–498, 497 drought  507–508, 509–510, 510 frost 500–501, 500 hail 502–503, 503 heat 505–506, 506 sunburn 507, 507, 508, 509 wind  488 fruit flies  303 fungal scald  285 fungi as biocontrol agents  110, 127, 141, 232, 437, 448

fungicides black leaf streak  95–96 application methods  96–98, 106–107 choice of fungicide  96, 98–102 costs 51–53 environmental impact  105–107 plant extracts  111, 114 resistance  97, 98–100, 101, 104–105 timing of application  102–104 Cladosporium leaf speckle  156 Cordana leaf spot  138 damage to leaves/fruit caused by  527–528, 528, 529, 529 eumusae leaf spot  131–132 freckle 171 fruit diseases postharvest  272, 277–279, 284, 529 preharvest  261, 270 health and safety issues  106, 278, 278 Malayan leaf spot  144 Marasmiellus rot  237 Mycosphaerella leaf speckle  159 Sigatoka leaf spot  125–126 Taiwan leaf speckle  161 fusaric acid  222 Fusarium spp.  268, 273, 288 Fusarium incarnatum  267, 268, 274, 285 Fusarium musae 273 Fusarium oxysporum f. sp. cubense 213 in culture  219–220 diagnosis 223–224 genetics 220–221 host range  213–215 pathogenicity 221–222 races 215 vegetative compatibility groups  215–219, 220 see also Fusarium wilt Fusarium solani  234, 267, 268 Fusarium verticillioides (F. moniliforme) 239, 261–262, 274, 286 Fusarium wilt  207–228 causal agent see Fusarium oxysporum f. sp. cubense compared with bacterial wilts  213, 298–299 control in Australia  224, 572 cultural 225–226, 225 disinfectants 226 resistant cultivars  226–228, 550 distribution and spread  208, 209–212, 222–223 TR4  219, 571–572 economic impact  207–208, 218–219, 568 pathophysiology 221–222 post-flooding  492, 492 suppressive soils  222



General Index

symptoms  208, 213 devastation  217 leaves  214, 215, 216 pseudostem  213, 215, 217 TR4 outbreaks  207–208, 215, 218–219, 223–224, 571–572

genetically engineered crops resistance to bunchy top  375–376 resistance to Fusarium wilt  228 resistance to Xanthomonas bacterial wilt  312–313 genetics black leaf streak resistance  87–88 fungi Colletotrichum spp.  282 Fusarium oxysporum 220–221 Pseudocercospora eumusae 129 Pseudocercospora fijiensis  59–63, 64, 85–86 Pseudocercospora musae  120, 129 hybridization of Musa species/subspecies  9–10, 29–30 ploidy of Musa spp.  10–12, 553–554, 558–559 Sigatoka leaf spot resistance  125 variant forms of banana  546–547, 553–554, 561–562 chimeras 558–559, 558 dwarfs 547–551, 549, 550, 551 flowers 559, 559, 560, 562 foliage 551–556, 554, 555, 556, 557 fruit  556, 558, 559–561, 560, 561, 562 giants 551, 552, 553 pseudostem pigmentation  556–558, 557, 558 viruses ABTV 377 ampeloviruses 413 BanMMV 411 BBrMV 381–382 BBTV 367–368 BSV  396–398, 401, 402 CMV 386 Xanthomonas bacterial wilt pathogen  300 genomic groups AA/AB 13 AAA 13–19 AAB 19–25 ABB 25–27 BBB 27–29 S/T components  27 and susceptibility to anthracnose 284 bacterial rhizome rot  333 bacterial soft rot  335 BBTV 373–374 black cross leaf spot  134 black leaf streak  80–82, 112 blood bacterial wilt  327

607

brown spot  257 Cladosporium leaf speckle  155–156 crown rot  275 Deightoniella leaf spot  140–141 drought 510–511 eumusae leaf spot  131 freckle 171 frost damage  501 Fusarium wilt  215, 226 Moko bacterial wilt  320–321 Mycosphaerella leaf speckle  159 nematodes  434, 441, 443, 447 Phaeoseptoria leaf spot  147 pseudostem wet rot  338 rust 172 Sigatoka leaf spot  123–124 Taiwan leaf speckle  160–161 giant variants of Cavendish cultivars  15, 551, 552, 553 gibberellic acid  548–549 global warming  74, 516–517 Gloeosporium musarum see Colletotrichum musae Glomerella cingulata (tip-end rot)  263, 263 glufosinate  531, 532, 533 glycosylphosphatidylinositol (GPI) protein family  86 glyphosate  310, 530–532, 532 greenhouses 488–490, 490, 501–502, 502 ground spraying of chemicals  107, 530 Guignardia spp. (freckle)  168 gumming 340

hail damage  502–503, 503 handling and transport of fruit  8–9 chilling injury  515–516, 515 dark centre  466, 466 fungal scald  285 modified atmosphere  272, 276–277, 285 hands (fruit)  3 malformed 467–468, 467 Haplobasidion musae (Malayan leaf spot)  142–144, 143 hard lump  466–467 harvesting abacá 33 banana 8 enset 36 see also postharvest diseases Hawaii  22, 171, 571 head rot (rhizome rot and tip-over)  330–334, 331, 332 health and safety fungicides  106, 278, 278 herbicides 310 heart leaf unfurling disorder of plantain  462 heart rot of abacá (bacterial)  339 banana mosaic  384 boron deficiency  484 fungal 238–239, 238

608

General Index

Heliconia spp.  214, 322, 571 Helicotylenchus spp. (spiral nematodes)  442–443, 442 herbicides  310, 530 damage caused by  530–535, 532, 533, 534, 535 Heterodera oryzicola (nematode)  449 high mat  462–463, 463 Honduras  46, 49, 266, 330, 464 Hoplolaimus pararobustus (nematode)  449 hot-water treatment fruit  279, 284–285 suckers/corms  435–436, 441, 447 humidity and black leaf streak  91 postharvest storage  277 hybridization events  10, 29–30 hypersensitive response to black leaf streak  84

imazalil  126, 278 imidacloprid  375, 528 indexing of viruses  408, 586–587 India bacterial diseases  330, 334, 336 Fusarium wilt  572 phytoplasma diseases  342 Sclerotium rot  241 Sigatoka leaf spot  127, 131 viral diseases  378, 394 Indonesia  19, 46, 324, 329 inflorescence see flowers inflorescence spot of enset  241 InPAct technique  376 insect transmission vectors abacá mosaic  392–393 banana mosaic  387–389, 389, 390 banana streak  404–406, 406 blood bacterial wilt  327 bract mosaic  383 bunchy top  370–371, 370, 375, 377 Moko bacterial wilt  315, 319, 320 phytoplasmas 342 Xanthomonas bacterial wilt  303–304 insecticides  375, 390 damage to flowers/fruit  528 intercropping black leaf streak control  90 nematode control  447, 449 as source of CMV  387, 388, 390 International Musa Testing Program (IMTP)  76 International Network for the Improvement of Banana and Plantain (INIBAP)  76, 578, 579, 580, 582 iron and anthracnose infection  283 deficiency 482, 482, 555, 557 excess 539–540, 540

irradiation of fruit  279 irrigation  7, 17, 508–509 boron 540 chlorine treatment  225, 535, 536 drip or under-canopy systems better  91, 114–115, 334 enset 35 salinity 537 Israel  262, 263 ITC (INIBAP Transit Centre)  578, 579, 580, 582

Javanese vascular disease  329 juglone 85

kerosene 535–536 Klebsiella pneumoniae (leaf spot)  339 kocho 36–37 kokkan  379

LAMP assays  585 Lasiodiplodia theobromae (Botryodiplodia theobromae)  268, 273–274, 286 latent infections anthracnose 282–283 crown rot  274 Xanthomonas bacterial wilt  297, 310 Latin America  568 bacterial diseases  330, 334, 336 black leaf streak  43–44, 46–47, 49–51, 52, 62 fruit diseases  256, 265, 266, 269 Fusarium wilt  210 nematodes 444 phytoplasma diseases  342–343 Sigatoka leaf spot  116 see also individual countries leaf disorders agrochemical damage disinfectants 535, 536 excess fertilizer  527 herbicides 530–535, 532, 533, 534, 535 mineral oils and fungicides  528, 528 bacterial infections abacá wilt  328 blood bacterial wilt  324, 324 leaf blight  339 leaf spot  339 Moko bacterial wilt  315, 315, 316 Xanthomonas bacterial wilt  298–299, 299, 300, 301 fungal infections black cross leaf spot  132–135, 132, 134 black leaf streak see black leaf streak Cladosporium leaf speckle  150–156, 151, 152



General Index

Cordana leaf spot  55, 133, 135–138, 135, 136 Deightoniella leaf spot  138–141, 139 eumusae leaf spot  127–132, 130 eyespot 141–142, 141, 142 freckle 166–171, 167, 574–575 Fusarium wilt  214, 215, 216, 222 Malayan leaf spot  142–144, 143 Mycosphaerella leaf speckle  156–159, 157, 158 Pestalotiopsis leaf spot  144–145 Phaeoseptoria leaf spot  145–147, 146 Pyricularia leaf spot  148–150, 148, 149, 150 rust 171–172, 171 Sigatoka leaf spot see Sigatoka leaf spot Taiwan leaf speckle  159–161, 160 tropical leaf speckle  161–166, 162, 163 nematode black leaf streak of enset  449, 449, 450 non-infectious heart leaf unfurling disorder  462 leaf-edge chlorosis  463, 463, 479 Roxana disease  463–464 yellow mat  464 see also mineral deficiency; mineral excess genetic variants deformed lamina  555, 556 drooping/erect 554, 554, 555 mosaic 551–553, 554 other 555–556 variegation 554–555, 555 viral infections abacá mosaic  391, 391, 392 banana mild mosaic  410–412, 410, 411, 586 banana mosaic  384–385, 385, 386 banana streak  394–395, 395, 396, 397, 577 bract mosaic  379, 380, 381 bunchy top  365, 365, 366, 366 enset streak  409–410, 409 weather damage cold  494, 498, 499 drought 509, 510, 511–512 flooding  492, 513, 513 frost 499, 500 hail 502, 503 heat 503–504, 504, 505, 506 lightning strike  514, 515 wind 487, 488 leaf-edge chlorosis  463, 463, 479 leaves  1, 3 abacá 33 photosynthesis  116, 217 as a source of disease  570

temperatures required for growth  493 unfurling  76 lightning strike  514–515, 515 lime (calcium)  479

macropropagation abacá 33 banana  4, 311, 435–436, 435 enset 36 magnesium deficiency  479–481, 480 main-stalk rot  286 Malayan leaf spot  142–144, 143 Malaysia  44, 46 mancozeb environmental impact  105–6 leaf diseases  98, 105, 125, 132, 159, 171 manganese and anthracnose  283–284 deficiency 481–482 excess  106, 539 Manila hemp see abacá manure 94 Marasmiellus pseudostem and root rot  235–237, 236, 237 maturity stain on fruit  468–469, 468 ‘May flowering’ (‘November dump’)  340, 466, 495–497, 497, 498 mealybug vectors of BSV  404, 406 melanin 85 Meloidogyne spp. (root-knot nematodes)  443–448, 445 meristem tip culture  390, 407, 583 methyl bromide soil fumigation  229, 322, 435 Metulocladosporiella spp. (Cladosporium leaf speckle) 150–156, 151, 152 Mexico  50, 116 micropropagation see tissue culture material mineral deficiency  475 and black leaf streak  95 boron 484, 484 calcium  463, 478–479, 479 copper 484–485 iron 482, 482, 555, 557 magnesium 479–481, 480 manganese 481–482 nitrogen 475–477, 477 phosphorus 477–478, 477 potassium  94, 465, 471, 478, 478 sulfur 481, 481 zinc 482–484, 483 mineral excess aluminium 541 arsenic 541 boron 540–541, 540 calcium 539, 539 copper 541

609

610

General Index

mineral excess (continued ) fluorine 541–542, 541 iron 539–540, 540 manganese  106, 539 sodium chloride (salinity)  527, 536–539, 537, 538 mineral oil black leaf streak  95, 102, 111, 114 damage to leaves/fruit  527–528, 528 Malayan leaf spot  144 Sigatoka leaf spot  125, 126 modelling of diseases  313–314 modified atmosphere storage and transport  272, 276–277, 285 Mokillo (bacterial finger-tip rot)  339–341, 340 Moko bacterial wilt  314–323 causal agent  317–319 compared with bugtok  314–315 control  321–323, 571 disease cycle  319–320 host susceptibility  320–321 symptoms 315–316, 315, 316, 317 morpholine fungicides  100 morphology abacá 33 banana  1, 3–5, 76 enset  34, 35 mosaic diseases see banana mild mosaic; banana mosaic; bract mosaic mosaic genetic variants  551–553, 554 Mozambique 571 multi-bunch genetic variants  559–560, 560 Musaceae family  1–2 Musicillium theobromae black-tip 261 cigar-end rot  257–258, 258, 259, 259, 260–261 mutant forms  546–562 BBTV resistance  375 chimeras 558–559, 558 dwarfs 547–551, 549, 550, 551 flowers 559, 559, 560, 562 foliage 551–556, 554, 555, 556, 557 fruit  556, 558, 559–561, 560, 561, 562 giants 551, 552, 553 pseudostem pigmentation  556–558, 557, 558 somaclonal variation  546–547 Mycosphaerella spp.  158 M. fijiensis see Pseudocercospora fijiensis M. musae 156–169, 157, 158 M. musicola (Pseudocercospora musae)  118–120, 120 Mycosphaerella leaf speckle  156–159, 157, 158

neem 437 neer vazhai  469, 469

nematicides burrowing nematode  436, 436–437 on planting material  436, 447 root-knot nematodes  447–448 root-lesion nematode  441 nematodes 429 black leaf streak of enset  449, 449, 450 burrowing 429–438, 430, 431, 432 and fungal root rots  234, 433, 446 Heterodera oryzicola 449 Hoplolaimus pararobustus 449 Paratylenchus spp.  449 root-knot 443–448, 445 root-lesion 438–442, 439 Rotylenchulus reniformis 448–449 spiral 442–443, 442 Neocordana spp. (Cordana leaf spot)  55, 133, 135–138, 135, 136 Neofusicoccum ribis (Dothiorella tip-end rot)  262 Neoscytalidium dimidiatum (Dothiorella tip-end rot)  262–263 Nigeria  127, 430 Nigrospora spp. (squirter)  287 nitrogen and black leaf streak  94 deficiency 475–477, 477 excess  470, 527 nomenclature fungi  xv, 41 Musa  1–2, 13, 32 see also classification; taxonomy ‘November dump’ (‘May flowering’)  340, 466, 495–497, 497, 498 nutrition as a cultural control measure  92–95, 114, 126, 302 mineral requirements  475

Oceania  22–23, 568 BBTV  362–363, 364 black cross leaf spot  132 black leaf streak  42, 44–45, 49, 112 Fusarium wilt  212 Malayan leaf spot  142 phytoplasma diseases  343–346 see also Australia; Papua New Guinea off-types see genetics, variant forms oils see mineral oil; organic oils organic farming  113–115, 113, 279 organic oils  111 oxamyl 448 oxyfluorfen  531, 533, 534 oxygen low levels delay ripening  276 root requirements  512–513



General Index

packing procedures  8, 467 and crown rot  275, 278–279 fungicide applications  278–279, 529 Pakistan  363, 572 Panama  49, 464 Panama disease see Fusarium wilt Pantoea agglomerans 341 Papua New Guinea  5, 6, 13, 31 black leaf streak  45 freckle 171 leaf spot diseases  134–135, 138, 142 phytoplasma diseases  343–345 paraquat  530, 531, 532 Paratylenchus spp. (nematodes)  449 parthenocarpy  9, 10 Pasteuria spp.  437, 448 PCR assays see polymerase chain reaction (PCR) assays pedicel withering  472 peduncle 3 main-stalk rot  286 peduncle rot  268, 269 peepers 4 bract mosaic  382 Pentalonia nigronervosa (black banana aphid)  370–371, 370, 377, 383, 388–389, 389 perithecia black cross leaf spot  132 Mycosphaerella leaf speckle  157 see also pseudothecia Peru  47, 113–114, 329 Pestalotiopsis leaf spot  144–145 Phaeoseptoria leaf spot  145–147, 146 phenylphenalenones 86–87 Philippines abacá  32, 33, 328, 376, 391 black leaf streak  45 bugtok 314 nematodes 444 typhoons 491–492 viral diseases  376, 378, 391 phosphorus deficiency  477–478, 477 photosynthesis  116, 217 Phyllachora musicola (black cross leaf spot)  132–135, 132, 134 Phyllosticta spp. (freckle) eradication measures in Australia  169, 574–575 on fruit  166, 267–268, 268 on leaves  166–171, 167 P. capitalensis 169 P. cavendishii  168–169, 170, 171 P. maculata  168, 170, 171 P. musaechinensis 169 P. musarum 169 phytoplasma diseases  342–346 banana wilt associated phytoplasma  343–345, 344, 345, 346 elephantiasis 342–343, 343

phytoprotectants  86–87, 283 phytosanitary measures  582–583 phytotoxins  84–85, 222 pitting of fruit  269–270, 270 Planococcus spp. (mealybugs)  404, 406 plant density  91, 95, 234, 467, 501, 507 plant extracts as biocontrol agents for fungi  111, 114, 132, 285 for nematodes  437, 448 as disinfectants  308–309 plantations  7, 9 planting material, as a source of disease  304, 569–570, 576, 577 polyketide synthase (PKS) genes  86 polymerase chain reaction (PCR) assays ABTV 584 BanMMV  412, 586 BBrMV  383, 584 BBTV  369, 584 BSV  403, 585 CMV 387 Fusarium wilt  224 Xanthomonas bacterial wilt  307 postharvest chilling injury  515–516, 515 postharvest fungal diseases  271–272 anthracnose  51, 280–285, 280 Botryodiplodia finger rot  286 cigar-end rot  260 crown rot  51, 272–280, 273, 529 fungal scald  285 main-stalk rot  286 ring rot  287, 287 soft rot  288 squirter 286–287, 287 stem-end rot  285–286, 285 Trachysphaera finger rot  288 postharvest treatments  275, 278, 279, 284, 529 potassium deficiency  94, 465, 471, 478, 478 pralinage 436 Pratylenchus spp. (root-lesion nematodes)  438–442, 439 preharvest diseases of fruit  255 anthracnose fruit rot  255–256, 256 brown spot  256–257, 257 cigar-end rot  257–261, 260 creamy pulp  52, 90 diamond fruit spot  266–267, 266, 267 freckle  166, 267–268, 268 peduncle rot  268, 269 pitting 269–270, 270 sooty mould/sooty blotch  270–271, 271 speckle 264–266, 264, 265 tip-end rots  261–263, 261, 263 premature ripening of fruit  48, 52, 90, 116, 298, 470 prochloraz  278, 284

611

612

General Index

propagation see macropropagation; tissue culture material propiconazole anthracnose 284 leaf diseases  100–101, 126, 159, 171 resistance 104 protective clothing  278 pruning 89–91, 89, 92 leaves left on the ground  70, 91 see also tools, contaminated Pseudocercospora eumusae (emusae leaf spot)  115–116, 127–132, 128 Pseudocercospora fijiensis (Mycosphaerella fijiensis)  56–64 diagnosis 59 genetics  59–63, 64, 85–86 growth and development  57–59, 58, 65 mating types  59, 60 pathogenicity  63–64, 84–86 taxonomy 56–57 see also black leaf streak Pseudocercospora longispora 158 Pseudocercospora musae (Mycosphaerella musicola) 118–120, 120 see also Sigatoka leaf spot Pseudomonas spp.  340 pseudostems abacá 33 bacterial infections blood bacterial wilt  325 Moko bacterial wilt  315, 316 rhizome rot/head rot/snap-off   330, 331 soft rot  334–336 wet rot  336–338, 337 Xanthomonas bacterial wilt  301 banana 3 drought-weakened  511 fungal infections damping-off of Musa seedlings  233 Deightoniella rot of abacá  233–234 Fusarium wilt see Fusarium wilt heart rot  238–239, 238 Marasmiellus rot  235–237, 236, 237 Sclerotium rot  240–241, 241, 242 injection of fungicides  97 lightning strike  514, 515 phytoplasma infections  342–343, 343, 345 pigmentation variants  556–558, 557, 558 viral infections banana streak  396, 396, 397, 398 bract mosaic  379, 380 wind tolerance  492 yellow mat  465 pseudothecia black leaf streak  57–8, 69–72 eumusae leaf spot  129

Sigatoka leaf spot  120, 121 see also perithecia pycnidia Dothiorella tip-end rot  262, 263 freckle  167, 168, 169, 170 Phaeoseptoria leaf spot  146, 147 Pyricularia angulata (pitting disease of fruit)  269–270, 270 Pyricularia leaf spot  148–150, 148, 149, 150 pyrimethanil 101 Pythium sp.  235

quarantine 568–569 in Australia  571, 572, 573, 587 for BBTV  579 for black streak in Costa Rica  49 facilities  587 for Fusarium wilt  225 for Heliconia 571 for Xanthomonas bacterial wilt  307–308

Radopholus similis see burrowing nematode rainfall black leaf streak  67, 71, 103, 113–114 drought  17, 507–512, 510, 511, 512 water requirements abacá 33 banana  7, 17, 508 enset 35 Ralstonia solanacearum species complex abacá bacterial wilt  328–329 blood bacterial wilt  325–326 Moko bacterial wilt  317–319, 571 Ralstonia syzygii  318–319, 326 Ramichloridium australiense (Zasmidium musae-banksii) 165 Ramichloridium biverticillatum (Zasmidium biverticillatum) 163–165, 165 Ramichloridium ducassei (Taiwan leaf speckle)  159–161, 160 Ramichloridium musae (Zasmidium musigenum) 163, 164 ratoon crop  5, 48 rayadilla disease  483, 483 reactive oxygen species  85 Rhizoctonia spp. (root rot)  234–235 rhizomes (corms)  3 Armillaria rot  228–229 banana weevil  431 burrowing nematode  430, 430, 432 Ceratocystis rot  239–240, 240 Cylindrocladium root rot  229–232, 433 high mat  462–463, 463 injection of fungicides  97 pathogen spread via  569–570 rhizome rot and tip-over  330–334, 331, 332



General Index

rhizome rot/head rot/snap-off   330–334, 331, 332 Rosellinia rot  232–233 Sclerotium rot  240–241, 241, 242 soft rot  334–336 Xanthomonas bacterial wilt control  310–311 yellow mat  465 see also Fusarium wilt Rhizopus spp.  288 ring rot  287, 287 ripening  8–9, 276 controlled  272, 276–277 mixed 469 premature leaf spot diseases  48, 52, 90, 116 non-infectious 470 Xanthomonas bacterial wilt  298 and temperature  505–506 uneven de-greening  471–472, 471 RNA interference/RNA silencing BBTV  367–368, 376 BSV 403–404 root-knot nematodes (Meloidogyne spp.)  443–448, 445 root-lesion nematodes (Pratylenchus spp.)  438–442, 439 roots aluminium toxicity  541 Armillaria corm rot  228–229 Ceratocystis corm rot  239–240, 240 Cylindrocladium root rot  229–232, 433 damping-off of Musa seedlings  233 fungal root rots  234–235 Fusarium wilt  208, 221 growth and development  4–5, 493 heat damage  504 Marasmiellus root rot  237 nematode pests see nematodes Rosellinia root rot  232–233 Sclerotium corm rot  240–241, 241, 242 sodium toxicity  538–539 waterlogged 512–514 yellow mat  465 Rosellinia root and corm rot  232–233 Rotylenchulus reniformis (nematode)  448–449 Roxana disease  463–464 rust 171–172, 171

safe movement of Musa germplasm  577–587 safety fungicides  106, 278, 278 herbicides 310 salinity, excess  527, 536–539, 537, 538 sanitation measures Armillaria corm rot  228–229 black streak  89–91, 89, 92, 93, 114

613

bract mosaic  384 cigar-end rot  261 Moko bacterial wilt  322–323 Sigatoka leaf spot  126 Xanthomonas bacterial wilt  308–311 see also disinfection Santa Maria stem-end rot  286 scald 285 SCBV (sugarcane bacilliform viruses)  399–400 Sclerotinia fruit rot  263 Sclerotium corm and pseudostem rot  240–241, 241, 242 SCMV-Ab (sugarcane mosaic virus, abacá strain) 392, 392, 584 Scolecobasidium humicola 133 screenhouses 488–90, 491, 501–502, 534 seedlings, damping-off of Musa 233 seeds  1, 14 abacá 33 M. acuminata ssp. banksii 570 as a source of disease  390, 570, 577, 583 serological tests  307 ABTV 584 BanMMV  411–412, 586 BBrMV  383, 584 BBTV  369, 579, 583–584 BSV  403, 585 CMV 387 SCMV/AMV  392, 584 Serratia marcescens  108, 109, 110 shade (effect on banana)  506–507 shade (effect on black leaf streak)  67, 115 Sigatoka leaf spot (yellow Sigatoka)  41 causal agent  118–120, 120 compared with black leaf streak  53, 55, 73–74 compared with eumusae leaf spot  115–116 control 125–127 disease cycle  120–122 distribution 115–116 economic impact  116–117 host susceptibility  121, 122–125 symptoms  83, 117–118, 117, 118 silicon 232 simazine  531, 535 single diseased stem removal (SDSR)  309–310 sinkers 470–471 SIX genes  221 snap-off (rhizome rot and tip-over)  330–334, 331, 332 sodium chloride excess  527, 536–539, 537, 538 sodium hypochlorite see bleach soft rot of fruit  288 soft rot of rhizome and pseudostem  334–336 soil temperature  501, 504–505 soil treatments chemical fumigation  229, 322, 435 fallow periods  311, 322, 434–435, 447

614

General Index

soil treatments (continued ) fungicide applications  97 for nematodes  434–435 sterilization by fire  225, 225 soil types and Cylindrocladium root rot  230 and Fusarium wilt  222, 226 and Rosellinia root rot  232 and Xanthomonas bacterial wilt  302 and yellow mat  465 solar radiation  506–507, 506, 507, 508, 509 Solomon Islands  345–346 somaclonal variation  546–547 see also genetics, variant forms sooty mould/sooty blotch  270–271, 271 South Africa  159, 440, 444 South America see Latin America spacing of plants  91, 95, 234, 467, 501, 507 speckle of fruit  264–266, 264, 265 spermogonia black leaf streak  57, 69 eumusae leaf spot  129 freckle  168, 169 Sigatoka leaf spot  118–120 spiral nematodes (Helicotylenchus spp.)  442–443, 442 spiroxamine 100 split peel  471, 471, 559 sporodochia Fusarium wilt  220 Sigatoka leaf spot  118, 119 spray equipment  96–97, 106–107, 530 spread of pathogens  567–571 banana streak  394, 576–577 black leaf streak  42–48, 60–62, 73–74, 573–574 bunchy top  362–364, 576 freckle 574–575 Fusarium wilt  208, 209–212, 222–223 TR4  219, 571–572 Xanthomonas bacterial wilt  296, 303–305, 575 squirter 286–287, 287 Sri Lanka  378 star-like wound disease  232 stature 15 dwarfs 547–551, 549, 550, 551 giants 551, 552, 553 stem-end rot  285–286, 285 stems 3–4 see also pseudostems storage of fruit see transport of fruit storm damage  490–492, 491 hail 502–503, 503 lightning strike  514–515, 515 Streptomyces galilaeus 110 strobilurin fungicides  101 resistance  104, 126

suckers 4, 7 abacá 33 de-suckering agents  535–536 enset 36 as source of planting material  311–312, 569, 569, 582–583 sugarcane bacilliform viruses (SCBV)  399–400 sugarcane mosaic virus, abacá strain (SCMV-Ab) 392, 392, 584 sulfur deficiency  481, 481 Sumatra disease of clove  326 sunburn  506, 507, 507, 508, 509 symptom evolution time  67–68

Taiwan 218–219 Taiwan leaf speckle  159–161, 160 taxonomy banana streak viruses  399, 401 Cladosporium leaf speckle  152 Cordana leaf spot  135, 137 Cylindrocladium spp.  229–230 freckle 168 fungi xv 41 Fusarium oxysporum 213 Musa spp.  1–2, 12–13, 32 Mycosphaerella spp.  158 Pestalotiopsis spp.  145 Pseudocercospora spp.  56–57, 118, 128 pseudostem heart rot  239 Ralstonia solanacearum  317, 318–319, 325 rust 172 tropical leaf speckle  163 Xanthomonas causing bacterial wilt  299–300 see also classification Technical Guidelines for the Safe Movement of Musa Germplasm  579, 580, 581, 581 temperature growth requirements abacá 33 banana 492–494 enset 35 high-temperature damage  503–506, 504, 505, 506 low-temperature damage  494–498, 494, 495, 496, 497 chilling 498, 499 frost  35, 498–502, 500 postharvest 515–516, 515 during transport  276, 277 see also climate tetraploidy 12 Thailand  26, 46, 127, 514 thiabendazole  277–278, 284 Thielaviopsis paradoxa Ceratocystis corm rot  239–240, 240 fruit diseases (postharvest)  277, 286



General Index

thiophanate-methyl damaged fruit  529, 529 tip-end rots  261–263, 261, 263 tip-over rot  330–334, 331, 332 tissue culture material  5, 578 in the control of banana mosaic  390, 585–585 banana streak  407, 585 Fusarium wilt  225–226 nematodes  435, 436, 447 Xanthomonas bacterial wilt  311 high mat  463, 463 off-types/mutants  546–547, 553–554, 558–559, 561–562 safe movement of pathogen-free germplasm 577–583 sensitivity to herbicides  530, 532 soft rot  330 tools, contaminated  226, 304, 308–309, 309, 323, 529 tooth-comb chlorosis  481 TR4 (tropical race 4, Fusarium wilt)  207–208, 215, 218–219 diagnosis 223–224 resistance  226–228, 550 spread  219, 571–572 Trachysphaera fructigena cigar-end rot  257–258, 258, 259–261, 260 finger rot  288 transgenic crops resistance to bunchy top  375–376 resistance to Fusarium wilt  228 resistance to Xanthomonas bacterial wilt  312–313 transmission vectors insects see insect transmission vectors other 305 transport of fruit  8–9 chilling injury  515–516, 515 in controlled atmosphere  272, 276–277, 285 fungal scald  285 see also postharvest fungal diseases triadimenol 97 triazole fungicides  100–101, 106, 126 see also propiconazole Trichoderma spp.  110, 127, 141 tridemorph  100, 125 damage caused by  528 triploidy 10–12 tropical leaf speckle  161–166, 162, 163 tropical race 4 (TR4 , Fusarium wilt)  207–208, 215, 218–219 diagnosis 223–224 resistance  226–228, 550 spread  219, 571–572 2,4-D (2,4-dichlorophenozyacetic acid)  310, 471 damage caused by  535, 535 2,4,8-THT (2,4,8-trihydroxytetralone)  85

615

Uganda  8, 17, 156, 296, 432, 441, 575, 577 ultraviolet irradiation  279 see also sunburn under-peel discolouration  515–516, 515 urea fertilizer 477 treatment of diseased material  93 Uredo musae (rust)  171–172 Uromyces musae (rust)  171–172

variegation of leaves and fruit  554–555, 555, 556, 558, 558 vascular wilts abacá bacterial wilt  328–329 blood bacterial wilt  323–328, 324, 325 Fusarium see Fusarium wilt Javanese vascular disease  329 Moko bacterial wilt  314–323, 315, 316, 317 Xanthomonas bacterial wilt see Xanthomonas bacterial wilt vegetative compatibility groups (F. oxysporum)  215–219, 220 Venezuela  47, 50 Vietnam  46, 572 virus indexing  586–587 vitamin A  32

water contamination with chlorine  535, 536 with fungicides  106 water requirements abacá 33 banana  7, 17, 508 drip or under-canopy irrigation better  91, 114–115, 334 drought 507–512, 510, 511, 512 enset 35 waterlogging  492, 512–514, 513 prevention  514 waxed fruit  280 weather damage cold 494–498, 494, 495, 496, 497 chilling 498, 499 frost  35, 498–502, 500 drought 507–512, 510, 511, 512 flooding  492, 512–514, 513, 514 hail 502–503, 503 heat 503–506, 504, 505, 506 lightning strike  514–515, 515 solar radiation  506–507, 506, 507, 508, 509 wind 487–492, 488, 489, 491, 493 see also climate weeds and black leaf streak  91–92 and CMV  387, 387, 390 control  91, 323

616

General Index

weeds (continued ) herbicide use  530–536 and Moko bacterial wilt  319, 322, 323 and SCMV  393 West Africa  20 black leaf streak  47–48, 48, 63, 112 Cladosporium leaf speckle  150, 154 Fusarium wilt  209 nematodes  430, 440, 444 wild finger  467–468 wild Musa species  9, 30 wilts bacterial see bacterial wilts phytoplasma (BWAP)  343–345, 344, 345, 346 wind and cultivar selection  492, 493 cultural management  492 damage 487, 488, 490–492, 491 physical protection  487–490, 490, 491 Windward Islands  7, 50, 278, 510

Xanthomonas bacterial wilt  296–314 causal agent  299–301 control 306–314 diagnosis 306–307 disease cycle  301–305 distribution and spread  296, 303–305, 575

economic impact  296–297 on enset  296, 297, 299, 302, 303, 304, 305–306, 575 host susceptibility  305–306, 312–313 and nematodes  433 symptoms 297–299 enset 299, 302, 303 flowers  297, 298, 298, 299 fruit  298, 299 leaves 298–299, 299, 300, 301 pseudostems  301

yellow mat  464–465 yellow pulp  472 yellow Sigatoka see Sigatoka leaf spot youngest leaf spotted (YLS)  68, 75, 76, 122

Zasmidium biverticillatum (Ramichloridium biverticillatum) 163–165, 165 Zasmidium musae-banksii (Ramichloridium australiense) 165 Zasmidium musigenum (Ramichloridium musae)  163, 164 zimmu 132 zinc deficiency  482–484, 483 Zythia sp.  235