Sucking Pests of Crops [1st ed.] 9789811561481, 9789811561498

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Omkar  Editor

Sucking Pests of Crops

Sucking Pests of Crops

Omkar Editor

Sucking Pests of Crops

Editor Omkar Department of Zoology University of Lucknow Lucknow, Uttar Pradesh, India

ISBN 978-981-15-6148-1 ISBN 978-981-15-6149-8 https://doi.org/10.1007/978-981-15-6149-8

(eBook)

# Springer Nature Singapore Pte Ltd. 2020 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Springer imprint is published by the registered company Springer Nature Singapore Pte Ltd. The registered company address is: 152 Beach Road, #21-01/04 Gateway East, Singapore 189721, Singapore

Preface

The increase in human population in the twentieth century at a pace much beyond that in any of the previous centuries is attributed to multiple factors including more scientific breakthroughs, rapid industrialization, better health care, better understanding of hygiene, and increased food availability. The increased food availability can be attributed to both the increase in agricultural land and the development of high yielding seeds and more efficient means of pest management. But while the overall agricultural yield has managed to keep pace with the rapidly increasing human population, it has also led to a rapid increase in yet another population of organisms, the insect pests. Pests reduce crop productivity in various ways. Without preventive measures using pesticides, natural enemies, host plant resistance, and other nonchemical controls, 70% of crops could have been lost to pests. Weeds produce the highest potential loss (30%), compared to animal pests (23%) and pathogens (17%). However, the control measures for pathogens and animal pests show efficacy of 32 and 39%, respectively, compared to almost 74 % for weed control. Herbivorous insects are known to cause damage up to 20% of the crop yield despite massive global inputs in pest control measures. Globally, arthropods destroy an estimated 18–20% of annual crop production worldwide, at a value of more than US$ 470 billion. The greater proportion of these losses (13–16%) occurs in the fields, before harvest, and losses have been heaviest in developing countries. An overview of recent studies on global food loss and waste magnitudes shows a range from 27 to 32% of all food produced in the world. In fact, multiple studies have shown that the global crop losses due to insect pests have increased considerably in the post-green revolution era, in almost all the crops except cotton and rice. Of these pests, sucking pests, viz. aphids, scale insects, mealy bugs, thrips, whiteflies, leafhoppers, and mites, form one of the major concerns for agricultural crop yield as they inflict both quantitative and qualitative losses. Sucking pests pierce plant parts with slender, sharp-pointed mouthparts and suck the plant sap. Withdrawal of the sap results in minute white, brown, or red spotting on the leaves, fruits, or stems of the plant. It may also cause curling leaves, deformed fruit, general wilting, browning, and drying of the entire plant. Many sucking pests promote fungal growth due to their exudates. Many of these pests are also resistant to

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Preface

pesticides. Some of these are important virus vectors, transmitting a range of plant viruses. Because of the multipronged attack of sucking pests, as well as their cosmopolitan distribution and polyphagous nature of many of them such as aphids and whiteflies, sucking pests have turned into a serious affliction of numerous agricultural and horticultural crops. It is also predicted that the increase in temperatures accompanied with higher humidity owing to global warming-induced climate change is likely to increase the incidence of sucking pests. Thus, an understanding of the biology of sucking pests, their damaging stages, nature of damage, and control measures is essential for furthering agricultural yield. Keeping these facts in mind, the present book Sucking Pests of Crops was proposed. The present form of the book contains 16 chapters under two sections: (1) Agricultural Pests and (2) Horticultural Pests. It is hoped that it will cater to the needs of PG students of Agricultural Zoology, Entomology, and Zoology with specialization in Entomology along with faculty members and also the researchers of this field. In addition, it is also likely to be more useful to the policy planners involved in agriculture and plant protection. Lucknow, India March 16, 2020

Omkar

Acknowledgements

At the very outset, I take this opportunity to express my gratitude to the contributors of different chapters contained in this book for sparing time from their routine to prepare their respective chapters to the level they could. I am also thankful to Dr. Geetanjali Mishra, Associate Professor, Department of Zoology, University of Lucknow for her intellectual inputs and Ms. Apooorva Shandilya for her great assistance. I would also like to extend my thanks to my research students Dr. Shashwat Singh, Dr. Swati Saxena, and M/S Priya Singh, Chandni Verma, Tripti Yadav, Shriza Rai, Lata Verma, Deepali Gupta, and Gauravanvita Singh for their assistance and support in various ways. Grateful thanks are also due to my wife, Mrs. Kusum Upadhyay, for her constant encouragements and support by sparing me from household activities for this work. I also express my thanks to Dr. Mamta Kapila and Ms. Akanksha Tyagi from Springer Nature India, New Delhi for taking keen interest in this project and for their support for publishing this work in time.

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Contents

Part I

Agricultural Crops

1

Sucking Pests of Cereals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Omkar and Arun Kumar Tripathi

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2

Sucking Pests of Rice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Swoyam Singh and S. N. Tiwari

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3

Sucking Pests of Pulse Crops . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 107 Hem Saxena, Sanjay M. Bandi, and Revanasidda

4

Sucking Pests of Sugarcane . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135 Omkar and Arun Kumar Tripathi

5

Sucking Pests of Oilseed Crops . . . . . . . . . . . . . . . . . . . . . . . . . . . . 153 Sundar Borkar, Navya Matcha, Duraimurugan Ponnusamy, and M. Surya Prakash Reddy

6

Sucking Pests of Rapeseed-Mustard . . . . . . . . . . . . . . . . . . . . . . . . 187 Sunita Yadav and Mandeep Rathee

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Sucking Pests of Soybean . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 233 Neeta Gaur and Rashmi Joshi

8

Sucking Pests of Cotton . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 249 P. S. Shera, Vijay Kumar, and Vikas Jindal

9

Sucking Pests of Forage Crops . . . . . . . . . . . . . . . . . . . . . . . . . . . . 285 N. S. Kulkarni

Part II

Horticultural Crops

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Sucking Pests of Vegetable Crops . . . . . . . . . . . . . . . . . . . . . . . . . . 307 A. T. Rani, K. Vasudev, K. K. Pandey, and B. Singh

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Sucking Pests of Temperate Vegetable Crops . . . . . . . . . . . . . . . . . 341 Akhtar Ali Khan, Meinaz Nissar, A. A. Kundoo, and Irham Rasool

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Sucking Pests of Temperate Fruits . . . . . . . . . . . . . . . . . . . . . . . . . . 369 Akhtar Ali Khan, A. A. Kundoo, Meinaz Nissar, and Muntazir Mushtaq

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Sucking Pests of Mango . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 411 P. Venkata Rami Reddy, M. A. Rashmi, K. Sreedevi, and Sandeep Singh

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Sucking Pests of Grapes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 425 N. S. Kulkarni

15

Sucking Pests of Banana . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 451 B. Padmanaban and M. Mani

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Sucking Pests of Citrus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 481 Sandeep Singh, P. V. R. Reddy, and Sikha Deka

Editors and Contributors

About the Editor Omkar has over three decades of teaching experience and has been actively involved in research for nearly four decades. He is currently a Professor and Former Head of the Department of Zoology, University of Lucknow, Lucknow 226007, India. He is a fellow of the National Academy of Sciences, India, and several other professional bodies. He was also a recipient of the Young Indian Zoologist of 20th Century Gold Medal (2000) by the Zoological Society of India; Prof. T. N. Ananthakrishnan Foundation Award (2012); Rescholar Award of Excellence in Agricultural Entomology by the Association of Entomologists, Patiala (2014); Prof. G. S. Shukla Gold Medal by the Academy of Environmental Biology, India (2014); and the Prestigious Saraswati Samman by Govt. of Uttar Pradesh (2017). His research focuses on identifying and harnessing the potential of beneficial insects, in particular ladybird beetles, for which he is globally recognized. He has completed ten research projects funded by state and central government agencies. He is author of several books and journal articles on this theme.

Contributors Sanjay M. Bandi Division of Crop Protection, ICAR-Indian Institute of Pulses Research, Kanpur, India Sundar Borkar Jawaharlal Nehru Krishi Vishwa Vidyalaya, Jabalpur, Madhya Pradesh, India Sikha Deka Citrus Research Station, Tinsukia, Assam Agricultural University, Jorhat, Assam, India P. Duraimurugan ICAR-ICAR-Indian Institute Rajendranagar, Hyderabad, Telangana State, India

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Oilseeds

Research,

Neeta Gaur Department of Entomology, College of Agriculture, GBPUA&T, Pantnagar, Uttarakhand, India xi

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Editors and Contributors

Vikas Jindal Department of Entomology, Punjab Agricultural University, Ludhiana, Punjab, India Rashmi Joshi Research Scholar, Department of Entomology, College of Agriculture, GBPUA&T, Pantnagar, Uttarakhand, India Akhtar Ali Khan Division of Entomology, Sher-e-Kashmir University of Agricultural Sciences and Technology of Kashmir, Srinagar, Jammu and Kashmir, India N. S. Kulkarni IGFRI, SRRS, Dharwad, Karnataka, India Vijay Kumar Department of Entomology, Punjab Agricultural University, Ludhiana, Punjab, India A. A. Kundoo Division of Entomology, Sher-e-Kashmir University of Agricultural Sciences and Technology of Kashmir, Srinagar, Jammu and Kashmir, India M. Mani ICAR-Indian Institute of Horticultural Research, Bengaluru, Karnataka, India Navya Matcha Jawaharlal Nehru Krishi Vishwa Vidyalaya, Jabalpur, Madhya Pradesh, India Muntazir Mushtaq Division of Biotechnology, Sher-e-Kashmir University of Agricultural Sciences and Technology of Kashmir, Jammu, Jammu and Kashmir, India Suresh M. Nebapure ICAR Indian Agricultural Research Institute, New Delhi, India Meinaz Nissar Division of Entomology, Sher-e-Kashmir University of Agricultural Sciences and Technology of Kashmir, Srinagar, Jammu and Kashmir, India Omkar Department of Zoology, University of Lucknow, Lucknow, Uttar Pradesh, India B. Padmanaban ICAR-National Research Centre for Banana, Tiruchirappalli, Tamil Nadu, India K. K. Pandey ICAR-Indian Institute of Vegetable Research, Varanasi, Uttar Pradesh, India A. T. Rani Scientist (Agril. Entomology), Division of Crop Protection, ICAR-Indian Institute of Vegetable Research, Varanasi, Uttar Pradesh, India M. A. Rashmi Division of Entomology and Nematology, ICAR-Indian Institute of Horticultural Research, Bengaluru, India Irham Rasool Division of Entomology, Sher-e-Kashmir University of Agricultural Sciences and Technology of Kashmir, Srinagar, Jammu and Kashmir, India

Editors and Contributors

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Mandeep Rathee Department of Entomology, CCS Haryana Agricultural University, Hisar, Haryana, India M. Surya Prakash Reddy Jawaharlal Nehru Krishi Vishwa Vidyalaya, Jabalpur, Madhya Pradesh, India P. Venkata Rami Reddy Division of Entomology and Nematology, ICAR-Indian Institute of Horticultural Research, Bengaluru, India Revanasidda Division of Crop Protection, ICAR-Indian Institute of Pulses Research, Kanpur, India Hem Saxena Division of Crop Protection, ICAR-Indian Institute of Pulses Research, Kanpur, India P. S. Shera Department of Entomology, Punjab Agricultural University, Ludhiana, Punjab, India B. Singh ICAR-Indian Institute of Vegetable Research, Varanasi, Uttar Pradesh, India Sandeep Singh Punjab Agricultural University, Ludhiana, Punjab, India Swoyam Singh Department of Entomology, G.B. Pant University of Agriculture & Technology, Pantnagar, Uttarakhand, India K. Sreedevi ICAR-National Bureau of Agricultural Insect Resources, Bengaluru, Karnataka, India S. N. Tiwari Department of Entomology, G.B. Pant University of Agriculture & Technology, Pantnagar, Uttarakhand, India Arun Kumar Tripathi CSIR-Central Institute of Medicinal and Aromatic Plants, CIMAP, Lucknow, India K. Vasudev Department of Food and Public Distribution, Ministry of Consumer Affairs and Food and Public Distribution, Govt. of India, Krishi Bhawan, New Delhi, India S. S. Yadav Department of Entomology, CCS Haryana Agricultural University, Hisar, Haryana, India Sunita Yadav Department of Entomology, CCS Haryana Agricultural University, Hisar, Haryana, India

Part I Agricultural Crops

1

Sucking Pests of Cereals Omkar and Arun Kumar Tripathi

Contents 1.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 1.2 Systematics of Major Insect-Pests Attacking Cereal Crops . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 1.3 Major Insect Pests of Cereal Crops . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 1.4 Integrated Pest Management Approaches for Sucking Insect-Pests of Cereals . . . . . . . . . . . 47 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 50

Abstract

Cereal crops are grasses belonging to monocot plants under the family Poaceae. They are grown primarily for the harvesting of mature grains which are used or processed into staple food and animal feed. They are also processed into various products, such as starch, malt, biofuel (alcohol), and sweetener (i.e., high fructose corn syrup). They are also a rich source of carbohydrates. The top five cereals in the world ranked on the basis of production tonnage are maize (corn), rice (paddy), wheat, barley, and sorghum. The global losses due to various categories of pests vary with the crop and agroclimatic conditions. Total yield losses from different pests of all crops have been estimated to be US$ 500 billion worldwide. Animal pests account for 15.6% loss of production, pathogen 13.3%, and weeds 13.2%. The insects also cause indirect loss as a vector of various plant pathogens. The present chapter describes about major sucking insect-pests of cereal crops like barley, finger millet, maize, oat, rice, sorghum, and wheat. Common name, scientific name, host range, life cycle, and nature of damage are given for each insects covered under different cereal crops along with economic importance of Omkar (*) Department of Zoology, University of Lucknow, Lucknow, India A. K. Tripathi CSIR-Central Institute of Medicinal and Aromatic Plants, Lucknow, India # Springer Nature Singapore Pte Ltd. 2020 Omkar (ed.), Sucking Pests of Crops, https://doi.org/10.1007/978-981-15-6149-8_1

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the concerned crop. Integrated pest management approach for sucking insectpests is given separately in the last section of this chapter.

1.1

Introduction

The cereals are members of the grass family (a monocot family Poaceae, also known as Gramineae), which usually have long, thin stalks, such as wheat, rice, maize, sorghum, millet, barley, and rye. Their starchy grains are used as food. The term cereal is not limited to these grains, but, also refers to foodstuff prepared from the starchy grains of cereals like flours, breads, and pasta. On a worldwide basis, wheat, and rice are the most important crops, accounting for over 50% of the world’s cereal production. All of the cereals share some structural similarities and consist of an embryo (or germ), which contains the genetic material for a new plant, and an endosperm, which is packed with starch grains. Cereals are staple foods, and are important sources of nutrients in both developed and developing countries. Cereals and cereal products are an important source of energy, carbohydrate, protein, and fibre, as well as containing a range of micronutrients such as vitamin E, some of the B vitamins, magnesium, and zinc. Crop losses are usually defined as the reduction in either quantity or quality of yield (Zadoks and Schein 1979) and these may be caused by abiotic and biotic factors, leading to the reduction in crop productivity. Losses can occur at any stage of crop production in the field (preharvest) or even during storage (postharvest) (Oerke 2006). Direct yield losses caused by pathogens, animals, and weeds are altogether responsible for 20–40% loss of global agricultural productivity (Sharma et al. 2017; Kalsa et al. 2019). The limited data available indicate that arthropods may be destroying an estimated 18–20% of the annual crop production worldwide. Further, the losses are considerably higher in the developing tropics of Asia and Africa, where most of the future increase in world population is expected during the next 50 years. There is an urgent need to precisely estimate the extent of food loss and waste at different stages from the agricultural fields to human consumption with emphasis on the developing countries. This is the necessary first step towards development of safe, economical, and sustainable methods of pest management, as well as food security for the future (Sharma et al. 2017). The present chapter deals with sucking insect-pests of some economically important cereals like barley, finger millet maize, oat, rice, sorghum, and wheat in detail covering crop importance, list of major sucking insects attacking each crop along with their respective systematics, life histories, and their economic impacts on the cereal crops. Systematics of insects recorded on selected cereal crops has been tabulated at one place (Table 1.1) and detailed descriptions of major pests are given in detail for each crop.

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Table 1.1 Systematic of major sucking insect-pests attacking cereals Damaging insects

Plant name 4.1. Hordeum vulgare

Common name 4.1.1. Russian wheat aphid 4.1.2. English grain aphid 4.1.3. Green bug 4.1.4. Rose grass aphid

4.2. Eleusine coracana

4.1.5. Corn leaf aphid 4.1.6. The bird cherryoat aphid 4.1.7. Barley mealybug 4.1.8. Barley thrips 4.2.1. Rusty plum aphid 4.2.2. Root aphid 4.2.3. Ragi jassid

4.3. Zea mays

4.4. Avena sativa

4.3.1. Corn leaf aphid 4.3.2. Black bean aphid 4.4.1. The bird cherryoat aphid 4.4.2. Russian wheat aphid

Scientific name Diuraphis noxia (Mordvilko)

Order Hemiptera

Family Aphididae

Pest status (major/ minor) Major

Macrosiphum avenae (F.)

Hemiptera

Aphididae

Major

Schizaphis graminum (Rondani) Metopolophium dirhodum (Walker) Rhopalosiphum maidis (Fitch) Rhopalosiphum padi (L.)

Hemiptera

Aphididae

Major

Hemiptera

Aphididae

Major

Hemiptera

Aphididae

Major

Hemiptera

Aphididae

Major

Trionymus haancheni (Mc Kenzie) Limothrips cerealium (Haliday) Hysteroneura setariae (Thomas) Tetraneura nigriabdominalis (Sasaki) Cicadulina bipunctella bipunctella (Melichar) Rhopalosiphum maidis (Fitch) Aphis fabae (Scopoli) Rhopalosiphum padi (L.)

Hemiptera

Pseudococcidae

Major

Thysanoptera

Thripidae

Major

Hemiptera

Aphididae

Major

Hemiptera

Pemphigidae

Major

Hemiptera

Cicadellidae

Major

Hemiptera

Aphididae

Major

Hemiptera

Aphididae

Major

Hemiptera

Aphididae

Major

Hemiptera

Aphididae

Major

Diuraphis noxia (Mordvilko)

(continued)

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Table 1.1 (continued) Damaging insects

Plant name

4.5. Oryza sativa

4.6. Sorghum

Common name 4.4.3. Green bug 4.4.4. Barley thrips 4.5.1. Rice stink bug 4.5.2. Chinch bug 4.5.3.1. White rice leaf hopper 4.5.3.2. Sugarcane leaf hopper 4.5.3.3. Brown plant hopper 4.5.3.4. White backed plant hopper 4.5.3.5. Green leafhopper 4.5.4. Rice delphacid 4.5.5. Rice black bug 4.5.6. Earhead bug 4.6.1. Green bug 4.6.2. Corn leaf aphid 4.6.3. Chinch bug 4.6.4. Yellow sugarcane aphid

Order Hemiptera

Family Aphididae

Pest status (major/ minor) Major

Thysanoptera

Thripidae

Major

Hemiptera

Pentatomidae

Major

Hemiptera

Blissidae

Major

Hemiptera

Cicadellidae

Major

Pyrilla perpusilla (Walker)

Hemiptera

Lophopidae

Major

Nilaparvata lugens (Stal)

Hemiptera

Delphacidae

Major

Sogatella frucifera (Horvath)

Hemiptera

Delphacidae

Major

Nephotettix virescens (Distant) Tagosodes orizicolus (Muir) Scotinophora lurida (Burmeister) Leptocorisa oratorius (F.) Schizaphis graminum Rhopalosiphum maidis Blissus leucopterus Sipha flava

Hemiptera

Cicadellidae

Major

Hemiptera

Delphacidae

Major

Hemiptera

Podopidae

Major

Hemiptera

Alydidae

Major

Hemiptera

Aphididae

Major

Hemiptera

Aphididae

Major

Hemiptera

Blissidae

Major

Hemiptera

Aphididae

Major

Scientific name Schizaphis graminum (Rondani) Limothrips cerealium (Haliday) Oebalus pugnax (F.) Blissus leucopterus (Say) Cofana spectra (Distant)

(continued)

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Table 1.1 (continued) Damaging insects

Plant name 4.7. Triticum

Common name 4.7.1. Russian wheat aphid 4.7.2. English grain aphid 4.7.3. Green bug 4.7.4. Corn leaf aphid 4.7.5. The bird cherryoat aphid

Scientific name Diuraphis noxia

Order Hemiptera

Family Aphididae

Pest status (major/ minor) Major

Macrosiphum avenae

Hemiptera

Aphididae

Major

Schizaphis graminum Rhopalosiphum maidis Rhopalosiphum padi

Hemiptera

Aphididae

Major

Hemiptera

Aphididae

Major

Hemiptera

Aphididae

Major

1.2

Systematics of Major Insect-Pests Attacking Cereal Crops

1.3

Major Insect Pests of Cereal Crops

1.3.1

Hordeum vulgare L. (Barley)

Crop Importance Barley is an edible grain cereal, which is the fourth largest grain crop globally after wheat, rice, and corn. Barley is grown for many purposes, but the majority of all barley is used for animal feed, human consumption, or malting (Kling 2004). High protein barleys are generally valued for food and feeding, and starchy barley for malting. The major sucking insect-pests recorded on this plant are described below.

1.3.1.1 Common Name: Russian Wheat Aphid (RWA) Scientific name: Diuraphis noxia Mordvilko (Hemiptera: Aphididae) Host Plants Russian wheat aphid affects cereal crops throughout the world, primarily barley (Hordeum vulgare) and wheat (Triticum aestivum) (Miller et al. 2001). RWA attacks most of the cereals including wheat, barley, triticale, rye, and oat. Other primary hosts include durum wheat (Triticum durum), field brome grass (Bromus arvensis), Elymus sp., and jointed goat grass (Triticum cylindricum).

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Fig. 1.1 Wingless adult of Diuraphis noxia, showing the presence of a “double tail”

Fig. 1.2 Winged (alate) adult Diuraphis noxia

Primary hosts for RWA support the entire life cycle and allow for reproduction to occur. All instars and adults can feed on these plants. Secondary hosts are plants that only support adults and final instars. They allow the aphid to survive but not reproduce. Secondary hosts include cereal rye (Secale cereale), triticale (Triticum aestivum x Secale cereale), and various grasses in the Poaceae family, such as oats (Avena sativa), tall wheat grass (Agropyron elongatum), and Indian ricegrass (Oryzopsis hymenoides). Life Cycle RWA is a small insect, 1.5–1.8 mm in length. The body is light green in colour, and an elongated spindle-shaped. They have short antennae (about one-quarter of body length), and a distinctive double-tailed (cauda) appearance when viewed from the side (Fig. 1.1). They also lack the visible siphunculi (special tubes or pores in the abdomen of aphids for extruding waxy defensive fluids), which are present on other cereal aphids. This characteristic distinguishes RWA from other cereal aphids. Instars look similar to apterous adults but do not develop the characteristic caudal features until the fourth and fifth instars. This species also has alate (winged) adult morphs (Aalbersberg et al. 1987) (Fig. 1.2).

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Fig. 1.3 Colony of Diuraphis noxia Damage: yellow and white streaks along the leaf surface

RWA spend their entire life on cereals and grasses, and have the ability to reproduce both sexually and asexually. Yet, RWA is only known to reproduce sexually in Russia and central Asia, and male aphids have not been observed in many parts of the world (Tolmay 2006). Sexual reproduction allows the aphids to overwinter as eggs, in contrast to areas where asexual reproduction occurs that requires the aphid to continue to feed over the winter period. All invasive populations of RWA outside of its natural range are parthenogenetic (i.e., reproduce asexually). Asexual Reproduction of RWA Asexually reproducing populations of RWA are all female and adults give birth to live nymphs. After the fourth moult, aphids develop into either wingless (apterous) or winged (alate) adults. Wingless adults have a higher reproductive capacity and can produce 4–5 nymphs per day for 3–4 weeks. Reproduction rates increase as the temperature increases with generation times becoming shorter and more young produced by each female. In general, maturation is completed within 7–10 days. Sexual Reproduction of RWA RWA are holocyclic, and, therefore, they can reproduce both sexually (usually for overwintering as eggs) and asexually (mostly during the warmer months). After mating, females lay 8–10 eggs on young cereal plants and die a few days afterwards. The eggs hatch in early spring and aphid population increases rapidly by parthenogenetic reproduction. Nature of Damage Russian wheat aphid feeding produces strong plant symptoms due to the injection of saliva into the plant during feeding (Kazemi et al. 2001). Symptoms include rolled leaves, chlorotic spots, leaf streaking, trapped awns giving a hooked appearance and a stunted appearance under heavy infestation (Kazemi et al. 2001). Heavily infested plants may typically look stunted with yellow or whitish streaks on leaves (Fig.1.3). These streaks, basically, are formed due to the saliva injected by the RWA. RWA

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can act as a vector for viruses, including Barley yellow dwarf virus and Barley stripe mosaic virus (Elsidaig and Zwer 1992; Kazemi et al. 2001).

1.3.1.2 Common Name: English Grain Aphid Scientific name: Macrosiphum (Sitobion) avenae F. (Hemiptera: Aphididae) Host Range English grain aphid is widespread throughout Europe, Asia, West Africa, America, and Japan. It prefers both cultivated and wild cereal grasses. Among the wild cereal grasses, it prefers Phleum pratense L., Avena fatua L., Agropyrum repens P.B., Dactylis glomerata L., Bromus mallis L., Bromus secalinus L., and Festuca pratensis Huds., etc. Life Cycle English grain aphids are pale green in colour with black antennae and black cornicles. They are up to 2.5 mm in length. Apterous female have green or yellow-brown Fig. 1.4 Macrosiphum avenae wingless adult

Fig. 1.5 Macrosiphum avenae winged adult

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fusiform body and long legs (Fig. 1.4). Antenna is longer than body. Siphunculi are black (1.5 times as long as light green tail). Winged female has red-brown thorax and green abdominal segments (Fig. 1.5). The female produces 6–12 eggs, which are oval and black. Overwintering takes place during the egg phase on winter cereals, and also on cereal weeds. Larval period lasts 8–12 days. Life span of apterous parthenogenetic females is about 30–60 days and they produce 20–40 larvae. Most populations are anholocyclic giving birth parthenogenetically only to asexual morphs. A female produces 20–40 nymphs which overwinter; sometimes already as apterous adults. Their life span takes about 4–10 weeks. A small part of the population is holocyclic. Some of these individuals give birth to males and females that mate, producing about a dozen eggs which overwinter and hatch during next spring. There are 15–20 annual generations (Gaur and Mogalapu 2018). Nature of Damage They usually prefer the upper parts of the plant and are commonly found in wheat heads where the accumulation of honeydew and sooty mould are sometimes observed. The pest causes much damage to wheat, barley, rye, oats, sorghum, and maize, resulting in great yield reductions. This aphid excretes honeydew which is colonized by sooty mould fungi. It is also a vector of the barley yellow dwarf virus (BYDV) and affects crops worldwide (Leybourne et al. 2020).

1.3.1.3 Common Name: Green Bug Scientific name: Schizaphis graminum (Rondani) (Hemiptera: Aphididae) Host Range Greenbug was first reported on oats during early twentieth century and also has colonized successfully in sorghum during 1960 (Harvey and Hackerott 1969). Greenbug is known to be originated from Virginia, North America (Hunter 1909) with a contradictory report that it might have originated from Italy. There are more than 80 grass species, including several cultivated cereals, millets, and turfgrasses, that support the survival of S. graminum.

Fig. 1.6 Schizaphis graminum wingless adult

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Fig. 1.7 Schizaphis graminum winged adult

Life Cycle Greenbugs are small (1.3–2.1 mm), elongate oval-shaped aphids with head and first part of thorax straw to pale green and with light to medium green abdomen (Fig. 1.6). A darker green stripe down the middle of the top surface of the abdomen is most visible on last instar nymphs and adults. The antennae are uniformly dusky. The cornicles or siphunculi are pale with slightly flared and darkened tips. It is facultatively holocyclic, i.e., in cold climates. It may reproduce sexually in the cool season and lay eggs that are able to withstand low temperatures, but where winters are mild, it will propagate parthenogenetically throughout the whole year. After mating, the winged female (Fig. 1.7) lays eggs into leaf sheaths of winter cereals or other grasses: 10–12 in groups of 2–4 over a period of 4–5 weeks. In contrast to other aphids, S. graminum has no change of host. In the spring, wingless females hatch from the eggs. After 1–3 weeks, they give birth to live young and asexual reproduction starts again (Shehata et al. 2018). There are several biotypes of greenbug that differ greatly in terms of host preference, temperature tolerance, and their ability to overcome plant resistance. Several biotypes (C, E, I, and K) have been identified of which biotype I is the most predominant and severe (Punnuri et al. 2013). There are eleven documented S. graminum biotypes, designated by letters from A to K, although only eight have any relation to HPR varieties (Porter et al. 1997). The term ‘biotype’ here will be used to designate strains of insects differing in their capability of infesting certain Host Plant of Resistant varieties (Diehl and Bush 1984). S. graminum has a bacterial endosymbiont (Buchnera aphidicola) that plays an important role in the insect’s amino acid metabolism.

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Fig. 1.8 Schizaphis graminum damage on barley

Nature of Damage Greenbug ingests phloem sap with its piercing-sucking mouthparts. This weakens the plant due to loss of nutrients and water. The necrotic lesions appear as small yellow or reddish spots around the punctures. As feeding continues, these soon coalesce into larger patches. Eventually, the whole leaf will wilt and the plant may be killed off completely. High densities of greenbugs significantly lower the plant’s photosynthetic activity, resulting in stunting and reduced yield. Infestation is particularly dangerous during stem elongation. The aphid prefers to settle on the undersides of older, lower leaves, which their colonies may cover completely in severe cases. On expanding, the colonies gradually move upwards on the plant. Greenbug saliva has enzymatic activity that breaks down cell walls and chloroplasts in susceptible plants (Al-Mousawi et al. 1983). Greenbug damage is often apparent in the field as circular patches that slowly grow in size (Fig. 1.8). The greenbug is the vector of several plant viruses including barley yellow dwarf virus, sugarcane mosaic virus, maize dwarf mosaic virus, and millet red leaf virus (Harvey et al. 2005; van Emden et al. 2007).

1.3.1.4 Common Name: Rose Grass Aphid Scientific name: Metopolophium dirhodum Walker (Hemiptera: Aphididae) Host Range M. dirhodum is of palaearctic origin, but is nowadays found in almost all grainproducing regions of the world. The rose grass aphid host alternates from rose as the primary host in spring and early summer to cereals and grasses especially wheat, barley, and maize as the secondary hosts (Honek et al. 2018).

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Fig. 1.9 Metopolophium dirhodum wingless adult

Fig. 1.10 Metopolophium dirhodum winged adult

Life Cycle Rose grass aphid has a spindle-shaped body and is green, with a noticeably lighter stripe along the back (Fig. 1.9). The apterae are up to 3 mm in length, the oviparous females less than 2 mm. The length of the antennae is about three-fourths that of the whole body. It usually undergoes an alternation of generations and hosts. The summer generation (virginoparae), which lives on the secondary host, is parthenogenetic, and consists solely of females. Only in the autumn morphs (sexuales) of both sexes appear, mate, and produce eggs, which are laid on the primary host (Rosa spp.). The colony founders (fundatrices) hatch in the spring, and adults of the subsequent second or third generations migrate to the summer host (Poaceae). However, in regions with mild, temperate climates, this aphid can be anholocyclic, i.e., without a sexual generation and with aphids overwintering on the secondary host as well. Mild autumns and winters, an early rise in temperatures in spring, and warm, dry weather in summer are conditions favourable for rapid population increase. Peak densities tend to more or less coincide with the milky stage of cereal development.

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As the plant grows, the aphid colonies climb from ageing to younger parts. When the host becomes overcrowded, winged morphs (alatae) (Fig. 1.10) develop and then spread to other plants. Nature of Damage The preferred feeding sites are the undersides of leaves, and less often, the ears. M. dirhodum feeds on plant sap, which it obtains by puncturing phloem vessels. Its saliva does not contain substances that are toxic to the plant, but the drain of nutrients and water can lead to yield losses, if the aphid occurs in large numbers. Plants that are already under water stress may start to yellow and become stunted. It is also a vector of several plant viruses, most importantly barley yellow dwarf virus, maize mosaic virus, and potato viruses.

1.3.1.5 Common Name: Corn Leaf Aphid Scientific name: Rhopalosiphum maidis Fitch (Hemiptera: Aphididae) Host Range The corn leaf aphid is a serious pest of maize and barley with Asiatic origin but it is now distributed throughout the tropics and temperate regions of the world (Blackman and Eastop 2000; Kuo et al. 2006). Host plants include many common grass weeds (barnyard grass, crabgrass, and foxtail) and most cereal crops (corn, barley, rye, oats, wheat, sorghum, and millet). Life Cycle Corn leaf aphids are oval-shaped, with soft bodies and a pair of cornicles protruding from end of their abdomen (Fig. 1.11). This aphid is bluish-green or black, with black legs and short antennae. Corn leaf aphids overwinter as females on the host plant. Their average body length is 2.56 mm. Both wingless and winged forms occur. They are polymorphic in nature. No males or egg stages occur. Females give birth to live youngs via parthenogenesis. Offsprings develop through four nymphal instars, each instar lasts for 2 days. Total life cycle is completed in 7–8 days (Kuo Fig. 1.11 Adult and nymph of Rhopalosiphum maidis

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et al. 2006). There are up to nine generation per year. These aphids are usually found in the whorls and on the tops of newly emerged leaves of the plants. Nature of Damage This aphid is a polyphagous pest and can cause damage to many host plant species and weeds from Poaceae and occasionally Cyperaceae and Typhaceae. R. maidis damages its host plants by feeding, viral disease transmission, and honeydew production. Aphid infestation occurs on seedlings, leaves, inside the whorl, the covers inflorescence of plants and produces plentiful honeydew (Blackman and Eastop 2000), which may result in deformed leaves as well as the sterilization of inflorescences (Hill 1987). In addition, R. maidis is a vector of plant viruses and may transmit ten viral diseases to cereals (Blackman and Eastop 2000). It interferes with pollen production and fertilization, resulting in poor kernel fill of the ears. Infestation also can cause a delay in plant maturity and reduced plant size (Bing and Guthrie 1991). Honeydew secreted by the aphids supports growth of sooty mould fungus, causing an unsightly appearance of the ears.

1.3.1.6 Common Name: The Bird Cherry-Oat Aphid Scientific name: Rhopalosiphum padi Linnaeus (Hemiptera: Aphididae) Host Range The bird cherry-oat aphid is found almost worldwide (except the subtropical and tropical regions). It attacks almost all cereal crops. The primary hosts are Prunus spp., and the secondary hosts are many species of Poaceae, Cyperaceae, and Typhaceae. Life Cycle The body of the apterous females is about 2.5 mm long (Fig. 1.12) and green-brown in colour. The head, siphunculi (which are swollen) and cauda are brown-black. The alate females (Fig. 1.13) are mostly green, siphunculi brown, body length about Fig. 1.12 Rhopalosiphum padi showing siphunculi (darker brown parts protruding from abdomen) and single tail

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Fig. 1.13 Rhopalosiphum padi winged adult

2.4 mm. Female lays egg in the narrow gap between the axillary buds and the stem. After egg hatch, the newly emerged nymphs move to bird cherry leaves, where they feed and develop. Nymphs develop rapidly into very large light green fundatrices. The fundatrix is pale green and 2.5–3.0 mm in length. These fundatrices give rise to a second, wax covered, generation. The bird cherry-oat aphid often reproduces exclusively by parthenogenesis from spring to late summer. All individuals are females, and they give birth to live young, which are also female. Most of them remain wingless. Each female is able to produce 60–80 larvae during its reproductive period of 3–4 weeks. Under stress conditions, winged forms appear and start to colonize other plants. In warm weather, the generation cycle can be completed in about a week. Each female is able to produce 60–80 larvae during its reproductive period of 3–4 weeks. Accordingly, population growth can be very rapid. In climates with mild winters, parthenogenesis remains the only mode of reproduction throughout the year (Luo et al. 2019). In colder regions, however, sexual forms appear in the autumn, which then migrate to the primary (winter) host (e.g., the bird cherry), mate, and produce eggs, which are laid near the buds. The overwintering eggs are very frost-resistant. In the spring, female founders (the so-called fundatrices) hatch and start to produce offspring. After several generations (usually three) on winter hosts, migration to the summer host starts. The summer (secondary) hosts are grasses, including most cereal crops. Nature of Damage R. padi uses its piercing-sucking mouthparts to penetrate plant tissues in order to reach a vascular bundle and ingest phloem sap. Strong infestations can sometimes lead to contortion of leaves. The insect causes most damage by transmitting a number of viruses, especially barley yellow dwarf virus (BYDV), the cereal yellow dwarf virus-RPV, filaree red leaf virus, maize leaf fleck virus, and rice giallume

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virus, oat yellow leaf disease, and the onion yellow dwarf virus. Infection with BYDV causes barley and wheat to turn yellow, whereas oat becomes reddish in colour. Affected plants are generally severely stunted and non-productive (Stern 1967). During severe infestation, honeydew produced, can create a sticky film on plant surfaces that can reduce photosynthesis and promote the growth of sooty mould.

1.3.1.7 Common Name: Barley Mealybug Scientific name: Trionymus haancheni McKenzie (Hemiptera: Pseudococcidae) Host Range The Haanchen barley mealybug was discovered for the first time on barley in Idaho in June 2003 (Alvarez 2004). Life Cycle Mealybugs are named for the waxy secretions that cover the soft bodies of these insects. They are pink, oval-shaped females with body length up to 5 mm. They have well-developed legs and are covered with a distinctive white, waxy secretion (Fig. 1.14). Egg masses are laid in a sac under a leaf sheath at the base of the plant and covered with cottony wax. A single female can lay up to 256 eggs. After hatching, the immature crawlers disperse to protected feeding sites under the leaf sheath. Crawlers moult into successive instars, each resembling small adults, becoming less mobile with each instar (McKenzie 1967). The Haanchen mealybug is apparently able to survive winter where it is protected by soil and plant material. Although winged male forms occur, they are rarely detected and do not appear to feed or damage plants. The number of generations is not known, but all stages have been found coexisting on infested plants. Fig. 1.14 Trionymus haancheni

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Nature of Damage Both immatures and females damage crops. The first signs of mealybug presence are ovisacs (cottony clusters of eggs) at the base of the plants. They feed with sucking mouthparts and reduce the amount of chlorophyll in the leaves, causing extensive yellowing and browning of the foliage. In addition to direct feeding injury to barley plants, the Haanchen barley mealybug can indirectly damage the crop by producing honeydew, which has the potential to reduce grain quality. Mealybug infestations cause yellowing of the foliage, reduced vigour, and root damage.

1.3.1.8 Common Name: Barley Thrips Scientific name: Limothrips cerealium (Haliday) (Thysanoptera:Thripidae) Host Range Barley thrips cause damage to cereal crops like barley, maize, oats, and wheat. Life Cycle Their bodies range from a pale yellow to darker brown and they have slender wings fringed with fine hairs (setae). The life cycle consists of five stages: egg, larva, prepupa, pupa, and adult. Adults are straw-coloured, yellowish-brown, and elongated measuring 1mm in length. Antennae have seven segments with the first segment paler and the second is usually dark. A brown band marks anterior edge of the abdominal tergites. There is a single pair of pores on tergite nine (Fig. 1.15). Females deposit eggs directly in the host tissue. Eggs are minute, kidney shaped laid in slits in leaf tissues. Nymphs are creamy to pale yellow in colour, resemble adults but wingless. Nymphs hatch from eggs after 5 days and begin to feed on the plants. Occasionally, wing buds are visible during pre-pupal stages. After two moults, the nymphs Fig. 1.15 Limothrips cerealium

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enter the pre-pupal stage (lasts about 1 day) during which wing buds are developing externally. Gradually the immobile pupa forms with the antennae folded over its back (occurs in the soil or curled leaves and is rarely seen). Nymphal and pupal period last for 5 and 4–6 days, respectively. The preimaginal stage is spent in soil without feeding. The adults survive for 2–4 weeks. Life cycle from egg to adult lasts for 30–35 days and they have about 10–12 overlapping generations. Adults emerge from the pupa after 2–3 days. Males are rare and the reproduction is parthenogenetic. Nature of Damage Nymphs and adults lacerate the tissue and suck the sap from the upper and lower surfaces of leaves. They inject saliva and suck the lyzed contents of plant cells resulting in silvery or brown necrotic spots. Seedlings infested with thrips grow slow and the leaves become wrinkled, curl upwards, and distorted with white shiny patches. Rusty appearances in patches develop on undersurface of leaves. Thripsinjured plants have oviposition scars on blades, ears with whitened spikelets, either empty or with shrivelled grains (Koppa 1970). They breed on the leaves, in the florets and cause dead tips in various grasses and young cereal crops (Lewis 1997). Damage causes yield reduction in cereal crops and also reduced germination of barley seed produced for the brewing industry. It is one among the few thrips species, whose biting of humans have been found to cause dermal pseudo-delusory syndrome (Guarneri et al. 2006).

1.3.2

Finger Millet: Eleusine coracana L. Gaertn

Crop Importance The grains of small millets are nutritionally superior to rice and wheat. They provide cheap proteins, minerals, and vitamins to poorest of the poor, where the need for such ingredients is the maximum. Millets are small grained cereals, the smallest of them include finger, kodo, foxtail, proso, little and barnyard millets. Nutritional superiority of millets is considered as nutri-cereals (Nutritious grains). Sucking pests of finger millet are responsible for considerable economic loss to the crop. These include species of Hemiptera and of the families Cicadellidae, Pimphigidae, and Aphididae. Details about some of the sucking insects causing damage to finger millets are given below.

1.3.2.1 Common Name: Rusty Plum Aphid Scientific name: Hysteroneura setariae (Thomas) (Hemiptera: Aphididae) Host Range Rusty plum aphid is known to attack all grasses belonging to plant family Poaceae, Cyperaceae, Poaceae, and Liliaceae. It has been recorded all over India, Australia, South Africa, Japan, Korea, Philippines, and Taiwan (Blackman and Eastop 2000). It was reported for the first time in India on rice, sorghum, Italian millet, and other crops in Madras (Mishra et al. 2010).

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Fig. 1.16 Hysteroneura setariae

Life Cycle It is a small, brown aphid with dark siphunculi and a pale cauda (Fig. 1.16). It usually forms colonies at the bases of spikelets of plants belonging to Poaceae, sometimes on leaves or unripe seeds. Alatae have greenish-grey abdomen. Length of body is around 1.85 mm. Life cycle is completed within 7–8 days and it reproduces for 13–14 days. The rate of reproduction on an average was 2.84–4.55 nymphs/day. There are two moultings at an interval of 2 days and first nymph is produced within 7–8 days. Nature of Damage Adult and nymphs suck the sap of the plant from the base of the spicklets, and spread to the entire plant. This results in reduced vigour of the plant and stunted growth. In severely infested plants, even the earheads are fully covered by aphids. The rusty plum aphid can also indirectly damage plants by vectoring various plant viruses, including cucumber mosaic virus (CMV), watermelon mosaic virus (WMV) (Coudriet 1962), sugarcane mosaic virus (SCMV) (Harborne 1988), bermuda grass mosaic virus (BgMV) (Masumi et al. 2011), barley yellow dwarf virus (BYDV) (Wangai et al. 1991), soybean mosaic virus (SMV) (Quimio and Calilung 1993), guinea grass mosaic virus (GGMV) (Kukla et al. 1984), onion yellow dwarf virus (OYDV), maize dwarf mosaic virus (MDMV), zucchini yellow mosaic virus (ZYMV) (Blackman and Eastop 2000), and peanut stripe virus (PStV) (Saleh et al. 1989). However, the aphid is not known to transmit any viral disease of rice (Jahn et al. 2005).

1.3.2.2 Common Name: Root Aphids Scientific name: Tetraneura nigriabdominalis (Sasaki) (Hemiptera: Pemphigidae) Host Range Root aphids are reported from Bangalore, Kolar, and Tumkur districts of Karnataka state. The eighteen host plants of these aphids belong to families Poaceae, Cyperaceae; Commelinaceae, and Rubiaceae.

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Fig. 1.17 Tetraneura nigriabdominalis adult wingless

Fig. 1.18 Tertaneura nigriabdominalis (alate)

Life Cycle They are small, greenish, or brownish white, plump, oval-bodied apterae (body length, 1.5–2.5 mm) clustered on roots of the plant (Fig. 1.17). Alatae (body length, 1.5–2.3 mm) have a shiny black head and thoracic lobes and a brown abdomen (Fig. 1.18). Antennae six-segmented, 0.17 times as long as body. Siphunculi have dark sclerotic rim. Reproduction is normally viviparous. There are four nymphal instars with the total nymphal period of 7–9 days. Adult female longevity varies 5–11 days and may produce 10–35 offspring each. Nature of Damage These aphids assume serious pest status on ragi. They suck the sap of the plant from the roots. As these aphids are confined to the root zone, their presence is rarely noticed until the plants show symptoms of wilting, excess tillering, stunted growth, and early maturity and occasionally caused drying of the roots. With consequent ill effect on the plant, like yellowing, stunting, and some time poor seed setting or produce shrivelled grains are the other symptoms. Infested plants turn pale yellow and become stunted. Its presence on the roots of some hosts is indicated by a reddish-

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purple discolouration of the leaves (Blackman and Eastop 1984). Sometimes, it causes indirect damage by transmitting plant viruses.

1.3.2.3 Common Name: Ragi Jassid Scientific name: Cicadulina bipunctella bipunctella (Melichar) (Hemiptera: Cicadellidae) Host Range Species of Cicadellidae are commonly known as leaf hoppers. The maize orange leafhopper is distributed widely from Africa to Asia including Japan, and northern Australia. Several cereal crops like maize and rice act as host plants. Life Cycle This hopper is free-living, multivoltine, and polyphagous in nature (Matsukura et al. 2010). Adults are small brown coloured, and wedge shaped (Fig. 1.19). The hoppers lay eggs inside the plant tissue and eggs hatch in a week time depending on temperature and humidity. The nymphs on hatching start sucking sap from the tender parts of the plant. Nymphs are pale greenish almost translucent and walk diagonally. Nymphal period is 7–9 days. Nature of Damage The maize orange leafhopper induces galls on various plants of the Poaceae. Both adults and nymphs of Cicadulina bipunctata have the ability to induce galls on their host plants, a unique feature among gall inducing insects. Galls produced by the leafhopper are characterized by growth stunting of the host plant and swelling of leaf veins, symptoms commonly referred to as ‘wallaby ear disease’ (Matsumura and Tokuda 2004).

Fig. 1.19 Cicadulina bipunctella adult

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Recent studies have recognized that some chemicals injected by Cicadulina bipunctata during feeding are responsible for gall induction (Matsukura et al. 2012). Galls are induced only on leaves which are newly expanding during leafhopper feeding, but not on leaves already developed or those which expand after the removal of leafhoppers (Matsukura et al. 2009). The degree of gall induction depends on the density and duration of feeding by Cicadulina bipunctata (Tokuda et al. 2013). They appear during early crop growth stage. Both adult and nymphs cause damage to seedlings in the nursery and main field by sucking sap from tender leaves. Signs of attack on damaged plant show white dots and specks. In severe stage, these specks give burnt appearance. These pests pose severe problem by acting as vectors of streak and mottle streak virus disease. They are key agents in causing epidemics of viral diseases on finger millet.

1.3.3

Maize: Zea mays L.

Crop Importance Maize is the world’s top ranking food crop followed by wheat and rice. Maize is sown and harvested somewhere in the world in every month of the year (Sharma and Dass 2012). Globally, maize is grown over an area of 183.57 million hectare with 1068.79 million metric tons production and productivity of 5.8 metric tons/hectare. India produced 23.70 million tonnes of maize grains from an area of 9.30 million hectare having an average productivity of 2.57 tons per hectare (ESIAPYMC 2015–2016). Maize crop is versatile in economic uses as well and as such referred to as ‘a miracle crop’ (Singh et al. 2012). It provides the industrial raw material for the production of glucose, starch, dextrin, cornflakes, corn oil, etc., along with nutritional needs (Bibi et al. 2010). Besides a large number of pharmaceutical products, alcoholic beverages are also commercially prepared from maize. The cobs are used for cleaning, brushing, polishing, abrasives for soaps, ceramics, glues and adhesives, as a carrier for pesticides, rubber compounds, and tyres (Ashfaq and Ahmad 2002). Maize plant is attacked by 140 species of insect pests causing a varying degree of damage. Details about some of the sucking insects damaging maize are described below.

1.3.3.1 Common Name: Corn Leaf Aphid Scientific name: Rhopalosiphum maidis Fitch (Hemiptera: Aphididae) (for details please see Sect. 1.3.1.5.) 1.3.3.2 Common Name: Black bean aphid Scientific name: Aphis fabae Scopoli (Hemiptera: Aphididae) Host Plants The black bean aphid is one of the most important pests of several cultivated crops throughout the world (Volkl and Stechmann 1998). It has been recorded on more

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Fig. 1.20 Aphis fabae (nymph and adult)

Fig. 1.21 Aphis fabae (winged)

than 200 host plant species in the world and about 50 plant species are susceptible to attack by this aphid in Iran (CAB International 2000). Its primary hosts on which the eggs overwinter are shrubs such as the spindle tree (Euonymus europaeus), Viburnum species, or the mock-orange (Philadelphus species). Its secondary hosts, on which it spends the summer, include a number of crops including sugar beets, celery, potatoes, tomatoes, etc. Life Cycle The body of the apterous female is brown to black and the cauda and siphunculi are black (Fig. 1.20). Alate females are mostly black, with a greenish area on the

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abdomen (Fig. 1.21). The length of the female body is 1.7–2.8 mm. The antennae are less than two-thirds of the length of the body, and both they and the legs are pale yellow in colour with black tips. Near the rear of the abdomen is a pair of slender, elongated tubes known as cornicles or siphunculi (Fig. 1.15). Their function is the production of a defensive waxy secretion. They are twice as long as the finger-like tail and both are brownish-black. The black bean aphid has both sexual and asexual generations in its life cycle. The primary host plants are woody shrubs, and eggs are laid on these by winged females in the autumn. The adults then die and the eggs overwinter. The aphids that hatch from these eggs in the spring are wingless females known as stem mothers. These are able to reproduce asexually, giving birth to live offspring, nymphs, through parthenogenesis. The life span of a parthenogenetic female is about 50 days and during this period, each can produce as many as 30 young ones. The offspring are also females and able to reproduce without mating, but further generations are usually winged forms. In the autumn, winged forms move to different host plants, where both males and females are produced. These mate and the females lay eggs which overwinter. Nature of Damage They suck sap from stems and leaves and cause distortion of the shoots, stunted plants, reduced yield, and spoiled crops. To obtain enough protein, aphids need to suck large volumes of sap. The excess sugary fluid, honeydew, is secreted by the aphids. It adheres to plants, where it promotes growth of sooty moulds. These are unsightly, reduce the surface area of the plant available for photosynthesis, and may reduce the value of the crop. These aphids are also the vectors of about 30 plant viruses, mostly of the non-persistent variety. The aphids may not be the original source of infection, but are instrumental in spreading the virus through the crop.

1.3.4

Oat: Avena sativa L.

Crop Importance Oats rank sixth in the world cereal production statistics following wheat, maize, rice, barley, and sorghum. It is an important livestock feed and is a good source of protein, fibre, and minerals. This crop is considered to be a rich source of protein, equal to meat, milk, and egg protein. As food, oats are mostly preferred in breakfast. Out of cereals, the highest amounts of β-glycan are found in barley and oats grains (Ahmad and Zaffar 2014). It has wider adaptability because of its excellent growth habits, quick re-growth, and better yield potential and provides palatable, succulent, and nutritious green fodder (Ahmad et al. 2014). It is cultivated in Punjab, Haryana, West Bengal, Jammu and Kashmir, Himachal Pradesh, Uttar Pradesh, Madhya Pradesh, Rajasthan, and Maharashtra.

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Oat crops are heavily attacked by wheat aphid, oat thrips apart from other insects. Details about sucking insects damaging oats are described below.

1.3.4.1 Common Name: The Bird Cherry-Oat Aphid Scientific name: Rhopalosiphum padi Linnaeus (Hemiptera: Aphididae) For details, please see Sect. 1.3.1.6 1.3.4.2 Common Name: Russian Wheat Aphid (RWA) Scientific name: Diuraphis noxia Mordvilko (Hemiptera: Aphididae) For details, please see Sect. 1.3.1.1 1.3.4.3 Common Name: Green Bug Scientific name: Schizaphis graminum (Rondani) (Hemiptera: Aphididae) For details, please see Sect. 1.3.1.3 1.3.4.4 Common Name: Barley Thrips Scientific name: Limothrips cerealium (Haliday) (Thysanoptera: Thripidae) For details, please see Sect. 1.3.1.8

1.3.5

Rice: Oryza sativa L.

Crop Importance Rice is a staple food crop and more than 90% of the world’s production is consumed in Asia. Three billion people depend on it as a major source of their subsistence diet (Cantrell and Reeves 2002). In India, 221 species of insects feeding on rice were reported (Arora and Dhaliwal 1996). Among these rice pests, about 20 of them are of economic importance. Details about some of the sucking insects damaging rice are described below.

1.3.5.1 Common Name: Rice Stink Bug Scientific name: Oebalus pugnax F. (Hemiptera: Pentatomidae) Host Plants This agricultural pest is known to attack cereal crops with small seeds, particularly wheat, sorghum, and rice. The insect also lives in wild grasses such as Johnson grass, barnyard grass, and sedge. Non-food crop hosts of the rice stink bug include: Echinochloa crusgalli, E. colona, Digitaria sanguinalis, Panicum dichotomiflorum, Phalaris minor, Paspalum urvillei, and Sporobolus poiretti. Life Cycle The adult Oebalus pugnax measures 9.5–13.0 mm in length. The rice stink bug is easily distinguished from other stink bugs because of its narrower profile and lighter colour. These true bugs are typically straw-coloured with sharp points on the apex of the shield and a yellow triangle exhibited on centre of the shield. Some adults have

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Fig. 1.22 Oebalus pugnax adult

Fig. 1.23 Oebalus pugnax eggs

grey colouring near the yellow triangle, while others may be a darker brown rather than straw-coloured (Fig. 1.22).

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Fig. 1.24 Life cycle of Oebalus pugnax

The rice stink bug lays its eggs in masses (two rows per mass) on rice foliage (Fig. 1.23). After egg hatch, nymphs (immature rice stink bugs without wings) complete five instars before becoming adults (Fig. 1.24). Each instar is a little bigger than the previous instar. The later instars (4th and 5th) and adults cause the most severe damage. Nature of Damage Rice growth and development has been categorized into four phases: the seedling, vegetative, reproductive, and ripening phases (Chaudhary et al. 2002). As a late season pest of rice, the reproductive and ripening phases are utmost concern to O. pugnax injury. The reproductive phase includes but not limited to the boot (pre-flowering) and heading (flowering) stages. The ripening phase is divided into the milk, soft dough, hard dough, and grain maturity stages (Chaudhary et al. 2002). O. pugnax can cause rice injury from flowering through grain maturation, resulting in direct and indirect losses (Patel et al. 2006). The rice stink bug has piercing-sucking mouthparts which the insect inserts in rice grains and extracts the contents. Damage results in ‘pecky’ rice which is discoloured rice, an empty shell or shriveled kernels. In addition, upon milling, ‘pecky’ rice tends to break so head rice (% whole grain rice after milling) is reduced (Krinski and Foerster 2017).

1.3.5.2 Common Name: Chinch Bug Scientific name: Blissus leucopterus leucopterus Say (Hemiptera: Blissidae)

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Fig. 1.25 Blissus leucopterus adult

Fig. 1.26 Blissus leucopterus life stages

Host Range The chinch bug is a sporadic pest of rice. They are found throughout the United States of America, southern Canada, Mexico, and Central Americas. They feed on plants, both wild and cultivated, belonging to the grass family. Some of the cereal crops damaged are wheat, rye, barley, oats, and corn. Life Cycle Chinch bugs are approximately 4 mm long when fully developed. The adult bodies vary in colour from dark red to brown with white wings and red legs (Fig. 1.25). Young nymphs are usually bright red and half the size of adults. A distinguishable feature is the white band found on the nymph’s abdomen. The wingless nymph is smaller but similar in shape to the adult. The head and thorax are brown, the eyes are dark red, and the abdomen is pale yellow or light red with a black tip. Adults are winged and are black and white. Adult females lay their orange eggs singly in soil cracks or on rice stems. Eggs hatch and nymphs begin feeding on rice stems usually near the soil surface. The insect completes five nymphal instars before becoming an adult (Fig. 1.26). The life span of chinch bug is a typically less than one year. The eggs of two generations are laid down from spring to summer, when they develop into adults. During the fall, the adults from the first generation die off, while the adults from the second generation retreat from the crops to look for overwinter shelters. The adults

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overwinter in any type of shelters. Once they emerge from their hibernation, they fly to fields with small grains growing (such as wheat) and start sucking the sap out of them. They continue to mate while the females begin laying eggs on the lower leaves of plants, or on the roots. The nymphs develop for the next 30 days until they mature into adults. The nymphs undergo six developmental stages, the last being the adult stage. They are wingless, reddish in colour, and progressively become darker with each moult until they have reached their adult stage with their wings fully developed. The nymphs also feed on the same growing plant until they start to ripen. This causes the population to seek out other growing crops. The population of chinch bugs feast and breed on the new growing plant (such as corn), giving birth to a 2nd generation. They continue to feed on the crops, and the 2nd generation of chinch bugs become fully mature. The chinch bugs retreat from the crops to look for shelter for overwinter (Capinera 2008). Two to three generations occur per year. Nature of Damage Chinch bugs feed by piercing the plant and sucking out the plant juices. Chinch bugs are often considered soil insects due to the location of their injury which is at or just below soil level. Damaged plant tissue will often rot due to secondary injury from digestive enzymes and pathogens that are present during feeding. Stunted, yellow, wilting, and dying plants are all symptoms of chinch bug injury. Damaging populations of chinch bugs that develop in grain and forage systems are nearly impossible to control. Conservation tillage systems enhance chinch bug problems in grains. Chinch bugs are typically more of a problem in dry years.

1.3.5.3 Common Name: Planthoppers Sixty-five species of planthoppers associated with rice agroecosystems are reported in tropical Asia (Susilo et al. 2017). Of the total, most notably are the leafhoppers such as Cofana spectra (white rice leafhopper) (Hemiptera: Cicadellidae), Pyrilla perpusilla (sugarcane leafhopper) (Hemiptera: Lophopidae), Nilaparvata lugens (brown planthopper (BPH)), and Sogatella furcifera (whitebacked planthopper) (Homoptera: Delphacidae), the green planthopper (GLH) Nephotettix nigropictus, and Nephotettix virescens (Homoptera: Cicadellidae).

Common Name: White Rice Leaf Hopper Scientific name: Cofana spectra Distant (Hemiptera: Cicadellidae) Host Range The Cofana spectra is common within the rice fields and also on variety of economic grass species nearby rice fields. It is widespread in Africa, Pacific, Australia, and Asian countries like India, Indonesia, Malaysia, Philippines, Sri Lanka, and Taiwan (Meshram and Ramamurthy 2014). Rice, sugarcane, wheat, sorghum, barley, grasses, etc., are preferred host plants.

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Fig. 1.27 Cofana spectra adult

Life Cycle C. spectra is characterized by a large central black spot on the head vertex towards the posterior margin and has brown lines on the forewings. Body length is 7–9 mm. The body is yellowish, the forewings grey-white with prominent veins and the head bears four black spots (Fig. 1.27). It is the largest among the species of leafhoppers and planthoppers. Adults rest on the lower surface of the leaves or at the base of the plant. Females oviposit by making a cut parallel to the long axis of the leaf with their saw-like ovipositors. The eggs are laid in rows of 10–15 across the slit at the base of the plant above the water level. The number of eggs laid per female is about 50 and they hatch in 5–12 days. Nature of Damage The white leafhopper sucks sap from the leaves and results in drying of leaf tips leading the leaf flip orange and curl. The damaged part of paddy is also exposed to secondary infection of fungal and bacterial on the affected part. This species has also been reported as vector of pathogenic viruses, such as rice yellow mottle virus (RYMV) that causes yield loss (Mitra et al. 2014). Common Name: Sugarcane Leaf Hopper Scientific name: Pyrilla perpusilla Walker (Hemiptera: Lophopidae) Host Range Sugarcane leaf hopper is a pest of sugarcane in north-eastern Uttar Pradesh (Butani 1964) but occasionally it also infests paddy and wheat crops. Apart from India, it is distributed in Afghanistan, Bangladesh, Borneo, Myanmar, Cambodia, Indonesia, Java, Nepal, Laos, Pakistan, South China, Sri Lanka, Sumatra, Thailand, and Vietnam (Kumarasinghe and Wratten 1996). Life Cycle The adults are white immediately after moulting but gradually turned strawcoloured, with pale green eyes, snout-like head with black spot positioned posteriorly (Fig. 1.28). The apical area and the outer cleval wing margins have minute black

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Fig. 1.28 Pyrilla perpusilla adult

Fig. 1.29 Eggs of Pyrilla perpusilla

spots. Both the adults and nymphs are very active, jumping from leaf to leaf on slight disturbance. Female measures 10.0 mm length and 2.2 mm breadth, whereas male measures 8.0 mm length and 3.5 mm breadth. Females lay eggs on the lower, shady, and concealed side of the leaves near the midrib in clusters of 30–40 in number in rows of 4–5. They are covered by pale waxy material. Eggs are oval-shaped, pale whitish to bluish-green when laid and turn brown just before hatching (Fig. 1.29). A female lays 600–800 eggs in her lifetime. Nymph passes through five nymphal instar stages to reach adult stage. First instar nymph is greenish white and colour changes in subsequent instars as pale to dark brown and total nymphal period varies between 40–50 days (Ganehiarachchi and Fernando 2006). Nature of Damage Both adult and nymph suck sap from the leaves of rice plant but most of the damage is caused in the nymphal stages. Leaves turn yellowish white and wither away due to heavy infestation. This infection causes great loss to the yield to poor growth of seed sets. The hoppers exude a sweet sticky fluid called ‘honeydew’ which promotes a quick growth of fungus Capnodium sp. Complete coverage of leaves by the sooty mould affects photosynthesis. Common Name: Brown Plant Hopper Scientific name: Nilaparvata lugens Stal (Hemiptera: Delphacidae)

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Fig. 1.30 Nilaparvata lugens adult

Host Range It is distributed in Orissa, Andhra Pradesh, Tamil Nadu, Karnataka, West Bengal, Maharashtra, Madhya Pradesh, Uttar Pradesh, Haryana and Punjab in India, South East Asia, China, Japan, and Korea. Preferred host plants are rice, sugarcane, grasses, etc. Life Cycle Adults are ochraceous brown in colour dorsally and deep brown ventrally. Females are larger (5–6 mm) than the males (4–5 mm). The females are dimorphic—fully winged macropterous and truncated-winged brachypterous (Fig. 1.30). The females are little longer than males. It is dimorphic, with fully winged ‘macropterous’ and truncate-winged ‘brachypterous’ forms. The macropters are potentially migrants and are responsible for colonizing new fields. At the time of colonization, the macropterous forms dominate in a rice field. After settling down on rice plants, they produce the next generation in which most of the female insects develop as brachypters and males as macropters. Copulation takes place just after the emergence and female starts laying eggs by making an incision in the leaf sheath. The eggs, in bunches of 3–10, are thrust within the tissues of the plant along with mid region of the leaf sheath and mid rib of leaves. The average number of eggs laid by a female varies between 200 and 250. The eggs are white, elongated and look like a curved club (Fig. 1.31). They hatch in about 8–10 days. The nymph (Fig. 1.32) undergoes five instars within a period of 15–20 days and the total life cycle is completed in about 20–25 days, which varies in the two sexes. The life span of male is 15–20 days and that of female is 15–30 days (Bae and Pathak 1970).

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Fig. 1.31 Nilaparvata lugens eggs

Fig. 1.32 Nilaparvata lugens nymphs

Nature of Damage The honeydew excreted by the nymphs and adults at the base of the plants is covered with sooty mould. It infests the rice crop at all stages of plant growth. As a result of feeding by both nymphs and adults at the base of the tillers, plants turn yellow and dry up rapidly. At early infestation, round yellow patches appear which soon turn brownish due to the drying up of the plants. This condition is called ‘hopper burn’. The patches of infestation then may spread out and cover the entire field. Crop loss is usually considerable and complete destruction of the crop occurs in severe cases.

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N. lugens is a vector of the virus diseases, e.g., glassy stunt, ragged stunt, and wilted stunt (Chen and Chiu 1981). Common Name: WhiteBacked Plant Hopper Scientific name: Sogatella furcifera Horvath (Hemiptera: Delphacidae) Host Range It is widely distributed in India, Myanmar, Sri Lanka, China, Pakistan, Japan, Indonesia, and Korea. Preferred host plants are rice, maize, millets, sugarcane, and grasses. Life Cycle The adults are yellowish-brown to black, with a typical white spot on the middle of the thorax. They are 2.5–3 mm long and 1.2–1.3 mm broad. Forewings are almost uniformly subhyaline with dark veins. There is a prominent white band between the junctures of the wings. The body is creamy white with the mesonotum and abdomen black dorsally and the legs, ochraceous brown. There is a conspicuous black dot at the middle of the posterior margin of each forewing which meets when the forewings come together (Fig. 1.33). Macropterous males and females and brachypterous females are commonly found in the rice crop, whereas brachypterous males are very rare. The planthoppers, especially adults, prefer to stay at the upper portion of rice stems. After a pre-oviposition period of 3–8 days, the female lays 100–350 eggs into stems or along the midribs of leaves (Fig. 1.34). The nymphs hatch 5–10 days after oviposition and immediately start feeding, preferably at the base of the plant. Young nymphs are creamy-white but as they mature, they develop distinct dark-grey and black markings on their abdomens (Fig. 1.35). During the next 12–18 days, they pass through five stages, each resembling the adult a bit more than the previous one (incomplete metamorphosis). They are very active and instantly jump away (hence Fig. 1.33 Sogatella furcifera adult

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Fig. 1.34 Sogatella furcifera eggs

Fig. 1.35 Sogotella frucifera nymph

the name) when disturbed. Adult male was short lived 14.4 days as compared to female (15.9) days (Kumar et al. 2015). Nature of Damage Both nymphs and adults suck the sap and cause stunted growth and ‘hopper burn’ leading to yield loss. ‘Hopper burn’ is caused in irregular patches, which leads to yellowing of the leaves, reduced vigour, and stunting of plants. Nymph falls on water keeping its legs stretched. The number of grains and the panicle length decrease if the infestation is at the panicle initiation stage. But when attacked later, during the maturation period, grains do not fill fully and ripening is delayed. When the hoppers are present in large numbers late in the crop growth stage, they are seen infesting the flag leaves and panicles. Gravid females cause additional damage by making oviposition punctures in leaf sheaths (Sharma et al. 2018). Feeding points and wounds caused by egg laying may later become potential sites for the invasion of bacteria and fungi.

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Moreover, the honeydew produced by the hoppers serves as a medium for mould growth.

Common Name: Green Leafhopper Scientific name: Nephotettix virescens Distant (Hemiptera: Cicadellidae) Host Range It is distributed all over the rice growing tract of India and sporadically occurs in an epidemic form, causing considerable damage to rice crop. Life Cycle The adult insects are small and bright green in colour (Fig. 1.36). Nymphs are yellowish-green in colour (Fig. 1.37). The female inserts the eggs in rows under the epidermis of leaf sheath and may lay up to 40–50 eggs. The egg and nymphal periods are 6–7 and 18 days, respectively. The nymph undergoes five successive moults between a period of 15–22 days. The insect completes its life cycle in about 20–25 days, depending upon the ecological conditions. The life span of adult insect Fig. 1.36 Nephotettix virescens adult

Fig. 1.37 Nephotettix virescens nymph

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is about 30 days. Under favourable conditions, there are about 10 generations per year. Nature of Damage Both adults and nymphs feed on the culms by sucking sap. The infected plant turns yellow and dies. The first symptoms of infestation are yellow, transparent spots that appear mainly on the tips and along the midribs of the leaves. These are soon followed by greyish-white, and later brown spots on the leaves and leaf sheaths of the young plants. Deposits of honeydew gradually cover the plant surface, creating a basis for the development of sooty moulds. The soft, whitish exuviae can often be found adhering to the sticky surface of the leaves. As a rule, the damage first becomes noticeable around the edges of the paddy field. The insects tend to prefer the peripheral leaves of a plant over the central ones, and are found mainly on the dorsal leaf surface. It is also a vector of virus diseases of rice, such as rice transitory yellowing and rice yellow dwarf. They are more active during July to September in north-eastern Uttar Pradesh.

1.3.5.4 Common Name: Rice Delphacid Complete Scientific name: Tagosodes orizicolus Muir (Hemiptera: Delphacidae) Host Range The planthopper, T. orizicolus (known previously as Sogata orizicola), is one of the most important pests of rice throughout rice growing areas of tropical America including Mexico, the Caribbean, and Central and South America (Morales and Niesses 1985). Host plants include rice, barnyardgrass (Echinochloa spp.) and other Poaceae (Asche and Wilson 1990). Life Cycle The female adult (Fig. 1.38) lays tiny, white eggs (~160 in a lifetime) in the midribs of rice leaves (ARS 1960). Nymphs (Fig. 1.39) emerge from eggs in 8–11 days and start feeding on leaf blades. Nymphs develop through five instars (3 days each) and into adults that are brown and about 0.4 cm long. Adults include both short and long winged forms and some wingless females. The average life span of an adult male is 25 days and that of a female is 34 days. Nature of Damage It is a phloem feeder, with piercing-sucking mouthparts damages a rice plant by feeding and ovipositing. A symptom commonly associated with feeding by T. orizicolus is yellow to reddish discolouration called ‘hopper burn’ of young leaves. It is also an important vector of tenuivirus Rhbv that causes hoja blanca disease [Rice hoja blanca virus (RHBV)], meaning ‘white leaf rice virus’, a plant virus in the family Phenuiviridae. This virus affects the leaves of the rice plant, stunting growth of the plant or killing it altogether (Asche and Wilson 1990).

40 Fig. 1.38 Tagosodes orizicolus adult

Fig. 1.39 Tagosodes orizicolus nymph

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1.3.5.5 Common Name: Rice Black Bug Scientific name: Scotinophora lurida (Burmeister), S. coarctata (Hemiptera: Podopidae) Host Range The rice black bug (RBB) occurs in Asian countries, such as South China, Vietnam, Brunei, Indonesia, Malaysia, Cambodia, Sri Lanka, Thailand, Myanmar, India, Bangladesh, and Pakistan (Reissig et al. 1986). Preferred host plants are rice and millet. Life Cycle Adults are brownish-black, oval-shaped, and about 8–9 mm long with a prominent scutellum and pronotum having a spine on either side (Fig. 1.40). It lives from 3 to 7 months. The female lays about 200 greenish eggs (1 mm long) during her lifetime and guards the egg until hatching (Reissig et al. 1986). It deposits its eggs on the lower part of the leaves or on the basal part of the rice plant near the water surface. The eggs are laid in masses of 40–60 individual eggs in several parallel rows. During dry conditions, the female bug deposits its eggs on the leaves and stem. Eggs are also laid in cracks on the soil and on roots. Eggs turn pinkish during hatching (Fig. 1.41). Brown nymphs have yellowish-green abdomen and 2–3 black scent glands (Fig. 1.41). Egg incubation of RBB is 3–7 days. The nymphs are brown and yellow with black markings (Reissig et al. 1986). Six nymphal instars are completed in 29–35 days.

Fig. 1.40 Scotinophora lurida adult

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Fig. 1.41 Scotinophora lurida adult, egg mass and nymph

Fig. 1.42 Chlorotic lesions on rice leaves due to Scotinophora lurida damage

Nature of Damage Both nymphs and adults suck plant sap from the culm during tillering to flowering at the base of the plant. It also sucks the sap from leaf sheath, leaf, and panicle. The affected plants turn reddish brown or yellow. During tillering stage, it causes drying up of central shoot (dead heart), stunted growth, and reduced tillers. During reproductive stage, it affects the panicle development and causes chaffy grains (white ears). In severe cases, plants wilt, dry, and turn bug burned, similar to hopper burn damage of brown plant hopper. The nymphs are destructive as it can feed at the basal part of the rice crop for up to 42 days. The insects prefer stem nodes or the base of the stem as feeding sites because of the large sap reservoirs (Reissig et al. 1986). Feeding by large number of bugs can cause plants to be stunted and the leaves turn reddish brown, a condition called ‘bug burn’ (Fig. 1.42).

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1.3.5.6 Common Name: Earhead Bug Scientific name: Leptocorisa oratorius F. (Hemiptera: Alydidae) Host Range It is widespread in nature and mainly recorded from Asia, Central America, and Australia, etc. Rice and grasses are the main hosts. It is also recorded on beans, breadfruit, guava, mango, millet, and more, as a minor pest. Life Cycle The adults are about 15 mm long, brown above, whitish-green below, with long legs and antennae (Fig. 1.43). Female lays 200–300 flat, dark, reddish brown eggs 10–20, in 2–3 rows on the leaves or panicles (Fig. 1.44). The eggs hatch into green-coloured nymphs, which gradually turn brown as they grow into adults. There are five nymphal instars with a total nymphal period of 13–17 days. Nymphs are wingless. The life cycle takes about 30 days. Nature of Damage Both nymphs and adults suck the sap from individual grains at milky stage. Affected grains become chaffy with black spots at the site of feeding puncture. Yield loss may be 10–40%. Obnoxious odour emanates on disturbing the bugs in the field. They Fig. 1.43 Leptocorisa oratorius adult

Fig. 1.44 Leptocorisa oratorius eggs

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spoil the seeds by sucking out their contents when they are still soft, i.e., from flowering to the dough stage. Microorganisms enter the damaged seed and cause them to dry out and discolour. Infestations of the rice bug are highest at the beginning of the rainy season.

1.3.6

Sorghum: Sorghum bicolour L. Moench

Crop Importance Sorghum is one of the important cereal crops of the world. Sorghum ranks fourth among the world cereals in the order of wheat, maize, and rice. It is the major source of food and fodder for millions of people in tropics and semi-arid tropics (Mundia et al. 2019). The stem and foliage are used as green fodder, hay silage, and pasture apart from being used as fuel and building material. Sweet sorghum is used in the preparation of jaggery, syrup, ethanol (vehicle fuel), and biscuits. Beer is prepared from sweet sorghum in many parts of Africa. Besides these products, popped and sweet sorghum are also very popular all over the world (House 1980). Details about some of the major sucking insects damaging sorgum are given below.

1.3.6.1 Common Name: Green Bug Scientific name: Schizaphis graminum (Rondani) (Hemiptera: Aphididae) For details please see Sect. 1.3.1.3. 1.3.6.2 Common Name: Corn Leaf Aphid Scientific name: Rhopalosiphum maidis Fitch (Hemiptera: Aphididae) For details please see Sect. 1.3.1.5. 1.3.6.3 Common Name: Chinch Bug Scientific name: Blissus leucopterus leucopterus Say (Hemiptera: Blissidae) For details please see Sect. 1.3.5.2. 1.3.6.4 Common Name: Yellow Sugarcane Aphids Scientific name: Sipha flava Forbes (Hemiptera: Aphididae) Host Range It was first described as Chaitophorus flavus Forbes (Forbes 1884) and later moved to Sipha flava (Forbes) (Davis 1909). It has an extensive geographic range, which encompasses South, Central, and North America, the Caribbean, Puerto Rico, and Hawaii (Blackman and Eastop 2000). It feeds on many plant species, which incorporate several diverse families, such as Cyperaceae, Poaceae, and Commelinaceae (Blackman and Eastop 2000). Economically important crops that are fed on by this aphid include sorghum, sugarcane, wheat, barley, rye, and other cereal crops, as well as many species of pasture grasses (Blackman and Eastop 2000).

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Fig. 1.45 Sipha flava aphid

Fig. 1.46 Winged and wingless Sipha flava aphids

Life Cycle Yellow sugarcane aphids are usually lemon yellow but sometimes pale green. They are covered with small spines and have two double rows of dark spots on the back. The cornicles (tail-pipes at the end of the abdomen) are reduced to slightly elevated pores (Fig. 1.45). There are winged and wingless forms (Fig. 1.46). Sipha flava reproduces without mating (i.e., parthenogeneticly) in warm climates and produce live young ones. Females mate with wingless males in areas with cold winters. Nymphs go through four instars before moulting directly into the adult stage (i.e., no pupal stage). Development from nymph to reproducing adult takes about 8 days on S. bicolor (Hentz and Nuessly 2004), but 18–22 days on sugarcane.

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Females produce one to five nymphs per day for about 22 days on average on maize and sugarcane. Adult females can be separated from large third instar nymphs by the presence of a small, rounded, knobby cauda adorned with several hairs at the end of the adult abdomen. Winged and wingless adults are 1.3–2.0 mm long. Winged females have yellow abdomens with spots and darker head and thorax than wingless females (Fig. 1.46). Nature of Damage They feed on sorghum and inject toxin into leaves of seedlings and older plants. Aphid feeding on seedlings turns the leaves purple and stunts their growth. Damage often leads to delayed maturity and plant lodging, which may be increased by associated stalk rots. These aphids prefer to feed on the lower surfaces of leaves. Honeydew produced by the feeding aphids collects on lower leaves and supports growth of sooty mould fungi.

1.3.7

Wheat: Triticum sp.

Crop Importance Wheat is a cereal grass of the genus Triticum (Family, Poaceae). It is one of the oldest and most important cereal crops. Of the thousands of varieties known, the most important are common wheat (Triticum aestivum), used to make bread; durum wheat (T. durum), used in making pasta (alimentary pastes) such as spaghetti and macaroni; and club wheat (T. compactum), a softer type, used for cake, crackers, cookies, pastries, and flours. Additionally, some wheat is used by industry for the production of starch, paste, malt, dextrose, gluten, alcohol, and other products (Shewry and Hey 2015). After rice, wheat is the second most important food crop grown in India and it is the staple diet for millions of Indians, mainly in the north and north-western parts of the country. Wheat is the main source of protein, carbohydrate, and vitamins for people in the rural areas. Some of the sucking insect-pests damaging wheat are described below.

1.3.7.1 Common Name: Russian Wheat Aphid (RWA) Scientific name: Diuraphis noxia (Hemiptera: Aphididae) For detail please see Sect. 1.3.1.1 1.3.7.2 Common name: English Grain Aphid Scientific name: Macrosiphum (Sitobion) avenae F. (Hemiptera: Aphididae) For detail please see Sect. 1.3.1.2

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1.3.7.3 Common Name: Green Bug Scientific name: Schizaphis graminum (Rondani) (Hemiptera: Aphididae) For detail please see Sect. 1.3.1.3 1.3.7.4 Common Name: Corn Leaf Aphid Scientific name: Rhopalosiphum maidis Fitch (Hemiptera: Aphididae) For detail please see Sect. 1.3.1.5 1.3.7.5 Common Name: The Bird Cherry-Oat Aphid Scientific name: Rhopalosiphum padi Linnaeus (Hemiptera: Aphididae) For detail please see Sect. 1.3.1.6

1.4

Integrated Pest Management Approaches for Sucking Insect-Pests of Cereals

Sucking pests pierce plants with slender, sharp-pointed mouthparts and suck the plant sap. Withdrawal of the sap results in minute white, brown, or red spotting on the leaves, fruits, or stems of the plant. It may also cause curling leaves, deformed grains, general wilting, browning, and drying of the entire plant. Many of these pests are also resistant to pesticides. Some of these pests are important virus vectors, transmitting a range of plant viruses. Virus-infected plants cannot be cured. However, control measures can be used to prevent or reduce the levels of the disease in crops by removing, or avoiding the sources of virus infection, and minimizing spread by these sucking pests. Sucking pests include aphids, leafhoppers, thrips, whitefly, flies, bugs, and mites. Some of the management methods are described below. Cultural Methods Agronomic adjustments and modifying the crop microclimate are the main objective. Apart from these, fertilizer application must be done judiciously. High levels of nitrogen promote succulent, nutritious new growth, which is preferred by aphids and can help boost aphid reproduction. Over fertilizing a plant can enhance aphid population growth and make the problem worse. Using smaller amounts of fertilizer throughout the growing season can help to reduce potential aphid outbreaks. Trap Cropping Trap cropping is the planting of a trap crop to protect the main cash crop from a certain pest or several pests. The trap crop can be from the same or of different family group, than that of the main crop, as long as it is more attractive to the pest. Companion Plants A potential strategy for controlling pests is through the use of ‘companion plants’ within a crop system. A companion plant may be associated with a target crop for various reasons. Firstly, it can attract aphids and draw them away from their host

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plants. Secondly, it can alter the recognition of the host plant. This effect is mostly attributed to companion plant volatiles since they disturb the sucking insect host plant location, and additionally they may react chemically and physiologically with the host plant, making it an unsuitable host for insects (Ben-Issa et al. 2017). Aphid Banker Plants Banker plants are mobile habitats that provide alternative hosts, prey, or food for commercially available natural enemies. As a biological control strategy, banker plants offer a novel nonchemical approach to managing commonly encountered pests in the greenhouse. Most banker plants that target aphids consist of a graminaceous plant, a non-pest cereal grain aphid, and a parasitoid that attacks both the non-pest and pest aphids occurring on crop plants. The use of banker plants may provide more effective, long-term pest control than pesticide applications, but both may be combined effectively. Among the various species of banker plants, Aphidius colemani (Viereck) (Hymenoptera: Braconidae)—Rhopalosiphum padi (L.) (Hemiptera: Aphididae) system is best to manage aphid pests (Payton Miller and Rebek 2018). Bio Intensive Pest Management (BIPM) The pest management programme, where natural enemies of the crop pests form the core component is designated as Bio Intensive Pest Management BIPM. The most commonly used bio-agents in BIPM are broadly classified into three categories: (a) Parasitoids (b) Predators, and (c) Microbes. Sucking insects, especially aphids have several natural enemies that can be attracted or released to help keep populations in check. The most common one is ladybird beetles. Ladybird beetles and their larvae feed on different types of aphids. It is best to use plants that will attract ladybird beetles, such as sunflowers, clovers, and coreopsis. The convergent ladybird beetle (Hippodamia convergens) and the two-spotted lady beetle, Adalia bipunctata, are commercially available from many biological control suppliers. Another effective natural enemy is green lacewing larvae (Chrysoperla rufilabris). Their larvae are extremely aggressive and eat numerous aphids a day. Lacewings are extremely sensitive to insecticides and even drift from an application can be harmful. The green lacewing (Chrysopa rufilabiris and C. carnea) adults are active at night and feed on nectar, pollen, and honeydew. The predatory larvae feed upon different species of aphids as well as mites, whiteflies, mealybugs, scales, and thrips. Because larvae will feed upon each other, they must be released as far apart as possible to discourage cannibalism. Green lacewings may be less effective on plants with hairy leaves. Green lacewings are commercially available as eggs on cards, or as larvae shipped with a food source in an inert material in a small container or as larvae shipped in separate cells. Larvae may survive better than eggs and are quicker acting. Green lacewings are also available as adults and shipped in a small cardboard container.

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The predatory midge, Aphidoletes aphidimyza, can feed on more than 60 different species of aphids. This gall midge is nocturnal, and prefers dark and humid areas near the lower plant canopy. They require a period of darkness for mating and egg laying. Only the larvae stage is predaceous. Adults feed primarily upon pollen and honeydew. The bright orange larva kills aphids by biting their knee joints, injecting a paralyzing toxin and then sucking out their body fluids. Aphidoletes aphidimyza is typically sold as pupae in bottles or blister packs Another natural enemy is parasitic wasps (Aphidius species) that sting aphids and impregnate them with an egg. The egg then grows inside the aphid, killing and mummifying it, and a new adult wasp hatches out of the mummified aphid. If these mummified aphids are seen near active aphid populations, it indicates that the Aphidius wasps are nearby and actively parasitizing the current population. Another natural enemy are entomopathogenic fungus, Beauveria bassiana. This fungus is usually applied as a foliar spray and is parasitic to many soft body insects. The organism is available in both liquid or powder form, but the powder is more stable and has a longer shelf life. This can be used for control of aphids or other soft body insect. Apply the product as a preventative every 7–14 days to help keep pest populations low.

1.4.1

Chemical Control

When all other control measures failed to keep the populations under control, a chemical insecticide may be needed. The goal with insecticide use is to choose the chemical with minimal impact to pollinators and natural enemies, but chemical that is still effective on the insect causing the problem. The first effective choice to spray would be either insecticidal soap or horticultural oil. These insecticidal products coat the aphid’s exoskeleton (body) and cause it to suffocate. These insecticides can also kill beneficial insects upon contact, but they have no residual activity. So only beneficial insects and pollinators that were directly hit by the application will be affected. Pollinators and natural enemies that arrive after the spray solution has dried will not be impacted by these soaps or oils. Applications should be made when temperatures are cooler, such as the mid- to late evening to avoid any potential plant damage. Another effective botanically derived chemical is azadirachtin. This compound is a natural insect growth regulator that modifies the way insects grow by inhibiting the shedding of the exoskeleton. It can be mixed with an entomopathogenic fungi or bacteria to allow more contact time between the insect’s exoskeleton and the pathogenic organism. This ensures that the fungi or bacteria have time to grow, penetrate the exoskeleton, and kill the insect. Among the synthetic insecticides, following may be used for management of sucking insect-pests. Active ingredient

Concentration (%)

Trade name

Method of use (continued)

50

Dimethoate Phosphamidon Profenophos Lamda-Cyhalothrin 5% EC

Omkar and A. K. Tripathi

0.05 0.05 0.01 0.05

Rogor 30 EC Dimecron Kareena 50 EC Karat

1.5 mL/L water 2.0 mL/L water 1.0 mL/L water 0.75 mL/L water

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Porter DR, Burd JD, Shufran KA, Webster JA, Teetes GL (1997) Greenbug (Homoptera: Aphididae) biotypes: selected by resistant cultivars or preadapted opportunists? J Econ Entomol 90(5):1055–1065 Punnuri SM, Huang Y, Steets J, Wu Y (2013) Developing new markers and QTL mapping for greenbug resistance in sorghum (Sorghum bicolor (L.) Moench). Euphytica 191:191–203 Quimio GM, Calilung VJ (1993) Survey of flying viruliferous aphid species and population buildup of Aphis glycines Matsumura in soybean fields. Philipp Entomol 9:52–100 Reissig WH, Heinrichs EA, Litsinger JA, Moody K, Fiedler L, Mew TW, Barrion AT (1986) Illustrated guide to integrated pest management in rice in tropical Asia. International Rice Research Institute, Los Banos Saleh N, Middleton KJ, Balladi Y, Horn N, Reddy DVR (1989) Research on peanut stripe virus in Indonesia. In: Proc. 2nd coordinators’ Mtg. on peanut stripe virus, ICRISAT, Patancheru, Andhra Pradesh, India, p 9 Sharma AR, Dass A (2012) Maize. Textbook of field crops production, food grain crops, vol 1. Directorate of Knowledge Management in Agriculture, Indian Council of Agricultural Research, New Delhi, p 98 Sharma S, Kooner R, Arora R (2017) Insect pests and crop losses. Springer, Singapore, p 45 Sharma KR, Raju SVS, Jaiswal DK (2018) Influence of environmental effect on the population dynamics of brown plant hopper, Nilaparvata lugens (Stal) and White-Backed plant hopper, Sogatella furcifera (Hovarth) in Varanasi region. J Entomol Res 42(3):339–342 Shehata HFH, Abdel-Rahman MAA, Mahmoud AMA, Ahmed MHA (2018) Temperature effects on the development, survival and reproductive potential of the Greenburg aphid, Schizaphis graminum (Rondani) (Homoptera: Aphididae). J Entomol Zool Stud 6:211–215 Shewry PR, Hey SJ (2015) The contribution of wheat to human diet and health. Food Energy Secur 4(3):178–202 Singh C, Singh R, Singh P (2012) Maize. Modern techniques of raising field crops. Oxford/IBH Publishing Company Pvt. Ltd, New Delhi, pp 84–111 Stern VM (1967) Control of the aphids attacking barley and analysis of yield increases streak and Diuraphis noxia in Mexico. In: Burnett PA (ed) Barley yellow dwarf, a proceedings of a workshop. CIMMYT, Mexico, pp 157–163 Susilo FX, Swibawa IG, Indriyati H, Purnomo AM, Hasibuan R, Wibowo L, Suharjo R, Fitriana Y, Dirmawati SR, Solikhin S, Ruruh Anjar Rwandini RA, Dad Resiworo Sembodo DR, Suputa A (2017) The white bellied planthopper (Hemiptera: Delphacidae) infesting corn plants in south Lampung Indonesia. Jurnal Hama dan Penyakit Tumbuhan Tropika 17:96–103 Tokuda M, Jikumaru Y, Matsukura K, Takebayashi Y, Kumashiro S, Matsumura M, Kamiya Y (2013) Phytohormones related to host plant manipulation by a gall-inducing leafhopper. PLoS ONE 8:e62350 Tolmay VL (2006) Genetic variability for Russian wheat aphid, Diuraphis noxia resistance in South African wheat genotypes. PhD thesis. Faculty of Natural and Agricultural Science. Department of Plant Sciences. University of the Free State van Emden A, Fritz H, Harrington R (2007) Aphids as crop pests. CABI, London, p 19 Volkl W, Stechmann DH (1998) Parasitism of the black aphid (Aphis fabae) by Lysiphlebus fabarum (Hym., Aphidiidae): the influence of host plant and habitat. J Appl Entomol 122:201–206 Wangai A, Plumb R, Forde S, Herts H (1991) Incidence of barley yellow dwarf virus in Kenya. Regional Wheat Workshop: For Eastern, Central and Southern Africa, Nakuru (Kenya), pp 16–19 Zadoks JC, Schein RD (1979) Epidemiology and plant disease management. Oxford University Press, New York

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Sucking Pests of Rice Swoyam Singh and S. N. Tiwari

Contents 2.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Major Insect Pests of Rice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Management of Sucking Pests . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Management of Rice bugs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Management of Thrips . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 Management of Mites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

Rice is one of the major staple foods across the globe. It is known to suffer losses of up to 200 million tonnes globally owing to various abiotic and biotic factors. The biotic factors, which are largely insect pests, contribute to around 35.55 million tonnes of rice being destroyed every year. Around 800 species of insects are known to infest rice under field and storage conditions. In this chapter an attempt has been made to summarize the life cycle, nature of damage and management of the most important sucking pests causing economic loss in rice ecosystem.

2.1

Introduction

Rice is the most important cereal which is cultivated at large scale in several countries including India, China, Indonesia, Bangladesh, Thailand, Vietnam, Myanmar, the Philippines, Cambodia and Pakistan (Prasad et al. 2017). In all S. Singh · S. N. Tiwari (*) Department of Entomology, G.B. Pant University of Agriculture and Technology, Pantnagar, Uttarakhand, India # Springer Nature Singapore Pte Ltd. 2020 Omkar (ed.), Sucking Pests of Crops, https://doi.org/10.1007/978-981-15-6149-8_2

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these countries several abiotic and biotic factors are known to cause an annual loss of more than 200 million tonnes (Khan et al. 1991a, b). Among the biotic factors, insect pests are known to cause major losses of 35.55 million tonnes which may amount up to $8467.36 million in India itself (Dhaliwal et al. 2015). According to various estimates approximately 800 species of insects are known to infest rice under field and storage conditions (Ul Ane and Hussain 2015; Heinrichs and Muniappan 2017). Insects belonging to several order such as Lepidoptera, Coleoptera, Orthoptera, Diptera, Isoptera are known to feed rice on various parts of it; however, only Hemipterous and Thysonapterous insect feed on sap from aerial parts leading to a loss of several million tonnes globally (Cheng et al. 2003; Kumar et al. 2012; Bisen et al. 2019). The extent of loss is known to vary from region to region and species to species in different countries. Brar et al. (2010) estimated a yield loss of 2.7 million tonnes due to direct feeding damage by only brown planthopper which caused an additional loss of 0.4 million tonnes by transmitting viruses. In this chapter an attempt has been made to summarize the life cycle, nature of damage and management of the most important sucking pests causing economic loss in rice ecosystem.

2.2

Major Insect Pests of Rice

2.2.1

Planthoppers

2.2.1.1 Brown Planthopper, Nilaparvata lugens (Stål) (Delphacidae: Hemiptera) The brown planthopper (BPH), Nilaparvata lugens (Stål), has been recognized as a key pest of rice in so many countries (Bisen et al. 2019). This insect is distributed mainly in Asia, Australia and Pacific Islands (Fig. 2.1) while it is not found in America and Africa. In Australia and the Pacific Islands, it is found on the Caroline

Fig. 2.1 Distribution of Nilaparvata lugens (Stål) in the world (source: https://www.cabi.org/isc/ datasheet)

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Islands, Fiji, Mariana Islands, Papua New Guinea and Solomon Islands (CAB 1984). In Asia the pest has been reported especially from the southeast part of the continent which includes Bangladesh, Brunei, Myanmar, China, Hong Kong, India, Indonesia, Japan Korea, Laos, Malaysia, Nepal, Pakistan, Philippines, Singapore, Sri Lanka, Taiwan, Thailand and Vietnam. The pest is found throughout the year in all parts of Asia except for Japan and Korea where it migrates from the East China Sea during the beginning of summer every year. In India it was initially reported as a sporadic pest causing non-economic damage over years until 1973–1974 when its epidemic caused serious loss of worth ($12 million) in “Kole” lands of Trichur district and Kuttanad area in Kottayam and Alleppey districts of Kerala (Koya 1974; Nalinakumari and Mammen 1975). Subsequently, its outbreak was reported in other states including Andhra Pradesh (Ghose et al. 1960), Tamil Nadu, Karnataka, Madhya Pradesh, Odisha and West Bengal. These states harbour the insect all-round the year, from where it is thought to migrate to the northern states of India, viz. Uttar Pradesh, Punjab, Himachal Pradesh, Haryana and Rajasthan (Krishnaiah 2014) (Table 2.1).

Identification of Brown Planthopper (BPH) Eggs The eggs (0.99 mm long and 0.20 mm wide) are cylindrical in shape, white or transparent in colour, craved into the leaf sheath (Fig. 2.2a) and are not visible to naked eyes. The eggs are laid in a group of 2–20 marking a brown excavation in the leaf sheath or blade (Khaire and Dumbre 1981). Nymphs First instar nymphs are 0.97 mm long and 0.37 mm wide. The head is triangular with a narrow vertex while compound eyes are small and brownish with a black centre and long vertex with demarcation between the thorax. The abdominal segments are not distinct. Legs are light-brown with a movable spur on hind tarsus. The body is creamy white with a pale brown tinge. Second instar nymphs are 1.29 mm long and 0.53 mm wide. The eyes are slightly red. The thoracic area is narrower than the abdomen. The abdomen is uniformly brownish-white and slightly swollen. The legs are brown; the body is oval and brownish-white in colour. Third instar nymphs are 1.42 mm long and 0.67 mm wide. They are oval and brownish in colour. The eyes are dull red. There is a longitudinal midline from the base of the vertex to the tip of meta-thorax, wings are rudimentary but gradually becomes apparent. Fourth instar nymphs are 1.99 mm long and 1.00 mm wide. The compound eyes are bulged and dull red in colour. The longitudinal midline runs from the base of vertex to the end of meta-thorax, wing rudiments cover the first two abdominal segments and partially cover the third. The abdomen is swollen with irregular brown patches which are clearly seen from the first to the seventh segments (Fig. 2.2b).

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Table 2.1 Sucking pests of rice in different agroecosystem Order Hemiptera

Family Blissidae Cicadellidae

Pentatomidae

Cercopoidea Delphacidae

Pseudococcidae

Alydidae

Common name Southern chinch bug Chinch bug Maize leafhopper White leafhopper White leafhopper Sharp-headed leafhopper Common green sugarcane leafhopper Rice green leafhopper Rice green jassid Green leafhopper (GLH) Zigzag leafhopper Orange rice leafhopper White-spotted spined bug Stink bug Stink bug Stink bug Green shield bug Island stink bug Rice stink bug Small rice stink bug Rice stink bug Paddy bug Stink bug Subterraneous stink bug Subterraneous stink bug Black rice bug Black rice bug Brown stem bug Stink bug Yellow-sided froghopper Brown planthopper (BPH) Corn planthopper White-backed planthopper Rice planthopper Rice planthopper Rice mealy bug Grey sugarcane mealy bug Rice ear head bug Slender rice bug Bean bug

Scientific name Blissus insularis Blissus leucopterus Cicadulina mbila Cofana spectra Cofana unimaculata Draeculacephala clypeata Hortensia similis Nephotettix cincticeps Nephotettix nigropictus Nephotettix virescens Recilia dorsalis Thaia oryzivora Eysarcoris parvus Mormidea guerini Mormidea pictiventris Mormidea ypsilon Nezara viridula Oebalus insularis Oebalus ornata Oebalus poecilus Oebalus pugnax Oebalus ypsiloinoides Piezodorus guildinii Scaptocoris castaneus Scaptocoris castaneus Scotinophara coarctata Scotinophara lurida Thyanta perditor Tibraca limbativentris Aeneolamia flavilatera Nilaparvata lugens Peregrinus maidis Sogatella furcifera Sogatella vibix Tagosodes orizicolus Brevennia rehi Saccharicoccus sacchari Leptocorisa acuta Leptocorisa oratorius Riptortus clavatus (continued)

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Table 2.1 (continued) Order

Family Aphididae

Common name Grain aphid Green corn aphid Rice root aphid Yellow sugarcane aphid English grain aphid Cereal root aphid

Coreidae

Thysanoptera

Lophopidae Meenoplidae Thripidae

Grain aphid Coreid bug Black bug Black bug Sugarcane planthopper Grey planthopper Grass thrips Rice leaf thrips Melon thrips Taiwan flower thrips Cereal thrips

Scientific name Hysteroneura setariae Rhopalosiphum maidis Tetraneura nigriabdominalis Sipha flava Sitobion avenae Rhopalosiphum rufiabdominale Rhopalosiphum padi Leptoglossus gonagra Phthia obscura Phthia picta Pyrilla perpusilla Nisia nervosa Anaphothrips obscurus Stenchaetothrips biformis Thrips palmi Frankliniella intonsa Haplothrips aculeatus

Source: Bhatt et al. (2018)

Fig. 2.2 Different stages of N. lugens: (a) eggs, (b) nymph, (c) adult female, (d) adult male

Fifth instar nymphs are on an average 2.69 mm long and 1.25 mm wide. The entire body is dark-brown and robust. The eyes are greyish-blue. The wing rudiments fully cover the first three abdominal segments (Kaur 2011). Adults are ochraceous brown in colour dorsally and deep brown ventrally. Females are larger (5–6 mm) than the males (4–5 mm) (Fig. 2.4d). The females are dimorphic—fully winged macropterous and truncated-winged brachypterous. The wings are subhyaline with a dull yellowish tint (Fig. 2.2c).

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Life Cycle Eggs N. lugens lays cylindrical eggs in groups protruding out of the leaf sheath or leaf blades of the rice plant. The insect makes an incision with the help of its ovipositor and inserts the eggs into the parenchymatous leaf tissue (Hattori and Sogawa 2002). The eggs are laid in longitudinal rows in such a way that only the opercular region is visible from outside. The number as well as the size of the eggs is reported to be less when laid in rice seedlings than in leaf blades or leaf sheath. One female may lay 124 egg masses each consisting of 2–11 eggs (Panwar 1995). The brachypterous form (B-form) lays more eggs (300–700 eggs) as compared to macropterous form (100 eggs) (Manjunath 1977). The freshly laid eggs are white in colour which gradually turns darker with two distinct spots. The eggs are not visible to naked eyes; however, a dirty brown patch in the oviposition site marks the laying of egg. The site and size of egg depend on the stage of the plant where the younger plants are reported to accommodate more eggs on the leaf sheath (Mochida 1964). The incubation period is much influenced by the changing climatic parameters. The incubation period is 6–8 days and 11–14 days during June–October and November– January, respectively (Misra 1980; Mallikarjun et al. 2019). The egg laying may be done in the range of 25–30  C temperature. Maximum eggs are laid at 27  C, while 33  C is lethal for it. Nymphs There are five nymphal stadia and four moultings in case of BPH which takes 10–18 days (Fig. 2.3) to become adults. Brachypterous insects have a shorter nymphal period for both the sexes as compared to the macropterous form (Kisimoto 1956). The nymphal instars are whitish brown in colour having a sac like abdomen and can be distinguished only by the shape of mesonotum, metanotum and body size. Temperature, availability of food and the density of the insect are the important factors which govern the duration of nymphal period. The temperature at 25  C is most favourable for nymphal stage, while 31  C or more proves lethal to the initial nymphal instars (Bae 1966). Adults The adults are light to dark in colour and are categorized into two distinct forms, viz. macropterous (long-winged) and brachypterous (short-winged) adults. The process of formation of both forms is governed by the availability of food and the density of the population. High population density and limited food may lead to the development of macropterous form of N. lugens so that they could transport themselves to a new host habitat in the vicinity especially during morning hours (Mathur and Chaturvedi 1980). On the other hand low population and abundance of food result in the development of brachypterous form which has comparatively larger body, legs, ovipositors and higher fecundity. Immediately after emergence of adults, mating takes place between males and females for almost 24 h. The average mating period of an individual male varies

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Fig. 2.3 Life cycle of N. lugens

from 84.20  18.90 s but males are polygamous who can mate with a maximum of nine females for 24 h, whereas an individual female can copulate twice during its lifetime (Mochida and Okada 1979). There are specific acoustic signals of the female abdominal vibrations which attract the male for mating (Ishii and Ichikawa 1975). The mating leads to a gap of 4–6 days prior to laying of eggs which is termed as pre-oviposition period (Krishnaiah 2014). The longevity of adult females (14–30 days) is generally longer than the adult males (14–21 days) (Nalinakumari and Mammen 1975; Mallikarjun et al. 2019). There may be eight overlapping generations of N. lugens in a year where the time taken to complete a single generation ranges from 23 to 106 days depending upon the temperature, relative humidity and other environmental factors (Kaur 2011). The duration of generations occurring in a single year has been depicted in Table 2.2. Nature of Damage The brown planthopper is basically a phloem feeder but also feeds occasional on xylem. Phloem is more preferred since it is nutritionally rich with nutrients like carbohydrates, amino acids and peptides whereas occasional feeding in xylem occurs to satisfy the requirement of water and mineral like nitrates, potash, phosphates, calcium, magnesium, zinc, etc. (Krishnaiah 2014). The insects have piercing and sucking type of mouthparts, the piercing stylet being the most important organ for penetration through the plant cell. The stylet secretes saliva which solidifies, acts as a tool for penetration and penetrates through each and every cell that comes in the way while reaching the phloem tube (from epidermis to the phloem tube). While penetrating the cells it leads to the death of all affected cells. Furthermore, once it reaches the phloem tube, it sucks the sap and disrupts the activity of phloem, thereby causing a typical “Hopperburn” symptom (Spiller 1990).

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Table 2.2 Duration of generation (in days) occurring in a single year in N. lugens Generation 1st 2nd 3rd 4th 5th 6th 7th 8th

Period July–August August–September September–October October–Mid-November Mid-November–March March–Mid-April Mid-April–June June–July

Duration of generation (days) 23 27 32 49 106 52 46 29

Source: Kaur (2011)

Fig. 2.4 Hopperburn in rice field

The initial symptoms of N. lugens feeding cause yellowing of leaf blade which gradually spreads to other above ground part of the plant. When the chlorosis or yellowing reaches the flag leaf, grain filling ceases and eventually the plant dies. This condition is known as “Hopperburn” (Fig. 2.4). The insect is also known to secrete the extra sink sap collected in the form of clear liquid droplets known as honeydew which is excessively rich in sugar and amino acids (Sogawa 1982). N. lugens is believed to transmit two viruses: rice grassy stunt virus (RGSV) and rice ragged stunt virus (RRSV). RGSV was first reported in Philippines in 1963 (Rivera et al. 1966). The plant infected by the virus become stunted with formation of narrow, short and erect leaves which are pale green or yellow in colour giving a characteristic “mottled” appearance. RGSV was first reported in 1976 in Indonesia (Hibino et al. 1977). The rice plant infected with RGSV shows a typical symptom of stunting with dark coloured

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serrated or twisted leaf. Apart from that, there is occurrence of vein swelling and galls due to hyperplasia or hypertrophy of phloem tissues. These galls are seen in the underside of the leaf blades or on outer side of the leaf sheath (Hibino 1989). The insect gets infected by feeding on the infected plants and carries the infection until it dies. The route of infection by RGSV follows an initial infection of midgut epithelium, midgut visceral muscles, haemolymph and finally to the salivary glands of its BPH vector, sequentially (Zheng et al. 2014). The retention period can be as long as 40 days. The infection is restricted only to the particular individual that has fed the infected host plant and is not transmitted to the offspring (Ling 1972).

2.2.1.2 White-backed planthopper, Sogatella furcifera (Horvath) (Delphacidae: Hemiptera) White-backed planthopper (WBPH), Sogatella furcifera (Horvath) has a long history as an insect pest of rice. It was first reported and described as Delphax furcifera Horvath from Japan and later was given the synonyms, Delphacodes furcifera in Formosa (Schumacher 1915) and Megamelus furcifera in Nigeria. In 1963, the genus of these five species was changed to Sogatella by R G Fennah of the Commonwealth Institute, London (Khan and Saxena 1984). Its presence is confined mostly in Asia; however, the insect has distributed itself to the continents of Africa, Australia and South America (Fig. 2.5). This insect is widely distributed in Bangladesh, Taiwan, China, Japan, Korea, Saudi Arabia, Siberia, Micronesia, Philippines, Laos, Cambodia, Myanmar, Nepal, Vietnam, Thailand, India, Indonesia, Pakistan, Fiji and Seychelles (Heong and Hardy 2009). In India, the attack by S. furcifera was first reported in 1903. Earlier, it was considered as a minor pest of rice but now it has attained the major pest status all over India. For a long period it was reported to be a serious pest of rice only in the northern parts of India. However, after 1995 the pest showed its distribution to the

Fig. 2.5 Distribution of S. furcifera in the world (source: https://www.plantwise.org)

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southern tracts of the country leading to considerable damage and loss in rice production.

Identification Sogatella furcifera are slender, straw yellow or yellowish brown insects which are 2.5–4 mm long having a wing expanse of 3.8 mm and 4.2 mm for male and female, respectively. The forewings are uniformly sub-hyaline with dark veins and a conspicuous black dot at the middle of posterior margin of each forewing which meets when the forewings come together (Nasu 1967). There is a whitish band connecting the vertex to the mesonotum due to which the insect has been given the name of “White-Backed Planthopper”. Apart from that lateral areas of thorax are all black, while the abdomen is dark-brown. Early instar nymphs are 0.6–1 mm which are either white or strongly mottled with dark grey and white colour. The later nymphs bear a narrower head (white or creamy white in colour), whereas the thorax and abdomen are marked with various grey and white markings dorsally (TNAU Agriportal 2015).

Life Cycle Eggs The eggs are cylindrical in shape (Fig. 2.6a) and look similar to that of N. lugens eggs. The most preferred place for egg laying is the leaf sheath, midrib and stem of the plant (Atwal et al. 1967; Kumar et al. 2015). The eggs are too small to be seen in naked eyes but when viewed under a microscope, 110–150 eggs are seen to be laid in group or cluster of 5–30. Once the egg hatches, the yellowish brown ovipositional scar which was done by the adult female can be seen with naked eyes (Kumar et al. 2015). The incubation period varies from 6 to 7 days (Vaidya and Kalode 1981; Kumar et al. 2015), while with the decrease in temperature the incubation period increases and may reach up to 21 days in winter (Ammar et al. 1980). Nymphs The nymphs are greyish white in colour (Fig. 2.6b) and they pass through five instars by undergoing four moulting. The average nymphal period is 13–14 days in male and 15–16 days in female but it may increase with the decrease in temperature and may be 50–51 days in case of both male and female in winters. The nymphs become

Fig. 2.6 Different stages of S. furcifera: (a) eggs, (b) nymph, (c) adult female, (d) adult male

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motionless at the point of hatching, then move upwards to the drooping leaf to avoid sunlight where they start feeding. Adults Adults have brownish black pronotum with a yellowish brown body (Fig. 2.6c, d) and measure about 3.5–4.0 mm long. They possess very distinct white band between the junctions of wings and the apex of its front wing has an unbranched band. There is a conspicuous black dot at the middle of the posterior margin of each forewing which meets when the forewings meet together (Singh and Singh 2014). The hind tibia is noticeable because of its distinct movable spur. The adults exhibit two body forms, viz. macropterous or long-winged and brachypterous or short-winged. Macropterous males and females and brachypterous females are commonly found in the rice crop, whereas brachypterous males are very rare. The hoppers, especially adults, prefer to stay at the upper portions of rice stems. Adults are positively phototropic and are attracted to light (Singh and Singh 2014). Adults usually live for 10–20 days in summer and 30–50 days during autumn. The entire life cycle of S. furcifera is depicted in Fig. 2.7. Nature of Damage The WBPH is also a phloem feeder like BPH. Both the insects attack rice plant more or less simultaneously. To avoid the interspecific competition between themselves, they have developed some differences in their behaviour of feeding. WBPH feeds the plant on its vegetative stage (early tillering stage) (Sanchez et al. 2003), whereas BPH prefers more the reproductive stage of the plant (Sogawa 1995). Eventually, there also lies a difference in the place of feeding. The WBPH is concentrated more in the upper part of the leaf sheath or on leaves or on the junction of the leaf and stem (Mitai 1959) unlike BPH which is concentrated more on the base of the plant. The damage caused by WBPH is only prevalent when the population of the insect goes beyond five per hill. The symptoms include discolouration of outer leaf sheath of young rice seedlings due to excessive oviposition of the insect. Furthermore, the insect bears a capillary stylet which it inserts into the leaf sheath and suck the phloem sap, thereby leading to “hopperburn” finally (Fig. 2.8). At the latter stage of the plants, they feed on the ears and suck the glumes of rice plants leading to browning of the glumes and formation of sterile grains (Noda 1985). Southern rice black-streaked dwarf disease (SRBSDV) is transmitted by S. furcifera and it was discovered for the first time from China in 2001. Initially, the disease was prevalent in China only but the distribution range has now been spread to Vietnam and Japan subsequently (Xu et al. 2014). The virus enters into the insect body only when the insect feeds on an infected plant. The WBPH vector can acquire SRBSDV after only 5 min of feeding on infected plants and it can transmit the disease for 6–14 days, nymphs being more active in circulative transmission. The virus first gets transmitted from the infected plant through the stylet to midgut of the insect where it replicates and thereby it again disseminates itself to the salivary gland of the insect.

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Fig. 2.7 Life cycle of S. furcifera

Fig. 2.8 Damage symptom of S. furcifera on rice

The viruliferous insect is now ready to infect new healthy plants by inserting the stylet and secreting the viruliferous saliva. All the stages of the rice plant are susceptible to the disease but the extent of damage depends on the time of infection. The earlier the infection, the severe is the damage. When the plant is affected at the seedling stage it becomes dwarf with stiff leaves. In case of severe attack, there is excessive stunting (even shorter than one-third of normal height) and the seedlings finally wither off and die. When the infection occurs at tillering stage, there is significant dwarfing with numerous tiller formations but heading is prohibited, whereas at elongation stage there is no dwarfing, rather there is formation of small spikes, barren grains and deficient grain weight. The symptoms in mature plant include dark green foliage, aerial rootlets and branches on the stem nodes.

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2.2.1.3 Small Brown Hopper, Laodelphax striatellus (Delphacidae: Hemiptera) Laodelphax striatellus is one of the serious pests of rice (Sanada et al. 2011) which causes direct damage as well as acts as a vector of various viruses leading to huge economic loss in rice and other cereals. The insect has a wide distributional range which extends from Europe to entire Asia including certain parts of northern Africa (Fig. 2.9). The distribution includes Africa: Morocco and Tunisia; Asia: Armenia, China, India, Indonesia, Iran, Israel, Japan, Kyrgyzstan, Philippines, South Korea, Taiwan, Thailand, Uzbekistan, Turkmenistan and Vietnam; Europe: Austria, Belgium, Bulgaria, Czech Republic, Finland, France, Germany, Greece, Hungary, Italy, Latvia, Lithuania, Madeira, Netherlands, Poland, Romania, Russia, Slovakia, Spain, Sweden, Switzerland and Turkey; Oceania: Papua New Guinea (Mackesy and Moylett 2018). In India the insect was first reported from Punjab in 1970s. The insect is known to be polyphagous and apart from rice it has been reported from many other alternate hosts including wheat, barley and millets (Pathak 1975), sugarcane, bajra and maize (Mathur and Chaturvedi 1980).

Identification The genus Laodelphax is dorsally excavate having a quadrate vertex, very short pygofer, long and slender legs and a broad diaphragm. Eggs The eggs are white, cylindrical, usually laid in the leaf sheath or the midrib of the plant. The eggs are attached with each other anteriorly. They are laid in group of 60–260 and bear a flat and small egg cap (Dale 1994; Nasu 1967).

Fig. 2.9 Distribution of Laodelphax striatellus in the world (source: https://www.plantwise.org)

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Fig. 2.10 Different stages of L. striatellus: (a) nymph, (b) adult female, (c) adult male

Nymphs The nymphs are light to dark brown in colour. They are similar to N. lugens but are smaller in size. They undergo four moulting to produce five nymphal instars and the wing pads develop during the fifth instar of the nymph (Fig. 2.10a) (Nasu 1967). Adults The environmental factors and the genetic makeup of the insect play an important role in the development of winged adults. The adults may be macropterous or brachypterous (Mori and Nakasuji 1991) but the former is seen to be more common in L. striatellus (Wang et al. 2013). The adult possesses a dark-brown body, mesonotum and pterostigma whereas the areas between the carinae of the frons are deep black (Wilson and Turner 2010) but the abdomen is milky white to black in colour (Mori and Nakasuji 1991) and the wings are hyaline (Wilson and Claridge 1991). The adult males are longer (3.5 mm) (Fig. 2.10c) as compared to the females (2 mm) (Fig. 2.10b) (Nasu 1967).

Life Cycle Laodelphax striatellus is a polyphagous species, found in several types of habitats, from dry to moist and often in anthropogenic sites: fallow land, fertilized meadows, damp grassland, vineyards, swamps and road verges (Kirby 1992; Nickel 2003). It has been reported to feed on a variety of grasses including barley, maize, rice and wheat. The biology of L. striatellus is greatly influenced by the change in temperature. The average life span of L. striatellus adult decreases from 30.2 to 18.8 days in fluctuation of temperature from 20  C to 30  C (Chiu and Wu 1994); however, the longest span of adult is observed within 20–24  C (Wang et al. 2013). The female lays 71.75 eggs at 30  C but the fecundity is maximum at 25  C (186.17 eggs), which further decreases with the decrease in temperature, i.e. 146.04 eggs at 20  C. Female planthoppers make a slit in plant tissue with their ovipositors, insert their eggs in rows and cover them with a secretion that later solidifies. As far as the incubation period is concerned, the average duration for egg development at 20, 25 and 30  C is 14.02, 8.84 and 5.65 days, respectively (Chiu and Wu 1994). The nymphal period is also influenced by temperature fluctuation. The number of nymphs per female is highest at 28  C. The preferred range of temperature (18–32 C) bears majority of macropterous insects as compared to brachypterous ones (Wang et al. 2013).

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There are 4–7 generations of the insect per year (Wang et al. 2013) but the number of generations varies depending upon length of the growing season of crop. For instance, only two generations per year are observed in Sweden as compared to eight generations in Israel (Mackesy and Moylett 2018). In this case the insect undergoes diapause to synchronize their life cycle with the seasonal changes. Laodelphax striatellus undergoes nymphal diapause at low temperature and short photoperiods (Denno and Roderick 1990) unlike BPH and WBPH who migrate to escape the winter. Nature of Damage The insect sucks the sap from the plants and blocks the phloem tubes with their feeding stylets which prevents the translocation of the food materials within the plant system. The sucking of sap initially causes yellowing and wilting, which leads to “Hopperburn” with increase in severity of attack. L. striatellus transmits rice black-streaked dwarf virus (RBSDV) which is localized in the countries like China, Japan and Korea. There are many instances of outbreak of the virus in all these countries leading to considerable losses in production. Apart from rice, the virus has also caused economic failure in crops like maize, barley and wheat. However, the virus is still not reported from India (Lee et al. 1977). RBSDV is transmitted by the planthopper, L. striatellus and two other planthopper species (Hibino 1989; Shikata 1974). There is persistent transmission of the virus particles where the minimum inoculation feeding is 5 min and the incubation period is over 2 weeks. Once the virus is circulated into the host insect, the insect can transmit the virus for the rest of its life. However, the infectivity is reduced as the insects grow by age (Lida 1966). It is propagative in the vectors but virus passage via eggs is absent. The cycle of the RBSDV virus is maintained all around the year. Although it attacks cereal crops like rice, maize, wheat and barley, they are also known to attack many gramineous weeds (Obi and Kosuge 1963; Ruan et al. 1981). Once the major crops like rice are harvested (Sept–Oct), the infected L. striatellus moves to grasses temporarily and then infect barley and wheat by transmitting virus (Nov–Dec). The insect overwinters as nymph in winter cereals like wheat and barley and with the advent of suitable weather in March (before harvest of winter cereals), the insect overcomes diapause and becomes adult. These adults suck the infected sap of the winter cereals and attain infectivity to cause infection in further crops of rice and maize (May–June) (Obi and Kosuge 1963). In single-cropping rice areas, RBSDV incidence is generally high in early planted rice (Ishii and Yoshimura 1973), whereas in double-cropping areas, it is high in the second rice crop or late-planted rice (Ruan et al. 1981). Infected rice plants show pronounced stunting, darkening of leaves, twisting of leaf tips, splitting of the leaf margin and waxy white-to-black galls along the veins on the underside of leaf blades and the outer surface of sheaths and columns. The galls result from hyperplasia and hypertrophy of the phloem tissues. RBSDV is localized in the phloem and gall tissues. Furthermore, if panicles are formed, they are not

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developed completely and are partially covered with flag leaf sheath (Ling 1972; Liu et al. 2007).

2.2.2

Leafhoppers

2.2.2.1 Green leafhopper, Nephotettix virescens (Distant) (Cicadellidae: Hemiptera) Nephotettix virescens is a major pest of rice which is known to cause economic damage by sucking sap from the leaf of the plant. Earlier it was known to be a minor insect but over the years with the changing pattern of insecticide use, the insect has attained the major pest status. The insect is seen to be concentrated only in the south Asian patches of the world including India, Pakistan, Sri Lanka, China, Myanmar, Hong Kong, Vietnam, Thailand, Laos, Malaysia, Indonesia and Japan (Fig. 2.11). In India, it is reported from the states of Andhra Pradesh, Bihar, Karnataka, Kerala, Tamil Nadu, Assam, West Bengal, Odisha, Maharashtra, Madhya Pradesh (EPPO 2014). Identification The body of the adult hopper is yellowish green (Fig. 2.9c) and the size varies in males (4.70–5.22 mm in length and 1.38–1.55 mm in width) (Das and Devee 2017) as well as in females (4.90–5.50 mm in length) (Nielson 1968). The size of the head and the pronotum are almost similar, the vertex is pointed without any black markings on it and holds the ocelli at its anterior margin, the face is black in colour while the scutellum is green. In male insect the forewings are subhyaline and have a black patch which do not touch the claval region (Sashank 2009).

Fig. 2.11 Distribution of Nephotettix virescens in the world (source: https://www.cabi.org/isc/ datasheet)

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Life Cycle An average of 200–300 eggs (Pathak 1975) are laid in groups of 8–20 (Misra and Israel 1968) by making cravings in the leaf sheath with the help of the ovipositor. The eggs are cylindrical and whitish in colour with two distinct spots which later change to dark at the time of hatching (Mathur and Chaturvedi 1980). The total egg stage lasts for 5–10 days (Nielson 1968), whereas the incubation period is of about 6.5 days. The nymphs are green in colour (Fig. 2.12a) and undergo five moulting to become adults in a total span of 20–24 days (Misra and Israel 1968). There are six overlapping generations per year (from March to November) where the peak population and damage coincide with July–August prior to heavy rain in India. The abundance of Nephotettix spp. has been attributed to high temperature, low rainfall and abundant sunshine (Harukawa 1951). After the harvest of rice, the insect then feeds on numerous alternate hosts including Leersia hexandra, Ischaemum rugosum, etc. and overwinters as adult on the approach of winter (Atwal and Dhaliwal 1976). Nature of Damage Both nymph and adult of N. virescens insert their needle like stylet into the vascular bundle of the plant and block it by its salivary sheath while sucking the sap from the plant which leads to yellowing, reduced vigour, retarded growth, reduced productive tillers, withering and sometimes even complete drying of the plant (Fig. 2.13). Apart from that the insect causes indirect damage by transmitting the rice tungro virus, transitory yellowing, yellow dwarf and yellow-orange leaf (Dale 1994). Rice tungro virus is a combination of two viruses, namely rice tungro bacilliform virus (RTBV) and the rice tungro spherical virus (RTSV) where both RTBV and RTSV are stylet-borne. RTBV and RTSV are transmitted in a semipersistent manner by N. virescens, N. nigropictus, N. cincticeps, Recilia dorsalis and some other Nephotettix spp. but N. virescens is the most efficient vector of tungro disease (Hibino 1983; Khan et al. 1991a, b; Dey 2016). Green leafhopper becomes major insect pest in early transplanted coarse rice which transmits rice tungro virus from seedling stage itself. The high usage of nitrogenous fertilizers with high temperature triggers the flare-up of insect pest population that causes tungro virus (Dey 2016). Immediately after feeding on source plants infected with RTBV and RTSV, the

Fig. 2.12 (a) Nymph of N. virescens, (b) adult N. nigropictus, (c) adult N. virescens

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Fig. 2.13 Damage symptoms of N. virescens on rice

vector leafhopper transmits the viruses, either both viruses together or RTBV or RTSV alone. It retains RTBV for 4–5 days and RTSV for 2–4 days. The leafhopper readily acquires RTSV on source plants infected with RTSV alone, but it does not acquire RTBV on plants infected with RTBV alone (Hibino and Cabauatan 1987). Leafhoppers that have fed previously on RTSV-infected plants acquire RTBV from plants infected with RTBV. After moulting, leafhoppers lose both RTBV and RTSV infectivity (Ling 1972; Inoue and Hirao 1981). After rice is harvested, the density of N. virescens falls rapidly to a low level or to nil in the rice fields and then rice stubble serves as sources of RTBV and RTSV. N. virescens that feeds on source plants moves to newly transplanted fields in surrounding areas (Thresh 1991) and disperses the viruses. Rice plants infected with RTBV and RTSV together show tungro symptoms, including stunting, yellow or yellowish-orange discolouration and reduced tillering (Hibino et al. 1978). Discoloured leaves may show irregularly shaped dark-brown blotches (Dey 2016). The leaves, especially the younger ones, show the symptoms of striping or mottling and inter-veinal chlorosis. Plants infected with RTBV alone show similar but milder tungro symptoms. Plants infected with RTSV alone show no obvious symptoms except very mild stunting. In infected plants, RTBV is localized in the vascular bundles, whereas RTSV is restricted to the phloem tissues (Sta Cruz et al. 1993). Rice transitory yellowing virus (RTYV) is identical or very close to rice yellow stunt virus, which was first identified at about the same time in China (Faan et al. 1965). RTYV is transmitted in a persistent manner by N. cincticeps, N. nigropictus and N. virescens (Hibino 1989). It is propagative in the vectors but is not transmitted via eggs. Vector efficiency is high for N. cincticeps and N. nigropictus, but reported low in N. virescens. In infected leafhopper cells, RTYV particles are found in vacuolate structures of cytoplasm. RTYV naturally infects only rice. After harvesting of early planted rice, second- or third-generation adults of N. cincticeps move to the late-planted rice and spread the disease. Once the late-planted rice is harvested, RTYV overwinter in rice stubble or in its vector, N. cincticeps. Rice plants infected with RTYV show leaf yellowing, reduced tillering and mild stunting. The symptoms are very confusing since the infected plants temporarily

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develop normal-looking leaves and may appear healthy, but symptoms again reappear after this temporary recovery of the plant. In infected rice plants, RTYV is localized in the phloem tissues.

2.2.2.2 Zigzag Leafhopper, Recilia dorsalis (Motschulsky) (Cicadellidae: Hemiptera) Recilia dorsalis is a minor pest of rice distributed mainly on the south-eastern portion of the world (Fig. 2.14). Identification These are agile insects generally pale yellowish brown in colour and have a distinct zigzag w-shaped wedge on their forewing giving its common name as “Zigzag leafhopper” (Fig. 2.15b). The pronotum and its head are of similar width, whereas the vertex is moderately pointed, bearing a median sulcus extending more than half of the length of vertex from base. The anterior margins of vertex bear the ocelli very

Fig. 2.14 Distribution of Recilia dorsalis in the world (source: https://www.plantwise.org/isc/ datasheet)

Fig. 2.15 (a) Nymph of R. dorsalis, (b) adult of R. dorsalis

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close to the eyes. The pronotum is wider than its length. Tegmina is subhyaline with four apical cells and three anteapical cells (Sashank 2009). Life Cycle Eggs After 2 weeks of mating, female lays up to 90 eggs by inserting its ovipositor into the leaf sheath just below or above the juncture of the sheath. There is no specific pattern of egg laying. The incubation period is 6 days followed by hatching of eggs and production of nymphs (Misra and Israel 1968). Nymphs There are five nymphal instars within a duration of 120 days whereas the total life cycle is about 180 days per generation (Atwal and Dhaliwal 1976). Adults They look similar to nymphs but are bigger in size. The adults also suck sap from the plant. The total life span of an adult varies from 50 to 60 days. In sub-tropical and temperate areas where the temperature variation is maximum, the insect is seen to overwinter in the egg stage with the subsequent decrease in temperature (December– February). As soon as the temperature rises (March–April) the insect breaks the hibernation and hatching of the eggs takes place. Nature of Damage Both nymph and adult suck the sap from the leaf blade which leads to slight orange discolouration of the leaf from the margins to the entire leaf. Heavily infested plant with R. dorsalis may cause even death of the seedlings. Older leaves are affected first. Leafhopper nymphs and adults excrete honeydew which contains sugar and leads to the infection with sooty mould. R. dorsalis can transmit rice tungro bacilliform virus, rice tungro spherical virus, rice dwarf virus and rice orange-leaf virus. R. dorsalis transmits rice dwarf virus (RDV) which is distributed in China, Japan, Korea, Nepal, Mindanao and Philippines (Cabauatan et al. 1993). The occurrence of the disease though suspected in India due to the presence of its vectors is still not reported so far (John 1968). R. dorsalis and some other Nephotettix spp. (Hibino 1989) transmit the virus in a persistent manner to infect rice and few species of weeds. The insect when feeds infected plants for minimum 30 min (Mathur and Chaturvedi 1980) becomes capable of transmitting the virus to healthy plants and the virus can maintain its infectivity for 10–35 days. Furthermore, the viruses participate in transovarial transmission, i.e. they are transmitted from the parent female to their offspring via eggs. Infected rice plants show pronounced stunting, shortened internodes, increased tillering, inhibited root growth (only extend horizontally) and shorter leaves that are darker green in colour, with fine chlorotic specks. Since RDV is transmitted by

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leafhoppers, it is distributed to the vascular bundles and in parenchymatous cells in the portions of leaves which produce white specks. R. dorsalis also transmits orange-leaf disease (ROL) which is distributed in South East Asian countries including China, Japan, Philippines, Thailand, Malaysia, India, Sri Lanka, etc. The disease was first reported in Thailand, 1960 (Ou 1963), whereas from India the disease was reported in 2013 (Valarmathi et al. 2013). Recilia dorsalis is the vector of the virus which transmits the disease persistently. The shortest acquisition period is 5 h, whereas the incubation period of the insect is 6–7 days. The infected plant is the source of the virus transmission where the incubation period in the plants extends up to 13–15 days (Rivera et al. 1963). The typical symptoms of orange leaf include orange discolouration and inward rolling of one or two rice leaf which occurs after 14–20 days of infection, subsequently all the leaves undergo orange discolouration and severe infection may lead to death of the plant after 35–48 days of infection.

2.2.2.3 White Rice Leafhopper, Cofana spectra (Distant) (Cicadellidae: Hemiptera) It is the largest leafhopper infesting rice. The genus constitutes four species of economic importance, viz. C. lineata (Distant), C. nigrilinea (Stal), C. spectra (Distant), C. unimaculata (Signoret), of which C. spectra (Distant) is distributed in all the major rice growing areas including Southeast Asia and Africa (Fig. 2.16). Identification The insect is pale yellowish in colour having a black spot on its vertex, a central spot at the margin of face and two similar spots at the margin near the eyes. Furthermore, the insect bears a distinct white sub-hyaline tegmina with darker veins. The average length and width of the insect are 10.75 mm and 2.25 mm having the pronotum bigger than its head but smaller than the vertex. The pronotum bears distinct ocelli at

Fig. 2.16 Distribution of Cofana spectra in the world (source: https://www.plantwise.org/isc/ datasheet)

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the base of the vertex. Clypeus and clypellus are found to be swollen (Sashank 2009; Manurung et al. 2018). Life Cycle Eggs The insect restricts itself to the lower part of the plant and with the help of its ovipositor it makes parallel cut in the leaf sheath (Karim and Riazuddin 1999) where it lays greenish eggs (Atwal and Dhaliwal 1976) in a cluster of 5–19 with a fecundity range of 36–52 eggs in total. The eggs are cylindrical in shape (1.07–1.29 mm long and 0.31–0.39 mm wide) whose incubation period ranges from 9 to 10 days. Nymphs The egg on hatching leads to the emergence of new nymph which first feeds on the leaf sheath but later on moves to the leaf blade in the subsequent instars. There are five nymphal instars and the total nymphal duration is completed in 25–33 days, first instar and fifth instar being the longest and shortest instars comprising of 9 days and 3 days, respectively (Sashank 2009). Adults Both males and female adults are white in colour (Fig. 2.17). On an average the length of the body, wings and legs of female leafhoppers (7.93  0.5 mm, 6.61  0.58 mm and 6.93  0.35 mm) exceeds than that of the males (7.29  0.42 mm, 5.62  0.49 mm and 6.65  0.58 mm) (Manurung et al. 2018). The longevity of the adults also varies where the female lives longer (8–13 days) than the male (3–9 days). Nature of Damage Both nymph and adult suck sap from the rice plant. The initial symptom starts with drying up of the leaf tip which later turns orange and curls up (Karim and Riazuddin 1999). Furthermore, the insect causes yellowing as well as stunting of the plant Fig. 2.17 Adult of Cofana spectra: (a) male, (b) female

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leading to significant reduction in tillering. In case of severe damage, the leaves turn brown and the plants are devoid of producing ears (Atwal and Dhaliwal 1976).

2.2.3

Rice Bugs

2.2.3.1 Gundhi Bug, Leptocorisa spp. (Alydidae: Hemiptera) Rice bug or Gundhi bug is one of the most devastating pests of rice which infests its developing spikelet. It sucks sap from the grains and leads to significant loss in grain yield as well as grain quality. There are 14 species of Leptocorisa spp. reported mainly from Asia (Litsinger et al. 2015) out of which nine are reported to infest rice. Leptocorisa spp. which infest rice includes L. oratorius (F.), L. acuta (Thunberg), L. chinensis (Dallas), L. biguttata Walker, L. costalis (Herrich Schaffer), L. pseudolepida Ahmad, L. luzonica Ahmad and L. tagalica Ahmad (Ahmad 1965). Leptocorisa oratorius (F.) and L. acuta (Thunberg) are the most popular species distributed in different parts of Asia where the former is the most important species as far as the infectivity to rice is concerned, whereas L. acuta is the most widely distributed species infesting rice in the countries like Bhutan, Myanmar, Sri Lanka, China (Fukien), Hong Kong, India, Indonesia, Malaysia, Pakistan, Philippines, Taiwan, Thailand, Vietnam, Australia, Fiji, Papua New Guinea, Samoa and Solomon Islands (CIE 1966) (Fig. 2.18). This insect was first reported in Java in 1878 (Koningsberger 1878) but in India Gundhi bug outbreak was first noticed in 1886 (Lefroy 1908). Both the species, viz. L. oratorius and L. acuta are seen to be prevalent in India where the former has distributed itself to the south of India and the latter is dispersed in northern parts of India.

Fig. 2.18 Distribution of Leptocorisa spp. in the world (source: https://www.plantwise.org/isc/ datasheet)

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Identification Eggs Eggs of L. oratorius are oval, blackish and dull, whereas L. acuta have light-brown eggs with a shining lustre (Cobblah and Den Hollander 1992). The egg measures about 1–1.2 mm in length and 0.8–0.86 mm in width. The shape of the eggs is dorsally flat and ventrally convex. The eggs are laid in groups where each group consists of 18–20 eggs. Eggs are cemented to the leaf surface such that each egg is in physical contact with the neighbouring egg in the group (Fig. 2.19a) (Hosamani et al. 2009). Nymphs There are five nymphal instars in Leptocorisa sp. All the instars differ from each other in colour and size. The identification characters of the different instars of L. oratorius have been listed as follows: • First Instar The insect is pale greenish in colour and is smaller in size varying between 1.8 and 2.1 mm in length and 0.3 and 0.6 mm in width having a long reddish antenna with white bands and reddish-brown eyes. The antennae are generally longer than the body. • Second Instar The second instar nymphs have a similar external appearance as that of the first instar except the fact that it is bigger in size. The length and width of the instar vary from 5.8 to 6.2 mm and 0.60 to 0.90 mm, respectively (Fig. 2.19b). • Third Instar There is further increase in length (8.8–11 mm) and width (0.9–1.1 mm) and a slight change in colour of the insect to dark greenish was observed. The antennae turn brown and are just a little longer than the body now. This instar also marks the development of pale green wing pads. • Fourth Instar The fourth instar larva is greyish green in colour having reddish-brown eyes. The lateral side of the head bears various reddish-brown stripes. This instar is larger than the previous instars where the length and width vary from 12.5 to 14.5 mm and 1.2 to 1.4 mm, respectively. • Fifth Instar

Fig. 2.19 Different stages of Leptocorisa sp.: (a) eggs, (b) nymph, (c) adult of L. oratorius, (d) adult of L. acuta, (e) ventral abdomens of (i) L. acuta and (ii) L. oratorius

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These are pale brown insects which is 14–16 mm long and 1.45–1.75 mm wide. The wings pads are entirely developed. Adults The adults are green to brownish orange in colour having black spots on the ventrolateral part of the abdomen. The adults (Fig. 2.19c, d) are slender and robust where the female is more robust and smaller in size (17.5–18.5 mm long and 2.4–3 mm wide) as compared to the males (18–19 mm long and 1.95–2.50 mm wide). But in case of L. acuta the ventro-lateral spots are lacking and they are comparatively smaller than L. oratorius (Fig. 2.19e).

Life Cycle Eggs Leptocorisa spp. lays oval shaped eggs in batches of 19 eggs per batch after an incubation period of 6–8 days (Hosamani et al. 2009). The preference of egg laying varies with respect to species. For example, L. oratorius prefers fresh leaves for oviposition, while dead leaves and twigs are preferred by L. acuta. The flag leaf of rice is most favoured by Leptocorisa spp. for laying eggs (Kalshoven 1981). The insect lays eggs on different parts of the leaf and panicle but majority of the eggs are seen on the leaf tip (Cobblah and Den Hollander 1992). A specific pattern is followed for laying eggs. Eggs are generally laid longitudinally in 2–3 rows parallel to the midrib. The eggs are generally glued with a white substance after being laid to protect them from different adverse conditions such as strong winds and heavy rain (Lefroy 1908). The oviposition period may extend for an average of 48 days (Litsinger et al. 2015) where the average eggs laid per day is 4.6 (Corbett 1930). Nymphs The neonate nymphs are gregarious in nature. Soon after hatching (within 3–4 h) these neonate nymphs starts probing the egg shell or imbibe water droplets from the leaves or probe the leaf for plant sap. The first instar nymph is entirely confined to the leaves, whereas after moulting the next instars go wandering to the panicles to feed gregariously there. Earlier it was believed that both L. oratorius and L. acuta have five nymphal instars comprising a total developmental period of 19–21 days (Litsinger et al. 2015) but Dutta and Roy (2016) reported six nymphal instars of L. acuta with a total developmental period of 22–24 days (Fig. 2.20). Adults The adults suck sap from the panicle and the grain. The females are bigger in size than the males. The adult longevity is almost 3 months (Hosamani et al. 2009). When larger in number, the adults cause heavy yield loss by sucking sap from the developing grains. As far as the sex ratio is concerned, the females outnumber the male with the ratio 1.0:0.76 in case of L. oratorius, whereas in L. acuta the males dominate in number (1.0:1.12) (Sands 1977).

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Fig. 2.20 Life cycle of L. acuta

The adults undergo hibernation during chilling winters of December in grasses or on shrubs of uncultivated fields or in forests. Some of the bugs regain their activity with the increase in temperature towards March where it breeds in the grasses like Eleusine coracana and Echinochloa spp., whereas many undergo dormancy from March to July (Lefroy 1908). During this time the rice bugs derive its nutrition from the guttation water exuded by the grasses to survive (Sands 1977). As soon as rice attains booting stage, the adults then migrate to the rice plant.

Nature of Damage Both nymph and adult suck sap from the grain, leaf as well as shoot but the developing spikelet at milking stage was most preferred (Akbar 1958). The bugs bear piercing and sucking type of mouthparts and hence they pierce the stylet into the grains, thereby secreting a viscous fluid which hardens to form the salivary sheath. The salivary sheath helps in further penetration of the stylet. Apart from that the bugs secrete certain enzymes which predigest the endosperm to facilitate better piercing of stylet. Loss of sap from the developing gains makes them structurally deformed with several brown spots on them. The deformed grains with spots are called as “Pecky grains” (Fig. 2.21) which are easily broken while milling. The endosperm of the grain is spilled by the rice bugs which attracts several secondary pathogens to attack the grains (Kalshoven 1981). These secondary pathogen leads to discolouration of the kernel and glumes of the grains. This discolouration of the kernel and glumes is also referred to as “dirty panicle” (Fig. 2.22) (Agyen-Sampong and Fannah 1980). Furthermore, the brown spots in the grain are also the result of outgrowth of secondary pathogens around the feeding site of the rice bugs.

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Fig. 2.21 (i) Normal grains, (ii) pecky grains

Fig. 2.22 Secondary infections by pathogens

The earlier instars of rice bugs were found to imbibe guttation water from rice plants. On the other hand, the propagules of pathogens like bacterial leaf blight, Xanthomonas oryzae pv. oryzae and sheath rot disease Sarocladium oryzae were emitted through the guttation water. So, while imbibing the guttation water through the hydathodes of the leaf the rice bugs disperse the spores of X. oryzae pv. oryzae (Mohiuddin et al. 1976) and S. oryzae (Vivekananthan and Rabindran 2008) mechanically to the other plants and transmit it to them by piercing the proboscis into the leaf tissue.

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Rice Thrips

2.2.4.1 Stenchaetothrips biformis (Bagnall) (Thripidae: Thysanoptera) Thrips are distributed in all the South Asian countries like India, Thailand, Cambodia, Indonesia, Bangladesh, Taiwan, Myanmar, Japan, Malaysia, the Philippines, Vietnam and Sri Lanka. Apart from that the insect also marks its existence on the continents of Africa and some parts of Europe (Fig. 2.23). In India, S. biformis is found in the upland rice growing areas of Odisha, Andhra Pradesh, Punjab, Haryana, Uttar Pradesh, West Bengal, Assam, Tamil Nadu and Kerala. Identification The females are 1–2 mm in size having a brown body with paler tarsi and fore tibia. The antennae are 7-segmented and its segments were of varying colours. The head of the insect is as long as it is wide. The females are macropterous, having light-brown fringed forewings with a darker clavus. Males are similar to the females except to the fact that former are smaller in size (Bhatt et al. 2018). Life Cycle The female thrips select a tender young leaf and make incision on the abaxial surface of the leaf and lays 14–24 eggs singly. The eggs are translucent in nature (Fig. 2.24a). The eggs after 6–7 days hatch to the first instar nymphs which are pale in colour. The first instar nymphs start feeding on the leaf as soon as it hatches. After feeding for 3–4 days, the insect moults to second instar nymphs (Fig. 2.24b). The second instar nymphs feeds for 1–2 days to moult into a non-feeding pre-pupal instar. The pre-pupal instar and the pupal instar are completed in 2–3 days where the

Fig. 2.23 Distribution of rice thrips in the world (source: https://www.plantwise.Org/ knowledgebank)

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Fig. 2.24 Different stages of rice thrips: (a) eggs and first instar nymph, (b) second instar nymphs, (c) adult female

Fig. 2.25 (a) Rolling of rice leaf, (b) thrips damaged rice field

pupation occurs inside the rolled leaf caused by feeding of the insect. The entire life cycle is completed in 10–12 days (Nugaliyadde and Heinrichs 1984).

Damage The thrips bear asymmetrical mouthparts, i.e. the right mandible is absent. Thus, by continuous upward and downward movement of the head, it scrapes the upper epidermis of the leaf which causes oozing of plant sap (Lewis 1991). This sap is then sucked up by the thrips. Due to the scrapping, the upper epidermal cells die and cause upward curling of the leaf. This upward curling in the entire leaf causes the typical symptom of “leaf rolling” (Fig. 2.25a). The thrips mainly attack the plant at the seedling stage (Bhatt et al. 2018).

2.2.5

Rice Mites

There are about 61 species of mites which have been reported to feed on rice under field and storage conditions. Among them two mite species, viz. Panicle mite, Steneotarsonemus spinki Smiley (Acari: Tarsonemidae) and Leaf mite, Oligonychus oryzae Hirst. (Acari: Tetranychidae) are known to cause maximum yield loss in rice production.

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2.2.5.1 Panicle or Sheath Mite, Steneotarsonemus spinki Smiley (Acari: Tarsonemidae) The panicle or the sheath mite S. spinki has been recognized as one of the serious pest in major rice growing countries of Asia including China, India, Republic of Korea, Philippines, Sri Lanka and Taiwan since 1970. Apart from Asia, the pest has succeeded in establishing itself to parts of Central America (Caribbean Islands, Cuba, Dominican Republic, Haiti, Mexico, Costa Rica, Nicaragua, Panama and Puerto Rico) and Africa (Kenya and Madagascar) (Fig. 2.26). Identification The females of S. spinki are 263.0 mm and 92.4 mm in length and width, respectively. The females have an elongated body, white or yellowish white in colour and broadest in the region of hysterosoma. The legs are robust except for IV legs, which are typical tarsonemid female legs, terminating in a whip-like seta two times the length of the leg. The males are smaller in size and measure 196.5 mm in body length and 109.3 mm in width. The anterior ends of apodemes III extend further than apodemes IV. Femur IV has a large inner median lateral flange; and inner anterior and outer median setae are short and equal in length. The tarsal claw is stout and curved ventrally (Cho et al. 1999). Life Cycle Eggs The creamy or yellowish white elongated microscopic (0.131–0.211 mm in diameter) eggs are stacked singly or in cluster of 2–5 in the intercellular region in the inner surface of rice leaf sheath (Fig. 2.27a) (Chaudhari et al. 2019). The incubation period

Fig. 2.26 Distribution of rice panicle mite, Steneotarsonemus spinki Smiley (plantwise.org/ knowledgebank/datasheet/108962)

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Fig. 2.27 Different stages of rice mite, S. spinki: (a) eggs, (b) adult

of the eggs range from 1 to 3 days and its colour changes to transparent white at the time of hatching. The larva after hatching cuts open slits in the chorion with the help of its legs and comes out. Larvae The hatched larva is transparent white in colour and the females are bigger than the males. Larval males have three pairs of small setae besides the uropore, but, the females have two such setae. The larval period (2–4 days) is comprised of two stages, viz. the active stage (1 day) followed by the quiescence stage (2 days). The quiescence stage is the dormant stage of the larva since it restricts feeding till the emergence of the adult (Lakshmi et al. 2008). Adults The adults are also transparent. The males are broader having a dagger shaped setae, whereas the females were narrower and comparatively bigger than the males (Fig. 2.27b). The adult period generally ranges from 3 to 4.5 days in males but the females survive more (3.5–7 days) (Chaudhari et al. 2019). However, the adult longevity is highly governed by temperature. The female oviposit on the tissues they feed and can lay up to 78 eggs within a period of 1.5–3 days (oviposition period). The total life cycle is completed in 10.5–14.5 days in males, while females take 12.5–16 days to complete their development. S. spinki is facultative parthenogenetic in nature, i.e. the unfertilized females produce only males by parthenogenesis, while female can mate with the males and produce offspring of both the sexes (Xu et al. 2001). Damage The lower leaf sheaths of the rice plant harbour maximum sheath mites. Sometimes, they are also found on the basal part of the midrib or the leaf blade. During the vegetative stage of the rice plant, the pest is concerned with increase in its population. Once the plant attains reproductive phase, the mite migrates to the panicle and attacks the developing grain at the milky stage. The feeding of S. spinki leads to

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deformed inflorescence, leaf sheaths and discolouration of rice hulls. Continuous feeding leads to reduction in panicle size, less or partial grain filling and grain sterility. Damage of S. spinki is more severe on short duration varieties. The infestation initially starts at 80 DAT and attains its peak at 100–120 DAT. Summer rice is more prone to the attack of sheath mite (Rao and Prakash 1996). The pest overwinters in rice stubbles or on alternate host (Schoenoplectus articulatus, Cyperaceae) during the non-availability of the main crop. Steneotarsonemus spinki also act as a vector of the sheath rot fungus, Acrocylindrium oryzae. The damage caused by the mites enhances the chances of sheath rot disease in rice since the mite harbours the conidia of the fungus in its body itself. The damaged tissue by the mite becomes an easy target to the inoculation of sheath rot fungus. The attack of the fungus leads to browning of the grains and leaf sheath. However, the fungus does not contribute to grain sterility.

2.2.5.2 Leaf Mite, Oligonychus oryzae Hirst. (Acari: Tetranychidae) Life Cycle The entire life cycle of Oligonychus oryzae comprises five different stages, viz. egg, larva, protonymph, deutonymph and adult. There are three quiescence stages, viz. nymphochrysalis, deutochrysalis and teliochrysalis between egg and adult stage of the mite (Aswin et al. 2016). Eggs Spherical, white transparent eggs (Fig. 2.28a) are laid singly or in small clusters of 10–12 on the upper surface by the gravid female. The egg resembles tiny droplets of water, consisting of two reddish eye spots on them. The colour of the egg gradually changes to dull white prior to hatching. The incubation period for hatching of the eggs is 3–4 days. Larvae Just after hatching from the eggs, the larvae are spherical and white in colour (Fig. 2.28b). They bear three pair of legs and a pair of simple eyes at the dorsum of propodosoma. Once the larva starts feeding, its colour changes to yellowish green.

Fig. 2.28 Different stages of leaf mite, Oligonychus oryzae: (a) eggs, (b) nymph

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The duration of larval period is lesser for males (1.37 days) as compared to females (1.40 days). Protonymph The larva undergoes moulting followed by a short quiescence (nymphochrysalis) which leads to the formation of a nymphal instar called as the protonymph which is oval in shape, amber in colour and slightly bigger in size as compared to the larvae. The protonymph bears 4 pairs of legs. Deutonymph It is the second instar nymph which emerges when the protonymph undergoes moulting. It is an active instar generally greyish green initially, which subsequently changes to dark green. Sexual dimorphism exists with respect to body size, where females are larger. Adults Adult mites emerge after the teliochryslais. The males are tapering posteriorly to a blunt point and are generally smaller than the females. After emergence the males move in search of female and guard them from the deutonymph stage and mate with them. Oligonychus oryzae also exhibit facultative parthenogenesis where the unmated and mated female lays up to 21 eggs and 18 eggs, respectively. The unmated females produce only males (Aswin et al. 2016). Damage The mite colonies settle themselves on the under surface of the leaf and devour the palisade layer of the leaf, thereby severely affecting the photosynthesis of the plant. Severe feeding depletes the nutrient content and causes yellow pecks on the leaf (Bhatt et al. 2018).

2.3

Management of Sucking Pests

2.3.1

Cultural Methods

Cultural practices are the modification of agronomic production practices which makes the environment least favourable for pest invasion, reproduction, survival and dispersal but makes the environment favourable for crop production. Among the cultural practices, crop rotation, spacing of the plant, water management, proper sanitation, fertilizer management and synchronous planting have been found to play important role in management of sucking pests. (a) Crop rotation Rice hoppers mainly depend on rice for their growth and reproduction. In tropical environment rice is grown twice or even thrice in a year. In these cases a constant source of food is available to them so as to flourish throughout

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the year. When rice is not available, the hoppers (especially leafhoppers) adapt themselves to certain alternate hosts (weeds) which only act as a mode of survival while their reproductive rate is highly reduced. So rotating any annual crop which is a non-host of hopper in between two crops of rice breaks the life cycle of the pest. The crops that can be taken as an option for rotation are soybeans, mung beans and sweet potatoes (Oka 1979). (b) Sanitation Weeds play an important role in maintaining the population of rice hoppers. These weeds influence the survival of the rice hoppers for the next season of rice since after the harvest of the rice plants the hoppers usually find weeds as the alternate host to survive (Alam 1964; Pathak 1969). Furthermore, the hoppers prefer a dense and overcrowded condition for its growth and development which is provided by the presence of weeds in the rice field. A weedy rice field harbours more hopper and causes more economic damage as compared to a weeded plot (Fernando 1975). Apart from weeds, the rice stubbles and the ratoon crop also acts as a harbour point for the hoppers to extend its life cycle (Altieri 2019). These stubbles also act as source of inoculum for various transmissions of viruses by both leafhoppers and planthoppers. Realizing the detriments of weeds and stubbles, steps should be taken to overcome them. A planned approach for sanitation to abolish the weeds in the later stages of the crop, the stubbles and ratoon after harvest of the crop is the most effective way to destroy the breeding/hibernating sites and sources of food for hoppers (Babendreier et al. 2019). For example, the epidemics of the rice dwarf virus transmitted by the green leafhopper can be completely subdued within 2 years by winter ploughing of weed (Alopecurus aequalis Sobol), an alternate host for the green leafhopper (Nakasuji and Kiritani 1976). Burning the stubble and straw after harvesting of the crop is an excellent method of sanitation which has been recommended by many scientists (Israel 1969; Kulshreshtha et al. 1974). Burning not only kills the overwintering arthropods but also helps in rapid decomposition of the plant remains (IRRI 1973). However, the advantages of burning are supressed by its disadvantages in the present times due to which it should not be recommended. The harmful effects of burning include loss of nutrient status of the soil, reduction of soil organic matter, depletion of beneficial soil flora and fauna (Mandal et al. 2004). Apart from that, the main disadvantage seems to be the degradation of environment by emission of various greenhouse gases (GHG) including 0.7–4.1 g of CH4 and 0.019–0.057 g of N2O per kg of dry rice straw and emission of other gaseous pollutants such as SO2, NOx, HCl and, to some extent, dioxins and furans (Oanh et al. 2011; Jenkins et al. 2003). (c) Spacing The hoppers prefer a microclimate which is shaded, cool and humid where the sun rays rarely penetrate. This favourable microclimate is provided to them by making a dense planting (Satpathi et al. 2012). It also deters the development of natural enemies which adds an extra benefit for the increase of the hopper

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population (Nishida 1975). Furthermore, the solar and ultraviolet radiations of the sun adversely affect the hoppers (Suenaga 1963). (d) Fertilizer management Excess nitrogen makes the plant more succulent and makes the canopy of the crop thicker and denser which forms a microclimate much favourable to hoppers (Nishida 1975; Satpathi et al. 2012). Apart from that nitrogen leads to more production of protein and amino acid in rice plant which are considered as the essential requirements of the hoppers. However, a higher dose of phosphorus and potassium suppresses the hopper infestation (Satpathi et al. 2012; Rashid et al. 2016). There have been numerous reports of increase in hopper infestation with indiscriminate fertilizer application (Velusamy et al. 1975; Rashid et al. 2017). Under such condition proper dose and timing of fertilizer application should be practised to manage the pest. (e) Water management The hoppers are shade loving insects. They prefer a microclimate which is shaded with high humidity (Nishida 1975; Chaudhary et al. 2014; Sarkar et al. 2018). Hence role of irrigation plays a vital role in the management of hoppers. There have been numerous reports of maximum population of insect coinciding with the period of standing water in the field (Fernando 1975). Standing water in the field creates an environment which is much favourable for the hoppers to flourish. On the other hand, standing water in the rice field has its own importance since it minimizes weeds and many other insects. Under such circumstances, alternate drying and wetting of the field at peak population of the hoppers play an important role in their management. Draining of water from the field decreases humidity which forces the insect to move towards the soil in search of optimum moisture where they can lay eggs. Again flooding the field destroys all the laid eggs of the hoppers, thereby causing efficient management.

2.3.2

Biological Control

Natural enemies play a very important role in suppressing the pest population below economic injury level (Dainese et al. 2017). This component is even more effective if it is integrated with other eco-friendly ways of pest management. The planthoppers and leafhoppers are known to be the host 19 egg parasitoids, 16 nymphal and adult parasitoids, 37 predators (including insect and spiders), one nematode (Chiu and Lung 1975), 8 entomo-pathogenic fungi and one bacteria (Shimazu 1976). Some of the most promising natural enemies, viz. predators (Table 2.3), egg parasitoids (Table 2.4) and adult parasitoids are listed below (Table 2.5): There are eight fungi and one bacterium which have been reported to attack both planthoppers and leafhoppers in rice (Li 1985). Entomopthora delphacis is considered as the most prevalent fungi reported to be infecting the brown planthopper in field condition (Shimazu 1976). Apart from that many other fungus including Beauveria bassiana, Metarhizium anisopliae and M. flavoviride var. minus are also found to be effective against both leafhoppers and planthoppers. However,

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Table 2.3 Effective predators of leafhopper and planthopper Order Hemiptera

Hymenoptera Coleoptera

Family Nabidae Miridae Veliidae Formicidae Carabidae

Staphylanidae Coccinellidae

Araneae

Salticidae Lycosidae

Predator Nabis spp. Cyrtorhinus lividipennis Migrovelia douglasi Tetramorium guineense Acupalpus inornatus Bembidion semilunium Casnoidea cyanocephala Ophionea indica Paederus fuscipes Stenus cicindelloides Coccinella arcuata Hippodamia tredacimpunctata Micrapis discolor Plexippus paykulli Lycosa pseudoannulata

Source: Fahad et al. (2015) Table 2.4 Effective egg parasitoids of leafhopper and planthopper Order Hymenoptera

Family Eulophidae Mymaridae

Trichogrammatidae

Diptera

Dryinidae Pipunculidae

Parasitoid Ootetrastichus spp. Anagrus flaveolus Gonatocerus spp. Lymaenon spp. Anaphes spp. Polynema spp. Camptoptera spp. Aphelinoidea spp. Oligosita spp. Paracentrobia andoi Paracentrobia garuda Trichogramma spp. Echthrodelphax migratorius Tomosvaryella spp.

very less effort relating to its formulation and commercialization has been done in India. In Korea, 7.5  1012 conidia/ha of B. bassiana and M. anisopliae and 4  1012 conidia/ha of M. flavoviride var. minus are recommended for effective management of rice hoppers (Geng and Zhang 2004). As far as the mycelium is concerned, the recommended dose for suppression of brown planthopper is 200 and 2000 g/ha, respectively (Aguda et al. 1987).

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Table 2.5 Effective nymph and adult parasitoids of leafhopper and planthopper

91

Order Hymenoptera

Family Dryinidae

Encyrtidae

2.3.3

Strepsiptera Diptera

Elenchidae Pipunculida

Hymenoptera

Dryinidae

Parasitoid Echthrodelphax fairchildi Haplogonatopus sp. Pseudogonatopus flavifemur Chrysopophagus australiae Echthrogonatopus exitiosus Elenchus laponicus Dorylas sp. Pipunculus javanensis Pseudogonatopus spp.

Use of Resistant Varieties Against Hoppers

Several varieties (Table 2.6) have been reported to be tolerant/resistant against plant and leaf hoppers.

2.3.4

Chemical Management

Several insecticides have been used for the management of plant and leaf hoppers. As far as the hoppers are concerned, it is always difficult to manage the planthoppers as compared to the leafhoppers. The planthoppers are basically present on the base of the plant which prevents them from the direct exposure of the contact of insecticides. The solution to that was use of systemic insecticides. But systemic insecticides were translocated along the xylem and got accumulated majorly on the leaves and rarely on the leaf sheath (Sur and Stork 2003) where the planthoppers escape the adverse effect of insecticides. Furthermore, the sub-lethal doses cause resurgence of the hoppers demanding further increase of concentration for control of the insect and a very high reproductive rate of hoppers makes it very easy for them to develop resistance against the insecticides. It is for this reason hoppers have developed resistance to many insecticides irrespective of their mode of action. Therefore, application of insecticides must follow certain norms and precaution such that the above mentioned problems could be avoided which are as follows: (a) Timely application The insecticides when applied on the hoppers killed the insects but failed to kill the eggs which further developed to nymphs and maintained a rapid increase in the population, thereby overcoming the effect of insecticides. Therefore, it is advised to apply the insecticides coinciding with the maximum population of third and fourth instar of the hoppers so that the egg stage of the insect can be avoided (Heinrichs et al. 1982). (b) Rotation of Insecticides Repeated use of a particular insecticide had enabled the hoppers to modify themselves at biochemical level so as to develop resistance against that

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Table 2.6 Tolerant/resistant varieties against plant and leaf hoppers Name of variety ADT 44 ADT 47 Ajay Basundhara Daya DRRH-2

Tolerant/ resistant against GLH BPH, WBPH BPH, WBPH BPH GLH, BPH WBPH

Golak JKRH 401 Karjat 5 KRH-2

BPH WBPH GLH, BPH BPH

Lalat MDU 3 Naina

GLH, BPH BPH, WBPH Rice tungro virus, BPH Rice tungro virus, BPH

PA 6201

PA 6444

Rice tungro virus, BPH

R 636–405 Revathy Richa Samalei Shaktiman Sneha

BPH BPH WBPH GLH, BPH WBPH, BPH Rice tungro virus GLH WBPH, BPH BPH BPH BPH BPH BPH BPH BPH BPH

Sugandhamati Suraksha Vijetha BRRI dhan 35 Bg 379–2 CR203 Jingxian89 GMJ3 GMJ5 GMJ5

Source: Bhatt et al. (2018)

State/country Tamil Nadu Tamil Nadu Odisha Assam Orissa Tamil Nadu, West Bengal, Haryana, Uttarakhand, Assam, Chhattisgarh, Gujarat, Odisha, Madhya Pradesh, Uttar Pradesh Odisha, West Bengal Andhra Pradesh, Bihar, Jharkhand, West Bengal Maharashtra Andhra Pradesh, Assam, Bihar, Chhattisgarh, Karnataka, Jharkhand, Kerala, Madhya Pradesh, Maharashtra, Odisha, Tamil Nadu, Uttar Pradesh, West Bengal Odisha Tamil Nadu Andhra Pradesh Andhra Pradesh, Assam, Bihar, Chhattisgarh, Karnataka, Jharkhand, Madhya Pradesh, Odisha, Tamil Nadu, Uttar Pradesh, West Bengal Andhra Pradesh, Assam, Bihar, Jharkhand, Maharashtra, Odisha, Uttar Pradesh, Tripura, Karnataka, Uttarakhand, Gujarat Chhattisgarh Kerala Andhra Pradesh, Assam, Chhattisgarh Odisha, Madhya Pradesh Odisha, West Bengal Odisha Andhra Pradesh, Assam, Gujarat, Tamil Nadu Odisha, West Bengal West Bengal Bangladesh Sri Lanka Northern Vietnam Guangdong Province of China Indonesia Indonesia Indonesia

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Table 2.7 Insecticides against brown planthopper (N. lugens) and white-backed planthopper (S. furcifera) Pest/pesticides Buprofezin 25 SC Clothianidin 50 WG Flonicamid 50 WG Dinotefuran 20 SG Imidacloprid 70 WG Imidacloprid 30.5 SC Imidacloprid 17.8 SL Acetamiprid 20 SP Fenobucarb 50 EC (BPMC) Thiamethoxam 25 WSG Ethofenoprox 10 EC Fipronil 5 SC Monocrotophos 36 Phosphamidon 40 SL Carbaryl 50 WP Acephate 75 SP Carbosulfan 25 EC Carbofuran 3 CG Acetamiprid 0.4 + chlorpyrifos 20 EC Buprofezin 15 + acephate 35 WP

Dose (g a.i./ ha) 200 10–12 75 30–40 21–24 21–26 20–25 10–20 625 25 50–75 50–75 500 500 750 500–750 200–250 750 10 + 500

Formulation (g/mL/ ha) 800 20–24 150 150–200 30–35 60–75 100–150 50–100 1250 100 500–750 1000–1500 1400 1250 1500 666–1000 800–100 25,000 25,000

Waiting period (days) 20 12 36 21 7 37 40 7 30 14 15 32 – – 15 15 14 – 10

187.5 + 437.5

1250

20

Source: Bhatt et al. (2018)

particular insecticide at a very faster rate (Wu et al. 2018). Hence, it is always safe and smart to rotate the spray of insecticide with different mode of actions which will provide least chances for the insect to adapt to the insecticide biochemically. (c) Maintaining proper dose Sub-lethal dose leads to resurgence of hoppers (Bao et al. 2009; Liu et al. 2016). Resurgence further increases the concentration of the insecticides as an attempt to manage the hoppers. In the present day where the entire world is polluted with insecticides leading to various abnormalities, aberration in doses and its consequences cannot be afforded. The proper dose and formulations of different insecticides to manage hoppers are listed in Tables 2.7 and 2.8.

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Table 2.8 Insecticides against green leafhopper (N. virescens and N. nigropictus) Pest/pesticides Buprofezin 25 SC Flonicamid 50 WG Thiamethoxam 25 WSG Ethofenoprox 10 EC Fipronil 5 SC Acephate 75 SP Carbosulfan 25 EC Carbofuran 3 CG Carbosulfan 6 G

Dose (g a.i./ha) 200 75 25 50–75 50–75 500–750 200–250 750 1000

Formulation (g/mL/ha) 800 150 100 500–750 1000–1500 666–1000 800–100 25,000 16,700

Waiting period (days) 20 36 14 15 32 15 14 – 37

Source: Bhatt et al. (2018)

2.4

Management of Rice bugs

2.4.1

Cultural Method

(a) Trap crops A small area of early maturing variety should be sown early than the main crop such that attainment of its milking stage is earlier which would attract the rice bugs to the crop. These bugs were then managed by chemicals (Corbett 1930). (b) Sanitation The population of rice bugs builds up in the weeds near rice fields and as soon as rice attains the milking stage the rice bugs shift their preferences to the rice. A total of 76 weeds species in rice ecosystem have been reported to support the growth and development of rice bugs (Litsinger et al. 2015). It is advised to remove the weeds from the outskirts of the rice field prior to the flowering stage of rice which will deprive the rice bugs from an alternate host and will minimize damage to the main crop. Furthermore, rice stubbles along with weeds after the crop harvest should also be removed to minimize the pest incidence in the subsiding season (Pasalu et al. 2004). (c) Plant maturity The maturity status and the time of sowing of rice affect greatly the relative abundance of the rice bugs. Since rice is most susceptible to rice bugs at its peak flowering stage (Berg 2000), it is always advised to the farmers to grow late maturing varieties in rice bugs endemic areas. It was observed that the early maturing varieties harbour ten times more the population of the rice bugs (Rothschild 1970). In India, the late maturing or the late sown plants escape the bug’s damage because the flowering period in these plants does not coincide in June–October when the population of the rice bugs is maximum (Pasalu et al. 2004).

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(d) Synchronous flowering In case of staggered planting, i.e. when the sowing is done at different times, the rice bugs avail a constant supply of host from the earliest period of sowing to the last harvested plant. This causes severe damage to the crops. Instead, synchronous planting should be adopted over large areas. The idea behind this concept is that initiation of flowering in the rice plant will occur at the same time over the area which will shorten the duration of attack by the rice bugs. Even though the population of the bugs will be high, the damage to the plants as a whole will be less (Litsinger et al. 2015).

2.4.2

Mechanical Control

One of the most ancient ways of controlling insects was picking them by hand during their peak population and destroying them. In India, it is recommended to the farmers to collect the eggs and the adults of rice bugs during August and destroy them by burning or dipping them in kerosene to minimize the loss caused by rice bugs. This method is followed by almost all the rice growing areas where the collection of insects was carried out by various collecting devices including hand nets (Corbett 1930), cloth bags (Biswas 1953), adhesive ropes (Drieberg 1909), etc.

2.4.3

Physical Control

Light traps attract the insects towards it. It was initially used for monitoring the incidence of the pest but when they are installed at the rate of 6–8 per acre they provide efficient management against rice bugs. The light traps when installed at the above rates can trap almost 3000 bugs/night (Sen and Chaudhury 1959). These light traps were installed with a basin containing water mixed with kerosene or crude oil to kill the adult bugs.

2.4.4

Host–Plant Resistance

There are certain properties of the plant which confer resistance to the rice bugs. In India, certain varieties like Sathi, Soma and Mundagakutty have an enclosed panicle within the leaf sheath which resists the stylet penetration of the rice bugs. Apart from that certain more varieties including Badshabhog, CR44–82, CR44–35, W1263, GAR 1, GAR 2. IR 22 are also reported to be resistant to rice bugs (Kalode and Yadava 1975; Kakde and Patel 2018).

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Table 2.9 Natural Order enemies of Leptocorisa sp. Predators Orthoptera Odonata Coleoptera Hemiptera Araneae

Parasitoids Hymenoptera Insect Pathogens Hypocreales

Family

Scientific name

Gryllidae Tettigonidae Libellu Coenagrionidae Coccinellidae Reduviidae Tetragnathidae Argiopidae Thomisidae

Anaxipha spp. Conocephalus spp. Neurothemis fluctuans Agriocnemis sp. Vernia discolor Sycanus dichotomus Cosmolestes picticeps Eucta spp. Argiope spp. Thomisus spp.

Scelionidae

Gryon flaviceps

Cordycipitaceae Clavicipitaceae

Beauvaria bassiana Metarrhizium anisopliae

Rothschild (1970), Litsinger et al. (2015)

2.4.5

Biological Control

The tropical weather conditions during the milking stage of rice act as the most favourable condition for breeding of the insects. Apart from that there are several biotic agents who act as a limitation to the growth and development of the bugs which include various predators, parasitoids and pathogens. The details of the natural enemies of rice bugs are enlisted in Table 2.9.

2.5

Management of Thrips

Thrips are more prone to areas of water stress (Chiang 1977; Thomas et al. 1979; Nugaliyadde and Heinrichs 1984). Therefore, flooding the field to submerge it for almost 2 days decreases the population of thrips (Catling and Islam 2013). Thrips overwinters on several alternate hosts such as weeds (Madhusudhan and Gopalan 1989). A proper field preparation 2–3 times before planting destroys the weeds, thereby reducing the alternate hosts to check further damage of the insect. Apart from that use of early maturing varieties, synchronous planting also plays an important role in the management of the pest. However, use of chemicals for the management of thrips is most effective (Table 2.10). These chemical insecticides should be applied at a specified time and amount to minimize the environmental hazards. Nitrogen can be applied after infestation of the thrips which compensates the damage caused by them and improves the plant growth.

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Table 2.10 Insecticides against rice thrips, Stenchaetothrips biformis S. no 1 2 3

Insecticide Lambda cyhalothrin 2.5 EC Lambda cyhalothrin 5 EC Thiamethoxam 25 WG

Dose (g a.i./ ha) 12.5

Formulation (g/mL/ ha) 500

Waiting period (days) 15

12.5

250

15

25

100

14

Source: Bhatt et al. (2018)

Table 2.11 Acaricides against rice mites Dose (g a.i./ S. no Acaricide ha) Panicle Mite, Steneotarsonemus spinki 1 Spiromesifen 72 240 SC 2 Dicofol 18.5 EC 500 Leaf Mite, Oligonychus oryzae 3 Fenazaquin 10 EC 100 4 Abamectin 1.8 EC 5 5 Fenpropathrin 100 10 EC

Formulation (g/mL/ ha)

Waiting period (days)

300

–

2500

–

1000 250 1000

– – –

Source: Bhatt et al. (2018)

2.6

Management of Mites

Devoid the mites with continuous supply of its host by subsequent fallowing of fields and rotation with a non-host crop. After the crop is harvested, the mite overwinters on the stubbles. Hence, the stubbles should be ploughed after harvesting the crop to ensure no overwintering population. The migration of the rice mites from one field to other is majorly brought about by human interventions such as use of various machines for performing field operations. Hence, proper sanitization of the machinery before using in an un-infested field is much required to prevent the further spread of the pest. Apart from that there have been certain resistant varieties developed against panicle mite, viz. MTU-7029, IR64 (Karmakar 2008) and leaf mite, viz. Paiyur 1, ADT 39 (Thilagam and Jalaludin 2018) which restricts the attack of the pest. However, systemic miticides or acaricides are the best option for management (Table 2.11).

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Conclusions

The challenge of more production of rice is unanimously increasing with the everincreasing trend in population. But the need of the hour also includes safety and sustainability of the ecosystem. There have been numerous works reflecting the accomplishment of Integrated Pest Management (IPM). Still chemical insecticides are the first choice among farmers for the management of sucking pest of rice. This is due to relatively poor socio-economic condition and ignorance of the famers. The sucking pests of rice, viz. brown planthopper and white-backed planthopper have the capacity to resist these chemicals (insecticides) at a very faster rate creating havoc among crop growers. The only way to overcome these constraints is complete dependence on IPM. There has been detailed information regarding the various interventions of IPM, viz. cultural, botanical, physical, mechanical, biological, host–plant resistance and environmental safe chemicals which can be applied in combination at different phases of the crop growth for a maximum suppression of the pest with respect to ecological sustainability.

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Sucking Pests of Pulse Crops Hem Saxena, Sanjay M. Bandi, and Revanasidda

Contents 3.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Significance of Sucking Pests . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Sucking Pests of Pulses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4 Integrated Management of Sucking Pests in Pulses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

Pulse crops are important component of tropical cropping systems contributing significantly to the farming system sustainability. Besides, pulses have high biological value as they are the principle source of dietary proteins among the vegetarian populations in India. Pulse crops are infested by a large number of insect pests at all the stages of crop growth which cause an average yield loss of 18–20% annually. Introduction of high yielding varieties, agrochemicals and changed cropping patterns had significantly altered the pest status in pulse crops specifically the sucking pests causing significant impact on pulse production, which were otherwise recognised as pests of lesser economic significance. The information on sucking pest complex in pulse crops, their distribution, incidence, host range, and nature of the damage is a prerequisite for successful management of these sucking pests in pulse crops.

H. Saxena (*) · S. M. Bandi · Revanasidda Division of Crop Protection, ICAR-Indian Institute of Pulses Research, Kanpur, India # Springer Nature Singapore Pte Ltd. 2020 Omkar (ed.), Sucking Pests of Crops, https://doi.org/10.1007/978-981-15-6149-8_3

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Introduction

Pulse crops are important component in tropical cropping systems and they are principle source of dietary proteins among vegetarian population in India. The protein content of pulses is about 22–24%, which is almost twice the protein content in wheat and thrice that of rice (Gowda et al. 2013). Pulses along with cereals form a perfect vegetarian diet of high biological value as they complement the staple cereals in the diets with proteins, essential amino acids, vitamins, and minerals. Pulse crops can be grown on a range of soil and climatic conditions as well as included in crop rotation, mixed cropping, and intercropping, thus maintaining soil fertility through biological nitrogen fixation and contributing significantly to the sustainability of the farming systems. India is the largest producer (25.42 million tonnes) and consumer of pulses in the world. Pulses account for around 20% of the area under foodgrains and contribute around 8–10% of the total foodgrain production in the country (DAC and FW 2018). Among the pulse crops, chickpea and pigeonpea are the major crops grown in India. These two crops together occupy more than 50% of total area and contribute over 62% to the total pulse production. Of which, chickpea dominates with over 45% share of total pulse production followed by pigeonpea (17%), urdbean (14%), mungbean (8%), and lentil (6.37%). The major pulse growing states are Madhya Pradesh, Maharashtra, Rajasthan, Uttar Pradesh, Karnataka, and Andhra Pradesh, together contributing about 75–80% to the total pulse production (DAC and FW 2018). Biotic factors are the major constraints in achieving the potential yield of pulses. Among the biotic stresses, diseases and insect pests are the major yield limiting factors. In India, about 26–30% of the potential production of pulse crops is annually lost due to insect pests and diseases. The annual yield losses due to insect pests and diseases have been estimated to be 18–20 and 8–10%, respectively (Dhar and Ahmad 2004). The change in cropping pattern and introduction of high yielding varieties has tremendously affected the pest complex in pigeonpea. The sucking pests which were earlier recognised as minor pests in pulses with lesser economic significance are attaining a status of major pests (Saxena et al. 2018). Pulse productivity has been severely threatened by increasing difficulties in managing these sucking pests due to their ability to evolve resistance to insecticides, resurgence and their secondary outbreak due to indiscriminate and injudicious application of synthetic insecticides. To devise economically feasible, ecologically sound, and socially acceptable management strategies against sap feeding pests of pulses, the detailed information on pest complex, their status and temporal association with host plant, yield losses, nature of damage, and feeding symptoms is of great significance.

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Significance of Sucking Pests

Sucking pests or sap feeders often have a long-lasting physiological effect on the growth of the host plant and these physiological changes include changes in both plant nutrients (Masters and Brown 1992) and plant secondary metabolites (Karban and Myers 1989). Sap feeders remove the nutrients from xylem or phloem of the host plant, thereby decreasing photosynthetic rates and plant growth (Meyer 1993). The important sucking pests infesting field crops include thrips, leafhoppers, aphids, plant bugs, whiteflies, scales, mealybugs, and mites. These sucking pests are difficult to control due to their ability to evolve resistance to synthetic insecticide, resurgence or their secondary outbreak. These pests rebound quickly under favourable climatic conditions and often cause severe economic losses. The pulse crops are affected by a number of sucking pests causing either direct or indirect (acting as vectors of viral diseases) yield losses.

3.3

Sucking Pests of Pulses

Sucking insect pests infesting pulse crops are mainly belonging to the insect orders; hemiptera (bugs, leafhoppers, aphids, etc.) and thysanoptera (thrips) (Table 3.1, Fig. 3.1). They possess specialised mouthparts (proboscis) allowing them to pierce/lacerate and suck the content of the plant tissues mostly from the phloem. Apart from insects, the mites of the order, Acarina also feed on the plant sap. These sap feeders reduce the plant vigour by draining the plant sap from leaves and other tender parts of pulse crops. The typical symptoms of sap feeders in pulses include yellow or brown discolouration of leaves, deformation of leaves, abortion or shedding of flowers, stunting and drying of plants. Mite infestation in pulses leads to curling and distortion of foliage. Some hemipetran sucking pests secrete honeydew causing the growth of sooty mould on the leaves. The pod sucking pest complex cause the shrivelling and reduction in grain size of the affected seeds. Apart from direct feeding damage, sucking pests including mites act as vectors of viral diseases and causing significant yield losses in pulses.

3.3.1

Pigeonpea

The insect pests feed on all the parts of pigeonpea, although those feeding on flowers, pods, and seeds are the most important biotic constraint affecting pigeonpea yield. More than 200 species of insect pests have been reported to feed on pigeonpea (Lateef and Reed 1990). However, the pests which occur continuously or attack at the middle or end of the crop cycle are considered to be economically important (Table 3.2) (Shanower et al. 1999).

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Table 3.1 Sucking pest infesting pulse crops Sl. no. Pests Insect pests 1 Pod sucking bug complex

2

Budsucking bug complex

3

Aphids

Taxonomic details

Host range

Pest distribution

Riptortus bugs: Riptortus pedestris F. R. dentipes F. (Alydidae: Hemiptera) Spiny brown bugs: Clavigralla gibbosa Spinola C. scutellaris (Westwood) C. tomentosicollis Stal. (Coreidae: Hemiptera) Giant coreid bug Anoplocnemis curvipes F. (Coreidae: Hemiptera) Lab lab bug: Coptosoma cribraria F. (Plataspidae: Hemiptera) Green stink bugs: Nezara viridula (L.) Piezodorus sp. Dolycoris indicus Stal. (Pentatomidae: Hemiptera) Campylomma spp. Creontiades pallidus (Rambir) Eurystylus spp. Taylorilygus vosseleri (Poppius.) (Miridae: Hemiptera) Aphis craccivora Koch A. fabae Scopoli Myzus persicae Sulzer Acyrthosiphon

Pigeonpea, lab lab, mungbean, cowpea and black gram

African and Asian countries including India

Reed et al. (1989), Sharma et al. (2010), Rao and Shanower et al. (1999), Singh and Van Emden (1979), Shanower et al. (1999), and Lal et al. (2005)

Pigeonpea

Africa and India

Dialoke et al. (2014), Reed et al. (1989), Kerzhner and Josifov (1999), and Schaefer and Panizzi (2000)

Most of the pulses

Africa and India, south and Central America, and Australia

Saxena (1978), Singh and Van Emden (1979), Patel and Srivastava (1989), Sharma

References

(continued)

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Table 3.1 (continued) Sl. no.

Pests

Taxonomic details

Pest distribution

References

Pigeonpea, cowpea, blackgram, beans and cowpea

India, USA and Africa

et al. (2010), Reed et al. (1989), Bock and Conti (1974), Bos (1971), and Brunt and Kenten (1974) Singh and Van Emden (1979)

Cowpea, mungbean, black gram and beans etc. Pigeonpea

America, Europe, Africa, Australia and Asia African and Asian

Most of the pulses

Central and southern India

Pigeonpea

Africa and Asia

Pigeonpea, wild pulse, Rhynchosia rothii

India (Gujarat, Tamil Nadu)

Host range

pisum Harris Macrosiphum spp. (Aphididae: Hemiptera)

4

Leafhoppers

5

Whitefly

6

Thrips

7

Cow bugs

8

Scales

9

Mealybugs

Empoasca fabae (Harris) E. kerri Pruthi Jacobiasca lybica (de Beryeven) (Cicadellidae: Hemiptera) Bemisia tabaci (Genn.) (Aleyrodidae: Hemiptera) Megalurothrips usitatus (Bagnall) (Thripidae: Thysanoptera) Otinotus oneratus W. Oxyrachis tarandus F. (Membracidae: Hemiptera) Ceroplastodes cajani Maskell Icerya purchasi Maskell. (Coccidae: Hemiptera) Coccidohystrix insolita (green) (Pseudococcidae: Hemiptera)

Srivastava and Singh (2009) and Reed et al. (1989) Oparaeke (2006), Palmer (1987), and Ogah (2011) Rao and Shanower et al. (1999), Sharma et al. (2010), and Pandey and Das (2013) Rao and Shanower et al. (1999) and Reed et al. (1989)

Patil et al. (1985), Rai et al. (1988), and Durairaj and Ganapathy (2000) (continued)

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Table 3.1 (continued) Sl. no. Pests Non-insect pests 10 Eriophyid mite

11

Red spider mites

Taxonomic details

Host range

Aceria cajani Channabasavanna. (Acarina: Eriophyidae) Schizotetranychus cajani Gupta Tetranychus urticae Koch: Acarina: Tetranychidae

Pigeonpea

Pest distribution Eastern Africa and South Asia. UP, Bihar, Gujarat, Tamil Nadu and Karnataka in India

References Reed et al. (1989) and Kannaiyan et al. (1984)

3.3.1.1 Pod Sucking Bug Complex Host Crops, Major Area of Incidence and Distribution Several species of bugs belonging to the families Aleyrodidae, Coreidae, and Pentatomidae of Hemiptera feeding on pods are commonly referred to as pod sucking bugs. Bugs belonging to genus, Anoplocnemis, Clavigralla (¼Acanthomia), and Riptortus are considered as serious pests (Shanower et al. 1999). Pod bugs infesting pulses are widely distributed in tropical and subtropical regions of the world, mainly along Africa and Asia (Reed et al. 1989). Clavigralla spp. being one of the emerging, severe pests of pigeonpea in Maharashtra and Karnataka (Sharma et al. 2010) and combined losses due to C. gibbosa and C. scutellaris in India vary among regions and occasionally exceed 50% (Bindra 1965). Green stink bug, Nezara viridula (Linn.) is also reported from most of the pulse growing areas in the country (Sharma 2016; Patra et al. 2016; Yadav et al. 2016). Apart from pigeonpea, pod sucking bugs often infest beans, mungbean, urdbean, and cowpea. Appearance, Biology, Nature of Damage and Symptoms The genus Clavigralla appears blue-grey in colour. The species, C. gibbosa and C. scutellaris are often confused together in the field but the later can be differentiated from the former with its robust body size during the adult stage and colour pattern of femur, tibia, and abdomen during nymphal stage (Hegde et al. 2012). These bugs lay the eggs in clusters on the leaf and pod surface. C. scutellaris lays eggs in clusters of 18–20 smooth and shiny eggs, whereas C. gibbosa deposits clusters of 10–12 sculptured eggs (Rao and Shanower 1999). The species belonging to Nezara usually appears green in colour and species of Riptortus are brown and more slender than other species. The adults of Anoplocnemis spp. are usually black or brown and largest among the pod sucking bugs. Pod sucking bug species oviposit eggs in clusters on the surface of leaves and pods. Most of these species require 4–5 weeks to complete one generation. Since adults live for more than 3 months, all the stages of their life cycle can be simultaneously observed on the pulse crops. Detailed biology is mentioned in Sect. 3.5.5.

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Giant coreid bug, A. curvipes

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Cow bugs, O.oneratus

Green stink bugs, N. viridula

Spiny brown bugs, C. gibbosa

Riptortus bugs, Riptortus spp.

Scale insects

Cowpea aphids, A. craccivora

Fig. 3.1 Sucking insect pests infesting pulses

Pea aphids, A. pisum

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Table 3.2 Sucking pests infesting pigeonpea Sl. no. 1 1.1 1.2

1.3 1.4 1.5

2

3

4

Common name Scientific name Pod sucking bug complex Riptortus Riptortus pedestris F., bugs R. dentipes F. Spiny Clavigralla gibbosa brown bugs Spinola C. scutellaris (Westwood) C. tomentosicollis Stal. Giant coreid Anoplocnemis curvipes bug F. Lab lab bug Coptosoma cribraria F. Green stink Nezara viridula (L.) bugs Piezodorus sp. Dolycoris indicus Stal. BudCampylomma spp. sucking Creontiades pallidus bugs (Rambir) Eurystylus spp. Taylorilygus vosseleri (Poppius.) Leafhoppers Empoasca fabae (Harris) E. kerri Pruthi Jacobiasc alybica (de Beryeven) E. kraemeri ross & Moore Cow bugs Otinotus oneratus W. and Oxyrachis tarandus F.

5

Mealybugs

Coccidohystrix insolita (green)

6

Scales

7

Thrips

Ceroplastodes cajani Maskell; Icerya purchase Maskell Megalurothrips usitatus (Bagnall)

Nature of damage

References

Both adults and nymphs feed by piercing the pod wall of pigeonpea and sucking the sap from developing seeds.

Singh and Van Emden (1979)

Both nymphs and adults attack vegetative and flower buds, and feed on the plant sap

Reed et al. (1989)

Both stages suck plant sap from tender plant parts, leaves and inject toxic saliva

Singh and Van Emden (1979)

Both adult and nymphs suck the sap from tender surfaces of shoots, stem and leaves

Yadav et al. (2016) and Pandey and Das (2013) Durairaj and Ganapathy (2000)

Crawlers congregate on leaves, stems and terminal shoots and suck the plant sap. Scales congregate on stems and suck fluid from tender stem, young shots and leaves Adults and nymphs suck the sap from floral parts

Rao and Shanower et al. (1999) Palmer (1987), Oparaeke (2006), and Ogah (2011) (continued)

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Table 3.2 (continued) Sl. no. 8 8.1

Common name Mites Eriophyid mites

Scientific name

Nature of damage

References

Aceria cajani Channabasavanna

Reed et al. (1989)

8.2

Red spider mites Two spotted spider mites

Schizotetranychus cajani Gupta Tetranychus urticae Koch

Both stages feed on lower surface of leaves and transmits devastating pigeonpea sterility mosaic disease (SMD) Adult and nymphs colonise the lower side of leaves by making silk webbings and suck sap from leaves

8.3

Kannaiyan et al. (1984)

Both adults and nymphs feed by piercing the pod wall of pigeonpea and suck the sap from developing seeds. Early nymphal stages feed by remaining in groups and later spread across plant parts. Symptoms include necrotised patches on leaves and pods which later turn dark and dry prematurely upon extreme damage (Singh and van Emden 1979). Grains in damaged pods develop dark patches, shrivel and become unsuitable for human consumption and sowing. Among the natural enemies, egg parasitoids, Gryon clavigrallae, and G. fulviventre reported to parasitise more than 55% of eggs of Clavigralla spp. in pigeonpea (Lal et al. 2005).

3.3.1.2 Bud-Sucking Bug Complex Major Area of Incidence and Distribution Species belonging to Campylomma, Creontiades, and Eurystylus are reported to attack pigeonpea in Africa (Dialoke et al. 2014) and Indian subcontinent, whereas Taylorilygus vosseleri is reported to attack pigeonpea only from African fields and it was reported as a pest of cotton later shifting on pigeonpea (Reed et al. 1989). Appearance, Biology, Nature of Damage and Symptoms Species of Creontiades are larger, measuring 8 mm long with a pale brown body (Kerzhner and Josifov 1999). Campylomma species are smaller, measuring 2 mm with greenish elongate and ovoid body. Eurystylus spp. and T. vosseleri are medium sized bugs (4 mm) with a mottle brown body (Schaefer and Panizzi 2000). Both nymphs and adults attack vegetative and flower buds, and feed on the plant sap. The affected plant parts develop dark spot and during a severe infestation, deformation of leaves and abortion of flower buds are observed (Reed et al. 1989). These bugs deposit eggs into the soft tissue of the plants. One generation of these bugs can generally be completed in a short time of up to 2 weeks.

3.3.1.3 Leafhoppers Host Crops, Major Area of Incidence and Distribution Leafhoppers are reported to attack pulses cultivated around the world. In India, Empoasca kerri is more severe on pigeonpea. Similarly, E. fabae and J. lybica are

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reported from America and Africa, respectively. E. kraemeri is another species from central and south America reported to attack beans and is also a minor pest of cowpea and other pulses (Singh and van Emden 1979). Apart from pigeonpea, leafhoppers also attack cowpea, blackgram, beans, and cowpea. Appearance, Biology, Nature of Damage and Symptoms Leafhoppers are small (2.5 mm), greenish insects that congregate on both upper and lower parts of the leaves and other tender parts of the plants. Female deposits eggs in the leaf veins on the underside of young leaves, on petioles, in the stem of young seedlings and egg period lasts for 7–10 days. There are five nymphal instars and nymphal development takes place in about 10 days. The adults are wedge-shaped, pale green insects whose life expectancy varies from 30 to 60 days (Singh and Allen 1979). The adults are more active and move diagonally or sideways on the plant surfaces and fly upon disturbance. Both nymphs and adults suck plant sap from tender plant parts, leaves, and inject toxic saliva. Affected parts show yellow discoloration along the leaf edges, followed by cupping of leaves. The typical symptoms include downward curling of leaves. Severely infested plants loose vigour and appear wilted and dry.

3.3.1.4 Cow Bugs (Otinotus oneratus W. and Oxyrachis tarandus F.: Hemiptera: Membracidae) Host Crops, Major Area of Incidence and Distribution Cow bugs are widely distributed across central and southern India. They are sporadic and of least economic significance (Rao and Shanower 1999) and generally infestation is found when the crop is 2 months old (Pandey and Das 2013). Appearance, Biology, Nature of Damage and Symptoms The adults appear brown to black in colour with a small body size (7 mm). The most prominent identification character is the presence of horn like projections on the thorax (Sharma et al. 2010). Bugs attract a lot of ants due to their sugary honeydew excretion. Adult bugs lay about 15–20 eggs in clusters on the stem. Both adult and nymphs suck the sap from tender surfaces of shoots, stem, and leaves (Yadav et al. 2016). The life cycle is completed in about a month under optimum conditions. Heavy infestations during the early growth of the crop may result in stunting and reduction of plant vigour.

3.3.1.5 Mealybugs (Coccidohystrix insolita (Green): Pseudococcidae: Hemiptera) Host Crops, Major Area of Incidence and Distribution Several species of mealybugs are reported to infest pigeonpea as well as other pulses in India, Trinidad, Africa, and Ghana (Singh et al. 2016). Incidence of C. insolita on pigeonpea is reported in Gujarat and Tamil Nadu.

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Appearance, Nature of Damage and Symptoms Mealybugs are soft bodied insects covered with white waxy coatings. Numerous crawlers are found in aggregation on leaves, stems, and terminal shoots. The affected leaflets turn yellow and drop off. The plant becomes stunted initially and severe incidence causes wilting and drying of plants. Honeydew excretion leads to the growth of sooty moulds on leaves and shoots, giving blackish appearance on leaves (Durairaj and Ganapathy 2000).

3.3.1.6 Scales (Ceroplastodes cajani Maskell; Icerya purchasi Maskell: Hemiptera: Coccidae) Host Crops, Major Area of Incidence and Distribution Several species of scale insects are known to infest pigeonpea crop in Africa and Asia. In India, Ceroplastodes cajani is the most common insect specific to pigeonpea, whereas Icerya purchasi is polyphagous and widely distributed species (Reed et al. 1989). Appearance, Biology, Nature of Damage And Symptoms The nymphs are mobile and can be dispersed by winds whereas adult females are sedentary and are found in colonies on the stem and occasionally on leaves. Scales are not major pests when pigeonpea is grown as an annual, but they build up and attack perennials (Rao and Shanower 1999). One generation can be completed in about 2–3 weeks. Scales suck the fluid from the tender stem, young shoots, and leaves of pigeonpea, and deposit honeydew which attracts ants towards the infested pigeonpea plant.

3.3.1.7 Thrips (Megalurothrips usitatus (Bagnall): Thysanoptera: Thripidae) Host Crops, Major Area of Incidence and Distribution This species attacks flowers of pigeonpea and beans in African and Asian countries. It causes severe damage to flowers and losses may reach up to 80–100% in pigeonpea (Oparaeke 2006). Appearance, Biology, Nature of Damage and Symptoms Thrips are very tiny insects (less than 1 mm), black in colour which are found crawling in and around floral parts (Palmer 1987). These thrips complete one generation within 3 weeks. Adults and nymphs suck the sap by lacerating floral parts. The infested flowers appear distorted, malformed, discoloured, and severe shedding of flowers takes place during severe infestation (Ogah 2011). Biology of thrips is detailed in Sect. 3.5.3.

3.3.1.8 Mites [Eriophyid Mites: Aceria cajani Channabasavanna (Acarina: Eriophyidae)]. [Red Spider Mites: Schizotetranychus cajani Gupta];

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[Two spotted spider mites: (Tetranychus urticae Koch) (Acarina: Tetranychidae)] Host Crops, Major Area of Incidence and Distribution Both eriophyid and red spider mites are spread over major pulse growing areas of eastern Africa and South Asia. Red spider mite, S. cajani reported to cause severe damage in India. Eriophyid mite is of more concern in Uttar Pradesh, Bihar, Gujarat, Tamil Nadu, and Karnataka due to its involvement in the transmission of pigeonpea sterility mosaic disease (Reddy et al. 1989). Appearance, Biology, Biology, Nature of Damage and Symptoms Adults of red spider mite appears velvet red with oval body shape. Adult female lays yellowish eggs within the webbings made by them. These mites form wind balls, a mechanism of dispersion with the help of winds. Adults and nymphs colonise the lower side of leaves by making silk webbings and suck sap from leaves causing yellow or white spots. Heavy infestation leads to defoliation (Reed et al. 1989). Spider mites are minute arthropods and vary in colour (green, greenish yellow, brown, or orange-red) with two dark spots on the body. Egg period is 2–4 days. Upon hatching, spider mites will pass through a larval stage and two nymphal stages (protonymph and deutonymph) before becoming adults. The life cycle is completed in 1–2 weeks. There are several overlapping generations in a year. The adult lives up to 3–4 weeks (Srinivasan 2014). Eriophyid mites are microscopic (0.2 mm) and appear pinkish with spindle shaped body. Females lay milky white eggs on vegetative terminals. They spread from plant to plant via wind currents. Eriophyid mites have a very short life cycle of about 2 weeks comprising egg and two nymphal stages (Oldfield et al. 1981). Both stages feed on the lower surface of leaves and cause severe crop losses due to the transmission of Pigeonpea Sterility Mosaic Virus (PPSMV) of the genus Emaravirus causing a devastating pigeonpea Sterility Mosaic Disease (SMD). SMD reduces the greenish plants to light green, later they turn to chlorotic patches resulting in mosaic patterns and disease is dubbed as the ‘green plague’, as the infected plants remain in the vegetative state without flower production (Patil and Kumar 2015). Patches of bushy pale and green plants with smaller stunted leaves, without flowers or pods and profuse branchings are the common field symptoms. During early stages (50%) at 24 h after application at 16% concentration (Fig. 6.4).

6.4.6.2 Mechanical Control Yellow sticky trap installation @ 12 traps/ha to monitor the alate adult population buildup is helpful in the management of cabbage aphid. As per Southwood (1978) sticky traps measure aphid populations more readily than other labour-intensive absolute monitoring methods, as traps continuously catch and retain insects without regular human involvement. For kitchen vegetable growers, aphids can be repelled by planting crops with reflective mulch-covered beds and monitoring aphid by yellow water pan traps, if aphid population seems threatening (above ETL) additional control measures may be initiated (Griffin and Williamson 2012).

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6.4.6.3 Biological Control Parasites and predators are known to regulate aphid populations naturally. The parasitic wasp D. rapae lays eggs within half-grown nymphs (preferring second to fourth instars over first instar nymphs and adults) and mummifies them, forming a light brown and hard shell. Although D. rapae is a very common parasitoid it is not always effective in controlling aphids. When the wasp populations are large enough to control aphids, the aphid population has usually exceeded damage thresholds (Pal and Singh 2012). One more reason may be nontarget effects of pesticides or natural suppression of D. rapae by many hyperparasitoids (Pal and Singh 2013). Protecting habitat that will foster the population and survival of natural enemies can help reduce the need for pesticides (Natwick 2009). D. rapae has been reported as the most effective natural enemy against B. brassicae (George 1957) with parasitism as high as 72.0% in the Netherlands (Hafez 1961), 76.0% in Kenya (Bahana and Karuhize 1986) and 51.5% in India (Halder et al. 2014). Zhang and Hasan (2003) reported that the parasitoid D. rapae accepted all stages of the cabbage aphid with an average number of 42.8 mummies produced per female parasitoid and the parasitism rate was 6.7 and 1.4% in the treated and control plots, respectively. Vaz et al. (2004) reported high natural parasitism of B. brassicae by D. rapae (93.2%) and Aphidius colemani (4.5%). Duchovskien and Raudonis (2008) investigated that D. rapae reduced the populations of cabbage aphid by 23.9–26.2% on cabbage plants in Italy. Syrphid fly maggots, ladybeetle adults and larvae and lacewing larvae (aphid lions) are common predators of B. brassicae on canola (Nieto et al. 2006; Hainan et al. 2007; Gill et al. 2013). Sable et al. (2008) reported 76.43 and 82.33% parasitism of B. brassicae by Aphidius spp. in cauliflower. Coccinella septempunctata, Chrysoperla carnea, Ischiodon scutellaris and D. rapae and other biocontrol agents have been reported from various parts of India on mustard and cabbage aphids (L. erysimi and B. brassicae) (Agarwala and Raychowdhary 1981; Mathur 1983; Akhtar et al. 2006; Rana 2006; Sayed and Teilep 2013). Parmar et al. (2008) reported that the mortality in all nymphal instars of L. erysimi ranged from 44.7 to 75.2% after 10 days of spraying when fed with V. lecanii @ 4.0 g/l treated mustard leaves. Many entomopathogenic fungi like Beauveria bassiana, Metarhizium anisopliae and Verticillium lecanii have been reported effective against B. brassicae (Ujjan and Shahzad 2012). Kotwal and Bhalla (1985) utilized entomophthora in natural control of B. brassicae and found fungal infection to vary from 0.7 to 25.4% with a mean of 9.7%. B. bassiana proved much effective against B. brassicae with 100% mortality (Embaby and Lotfy 2016) compared to M. anisopliae (91.11% mortality) after 10 days of treatment. B. bassiana (Bb-5a) showed superior effect in the suppression of B. brassicae under field conditions (Ramanujam et al. 2017). Wagle et al. (2006) reported that Coccinella transversalis on average devoured 38.8 B. brassicae per day. Thakur and Sharma (2014) in organic cabbage cultivation studies reported that V. lecanii @ 108 conidia/ml resulted in 63.3% mortality compared to in untreated check after 72 h of exposure. Release of C. septumpunctata @ 90 eggs/m2 resulted in higher reduction of B. brassicae population (42.0–89.8%) compared to 30 eggs/m2 releases (31.7–81.3%) during winter months in alfalfa crop in Saudi Arabia (Al-Solami et al. 2016). Laboratory studies by Jesu Rajan et al. (2018) confirmed

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that C. transversalis consume 2695–2773 B. brassicae adults during its total life span.

6.4.6.4 Botanicals Botanical plants are readily accessible to marginal farmers in their locality and have been found to be environmentally safe having low toxicity to mammals, natural enemies and pollinators (Delvin and Zettel 1999). Rana Bhat and Yubak Dhoj (2005) indicated that the 1:5 concentrated solution of Bakain (Melia azedarach L.) extract at 5 day intervals effectively lowered the population of B. brassicae on cabbage in Nepal. Seed extract of the Chinaberry tree, Melia azedarach (Mekuaninte et al. 2011; Kibrom et al. 2012), leaf extract of peppermint, Mentha piperita, seeds and leaf extract of flowering lantana, Lantana camara (Baidoo and Adam 2012) have showed promising results against cabbage aphid. Tagetes minuta crude extract (100%) (Phoofolo et al. 2013) resulted in 25–80% mortality of adult forms of B. brassicae. Duchovskiene (2005) reported that Neem Azal-T/S @ 0.5% water spraying solution effectively controlled B. brassicae. Essential oils from various plants like Nepeta cataria (Motazedian et al. 2014), Veronica and Agrimonia (Gorur et al. 2008), Foeniculum vulgare and Laurus nobilis (Mustafa and Gorur 2009), Mentha piperita (Birhanu et al. 2011), Melia azedarach and Croton macrostachyus (Michael and Raja 2012) significantly reduced the reproduction potential of the cabbage aphid and resulted in higher mortality. Thakur and Sharma (2014) in organic cabbage cultivation studies reported that neem oil (@ 0.3%) resulted in 100% mortality followed by 81.1% mortality (24.3 aphid) in aqueous extract of Eupatorium (@10%) and 56.7% in each Panchagavya and Lantana extract. Spraying cabbage plants with garlic and pepper extracts (Baidoo and Mochiah 2016) reduced B. brassicae infestation by 42.1 and 26.4%, respectively. Shiberu and Negeri 2016 obtained 59.5, 57.2 and 52.5% mortality of B. brassicae by Azadirachta indica seed, Dodonaea angustifolia and Cymbopogon citrates, respectively, after third day observation in first spray. Peris and Kiptoo (2017) reported 69.5, 65.6 and 53.7% mortality of B. brassicae adults in kale crop by extracts of garlic, ginger and Mexican marigold, respectively, after 3 weeks. Priyanka (2017) reported 85.2% reduction in B. brassicae population third day after the spray of citronella (Cymbopogon citrates) oil in cabbage crop. 6.4.6.5 Host Plant Resistance All season, Red Drum Head, Sure Head, Express Mail are some B. brassicae tolerant varieties of cabbage in India. Three year experiments by Lal (1991) at Kullu, India revealed that none of the cabbage varieties was immune to B. brassicae and three of those (All Season, Red Drum Head, KK Cross) showed only moderate resistance. Jahan et al. (2013a, b) concluded that canola cultivars, namely Smilla and Buris increased the efficiency of D. rapae in the control of the B. brassicae. Studies on resistance of Brassica genotypes to B. brassicae have also been carried out by several workers (Ellis et al. 1998; Zandi-Sohani et al. 2004; Jamshidi-Kaljahi et al. 2006).

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6.4.6.6 Chemical Control Most of the farmers have continually used different types of synthetic pesticides that are systemic and broad spectrum to manage aphids, including methomyl, emamectin benzoate, pyrethroids and neonicotinoids (Gill and Garg 2014). Extensive application of these insecticides has led to development of resistance in different parts of the world (Garg et al. 1987; Sweeden and McLeod 1997; Ahmad and Aslam 2005). Nale et al. (2016) reported resistance factor of acetamiprid, imidacloprid, thiamethoxam and dimethoate was 85.06, 137.80, 13.50 and 44.17 folds, respectively, in B. brassicae infesting cabbage. Development of rapid resistance to insecticides (Ahmad and Akhtar 2013) plays significant roles in the ecological and economic importance of this species. To achieve maximum control with minimum efforts, proper surfactant proportion in combination with well-adjusted spray equipment is important (Kessing and Mau 1991) because of the waxy nature of the pest and crop. Moreover, application dose and timing are very important to keep aphids under control while conserving populations of natural enemies (Hines and Hutchison 2013). Furthermore, these bear high costs which make them unaffordable to middle class or marginal farmers. Insecticide resistance studies in cabbage aphid carried out by Ahmad and Akhtar (2013) in Pakistan report that B. brassicae has developed resistance to number of chemicals, namely Methomyl, Emamectin Benzoate, Cypermethrin, Lambda cyhalothrin, Bifenthrin, Deltamethrin, Imidacloprid, Thiamethoxam, etc. Hence, their judicious use is advocated. Abo El-Ghar and Abd El-Ghany (1989) reported that Fenvalerate (71 g/ha) showed satisfactory effectiveness in reducing B. brassicae adult population by 77% on treated cabbage plants and reduced the F1 progeny of D. rapae by only 36.4% compared to 70.5 and 58.3% reduction, respectively, by Methomyl and Cypermethrin at same dose. Hence, non-chemical methods need to be applied for better management of this pest. Thiamethoxam SP used by Ali and Zedan (2015) resulted in 93.6, 89.3 and 77.6% reduction in B. brassicae population on cauliflower crop in Egypt after 3, 5 and 7 days, respectively, with minimum negative effects on Coccinella undecimpunctata and Scymnus spp. Srivastava et al. (2016) reported that Acetamiprid 20%SP @150 g/ha showed best control of B. brassicae (83.05%) followed by Acetamiprid @100 g/ha (81.04%) and Acetamiprid @75 g/ha (77.97%) in cabbage. Khan et al. (2017) demonstrated that insecticide pyriproxyfen had very good efficacy in reducing the B. brassicae population in cabbage. 6.4.6.7 Integrated Pest Management Studies Farooq and Tasawar (2006) in an integrated approach reported 98.6, 93.0, 83.5 and 18.0% reduction in aphid population on canola in Pakistan by using methomyl, blank spray of water + release of both C. carnea and C. septempunctata, respectively, after 7 days of treatment. Mahmoud and Osman (2015) reported that seed treatment with Actara 25%WG (Thiamethoxam, 3.5 g a.i/kg seed) + Salicylic Acid (SA) (250 mg/lit) resulted in maximum B. brassicae reduction (0.8–30.4 mean number of aphids/10 cm terminal shoot length) followed by Gaucho70% WS (Imidacloprid, 5 g a.i/kg seed) + SA compared to untreated control (11–393 aphids/10 cm terminal shoot length) in canola crop. Sulfoxaflor treatments caused

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about 90% reduction in B. brassicae followed by pyrifluquinazon and B. bassiana, which caused about 80 and merely 19% reduction, respectively, in broccoli (Dara 2017).

6.5

Green Peach Aphid/Peach Potato Aphid Myzus persicae (Sulzer) (Hemiptera: Aphididae)

While these two above mentioned aphid species are specialist feeders, green peach aphid/peach potato aphid, Myzus persicae is a generalist feeder. It has a wide range of host plants, including very many economically important crops such as peach and potato. Among the brassicas, turnip seems to be a particularly favourite host, but the aphid is considered to be a very important pest of cabbage. Alternative hosts include capsicums and other solanaceous plants. It is virtually cosmopolitan, but it is more abundant in temperate regions than in the tropics. The greater economic importance of this species is due to its role as a vector of plant viruses. The species has been reported to transmit more than 100 plant viruses including potato virus Y and potato leaf roll virus to the plants from family Solanaceae and various mosaic viruses such as the persistent beet western yellow virus (Ponsen 1972; Eskanderi et al. 1979; Bwye et al. 1997). It is of minor significance to cruciferous oilseeds (Fig. 6.5). Adults are 1.2–2.3 mm long. Wingless forms are usually uniformly green in colour with a darker thorax. Antennae are two-thirds as long as the body. Cornicles are fairly long. Damage is by direct feeding and by virus transmission. Direct feeding by aphid colonies causes leaf curl, discolouration, stunted growth and even death of the infested plants. Seed set is also reduced. In heavy infestation, copious amounts of honeydew are produced on which sooty mould fungus grows. This reduces the quality of the crop. The threshold for M. persicae is correspondingly lower, at 25% of plants infested (Bailey 2007). Early sowing (25th October) has been reported as an escape method in gobhi sarson against M. persicae by Prasad et al. (2013) in Himachal Pradesh. Loureiro and Fig. 6.5 Myzus persicae nymph. Source: http://www. nbair.res.in/insectpests/index. php (assessed on 16.01.019)

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Moino (2006) reported cent per cent mortality of peach potato aphid by M. anisopliae and B. bassiana both at 107 and 106 spores/ml. In a study conducted by Sarwar (2013) in Pakistan, three genotypes of mustard, namely NM-1, NM-2 and NM-3, were adjudged completely resistant with no population of aphids. Repellent action of trap crops like Ocimum basilicum, Allium sativum, Lavandula latifolia, etc., has been utilized in Brassica against M. persicae which reduced the host finding ability (Lai et al. 2011; Potts and Gunadi 1991; Ben Issa et al. 2017). White et al. (1995) utilized Phacelia tanacetifolia as companion crop to attract hoverflies leading to reduced infestation by the M. persicae in cabbage crops. Early season populations of M. persicae on broccoli were lower when interplanted with living mulch of clovers (Trifolium spp.) compared with clean cultivation (Costello 1992; Costello and Altieri 1995). The most common natural enemies attacking aphids are the convergent ladybeetle Hippodamia convergens, seven-spotted ladybeetle C. septempunctata, green lacewing C. carnea and the parasitoid wasps Lysiphlebus testaceipes and D. rapae (Giles et al. 2003; Jones 2005; Jessie 2013). Etheric extract (10%) of leaves of three plants, namely Artemisia herba-alba, Eucalyptus camaldulensis and Rosmarinus officinalis, was effective and caused significant mortalities of M. persicae (100%, 53% and 60%, respectively) at the highest concentration (Nia et al. 2015). Faraone et al. (2015) revealed that the toxicity of imidacloprid against M. persicae was synergized 16- to 20-fold by L. angustifolia and T. vulgaris essential oils. Sulfoxaflor at high rate (2.0) caused a 73% reduction in M. persicae followed by sulfoxaflor low rate (1.5) (47%) and B. bassiana (40%) in broccoli (Dara 2017).

6.5.1

Clearcut Identification Clues for Three Aphis Species

The cabbage aphid is difficult to distinguish from the turnip aphid. The cabbage aphid is 2.0–2.5 mm long and covered with a greyish waxy covering, but the turnip aphid is 1.6–2.2 mm long and has no such covering (Carter and Sorensen 2013). The cabbage aphid and green peach aphid can be confused when they are both found feeding in the same field. However, they can be differentiated morphologically. For instance, the cabbage aphid is waxy with short cornicles. On the other hand, the green peach aphid lacks a waxy covering and possesses long cornicles (Opfer and McGrath 2013).

6.6

Painted Bug, Bagrada hilaris (Burmeister) (Hemiptera: Pentatomidae)

6.6.1

Distribution and Host Range

This insect often infests rapeseed-mustard. It is native to Africa. These are found throughout South-Eastern Africa, Southern Asia and Southern Europe (Hill 1975). It is an important pest of crucifer crops in India (Rai 1976; Patidar et al. 2013) and

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abroad throughout Asia, Africa, some Islands of southern Europe such as Malta and Pantelleria (Italy), California and Arizona (Palumbo and Natwick 2010; Palumbo et al. 2016), New Mexico (Bundy et al. 2012), Texas (Vitanza 2012), Nevada (Perring et al. 2013), Utah (Reed et al. 2013) and Hawaii (Matsunaga 2014). It has also been reported from Mexico (Sanchez-Pena 2014) and Chile (Faundez et al. 2016). The species feeds on various wild and cultivated plants, but it is a severe pest of cole crops in the genus Brassica within its native and invaded range (Howard 1907; Obopile et al. 2008; Abrol 2009; Perring et al. 2013; Reed et al. 2013; Palumbo et al. 2016) during winter where it causes considerable damage (Batra 1958; Sandhu 1975). This insect is more prevalent in lighter soils of drier region in the state of Rajasthan, Punjab, Haryana, Uttar Pradesh and Madhya Pradesh. Besides crucifers, it is known to feed on sugarcane, rice, indigo and coffee (Figs. 6.6 and 6.7).

6.6.2

Identification

6.6.2.1 Eggs Females lay oval or barrel, pale-yellow eggs singly or in groups of 3–8 undersides of the leaves, on stalks, pods and sometimes in soil near base of the plants (Palumbo and Natwick 2010). A single female can lay as many as 100 eggs within 2–3 weeks. The incubation period is 5–8 days. Eggs have an opaque, white or light red hue and Fig. 6.6 Bagrada hilaris infestation

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Fig. 6.7 Symptoms of Bagrada hilaris infestation

nymphs emerge after 3–4 days under optimal conditions (Reed and Perring 2012). Size of the eggs varies from 0.5 to 0.6 mm  0.8 to 1.0 mm (Rakshpal 1949; Verma et al. 1993).

6.6.2.2 Nymphs Bagrada bugs have five nymphal stages. Newly emerged first instar nymphs are bright red with slightly dark pronotum, legs, head and antennae. First instars remain near eggs and do not feed until after moulting into second instar. Later instars turn darker in appearance with prominent markings. Initially they do not have wings; wings develop gradually as the nymphs grow. Wing pads are visible in the last nymphal instar. The length and breadth of first, second, third, fourth and fifth instar nymphs were 1.12 and 0.77, 1.39 and 1.09, 1.50 and 1.45, 2.45 and 1.69 and 5.29 and 3.04 mm, respectively (Verma et al. 1993). 6.6.2.3 Adults The adult body is shield shaped. The adult bugs are 5–7 mm long and 3–4 mm wide. Females are slightly larger than males. They are subovate, black and have a number of orange or brownish spots, with red and yellow marking running lengthwise. The females measured 7.12 mm in length and 3.94 mm in width, whereas the male was 5.29 mm long and 3.04 mm across width (Verma et al. 1993).

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Life Cycle

A female bug may lay 37–102 eggs during its life span of 3–4 weeks. The eggs hatch in 3–5 days during summer and 20 days during winter months. The nymphs develop fully in five stages and transform themselves into adults in 16–22 days during the summer and 25–34 days during the winter. The entire life cycle is completed in 19–54 days and it passes through 9 generations per year. Tiwari and Saravanan (2009) recorded that the female bugs laid 98.6  14.1 eggs during its life span and incubation period was 5.8  0.6 days. The average duration of nymphal period was 16.6  1.3 days. Male and female longevity were 19.6  1.6 days and 25.5  2.0 days, respectively. The total life cycle completed within 22–40 days. Laboratory studies conducted by Nagar et al. (2011) on the biology of B. hilaris revealed that the mean number of eggs laid by female were 70.8  6.1 on soaked mustard seed. The average pre-oviposition period, oviposition period, incubation period and total developmental period were found to be 6.2  0.1, 3.6  0.1, 16.2  0.6 days and 18.6  0.36 days, respectively. As per Shrikhandia and Tara (2016) the duration of eggs lasted for an average of 6.0  0.8. There were five nymphal instars which completed their development in 17.8  3.2 days. The total life cycle from egg laying to adult emergence was completed in 19–28 days with an average of 23.8  3.5 days. The female lived longer (26.5  1.1 days) than the male (20.3  1.2 days). Pawar et al. (2009) indicated that maximum temperature and morning relative humidity were highly negative significant, whereas evening relative humidity and wind velocity showed highly positive significant correlation with the pest population. Adult activity peaks up when day temperature is above 25  C and below 41  C and they seek shelter near soil surface when temperatures are not in this range. Adults are known to overwinter on several cruciferous weeds and may survive summers likewise (Singh and Malik 1993; Reed et al. 2013). However, Bhati et al. (2015) observed the painted bug on Brassica crops at two distinct stages of crop growth, first at seedling stage and second at pod bearing to maturity stage, escaping flowering stage of the crop growth. Several studies on the biology of B. hilaris have been undertaken under diverse environmental conditions (Rakshpal 1949; Singh and Malik 1993; Singh 1996; Ghoshal et al. 2006; Nagar et al. 2011). It is found active during seedling stage (October–November) (Vora et al. 1985) and at harvest stage (March–April) (Singh and Malik 1993; Singh 1996). Painted bug exhibited highly significant negative association with maximum, minimum and mean temperature, whereas significant negative correlation was observed with evaporation at Junagarh, Rajasthan (Divya et al. 2015).

6.6.4

Nature of Damage

This insect attacks the crop at two stages in the season, i.e. at initial or seedling stage in October–November and at maturity in March–April. Both nymphs and adults suck sap from the plants at seedling stage and oil from seed at maturity stage of the crop. Bugs can also be seen feeding on the harvested material lying on the threshing

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floor. The painted bug is highly polyphagous pest and is more serious on young/ seedling stage. The infestation of this pest in the vegetative growth stage results in first whitening of leaves, then wilting leading to complete drying of the plant. In both the cases resowing of the crop becomes necessary. Loss of sap from the leaves and developing pods results in gradual wilting and drying, curling of the pods and shriveling of grains. Severe attack at seedling stage may even kill the plants and bear a brunt-up look. The pest incidence at seedling stage may result in complete failure of the mustard crop necessitating resowing (Bakhetia 1987; Singh et al. 1993a, b). These feed on both surfaces of the leaves, presumably inject saliva to break the inner leaf tissues (Palumbo and Natwick 2010). The nymphs and adult bugs also excrete a sort of resinous material, which spoils the pods.

6.6.5

Extent of Losses

A severe infestation of this pest was reported by Srivastava et al. (1972) in the plains of India which forced resowing in Haryana, Rajasthan and Delhi. Dhaliwal and Goma (1979) observed 30–72% damage in cauliflower seed crop due to painted bug feeding on leaves, buds, flowers and developing seed pods during 1973–1974 in Himachal Pradesh. The losses attributed at seedling stage due to painted bug attack varied from 26.8 to 70.8%. The attack at the pod formation and maturity stages is much more alarming as it results in losses to the tune of 30.1% in yield and 3.4% in oil content (Singh et al. 1980). Due to its infestation during seedling stage in rapeseed-mustard crops, the seed yield losses were reported to be 26.8–70.4% by Joshi et al. (1989) from Rajasthan. In all brassica crops tested, feeding damage increased with greater numbers of B. hilaris adults caged on cotyledon and 2-leafstage plants. Significantly more feeding damage occurred on the upper (younger) two leaves than on the lower (older) two leaves of the 4-true leaf plants for all host plants suggesting that B. hilaris feeds preferentially on newer leaf tissue. Significant reductions in leaf area, relative chlorophyll content and dry weight in all crops indicated negative impacts on plant growth by B. hilaris. Moreover, cotyledon and 2-leaf plants were more severely impacted by B. hilaris induced injury (Huang et al. 2014).

6.6.6

Integrated Pest Management

6.6.6.1 Monitoring Early detection is crucial to contain this pest as bagrada bug populations tend to increase rapidly under favourable conditions. Monitoring should begin before planting by scouting and inspecting areas around the field intended for production. Sweep netting and careful visual observation of the soil surface, weeds, grasses and other vegetation in the vicinity of field are essential to determine if bagrada bugs are present. Fields surrounded by weedy areas with an abundance of wild mustards or other host plants may be at a higher risk for bagrada bug infestation. Regularly

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inspect all plants in the field, as well as all plant shipments coming in or going out of the operation. Observations or collecting bagrada bugs in the field should be done during the day and done with care because disturbing the plants will cause bagrada bugs to drop to the soil surface or fly away. Blacklight trapping is not effective; however, visual observation at night with a fluorescent lamp is effective (Huang et al. 2014). Scouting should coincide with peak activity occurring during the warmer parts of the day to improve accuracy of field sampling.

6.6.6.2 Cultural Practices Deep ploughing in summer is essential to destroy the eggs and application of first irrigation at 3–4 weeks after sowing for effective control of this bug and irrigation reduces the chances of wilting of the crop due to desapping by bug. Timely sowing of crop to protect the crop at seedling stage; clean cultivation by weeding, hoeing and destroying of debris in and around the field is effective in the management of painted bug (Singh et al. 1993a, b; Singh and Malik 1993). Quickly thresh the harvested material and dispose of the crop refuge to reduce the infestation (Bakhetia and Sachan 1997). Covering well irrigated bare soil with a thin sheet of clear plastic for several weeks during warm weather will control hatching nymphs and may also control eggs if the soil temperature is high enough. 6.6.6.3 Biological Control There are known predators and parasitoids that attack B. hilaris (Bundy et al. 2018) but there are no established biological control programs in practice. In Pakistan, B. hilaris is only a sporadic pest, likely due to a suite of natural enemies like Trissolcus hyalinipennis Rajmohana & Narendran, Gryon sp. and Ooencyrtus sp. (Mahmood et al. 2015). Recently, Ganjisaffar et al. (2018) reported T. hyalinipennis parasitizing painted bug from North America. The tachinid fly, Alophora spp. has been reported from India and other countries. This egg parasitoid can be preserved by provision of alternate hosts, like weeds namely Lambsquarters, purple nutsedge, Euphorbia spp., perennial sowthistle and field bindweed; and crops like corn, sorghum, sunflowers, papaya, potato, cotton and some legumes. 6.6.6.4 Botanicals Leaf extract of Argemone mexicana effectively controlled painted bug both in field and laboratory conditions (Pandey et al. 1981). Chandel et al. (2011) revealed that extracts of Azadirachta indica and Vitex negundo irrespective of concentrations (0.5, 1.0 and 2.0%) were more effective, causing 80.9 and 74.9% mortality of nymphs and adults of B. hilaris, respectively. Guarino et al. (2018) revealed that diterpene hydrocarbon present in both of the preferred hosts (B. oleracea and B. napus) acts as a good candidate for use in lures for monitoring B. hilaris in the field. 6.6.6.5 Chemical Control There are several strategies that contribute to the integrated management of B. hilaris (Palumbo et al. 2016; Bundy et al. 2018), with chemical control being the anchor of these tactics. Contact foliar insecticide applications are the most effective during the

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afternoon and early evening during the height of insect activity on plants (Chauhan and Yadav 2007); however, because bagrada bugs fly away when disturbed, adults can easily escape before contact with the insecticides and return later (Reed et al. 2012). Additionally, bagrada bugs may drop to the soil to avoid contact with insecticides. At the time of crop germination if one nymph or one adult is present in 1 m row length of the crop area, spraying the crop with 200 ml Malathion 50 EC in 200 l of water per acre is quite effective. If required, at the time of harvesting spray the crop with 400 ml Malathion 50 EC in 400 l of water per acre. Studies conducted by Chauhan and Yadav (2007) revealed that Fenvalerate 0.4 D @ 8 kg per acre effectively checked painted bug infesting turnip. In another trial on painted bug infesting raya, Monocrotophos 0.04% was found to be the best followed by Endosulfan 0.07%. Ahuja et al. (2008) revealed that sowing of mustard seed treated with Imidacloprid @ 5–7 g/kg in second fortnight of October in dry soil followed by irrigation resulted in lower plant damage (4.9–5.8%) due to B. hilaris. Nagar et al. (2011) observed that Malathion 50 EC @ 500 ml/500 l of water was most effective in reducing damage by painted bug. Singh et al. (2011) revealed that seed treatment with Imidacloprid 70 WS @ 5 g/kg seed or Thiamethoxam 70 WS @ 5 g/kg seed with application of Endosulfan 4% dust @ 10 kg/ha provided significant reduction in bagrada population and higher yield. Ratnoo et al. (2018) reported minimum bug population in Triazophos 40 EC @ 1 l/ha treatment followed by Imidacloprid 17.8 SL @ 150 ml/ha and Acephate 75 SP @ 500 g/ha after two sprays. Thomas et al. (2018) tested mustard pods and young branches @ 1:10 concentration of nanoformulations of NSKE, copper, silver and manganese in comparison with conventional chemicals like Imidacloprid. The highest per cent mortality was found in the population treated with Imidacloprid as (97.8  0.7) over control (2.4  1.0). Among all nano-formulations, per cent mortality was recorded maximum (84.4  0.6) in the population treated with silver at 1:10 concentration on the fourth day of application followed by copper (77.8  0.6), manganese (73.3  1.0) and neem seed kernel extract (NSKE) (68.9  1.5). Integrated Pest Management of Sucking Pests in Nutshell • Timely sowing of the crop before 15th October helps to escape infestation of mustard aphid. • Apply balanced dosages of fertilizers, as application of only nitrogenous fertilizers makes the crop vulnerable to aphids. • Seed treatment with Imidacloprid 70 WS @ 5.0 g/kg of seed should be done for management of painted bug and other insects during early stage of crop. • Plough the field in summer season and follow the clean cultivation by weeding, hoeing and burning of debris in and around the field. • Plucking and destroying the infested twigs or dipping them in kerosenized/ insecticide treated solution from the border rows by 2–3 times at 10 days interval is very functional to avoid the further increase of insects in the crop season. • Foliar application of insecticides, such as oxydemeton methyl (Metasystox) 25 EC or dimethoate (Rogar) 30 EC @ 1 ml/l of water, is done when 26–28 aphids/plant is observed.

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• If the pest population builds up again, insecticidal spray can be repeated at 15 days interval or 5% NSKE and 2% neem oil sprays are also effective to manage mustard insects. • In case of severe infestation spray the crop with Endosulfan 35 EC, Phosphamidon 85SL, Quinalphos 25 EC, Malathion 50 EC @ 1–1.5 ml/l of water. • In eco-friendly management/Verticillium lecanii/@ 108 CS/ml of water in combination with 5% NSKE, 2% neem oil is effective in reduction of aphid population. • Dimethoate @ 1 ml/l followed by C. septempunctata @ 5000 beetle/ha or NSKE @ 5% + V. lecanii @ 108 CFU ml 1 or neem oil @ 2% + C. carnea @ 50,000 larvae/ha are very effective strategies for aphid control. • Spraying should be done in the afternoon for avoidance of toxicity to pollinators. • Thresh the crop as early as possible to avoid the further losses and dispose of infected plant debris immediately.

References Abo El-Ghar GES, Abd El-Ghany ME (1989) Impact of two synthetic pyrethroids and methomyl on management of the cabbage aphid, Brevicoryne brassicae (L.) and its associated parasitoid, Diaeretiella rapae (McIntosh). Pest Manag Sci 25(1):35–41 Abrol DP (2009) Plant-insect interaction. In: Gupta SK (ed) Biology and breeding of crucifers. CRC Press, Boca Raton, FL, pp 129–150 Adhab M, Schoelz JE (2015) Report of the turnip aphid, Lipaphis erysimi (Kaltenbach, 1843) from Missouri, USA. J Plant Prot Res 55(3):327–328 Afrin S, Latif A, Banu NMA, Kabir MMM, Haque SS, Emam Ahmed MM, Tonu NN, Ali MP (2017) Intercropping empower reduces insect pests and increases biodiversity in agroecosystem. Agric Sci 8:1120–1134 Agarwal N, Rohilla HR, Singh H (1996) Evaluation of rapeseed-mustard genotypes against mustard aphid, (Lipaphis erysimi Kalt.) at inflorescence stage. Ann Biol 12:93–95 Agarwala BK, Raychowdhary DN (1981) Notes on some aphid affecting economically important plants in Sikkim. Ind J Agric Sci 51:690–692 Aggarwal M, Haseeb M, Manzoor U (2014) Biology and seasonal incidence of aphid, Brevicoryne brassicae on cabbage. Ann Plant Prot Sci 22(2):275–277 Agnihotri A, Prem D (2007) Oils quality improvement in rapeseed-mustard, p 33–43. In: Souvenir, National seminar on changing global vegetable oils scenario: issues and challenges before India, held during January 29–31, 2007, at Hyderabad, p 91 Ahmad M, Akhtar S (2013) Development of insecticide resistance in field populations of Brevicoryne brassicae (Hemiptera: Aphididae) in Pakistan. J Econ Entomol 106:954–958 Ahmad M, Aslam M (2005) Resistance of cabbage aphid, Brevicoryne brassicae (Linnaeus) to endosulfan, organophosphates and synthetic pyrethroids. Pak J Zool 37(4):293–295 Ahmad S, Aftab A, Sabri MA, Ghazal A, Ullah Z, Ahmad MI (2017) Efficacy of four different insecticides with different mode of action against canola aphid (Lipaphis erysimi) under field condition. J Entomol Zool Stud 5(3):626–629 Ahmed S, Cheema SA, Zubair M, Abbas Q, Bashir MR, Malik K, Aslam A, Maan NA (2018) Comparative efficacy of insecticides against mustard aphid in Brassica juncea. Int J Entomol Res 3(3):34–37

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Tiwari SK, Saravanan L (2009) Life history and seasonal activity of painted bug, (Bagrada hilaris) (Burm.) infecting mustard in Allahabad Uttar Pradesh. Pestology 33(2):21–23 Tsaganou FC, Hodgson CJ, Athanassiou CG, Kavallieratos NG, Tomanovic Z (2004) Effect of Aphis gossypii Glover, Brevicoryne brassicae (L.), and Megoura viciae Buckton (Hemiptera: Aphidoidea) on the development of the predator Harmonia axyridis (Pallas) (Coleoptera: Coccinellidae). Biol Control 31:138–144 Ujjan AA, Shahzad S (2012) Use of entomopathogenic fungi for the control of mustard aphid (Lipaphis erysimi) on canola (Brassica napus L.). Pak J Bot 44:2081–2086 Ujjan AA, Shahzad S (2014) Insecticidal potential of Beauveria bassiana strain PDRL1187 and Imidacloprid to mustard aphid (Lipaphis erysimi) under field conditions. Pak J Zool 46 (5):1277–1281 Ulusoy MR, Olmez BS (2006) Effect of certain Brassica plants on biology of the cabbage aphid Brevicoryne brassicae under laboratory conditions. Phytoparasitica 34(2):133–138 Upadhyay S (1996) Influence of sowing dates and fertilizer levels on the incidence of aphid (Lipaphis erysimi Kalt.) on Indian mustard. Indian J Entomol 57:294–297 Vaz L, Tavares MT, Lomonaco C (2004) Diversity and size of parasitoid hymenopterans of Brevicoryne brassicae L. and Aphis nerii Boyer de Fonscolombe (Hemiptera: Aphididae). Neotrop Entomol 33:225–230 Verma AK, Patyal SK, Bhalla OP, Sharma KC (1993) Bioecology of painted bug (Bagrada cruciferarum) (Heteroptera: Pentatomidae) on seed crop cauliflower (Brassica oleracea var. botrytis sub var. cauliflora). Indian J Agric Sci 63(10):676–678 Verma SC, Sharma PL, Bhardwaj RK (2017) Spatial distribution of Brevicoryne brassicae (L.) in cabbage Brevicoryne brassicae in mid-hills of Himachal Pradesh, India. J Appl Nat Sci 9 (3):1587–1591 Vitanza S (2012) Texas Agrilife Extension El Paso County. IPM Program Newsl 37(6). http://www. tpma.org/_newsletters/_el_paso/_2012/08312012%20_6.pdf Vora VJR, Bharodia RK, Kapadia MN (1985) Pests of oilseed crops and their control: Rape and Mustard. Pesticides 19(1):38–40 Wagh VH, Kharat AP (2015) Influence of abiotic factors on the incidence of cabbage aphid Brevicoryne brassicae (L.) on red cabbage. Ecol Environ Conserv Paper 21(3):1281–1283 Wagle BKS, Saravanan L, Sudha JP, Gupta P (2005) Seasonal incidence of cabbage aphid, Brevicoryne brassicae (L.) and its natural enemies in relation to weather parameters on cabbage. In: National conference on applied entomology, RCA, Udaipur, 26–8 September 2005, pp 22–23 Wagle BKS, Saravanan L, Jacob PS, Anand P (2006) Biology and predation potential of aphidophagous predators, Coccinella septempunctata and Coccinella transversalis on cabbage aphid Brevicoryne brassicae Linnaeus. Allahabad Farmers 9:70–74 Webb SE (2010) Insect management for crucifers (cole crops) (broccoli, cabbage, cauliflower, collards, kale, mustard, radishes, turnips) ENY-464. Entomology and Nematology Department, Florida Cooperative Extension Service, IFAS, University of Florida, Gainesville, FL White AJ, Wratten SD, Berry NA, Weigmann U (1995) Habitat manipulation to enhance biological control of Brassica pests by hover flies (Diptera: Syrphidae). J Econ Entomol 88:1171–1176 Yadav S and Kumar A (2018) Response of various Brassica genotypes against mustard aphid, Lipaphis erysimi (Kaltenbach) under natural conditions in Haryana. p 54. In: Souvenir and abstracts: 2018. International conference on bio and nano technologies for sustainable agriculture, food, health, energy and industry, held during 21–23 February, 2018, at GJUS&T Hisar, Haryana, p 292 Yadav S, Singh SP (2015) Bio-intensive integrated management strategy for mustard aphid Lipaphis erysimi Kalt. (Homoptera: Aphididae). J Appl Nat Sci 7(1):192–196 Yadav MK, Patel JI, Pareek A (2017) Biology and predatory potential of lady bird beetle, Coccinella septempunctata (Lin.) on mustard aphid, Lipaphis erysimi (Kalt.). The Bioscan 12 (3):1363–1366

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Yadav SK, Singh RB, Gautam MP, Singh G, Giri SK (2018) Bio-efficacy of insecticides against mustard aphid (Lipaphis erysimi Kalt.) on mustard (Brassica juncea L.). Int J Chem Stud 6 (2):2704–2708 Yadava JS, Singh NB (1999) Strategies to enhance yield potential of rapeseed-mustard in India. In: Wratten N, Salisburry PA (eds) Proceedings of the 10th international rapeseed congress. Canberra, Australia, pp 113–115 Yadu YK, Dubey VK (1999) Effect of nitrogen application and spray time on grain yield, net profit and mustard aphid (Lipaphis erysimi) population. Adv Plant Sci 12(1):45–48 Yue B, Liu TX (2000) Host selection, development, survival, and reproduction of turnip aphid (Homoptera: Aphididae) on green and red cabbage varieties. J Econ Entomol 93(4):1308–1314 Zandi-Sohani N, Soleiman-Nejhadian E, Mohiseni A (2004) Study on the resistance of five canola (Brassica napus L.) cultivars to cabbage aphid, Brevicoryne brassicae (L.). Sci J Agric 27:119–127 Zhang WQ, Hasan A (2003) Use of the parasitoid Diaeretiella rapae (McIntosh) to control the cabbage aphid Brevicoryne brassicae (L.). J Appl Entomol 127(9–10):522–526

7

Sucking Pests of Soybean Neeta Gaur and Rashmi Joshi

Contents 7.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2 Important Sucking Pests of Soybean . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.3 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

Soybean is an important crop around the world and is also known as miracle crop. There are many arthropods associated with this crop which are responsible for economic damage to the crop. These insect pests cause damages by acting as borers, defoliators, sap sucking, pod and seed feeding, etc. Among all the pests, sucking pests have recently got more attention due to its widely destructive nature by acting as a vector of some virus diseases in soybean, such as yellow mosaic virus dissemination by Bemisia tabaci. The ability of these insects to show exceptional reproductive nature and physiological modification has been making its management a challenging task for researchers. This chapter provides information on diagnostic features, life cycle, nature of damage, management practices and integrated pest management modules of major sucking pest of soybean.

7.1

Introduction

Soybean is an important oilseed crop around the world containing 40% oil and 20% protein. The earliest known record of its domestication was found in eleventh century BC in China (Hymowitz 1970). Today soybean is grown globally around N. Gaur (*) · R. Joshi Department of Entomology, College of Agriculture, GBPUA&T, Pantnagar, Uttarakhand, India # Springer Nature Singapore Pte Ltd. 2020 Omkar (ed.), Sucking Pests of Crops, https://doi.org/10.1007/978-981-15-6149-8_7

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Table 7.1 List of sucking pests of soybean S. No. 1 2 3 4 5

Common name Whitefly Aphids Stink bugs Leafhoppers or Jassids Lygaeid bugs

6

Mealy bugs

7 8

Capsid bugs Thrips

9

Red spider mite

Scientific name Bemisia tabaci Gennadius Aphis glycines Matsumura Nezara viridula (L.) Empoasca spp.

Family Aleyrodidae Aphididae Pentatomidae Cicadellidae

Order Hemiptera Hemiptera Hemiptera Hemiptera

Chauliops fallax Scott C. nigrescens Distant Pseudococcus vastator Maskell Ceratonia pallidifer Walker Caliothrips indicus Bagnall Ayyaria chaetophora Karny Tetranychus cinnabarinus (Boisduval)

Lygaeidae

Hemiptera

Coccidae

Hemiptera

Miridae Thripidae

Hemiptera Thysanoptera

Tetranychidae

Acarina

the world, majorly in USA, Brazil, Argentina, India and China in about 121 million hectare area. Due to poor crop cultivation practices many limiting factors affect the yield of crop, insect pest being one of the major causes. A wide range of insect pests belonging to orders Coleoptera, Diptera, Hemiptera, Lepidoptera, etc. are known to affect soybean production. Different sucking pests occur in different agroclimatic regions of the world. Among these Bemisia tabaci is one of the devastating insects affecting many crops in many parts of India and world. Other sucking pests, such as aphids, sting bugs, leafhoppers, thrips, coreid bugs, etc. have also been observed causing damage sporadically, the list has been given in Table 7.1. Keeping in mind the importance of sucking pests of soybean, detailed study of some important sap sucking insects is reported in this chapter.

7.2

Important Sucking Pests of Soybean

7.2.1

Whitefly (Bemisia tabaci Gennadius—Aleyrodidae, Hemiptera)

Two species of whitefly play major role in infestation in many crops of the world, viz. Greenhouse whitefly (Trialeurodes vaporariorum) and sweet potato whitefly (Bemisia tabaci), also referred to as tobacco whitefly, poinsettia whitefly or silverleaf whitefly. Later is prevalent in Soybean crop of India due to its ability to act as vector of many potential viral diseases and development of resistance against pesticides. “Whitefly of the world” is an important document which provides a broad overview about this pest (Mound and Halsey 1978). Bemisia tabaci was first reported in India in 1905 (Husain 1931). Identification of whitefly distribution is not an easy task as it becomes more difficult due to facilitation of its movement by humans (Bryne and Bellows 1991).

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7.2.1.1 Diagnostic Features According to the classification done by David N. Bryne (1991), whitefly can be identified with help of three important physical characteristics, i.e. vasiform orifice, waxes and fourth nymphal instar. Broadly they can be identified as having opisthognathous head, membranous wings and incomplete metamorphosis. The Vasiform Orifice is located on the dorsum of ninth abdominal segments of male while in females it may extend up to eighth abdominal segment. So it is not the anus in which whitefly empties its honeydew but the ligula associated with vasiform orifice. In whiteflies, the anus is present dorsally and when the honeydew is excreted by the insect, it fills the ligula of vasiform orifice. The ligula is then released by the insect and it gets rid of the honeydew excreted which was otherwise hindering the movement of insect (Gupta 1972). Whiteflies produce extracuticular waxes in all its life stages except the egg. Byrne and Hadley (1988) described the process of production of curly threads of waxes in B. tabaci. The ventrolateral abdominal surface consists of wax plates which in turn consist of rows of mitochondria, which again is associated with wax canal. When the wax is secreted from these canals it is ripped off in between by the action of hind tibiae on wax plate and thus resulting in curly wax particles. There are two pairs of wax plates in comparison to males which have four pairs of them. There are different biotypes of whiteflies; however, the concept of biotype came into light with the invasion of whitefly in southern USA which was different from the indigenous population referred to as A biotype. The invader population with different esterase profile was known as B biotype (De Barro et al. 2011). After determination of these two biotypes there have been biotypes ranging from C to T added to the list (Perring 2001). 7.2.1.2 Life Cycle While looking into the literature available on whitefly, it can be determined that temperature (26–32  C) and humidity (60–70%) play an important role in life cycle of this insect. It can complete a generation in 20–30 days under favourable weather conditions (Saini 1998). The life cycle of whitefly can be broadly categorized in three stages, i.e. egg, four nymphal instars and adult (Fig. 7.1). Egg The eggs of whitefly are pyriform or ovoid having pedicel which is a peg like extension of chorion measuring about 0.2 mm long (Byrne and Bellows 1991). Eggs are inserted by the insect in leaf tissues. Hatching period is about 4–7 days and the eggs turn brown before hatching. Nymph There are four nymphal instars in life cycle of whiteflies. The first instar is mobile and is known as crawler. These nymphs usually settle in few hours but take longer time in cold weather and insert their mouthparts into the phloem tissues after settling (Avidov 1956). First nymphal instar is about 0.27 mm long and 0.15 mm wide. It generally takes 2–4 days for completion of this stage. The second and third instars are immobile and they take 2–3 days each for completion of these stages (Capinera 2004). Fourth nymphal instar is called pupa and it measures about 0.7 mm

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1st instar nymph, 0.27mm long,0.15mm wide, 2-4 days

Egg 0.2mm, 4-7 days

Life cycle of whitefly

Adult

4th instar nymph, 0.7mm length, 0.4mm width, 4-7 days

2nd instar nymph,0.36mm length,0.22mm width,2-3 days

3rd instar nymph, 0.49mm length, 0.29mm width,2-3 days

Fig. 7.1 Life cycle of whitefly

in length and 0.4 mm in width and takes about 10–14 days. Nechols and Tauber (1977) stated that last nymphal instar has three morphologically distinct stages. The first stage is flattened and translucent. In second stage apolysis takes place and in the last stage apolysis is complete. The last stage has red eyes and yellow body pigment and is considered as pharate adult. Adult At the time of emergence, wings of adult are clear and in time gets covered with white wax. The adult starts feeding by piercing within few hours and moves from old leaves to younger leaves. The adult generally feeds on underside of leaves. It measures about 1.0–1.3 mm in length and holds its wings vertical parallel to the body when viewed dorsally (Fig. 7.2). Adults live for 10–20 days and may produce up to 300 eggs (Reddy and Rao 1989). Copulation depends on the prevailing weather conditions. During summer months, it takes place within 1–8 h of eclosion and in fall and spring it takes place in usually 3 days following eclosion (Avidov 1956). If we take a general view it was found that B. tabaci prefers hot and dry conditions with temperature ranging from 26 to 30  C and 60 to 70% RH (Traboulsi 1995).

7.2.1.3 Nature of Damage The damage by whiteflies may result in greater than 50% yield losses by extracting large amount of phloem sap (Lloyd 1922). This insect is also an important vector of many viral diseases, such as Begomovirus (Geminiviridae), Circulative transmission; Ipomovirus (Potyviridae), Semipersistent transmission; Crinivirus (Closteroviridae), Semipersistent transmission; Carlavirus (Betafelaxiviridae),

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Fig. 7.2 Whiteflies adults and nymphs on soybean leaf

Semipersistent transmission and Torrodovirus (Secoviridae) (Tiwari et al. 2013). The losses due to viral diseases are heavy in terms of yield reduction.

7.2.1.4 Management Bemisia tabaci species complex is an important pest and causes considerable damage to crops. International Union for the Conservation of Nature and Natural Resources (IUCN) has designated as “100 of the World’s Worst Invasive Alien Species”. Therefore there is a need to manage whiteflies by combination of various manipulations as: Cultural Management Cultural control is one of the management practices which is widely accepted for management of any pest. Luko Hiljea et al. (2001) has mentioned various methods used for the management of this pest. They classified various cultural practices used in different subheadings. Here we will use this classification to describe various methods of biological control. 1. Avoidance in time These practices separate crops from sources of insects in time. (a) Crop-free periods It reduces the virus inoculum in the area and helps in checking complete spread of this pest.

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(b) Crop residue disposal Spread of infested crop residue can be checked by proper disposal of crop waste. This is also helpful is achieving the goals of crop-free periods. (c) Planting dates Planting early or late may help in evading pest or vector infestation. (d) Weed removal Removal of weeds may help in removal of source of pest or vector incidence. Avoidance in space These practices will help in reducing the contact of pest with crop by exclusion, barriers or high planting density. The basic principle of high density planting is to grow more plant so that a greater number of plants escape insect infestation and thus may compensate for crop loss. Behavioural manipulation These practices manipulate olfactory and visual cues and thus change the behaviour of whitefly. Intercropping and mulches help in manipulating the planting of different species in close proximity and reducing insect ability to find host plant respectively. Host suitability These practices induce change in host quality. Fertilization and irrigation are one of the major practices in cultural control which help in affecting the biological process of insect. Application of fertilizer and irrigation affect the growth and development of plant and increasing resistance of host plants and thus affecting the development of insects. Removal These practices such as overhead irrigation helps in reducing the pest/vector directly, like rainfall affects the insect population in an area.

Biological Management A wide range parasitoids and predators are known to attack B. tabaci in field conditions if their growth is not hampered with pesticide application. Among parasitoids, Encarsia formosa and Eretmocerus sp. limit the development of whiteflies. Mycopathogens, like Verticillium lecanii, Paecilomyces sp. and Beauveria bassiana are important biocontrol agents under field conditions (Ioannou 1997). Among predators, Chrysoperla carnea, C. scelestes, Geocoris bicolor and Mallada boninensis are found to control the egg and nymph stages of whiteflies.

Chemical Management Pesticide application has many direct and indirect effects on the growth and development of whiteflies. Although there are many cases of resistance reported in this insect against insecticides. Application of combination pesticides, like organophosphates + pyrethroids in rotation mode would be useful to combat resistance.

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IPM Module Swathi and Gaur (2018) studied the effect of border crop and insecticide for management of Whitefly. It was found in the study that two rows of maize as border crop + seed treatment with thiamethoxam 30% FS @ 10 mL/kg + foliar spray of imidacloprid 17.8 SL @ 500 mL/ha at 30–35 DAS + spray of triazophos 40 EC @ 800 mL/ha at 45–50 DAS was effective in management. Border maize crop acts as barrier for the movement of whiteflies. Other practices also help in reduction of whitefly population.

7.2.2

Aphids (Aphis glycines Matsumura—Aphididae, Hemiptera)

Soybean aphid is native to eastern Asia including China, Indonesia, Japan, Korea, Malaysia, the Philippines, Taiwan and Thailand (Blackman and Eastop 2000). Matsumura (1917) first described soybean aphid and although this pest was native to eastern Asia but it became a major source of economic loss in the USA.

7.2.2.1 Diagnostic Features Aphids show peculiar mode of development by exhibiting polymorphism in different generations. Soybean aphids occur both as winged or migratory and wingless. The most important morphological feature present is cornicles or honey tubes at the posterior end; these tubes produce waxy secretions consisting of mainly triglycerides. These are liberated when aphid finds any danger and act as alarm pheromone, and help them in scattering. Wingless aphids are pear shaped yellow to lime green in colour (Fig. 7.4), while winged aphids have dark thorax and transparent wings that pass past the abdomen. 7.2.2.2 Life Cycle Life cycle of soybean aphid is highly complex as this is heteroecious (hostalternating) and holocyclic (generating sexual morphs) (Fig. 7.3). There are mainly six types of individuals present in life cycle of soybean aphids (Richards and Davies 1977) 1. Fundatrices—These are apterous, viviparous and parthenogenetic females with sense organs, legs and antennae not so well developed. The reduction in these sense organs may be due to development of high reproduction capacity. 2. Fungatrigeniae—These are progeny of fundatrices which are apterous, parthenogenetic and viviparous females. They live on primary hosts. 3. Migrantes—These are winged parthenogenetic viviparous females and develop from later generations of fundatrigeniae. They develop on primary hosts and later on fly to secondary hosts. 4. Alienicolae—These are parthenogenetic viviparous females which develop mostly on secondary hosts. 5. Sexuparae—These are parthenogenetic viviparous females. They mainly develop on secondary hosts and later on migrate to primary hosts.

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Fig. 7.3 Life cycle of A. glycines (Wang et al. 1962)

6. Sexuales—These are produced from sexuparae and appear only once in life cycle. It consists of sexually reproducing males and oviparous females.

In nonmigratory species migrantes and alienicolae are not present. In summer and spring soybean aphids reproduce asexually on soybean which is considered a secondary host. The optimum temperature range required for soybean aphids is 8.6–34.9  C. Under most favourable environment soybean aphids have been reported to produce 18–20 generations in China (Li et al. 2000). Also because of this characteristic pattern of reproduction there are found different biotypes of aphids.

7.2.2.3 Nature of Damage Soybean aphids are phloem feeders and with the help of piercing and sucking type of mouth parts they suck the sap from phloem tissues of plant. Injury symptoms such as reduced plant height, lower pod set and fewer seeds within pods have been observed in moderate infestation. Heavily infested plants show reduced photosynthetic rate with stunted growth, since the plants are covered with sooty mould growing on honey dew secreted by aphids. High population density of aphids reduces oil content in soybean pods (Beckendorf et al. 2008). Soybean aphids have been known to transmit many plant viruses and are a potential vector of plant viruses such as

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Fig. 7.4 Soybean aphids feeding on young leaves of soybean

soybean mosaic virus, alfalfa mosaic virus, cucumber mosaic virus, potato virus Y, etc. (Fig. 7.4).

7.2.2.4 Management Cultural Control Tilmon et al. (2011) referred in their study that monoculturing of soybean in large fields play an important role in development of soybean population. It was found that “80% control of A. glycines, a yield increase of 16%, and more than twice the natural enemy densities were achieved when soybean and maize were cosown or interplanted, integrated with (organic and inorganic) fertilizer, fertilizer seed-coating and pesticide applications” (Wang and Ba 1998; Zhishan et al. 2004). It was also found that soybean varieties with more nitrogen content will be more attacked by aphids as compared to one with lower nitrogen content and higher lignin content.

Biological Control A wide range of natural enemies including predators and parasitoids have been found to be effectively controlling the population of soybean aphid under natural conditions. A wide range natural enemies are known to control aphid population of soybean in Asia (Liu et al. 2004) and this is one of the major causes that the population of this pest is under control in Asia, being a major pest in North America.

Chemical Management Hodgson et al. (2012) recommended for the management of soybean aphids in USA and for chemical control it has been suggested to apply insecticides only at ETL i.e. “250 aphids per plant with 80% of the plants infested” to avoid resistance development in aphids. Also rotation of insecticides with different modes of action is an important point to be considered while using chemical management. Seed treatment is also an effective way and clothianidin, imidacloprid and thiamethoxam can be used, with thiamethoxam as most effective up to 49 days after planting.

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7.2.3

Stink Bugs (Nezara viridula (L.) Pentatomidae, Hemiptera)

Nezara viridula is one of the most important insect pests which occur in various tropical and subtropical areas (Pherson and Cuda 1974). It was found to be originated from Ethiopian region of eastern Africa (Hokkanen 1983). It is polyphagous pest and feeds on a wide range of dicotyledonous plants and few monocotyledonous plants and fruit or pod stage is the most preferred stage of insect.

7.2.3.1 Diagnostic Features As discussed by Bundy and McPherson (2000) one of the important diagnostic feature of N. viridula is the characteristic pattern of eggs. The number of eggs laid is high in the middle and decreases on its side and gives it a hexagon like shape. 7.2.3.2 Life Cycle Todd (1989) and Miner (1966) in their paper discussed the life history of N. viridula in detail. The insect is active at high temperature i.e. when temperature exceeds 24  C (Underhill 1934). There are mainly three stages in life stages of green sting bugs—egg, nymph and adults (Fig. 7.5). Egg Eggs are pale yellow or cream colour at oviposition and turn light pink before hatching. Incubation period is around 5 days in summers and 2–3 weeks in early

Egg

1st nymphal instar

On an average 32 eggs are laid by a female, with mean incubation period 9-19 days

Brownish red colour, with average period ranging from 5-10 days

Adult Copulate in about 22.3 days with preoviposition period 30-66 days. Egg laying females live longer

2nd nymphal instar Black colour, avg. period ranging from 6-14 days

Life cycle of N. viridula

3rd nymphal instar 5th nymphal instar

Usually green with some yellow and black colour, average period from 6-11 days

Green and black, avg period from 18-20 days

4th nymphal instar green with little black colour, avg period ranging from 6-11 days

Fig. 7.5 Life cycle of N. viridula, information derived from (Miner), 1966

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spring. On an average 32 eggs are laid per egg mass packed in a barrel shape on underside of leaves. The incubation period ranges from 9 to 19 days. Nymph There are five nymphal instars present in sting bugs. First nymphal instar do not feed and cluster around egg mass. After moulting of first instar, the second and third instar begin to feed, differing in size with similar pattern of feeding. It is feeding by fourth and fifth nymphal instars which is responsible for much of the economic damage of crops. Adults Within few hours of moulting adult starts feeding avidly and then gradually decline interest in feeding until death. The adults of N. viridula show a characteristic “edge effect”, in this case invading adults are first found on the edge of the field. The bugs overwinter as adults in leaf litters and become active when temperature exceeds 24  C. Females start copulating after approximately 22 days and lay egg mass after 3 weeks, with frequency of laying new egg masses after every 8–10 days (Kamminga et al. 2012) and leaves the plant after oviposition is complete.

7.2.3.3 Nature of Damage In histolytic studies of damaged portion of soybean by adults of green sting bugs it was found that in damaged area cell wall was ruptured or completely missing. The bug injects its feeding stylet in cell and draws food through stylets after liquefying cell content with histolytic agents. Feeding by sting bug causes “green stem syndrome” which results in green stems even after attaining maturity. Feeding by sting bugs reduces the quality of seeds as well as feeding punctures made by insect may also lead to pathogen infection. It also transmits the yeast spot disease (McPherson and McPherson 2000) and results in economic loss of growers. 7.2.3.4 Management Management of sting bugs can be given under different subheadings such as cultural, mechanical, chemical, biological, etc. Monitoring Sweep nets and beat sheets methods of monitoring are recommended to sample presence of sting bugs in field. ETL level of sting bugs is one sting bug per 25 sweeps or 1 sting bug per two-row meters (Herbert 2012). Pheromone traps have also been used for monitoring of sting bugs. Yellow base pyramid trap with pheromone lure is commonly used for pheromone traps. Cultural Management One of the best cultural practices for management of sting bugs is use of trap crops, like early maturing soybean, buckwheat, sunflower, pearl millet, sorghum. When the insect gets trapped in these crops then either by destroying trap crops or using insecticides over them will kill the insect present on the trap crops.

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Biological Control This is one of the most important management practice of N. viridula and several biological control has been successfully implemented in the past. A wide range of predators and parasitoids have been found infesting sting bugs, like mermithid nematodes, lacewing larva; spined soldier bugs (Podisus maculiventris) and birds. The most successful biological control agent for sting bugs is a scelionid egg parasitoid Trissolcus basalis and the tachinid flies Trichopoda pennipes and T. pilipes attacking nymphs and adults of sting bugs. It was found that higher mortality was observed in early stages of insect due to parasitism by biocontrol agents (Kiritani and Sasaba 1969). Chemical Control Many organophosphates, pyrethroids and neonicotinoids have been used for management of sting bugs. It was found by Willrich et al. (2003) that nymphs were more susceptible to chemical control as compared to adults. Azadirachtin was found to have antifeedant effect as well as acts as important growth regulator disrupting the moulting of nymphs of sting bugs. Integrated Pest Management Correa-Ferreira et al. (2000) implemented integrated soybean management in Brazil including monitoring, ETL based management practices, mass release of T. basilis, insecticidal application on field borders. This management practice was highly successful resulted in the decrease in application of insecticides for management of sting bugs.

7.2.4

Leafhoppers or Jassids (Empoasca spp., Cicadellidae, Hemiptera)

A wide range of leafhoppers fauna is present in soybean in the USA and the most abundant species found there is E. fabae (Tugwell et al. 1973). In India detail account of leafhoppers fauna was given by H. S. Pruthi (1940) who described many species of Empoasca present in India.

7.2.4.1 Diagnostic Features The leafhoppers show burn type of symptoms in the field known as “hopperburn” and known as “V”-shaped leaf chlorosis where the outer edges and tip turn yellow in earlier stages before giving burn like effect. Hypertrophy due to salivary secretion in phloem cells causes an interruption in photosynthesis causing plasmolysis of parenchyma cells leading to hopperburn (Medler 1941). The morphological characters like venation, head structure, ocelli position, body shape and genital structures have also been used to classify different types of leafhoppers (Dwight 1971).

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7.2.4.2 Life Cycle The life cycle of leafhoppers is divided into three stages, i.e. egg, nymph and adults. The number of generations in a year depends upon factors such as physiology of host plant and prevailing temperature. Egg Eggs are laid by females either singly exceeding up to one to two inside leaf epidermis by making a slit with the help of its ovipositor. In some species, eggs are covered by thread like papillae and terminal leaflets are favoured by females to oviposit as compared subterminal leaflets (Miller and Hibbs 1963; Severin 1950). Number of eggs laid generally do not exceed 200–300, and the oviposition period varies from few days to few months, i.e. ranging from average 2.7 eggs per day. The incubation period varies according to temperature ranging from 2.5 to 6 days in summers and one to several months in case of overwintering eggs or eggs laid in late autumn or early spring. Nymph Leafhoppers generally pass through five nymphal instars. The nymphal period varies in different species ranging from 7–8 days to 10 months in case of some species. Rate of development of nymphs also depends upon nature of food plant intake by the insect. Adult Leafhoppers are usually bisexual and parthenogenesis is also found as a means of reproduction among them. Males appear before females and also disappear before females in field. One mating is usually found sufficient to fertilize eggs during lifetime (DeLong 1938) and it was mated females that live longer than males of same age (Severin 1921). Leafhoppers mainly prefer adult stage of their life cycle to overwinter.

7.2.4.3 Nature of Damage Leafhoppers show characteristic “hopperburn” symptoms in the field. Glabrous and pubescent characteristics of soybean and other beans are mainly responsible for infestation with leafhoppers (Ogunlana and Pedigo 1974). Leafhoppers are also known to be important vectors of diseases caused by phytoplasma. 7.2.4.4 Management Pest management of leafhoppers requires weekly monitoring. Scouting is done with the help of sweep net and monitoring is done keeping in mind to take sample if there is presence of dew or if raining or when the wind speed is >10 m/h (Cherry et al. 1977). Cultural Control Early harvesting is advised to control population of leafhoppers in field; it is because it avoids further damage by leafhoppers (Undersander et al. 2004). Intercropping with grasses is also very effective as compared to monoculture against leafhoppers. Glandular hairy varieties are also an important factor to resist plants from leafhopper

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damage. “Crowning” or spring pruning is also an important management practice when the crop is grown in spring (Calderwood et al. 2015). Biological Control There are many generalist natural enemies which are found to be effective against leafhoppers like predatory true bugs and flies, lacewings and lady beetles. However, lady beetles and spiders are the most abundant, next being lacewings followed by syrphid flies and finally minute pirate bugs (Chasen et al. 2014). Chemical Control Different foliar insecticides are registered for chemical management of leafhoppers. Pyrethroids, organophosphates and some insecticide premixes, such as organophosphate + pyrethroid; chlorantraniliprole + pyrethroid; neonicotinoids + pyrethroid were found to be effective. However, before applying broad spectrum pesticides other beneficial arthropods should be taken care of.

7.3

Conclusions

Sucking pests are one of the important causes of economic losses of soybean crop. Proper information on biology, its important diagnostic features and seasonal occurrence play an important role in deciding the management practices to combat losses caused by the sucking pests. Although the development of resistance and biotypes is one of the major issues concerning the management of sucking pest lately, but the use of integrated pest management evolved as promising tool to alleviate losses caused by the pest.

References Avidov Z (1956) Bionomics of the tobacco whitefly (Bemisia tabaci Gennad.) in Israel. Ktavim 7:25–41 Beckendorf EA, Catangui MA, Riedell WE (2008) Soybean aphid feeding injury and soybean yield, yield components, and seed composition. Agron J 100:237–246 Blackman RL, Eastop VF (2000) Aphids on the world’s crops: an identification and information guide, 2nd edn. Wiley, New York Byrne DN, Hadley NF (1988) Particulate surface waxes of whiteflies: morphology, composition and waxing behaviour. Physiol Entomol 1(3):267–276 Byrne DN, Bellows TS Jr (1991) Whitefly biology. Annu Rev Entomol 36:431–457 Bundy CS, McPherson RM (2000) Morphological examination of stink bug (Heteroptera: Pentatomidae) eggs on cotton and soybeans, with a key to genera. Ann Entomol Soc Am 93 (3):616–624 Calderwood LB, Lewins SA, Darby HM (2015) Survey of northeastern hop arthropod pests and their natural enemies. J Int Pest Manage 6(1):18. https://doi.org/10.1093/jipm/pmv017 Capinera JL (2004) Encyclopedia of entomology, vol I. Kluwer Academic Publishers, Dordrecht Chasen EM, Dietrich C, Backus EA, Cullen EM (2014) Potato leafhopper (Hemiptera: Cicadellidae) ecology and integrated pest management focused on Alfalfa. J Int Pest Manage 5(1):1–8. https://doi.org/10.1603/IPM13014

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Cherry RH, Wood KA, Ruesink W (1977) Emergence trap and sweep net sampling for adults of the potato leafhopper from alfalfa. J Econ Entomol 70:279–282 Corrêa-Ferreira BS, Domit LA, Morales L, Guimarães RC (2000) Integrated soybean pest management in micro river basins in Brazil. J Int Pest Manage 5:75–80 De Barro PJ, Liu SS, Boykin LM, Dinsdale AB (2011) Bemisia tabaci: a statement of species status. Annu Rev Entomol 56:1–19 DeLong DM (1938) Biological studies of the leafhopper, Empoasca fabae as a bean pest. U.S. Department of Agriculture, Beltsville, MD, pp 1–60 Dwight MD (1971) The bionomics of leafhoppers. Ann Rev Entomol 32:179–210 Gupta PC (1972) External morphology of Bemisia gossypiperda (M. & L.) a vector of plant virus diseases (Homoptera: Aleyrodidae). Zool Beitr 8:1–20 Herbert DA (2012) Insects: soybeans. In: Herbert DA, Hagood S (eds) Field crops 2012. Virginia Cooperative Extension Publication No. 456–016, pp 4-61–4-76. http://pubs.ext.vt.edu/456/ 456–016/Section_4_Insects-1.pdf Hiljea L, Costab HS, Stansly PA (2001) Cultural practices for managing Bemisia tabaci and associated viral diseases. Crop Prot 20:801–812 Hodgson EW, McCornack BP, Tilmon K, Knodel JJ (2012) Management recommendations for soybean aphid (Hemiptera: Aphididae) in the United States. J Int Pest Manage 3(1):2012. https://doi.org/10.1603/IPM11019 Hokkanen H (1983) Interspecific homeostasis. pest problems and the principle of classical biological pest control. PhD thesis, Cornell University, Ithaca, p 157 Husain MA (1931) A preliminary note on the white-fly of cottons in the Punjab. Agric J India 25:508–526 Hymowitz T (1970) On the domestication of the soybean. Econ Bot 24:408–421 Ioannou N (1997) Current status of whitefly - virus complex in near east. In: Ioannou N (ed) Management of the whitefly-virus complex. Proceedings of the FAO workshop on management of the whitefly virus complex in vegetable and cotton production in the Near East, 2–6 October, 1995, Larnaca, Cyprus. FAO, Rome, pp 137–157 Kamminga KL, Koppel AL, Herbert DA Jr, Kuhar TP (2012) Biology and management of the green stink bug. J Int Pest Manage 3(3):8. https://doi.org/10.1603/IPM12006 Kiritani K, Sasaba T (1969) The difference in bio- and ecological characteristics between neighbouring populations in the southern green stink bug, Nezara viridula L. lpn. J Ecol 9 (5):77–84 Li CS, Luo RW, Yang CL, Shang YF, Zhao JH, Xin XQ (2000) Biology and control of Aphis glycines. Soybean Sci 19:337–340 Liu J, Wu K, Hopper KR, Zhao K (2004) Population dynamics of Aphis glycines (Homoptera: Aphididae) and its natural enemies in soybean in northern China. Ann Entomol Soc Am 97 (2):235–239 Lloyd LL (1922) The control of the greenhouse white fly (Asterochiton vaporariorum) with notes on its biology. Ann Appl Biol 9:1–34 Matsumura S (1917) A list of the Aphididae of Japan, with description of new species and genera. J Coll Agric Sapporo Japan 7:387–388 Mc Pherson JE, Cuda JP (1974) The first record in Illinois of Nezara viridula (Hemiptera: Pentatomidae). Trans Ill State Acad Sci 67(4):461–462 McPherson JE, McPherson RM (2000) Stink bugs of economic importance in North America & Mexico. CRC, Boca RatonL Medler JT (1941) The nature of injury to alfalfa, caused by Empoasca fabae (Harris) Ann. Entomol Soc Am 34:439–450 Miller RL, Hibbs ET (1963) Distribution of eggs of the potato leafhopper, Empoasca fabae, on solanum plants. Ann Entomol Soc Am 56:737–740 Miner FD (1966) Biology and control of stink bugs on soybeans. Arkansas Agric Exp Station Bull 708:1–40 Mound LA, Halsey SH (1978) Whitefly of the World. Wiley, New York, p 340

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Nechols JR, Tauber M (1977) Age-specific interaction between the greenhouse whitefly and Encarcia formosa: influence of the parasite on host development. Environ Entomol 6:207–210 Ogunlana MO, Pedigo LP (1974) Pest status of the potato leafhopper on soybeans in central Iowa. J Eeon Entomol 67:201–202 Perring TM (2001) The Bemisia tabaci species complex. Crop Prot 20:725–737 Pruthi HS (1940) Description of some new species of Empoasca Walsh (Europterygidae, Jassoidea) from North India. Indian J Entomol 2(1):1–9 Reddy AS, Rao NV (1989) Cotton whitefly (Bemisia tabaci Genn.). Indian J Plant Prot 17:171–179 Richards OW, Davies RG (eds) (1977) Imms’ general textbook of entomology. Imperial College University of London, London Saini HK (1998) Effect of synthetic pyrethroids on biology of whitefly Bemisia tabaci (Gennadius) on Gossypium hirsutum (Linn.). M.Sc. Thesis, Punjab Agricultural University, Ludhiana, India Severin HHP (1921) Summary of life history of beet leafhopper (Eutetti.'!' tenella Baker). 1. Bean. Enlomol 14:443–436 Severin HHP (1950) Texananus incurllatus III life history on virus-infected and on healthy plants. Hilgardia 19:546–548 Swathi M, Gaur N (2018) Effect of border crops and insecticides on management of whitefly, Bemisia tabaci (Gennadius) transmitted yellow mosaic virus in soybean. Int J Curr Microbiol App Sci 6(5):613–617 Tilmon KJ, Hodgson EW, O’Neal ME, Ragsdale DW (2011) Biology of the soybean aphid, aphis glycines (Hemiptera Aphididae) in the United States. J Int Pest Manage 2(2):2011. https://doi. org/10.1603/IPM10016 Tiwari SP, Nema S, Khare MN (2013) Whitefly—a strong transmitter of plant viruses. Sci J Plant Pathol 02(02):102–120 Todd JW (1989) Ecology and behavior of Nezara viridula. Annu Rev Entomol 34(1):273–292 Traboulsi (1995) Bemisia tabaci: a report on the pest status with particular reference to the near east. FAO Plant Protect Bull 42:33–35 Tugwell P, Rouse EP, Thompson RG (1973) Insects in soybeans and a weed host (Desmodium sp.). Arkansas Agric Exp Sta Rep Ser 214:218 Underhill GW (1934) The green stinkbug. Virginia Agric Exp Station Bull 294:1–26 Undersander D, Becker R, Cosgrove D, Cullen E, Doll J, Grau C, Kelling K, Rice ME, Schmitt M, Sheaffer C et al (2004) Alfalfa management guide. In: NCR547 North Central Regional Extension Publication. Society of Agronomy, Madison Wang YZ, Ba F (1998) Study on optimum control of the soybean aphid. Acta Phyt Sinica 25:152–155 Wang CL, Xiang NY, Zhang GS, Zhu HF (1962) Studies on the soybean aphid, Aphis glycines Matsumura. Acta Entomol Sinica 11:31Ð44 Willrich MM, Leonard BR, Cook DR (2003) Laboratory and field evaluations of insecticide toxicity to stink bugs (Heteroptera: Pentatomidae). J Cotton Sci 7:156–163 Zhishan WU, Schenk-Hamlin D, Zhan W, Ragsdale DW, Heimpel GE (2004) The soybean aphid in China: a historical review. Ann Entomol Soc Am 97(2):209–218

8

Sucking Pests of Cotton P. S. Shera, Vijay Kumar, and Vikas Jindal

Contents 8.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2 Sucking Pests . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.3 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

The adoption of genetically modified cotton expressing genes from soil bacterium, Bacillus thuringiensis Berliner has changed the entire pest scenario in the cotton ecosystem. The pest status of bollworms and leaf feeding insects has declined, but the incidence of sap feeders including whitefly, Bemisia tabaci (Gennadius), leafhopper, Amrasca biguttula biguttula (Ishida), mealybug, Phenacoccus solenopsis Tinsley, thrips, Thrips tabaci (Lindeman), aphid, Aphis gossypii (Glover) and green mirid bug, Creontiades biseratense (Distant) has increased owing to reduction in insecticide use during reproductive phase of the crop. The outbreaks of mealybug in entire cotton belt and mirid bug in Central and South India were witnessed during the last decade. Similarly, recent occurrence of whitefly outbreak in North India (Punjab, Haryana and Rajasthan) in 2015 caused havoc to cotton crop, inflicting huge economic losses to farmers. The population of thrips early in the season, leafhoppers during vegetative–flowering phase and aphids at fag end of the crop season are also gaining importance. A brief description on sucking pests, their damage symptoms and management strategies have been discussed in the chapter.

P. S. Shera (*) · V. Kumar · V. Jindal Department of Entomology, Punjab Agricultural University, Ludhiana, Punjab, India e-mail: [email protected]; [email protected]; [email protected] # Springer Nature Singapore Pte Ltd. 2020 Omkar (ed.), Sucking Pests of Crops, https://doi.org/10.1007/978-981-15-6149-8_8

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Introduction

Cotton (Gossypium spp.) fibre has exercised a profound influence on humans from the time immemorial. With a history going back to ancient times, the fibre has maintained its immaculate purity and importance to this day. In India, cotton crop has the pride of place among cash crops from the earliest time. Cotton fabrics were found in the Mohenjo-daro relics from the ancient Indus Valley Civilization which flourished in India about 5000 years ago (Sethi et al. 1960). Even in modern times, cotton is an important crop of commerce in India and plays a key role in agricultural, industrial, social and monetary affairs of the country. About 60 million people of the country are involved directly or indirectly in cotton production, processing, textiles and related activities (Gopalakrishnan et al. 2007). All the four cultivated species of cotton, viz. Gossypium arboreum L., Gossypium hirsutum L., Gossypium herbaceaum L. and Gossypium barbadense L. along with intra-specific and inter-specific hybrids are grown in India under diverse agroclimatic conditions, varying from 8 to 32
N latitude and 70 to 80
E longitude. India ranks first in global scenario contributing about 37 and 24% of the world cotton area and production, respectively. However, the average productivity of cotton in India (504 kg ha1) is still far less than other major cotton growing countries of the world, viz. Australia (2202 kg ha1), China (1761 kg ha1), Brazil (1555 kg ha1), USA (1008 kg ha1), Egypt (718 kg ha1), Uzbekistan (671 kg ha1), Pakistan (638 kg ha1) and Turkmenistan (569 kg ha1) (AICCIP 2018). The insect pests are major bottleneck in achieving high productivity. Cotton crop harboured 1326 insect species from sowing to maturity in different cotton growing areas of the world (Hargreaves 1948) and 162 species have been reported on the cotton crop in India alone, of which 24 species have attained pest status (Sundramurthy and Chitra 1992; Arora et al. 2006). The insect pest complex of cotton crop is broadly divided into four categories, viz. sucking pests, foliage feeders, bollworms and lint stainers. The adoption of transgenic Bt cotton expressing genes from soil inhabiting spore forming bacterium, Bacillus thuringiensis Berliner in countries, such as India, has changed the entire pest scenario in the cotton ecosystem. The pest status of bollworms and leaf feeding insects has declined, but sap feeders are emerging as serious pests (Kumar et al. 2015). Changing Scenario of Sucking Pests The insect pest complex of cotton crop has undergone a tremendous change (Table 8.1) owing to many reasons, viz. increase in area under American cotton (upland cotton) replacing the Asiatic cotton, adoption of Bt cotton, change in ecological scenario, cultivation of large number of cotton cultivars with different varietal reaction to insect pests, unrecommended insecticides, excessive use of inputs, like fertilizers and insecticides, at over/under dosages, mixtures of synthetic pyrethroids with other conventional insecticides and faulty spray technology, etc. The indigenous cotton varieties of G. arboreum were predominant in early 1950s and were resistant to leafhopper, Amrasca biguttula biguttula (Ishida). Large scale

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Table 8.1 Changing status of cotton insect pests in India during last five decades Year Up to 1970

Major pest Leafhopper, pink bollworm

1971–1980

Leafhopper, pink bollworm, spotted bollworm Leafhopper, pink bollworm, spotted bollworm Leafhopper, whitefly American bollworm, spotted bollworm, pink bollworm Leafhopper, whitefly, mealybug, tobacco caterpillar, spotted bollworm, American bollworm Whitefly, leafhopper, thrips

1981–1990

1991–2000

2001–2010

2011–2019

Minor pest Whitefly, aphid, thrips Whitefly, aphid, thrips Whitefly, aphid, thrips Aphid, thrips

New pest Spotted bollworm

Pest outbreak American bollworm in some pockets (1978)

American bollworm

American bollworm (1983)

Tobacco caterpillar

American bollworm (1990, 1995)

Leaf miner, CLCuV

Whitefly (1995) and American bollworm (1997, 1998)

Aphid, thrips

Mealybug

American bollworm (2001), Tobacco caterpillar (2005), and Mealybug (2007)

Aphid, mealybug

–

Whitefly (2015)

Modified after Kumar et al. (2015)

adoption of high yielding G. hirsutum varieties along with use of heavy inputs, i.e. fertilizers and insecticides, has altered the pest scenario. Leafhopper was the only serious sucking pests on cotton upto mid-1970s (Dhawan 2001). Owing to excessive use of synthetic pyrethroids for the control of bollworms, whitefly attained status of key pest and is continuously inflicting huge losses to cotton crop. Moreover, whitefly as a vector of cotton leaf curl virus has attributed a lot in taking major toll of cotton production in North Zone of the country. From 2012 onwards, the whitefly has been on upsurge with population levels above ETL (6 adults per leaf in the upper canopy of plant before 10:00 am) throughout the cropping season (Singh et al. 2016). During 2015, severe incidence of whitefly (60–90 adults per leaf) was recorded in Abohar, Faridkot, Fazilka, Muktsar and Mansa districts of Punjab and yield losses varied from 40 to 100%. Besides economically important pests, like leafhopper and whitefly, other sucking pests like thrips hitherto occurring during May–June and aphids that occur at the end of the crop season are also gaining importance. During the last decade, a new sucking pest, mealybug Phenacoccus solenopsis Tinsley appeared in a few pockets of Bathinda, Ferozepur and Muktsar districts. The serious damage of this pest was observed in cotton fields near roads, field bunds and water channels having abundance of weeds flora. The maximum damage was observed in Sangat block of Bathinda district, but in 2007, it spread to other cotton growing areas of Punjab and emerged in a serious proportion causing huge loss to the cotton crop in Punjab (Dhawan et al. 2007). Similarly, mirid bug Creontiades biseratense (Distant), a

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minor pest appeared in epidemic form in Dharwad (Karnataka) and Coimbatore (Tamil Nadu) and has also gained importance as major pest of cotton (Udikeri et al. 2009; Surulivelu and Dhara Jothi 2007). Thrips tabaci (Lindeman) remained as minor pest of cotton in Northern India during the last decade but severe outbreak was recorded in a few cotton fields of Punjab and Haryana during August 2017.

8.2

Sucking Pests

8.2.1

Whitefly, Bemisia tabaci (Gennadius) (Hemiptera: Aleyrodidae)

8.2.1.1 Distribution Although B. tabaci was described from Greece (Gennadius 1889), it probably originated in India or Pakistan (Mound 1983). Whitefly has been widely distributed in various countries such as Portugal, Spain, France, Italy, Mexico, Hawaii, Turkey, Jordan, Venezuela, Brazil, Sudan, Cuba, Egypt, Puerto Rico, Spain, Tunisia, USA, Mexico, China, Japan, India, Netherland and several other countries/regions (Thompson 2011). 8.2.1.2 Host Plants Whitefly feeds on more than 900 diverse host plants, including species of economic importance belonging to the 63 families (Perring 2001). 8.2.1.3 Damaging Stage(s) Nymphs and adults. 8.2.1.4 Species Status Using molecular tools, it was demonstrated that B. tabaci is having a complex of atleast 43 cryptic biological species (Tay et al. 2017). The two most invasive and destructive species of whitefly now found in almost 60 countries are B. tabaci B (Middle East-Minor Asia 1) and B. tabaci Q (Mediterranean genetic group) (Dinsdale et al. 2010; De Barro et al. 2011; Wan and Yang 2016). In India, nine cryptic species has been reported. A new genetic group MEAM-K has been found in Karnataka state. AsiaII_1 cryptic species is predominant in northern region of India. 8.2.1.5 Diagnostic Features Whitefly lays eggs singly on the abaxial surface of leaves which are firmly attached to the leaf tissue by pedicel. The freshly laid eggs are yellowish white in colour, transparent and spindle shaped and then it becomes dark brown before hatching. First instar nymph is the only mobile nymphal stage, which becomes sedentary after finding a suitable feeding location on the lower surface of leaf. The fourth nymphal stage is termed as puparium, which is yellowish white in colour and oval in shape, has well-developed reddish brown coloured compound eyes which are distinctly visible. The adults are tiny, moth-like insects that feed on plant sap (Fig. 8.1).

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Fig. 8.1 Whitefly damage

8.2.1.6 Life Cycle Bemisia tabaci moults four times during its life cycle and has six different life stages, namely egg, three nymphal instars, pupa and adult. Female whitefly on an average lays 50–120 eggs. Hatching occurs after 4–5 days (Khodke and Dagaonkar 1992; Vennila et al. 2007a). The first three nymphal stages last for 9–14 days and pupa lasts for about 4–5 days after which the adults emerge through an inverted ‘T’ shaped slit in the dorsal surface of the pupal case. A female lives for 5–6 days and male for 4–5 days (Khodke and Dagaonkar 1992). B. tabaci remains active throughout the year and has 12–15 overlapping generations in a year (Aneja 2000). 8.2.1.7 Nature of Damage Whiteflies have been reported to cause damage to cotton by sucking sap by nymphs and adults from the leaves and producing honey dew on the leaves. Chlorotic spots appear at feeding site on leaf surface, followed by wilting and leaf shedding. Damage at early stages of growth affects the development of reproductive structures and consequently the yield may be greatly reduced. The leaves turn black due to sooty mould development (Fig. 8.1). In case of severe infestation, whitefly causes reduction in growth, boll formation and results in excessive shedding of fruiting forms and premature opening of bolls. 8.2.1.8 Management Options Host Plant Resistance With the help of appropriate screening techniques, several resistant/tolerant sources and varieties/hybrids to whitefly have been identified (Table 8.2). The resistant varieties or hybrids against whitefly (LK 861, Kanchana and Supriya) have been released for cultivation in India (Singh and Kairon 2001; Dhawan et al. 2008; Dhawan 2019). The resistance/tolerance to sucking pests has been attributed to both morphological and biochemical characters in cotton (Table 8.3). For example, hairy cotton varieties are prone to whitefly and glabrous cotton leaves significantly hinder whitefly oviposition. The varieties having thin leaf lamina, okra leaf type and

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Table 8.2 Resistant/tolerant sources (G. hirsutum and G. arboreum) identified against whitefly in cotton Insect pests Whitefly

Resistant/tolerant sources identified G 27 LD 327, LD 323 LD 327 TH 18, TH 19, IL 88, IL 90, IL 99, R1, R2, R23, R 27 USA-22 LD 694, Supriya, PA 183 DHH 419 and GSHV 97/1092

Reference(s) Sidhu and Dhawan (1980) Dhawan et al. (1991) Simwat and Gill (1992) Kular and Butter (1997) Butter and Vir (1989) Jindal et al. (2008), Jindal and Dhaliwal (2009) Jindal et al. (2007)

Table 8.3 Plant traits associated with resistance/susceptibility against sucking pests of cotton Plant traits Morphological characters Okra Rapid fruiting Hirsute Pilose Glabrous leaf Hairy leaves Red plant body Biochemical characters High gossypol Heliocides (H1 to H 4) High tannins High phenols Silicated leaf

Leafhopper

Whitefly

– S R R S R –

R R S S R S R

R – – – R

– R R R –

R relative resistance, S relative susceptible Source: Dhawan (2004)

high hair density should be included in insect resistant breeding program to develop resistant varieties against whitefly as it will help to reduce the menace of both whitefly and leafhopper in cotton agro-ecosystem (Dhawan 2019).

Cultural and Mechanical Control With the awareness of environmental problems, exploitation of different cultural and mechanical practices, viz. planting time, plant geometry, tillage, crop sanitation, fertilizer as well as water management, intercropping, crop rotation, manipulation of carryover sources, barrier/trap crops, etc. have been advocated as a vital approach to curb pest population, as these are simple and easy to follow. The extensive studies on these cultural and mechanical control practices have resulted in recommendation of best suited ones as a component of IPM module for cotton whitefly (Table 8.4).

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Table 8.4 Impact of different cultural and mechanical control strategies on whitefly Cultural practices Grow recommended cotton varieties/hybrids Non-judicious use of irrigation water Proper plant spacing and maintenance of plant population Uprooting and destruction of weeds during in and off season like Sida sp., Abutilon sp., Parthenium sp., Achyranthes sp., Trianthema sp., Xanthium sp., Tribulus sp., Digera sp., etc. Judicious use of nitrogenous fertilizers Avoid cultivation of adjoining crops in and around cotton crop—Okra, moong, clusterbean Installation of yellow sticky traps Optimum sowing time (April to mid-may in North India)

Impact Undescript material highly susceptible to sucking pests and poor yielder Conducive for population build-up Reduces incidence; facilitate spraying operations Reduces carry-over/inoculum and population build

Over dose conducive for population buildup Most preferred; increase in population build-up and incidence on cotton crop Monitoring and management More damage in late sown crop; escape mechanism

Modified after Kumar et al. (2015)

Biological Control Several bioagents from diverse groups have been recorded against cotton insect pests across India (Table 8.5). Predatory fauna comprising 16 species of insects in 5 families and 4 orders; 7 species of arachnids in 4 families constituting a total of 23 species have been recorded to be associated with sucking insect pests in cotton (Natarajan 1990; Kedar et al. 2014; Sangha et al. 2018; Dhawan 2019). The parasitoids, namely Encarsia lutea (Masi) (Hymenoptera: Aphelinidae), E. sophia (Girault & Dodd), E. shafeei Hayat and Eretmocerus sp. have been reported to parasitize B. tabaci under field conditions (Sharma et al. 2003; Kedar et al. 2014; Sangha et al. 2018; Dhawan 2019). Of the 65 pesticide products registered in India against sucking insect pests on cotton, only one entomopathogenic fungus, Lecanicillium (¼Verticillium) lecanii is recommended for the management of whitefly (CIBRC 2019). Against whitefly, myco-insecticides V. lecanii, Fusarium sp. and B. bassiana have been reported to reduce its population by 38.3, 37.1 and 36.0%, respectively (Sharma and Summarwar 2017). Other fungal pathogens like Aspergillus sp., Paecilomyces sp. and Fusarium sp. on Bemisia tabaci in cotton have also been recorded from cotton whitefly (Rao et al. 1989). Due to inconsistent performance, commercial products have not been popular with the cotton farmers. The technology is, however, invaluable and needs to be fine-tuned to the point at which mass production costs are low and efficacy is rapid and reliable before being popularized. Botanicals Neem products can be effectively used for pest management alone or in combination/alternation with recommended insecticides as best alternative to reduce the pesticide usage. The spray of Neemark, Indne and RD 9 was found significantly at

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Table 8.5 Arthropod predators and parasitoids of sucking pests on cotton crop Natural enemies Predators Coccinella septempunctata Linnaeus Coccinella transversalis Fabricius Cheilomenes sexmaculata (Fabricius) Brumoides suturalis (Fabricius) Hyperaspis maindroni Sicard Serangium parcesetosum Sicard Scymnus nubilis Mulsant Coelophora bissellata Mulsant Chrysoperla zastrowi sillemi (Esben-Peterson) Zanchius breviceps (Wagner) Deraeocoris indianus Carvalho Zelus sp. Geocoris sp. Antilochus sp. Ischiodon spp. Syrphus spp. Neoscona sp. Argiope sp. Oxyopes sp. Thomisus sp. Runcinia sp. Hyllus sp. Chrysilla sp.

Family

Order

Coccinellidae Coccinellidae Coccinellidae Coccinellidae Coccinellidae Coccinellidae Coccinellidae Coccinellidae Chrysopidae Miridae Miridae Reduviidae Geocoridae Pyrrhocoridae Syrphidae Syrphidae Araneidae Araneidae Oxyopidae Thomisidae Thomisidae Salticidae Salticidae

Coleoptera Coleoptera Coleoptera Coleoptera Coleoptera Coleoptera Coleoptera Coleoptera Neuroptera Hemiptera Hemiptera Hemiptera Hemiptera Hemiptera Diptera Diptera Araneae Araneae Araneae Araneae Araneae Araneae Araneae

Source: Natarajan (1990), Sharma et al. (2003), Kedar et al. (2014), Sangha et al. (2018), and Dhawan (2019)

par with dimethoate for the control of B. tabaci (Dhawan et al. 1992; Dhawan and Simwat 1992). Neem oil (0.5%) and NSKE (5%) were also found to be effective against whitefly (Natarajan and Sundaramurthy 1990; Uthamasamy and Gajendran 1992; Vidya et al. 1993). Mann et al. (2001) reported two azadirachtin enriched neem based insecticides, NeemAzal (1%) and RakshakGold (1%) to be effective for whitefly control during the flowering phase. Neem oil and neem seed extract @ 1.0 L/acre during the early part of the season have been advocated for sucking pests on Bt cotton and validated at farmers’ fields (Anonymous 2019). Insect Growth Regulators The chemicals that affect the growth and development of insects are called insect growth regulators (IGRs), a name in keeping with the familiar plant growth regulators (PGRs) that influence plant growth and physiology. The growth regulators that are of interest for pest control include insect hormones (moulting and juvenile hormones) and their analogues and chitin synthesis inhibitors (CSIs). IGRs, namely buprofezin and pyriproxyfen have been evaluated against sucking

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insect pests of cotton in India. Buprofezin acts by inhibiting cuticle deposition. It also suppresses egg laying in female adults with inhibition of prostaglandin synthesis and has effects on levels of hormones associated with moulting in nymphs. It has been used effectively on cotton crop against B. tabaci (Beevi and Balasubramanian 1995; Gogi et al. 2006). Pyriproxyfen is a potent juvenile hormone analog affecting the embryogenesis, metamorphosis and adult formation of various sucking insect pests of agricultural importance. It has been found to be effective for the management of whitefly and leafhopper on Bt cotton (Kumar et al. 2014, 2019). Biotechnological Approaches RNA interference (RNAi) is a new technique being used for analyzing the functions of individual genes by the use of loss-of-function analyses in many animals, including insects. Before the discovery of RNAi, it was almost impossible to analyze the functions of genes in insects for which genetic analyses are difficult. RNA interference (RNAi) is a cellular process by which a specific mRNA is targeted for degradation by introduced double-stranded RNA (small interfering RNA) that contains a strand complementary to a fragment of the target mRNA, resulting in sequence specific inhibition of gene expression. The technique was explored to study the gene function in whitefly, jassid, etc. in cotton. In whitefly, various gene like Actin ortholog, V-ATPase A subunit, BtCG5885, BtGATAd, BtSnap, hsp70, hsp90, GST, TLR7, Acetylcholinesterase (AChE), Ecdysone receptor (EcR), vitellogenin, Aquaporin, juvenile hormone esterase have been studied through RNAi and their phenotypic effects were recorded (Grover et al. 2018, 2019). Nicotiana tabacum transgenic plants expressing dsRNA corresponding to Acetylcholinesterase (AChE) and Ecdysone receptor (EcR) resulted in 50–90% mortality. Similarly, Lactuca sativa plants expressing dsRNA of v-ATPase caused 84–98% mortality and 95 fold decrease in fecundity of whitefly. The comprehensive review on RNAi in whitefly is done by Grover et al. (2018). Chemical Control The economic threshold level for whitefly is 20 nymphs/leaf or 6–8 adults per leaf or appearance of honeydew on 50% of plants. The insecticides belonging to different groups like organophosphates, carbamates, nionicotinoids, phenylpyrazoles, pyrethroids, insect growth regulator, inhibitors of mitochondrial ATP synthase, pyridinecarboxamide, ketoenols were recommended for management of whitefly (Table 8.8). Integrated Pest Management Keeping in view the ill effects of pesticide use in cotton, validation and promotion of IPM technology through farmer’s participatory approach became a necessity to sustain productivity of cotton with least disturbance to the ecosystem. In India whitefly appeared in epidemic form during the year 2015 in Punjab. As a result, the crop productivity fell substantially from 574 kg/ha of lint in 2014–2015 to only 197 kg/ha in 2015–2016. For effective management of whitefly, IPM strategy was implemented in Punjab. It includes regular surveillance of whitefly on alternate hosts

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(brinjal and cucurbits) from February and on cotton from April onwards, clean cultivation campaign during March–May to prevent the carryover of whitefly, availability and quality inputs, ensuring canal water availability for timely sowing, recommended dose of fertilizers, use of non-chemical approaches like traps, neem based botanicals, chemical insecticides based on economic threshold level, proper spray methodology, spray of potassium nitrate to improve crop health. The incidence of whitefly was monitored at weekly interval in cotton belt of Punjab. During 2015, the higher incidence of whitefly (22–42 adults/leaf) was recorded, whereas in 2016, whitefly appeared early in the season during 24th standard meteorological week. However, during rest of the period, it remained low due to adoption of IPM strategy. During 2017, higher population was recorded during second fortnight of July to first week of September. In addition, trainings for extension personnel, scouts and pesticides dealers and cotton growers were also organized. Literature pertaining to whitefly management was distributed to cotton growers. Live TV programmes were broadcasted on whitefly management. Interstate consultative and monitoring Committee constituted by Government under the Chairmanship of Vice Chancellor, PAU regularly reviewed the incidence of whitefly through meetings as well as during field visits. Wherever necessary mid-course modifications were made based on feedbacks from the fields. Farmers were advised for judicious use of pesticides. The impact of adoption of IPM strategy resulted in reducing the pesticide use in the State by INR 737.8 million (saving of INR2589/ha for the cotton growers) in 2016 over 2015. The IPM campaign in 2017and 2018 resulted in still further reduction in pesticide use worth INR 817.3 million (saving INR 2808/ha) and INR 884.9 million (saving INR 3060/ha) over the consumption in the year 2015. With the joint efforts of scientists, State Department of Agriculture and farmers, whitefly was managed successfully during 2016, 2017 and 2018 resulting in all time the highest productivity of 756, 750 and 778 kg/ha lint, respectively (Kumar et al. 2019).

8.2.2

Leafhopper, Amrasca biguttula biguttula (Ishida) (Hemiptera: Cicadellidae)

8.2.2.1 Distribution It is widely distributed in India and is the most destructive pest of American cotton in north-western regions. 8.2.2.2 Host Plants Cotton, okra, potato, brinjal and some wild plants, etc. 8.2.2.3 Damaging Stage(s) Nymphs and adults. 8.2.2.4 Diagnostic Features Female lays curved, elongated and yellowish white coloured eggs, which are embedded in the midribs of large veins on the undersurface of leaves. Nymphs

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remain confined to lower surface of leaves and are pale greenish in colour. They are wedge shaped and move diagonally in relation to their body. Adults have pale green body. The hind portion of the forewings has characteristic two black spots on the vertex.

8.2.2.5 Life Cycle The females lay eggs in the spongy parenchymatous tissue of the leaf veins between the vascular bundles and epidermis. A single female can lay a maximum of 29 eggs but on an average, it lays about 15 eggs. The incubation period of the eggs ranged from 4 to 11 days, depending upon the season. There were five nymphal instars, which completed their development in 7 days during summer and 21 days during winter. Mating took place 2–16 days after emergence and oviposition started 2–7 days after mating. The period of oviposition has been reported to vary from 2 to 14 days. The mated adults lived only for 36 days in summer and 49 days in winter, whereas unmated ones survived for 3 months. There were 11 overlapping generations of leafhopper in a year (Arora et al. 2006). 8.2.2.6 Nature of Damage Leafhopper remains active during July to September with the peak activity during mid-July to end August. It is deleterious during vegetative stage of the cotton plant and has the ability to build-up to serious proportions. Both nymphs and adults suck the sap from the leaves. Among different development stages, nymphs have been reported to be more injurious than the adults, while older nymphs are more damaging than the younger ones. In the early stages of attack, the leaves of cotton turn brownish, crinkled and finally assume an inverted cup shaped appearance, gradually turn brick red in colour and shed prematurely (Fig. 8.2). Besides causing physical injury, they inject toxic saliva into the plant tissue which is more injurious. The young seedlings succumb and older plants remain stunted. The cotyledons of cotton show curling and withering, whereas older leaves exhibit typical symptoms of hopper burn.

Fig. 8.2 Leafhopper damage

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8.2.2.7 Management Options Host Plant Resistance The resistant/tolerant lines of cotton have been identified against leafhopper (Table 8.6). The earlier work of Hussain and Lal (1940) has led to evolving hairy cotton varieties resistant to leafhopper and by 1943 resistant varieties such as Punjab 4F, LSS and 289F/43 covered extensive areas where leafhopper had posed a serious threat. The resistance/tolerance to sucking pests has been attributed to both morphological and biochemical characters in cotton (Table 8.3). For example, hairy cotton varieties are extremely resistant to cotton leafhopper but prone to whitefly and vice versa. Likewise, glabrous cotton leaves significantly hinder whitefly oviposition. In such circumstances, it is desirable to give greater emphasis on exploiting other common characters that may confer resistance against majority of key insects especially under North Indian conditions, where both leafhopper and whitefly are serious problems (Dhawan 2019). Cultural and Mechanical Control 1. Proper plant spacing and judicious use of fertilizers should be used to avoid conducive environment for population build-up of leafhopper. 2. The crops like okra being most susceptible to leafhopper should not be grown near cotton fields. Biological Control Lecanicillium lecanii was found to be most effective in reducing the population of leafhoppers with an average mortality of 69.17% (Manivannan et al. 2018). Table 8.6 Resistant/tolerant sources (G. hirsutum and G. arboreum) identified against leafhopper in cotton Insect pests Leafhopper

Resistant/tolerant sources identified J-129, J-150, 231-R 5197, line-91, line 129, 5201/6, 5281 G-27, F 414 G 27, LD 133 and LD 230 LD 327 SRT-1, Jhurar Pantese 4, Fateh, B 1007, Jhurar, SRT 1 AKH 9312, RS 992, Bar 7-8-2, BIOC 7, 666/102, F 520, F 882, WC stonvilla 213 Okra leaf mutant, Reba B-50, B 1007, SRT-1, Khandwa 2, Pantose 4 CCH 510-4, KR 13

Reference(s) Singh (1970) Singh et al. (1972) Sidhu and Dhawan (1980) Chakravarthy and Sidhu (1986) Simwat and Gill (1992) Dhillon et al. (1998) Dhillon et al. (1999) Anonymous (2000) Anonymous (2002) Jindal et al. (2007)

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Botanicals The spray of Neemark, Indne and RD 9 was found significantly at par with dimethoate for the control of A. biguttula (Dhawan et al. 1992; Dhawan and Simwat 1992). Neem oil (0.5%) and NSKE (5%) were also found to be effective against whitefly, leafhopper and aphid (Raju et al. 1992; Uthamasamy and Gajendran 1992; Vidya et al. 1993). Insect Growth Regulators Pyriproxyfen, a potent juvenile hormone analog affecting the embryogenesis, metamorphosis and adult formation, has been found to be effective for the management of leafhopper on Bt cotton (Kumar et al. 2014, 2019). Chemical Control The economic threshold level (ETL) for leafhopper has been established, viz. 1–2 nymphs/leaf or appearance of 2nd grade injury—curling & yellowing at margins in some of the fully formed leaves in the upper canopy on 50% of plants. The insecticides having label claim against leafhoppers on cotton are given in Table 8.8.

8.2.3

Aphid, Aphis gossypii (Glover) (Hemiptera: Aphidiidae)

8.2.3.1 Distribution Worldwide including tropical, sub-tropical, and temperate areas (Inaizum 1980). 8.2.3.2 Host Plants Cotton aphid, a cosmopolitan species is extremely polyphagous, polymorphic and breed incessant parthenogenetically. It has broad host range damaging various crops including Cucurbitaceae, Malvaceae, Solanaceae and Rutaceae as well as some ornamental plants such as chrysanthemum (Ebert and Cartwright 1997). 8.2.3.3 Damaging Stage(s) Nymphs and adults. 8.2.3.4 Diagnostic Features These are small pear shaped, soft-bodied and yellow or dark green in colour (Fig. 8.3). They may be apterous (wingless) or alate (winged). The dark cornicles or siphunculi are the main diagnostic structures in aphids, which are present on the posterior part of the abdomen (Minks and Harrewijn 1986). There is considerable overlap between instars, but there is no overlapping between nymphs and adults. First-instar and second-instar nymphs have four and five antennal segments, respectively. Third-instar nymphs have no setae on the margin of the genital plate, whereas they are present in the fourth instar nymphs. Developing wings are prominent in fourth-instar nymphs as compared to small wing pads in third-instar nymphs (Ebert and Cartwright 1997).

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Fig. 8.3 Aphid damage

8.2.3.5 Life Cycle Aphids have both apterous and winged forms. The nymphal period varied from 4 to 24 days and 5 to 22 days in apterous and winged form, respectively. In apterous form, the adult longevity ranged from 1 to 41 days, while it varied from 1 to 24 days in winged form. The total life cycle of female is completed in 28–44 days (Kataria and Kumar 2015). 8.2.3.6 Nature of Damage Both young and adult cotton aphids suck plant sap. Infested plants become weak and the tender shoots, leaves fade gradually and may become blighted due to appearance of sooty mould on middle canopy leaves in case of severe attack. Dry conditions favor rapid increase in pest population and younger plants are more susceptible than the older ones (Arora et al. 2006). 8.2.3.7 Management Options Cultural and Mechanical Control High nitrogen and water availability levels have shown to enhance aphid growth and development in cotton (Slosser et al. 2001). Avoid excessive use of nitrogenous fertilizers to reduce population build-up. Biological Control Aphids play an important role in regulating the food chains within agro ecosystems by serving as food for several parasitoid and predator species (Lawo et al. 2009). The role of insect species in the coccinellidae, chrysopidae, syrphidae and hemerobiidae as predators and aphelinidae as parasitoids in biological control of aphids has been widely recognized (Ebert and Cartwright 1997). However, biological control of cotton aphid is only possible, when the aphid population is low. But if exceeds the threshold level, insecticides are used to control aphid population to reduce economic losses (Ebert and Cartwright 1997).

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Few studies have been conducted to demonstrate the action of various entomopathogens, namely Beauveria bassiana, L. lecanii and Pseudomonas fluorescens against cotton aphids. Vu et al. (2007) reported that among different entomopathogenic fungi, L. lecanii performed best in controlling A. gossypii. Verticillium (¼ Lecanicillium) lecanii (Verticel) 1.15 WP @ 7.5 kg/ha did not differ significantly from chemical insecticide (Acetamiprid 20 SP) in reducing the population of aphids on cotton (Patil et al. 2012). Under field conditions, three sprays of L. lecanii @ 4 g/L or B. bassiana @ 4 g/L at fortnightly interval starting from initiation of aphid population have been found effective in Bt cotton (Raghunandan et al. 2018). Manjula et al. (2018) demonstrated the potential of P. fluorescens @ 1% as soil treatment and foliar application, which caused significant mortality of cotton aphid in Bt and non-Bt cotton.

Botanicals The aqueous extract of neem seeds has been found efficient against cotton aphid causing nymph mortality and reducing their survival period and fecundity (Santos et al. 2004). Spray of neem seed kernel extract (5%) or neem oil (1%) in combination with detergent/soap @ 1 g/L has been found to suppress the population of aphids (Vennila et al. 2007d). Similarly, neem seed kernel extract (5%) has been found to be effective against cotton aphid followed by Pongamia glabra seed kernel extract (5%), neem oil (3%) and Pongamia glabra oil (3%) (Vinodhini and Malaikozhundan 2011).

Insect Growth Regulators The juvenoid insect growth regulators, kinoprene and fenoxycarb and the insecticide synergist piperonyl butoxide (PBO) slows population growth, delays adult emergence and significantly alters population structures of A. gossypii (Satoh et al. 1995). Due to higher efficacy and lower toxicity on associated natural enemies, binary mixtures of insecticide flonicamid with pyriproxyfen or buprofezin could be promising candidates for the management of A. gossypii (Eldesouky 2019).

Chemical Control Need based sprays of chemical insecticides based on ETH level, i.e. 10 aphid nymphs/plant or appearance of honey dew on 50% plants have been suggested. Different insecticides belonging to various groups, viz. carbamates (carbaryl, carbosulfan), organophosphates (monocrotophos, chlorpyriphos, quinalphos, profenophos, methyl parathion, oxydemeton-methyl), neo-nicotinoids (imidacloprid, thiamethoxam, thiacloprid, acetamiprid, clothianidin, dinotefuran), phenylpyrazoles (fipronil), pyrethroids (fenvalarate), pyridine carboxamide (flonicamid) and miscellaneous (diafenthiuron) have been recommended against aphid on cotton (Table 8.8).

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Thrips, Thrips tabaci (Lindeman) (Thysanoptera: Thripidae)

8.2.4.1 Distribution It is a major pest of agricultural crops globally. Severe damage to different crops has been reported in Africa, Asia, Europe, North and South America and Australasia (Boateng et al. 2014). 8.2.4.2 Host Plants They have a wide host range, reportedly feeding on over 300 cultivated crops as well as uncultivated plants in at least 25 families. The principal crop hosts include onion, cotton, beans, broccoli, alfalfa, asparagus, cabbage, carnation, carrot, cauliflower, Chinese broccoli, cucumber, blackberry, kale, garlic, head cabbage, tomato, turnip, pepper, leek, melon, orchids, papaya, peas, pineapple, potato, pumpkin, lettuce, rose, squash, strawberry, tobacco, celery and mustard (Lewis 1997; Shelton et al. 2008). 8.2.4.3 Damaging Stage(s) Nymphs and adults. 8.2.4.4 Diagnostic Features Females lay white or yellow kidney-shaped eggs singly by inserting them into leaf tissue. As eggs mature, they develop an orange tinge and eventually reddish eye spots become evident. The first and second instars are active feeding stages. The first instar is semi-transparent and dull white, later changing to yellowish white. The second instar is larger and yellow having red eyes. The abdomen is divided into eight distinct segments and has a large posterior segment that is conical in shape. The prepupa and pupa are relatively inactive and nonfeeding stages. Adult thrips are pale yellow to grey in colour with darker transverse bands across the abdomen. The fore wings and hind wings are fringed and pale in colour (Patel et al. 2013). 8.2.4.5 Life Cycle The pest breeds on cotton during May–September. The adult female lays 50–60 kidney-shaped eggs, singly into green plant tissue, at the rate of 4–6 eggs per day (Arora et al. 2006). The average larval period varied from 5 to 7 days and total life cycle lasted for 38–62 days (Patel et al. 2013). 8.2.4.6 Nature of Damage Both the nymphs and adults first lacerate the surface tissues of the foliage and then feed on the exuding sap. The affected leaves curl and give silvery brown appearance (Fig. 8.4). Prolonged dry spell during May–June often results in high build-up of the pest on the young plants. The attacked plants give bushy appearance with lopsided growth during the seedling stage.

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Fig. 8.4 Thrips damage

8.2.4.7 Management Options Cultural and Mechanical Control 1. Maintain optimum plant stand for the management of thrips during very early crop growth stage. 2. Cotton field should be free from weeds right from the beginning of crop growth to reduce the population build-up of thrips. Botanicals Spray neem seed kernel extract @ 5% or crude neem oil spray @ 1% by adding detergent/soap powder @ 1 g/L of spray fluid for getting uniform spray suspension to suppress thrips population during pre-squaring crop stage (Vennila et al. 2007c). Chemical Control Spray of chemical insecticides, imidacloprid 200 SL and thiamethoxam 25 WG @ 1–1.5 g/L of water has been suggested when there is high degree of symptoms, indicative of high thrips population (Vennila et al. 2007c). Different ready-mix insecticides, acephate 50% + imidacloprid 1.8% SP and thiamethoxam 12.6% + lambda-cyhalothrin 9.5% ZC were found to be highly effective, whereas fipronil 5% SC and buprofezin 15% + acephate 35% WP were moderately effective against thrips on Bt cotton (Padaliya et al. 2018). Recently, a new chemical insecticide, Spinetoram in the spinosyn class of insecticides has been registered against thrips on cotton by Central Insecticide Board and Registration Committee (CIRBC 2019).

8.2.5

Mealybug, Phenacoccus solenopsis Tinsley (Hemiptera: Pseudococcidae)

8.2.5.1 Distribution It was first documented as a pest of cotton in Texas, USA (Fuchs et al. 1991) and now widely distributed in various ecological zones of the globe including Central

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America, South America, China, Africa and Australia (García Morales et al. 2016). During the last decade, it emerged as a serious threat to cotton in India and Pakistan, causing severe economic losses and is now widespread in almost all cotton growing states of India.

8.2.5.2 Host Plants Phenacoccus solenopsis is a polyphagous pest and has a wide morphological diversity, biological adaptations and ecological adjustments that allow it to feed on diverse host plants, including plant species of economic importance belonging to the families Malvaceae, Solanaceae, Compositae, Amaranthaceae, Asteraceae, Convolvulaceae, Euphorbiaceae, Verbenaceae, Zygophyllaceae, Cucurbitaceae, Lamiaceae and Fabaceae (Nagrare et al. 2011). 8.2.5.3 Damaging Stage(s) Nymphs and adults. 8.2.5.4 Diagnostic Features The eggs are whitish yellow, semi-transparent, oval to oblong in shape. They are laid within the white cottony ovisac that remains underneath the body of female initially and later on female moved leaving the ovisac on the leaf surface. The moulting occurs thrice in females and four times in males. The second instar nymphs are whitish yellow in colour, oblong in shape and secrete waxy material on their body. Wax secretion intensifies in third instar and whole body appears milky white. The adult males are slender and have well-developed mesothoracic wings, whereas adult females are wingless, oblong in shape and light to dark yellow in colour ventrally. Dorsally the whole body is well segmented, covered with waxy deposition except at the posterior abdominal region where blackish stripes on either side of mid-dorsal line are visible (Fig. 8.5). It possess a pair of brownish, short, filiform eight segmented antennae and three pairs of brownish red legs (Dhawan and Saini 2009; Pawar et al. 2017). 8.2.5.5 Life Cycle The life cycle of P. solenopsis females ranges from 27 to 38 days including adult longevity in warmer conditions, whereas the life cycle of male varies from 16 to 23 days. The duration of different development stages of female P. solenopsis is 1–2 days (eggs), 4–6 days (1st instar), 4–6 days (2nd instar), 5–7 days (3rd instar) and 13–17 (adult female longevity). The fecundity of adult female varies from 270 to 340 young ones. The females give birth to young ones by ovoviviparity (egg are retained within the body until ready to hatch and crawlers directly emerge) (Dhawan and Saini 2009). 8.2.5.6 Nature of Damage The nymphs and adults of P. solenopsis suck cell sap from leaves, twigs, stem, inflorescence and fruiting bodies (Fig. 8.5). Cotton plants infested during vegetative phase exhibit symptoms of distorted and bushy shoots, crinkled and/or twisted and

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Fig. 8.5 Mealybug damage

bunchy leaves, and stunted plants that dry completely in severe conditions. Lateseason infestations during reproductive crop stages result in reduced plant vigour and early crop senescence. While feeding, it injects a toxic substance into the plant parts resulting in chlorosis, stunting, deformation and death of plants. The development of sooty mould on honeydew secreted by them invariably affects photosynthesis of plants (Nagrare et al. 2009). The damage to cotton crop by this pest has resulted in huge economic losses to the cotton growing farmers. The estimated yield losses in cotton due to mealybug were around 12% in 2006 and almost 40% in 2007 in Punjab province of Pakistan (Kakakhel 2007). In Punjab, India, Dhawan et al. (2007) reported that highly infested plants gave an average seed cotton yield of 0.135 kg as compared to healthy plants (0.242 kg), recording 44.21% reduction over healthy plants. The avoidable loss was estimated to be Rs 37,413/- ha1 in monetary terms.

8.2.5.7 Management Options Cultural and Mechanical Control The utilization of different cultural and mechanical practices, viz. recommended varieties, crop sanitation, fertilizer management, manipulation of carryover sources, barrier/trap crops, etc. has been advocated as a vital approach to curb mealybug population. The extensive studies on cultural and mechanical control practices have resulted in recommendation of best suited ones as a component of IPM module for cotton mealybug (Table 8.7). Biological Control The role of coccinellids and chrysopids as predators (Suroshe et al. 2013; Shera et al. 2017) and encyrtids as parasitoids (Ram et al. 2011; Dhawan et al. 2011; Suroshe et al. 2013; Shera et al. 2017) in biological control of mealybugs has been widely

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Table 8.7 Impact of different cultural and mechanical control strategies on mealybug in cotton Cultural/mechanical practices Grow recommended cotton varieties/hybrids Uprooting and destruction of weeds during in and off season like Sida sp., Abutilon sp., Parthenium sp., Achyranthes sp., Trianthema sp., Xanthium sp., Tribulus sp., Digera sp., etc. Avoid cultivation of adjoining crops in and around cotton crop—Okra, moong, clusterbean Do not throw uprooted infested plants/weeds in cotton fields/water channels Growing barrier crops in and around cotton crop— Bajra, maize, sorghum After last pick, beating of infested sticks against ground Stacking the infested sticks separately and using them on priority for fuel purpose up to end of February Do not allow the movement of farm animals in infested fields Restrict the movement of workers in infested fields Prevent the movement of sticks from the infested areas to the new areas

Impact Undescript material highly susceptible to sucking pests and poor yielder Reduces carry-over/inoculum and population build

Most preferred; increase in population build-up and incidence on cotton crop Check further spread Least preferred so act as barrier; and restricts its migration to cotton crop Collection and destroying dislodged mealybugs by burying reduces carry over Reduces carry over

Check further spread Check further spread Check further spread from infested areas to new areas

Modified after Kumar et al. (2015)

recognized. Among these, an endoparasitoid Aenasius arizonensis (Girault) (¼Aenasius bambawalei Hayat) has been reported to be the most prominent and key mortality factor of P. solenopsis under field conditions across India. This parasitoid has neither been imported purposefully nor released artificially and is a classic example of fortuitous biological control. Field parasitization to the extent of 95% has been reported in P. solenopsis by this parasitoid (Ram and Saini 2009). Fallahzadeh et al. (2014) from Iran reported that this species fall closer to A. arizonensis (Girault) and reported A. bambawalei to be a junior synonym of A. arizonensis. It is an important parasitoid due to its adaptability to environmental conditions, faster multiplication than the host, female biased sex ratio, high host searching capacity, ease of culturing in the laboratory, high dispersal capacity and synchronized lifecycle with the host (Nagrare et al. 2011) (Table 8.8). Foliar spray of L. lecanii or B. bassiana (2  108 cfu/mL) @ 5 g or mL/L of water found to be effective during high humid months in reducing the population of mealybugs (Tanwar et al. 2007). Under field conditions, entomopathogens, B. bassiana, L. lecanii and M. anisopliae reduced the mealybug population by 37.7, 39.8 and 41.4%, respectively, and were least toxic to generalist predators (Rishi et al. 2012).

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Table 8.8 Major insecticides having label claim against sucking pests on cotton Insecticide group/mode of action (IRACa) Organophosphate (acetyl cholinesterase (AChE) inhibitors)

Carbamates (acetyl cholinesterase (AChE) inhibitors)

Neonicotinoid (nicotinic acetylcholine receptor (nAChR) competitive modulators)

Insecticide Acephate 75 SP Chlorpyriphos 20 EC Ethion 50 EC Methyl parathion 50 EC

Dose (g or mL/ha) 390 1250 1500–2000 500–1000

Monocroptophos 36 SL

500

Monocroptophos 15 SG

1333

Oxydemeton methyl 25 EC Phorate 10 G

1200 10,000

Profenophos 50 EC

1500–2000

Triazophos 40 EC Carbaryl 50 WP

1500–2000

Carbaryl 85 WP Carbosulfan 25 DS

1411 60 g/kg seed

Acetamiprid 20 SP

50

Clothianidin 50 WDG Dinotefuran 20 SG

30–40 40–50 125–150

Imidacloprid 17.8 SL

100–125

Imidacloprid 70 WG

30–35

Imidacloprid 48 FS

500–900

2000

Target pests Leafhopper Whitefly, aphid Whitefly Aphid, leafhopper, thrip Aphid, leafhopper, thrips, whitefly Aphid, leafhopper, thrips, whitefly Aphid, leafhopper Aphid, leafhopper, thrips, whitefly Aphid, leafhopper, thrips, whitefly Whitefly Aphid, leafhopper, thrips Thrips, whitefly Aphid, leafhopper, thrips Aphid, leafhopper, whitefly Leafhopper Whitefly Aphid, leafhopper, thrips, whitefly Aphid, leafhopper, thrips, whitefly Aphid, leafhopper, Thrips Aphid, leafhopper, thrips, whitefly

500–1000 (continued)

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Table 8.8 (continued) Insecticide group/mode of action (IRACa)

Dose (g or mL/ha)

Target pests Aphid, leafhopper, thrips, whitefly Aphid, leafhopper, thrips Aphid, leafhopper Aphid, leafhopper, thrips, whitefly Leafhopper, thrip Aphid, leafhopper, thrips, whitefly Aphid, leafhopper, thrips Whitefly Aphid, leafhopper, thrips, whitefly Aphid, leafhopper, thrips, whitefly Whitefly Aphid, leafhopper, red cotton bug Aphid, leafhopper, thrips, whitefly Whitefly

Imidacloprid 30.5 SC

60–75

Thiamethoxam 30 FS Thiamethoxam 70 WS

10

Thiamethoxam 75 SG Thiamethoxam 25 SG

125

Thiacloprid 21.7 SC

100–125 500–600

Fipronil 5 SC

1500–2000

Fipronil 18.87 SC

375

Pyrethroids (sodium channel modulators

Bifenthrin 10 EC Fluvalinate 25 EC

800 200–400

Insect growth regulator (chitin synthesis inhibitor)

Buprofezin 25 SC

1000

Insect growth regulator (juvenile hormone mimic) Inhibitors of mitochondrial ATP synthase

Pyriproxyfen 10 EC Diafenthuron 50 WP

500

Pyridinecarboxamide (Chordotonal organ modulators)

Flonicamid 50 WG

150

Spinosyns (nicotinic acetylcholine receptor (nAChR) allosteric modulators) Ketoenols (inhibitors of acetyl CoA carboxylase—Lipid synthesis, growth regulation)

Spinetoram 11.7 SC Spiromesifen 22.9 SC

420

Aphid, leafhopper, thrips, whitefly Aphid, leafhopper, thrips, whitefly Thrips

600

Whitefly, mite

Phenylpyrazoles (GABA-gated chloride channel blockers)

Source: CIBRC (2019) IRAC (2019)

a

Insecticide Imidacloprid 48 WS

430

100 200

600

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Botanicals Among different botanicals, Ocimum sanctum @ 0.6% has been found to be effective against mealybug as compared to Azadirachta indica, Calotropis gigantea, Nicotiana tabacum and Allium sativum which showed different levels of insecticidal activities (Prishanthini and Vinobaba 2014). Bharathi and Muthukrishnan (2017) observed that fish oil rosin soap @ 2.5%, neem oil @ 3% and NSKE @ 5% are found effective in reducing the population of mealybug in both preliminary and confirmatory field trials. Insect Growth Regulators Buprofezin acts by inhibiting cuticle deposition. It also suppresses egg laying in female adults with inhibition of prostaglandin synthesis and has effects on levels of hormones associated with moulting in nymphs. It has been used effectively on cotton crop against mealybug (Dhawan and Kumar 2012). Chemical Control Insecticides belonging to three major groups, viz. organophosphates (profenophos 50 EC, acephate 75 SP, chlorpyriphos 20 EC and quinalphos 25 EC) and carbamates (thiodicarb 75 SP and carbaryl 50 WP) have been recommended for the management of cotton mealybug (Anonymous 2019).

8.2.6

Mirid Bug, Creontiades biseratense (Distant) (Hemiptera: Miridae)

8.2.6.1 Distribution Green mirid bug, considered as minor pest, gained importance during the last decade in Southern states of India including Karnataka (Patil et al. 2006), Tamil Nadu, Andhra Pradesh and Maharashtra (Surulivelu and Dhara Jothi 2007). Besides this, Hyalopeplus lineifer (Walker) and Campylomma livida (Reuter) also occurred on Bt cotton in Karnataka, however their population was low in number as compared to C. biseratense (Udikeri et al. 2009). 8.2.6.2 Host Plants Polyphagous with wide range of host plants including cotton, amaranthus, bajra, castor, dhaincha, lucerne, Indian mallow, mungbean, pigweed, pigeonpea, peanut, safflower, sorghum, soybean, sunflower (Sankar et al. 2010). 8.2.6.3 Damaging Stage(s) Nymphs and adults. 8.2.6.4 Diagnostic Features Eggs of green mirid bug are transparent nacreous white, which turns to pink colour before hatching. They are laid singly or in groups on squares, tender bolls and leaf tissues and are cigar shaped (Ravi et al. 2015). The first instar nymph has prominent

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filiform antennae; the last segment of the antenna is pinkish in colour and the rest of the body is transparent yellow. The second- and third-instar nymphs are light yellow or green in colour. Wing pad development starts in second instar, which is yellowish brown in colour and turns blackish and extended over abdomen in third instar. In fourth instar, the nymphal wing pads become prominent and appear light yellow green or brown in colour. The fifth-instar nymphs are green or pale yellow or brownish in colour. The wing pads become conspicuous with elongation over abdomen and colour changed to transparent creamish with brown tinge. The freshly emerged adults have light green transparent wings. Body colour of adult turns to brown after 10–14 h of moulting (Udikeri et al. 2010).

8.2.6.5 Life Cycle The eggs of mirid bug hatch in 5–7 days. There are five nymphal instars and nymphal period is 12–18 days (Ravi et al. 2015). The total life cycle lasts for 34–38 days for males and 34–45 days for females (Udikeri et al. 2010). 8.2.6.6 Nature of Damage The mirids suck sap by piercing their sharp stylet into the plant tissues, squares and small tender bolls leading to premature shedding of squares and tender bolls as well as deformation of mature bolls (Ravi 2007). The avoidable losses due to mirid bugs have been reported to be 11.69% (Udikeri et al. 2009). 8.2.6.7 Management Options Cultural and Mechanical Control 1. Avoid close spacing to reduce population build-up. 2. Uproot and destroy the host plants. 3. Grow strips of lucerne as trap crop within field or in borders to reduce its population on cotton crop (Sankar et al. 2010). Biological Control Conservation of natural predators like damsel flies, big eyed bugs, assassin bugs, ants and spiders helps to control the mirid bug population (Ravi 2007). The biopesticides, Metarhizium anisopliae @ 1.0 g/L (17.6–29.3% reduction), Pochonia (Verticillium) lecanii @ 1.0 g/L (15.9–32.5% reduction) and nimbecidine @ 3.0 mL/ L (23.2–30.0% reduction) have been found to be less effective in suppressing the mirid population as compared to chemical insecticides. However, the cost benefit ratio was comparatively more in M. anisopliae (16.95:1.00) and P. lecanii (15.33:1.00) as compared to tested chemical insecticides except acephate 19.80:1.00 (Sugandi 2009). Chemical Control To ensure the need based application of insecticides, economic threshold values (ETH) have been established for mirids on cotton. Due to plant ability to compensate the damage, threshold values may vary between warm and cool season areas. In

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warm season, ETH level is 1.0 mirids/m by visual method, while 3.0 mirids/m by beat sheet method; however in cool season, ETH level is 0.5 and 1.5 mirids/m by visual and beat sheet methods, respectively (Sankar et al. 2010). The insecticides, acephate 75 WP @ 1.0 g/L, profenophos 50 EC @ 2.0 mL/L, indoxacarb 14.5 SC @ 0.5 mL/L, buprofezin 25 SC @ 0.5 mL/L and fipronil 5 SC @ 1.0 mL/L have been found effective against mirid bug on cotton (Sugandi 2009). The superiority of acephate 75 SP fipronil 5 SC profenophos 50 EC has also been reported against mirid bug on cotton crop (Udikeri et al. 2009; Bheemanna et al. 2010). Integrated Pest Management In Tamil Nadu, IPM technology including lucerne as a trap crop, four foliar sprays of acephate 75 WP and azadirachtin 10,000 @ 2 mL/L alternatively between 15 days interval have been validated through farmer’s participatory approach on Bt cotton. The population of mirid bugs was significantly lower in IPM, besides increase in seed cotton yield (63.4%) as compared to farmers’ practice. The population of coccinellids, green lacewing and spiders was also higher in IPM as compared to farmer’s practice (Birah et al. 2018)

8.2.7

Red Cotton Bug, Dysdercus koenigii (Fabricius) (Hemiptera: Pyrrhocoridae)

8.2.7.1 Distribution India, Pakistan and Southeastern Asia. 8.2.7.2 Host Plants Plant families, Malvaceae and Bombacaceae (Kamble 1971; Kohno and Ngan 2004). 8.2.7.3 Damaging Stage(s) Nymphs and adults. 8.2.7.4 Diagnostic Features Eggs are spherical in shape and laid in moist soil or in crevices in the ground. The colour of eggs is creamy white, turning to yellowish orange prior to hatching. There are five nymphal instars. The first- and second-instar nymphs are orange/red in colour. Initially, third-instar nymphs are orange red, which turns to reddish colour and there is emergence of wing pads on thorax. The fourth- and fifth-instar nymphs are crimson red in colour and cylindrical in shape. Prominent wing pads appeared in fifth instar, the posterior tips of wing pad are darker in colour than proximal part. Proboscis remains crimson red in colour while antennae and legs are black in colour. Adults are also crimson red in colour with a pair of black spots on the forewings (Fig. 8.6). The membranous hind wings are concealed under the forewings when the insect is at rest (Jaleel et al. 2013).

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Fig. 8.6 Red cotton bug

8.2.7.5 Life Cycle The female lays 100–130 eggs in moist soil, in crevices in loose irregular masses of 70–80 eggs each (Arora et al. 2006). There are five nymphal instars which complete their development in 23–25 days. Female lives longer (20.85  6.12 days) than the male (16.18  6.06 days). Each female mates three times in her life time; fecundity and hatching are more after first time mating and it decreases after subsequent second and third time mating (Jaleel et al. 2013). 8.2.7.6 Nature of Damage Red cotton bug is a minor pest and occurs at the boll bursting stage. Heavily attacked bolls open badly and the lint is of poor quality. The bugs stain the lint with their excreta or body juices as they are crushed in the ginning factories. The staining of lint by the growth of certain bacteria inside the bolls is also believed to be initiated by these bugs (Vennila et al. 2007b). 8.2.7.7 Management Options Cultural and Mechanical Control 1. Plough the field to expose the eggs to abiotic and biotic factors. 2. Uproot and destroy the alternate weed hosts. 3. Hand picking of the bugs during early stages and destroy them so as to reduce population build-up. 4. Dislodge the gregarious population of bugs in a container containing water with a thin film of kerosene (Vennila et al. 2007b). Biological Control Antilochus conquebertii Fabricius is the natural predator, which feeds predominantly on the cotton stainers of the genus Dysdercus and other members of the Pyrrhocoridae family (Evangelin et al. 2015). It has a substantial potential for the biological control of the red cotton bug, D. koenigii (Sahayaraj and Fernandez 2017). The predatory behaviour of Hemipteroseius indicus (Krantz & Knot) and their

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possible use as biological control agents against D. koenigii (F.) has also been studied (Banerjee and Datta 1980). The mites prefer the adults with wings intact as compared to nymphs without wings. For the use of microbial control in pest management, three different entomopathogenic fungi, B. bassiana, Isaria fumosorosea and M. anisopliae have been found effective against adults of D. koenigii under laboratory conditions with possibilities of using these entomopathogens as a substitute to chemical pesticides (Khan et al. 2014), The potential of mycoinsecticide, Aspergillus Niger has also been proved as a biocontrol agent against D. koenigii by compromising its defense system (Kumari et al. 2019). Chemical Control Among the insecticidal groups, pyrethroids provide effective control of red cotton bug followed by carbamates, neo-nicotinoids and organophosphates after 24 h of application, while, after 72 h, carbamates give maximum pest mortality followed by organophosphates, pyrethroids and neo-nicotinoids (Rafiq et al. 2014). Akhtar et al. (2016) observed that chlorpyriphos 40 EC gives maximum population reduction when applied after the last picking with irrigation water, while carbosulfan 20 EC and bifenthrin 2.5 EC are most effective in case of foliar application. As it occurs very late in the crop season, chemical management is not recommended as lateseason insecticidal applications may leave residues in the harvested lint in addition to being uneconomical.

8.2.8

Dusky Cotton Bug, Oxycarenus laetus Kirby (Hemiptera: Lygaeidae)

8.2.8.1 Distribution Dusky cotton bug, also known as ‘lint stainer’ is a cosmopolitan pest and widely distributed in Africa, Europe, Tropical and Sub-tropical Asia, Philippine Islands, South America, Brazil, Argentina, Bolivia and West Indies (Slater and Baranowski 1994). 8.2.8.2 Host Plants Abutilon indicum, Sida acuta, Gossypium hirsutum, Thespesia populnea, Hibiscus sabdariffa, H. vitifolius, H. esculentus, Abelmoschus esculentus, Parthenium hysterophorus, Tridax procumbens, Azadirachta indica, Prunus cerasoides, Murraya koenigii, etc. 8.2.8.3 Damaging Stage(s) Nymphs and adults. 8.2.8.4 Diagnostic Features The oval or cigar-shaped eggs are laid either singly or in groups of 2–10 in between calyx and the rind of half opened bolls or in the lint of half or fully opened bolls or on

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Fig. 8.7 Dusky cotton bug

the seeds. Initially they are whitish turning pale and finally becoming light pink before hatching (Srinivas and Patil 2003). There are five nymphal instars. Early instar nymphs are orange in colour when about to moult. After the first moult, the nymphs become reddish brown and then darker after each moult (Vennila et al. 2007b). The adults are small, flat with pointed head and uniform dark brown body with dirty white or dusky brown transparent hemielytra (Fig. 8.7) (Srinivas and Patil 2003).

8.2.8.5 Life Cycle The eggs are usually laid in the lint of half opened bolls, either singly or in small clusters of 3–18 each. The egg stage lasts for 6–10 days and the nymphs, on hatching, pass through five stages, completing the development in 15–23 days. Adult females live longer than males. The life cycle lasts for 33–49 days for males and 35–51 days for females and a number of generations are completed in a year (Srinivas and Patil 2003; Arora et al. 2006).

8.2.8.6 Nature of Damage Both the nymphs and adults suck the sap from the immature seeds. Infested seeds do not ripe and become lighter in weight. Adults of dusky cotton bug get crushed at the time of ginning and stain the lint. Being associated with open bolls they cause nuisance to workers during picking (Vennila et al. 2007b). The population of dusky cotton bug maintained as 25 pairs per boll resulted in maximum lint discoloration (slightly dark yellow), 28–32% reduction in seed germination and 18–21% reduction in seed weight (Ahmed et al. 2015). The quality of lint was reduced with increase in its population. The colour of lint changed from pure-white to white when bolls were exposed to 10 and 15 pairs of bugs, while it changed to light-yellow and slightly yellow when bolls were exposed to 20 and 25 pairs of bugs, respectively (Khan et al. 2014). However, 5-pairs per boll do not reduce the quality of cotton lint (Ahmed et al. 2015).

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8.2.8.7 Management Options Cultural and Mechanical Control 1. Remove alternate host plants like okra, etc. before and near the main crop. 2. Proper management of dusky cotton bug on Psidium guajava, Eucalyptus camaldulensis, Mangifera indica, A. indica and Syzygium cumini, which act as its major hibernating sites so as to prevent its spread from alternate host plants to the cotton crop (Abbas et al. 2015). 3. Dislodge the gregarious population of bugs in a container containing water with a thin film of kerosene (Vennila et al. 2007b). Biological Control Three different entomopathogenic fungi, B. bassiana, I. fumosorosea and M. anisopliae have been found effective against adults of dusky cotton bug with accumulative mortality of 90.0, 75.0 and 75.0%, respectively, over a period of 7 days at the concentrations of 3  108 spores/m under laboratory conditions (Khan et al. 2014). Botanicals Amongst six plant extracts, i.e. neem (A. indica), milkweed (Calotropis procera), moringa (Moringa oleifera), citrus (Citrus sinensis), tobacco (Nicotiana tabacum) and castor (Ricinus communis), tobacco and milkweed @ 2.5 and 5% gave significant control of the dusky cotton bug (Abbas et al. 2015). Similarly Saleem et al. (2018) also observed that N. tabacum exhibits competent insecticidal properties with 53.13, 70.83 and 96.91% mortality at 1.5, 2.5 and 5% concentrations, respectively. Chemical Control The chemical insecticides, monocrotophos 36 SL (700 g a.i./ha), carbaryl 50 WP (2000 g a.i./ha and acephate 75 SP (750 g a.i./ha.) have been found effective for the control of dusky cotton bug. However, botanical (neemguard; 1500 ppm) and IGR (novaluron 10 EC) were least effective (Srinivas and Patil 2004). Under field conditions, Triazophos 40 EC (triazophos), Nurelle-D 505 EC (cypermethrin + chlorpyriphos), Curacron 500 EC (profenophos), Fiprox 5 SC (Fipronil), Adder Plus 360 EC (deltamethrin + triazophos) and mixture of Lancer 2.5 EC (Lambda-cyhalothrin) + Triazophos 40EC (triazophos) have been found effective in reducing dusky cotton bug population after 3 (23.8–55.9%) and 7 days of spray (27.5–54.0%) (Akram et al. 2013).

8.3

Conclusions

Genetically modified cotton having genetic resistance provides innate control of bollworms and tobacco caterpillar but sucking pests are still posing a great threat. For achieving sustained production in the cotton ecosystem, it is emphasized that sucking pest problems should be tackled with IPM approach based on sound

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ecological principles with minimum use of insecticides so that pest population can be managed below damaging level. The remarkable examples are successful management of whitefly and mealybug on cotton crop in Punjab by implementing IPM through farmer’s participatory approach. Further, the study on ecological impact of transgenic cotton and impact analysis of IPM approach on socio-economic status of farmers is of utmost importance.

References Abbas M, Hafeez F, Farooq M, Ali A (2015) Dusky cotton bug Oxycarenus spp. (Hemiptera: Lygaeidae): hibernating sites and management by using plant extracts under laboratory conditions. Pol J Entomol 84:127–136 Ahmed R, Nadeem I, Jawwad YM, Niaz T, Ali A, Ullah Z (2015) Impact of dusky cotton bug (Oxycarenus laetus Kirby) on seed germination, lint color and seed weight in cotton crop. J Entomol Zool Stud 3:335–338 AICCIP (2018) Annual report 2017–18. All India coordinated cotton improvement project. Central Institute for Cotton Research, Regional Station, Coimbatore Akhtar FA, Tariq H, Raza A, Nadeem I, Yousaf J, Ahmed R, Niaz T (2016) Evaluation of different insecticides for the management of red cotton bug Dysdercus spp via flooding and foliar methods of application. Int J Entomol Res 1:16–18 Akram M, Ramzan M, Mehfooz-ul-Haq MA, Saleem MS (2013) Bioefficacy of organophosphates, pyrethroids and new chemistry insecticides against a field population of dusky cotton bug, Oxycarenus spp. (Hemiptera: Oxycarenidae) in Bt cotton ecosystem. Pak J Life Soc Sci 11:48–52 Aneja AK (2000) Studies on the biology of cotton whitefly Bemisia tabaci (Genn.) on American cotton Gossypium hirsutum (Linn.). M.Sc. thesis, Punjab Agricultural University, Ludhiana, India Anonymous (2000) QRT report (1995–2000). All India coordinated cotton improvement project. Punjab Agricultural University, Ludhiana Anonymous (2002) Report on cotton research in Punjab. Punjab Agricultural University, Ludhiana Anonymous (2019) Package of practices for kharif crops in Punjab. Punjab Agricultural University, Ludhiana Arora R, Jindal V, Rathore P, Kumar R, Singh V, Bajaj L (2006) Integrated pest management of cotton in Punjab, India. In: Radcliffe’s IPM world textbook. University of Minnesota, St. Paul, MN. http://ipmworld.umn.edu Banerjee P, Datta S (1980) Biological control of red cotton bug, Dysdercus koenigii Fabricius by mite, Hemipteroseius indicus (Krantz and Khot). Indian J Entomol 42:265–267 Beevi SP, Balasubramanian M (1995) Effect of buprofezin, a novel insect growth regulator, against cotton whitefly Bemisia tabaci Genn. Entomon 20:11–14 Bharathi K, Muthukrishnan N (2017) Evaluation of botanicals against cotton mealybug, Phenacoccus solenopsis Tinsley (Psuedococcidae: Hemiptera). Int J Curr Microbiol App Sci 6:1055–1061 Bheemanna M, Hosamani AC, Hanchinal SG (2010) Bioefficacy of insecticides against mirid bug, Creontiades biseratense (distant) in irrigated Bt cotton. Karnataka J Agric Sci 23:135–136 Birah A, Tanwar RK, Kumar A, Divya S (2018) Development and validation of pest management strategy against mirid bug, Creontiades biseratense in Bt cotton. Indian J Agric Sci 88:1248–1252 Boateng CO, Schwartz HF, Havey MJ, Otto K (2014) Evaluation of onion germplasm for resistance to Iris yellow spot virus and onion thrips, Thrips tabaci. Southwest Entomol 39:237–260 Butter NS, Vir BK (1989) Morphological basis of resistance in cotton to whitefly Bemisia tabaci. Phytoparasitica 17(4):251–261

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9

Sucking Pests of Forage Crops N. S. Kulkarni

Contents 9.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.2 Cultivated Grasses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.3 Cereal Forage Crops . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.4 Forage Legumes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

In developing countries like India, livestock is a major source of additional income to the poor. India has the largest livestock population in the world. These livestock require regular feed which is often obtained through forage crops. The forage crops, especially the cultivated ones are susceptible to attack by multiple pests, which in turn reduces the food of the livestock, thereby causing an indirect loss to the farmers. This chapter discusses the various pests of forage crops, the nature of their damage and their management measures.

9.1

Introduction

Livestock is a major source of livelihood security for the poor in most of the developing countries like India. Livestock has a direct influence on agricultural production. A wide range of products generated from livestock enable farmers to improve their sources of income and to mitigate risk factors. In arid and semi-arid regions, livestock is the only source of livelihood, particularly when expected rainfall fails. In India where over 75% farmers are small and marginal holders, N. S. Kulkarni (*) IGFRI, SRRS, Dharwad, Karnataka, India # Springer Nature Singapore Pte Ltd. 2020 Omkar (ed.), Sucking Pests of Crops, https://doi.org/10.1007/978-981-15-6149-8_9

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livestock is the main source of livelihood for a majority of the rural population. Livestock have been contributing about 15–20% to the household income of farmers, which has been steadily increasing during recent years. The demand for milk and meat is likely to increase by several folds in the coming years. India has the largest number of livestock, representing over 17% of the world population. Among four important species of livestock, cattle represent over 43% of the population followed by buffaloes (19%), goats (26%) and sheep (10%). While cattle and buffaloes are maintained for milk and animal power, sheep and goat are maintained mainly for meat, with milk and wool as secondary sources of income. Cattle and buffaloes, which are considered as milch animals, are large in size, partly stall fed and require substantial quantity of feed and fodder for economic management. However, in case of sheep and goats, most of the population is maintained exclusively on free grazing, although supplementary feeding can significantly benefit their growth, production and reproduction. Fodder crops are the cultivated plant species that are utilized as livestock feed in the form of silage or hay. Numbers of fodder crops are grown across India for various purposes. However, only important forage crops are given in this chapter. Forage crops are mainly divided into two groups, namely grasses and legumes.

9.2

Cultivated Grasses

Grasses are cultivated for grazing as well for cut-and-carry systems. Cultivated grasses include both annual and perennial types. In general, annual types are propagated by seeds only. However, in addition to seeds, perennial types are usually amenable to vegetative propagation. Compared to legumes, insect pest incidence on grasses seems to be less serious. However, due to dwindling land resources for cultivating forage grasses, losses caused by any means or any extend can longer be ignored.

9.2.1

Napier and Napier Hybrid Pennisetum Spp.

It possesses more tillers and leaves than Napier Pennisetum purpureum and is more vigorous and higher in fodder yield and quality. Crude protein ranges from 8 to 11%. Napier grass is also called as elephant grass due to its tallness and vigorous vegetative growth. The plants tiller freely and a single clump may produce more than 50 tillers under favourable climatic and soil conditions. Hybrid Napier is a cross between Bajra
Napier (Pennisetum americanum
Pennisetum purpureum) and it is a perennial grass which can be retained on field for 2–3 years. Compared to Napier grass, Hybrid Napier produces numerous leaves. It has larger leaves, softer and less persistent hairs of leaf blades and sheaths and less sharp leaf edges. The stems are also less fibrous than Napier. The tillers are more numerous and grow faster. Napier grass has a natural built mechanism on the leaves and stems of hairiness which protects it from the insect pests, since the hair stick to the body of the insects

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and paralyses them. Only very hard insects like grasshoppers and stem borers occasionally attack this crop. Grasshoppers can be repelled by spraying 5% NSKE (Neem Seed Kernel Extract) or 2 mL readily available neem formulations and 200 L of spray solution is required per hectare. The sprayed fodder should not be fed to animals for at least for 3 days. If stem borer incidence is severe, apply carbofuran 3G granules @ 10 kg/ac and the treated fodder should not be fed to animals for at least one month or more.

9.2.2

Guinea Grass Panicum maximum

Guinea grass is the most popular fodder grass grown under irrigated conditions in the tropics. It is a highly valued grass because of its wide adaptation, quick growth, ease of establishment, palatability, herbage yield, good persistence and good response to fertilizers (Thomas 2003). It is a tall (1–4.5 m), tufted and fast growing highly palatable perennial grass. It has short creeping rhizome. Planting of guines grass though root slips is preferred over seeds. Crude protein ranges from 4 to 14%. This grass is almost free from insect pests and does not require any plant protection measures.

9.2.3

Brachiaria Spp.

Brachiaria is the single most important genus of forage grass for pastures in the tropics Brachiaria cultivars can grow in infertile and acidic soils. This grass is also called as signal grass and there are several species of Brachiaria exists. Important ones are B. decumbens (Signal grass), B. Brizantha and B. ruziziensis and B. mutica (Para grass). Para grass B. mutica is suitable for cultivation in humid areas. It is grown in seasonally flooded valleys and lowlands and can withstand water logging and long term flooding. It cannot grow on dry lands in arid or semi-arid areas. Waterlogged soils are best suited for this crop. It can be grown on sandy soils also, provided water supply is sufficient. Seed setting is very poor in this grass. It is propagated exclusively by stem cuttings. It can be planted at any time in South Indian conditions, but June–July planting is advisable under rainfed. Brachiaria spp. are infested by few insect pests namely spittle bugs (Homoptera: Cercopidae), burrowing stink bugs (Hemiptera: Cydnidae) and leaf cutting ants (Hymenoptera: Formicidae), etc. Pest management strategies are rarely required in Brachiaria.

9.2.4

Buffel Grass

Cenchrus is a promising green grass type which performs well in dry lands cultivation under rainfed conditions. Cenchrus ciliaris (Anjan grass) and C. setigerus (Black anjan grass) are the two commonly grown species but low yielding in nature.

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One major pest buffel grass caterpillar Mampava rhodenuera is noticed in Australia but not in Indian conditions.

9.3

Cereal Forage Crops

Cereal forage crops are similar to cereal food crops except for the reason that vegetative growth is more important in forage crops rather than grains in cereal food crops. Most of the cereal forage crops are of annual type. However, in case of sorghum perennial sorghum types are gaining more importance than annual forage crops for their vegetation, input requirement and ease of maintenance.

9.3.1

Fodder Sorghum/Perennial Sorghum

Fodder sorghum varieties are annual type and mainly suited for single cut, whereas perennial sorghum is of perennial type and suited for multi-cut. Annual fodder sorghum varieties are as vulnerable as grain sorghum to insect pests. However, perennial sorghum is almost resistant to insect pests. Plant protection measures are required in only annual type varieties. Insect pests of annual type fodder sorghum are same as of grain sorghum and only few among them are important.

9.3.1.1 Aphid Complex Schizaphis graminum, Rhopalosiphum maidis and Melanaphis sacchari Description The apteral Schizaphis graminum are small aphids with a rather elongate oval shape. The head and prothorax are yellowish or greenish straw coloured. The rest of the thorax and the abdomen are yellowish green to bluish green with a darker spinal stripe. The apteral of Rhopalosiphum maidis are rather elongate aphids. The antennae are short. The colour is yellow green to dark olive green or bluish green. Sometimes they are dusted with wax. Around the base of the siphunculi there are dark purplish areas. The siphunculi are dark and short. The length is 0.9–2.4 mm. R. maidis is bluish green in colour with dark green legs. Melanaphis sacchari is yellow coloured aphid and called as sugarcane aphid. Life Cycle Populations are primarily composed of females that reproduce without mating. They do not lay eggs but instead give birth to youngones directly. There are 3–4 nymphal stages seen in aphids. Winged individuals are produced in response to changes in weather, population density and host plant quality. Damage The aphids are found in colonies on the plants. Sap sucking on the leaves often causes yellowing and other phytotoxic effects. Schizaphis graminum are known to be a vector of virus diseases. Leaves, leaf sheaths and inflorescences are infested with colonies of aphids. The leaves may become mottled and distorted. Inflorescences can become sterile. New growth may remain dwarfed. Aphids live

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inside the leaf whorl and suck the sap. They also feed on the panicles and produce honey dew on which sooty mould grows. However, their infestation rarely reaches damaging proportion. The infestation may result in the yellowing, tanning and drying up of leaves. Heavy attacks cause the plant to wilt and die. These aphids are known to be a vector of virus diseases. Management Spraying of insecticides like Dimethoate 0.03% or botanical preparation of NSKE (Neem Seed Kernel Extract) 0.05% + soap. Usually natural enemies such as ladybeetles, Chrysopa, hover fly larvae, parasitic wasps and others control aphid infestations.

9.3.2

Fodder Oats

Oat is grown for grains and for fodder. It is a crop of the temperate region, and in India, it is grown during the rabi season. Among insect pests reported on oats, bird cherry aphid Rhopalosiphum maidis in important one.

9.3.2.1 Bird Cherry Aphid Rhopalosiphum maidis Fitch Description Bird cherry aphid R. maidis is the most common aphid found on cereals. Its colour ranges from orange green to olive green to dark olive green, and sometimes greenish black. It has long antennae and long tube-shaped cornicles arising from the side of the abdomen near the rear end. Wingless forms frequently have a reddish orange patch around the base of the cornicles. Bird cherry-oat aphid may be found any time after seedling emergence. Life Cycle Nature of Damage: Bird cherry-oat aphid attacks all small grains including wheat, barley, oats, rye and triticale. It may also be found on sorghum and corn. Heavy populations may cause a golden yellow streaking on the leaves, unlike white streaks caused by Russian wheat aphid. Occasionally heavy populations cause the flag to curl up in a tight corkscrew fashion that may trap the awns, resulting in a fish-hook appearance to the head. Leaf curl caused by the bird cherry-oat aphid resembles a corkscrew, while that by the Russian wheat aphid resembles an upright soda straw. Management Spray of insecticides like Dimethoate 0.03% or botanical preparation of NSKE (Neem Seed Kernel Extract) 0.05% + soap. Usually natural enemies, such as lady beetles, Chrysopa, hover fly larvae, parasitic wasps and others also control aphid infestations.

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Forage Legumes

Forage legumes belong to the legume or bean family (Fabaceae or Leguminosae). Legumes are second only to grasses in importance to human and livestock nutrition. Important forage legumes include lucerne, cowpea, Stylosanthes, cluster beans and Centrosema. Numbers of insect pests are known to attack these forage legumes and thereby reduce the forage and seed yield qualitatively as well as quantitatively. Details of these insect pests with brief description of their life cycle, damage along with management options are discussed in this article.

9.4.1

Lucerne (Medicago sativa L.)

Lucerne is considered an insectary due to the large number of insects it attracts. Over 1000 species of arthropods have been reported from lucerne fields. Of these fewer than 20 causes injury, and fewer still are serious pests. Although only a few species infest lucerne, they can cause substantial yield and quality losses if present in high numbers. An effective pests management programme can significantly reduce the losses caused by these pests (Summers et al. 2007). Among pests of lucerne Homoptera species (prevalently aphids) were 33.1%, Heteroptera 18% (Lygus rugulipennis being dominant), Coleoptera, 12.9% (Hypera postica), Thysanoptera (thrips), 8.1%, Lepidoptera (caterpillars) 0.6%; and Diptera (flies) and Orthoptera (grasshoppers), 0.1%. Beneficial Arthropods represented 17.2%, and consisted of species belonging to the following groups, in decreasing order: Heteroptera, 6.4%; Hymenoptera, 4.2%; Coleoptera, 3.8%; Aranea, 2.2%; Neuroptera, 0.5%; Diptera, 0.1% (Searpe 1997). Lucerne seed growers must manage the insect pest populations to obtain increased seed production while simultaneously protecting pollinators and other beneficial insects. The major aim of management is to combine mechanical, cultural, biological and chemical control methods with management practices that allow the beneficial insect populations to increase.

9.4.1.1 Aphid Complex (Pea Aphids Acyrthosiphon pisum, Blue Green Aphids A. kondoi, Spotted Lucerne Aphids Therioaphis trifolii and Cowpea Aphids Aphis craccivora Four species of aphids T. trifolii, A. pisum, A. kondoi and A. craccivora were found in India and cause damage on Lucerne. Aphids feed in groups, often on the growing tips of plants. They have piercing-sucking mouthparts that extract the plant sap (phloem). Excess plant sap is excreted as sticky material called honeydew. Severe aphid infestations can retard growth, reduce hay yield, and may even kill lucerne plants. Damage can also reduce the lucerne’s feed value. Black fungus called sooty mould grows readily on the honeydew excreted by aphids reduces palatability of the lucerne hay. Aphids were found responsible for spreading of Verticillium wilt (Verticillium alboatrum) in Egypt (Adawy et al. 2001).

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Fig. 9.1 Pea aphids Acyrthosiphon pisum

Fig. 9.2 Lucerne field damaged by Pea aphids

The Pea Aphids A. pisum (Fig. 9.1) Description A. pisum, commonly known as the pea aphid, is a sap-sucking insect in the Aphididae family. It feeds on several species of legumes (plant family Fabaceae) worldwide, including forage crops, such as peas, clover, lucerne, and broad beans and ranks among the aphid species of major agronomical importance. Pea aphids are capable of asexual reproduction. Each female produces live offspring at a rate of up to 12 per a day, but on average five to six individuals per a day, depending on temperature. Aphids produce many generations per year. Damage The pea aphid is capable of producing 12–15 generations per year. The pea aphid is the least serious pest of this complex, because it does not inject toxin into lucerne plants as it feeds. Pea aphid damage is usually limited to cupping and curling of the leaves and severe burning of the foliage when populations are large (Fig. 9.2).

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Life Cycle Female pea aphids lay fertilized eggs and the nymphs that hatch from these eggs are all females, which undergo four moults before reaching sexual maturity. They will then begin to reproduce by viviparous parthenogenesis, like most aphids. Each adult female gives birth to four to 12 female nymphs per day, around a hundred in her lifetime. These develop into mature females in about 9–10 days. The life span of an adult is about 30 days. Spotted Lucerne Aphids Therioaphis trifolii (Family—Aphididae, Order— Hemiptera) Description Spotted lucerne aphids inflict more serious damage because they inject toxins into the plant as they feed. Toxins injected by the spotted lucerne aphid can stunt growth and cause yellowing of the entire plant. Infested plants have smaller leaves and shorter internodes than normal. Susceptible plants can be killed when populations are heavy. Lucerne that is stressed by lack of water or by cutting is not able to withstand as large an aphid population as healthy unstressed lucerne. Colour of the aphids is right yellow to green, body measures up to 1.5 mm long and abdomen with 6–8 rows of distinctive black spots. Both winged and wingless forms are seen. Life Cycle The Spotted lucerne aphid occurs only as asexually reproducing females. They breed almost continuously throughout the year, but are most favoured by warm, dry weather. Under optimal conditions they are capable of rapid rates of reproduction. Over summer they feed on summer green legumes and lucerne. Spotted Lucerne aphid can produce as many as 35 generations per year. The immature aphids, which closely resemble the adults, moult four times before reaching maturity. The developmental time from newborn nymph to adult is approximately six days depending on the species.The population development of aphid Therioaphis maculata [T. trifolii form maculata] on lucerne was studied in Uttar Pradesh, India, from November to April 1978–1981. The aphid first occurred in mid-November and its population increased up to the 2nd week of January and then declined. There was a negative correlation between temperature and the aphid population, while higher relative humidity was favourable to the aphid. A maximum temperature of 20–25  C and a minimum of 9–10  C and a relative humidity of 80–85% in the morning and 40–55% in the evening were the most conducive conditions for population build-up. Damage Medic plants become stunted, yellow and sticky, eventually dying if under heavy aphid infestation. Seedlings under Spotted lucerne aphid attack become stunted with outer leaves becoming mottled and reddish grey, while younger inner leaves become pale and yellow. Surviving plants can remain unhealthy and unproductive.In lucerne spotted lucerne aphids congregate on the lower parts of the plant and spread to the stems as infestations increase. The leaves turn yellow, wilt and drop off progressively from the bottom of the plant. Heavy infestations left untreated can reduce lucerne stands to stalks. These become sticky with honey dew exuded by

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the aphids and then black sooty mould grows on the honey dew. Attempts to make hay from heavily infested lucerne stands have been unsatisfactory. Hay bales have been severely damaged because the sticky nature of the honeydew prevents free flow of material through the baler. Blue Green Aphid A. kondoi Description Blue green aphids (BGA) are relatively large (up to 3 mm), with a pair of slender tubes like exhaust pipes (cornicles), projecting from the back to beyond the tip of the abdomen. Winged aphids fly into pastures or crops and start colonies of wingless aphids, which cause damage. Overcrowding or plant deterioration triggers the development of new winged aphids which migrate to establish new colonies. Winged aphids can spread viruses. Lifecycle Like other aphids, all BGA are females and give birth to live youngs, without mating. Reproduction rates are very high, so numbers increase rapidly when conditions are favourable. BGA survive hot dry summers in low numbers on sheltered host plants, usually as winged aphids. During heavy infestations, plants can be covered with white speckles, which are cast-off aphid skins. Nature of Damage In lucerne and medics, heavy infestations cause stunted growth, leaf curling and leaf drop. Dry matter production can be reduced. In subterranean clover, leaves wilt before turning grey-brown and dying, becoming dry and “crisp”. Pastures take on a patchy burnt appearance. Seed yield of annual species can be reduced by 20–80%. Forage quality (including phenolic monomers) of germplasms KS153P4 and KS153BA3P4 (susceptible and resistant to A. kondoi, respectively) with and without A. kondoi infestation was compared. Leaves infested with A. kondoi had lower concentration of crude protein and true in vitro digestible dry matter than uninfested leaves of controls. Infestation of lucerne for short time periods by A. kondoi decreased the nutritive value of leaves and stems (Lenssen et al. 1991). Cowpea Aphid Aphis craccivora (Fig. 9.3) Description Cowpea aphid is readily distinguishable from other aphids inhabiting lucerne because it is the only black aphid found infesting the crop. It is a relatively small aphid and the adult is usually shiny black while the nymph is slate grey. The appendages are usually whitish with blackish tips. Cowpea aphid has been a long time resident of lucerne in most part of the world including India. In southern India, populations are highest from August to September; numbers peak in second fortnight of August. This aphid has an extensive host range. In addition to lucerne, it infests many other legumes including cowpea. Life Cycle A. craccivora is a shiny black aphid, feeding on the undersurface of the lucerne leaves, young stems and pods. Parthenogenesis reproduction occurs all year round. The aphid is ovoviviparous. The females retain eggs inside their bodies and

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Fig. 9.3 Cowpea aphid in lucerne Aphis craccivora

give birth to small larvae. Young colonies of this small aphids are found on growing points of plants in association with ants. Nature of Damage Cowpea aphid injects a powerful toxin into the plant while feeding and, when populations are large, this can stunt or kill plants. While feeding, this aphid produces a considerable amount of honeydew upon which sooty mould grows. The black sooty mould reduces photosynthesis and may make leaves unpalatable to livestock. The honeydew also makes the lucerne sticky, which causes problems with harvest. Management of Aphids Monitoring: When the blue green aphid and the pea aphid are present in the same field, it is important to determine the relative proportion of each species, because the blue green aphid can cause more serious damage than the pea aphid. Record the total number of aphids counted for each stem, and calculate the average number of aphids per stem. Aphid infestations in a field are typically patchy, especially an early infestation. Stems on lucerne plants in infested areas are often completely covered with aphids whereas plants in other areas of the field may appear aphid-free. Because of the spotty distribution of cowpea aphid infestations, spot treatments may be feasible, especially if the infestation is on the field border. No guidelines or economic threshold levels have been established for cowpea aphid in lucerne. However, as thumb rule under 10 in. of plant height 10–12 aphids/stem, from 10 to 20 in. plant height 40–50 aphids/stem and over 20 in. 40–50 aphids/stem may be considered. When to Treat? There are a number of factors that should be considered when determining an action/economic threshold for lucerne aphids which includes 1. Value of the commodity—when lucerne hay prices are higher, economic thresholds are lower. 2. Plant height and age of the stand—the shorter the plant, the less aphid pressure it can withstand.

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3. Beneficial insects—high numbers of ladybeetle larvae, lacewing larvae and parasitic wasps may provide adequate biological control. The presence of ladybeetle larvae or adults at a ratio of approximately one lady beetle to 10 aphids can provide adequate natural control. 4. Weather and irrigation scheduling—irrigation and high relative humidity in the crop encourages the natural development of a fungus disease that can completely control aphids. Rain and overhead sprinklers can also reduce aphid numbers in lucerne. 5. Cutting schedule—most aphidicides have restrictions pertaining to harvest interval and number of applications per cutting. 6. Pest complex present—many times Egyptian Lucerne weevil (Hypera postica) populations are coincident with aphids and further the need for insecticidal control. 7. Species of aphid present—the three aphid species have different thresholds depending on their potential severity. The spotted lucerne aphid has the lowest treatment threshold, followed by the blue lucerne aphid, and the pea aphid, respectively. If both blue lucerne and pea aphids are present, use blue lucerne aphid treatment thresholds. Resistant Varieties The most effective means of controlling pea and spotted lucerne aphids is planting resistant varieties. In a field trial conducted at New South Wales, 24 cultivars of lucerne were naturally infested by aphids and their establishment on these cultivars was found to be related to anti-xenosis (influencing the frequency of alates alighting on plants as well as the numbers remaining) and antibiosis (indicated by differences in aphid fecundity) (Holtkamp and Clift 1993). Glandular-haired species were compared with perennial M. sativa clones having resistance or susceptibility to Therioaphis maculata: M. sativa praefalcata (perennial tetraploid), M. blancheana, M. disciformis (both annual diploids), M. rugosa and M. scutellata (both annual tetraploids). All were more resistant (¼ less attractive) than susceptible M. sativa. M. disciformis was significantly more resistant than the other annual species, possibly as a result of possessing dense, simple hairs (in addition to glandular hairs) on both leaf surfaces that may hinder insect feeding (Ferguson et al. 1982). In tests of 270 lines of medic sown in 1978, aphid tolerance was closely related to the plant species. Nearly all the lines of snail medic (M. scutellata) and gama medic (M. rugosa) tested were highly tolerant to aphids (Brownlee and Wetherall 1979). Significant reduction of dry production in aphid susceptible varieties than in aphid resistant varieties was observed (Lodge 1980). Cultural Control Use of border-strip cutting during harvest to help maintain populations of parasites and predators within the field. Border-strip harvesting involves leaving uncut strips of lucerne at various intervals across the field. These serve as a refuge for natural enemy species and to retain predators in the lucerne where they do no harm, thus keeping them out of neighbouring crops, such as cotton or beans, where they cause significant damage. Research has shown that this practice

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significantly increases populations of parasites and predators of aphids, caterpillars and other lucerne insect pests. Biological Control Although parasitism as high as 95% has been documented, aphid population levels can become so high that enough non-parasitized individuals remain to cause significant injury. This aphid is also susceptible to the usual complement of aphid predators including ladybeetles, lacewings, big eyed bugs, damsel bugs and syrphid flies. Early in the season many of these predators are generally not active, but they become active as season progresses. The ladybeetle Coccinella septempunctata and C. transversalis (Fig. 9.4) are abundant and commonly noticed predators feeding on the aphid. Parasitic wasps, hover flies, ladybirds, lace wings and fungal disease are biocontrol agents. Biocontrol agents are only effective when aphid numbers are low. Larvae of common ladybeetles (primarily the convergent ladybeetle, Hippodamia convergens) can attack and consume large numbers of aphids in lucerne. The predominant aphid predator in area lucerne is the convergent ladybeetle. Parasitic wasps that are somewhat host specific also prey on lucerne aphids. The predominate parasitic wasp species of the pea aphid is Aphidius smithi, while the major parasite of the blue lucerne aphid is A. ervi, and the major parasite of the spotted lucerne aphid is Trioxys complanatus. Golden brown aphid mummies attached to the leaves and stems of Lucerne plants indicate parasitic wasps are present and active. Lacewing larvae (Chrysopa spp.) and other generalist predators including Syrphid or hover fly larvae, big eyed bugs (Geocoris spp.), damsel bugs (Nabis spp.) and minute pirate bugs (Orius spp.) also feed on aphids. Aphids also may be controlled by a naturally occurring fungal disease favoured by high relative humidity and cool conditions. Predators and parasitoids are capable of making a significant contribution to lucerne aphid control in a pest management programme (Milne and Bishop 1987). Four species of aphids Therioaphis trifolii, A. pisum, A. kondoi and A. craccivora and predatory insects

Fig. 9.4 Ladybird beetle Coccinella transversalis

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belonging to the families Coccinellidae, Syrphidae, Anthocoridae, Geocoridae and Chrysopidae were found in Brazil (Mendes et al. 2000). Predators C. septempunctata and Harmonia axyridis prefer consumption of aphids over grubs of weevil (Kalaskar and Evans 2001). Important predators of Spotted lucerne aphid were Episyrphus balteatus and the coccinellids C. septempunctata, C. repanda [C. transversalis], Menochilus sexmaculatus and Scymnus nubilus. The syrphid occurred in the fields earlier than the coccinellids which appeared to require higher temperatures (Faruqui et al. 1986). Pathogenicity and virulence of entomogenous fungus, Verticillium lecanii, was studied on the pea aphid, A. pisum. Vertalec, a commercial product of V. lecanii, was evaluated under controlled conditions. Results indicated that Vertalec can be an effective agent against pea aphids. Further studies are recommended for its evaluation under natural conditions (Safavi et al. 2002). Organically Acceptable Methods The uses of biological and cultural controls are acceptable on organically certified crops. Organically certified insecticides such as Azadirachtin, neem oil and mineral oils can be used on lucerne to control aphids. However, studies have shown that at best they provide some suppression of populations but do not control them. Neem was found to be more effective than Malathion in reducing aphid population in Lucerne. Chemical Control There are a number of insecticides available for spotted lucerne aphid control. Care must be taken in selecting the insecticides to avoid the detrimental effects on the biocontrol agents. Following serious outbreak of Therioaphis trifolii on lucerne at Ludhiana, Punjab, India, in May–June 1977, oxydemetonmethyl, dimethoate, phosphamidon and malathion were tested for its control. Five days after spraying, oxydemeton-methyl at 75 and 150 g a.i./ha and dimethoate and phosphamidon at 150 g a.i./ha were effective. Malathion was relatively ineffective. The insecticides did not appear to be very toxic to adults of the predatory coccinellids Menochilus sexmaculatus, Brumus suturalis [Brumoides suturalis] and C. septempunctata, which migrated to untreated plots following insecticide application (Sandhu 1986).

9.4.1.2 Leafhopper Empoasca fabae Description The leafhopper adult is a pale green, wedge-shaped, winged insect about 3 mm long with piercing and sucking mouthparts. It is most broad towards the head, tapering evenly to the wing tips. It has a row of six rounded, white spots behind the head. Nymphs are smaller than adults and are wingless. Life Cycle Both nymphs and adults damage the lucerne by sucking on plant juices. Females lay their eggs in the tissue of main veins and petioles of leaves. Development from egg to adult takes approximately 4 weeks. Nature of Damage Damage is most severe in new seedlings and young regrowth. While leafhopper nymphs and adults suck juices from plant foliage, they inject a

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protein that blocks veins. This causes the edges to become yellow and puckered, with a characteristic yellow “V” shape beginning at the tip of the leaves. When severe, the leaves appear burned, which is called “hopperburn”. Leafhoppers feeding causes reduced stem elongation, reduced root development, leaf cupping and stunting. Yields can be lowered by as much as 50% with a severe infestation, accompanied by a reduction in protein levels of 2–3%. Border areas are usually affected first. The symptoms of leafhopper are commonly confused with herbicide injury problems and nutrient deficiency. High-risk factors include hot, drier-thannormal seasons. Symptoms are sometimes confused with nutrient deficiency or herbicide injury, and are often dismissed as “drought damage”. Monitoring Economic losses occur before plant symptoms develop. So it is important to identify the presence of large leafhopper populations before the damage occurs. It is particularly important to monitor new seedlings. Management Strategies 1. Resistant varieties are available that use glandular hairs as the resistance factor. These glandular hairs, both on the leaves and stems, act as mechanical barriers to leafhopper feeding. The glandular hairs are not fully expressed the first year. 2. Treat new seedlings of leafhopper resistant varieties the same as regular lucerne. 3. Cutting lucerne early will potentially reduce egg, nymph and adult populations. A naturally occurring fungal pathogen helps reduce the populations of the leafhopper under cool and moist conditions. 4. Predators and parasites appear to play a minor role in controlling the pest. If populations exceed the action thresholds, an insecticide may be necessary. 5. Adult leafhopper Empoasca fabae densities per square metre were reduced in the intercrop treatment with oats compared with the lucerne monoculture. Factors that may cause the observed reduction in leafhopper density may be associated with the host plant or the habitat condition (Lamp 1991).

9.4.2

Cowpea Vigna unguiculata (L.)

9.4.2.1 Cowpea Aphids Aphis craccivora (Fig. 9.5) Description Adults are small (up to 2.5 mm long) and are shiny black, while the nymphs are slate grey. Life Cycle Females produce live youngs which grow through wingless nymph stages. Adult males and females may be wingless or winged. All stages may be present in aphid colonies. Colony development is dependent on temperature; it is retarded by cold temperature and increases by hot summer temperatures. Aphids A. craccivora is a medium sized, shiny black aphid whose biology varies depending on climate and soil. Under favourable conditions a generation may take only 13 days. Adults live from 6 to 15 days and may produce more than 100 progeny.

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Fig. 9.5 Cowpea Aphid Aphis craccivora on cowpea pod

Parthenogenesis reproduction occurs all year round. The aphid is ovoviviparous; the females retain eggs inside their bodies and give birth to small larvae. Young colonies of these aphids are found on growing points of plants in association with ants (Soans and Soans 1971; Patro and Behera 1991). Nature of Damage This species of aphid not only causes direct damage to its host but also transmits cowpea aphid-borne mosaic virus. Cowpea aphids inject toxins into the plant while feeding; they most likely reduce vigour and yields. Aphid-feeding also produces honeydew, which grows sooty mould that reduces photosynthesis and makes harvesting difficult. Cowpea aphids may contribute to poor productivity of medic pastures in dry years. Cowpea aphids normally feed on the under surface of young leaves, on young stem tissue and on pods of mature plants. When present in large numbers, they cause direct feeding damage. The plants become stunted, leading to leaf distortion, premature defoliation and death of seedlings. An indirect and generally more harmful effect, even of small populations, is the transmission of cowpea aphid-borne mosaic virus. The aphid while feeding removes sap from the leaves, pods, seeds and other aerial plant parts causing damage to the plant resulting in yield reductions. Infestation with A. craccivora causes significant reductions in seed yield (Ofuya 1989). In a Chinese study A. craccivora infestation resulted in a reduction in plant height to 41.9%, green leaf area and delayed production of harvestable pods by 30 days (Chang and Thrower 1981). Infestations of A. craccivora on cowpea caused reduction in growth and losses in yield (Annan et al. 1997; Attia et al. 1986). Bishara et al. (1984), reported that A. craccivora was the most damaging pest of cowpea in Egypt, particularly early in the growing season. In addition to loss due to damage caused by the aphid, A. craccivora is known to be an important vector of plant viral disease, transmitting over 30 plant viruses (Wightman and Wightman 1994). The aphid also produces honeydew, a substrate which attracts fungi (Mayeux 1984). The aphids attack all growing stages and parts of the plant: flowering, seedling and vegetative growing points including the leaves

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and the plant as a whole. Symptoms of aphids damage on leaves shows distortion, stunting of leaflets, lesions, abnormal colours, premature defoliation and sooty mould. Seeds are shrivelled and growing points show symptoms of rosette (Bottenberg et al. 1998).Whole plant symptoms show stunting, deformities and yield reductions (Annan et al. 1997). Monitoring Inspect crop and sample leaves for winged aphids from early winter onwards and for aphid colonies in spring and early summer. The presence of ladybirds is an indication that aphids are present. Management Biological Control: Pesticides that kill aphid predators should be avoided. Parasitoids are often found in aphids. Heavy rainfall can reduce populations significantly. Chemical Control: This pest is easily controlled with systemic insecticides like dimethoate 30 EC @ 2 mL/L or acephate 1 g/L or imidacloprid 200 SL @ 0.3 mL/L. Normally four applications of insecticide are adequate to control these insect pests. Seed treatment using systemic insecticide or seed fumigation is advised.

9.4.2.2 Leafhoppers Empoasca kerri Description The jassids or leafhoppers are widely distributed throughout the country. It is one of the destructive pests in northwestern region of the country. Damage is caused by the nymphs and adults both which are very agile and move briskly, forward and sideways. Adults are about 3 mm in length and are of greenish colour. The winged adults jump at the slightest disturbance and are positively phototactic in nature. Life Cycle The biology of the several species of Empoasca, which closely resemble each other in appearance is generally similar. Eggs, are laid on the underside of leaves, hatch into nymphs within 7–10 days. There are five stages (instars) in nymphal development which last about 10 days before the adult appears. The adults’ life expectancy varies from 30–60 days. Leafhoppers infest cowpeas at the seedling stage. Nature of Damage The symptoms of damage are yellow discoloration of the leaf veins and margins, followed by cupping of the leaves. Severely infested plants become stunted, so leading to confusion with virus symptoms and may dry prematurely. Management Resistant varieties or genotypes of IGFRI include CL-324, CL-331, CS-55, CS-98, IFC-9102, IFC-9201, UPC-9203, UPC-9204, UP-93-1, UP-93-2, UP-93-3, UP-93-4, UP-9201. Shaking the infested plants over the vessels of oil and water or oily cloth gives effective control. Use of sticky traps also helps in

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suppressing this pest. Spraying of dimethoate 30 EC @ 2 mL/L or Imidacloprid 200 SL @ 0.3 mL/L is advised to control this pest.

9.4.3

Berseem Trifolium alexandrinum

It is an annual leguminous fodder crop. It is one of the most suitable fodder crops for northern India. It remains soft and succulent at all stages of growth. It can be grown without irrigation in areas with high water table and under water-logged conditions. Berseem is usually infested with caterpillars gram caterpillar Helicoverpa armigera and Bihar hairy caterpillar Spilosoma obliqua.

9.4.4

Cluster Bean/Guar: Cyamopsis tetragonoloba

Insect pests like Yellow mite (Polyphagotarsonemus latur and P. datus) and Leaf hoppers (Empoasca sp., Exitianus indicus) cause damage in cluster bean/guar. Yellow mites can considerable damage and affect the overall growth and seed yield of cluster bean.

9.4.4.1 Yellow Mite Polyphagotarsonemus latus Description Female mites are about 0.2 mm long and oval in outline. Their bodies are swollen in profile and a light yellow to amber or green in colour with an indistinct, light, median stripe which forks near the back end of the body. Males are similar in colour but lack the stripe. The two hindlegs of the adult females are reduced to whip-like appendages. The male is smaller (0.11 mm) and faster moving than the female. The male’s enlarged hindlegs are used to pick up the female nymph and place her at right angles to the male’s body for later mating. Life Cycle The eggs are colourless, translucent and elliptical in shape. They are about 0.08 mm long and are covered with 29–37 scattered white tufts on the upper surface. Young broad mites have only three pairs of legs. After one day, the larva becomes a quiescent nymph that is clear and pointed at both ends. The nymphal stage lasts about a day. Nymphs are usually found in depressions on the fruit, although female nymphs are often carried about by males. Damage This destructive pest causes terminal leaves and flower buds to become malformed. The mite’s toxic saliva causes twisted, hardened and distorted growth in the terminal of the plant. Mites are usually seen on the newest leaves and small fruit. Leaves turn downward and turn coppery or purplish. Internodes shorten and the lateral buds break more than normal. The blooms abort and plant growth is stunted when large populations are present. On fruit trees the damage is usually seen on the shaded side of the fruit, so it is not readily apparent. Fruit is discoloured by feeding

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and in severe cases premature fruit drop may occur. Severely damaged fruit is not saleable in the fresh market but may be used for processing. Management While a number of miticides are labelled for control of this pest, insecticidal oils or soaps are usually quite effective and less toxic to the environment. Most common acaricide dicofol @ 1 mL/L is effective for the management of this pest. For large area or greenhouse control, biological control agents are available, including several species of predatory mites.

9.4.5

Stylosanthes Spp.

Important insect pests of Stylosanthes include Pod borer (Helicoverpa armigera), Hairy caterpillars (Spilosoma obliqua), Leafhopper (Empoasca kern), Black aphid (Aphis craccivora). Among these insect pests, pod borer (Helicoverpa armigera) is of economic importance where seed production is practiced.

9.4.6

Centrosema: Centrosema pubescens

Several insect pests like thrips, leaf eating beetles, caterpillars and polyphagous red mites cause damage on different parts of Centrosema and affect the seed production. Among these polyphagous mites are important one. Very little information is available concerning pest biology, distribution, population dynamics and appropriate control measures.

9.4.6.1 Red Spider Mite: Tetranychus urticae Adults and nymphs of spider mites suck the sap from the leaves, causing the area around the feeding punctures to become chlorotic and appear as conspicuous whitish to yellowish stippling on the upper surface of the leaf. Under heavy infestation, photosynthesis is greatly reduced and the chlorotic areas may coalesce forming mottled yellowish interveinal patches. The leaves eventually turn yellow and may become brown and scorched and drop prematurely. Severe infestation affects the overall plant and thereby affects the seed production.

References Adawy EL, Barogy ES, Gaafer EM, Essa MA, El-Sharkawy TA (2001) Role of the aphid in transmission of Verticillium wilt to lucerne. Egyptian J Agri Res 79(4):1329–1338 Annan IB, Ampong-Nyarko K, Tingey WM, Schaefers GA (1997) Interactions of fertilizer, cultivar selection, and infestation by cowpea aphid (Aphididae) on growth and yield of cowpeas. Int J Pest Manage 43(4):307–312 Attia AA, El-Heneidy AH, El-Kady EA (1986) Studies on the aphid, Aphis craccivora, Koch. (Homoptera: Aphididae) in Egypt. Bulletin de la Société Entomologique d'égypte 66:319–324

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Bishara SI, Fam EZ, Attia AA, El-Hariry MA (1984) Yield losses of faba bean due to aphid attack. FABIS Newsletter, Faba Bean Information Service, ICARDA 10:16–18 Bottenberg H, Tamò M, Singh BB (1998) Occurrence of phytophagous insects on wild Vigna sp. and cultivated cowpea: comparing the relative importance of host-plant resistance and millet intercropping. Agri Ecosyst Environ 70(2/3):217–229 Brownlee WP, Wetherall R (1979) Insecticides and varieties to combat lucerne aphids. Wool Tech Sheep Breed 27(4):11–14 Chang LH, Thrower LB (1981) The effect of Uromyces appendiculatus and Aphis craccivora on the yield of Vigna sesquipedalis. J Phytopath 101(2):143–152 Faruqui SA, Pandey KC, Patil BD (1986) Field population studies and natural control of spotted lucerne aphid. Ind J Eco 13(1):120–122 Ferguson S, Sorensen EL, Horber EK (1982) Resistance to the spotted lucerne aphid (Homoptera: Aphididae) in glandular-haired Medicago species. Environ Ent 11(6):1229–1241 Holtkamp RH, Clift AD (1993) Establishment of three species of lucerne aphids on 24 cultivars of lucerne. Australian J Agri Res 44(1):53–58 Kalaskar A, Evans EW (2001) Larval responses of aphidophagous lady beetles (Coleoptera: Coccinellidae) to weevil larvae versus aphids as prey. Ann Ent Soc Am 94(1):76–81 Lamp WO (1991) Reduced Empoasca fabae (Homoptera: Cicadellidae) density in oat-lucerne intercrop systems. Environ Ent 20(1):118–126 Lenssen AW, Sorensen EL, Posler GL, Blodgett SL (1991) Depression of forage quality of lucerne leaves and stems by Acyrthosiphon kondoi (Homoptera: Aphididae). Environ Ent 20(1):71–76 Lodge GM (1980) Effects of spotted lucerne aphids and blue-green aphids on the dry matter production of some lucerne varieties. In: Wood IM (ed) Proceedings of the Australian agronomy conference “pathways to productivity”. Queensland Agricultural College, Lawes, p 261 Mayeux A (1984) The groundnut aphid. Biology and control. Oléagineux 39(8/9):425–434 Mendes S, Cervino MN, Bueno VHP, Auad AM (2000) The diversity of aphids and their parasitoids and predators in lucerne crops. Pesuisa Agri Brasil 35(7):1305–1310 Milne WM, Bishop AL (1987) The role of predators and parasites in the natural regulation of lucerne aphids in eastern Australia. J App Eco 24(3):893–905 Ofuya TI (1989) The effect of pod growth stages in cowpea on aphid reproduction and damage by the cowpea aphid, Aphis craccivora (Homoptera: Aphididae). Ann App Bio 115(3):563–566 Patro B, Behera MK (1991) Mutualism between the bean aphids (Aphis craccivora Koch) and ants. Orissa J Agri Res 4(3–4):238 Safavi SA, Rassulian GR, Askary H, Pakdel AK (2002) Pathogenicity and virulence of entomogenous fungus, Verticillium lecanii (Zimm.) Viegas on the pea aphid, Acyrthosiphon pisum (Harris). J Sci Tech Agri Natur Res 6(1):245–255 Sandhu GS (1986) Chemical control of spotted lucerne aphid Therioaphis trifolii (Monell) on lucerne with reference to conservation of Coccinellid predators. Ind J Pl Prot 13(2):125–127 Searpe DC (1997) Structure and abundance of Arthropods in seed lucerne crops at Dabuleni. Analele-Institutului-de-Cercetari-pentru-Cereale-Protectia-Plantelor 28(1):69–78 Soans AB, Soans JS (1971) Proximity of the colonies of the tending ant species as a factor determining the occurrence of aphids. J Bom Nat His Soc 68(3):850–851 Summers CG, Godfray LD, Natwick ET (2007) Managing insects in lucerne, vol 8295. Division of Agriculture and Natural Resources, University of California, Davis, p 24 Thomas CG (2003) Forage crop production in the tropics. Kalyani Publishers, New Delhi, p 333 Wightman JA, Wightman AS (1994) An insect, agronomic and sociological survey of groundnut fields in southern Africa. Agr Eco Environ 51(3):311–331

Part II Horticultural Crops

Sucking Pests of Vegetable Crops

10

A. T. Rani, K. Vasudev, K. K. Pandey, and B. Singh

Contents 10.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.2 Major Group of Sucking Insect Pests Infesting Vegetable Crops in India . . . . . . . . . . . . . 10.3 Integrated Management of Sucking Pests of Vegetable Crops . . . . . . . . . . . . . . . . . . . . . . . . . 10.4 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

308 314 333 339 340

Abstract

Vegetable crops are cultivated worldwide for its nutritional benefits and neutraceutical properties. It is the only source to meet the goal of nutritional security. Sucking insect pests are considered as one of the major biotic constrains for vegetable production in India. These pests cause direct damage by sucking the sap via specially adapted mouthparts and secrete the sugar rich honeydew deposit on plant surface and create the black sooty mould, thereby hindering the normal photosynthesis of the plants. Apart from direct damage, they also act as vectors for several viral diseases. In recent past, some of the insect pests of vegetable crops become major and are gradually attaining the major pest status in different regions of the country due to changes in the cropping pattern, ecosystems and habitat, climate and wider use of high input intensive vegetable varieties/hybrids. Sucking pests like whitefly (Bemisia tabaci); leafhopper (Empoasca motti) on bitter gourd; red spider mite (Tetranychus spp.) on okra, brinjal, cowpea, and Indian bean; yellow mite (Polyphagotarsonemus latus) on chilli; and mealybug (Phenacoccus solenopsis) on okra, chilli, brinjal, and tomato, especially in A. T. Rani (*) · K. K. Pandey · B. Singh ICAR-Indian Institute of Vegetable Research, Varanasi, Uttar Pradesh, India K. Vasudev Department of Food and Public Distribution, Ministry of Consumer Affairs and Food and Public Distribution, Government of India, Krishi Bhawan, New Delhi, India # Springer Nature Singapore Pte Ltd. 2020 Omkar (ed.), Sucking Pests of Crops, https://doi.org/10.1007/978-981-15-6149-8_10

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protected conditions have intensified the severity of occurrence in different parts of country. Success of management of insect pests highly depends on correct identification and choice of proper control measures. An attempt has been made in this book chapter to compile information on pest identification, its biology, and nature of damage and integrated management of sucking pests for sustainable vegetable production.

10.1

Introduction

The horticulture production has become a key source for economic development in the country. Globally, India is the second largest producer of fruits and vegetables. In the foreign trade, export growth of fresh fruits and vegetables in term of value is 14% and of processed fruits and vegetables is 16.27%. Production has come down around 30% of the total economy of the country only due to attack of insect pests. Pest associated losses increased from an average of 7.2% during the pre-green revolution period (early 1960s) to 23.32% during the post-green revolution period (early 2000s) in different crops in India in spite of advancement in technology (Dhaliwal et al. 2010). Insect pests on an average are estimated to cause 15–20% yield losses in principal food and cash crops (Chakrabarty 2015). Vegetable crops are cultivated worldwide for its nutritional benefits and neutraceutical properties. Consumption of these items provides taste, palatability, increases appetite and provides fibre for digestion and to prevent constipation. They also play key role in neutralizing the acids produced during digestion of pretentious and fatty foods and also provide valuable roughages which help in movement of food in intestine. Some of the vegetables are good sources of carbohydrates (leguminous vegetables, sweet potato, potato, onion, garlic and methi), proteins (peas, beans, leafy vegetables, and garlic), vitamin A (carrot, tomato, drumstick, leafy vegetables), vitamin B (peas, garlic, and tomato), vitamin C (green chillies, drumstick leaves, Cole crops, leafy vegetables, and leaves of radish), minerals (leafy vegetables, drumstick pods). As per dietician, daily requirement of vegetables is 75–125 g of green leafy vegetables, 85 g of other vegetables, and 85 g of roots and tubers with other food. Insect pests are the major biotic constraints to vegetable production in India. Among the different insect pests of vegetables, sucking pests are causing serious damage to agricultural and horticultural crops reducing both quality and quantity of produce. Like many hemipteran pests of crops, they are primarily phloem feeders, abstracting sap via specially adapted mouthparts and also secrete the sugar rich honeydew which deposits to the plant surface and creates the black sooty mould, and thereby hindering the normal photosynthesis of the plants. Apart from direct damage, they also act as vectors for several viral diseases (Table 10.1). The crop losses to the tune of 30–40% due to insect pests have been reported in vegetable crops (Table 10.2). The severity of pest problems has been changing with the developments in agricultural technology and modifications of farming practices.

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Table 10.1 Viruses transmitted by sucking insect pests of vegetable crops Insect species Bemisia tabaci

Disease/virus transmitted Tomato leaf curl virus (ToLCV)

Yellow vein mosaic virus (YVMV), enation leaf curl virus Golden mosaic disease/yellow mosaic disease Squash leaf curl virus (SLCV)

Myzus persicae, Aphis gossypii, A. craccivora Myzus persicae, Aphis gossypii

Curly shoot disease Papaya ring spot virus (PRSV)

Zucchini yellow mosaic virus (ZYMV)

Aphis spp.

Cucumber mosaic virus (CMV)

Frankliniella occidentalis, Thrips palmi Thrips palmi

Groundnut bud necrosis virus/ Peanut bud necrosis virus (GBNV/PBNV) Capsicum chlorosis virus (Gloxinia tospovirus) (CaCV) Iris yellow spot virus (IYSV) Watermelon bud necrosis virus (WBNV)

T. tabaci T. palmi

Crop Tomato, eggplant, cucurbits, cotton, chilli, pumpkin, cucumber and melon, sponge gourd, chayote, squash, bottle gourd, cucumber, muskmelon, and weeds Okra

Cowpea, dolichos bean, French bean Pumpkin, bitter gourd, ridge gourd, ash gourd, squash, chayote, summer squash French bean Cucurbits

Zucchini, squash, muskmelon, pumpkin, watermelon, yellow squash, bottle gourd, gherkins, snake gourd, and bitter gourd Artichoke, beans (broad, lima, snap), sugar beet, carrot, celery, parsley, lettuce, pea, pepper, potato, sweet potato, spinach, brinjal and tomato, chickpea lentil, soybean, and yams Tomato, brinjal, chilli, watermelon, bitter gourd, cowpea, peas, potato, green gram, groundnut, sunflower Tomato, chilli

Watermelon, ridge gourd, cucumber, muskmelon, bitter gourd, pumpkin, chilli, tomato

(Source: Nagendran et al. 2017) Table 10.2 Yield losses due to major insect pests in major vegetables in India Crop Tomato Chilli Okra

Pest Whitefly (Bemisia tabaci) Thrips (Scirtothrips dorsalis) Mites (Polyphagotarsonemus latus) Leafhopper (Amrasca biguttula biguttula) Whitefly (Bemisia tabaci)

Yield loss (%) 60–90 12–90 34 54–66 54

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The changing scenario of insect pest problems in agriculture as a consequence of green revolution technology has already been well documented. There has been further shift in the status of several insect pests after the introduction of transgenic crops and the current scenario of climate change (Dhaliwal et al. 2010). In recent past, some of the sucking insect pests of vegetable crops become major and are gradually attaining the major pest status in different regions of the country due to changes in the cropping pattern, ecosystems and habitat, climate and wider use of high input intensive vegetable varieties/hybrids. Sucking pests like whitefly (Bemisia tabaci), leafhopper (Empoasca motti) on bitter gourd, red spider mite (Tetranychus spp.) on okra, brinjal, cowpea, and Indian bean yellow mite (Polyphagotarsonemus latus) on chilli especially in protected conditions have intensified the severity of occurrence in different parts of the country. More recently mealybug (Phenacoccus solenopsis) an invasive, emerging, polyphagous pest has been observed in serious proportion on a number of solanaceous (chilli, brinjal, and tomato), malvaceous (okra), and cucurbitaceous vegetables and other crops including many weeds. The important sucking pests of vegetable crops are listed below in Table 10.3. Success of management of insect pests highly depends on correct identification and selection of appropriate control measures. Among various methods of pest control, chemical method still enjoys first choice because of its quick action and easy availability. Most of the plant protection recommendations in vegetables so far indicated the calendar based application of pesticides. This has become a common practice over the years by most of the farmers growing vegetables in the country. Vegetables consume about 13–14% of the total pesticide consumption in India. Presently, the maximum pesticide usage is in chilli (5.13 kg a.i./ha) followed by brinjal (4.60 kg a.i./ha), cole crops (3.73 kg a.i./ha), and okra (2–3 kg a.i./ha) (Rai et al. 2014b). Chemical method of management has its own demerits like resistance to insecticides, resurgence of target insects, and secondary pest outbreak in addition to these residues to food and beverages, contamination of groundwater, adverse effect on environment and human and animal health’s, and wide spread killing of nontarget organisms. Unlike cereals, the green pods and fruits harvested at shorter intervals and used as vegetables are prone to retain pesticide residue. The residue problem becomes more severe when the good agricultural practices (GAPs) are not followed. In the present context of food and environmental safety, biological and non-chemical approaches for the management of pests are being emphasized in developing biointensive integrated pest management (IPM) technologies to reduce the toxic residues in the produce and cost of inputs. However, field efficacy of biocontrol agents in many cases has failed to combat the pests due to evolution of new biotypes/races in the insect population. Hence, it is imperative to develop, validate, and implement multifaceted approaches consisting of biological, host plant resistance/tolerance, cultural practices, etc., to manage the pests. In the last couple of decades, researches on vegetable pest management have been reoriented in this direction that has resulted in the generation of extensive information on these aspects. An attempt has been made in this book chapter to compile all the

Sharpshooter leafhoppers Indian wax scale

Brevicoryne brassicae (Linnaeus) Bothrogonia albidicans (Walker) Ceroplastes ceriferus (Fabricius) Cletomorpha hastata (Fabricius)

Bagrada hilaris (Burmeister) Bemisia tabaci (Gennadius)

Cicadellidae Coccidae Coreidae

Hemiptera Hemiptera Hemiptera

Aphididae

Aleyrodidae

Hemiptera

Hemiptera

Pentatomidae

Aphididae

Aphididae

Aphididae

Hemiptera

Hemiptera

Aphis spiraecola Patch

Harlequin bug, painted bug Tobacco whitefly, sweet potato whitefly, cotton whitefly Mustard aphid

Hemiptera

Aphis craccivora Koch

Groundnut aphid, cowpea aphid Spirea aphid, green citrus aphid

Hemiptera

Coreidae

Hemiptera

Aphis gossypii Glover

Cicadellidae

Aleyrodidae

Hemiptera

Hemiptera

Melon aphid, cotton aphid

Cotton leafhopper, okra leafhopper Pod bug, squash bug

Spiralling whitefly

Mango, ber, citrus, cucurbits, coffee, ficus, amaranthus, rose Brinjal, amaranthus, and many other plants

Ragi, rice, okra, horse gram

Cotton, tomato, sweet potato, tobacco, cassava, pulses, chillies, brinjal, okra, ornamentals, etc. Mustard, cabbage, cauliflower, knol-kohl

Cotton, hibiscus, okra, watermelon, cucumber, pumpkin, tomato, potato, brinjal Cowpea, beans, peas, groundnut, several other leguminaceous plants Citrus, crucifers, potato, pepper, tobacco, apple, Spiraea, Prunus, and a variety of ornamentals Mustard, cabbage, cauliflower

Pigeon pea, cowpea, field bean, brinjal

Agricultural and horticultural crops, ornamentals Cotton, okra, brinjal

Crop(s) damaged Brinjal and other Solanum spp.

Minor

Minor

Minor

(continued)

Minor, serious in protected cultivation Major

Major

Major

Major

Minor, occasionally serious Major

Major

Major

Pest status Minor

Common name Brinjal bug

Family Coreidae

Table 10.3 List of sucking pests of major vegetable crops Order Hemiptera

Sucking Pests of Vegetable Crops

Scientific name Acanthocoris scabrator (Fabricius) Aleurodicus dispersus Russell Amrasca biguttula biguttula (Ishida) Anoplocnemis phasiana (Fabricius)

10 311

Tomato suck bug, tobacco capsid/mirid

Giant red bug

Egyptian cottony cushion scale Lantana bug, croton bug

Lygus sp. Macrocheraia grandis (Gray) Nesidiocoris tenuis (Reuter)

Frankliniella occidentalis (Pergande), F. schultzei (Trybom) Icerya aegyptiaca (Douglas) Insignorthezia insignis (Browne)

Hemiptera

Drepanococcus chiton (Green) Eurydema sp. Ferrisia virgata (Cockerell)

Stink bug

Painted bug Striped mealybug, guava mealybug, grey mealybug Western flower thrips

Hemiptera

Dolycoris indicus Stal

Cucurbit stink bug, red pumpkin bug

Hemiptera

Miridae

Miridae Largidae

Ortheziidae

Hemiptera

Hemiptera Hemiptera

Monophlebidae

Thripidae

Pentatomidae Pseudococcidae

Coccidae

Pentatomidae

Dinidoridae

Coccidae

Family Pseudococcidae

Hemiptera

Thysanoptera

Hemiptera Hemiptera

Hemiptera

Hemiptera

Coccus hesperidum Linnaeus Coridius janus (Fabricius)

Brown soft scale

Order Hemiptera

Scientific name Coccidohystrix insolita (Green)

Common name Eggplant mealybug

Table 10.3 (continued)

Brinjal Cotton, okra, hibiscus, Trewia nudiflora, Bombax ceiba Tomato, tobacco, brinjal, sesame, cucurbits, sorghum

Cucurbits, cabbage, eggplant, tomato, carrot, peas, beans, capsicum, tobacco, cotton, ground nut, grain legumes Tomato, capsicum, grapes, sapota, apple, banana, guava, jack, hibiscus, lantana Lantana, croton, coffee, brinjal, ornamentals

Cabbage, mustard, and cauliflower Cotton, guava, custard apple, papaya, beans, crotons, pepper

Citrus, mango, custard apple, lychee, chilli, coffee, cotton, hibiscus Cucurbits (pumpkin, bottle gourd), brinjal, Calotropis, lablab, Abutilon muticum Pigeon pea, maize, sorghum, sunflower, amaranthus Pigeon pea, papaya, brinjal, tea, cocoa

Crop(s) damaged Pigeon pea, brinjal, other Solanum spp.

Minor

Serious on several ornamentals Minor Minor

Major as vector, occasionally serious Minor

Minor Occasionally serious

Minor

Minor

Minor

Pest status A common and often serious pest of brinjal Minor

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Cotton white scale, hibiscus snow scale, lesser snow scale Red spider mite, two-spotted spider mite Brinjal lace wing bug, eggplant lace bug

Red banded shield bug, legume stink bug, lucerne shield bug

Sweet potato bug

Solenopsis mealybug

Lantana mealybug

Madeira mealybug

Green stink bug

Tetranychidae Tingidae

Acari Hemiptera

Tetranychus urticae Koch Urentius hystricellus (Richter)

Diaspididae

Pentatomidae

Coreidae

Pseudococcidae

Pseudococcidae

Pseudococcidae

Pentatomidae

Hemiptera

Hemiptera

Hemiptera

Hemiptera

Hemiptera

Hemiptera

Hemiptera

Pinnaspis strachani (Cooley)

Phenacoccus solenopsis Tinsley Physomerus grossipes (Fabricius) Piezodorus hybneri (Gmelin)

Nezara viridula (Linnaeus) Phenacoccus madeirensis Green Phenacoccus parvus Morrison

Cowpea, soybean, lucerne, lentil, and other pulses; also reported on cotton, sorghum, potato, tomato, chillies, brinjal, linseed Castor, mango, pumpkin, drumstick, pomegranate, capsicum, brinjal, black pepper, and curry leaf Pepper, tomato, potato, beans, corn, strawberries, rose, castor Brinjal

Sorghum, rice, pigeon pea, cotton, okra, brinjal, castor, vegetables Cotton, hibiscus, tomato, potato, brinjal, acalypha Naga King chili, Capsicum chinense, C. annuum, Amaranthus sp., tomato, China aster, Callistephus chinensis, Chrysanthemum sp. Cotton, okra, tomato, potato, hibiscus, parthenium Sweet potato, tamarind

Minor

Major

–

Minor

Minor

Minor

Minor

Minor, rarely serious Minor

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information on pest identification, its biology, and nature of damage and integrated management of sucking pests for sustainable vegetable production.

10.2

Major Group of Sucking Insect Pests Infesting Vegetable Crops in India

10.2.1 Whiteflies Taxonomic Position Phylum: Arthropoda. Class: Insecta (Insects). Order: Hemiptera (True Bugs, Hoppers, Aphids, and Whiteflies). Suborder: Sternorrhyncha (Plant-parasitic Hemipterans). Superfamily: Aleyrodoidea. Family: Aleyrodidae (Whiteflies). Amongst 1200 species of whiteflies, Bemisia tabaci (Gennadius) is considered as the most important one. It is a cosmopolitan; highly polyphagous with remarkable ability to adapt to new hosts (Gerling and Kravchenko 1995). It is widely distributed in tropical and subtropical regions. In temperate regions this pest is majorly found in greenhouses and throughout the northern and western regions of the Indian subcontinent. It has been reported to attack more than 600 host plant species (Secker et al. 1998) belonging to 77 families (Basu 1995) in different parts of the world. Hot and dry conditions are suitable for the infestation of whitefly, while heavy rain drastically reduces build-up of its population. This insect is active during the day time and settles on lower leaf surfaces at night. The economic damage may result from direct feeding leading to devitalization of the plants, yellowing, and finally death of the plant. They also serve as vector for many dreaded viral diseases. This pest was earlier considered as a minor pest and later they were found to attack many vegetable crops and cause havoc loss both in field and greenhouse conditions. In recent times, severe incidence of leaf curl virus and yellow vein mosaic disease transmitted and spread by whitefly, has evoked wide spread concern in northern India. Similarly, the detection of Biotype-B of this pest in Karnataka on tomato causing severe incidence of leaf curl has sent a scare among the tomato growers in the country (Rai et al. 2014a). However, efforts are on to develop suitable management strategy for B-Biotype of whitefly as a pest and vector is the need of the hour.

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10.2.1.1 Cotton Whitefly, Bemisia tabaci (Gennadius) and Greenhouse Whitefly, Trialeurodes vaporariorum Diagnostic Features B. tabaci The eggs are stalked, sub-elliptical, and light yellow at first and turns brown later on. Nymphs are oval, scale like, elliptical, greenish-yellow and remain attached to the leaf surface. Adults are tiny, with yellowish body and wings coated with milky white waxy powder (Fig. 10.1a). Their pale bodies make them stand out against the green background. They are sluggish creatures, clustered together on the underside of the leaves. T. vaporariorum Eggs are stalked and remain suspended on the lower surface of the leaf. Newly emerged nymphs are light yellow. Greenhouse whiteflies are small insects with white coloured wings. Biology B. tabaci The insect breeds throughout the year. The females mostly lay eggs singly near the veins on the underside of the leaves. Each female can lay about 300 eggs in its lifetime. Eggs are small (about 0.25 mm) and vertically attached to the leaf surface through a pedicel. The eggs are not visible to the naked eye and must be observed under a magnifying lens or microscope. Egg period is about 3–5 days during summer and 5–33 days in winter. On hatching, the first instar larvae nymphs move on the leaf surface to locate a suitable feeding site. Hence, it is commonly known as crawler. Nymphs grow through three stages. The first instar nymph has antennae, eyes, and three pairs of well-developed legs. The legs and antennae are atrophied during the next three instars and become stationary throughout the remaining stages. The last nymphal stage has red eyes. This stage is sometimes referred to puparium, although insects of this order (Hemiptera) do not have a perfect pupal stage (incomplete metamorphosis). Nymphal period is about 9–14 days during summer and 17–73 days in winter. Adults emerge from puparia through a T-shaped slit, leaving behind empty pupal cases or exuviae. The whitefly adult is a soft bodied, moth-like fly. White flies are minute insects with yellow body covered with a white waxy bloom. The wings are held over the body like a tent. The adult males are slightly smaller in size than the females. Adults live from 1 to 3 weeks. Total life cycle is completed in 14–122 days with 10–15 generations in a year. T. vaporariorum Eggs are stalked, laid on the lower surface of leaf. Incubation period is 3–8 days. There are four nymphal stages. Development of first, second, third, and fourth instar nymph is completed in 2–6, 5–8, 3–5, and 3–6 days, respectively. Full grown nymph pupates in a yellow pupal case surrounded by a waxy palisade and waxy fringe. Adults are minute, pale coloured body, and remain hidden on the under surface of leaves. The total life cycle is completed in 15–32 days.

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Fig. 10.1 Major sucking pest of vegetables: (a) Bemisia tabaci (Gennadius); (b) Aphis gossypii Glover; (c) Brevicoryne brassicae (Linnaeus); (d) Lipaphis erysimi (Kaltenbach); (e) Aphis craccivora Koch; (f) Amrasca biguttula biguttula (Ishida); (g) Urentius hystricellus (Richter); (h) Dysdercus sp.; (i) Bagrada hilaris (Burmeister); (j) Coridius janus (Fabricius); (k) Nezara viridula (Linnaeus); (l) Coccidohystrix insolita (Green); (m) Phenacoccus solenopsis Tinsley; (n) Frankliniella schultzei (Trybom), and (o) Tetranychus urticae Koch. (Source: NBAIR database (n.d.), http://www.nbair.res.in/insectpests/index.php)

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Host Range and Disease Transmission B. tabaci is highly polyphagous and is known to feed on several vegetables including tomato, brinjal, chillies, cow pea, field bean, cucumber, okra, peas, melon, potato, leguminous vegetables, on field crops and some weeds. A major vector of viral diseases and is known to transmit more than 60 plant viruses, like cassava mosaic virus, tomato yellow leaf curl virus, cotton leaf curl geminivirus, tobacco leaf curl, okra yellow vein mosaic virus, etc. T. vaporariorum is known to feed on tomato and cucurbits under greenhouse condition. Nature of Damage Tomato Nymphs and adults suck the sap usually from the under surface of the leaves. The severely infested plants show yellowing, downward curling, and drying of leaves. Brinjal Both the adults and nymphs suck the plant sap and reduce the vigour of the plant. In severe infestations, affected plants show numerous chlorotic spots on leaves/yellowing, the leaves turn yellow and drop off. When the populations are high they secrete large quantities of honeydew, which favours the growth of sooty mould on leaf surfaces and reduces the photosynthetic efficiency of the plants. Okra Nymphs and adults suck the cell sap and lower the vitality of the plant. Chlorotic spots on the leaves, which latter coalesce forming irregular yellowing of leaf tissue, extend from veins to the outer edges of the leaves. Severe infestation results in premature defoliation. They also excrete honeydew on which sooty mould grows which interferes with the photosynthesis of the plants. Affected plants give a sickly black appearance. B. tabaci transmits a vein clearing disease of okra.

10.2.2 Aphids Taxonomic Position Phylum: Arthropoda. Class: Insecta (Insects). Order: Hemiptera (True Bugs, Hoppers, Aphids, and Whiteflies). Suborder: Sternorrhyncha (Plant-parasitic Hemipterans). Superfamily: Aphidoidea. Family: Aphididae (Aphids). Aphids are small, succulent, pear shaped insects that vary in colour from yellow to green to black. This is a cosmopolitan pest and highly polyphagous. Both the nymphs and adults possess piercing and sucking mouthparts. It prefers to feed on cotton, cucurbits, eggplant, and okra. Cool, dry, and humid conditions are favourable for multiplication of aphids while heavy rains wash away the aphid colonies. The

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yields are also reduced directly by sucking sap and more through the spread of virus diseases acting as vectors indirectly.

10.2.2.1 Cabbage Aphid, Brevicoryne brassicae Linnaeus, Mustard Aphid, Lipaphis erysimi (Kaltenbach), Cotton Aphid, Aphis gossypii Glover, Green Peach Aphid, Myzus persicae (Sulzer), Pea Aphid, Acyrthosiphon pisum (Harris), and Black Bean Aphid, Aphis craccivora Koch Diagnostic Features B. brassicae Eggs are pale-yellow with greenish tinge. Nymphs are 1–1.5 mm long and yellow-green with light ash grey tinge. Apterae are green and covered with a greyish white mealy wax that is also secreted on the plant and spreads throughout the colony (Fig. 10.1c). The head, tips of the antennae, and the legs are dark. Their siphunculi are thick and very short, 0.06–0.07 times the body length and 0.8–1.0 times the length of the cauda. The cauda is triangular and broad. L. erysimi The wingless female is pale green or whitish green with two rows of dark bands on the thorax and abdomen which unite into a single band near the tip of the abdomen. The antennae are dark, the legs are pale with dark joints, and the cornicles are pale with dark tips. The body is faintly dusted with a white powder. The winged female is a similar size and has a black head and thorax and a pale green abdomen with black bands near the tip and black patches on the sides (Fig. 10.1d). The antennae and legs are dark, and the cornicles are black at the base and yellowish towards the tips. Wingless males have occasionally been seen. These are smaller than the females and olive-green to brownish in colour. A. gossypii Eggs are very tiny, shiny black and are found in the crevices of bud, stems, and barks of the plant. Nymphs are greenish brown or yellowish, found on the undersurface of leaves. Adults are yellowish-green to dark green, 1–4 mm long, soft bodied insects with two long antennae that resemble horns (Fig. 10.1b). They possess a pair of siphunculi (cornicles) near the posterior side of the abdomen. Wings when present are transparent with black veins. A. craccivora Adults are greenish black in colour. The female has a glossy black or dark brown body with a prominent cauda (tail-like protrusion), and legs are brown or yellow in colour (Fig. 10.1e). The antennae have six segments and the limb segments, cauda and cornicles are pale proximally (close to the body) and dark distally (further from the body). The adults do not have wax on their dorsal surface but the nymphs are lightly dusted with wax. Winged females are up to 2.2 mm (0.1 in.) long and have crossbearing on the abdomen. Wingless females are a little smaller.

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M. persicae Eggs are elliptical, initially yellow or greenish, but soon turn black. Nymphs initially are greenish, but soon turn yellowish, greatly resembling viviparous adults. Adults of M. persicae are usually green in colour but may be pale brown to pinkish, 1.5–2.5 mm long with abdomen having a pair of siphunculi. Acyrthosiphon pisum Adult aphids are large pear shaped, green, yellow, or pink in colour with long conspicuous cornicles. Both alates as well as apterous forms are present. Biology B. brassicae This pest infests crucifers in cold season. The pest is active from October to April. In mid hills of Himachal Pradesh, it appears in the last week of January with peak during first week of April. It reproduces through parthenogeneticviviparous; however, during severe winter, sexual reproduction also occurs. There are four nymphal instars. The nymphs mature in 10–15 days and immediately start laying eggs. Overcrowding coupled with high temperature and low humidity results in appearance of alates for migration. A single female can produce 40–45 young ones. Total life cycle is completed in 10–45 days. There are many generations in a year. A. gossypii Unlike many insects, most aphids do not lay eggs. They usually reproduce through parthenogenetic-viviparous but during cooler winter, eggs are laid. When the temperature rises, the eggs hatch. Nymphs are young aphids; they look like the wingless adults but are smaller. They mature in 3–4 days and reproduce parthenogenetically. The adult colour is highly variable and it varies from light green to greenish brown. Both wingless and winged forms occur. Winged forms are produced predominantly under high population density conditions, inferior host plant quality and when temperature rises which migrate to other crops. The wingless forms are more common. They possess a pair of black-coloured cornicles on the dorsal side of the abdomen. Aphids mostly are found in groups. Each female produces about 20 nymphs a day, which become adults in a week. Acyrthosiphon pisum Females are common and males are rare. Males have been reported from Europe and USA but not from India. Reproduction is parthenogenic and viviparous. One generation will be completed in a week. Undergo several overlapping generations in a year. Host Range and Disease Transmission B. brassicae feed on all cruciferous vegetables. L. erysimi feed on all cruciferous vegetables, leafy vegetables, spinach, etc. A. gossypii is known to feed on brinjal, chilli, tomato, okra, cucurbits and cotton, tobacco. A. craccivora: Leguminous vegetables, peas, beans, cowpea, etc.

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M. persicae is found to feed on tomato, brinjal, chilli, cabbage, cauliflower, cucurbits, leafy vegetables, potato, spinach, etc. Acyrthosiphon pisum: Peas. Nature of Damage Solanaceous Vegetables Both nymphs and adults of A. gossypii and M. persicae appear in large numbers on the tender shoots, leaves, and on the lower surface of the leaves. They cause damage by sucking the cell sap from leaves and tender apical shoots and reduce the vigour of the plant. The affected parts turn yellow, get deformed, curling and crinkling of leaves and dry away. Excrete copious quantity of honeydew on which sooty mould grows, covering the affected parts with a thick black coating which hampers the photosynthetic activity. The infested plants become weak, pale, and stunted in growth and consequently bear small sized fruits. The fruits that develop black colour due to sooty mould lose quality and fetch low price. The yields are also reduced by aphids directly and more through the spread of virus diseases acting as vectors indirectly. Cruciferous Vegetables Nymphs and adults of B. brassicae and L. erysimi suck the cell sap from tender leaves/shoots. Affected plants show stunted growth and poor head formation. Under severe infestation the entire plant may dry up. When seedlings are infested they lose vigour, gets distorted, and become unfit for transplanting. Aphids excrete honeydew which attracts sooty mould and interferes with photosynthesis. If infestation starts early, heavy losses can occur. Leguminous Vegetables Adults and nymphs of A. craccivora pierce plant tissues to feed on plant sap. Their feeding causes rolling, twisting, or bending of leaves. Heavily attacked leaves turns yellow and eventually wilt. Aphids feeding on flower buds and fruits may cause malformed flowers and fruits. Aphids excrete honeydew that accumulates on leaves and branches. Sooty moulds growth on honeydew causes turning leaves and branches black. Heavy coating with honeydew and sooty moulds reduce photosynthesis, affecting plant growth and yield. Peas Both nymphs and adults of Acyrthosiphon pisum suck the sap from young plant parts, like shoot tips, flower, flower buds, and pods. Affected leaves get cupped or become irregularly distorted. Shoots become stunted and malformed. They also excrete large quantities of honeydew which encourages the growth of black sooty mould fungi and that affects photosynthesis. Plants become weak and the pod formation is adversely affected.

10.2.3 Jassids/Leaf Hoppers Taxonomic Position Phylum: Arthropoda. Class: Insecta (Insects).

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Order: Hemiptera (True Bugs, Hoppers, Aphids, and Whiteflies). Suborder: Auchenorrhyncha (Free-living Hemipterans). Superfamily: Cicadoidea (Cicadas, Leafhoppers, and Treehoppers). Family: Cicadellidae (Leafhoppers).

10.2.3.1 Okra Leaf Hopper Amrasca biguttula biguttula Ishida, Amrasca devastans, and Empoasca kerri Pruthi This is a more common, cosmopolitan, polyphagous, and highly destructive pest. It occurs in several countries including India, Bangladesh, China, Myanmar, North Africa, Pakistan, Philippines, Sri Lanka, and Taiwan. Relatively dry and humid weather condition favours population build-up. Diagnostic Features A. biguttula biguttula The eggs are pear shaped, elongated, and yellowish white in colour. The nymphs resemble the adults, but lack wings. Instead, they have slightly extended wing pads. They are pale green in colour. They tend to move sideways when disturbed. Adults are wedge shaped, 2–3 mm long, pale green in colour. They have fully developed wings with a prominent black spot on posterior portion of each forewing. Adults of winter generation are slightly reddish (Fig. 10.1f). Biology A. biguttula biguttula Adult females lay eggs singly along the midrib and lateral veins on the under surface of leaves. Incubation period is 4–10 days. The nymphal period varies from 1 to 4 weeks depending on the temperature. The adults may live for 1–2 months. Mating takes place 2–16 days after emergence. Oviposition begins 2–7 days after copulation. Each female lays 15–30 eggs. There are 10–12 overlapping generations in a year. There is no true hibernation or diapauses but the adults have the ability to tide over the adverse climatic conditions. The pest appears with the onset of cloudy weather and their population is adversely affected after heavy monsoon showers. Host Range and Disease Transmission A. biguttula biguttula Cotton, okra, eggplant, tomato, castor, cucurbits, hollyhock, potato, sunflower, and many other malvaceous plants. Transmit the phytoplasmal disease known as little leaf of brinjal. E. kerri Leguminous vegetables, tomato, brinjal, okra. Nature of Damage Okra Both nymphs and adults suck the sap from the leaves and also inject toxic saliva into plant tissues. The affected leaves become yellow, crinkled, curled and

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show marginal browning. In case of heavy infestation, the leaves turn dark brick red, become brittle, crumble and cause serious hopper burn/drying of leaves, resulting in stunted growth. Brinjal Both nymphs and adults are found between the veins of leaves on the undersurface and suck the sap from the lower leaf surfaces through their piercing and sucking mouthparts. While sucking the plant sap, they also inject toxic saliva into the plant tissues, which leads to yellowing. When several insects suck the sap from the same leaf, yellow spots appear on the leaves, followed by crinkling, curling, bronzing, and hopper burn or drying of leaves. The infested leaves curl upward along the margin, which may later turn yellowish, crinkle and show burnt up patches. Plants become stunted and may be killed in severe cases and fruit set is adversely affected by the infestation.

10.2.4 True Bugs Taxonomic Position Phylum: Arthropoda. Class: Insecta (Insects). Order: Hemiptera (True Bugs, Hoppers, Aphids, and Whiteflies). Suborder: Heteroptera (True Bugs).

10.2.4.1 Brinjal Lacewing Bug, Urentius sentis, U. hystricellus (Family: Tingidae) This pest is distributed in north western parts of India. The pest is active from April to October. Hibernates as an adult in cracks and crevices in the soil. Diagnostic Features Nymphs are about 2 mm, pale or black or brownish, stoutly built with prominent spines. Adults are about 3 mm, straw coloured dorsally and dark brown to black ventrally. Females are oval and males are elongated. Pronotum and elytra are reticulated. Coastal area is hyaline with strong spines on the outer margins (Fig. 10.1g). Hindwings are whitish and transparent. Biology Eggs are laid singly in the tissue on the underside of the leaves. Each female lays 35–44 eggs. The eggs hatch in 3–12 days. Nymphs pass through 5 instars. Young nymphs feed gregariously. The older nymphs feed individually. Nymphal period is 10–23 days. Adult longevity is 30–40 days. There are eight overlapping generations in a year. Host Range and Disease Transmission It is a monophagous pest, feeds mainly on brinjal crop.

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Nature of Damage Brinjal Both adults and nymphs are usually found on the ventral surface of leaves and cause damage by sucking the cell sap from brinjal leaves. Young nymphs feed gregariously on the lower surface of the leaves and also inject some toxic saliva into the plant tissues. Symptoms of damage are leaves with yellow patches, soiled with exuviae and excreta, presence of yellowish brown, flat, lace-like adults on the dorsal surface and spiny black nymphs on the under surface of leaves. Excreta impart mottled appearance to the infested leaves. Under severe infestation up to 50% of the crop may be destroyed.

10.2.4.2 Red Cotton Bug, Dysdercus koenigii, Dysdercus cingulatus (Fabricius) (Family: Pyrrhocoridae) The pest is distributed all over the Indian subcontinent. The insect is active throughout the year. Passes winter in the adult stage and the bugs become active during spring. Diagnostic Features The eggs are spherical, bright yellow in colour. Young nymphs have flabby slender abdomen and develop black markings on the body. Adults are reddish bugs with white bands on the abdomen and black markings on the wings (Fig. 10.1h). Biology The adult female bug lays on an average 100–300 eggs in moist soil or in cracks and crevices in the ground in clusters. Eggs hatch in 7–8 days to become nymphs. Nymphs have five stages and complete their development in 49–89 days. Adult longevity is variable and may live up to 3 months during winter. Host Range and Disease Transmission The pest is known to feed on cotton, okra, hollyhock, maize, sorghum, millets, musk melon, hemp, rose, and other malvaceous plants. Nature of Damage Okra Both nymphs and adults suck the sap from the tender shoots, fruits, and leaves of okra. Damage causes devitalization of the plants. Feeding deprives the plants of carbohydrates, free amino acids, and proteins, reducing both quality and quantity of fruits.

10.2.4.3 Dusky Cotton Bug, Oxycarenus laetus (Family: Lygaeidae) Diagnostic Features It is distributed all over the Indian subcontinent. The pest is active throughout the year, but during winter only adults are found. During spring cigar shaped eggs are

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laid on the leaves of okra or Hibiscus. Eggs initially are whitish but turn light pink before hatching. Biology Adult female bugs lay cigar shaped eggs on the leaves of okra or hibiscus during spring. Eggs initially are whitish but turn light pink before hatching. Eggs hatch in 5–10 days. There are seven nymphal stages and nymphal period is 30–40 days. Total life cycle is completed in 36–50 days. Host Range and Disease Transmission The pest prefers to feed on okra, hollyhock, cotton, and other malvaceous plants. Nature of Damage Okra Nymphs and adults are damaging. They suck the cell sap from leaves, fruits and reduce vitality of the plants.

10.2.4.4 Painted Bug, Bagrada hilaris (Burmeister), Eurydema pulchrum, Pumpkin Stink Bug, Coridius janus (Fabricius), C. brunneus and Green Stink Bug, Nezara viridula (Family: Pentatomidae) Painted bugs are important pests of cruciferous crops at flowering and pod formation stage. Appears during March–April and peak activity coincides with pod formation stage. In Haryana the pest remains active throughout the year with two peaks (October–November and March–April). The incidence is negatively correlated with RH and positively correlated with temperature. The pest is reported to be distributed in India, Sri Lanka, Pakistan, Afghanistan, East Africa, and South East Asia. The pumpkin and green stink bugs are distributed across India. Diagnostic Features B. hilaris The adult bug is 5–7 mm in length, shield-shaped, and black with white and orange markings (Fig. 10.1i). The female is larger than the male, lays up to 100 oval or barrel-shaped eggs. The eggs are white when freshly deposited and turn orange over time. Nymph is bright orange-red and turns darker as it develops, becoming black by the last instar. Coridius spp. Adults are flat, medium sized bugs. C. janus is about 30 mm long, pronotum and base of elytra are bright red while head and wings membrane are black. C. brunneus is pale brown in colour and slightly smaller in size (Fig. 10.1j). N. viridula The adult males can reach a body length of about 12.1 mm from front to elytral apex, while females are bigger, reaching a size of about 13.1 mm. The body is bright green and shield-shaped and the eyes are usually reddish, but they may also be black (Fig. 10.1k). Scent gland openings are short and wide. Several distinct morphs

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can be distinguished by the pattern of their exoskeleton colouration, which is predominantly green. Biology B. hilaris The eggs are laid singly or in batches of 2–12 on leaves, stems, and flower buds or in soil beneath plants. Each female can lay on an average 230 eggs at 15–20 eggs per day. Incubation period is 2–5 days. Nymphal period is 18–20 days. Mating takes place immediately after final nymphal molt. Pre-oviposition period is about 1 week. Adult longevity is 16–18 days. Total life cycle was completed in 21–27 days. There can be 6–8 generations in a year. Coridius spp. The eggs are deposited in long rows clinging to the leaves and tender shoots. Incubation period is 9–10 days. Nymphal duration lasts for 24–28 days. The bugs emit characteristic buggy smell, hence the common name is stink bug. N. viridula The female lays 30–130 eggs at a time, in the form of an egg mass glued firmly to the bottom of a leaf. The eggs are barrel-shaped, with an opening on the top. The eggs take between 5 and 21 days to develop, depending on the temperature. The newly emerged nymphs gather near the empty eggs and do not feed until 3 days. They moult five times before reaching maturity, increasing in size each time. Each nymphal instar lasts about a week, except for the last instar, which is a day longer. There can be four generations in a year. Total life cycle is completed in 35 days. Host Range and Disease Transmission B. hilaris The pest prefers to feed on all cruciferous vegetables. It has also been reported from Maize, Bajra, Black gram, Vigna mungo, etc. Coridius spp. It is known to feed on cucurbits especially pumpkins and gourds. Nezara viridula Cosmopolitan and polyphagous pest feeds on sorghum, rice, pigeon pea, cotton, okra, brinjal, castor, vegetables, and several other hosts. Nature of Damage B. hilaris Both nymphs and adults suck cell sap from the leaves and the developing pods which gradually wilt and dry up. On leaves pale or whitish markings appear and later on these leaves turn brown. In case of severe infestation, the seed formation is reduced. The nymphs and adults also excrete a sort of resinous material which spoils the pods. Coridius spp. Both nymphs and adults suck the cell sap from cucurbit plants and thereby devitalizing the plant and retarding their growth.

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10.2.4.5 Black Bean Bug, Chauliops nigrescens (Family: Lygaeidae) This pest is distributed in India, Sri Lanka, Japan, south-eastern Australia, etc. In India it is serious in northern states on many beans. Diagnostic Features The eggs are cylindrical, dark brown in colour. There are five nymphal instars. Newly emerged nymphs are light orange in colour. Fourth instar nymphs are dull red and fifth stage nymphs are dark brown colour. The average size of female and male is 2.75  1.00 and 2.85  1.25 mm, respectively. Biology The adult bugs appear during the onset of monsoon. The eggs are laid singly or sometimes in pairs glued to the leaf surface. Fecundity is 45 eggs/female. Incubation period is about 8 days. Nymphs mature through 5 instars. The total nymphal period is completed in 18–23 days. Pre-oviposition period is about 9 days, oviposition period is 15 days. Adult longevity is about 25 days in females and 21 days in males. There are three overlapping generations in a year. Host Range and Disease Transmission The pest is reported to feed on soybean, cowpea, moth bean, French bean, horse gram, and green gram. Nature of Damage Leguminous Vegetables Both nymphs and adults suck the cell sap. They cause reduction of chlorophyll content resulting in less photosynthetic activity. Quality and the yield both are greatly reduced. Severely damaged leaves show several minute yellow specks and small black pustules of excreta. Leaves gradually wither and fall of.

10.2.5 Mealy Bugs Taxonomic Position Phylum: Arthropoda. Class: Insecta (Insects). Order: Hemiptera (True Bugs, Hoppers, Aphids, and Whiteflies). Suborder: Sternorrhyncha (Plant-parasitic Hemipterans). Superfamily: Coccoidea (Scales and Mealybugs). Family: Pseudococcidae (Mealybugs).

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10.2.5.1 Solenopsis Mealybug Phenacoccus solenopsis Tinsley, P. madeirensis Green, Ferrisia virgata, and Brinjal Mealybug Coccidohystrix insolita (Green) (Family: Pseudococcidae) Solenopsis mealybug, P. solenopsis previously known as a pest of cotton but now possess a new threat to most of the cultivated crop plants. There are 84 plants across 50 families are recorded as hosts at Central cotton growing zone of India, out of that 60 plant species from 22 families belonged to weeds (Vennila et al. 2010). Presently, they feed on the host plants covering cereals, pulses, oil seeds, fruits, vegetables, ornamental crops as well as many weeds including Parthenium. Amongst vegetable crops, they found to attack on variety of plants belonging to malvaceae (ladies finger), solanaceae (tomato, brinjal, potato, chilly), leguminaceae (field bean), and cucurbitaceae (pointed gourd, cucumber, melons, and gourds) (Halder et al. 2013). Diagnostic Features P. solenopsis The adult female body is yellow, covered with a powdery, waxy secretion, dorsum with dark bars on the thorax and abdomen, with a pair of long terminal waxy filaments (Fig. 10.1l). Body with 18 pairs of cerarii, each with two spinose setae, located around the marginal area. Antennae 9-segmented. The first instar nymphs have a 6-segmented antennae and lack the circulus, and the third instar nymph has 7-segmented antennae and a circulus. C. insolita Adult females are light yellowish-green in colour, with many long, glassy filaments. These are small, oval, soft bodied insects measuring 3–4 mm in length, covered in white, mealy wax. Very little dorsal wax on adult female and secretes a white, waxy ovisac up to 6 times as long as the body of the female (Fig. 10.1m). The immature stages do not secrete a thick layer of mealy wax, the body being shiny yellow-green with submedian grey spots on 2 abdominal and 1 thoracic segments. Body with 17 pairs of ceraii, numerous dorsal cerarii present also, each cerarius consisting of 1–15 large conical setae situated on a sclerotized prominence, without any associated trilocular pores; legs well developed, each claw with a denticle present on plantar surface; circulus absent; anal lobes well developed, each with a sclerotized ventral bar. Posterior ostioles present, anterior ostioles absent. Biology P. solenopsis A female produces about 200–500 eggs in an ovisac. The emerging nymphs (crawlers) settle on the leaves, stems, and leaf petioles as well as around okra fruits. In dry regions the mealybugs are often found on the lower stems, foliage, and roots, whereas in more humid areas they settle on the upper plant parts. The pest excretes much honeydew and move between sites, thus spreading rapidly under favourable conditions. Adult females can live for up to 3 months and withstand starvation up to 12 days at 28  C and 50.6% RH. They can be dispersed by ants, visiting birds, and rodents.

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C. insolita They have reproductive potential of laying 200–300 eggs, majority of which are females, resulting in explosive outbreaks. Eggs incubate beneath their body cavity for about 4–5 days. There are three nymphal instars which last for 22–25 days. Their total life span under normal conditions from egg to adult under is 26–30 days. Host Range and Disease Transmission P. solenopsis Highly polyphagous with hosts in over 50 families and common hosts include cotton, okra, tomato, brinjal, okra, potato, pomegranate, hibiscus, etc. P. madeirensis Highly polyphagous and found on numerous host plants. Some of the common hosts observed include cotton, hibiscus, tomato, potato, brinjal, acalypha, etc. C. insolita It is a common and often serious pest of brinjal. This pest is polyphagous and reported to be feed on Pigeon pea, brinjal, tomato, capsicum, croton, stored potato and other Solanum spp. C. insolita is recorded from the following families of host: Acanthaceae, Amaranthaceae, Apocynaceae, Araceae, Arecaceae, Aristolochiaceae, Asteraceae, Chenopodiaceae, Cucurbitaceae, Euphorbiaceae, Fabaceae, Malvaceae, Menispermaceae, Moraceae, Poaceae, Rhamnaceae, Rubiaceae, Solanaceae, Sterculiaceae, Tiliaceae, Zygophyllaceae. Nature of Damage Both nymphs and adults suck sap from leaves and tender shoots. Heavy clustering of mealybugs is usually seen under the surface of leaves as a thick mat, with a waxy secretion. The extraction of sap by the mealybug results in the leaves of the plant turning yellow and becoming crinkled or malformed, which leads to loss of plant vigour, foliage and fruit-drop, and potential death of the plant, if not treated. They also excrete copious amounts of honeydew on which the sooty mould fungus grows. Affected plants appear sick and black, resulting in reduced fruiting capacity. Phloem feeding by P. solenopsis mealybug affects the growing regions of the plant often resulting in bunched and stunted growth with plants producing smaller fruit or flowers, which ultimately lead to a reduction in seed or fruit yields. Old plants are usually affected; leaves and tender shoots are covered by a large number of mealybugs, attended by small, brown ants.

10.2.6 Thrips Taxonomic Position Phylum: Arthropoda. Class: Insecta (Insects). Order: Thysanoptera (Thrips).

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Thrips are minute and slender insects with fringed wings and unique asymmetrical mouthparts. They appear in nursery as well as in main field and affect the crop throughout its life period. Pest infestation increases at high temperature. Many thrips species are pests of commercially important crops, feed on plants by puncturing and sucking up the cell contents. A few species serve as vectors for over 20 viruses that cause plant disease, especially the Tospoviruses.

10.2.6.1 Cotton Thrips, Thrips tabaci Lindeman, Blossom Thrips, Frankliniella schultzei, F. occidentalis, Chilli Thrips, Scirtothrips dorsalis Hood and Groundnut Thrips, Caliothrips indicus All the species of thrips are widely distributed throughout India. The pest is active throughout the year and breeds on different hosts during different seasons. Diagnostic Features T. tabaci Eggs are tiny, kidney shaped, and white in colour. Nymphs and adults are slender, fragile, and yellowish in colour. Adults have fringed wings heavily with fine hairs. Males are 0.8–1.0 mm long while the females are 1.0–1.2 mm long. S. dorsalis Eggs are minute and dirty white. Nymphs and adults are small, slender, fragile, and yellowish-straw in colour. Fringed wings are uniformly grey in colour. F. schultzei The adult common blossom thrips is a very small insect with a length of between 1 and 1.6 mm. There are two colour morphs, a dark form and a pale form, both colour morphs are being reported from Egypt, India, Kenya, New Guinea, Puerto Rico, Sudan, and Uganda. Identification of thrips species is dependent on the colour, the number and arrangement of the bristles on the body, and the details of the comb on the eighth abdominal segment (Fig. 10.1n). Biology T. tabaci Eggs are laid singly in slits made in leaf tissue. Eggs hatch in 4–9 days. Nymphs pass through four instars. Nymphs become full grown in 4–6 days. Full grown nymphs fall onto the ground and pupate in soil at a depth of about 2.5 cm. Pre-pupal and pupal periods are 1–2 and 2–4 days. Adult longevity is 2–-4 weeks. They undergo several overlapping generation in a year. F. schultzei Parthenogenesis is the main mode of reproduction, though sexual reproduction also occurs. A female lays on an average 50 eggs and the life cycle is completed in about 13–33 days. Females live longer than males. There are several overlapping generations in a year. S. dorsalis Reproduction is both sexual and parthenogenetic. Eggs are laid on just under the leaf tissues. Oviposition period is about 1 month. Each female lays about

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100 eggs. Total life cycle is completed in 14–18 days. There exists many overlapping generations in a year. C. indicus The pest is active at flowering. On garden pea the pest has been recorded from germination stage till harvest. Life cycle is completed in 2–4 weeks. Several overlapping generations are found in a year. Host Range and Disease Transmission T. tabaci It is a cosmopolitan and polyphagous pest known to feed on onion, garlic, cotton, cabbage, cauliflower, potato, tobacco, tomato, cucumber, peas, pine apple, etc. F. schultzei Cosmopolitan, polyphagous pest, vector for tomato spotted wilt virus in tomato. S. dorsalis Found in almost all chilly growing areas. It is a polyphagous pest. Besides chilli, it also infests brinjal, cotton, groundnut, castor, bottle gourd, guava, tea, and grapevine. It is more common on un-irrigated chilli crop than irrigated one. Known to transmit chilli leaf curl disease. C. indicus Reported to feed on peas, cowpea, Indian bean, etc. Nature of Damage Onion and Garlic On onion and garlic they are usually congregated at the base of the leaf or in the flower. Adults and nymphs of T. tabaci lacerate the epidermis of the leaf and suck the exuding sap. The affected leaves show silvery white blotches which later become brownish and cause retardation of growth. Bulbs remain undersized and get distorted. It also transmit viruses. Tomato Both adults and larvae of the common blossom thrips feed on flowers and pollen. They also lacerate the leaf tissue and imbibing the oozing sap. Severely infested flowers wilt, fade, and drop prematurely without bearing fruits. Pale and silvery sheens appear on the affected leaves. Secondary damage is caused by the viruses that can be transmitted between plants. Chilli Nymphs and adults of S. dorsalis lacerate the host tissue and imbibe on the oozing sap. Tender leaves and growing shoots are more preferred. Sometime buds and flowers are also attacked. The infested leaves curl upward, crumble, and shed; infested buds become brittle and drop down. Affected fruits show light brown scars. Incidence is more in dry weather condition. In severe condition, 30–50% crop may be destroyed. In mixed cropping of onion and chillies, both crops suffer badly. Early stage infestation leads to stunted growth and flower production, fruit set are arrested.

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Leguminous Vegetables Damage is caused by both nymphs and adults of C. indicus by feeding on cell sap. They lacerate the leaf surface and suck the oozing sap. White patches develop on the infested leaves. The pest is active at flowering and both the yield and viability of the seeds are reduced. A severe infestation results in the formation of white silvery sheens all over the leaf surface.

10.2.7 Mites Taxonomic Position Kingdom: Animalia. Phylum: Arthropoda. Class: Arachnida. Subclass: Acari (Mites and Ticks). The predominant phytophagous mite species associated with vegetables are Tetranychus neocaledonicus Andre, T. urticae Koch., T. cinnabarinus (Boisduval) (Tetranychidae: Acarina), and Polyphagotarsonemus latus (Banks) (Tarsonemidae: Acarina). The activity of these tetranychid mites has been observed during the postmonsoon and dry period. The activity declines with the drop of temperature (Puri and Mote 2004).

10.2.7.1 Red Mites Tetranychus urticae Koch, T. cinnabarinus (Boisduval) and T. neocaledonicus Andre (Family: Tetranychidae) Diagnostic Features T. urticae Commonly known as red spider mite or two spotted spider mite. They are minute in size and vary in colour (green, greenish-yellow, brown, or orange-red) with two dark spots on the body. Eggs are round, white, or cream-coloured. Nymphs yellowish and feed on lower leaf surface. Adult red coloured small size, form silken webbing on underside of leaves and on young shoots (Fig. 10.1o). Biology T. urticae Eggs are laid in mass mostly along the midrib and side margins on the lower surface of the leaves. Eggs hatch in about 2–4 days. Upon hatching, it will pass through a larval stage and two nymphal stages (protonymph and deutonymph) before becoming adult. Newly emerged larva becomes protonymphs in about 2 days. Protonymph stage is 2–3 days and deutonymph stage lasts for 1–3 days. Male longevity is 9–13 days and females live for 14–20 days. The adult lives up to 3–4 weeks. The life cycle is completed in 1–2 weeks. There are several overlapping generations in a year. Weather factors play an important role in the life cycle of spider mites. Under dry and hot conditions the multiplication of these mites is very

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high and the infestation is also severe. High humidity and temperature reduces the reproduction of the pest and hence the incidence is low. Host Range and Disease Transmission T. urticae is highly polyphagous pest known to feed on more than 100 species of plants including ladies finger, brinjal, beans, cow pea, and cucurbits. It is one of the most common and destructive species of mite and is cosmopolitan in distribution. T. cinnabarinus mainly feeds on cucurbits like pointed gourd, cucumber, melons, gourds but is also found on a number of other crops including brinjal, beans, onion, peas, cole crops, tomato, sweet potato, and various ornamental crops. Nature of Damage Brinjal Spider mites usually extract the cell contents from the leaves using their long, needle-like mouthparts. This results in reduced chlorophyll content in the leaves, leading to the formation of white or yellow speckles on the leaves. In severe infestations, leaves will completely desiccate and drop off. The mites also produce webbing on the leaf surfaces in severe conditions. Under high population densities, the mites form ball-like mass using strands of silk, which will be blown by winds to new leaves or plants, in a process known as ballooning. Tomato Damage is caused by the larvae, nymphs, and adults by sucking the cell sap from under side of leaves, flower buds, and flowers. When population is high, it results in bronzing and curling of leaves and discolouration of flowers and leaves. Webbing of leaves, sepals, and petals occur which gives untidy look to the plant. The infestation is more severe under polyhouse conditions. Okra Affected leaves become reddish brown and bronzy. Under severe infestation, larvae form silken webbing on the leaves. Cause withering and drying of leaves. Flower and fruit formation get affected. Leguminous Vegetables The puncture-and-suck mode of feeding of spider mites produces yellow or white stippling and leads to reddish or pale discolouration of leaves. Plants lose vigour and become unthrifty. Heavy mite infestation causes stunting of plants and premature leaf drop may occur. When mite populations are high, there is visible webbing on leaves.

10.2.7.2 Yellow Mites: Polyphagotarsonemus latus (Banks) P. latus is known as broad mite or yellow mites. In recent year’s major bottle neck of chilli growers in Varanasi region, Uttar Pradesh is mainly due to heavy infestation of this mite (Singh 2004). Nicotina and Cioffi (2000) as reported the dangerous diffusion of this broad mite on vegetables and floral crops especially in greenhouses. At cellular level, the typical symptoms of mite can be recognized by thickening the lower epidermal cells, elongation of the palisade cell layer, shrinkage and deformation of parenchymatous tissues (Rai et al. 2007).

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Diagnostic Features Female mites are about 0.2 mm long and oval in outline. Their bodies are swollen in profile and a light yellow to amber or greenish with an indistinct, light, median stripe that forks near the back end of the body. Males are similar in colour but lack the stripe. The two hindlegs of the adult females are reduced to whip-like appendages. The eggs are colourless, translucent, and elliptical in shape. Young broad mites (larvae) have only three pairs of legs. They are slow moving and appear whitish due to minute ridges on the body. After one day, the larva becomes a quiescent nymph that is clear and pointed at both ends. The nymphal stage lasts about a day. Adults are large, oval, broad, and yellowish in colour. Biology The broad mite has four stages in its life cycle: egg, larva, nymph, and adult. Adult females lay 30–76 eggs (averaging five per day) on the undersides of leaves and in the depressions of small fruit over an eight- to 13-day period. Adult male lives for 5–9 days. While unmated females lay eggs that become males, mated females usually lay four female eggs for every male egg. The eggs hatch in 2 or 3 days and the larvae emerge from the egg to feed. After 2 or 3 days, the larvae develop into a quiescent larval (nymph) stage. Quiescent female larvae become attractive to the males which pick them up and carry them to the new foliage. Males and females are very active, but the males apparently account for much of the dispersal of a broad mite population. Host Range and Disease Transmission Broad mite or yellow mite has also been identified as serious pest on chilli and beans. This mite infestation causes Murda disease in chilli. Nature of Damage Chilli/Capsicum Both nymphs and adults suck the sap and devitalize the plant causing ‘Murda’ disease of chillies. The affected leaves becoming inverted boat shaped. Infestation results in downward curling of leaves along the margin with elongation of petioles. Affected leaves turning dark green in certain cases. It also causes clustering of young leaves at the tip of branch. Due to infestation of this mite, the underside of the leaves turns reddish and plants become rosette.

10.3

Integrated Management of Sucking Pests of Vegetable Crops

Insect pests are major constraints in achieving full yield potential in vegetables. Indiscriminate use of chemical pesticides has resulted in emergence of more aggressive pests due to resistance development, residual problems in food and drinking water, and ecological imbalance due to elimination of beneficial insects, and finally outbreak of secondary pests like sucking insects. Therefore, for sustainability of

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vegetable crops, these insect pests are need to be managed through eco-friendly measures supported by need based and judicious use of chemicals without disturbing environmental balance and serenity. Integrated pest management (IPM) is one of the economically viable and environmentally safe key technologies to increase vegetable productivity in the country. Some of the important components of IPM practices in vegetables are given below. 1. 2. 3. 4. 5. 6.

Use of resistant varieties/hybrids/genotypes (host plant resistance). Use of healthy seeds obtained from a reliable source. Crop rotations, intercropping, trap/barrier crops. Optimum dates for sowing, planting, and harvesting. Use of biocontrol agents, biofumigants, and botanicals. Need based application of safer and label claim chemical pesticides.

10.3.1 Host Plant Resistance Sucking pests can be combated to a greater extent with the adaption of resistant varieties/genotypes. Keeping in mind the diversity and intensity of pests in particular place, selection of resistant/less susceptible varieties/hybrids/genotypes holds well in pest management. Unlike cereals, in vegetables very less number of resistant/less susceptible genotypes have been developed for insect pest management (Table 10.4). Being completely safe, host plant resistance fits well with all other components. Use of tolerant/resistant cultivars is an important component in the integrated pest management strategy. Some of the varieties developed for management of sucking pests in different vegetable crops are listed in Table 10.4.

Table 10.4 List of resistant varieties identified against the sucking pests of vegetable crops Crop Brinjal

Cabbage

Pest species Leafhopper (Amrasca biguttula biguttula) Whitefly (Bemisia tabaci) Yellow mite or broad mite (Polyphagotarsonemus latus) Aphid (Brevicoryne brassicae)

Okra

Jassid (Amrasca biguttula biguttula)

Onion

Whitefly (Bemisia tabaci) Thrips (Thrips tabaci)

Chilli

Resistant genotypes Chaklasi Doli, Doli 5, Pusa purple Pusa purple Japani Longi, GKC-29, Kashi Gaurav, VR-339 All season, red drum head, sure head, express mail IC-7194, IC-13999, new selection, Punjab Padmini Arka Anamica, Hisar Unnat, Varsha Uphar PBR-2, PBR-6, Arka Niketan, Pusa Ratnar, PBR-4, PBR-5, PBR-6

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10.3.2 Cultural Control Cultural practices followed for crop cultivation strongly influence build-up of insect population. Pests incidence can be reduced by raising the crop when the most sensitive stage is least invaded by insects. In that context, the following general aspects need to be followed for the sucking pest management. 1. Nursery should be maintained weed free, away from the cropping field and monitored periodically for insect population to ensure insect free seedling production. 2. Seed treatment with imidacloprid 70% WS @ 5 g/kg for management of sucking pests of vegetable crops, such as chilli, cabbage/cauliflower, etc. 3. Before transplanting, roots of seedlings should be dipped for 15 min in imidacloprid 17.8% SL @ 7.0 mL/L to protect against sucking insects including whitefly in tomato. 4. Destruction of weeds before flowering is important as the insects are attracted towards the pollen for feeding. Also weeds should be cleared from cropping area. 5. All crop residues in previously infested fields should be removed and burnt, as they harbor mealybug populations which may invade the new crop. 6. Sowing of crops in such a way that early stage (sensitive stage) of crop is least invaded by insects. 7. Intercropping vegetable crops with two rows of border/barrier crops, such as pearl millet, maize, and sorghum, significantly reduces the incidence of sucking insects like whiteflies, on okra and thrips on onion by limiting their movement, which ultimately delay the initial incidence up to 10–15 days. 8. Growing of Indian mustard as trap crop along with cabbage has been successfully utilized for the management of aphids (Brevicoryne brassicae and Lipaphis erysimi) on cabbage. Later mustard is sprayed with systemic insecticides to manage aphids. 9. Growing of okra as trap crop along with brinjal for the management of leaf hoppers. 10. Crop rotation should be followed for the management of sucking pests including mealybugs and brinjal lacebug. 11. Use of soap oil or fish oil resin soap twice at an interval of 15–20 days for the control of mealybugs. Mites Grow nurseries away from infested crops and avoid planting next to infested fields. 1. Grow healthy crops, avoid water and nutrient stress. 2. Apply mulch and incorporate organic matter into the soil to improve the water holding capacity and to reduce evaporation. 3. Keep perennial hedges, such as pigeon peas, as they encourage population of predatory mites.

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4. Uproot and burn infested plants. This can be successfully done during the early stages of infestation when the mites concentrate on a few plants. 5. Keep the field weed free. 6. Remove and burn infested crop residues immediately after harvest.

10.3.3 Physical Control 1. Use of insect proof net with a pore size of 2%). They may affect fruit finish on some varieties (Anonymous 2003a, b). The use of neem based bio-insecticides (Neem Azal-T/S) before full blooming will reduce the aphid infestation without interfering the biological control. There are a number of predators that often control rosy apple aphid, so distorted leaves should be opened to determine if the aphids or predators are still present before making control decisions. The most common natural enemies of rosy apple aphid are Harmonia axyridis Pallas (Brown and Mathews 2007) and Adalia bipunctata L., the syrphid Episyrphus balteatus (DeGeer), the midge Aphidoletes aphidimyza (Rondani), and the braconid Ephedrus persicae Froggatt (Wyss et al. 1999). Their population could be increased either by weed strips (Wyss 1995; Dixon et al. 1994) or augmentative releases of indigenous natural enemies (Wyss et al. 1999).

12.2.3.4 Peach Leaf Curl Aphid Brachycaudus helichrysi (Kaltenbach) Leaf curl aphids are common problems for both trees and shrubs. These have soft pear-shaped bodies with long legs and antennae and may be green, yellow, brown, red, or black depending on the species and the plants they feed on. Life Cycle Female aphids lay eggs in protected crevices in trees and shrubs in late fall. The eggs hatch into nymphs in the spring soon after the host plants start to grow. These aphids Fig. 12.13 Damage of Peach leaf curl aphid (Brachycaudus helichrysi)

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Fig. 12.14 Peach leaf curl aphid (Brachycaudus helichrysi)

mature in 10–14 days. Adults live for 2–3 week, and then eggs are produced parthenogenetically which hatch inside the body of mother. Each viviparous female produces about 50 nymphs in 13 days of life span. Entire life cycle takes 22–25 days. After completing 3–4 asexual generations, the aphid migrates to its alternate host to pass summer, where it completes 4–5 generations. The last generation of aphids contain males and females that mate before the females lay eggs. Damage and Symptoms Damage is severe because the aphid colonises young shoots, buds, grafts, and young plants (Fig. 12.13). Nymphs and adults suck the sap from leaves, shoots, and fruits (Fig. 12.14). The sucking of sap causes the leaf to “curl” around the insect. Leaf curling affects the plant ability to collect sunlight for energy production and robs the plant of valuable nutrients. The aphids also produce a large amount of honeydew that will allow the growth of sooty mould that will affect the appearance of the affected part. In addition, B. helichrysi can transmit the Plum pox virus (PPV), which causes sharka, a debilitating disease of peaches, and potato viruses. Management Where aphid populations are localised on a few curled leaves or new shoots, the best control may be to prune out these areas and dispose of them. In large trees, some aphids thrive in the dense inner canopy; pruning out these areas can make the habitat less suitable. In early stages, aphid populations should be washed with a sharp spray of water from a hose pipe. Many predators feed on aphids. The most well-known are lady beetle adults and larvae, lacewing larvae, soldier beetles, and syrphid fly larvae. Substantial numbers of any of these bioagents may reduce the aphid population rapidly. When aphids cannot be controlled by these natural methods, insecticide like oxydemeton–methyl 25% EC @ 13320 g/acre, Carbofuran 3% CG @ 600–800 mL/600–800 L water/acre may be applied.

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Ants are often associated with aphid populations for obtaining honeydew. To protect their food source, ants ward off many predators and parasites of aphids. Managing ants is a key component of aphid management.

12.2.3.5 Green Peach Aphid Myzus persicae The Green Peach Aphid (Myzus persicae) is a species of aphid so-named due to the fact that it is a significant aphid pest of peaches in many parts of the USA, but is also found worldwide. Green peach aphid feeds on hundreds of host plants in over 40 plant families. In temperate regions the primary or overwintering hosts are trees of the genus Prunus, particularly peach and peach hybrids, but also apricot and plum. During the summer months the aphids abandon their woody hosts for secondary or herbaceous hosts, including vegetable crops in the families Solanaceae, Chenopodiaceae, Compositae, Brassicaceae, and Cucurbitaceae. Thus, it is sometimes known as the peach-potato aphid, reflecting two of its most common hosts Vegetables that are reported to support green peach aphid include artichoke, asparagus, bean, beets, broccoli, Brussels sprouts, cabbage, carrot, cauliflower, cantaloupe, celery, corn, cucumber, fennel, kale, kohlrabi, turnip, eggplant, lettuce, mustard, okra, parsley, parsnip, pea, pepper, potato, radish, spinach, squash, tomato, turnip, watercress, and water melon. Numerous flower crops and other ornamental plants are suitable for green peach aphid development. Stone fruit crops, such as peach, are sometimes damaged before the aphids leave for summer hosts (Heathcote 1962). Life Cycle The green peach aphid overwinters as an egg stage on the bark of fruit trees, specifically peach, cherry, apricot, and plum. Eggs hatch and young nymphs develop into stem mothers, which produce living young nymphs. The green peach aphid has both a sexual and asexual form. The majority of reproduction occurs asexually by a process called parthenogenesis where live young ones are produced. Development occurs very quickly, growing from neonate to adult in as few as 5 days. After several generations of wingless adults on fruit trees, winged aphids appear during June, and all aphids leave fruit trees during June and July, which disperse to other hosts including many vegetable crops. Generations developing on vegetable crops will have both winged and wingless adults and reproduce asexually. In late August winged forms migrate back to fruit trees. Near the end of the growing season on fruit trees sexual forms of the green peach aphid appear for the first time. After mating the female green peach aphid oviposits overwintering eggs on the bark of fruit trees. In total the green peach aphid may have 10–15 generations in a growing season. Damage and Symptoms Adults and nymphs suck the sap from leaf undersides, causing curling, shrivelling of leaves, and yellowing of foliage. Flowers and fruits may also be fed upon, resulting in distortion and discolouration. When abundant, aphid feeding results in excretion of large amounts of honeydew which supports the growth of a black sooty fungus that though not directly harming the plants, may block out sufficient light to reduce

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yield besides causes spotting of leaves and fruit. This species transmits over 100 plant viruses, with both persistent and non-persistent transmission. Both adults and nymphs can transmit viruses, but winged adults are of greatest importance because of their mobility. This aphid may also serve as a vector of virus diseases to stone fruits. Green peach aphid is one of the several aphids that can transmit plum pox virus. Plum pox disease is one of the most devastating diseases of stone fruit in Europe. Management Biological and cultural controls can be useful for limiting damage from this aphid. For instance, removing old crop debris from the field chopping or discing of crop debris immediately after harvest and destruction of alternate host plants reduce sources of virus and thereby its transmission by aphids, and using reflective mulches early in the season repel aphids from young plants. Heavy infestations on seedling and young plants may require treatment with insecticides. The green peach aphid is attacked by a number of common predators, including lacewings, ladybird beetles, syrphid flies, and parasitoids, including the parasitic wasps Lysiphlebustes taceipes Aphidius matricariae Aphelinus semiflavus, and Diaeretiella rapae; their population should be conserved and encouraged. When pesticides are applied care should be taken to select pesticides that are not damaging to natural enemies of aphids. Because aphid’s reproductive rate is greater than their natural enemies, incorrect pesticide use may contribute to aphid outbreaks by removing the natural enemies. Trees should be inspected weekly from petal fall until the terminals harden off. For nectarines the treatment threshold is one colony per tree, and for peaches the threshold is five or more colonies per tree. If aphid population are detected consider an insecticide application, besides, a dormant oil application suppresses the overwintering egg hatch. Motile forms can be treated with an insecticide as they appear in the spring.

12.2.3.6 Plum Aphid Hyalopterus pruni (Geoffroy) The adult is pale bluish green and has a white, powdery coating that makes it look grey to light green. Plum, almond, apricot, peach, and other Rosaceae plants are some of the important hosts plants of this pest. It is cosmopolitan in distribution. Life Cycle Mealy Plum Aphids overwinter as eggs on the primary host: plum, hawthorn, and sometimes on apricot and peach. They hatch in March/April and form dense colonies feeding on the underside of leaves. The resultant females reproduce by viviparous parthenogenesis until early summer. After two or three generations, winged aphids appear in summer and migrate to secondary host (reeds), where they produce several generations, but many remain on plum. In autumn winged males and females appear that fly to almond, apricot or peach trees whereon they mate and lay overwintering eggs on the primary host. These are laid near the base of flower buds and hatch when conditions are favourable during spring.

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Damage Vegetative growth on the trees may be stunted by high aphid numbers, but the principal damage caused by mealy plum aphid is the development of the black sooty mould that grows on the aphid’s honeydew. When the aphid occurs in large numbers on young leaves it causes significant feeding damage, but leaf curling only seems to occur when very young leaves are attacked. Infested trees are also contaminated with abundant honeydew and subsequent sooty mould. It reduces the plants ability to carry out photosynthesis and thus reduces productivity and yield. Management It is important to monitor for these aphids shortly before bud break, as management decisions must be made at that time. The aphids are best controlled when small and exposed. Several insect pest predators can keep the aphid populations below economic damage. These include ladybird beetles, lacewing larvae, syrphid fly larvae, soldier beetles, and predacious midges; their population should be encouraged and conserved. The use of delay dormant spray oil is the most effective way to control aphids. After the aphids become active and leave begin to curl they are more protected and harder to control. Attempts at late season control can disrupt predators.

12.2.3.7 Black Bark Aphid Pterochloroides persicae (Cholodkovsky) The aphid is commonly known as Black bark aphid, black peach aphid, cloudy winged peach aphid, and brown peach aphid. It is geographically distributed in Middle East, Eastern and Southern Europe, North Africa, and India. The aphid infests almond, peach, plum, and apricot. Peach appears to be its preferred host. Pterochloroides persicae is a single species recorded under genus Pterochloroides on Prunus persica (Peach) in fruit ecosystem of Kashmir (Khan and Shah 2017). Life Cycle The life cycle of the species is complex, having alteration of parthenogenetic and sexual generations, apterous and alate forms exhibited and persistently overlapping. The pest lives and feeds on the bark and branches of the host trees, whereas in warmer regions, it reproduces parthenogenetically the year around. Environmental conditions predominately influence the population dynamics of the species. In cooler areas males and oviparous females appear in autumn, and winter eggs are laid during winter. The eggs are laid on the stem and are light brownish in colour. The eggs begin to hatch from the first week of March. Newly emerged nymphs start feeding on small twigs. These grow and attain full maturity and start to produce young ones parthenogenetically from the second week of April. After producing several generations in a same manner up to autumn, males and oviparous females begins to appear, mate, and produce overwintering eggs. Probably six to eight generations of the species are prevalent in a season from temperate regions. At 27
C the pest can complete a generation in 2 weeks; it may rise about 20 annual generations. The species was seen to be active at temperatures as low as 3 while as most prolific at mean temperature of 20–22
C.

392 Fig. 12.15 Colony of Black bark aphid Pterochloroides persicae (Cholodkovsky) on Peach

Fig. 12.16 Colony of Black bark aphid Pterochloroides persicae (Cholodkovsky) damage on Peach trunk

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Damage and Symptoms It is a serious pest of peach, almonds, and apricots. It feeds on the phloem which results in general weakness of a tree, withered branches and reduced yield (Stoetzel 1994). Heavy, ongoing infestations by thousands of aphids kill trees (Figs. 12.15 and 12.16). The species is not listed as transmitting virus but high densities results in premature fruit drop, leaf curling an irregular curvature of twigs and stunted growth. In addition, the large amounts of excreted honeydew that is colonised by sooty mould add to the damage. Management Due to the aphids’ exposed position on bark and branches, their control is relatively straightforward, and can be achieved with pesticides (like imidacloprid) or certain plant extracts, provided they do not affect pollinating bees or the natural enemies. Predators recorded to prey on P. persicae were Harmonia eucharis (Mu.), Coccinella septempunctata L, Hippodamia variegate (Goe.), and Adalia tetraspilota (Hope), Chrysoperla Sillemi E. & P, spiders and syrphid flies (Khan et al. 2009; Khan and Shah 2017). These predators can be utilised for the management. The larvae of Cecidomyiidae, Chamaemyiidae, and Syrphidae prey on P. persicae and greatly reduce its numbers (Khan and Shah 2017). Several entomopathogenic fungi infect the aphid in Tunis and Israel, and may at times decrease its populations.

12.2.3.8 Walnut Aphids Walnut green aphid: Chromaphis juglandicola (Kaltenbach). Dusky-Veined Aphid: Callaphis juglandis (Goetze). Walnut aphids are destructive pests that attack walnut in many countries. The seasonal developments of both the species are very similar, but their appearance and behaviour are quite different. Walnut green aphid nymphs are yellow and the adults are yellow-green with a pair of dark spots on each abdominal segment. A yellowwhite population has been reported from India and California. They are much smaller than the dusky-veined aphid (Khan and Kundoo 2018). Nymphs of the dusky-veined aphid have dark, banded spots on the back. These spots are much less pronounced or absent on the nymphs of the walnut aphid. Life Cycle The life cycle of both these aphid species is basically the same. Chromaphis juglandicola is a holocyclic monoecious aphid that spends all of its life upon walnut trees. Both spend the winter conditions as hibernating eggs, which hatch in the spring. Eggs hatch as soon as leaf buds of early cultivars begin to open. Soon stem mothers or fundatrices start new colonies. Stem mothers are vivaparous and parthenogenetic, that is, they give birth to live young without need to be fertilised. These aphids settle on the leaflets. Depending upon temperature, the population densities may swell to several hundred individuals per leaf. The aphids pass through many generations in a year. However, when the days get shorter and the weather cools down, they start producing two new forms. Winged males and wingless females

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Fig. 12.17 Dusky-Veined Aphid: Callaphis juglandis

Fig. 12.18 Walnut aphids Chromaphis juglandicola (Kaltenbach)

appear again in the fall, typically in September. The population densities may swell to several hundred individuals per leaf its depend upon favourable temperature. These mate and deposit their overwintering eggs in the basal cracks of host-tree bark and buds. Damage The colonies of walnut aphids suck the sap confining themselves to ventral surface of the leaves and tender shoots (Khan 2015; Shah 2015). Dusky-veined soft bodied aphids (Callaphis julandis) prefer to feed on the upper surface of leaves along the midribs and suck the sap and devitalise them (Fig. 12.17). Walnut aphids Chromaphis juglandicola (Kaltenbach) are much smaller than the dusky-veined aphid and can be further distinguished by their yellow colour (Fig. 12.18). In each case, plant growth becomes stunted. The affected part become dry, pale, curl, and disfigured. In case of severe infestation, even the growth of the tree is affected and

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plant is stunted. Aphid populations of over 15 individuals per leaf laterally in the season can reduce nut yield and quality and cause an increase in nuts with perforated shells. An infestation in summer lowers the nut quality (Khan and Kundoo 2018). Walnut aphid infestation results in reduced tree vigour, nut size, quality, and yield. The aphids also secrete honeydew, on which sooty mould develops causing hindrance in photosynthetic activity of the plant and further retarding its growth. Sooty moulds grow on the nutrient rich honeydew turning the husk surface black, and increasing the chance of sunburn on exposed nuts (Faostat 2011). Management 1. Management decisions should be taken on the basis of monitoring of aphid population starting from May onwards by taking five first sub-terminal leaflets (each compound leaf has five leaflets) from ten trees for a total sample of 50 leaflets from the upper surface of each leaflet for dusky-veined aphids and the lower surface for walnut aphids. 2. Chemicals should be applied in such a way that should not interfere with the biological control of aphids. 3. The use of oil during the growing season has also been shown to be destructive against Trioxys. Predators, such as ladybird beetles (Kundoo et al. 2018a, b; Khan 2010b), lacewings (Khan and Zaki 2008), spiders (Khan 2011), and syrphid flies, play an important role in the natural control of the aphids (Khan 2017; Khan and Riyaz 2017). Reliance on biological control is the main management method in an organically certified crop. Pesticides which are effective for the management of both aphid species are as follows: 4. Horticultural mineral oil at 2% or diesel oil + fish oil soap at 6.33% (potash based) in the ratio of 1:7 (stock solution/water) plus ethion 50 EC at 100 mL/100 L of water is effective for the management of both aphids. 5. Dimethoate 30 EC (100 mL) or methyl-o-demeton 25 EC or phosolone 35 EC (140 mL) or chlorpyriphos 20 EC at 100 mL/100 L of water is effective for the management of aphids or Spray 0.02% chlorpyriphos 20EC or 0.07% endosulfan 35 EC during late May or early June. If infestation is heavy, repeat the spray after 3 weeks. 6. The parasitoids Trioxys pallidus and Diaeretiella rapae are major enemies of the pest. The former was introduced from Iran into California and provided good control. Predatory Coccinellidae along with Syrphidae are at times key factors that regulate the pest’s numbers, their population should be encouraged and conserved (Khan et al. 2016). The parasitic wasp, Trioxy pallidus, has reduced the need for chemical control of walnut aphid. Only a small percentage of orchards require treatment for walnut aphid, except when the parasite is disrupted by treatments for other pests.

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12.2.4 Psyllids 12.2.4.1 Pear Psylla, Psylla pyri (Linnaeus) Psylla pyri is commonly known as pear psylla or pear psyllid. It completes its development only on pears. It is native to Western Europe, found in Asia, North America, etc. The colour is variable ranging between orange-red and black, the wings are transparent with dark veins. The younger nymphs are yellowish with red-purple eyes. Large hard shelled nymphs are darker, with black areas interspersed with green or brown colouration. These forms have noticeable wing pads and are free living. Adults resemble very small cicadas and can be reddish brown (overwintered generation) or tan to light brown (summer forms) (Fig. 12.19). Life cycle The pear psylla overwinters as adult stage, which is somewhat larger and darker than the summer adult. The newly emerging overwintering females emerge in spring season and deposit their eggs on fruit spurs, but as the buds open, eggs are laid on exposed leaf or fruit tissues. Eggs are laid in lines or rows. These eggs hatch when foliage appears. The pear psylla passes through five nymphal stages. The fifth instar is referred to as the hardshell stage and is dark brown to black in colour and bears prominent wing pads (Figs. 12.20 and 12.21). This stage can move actively and is often found at the base of leaf petioles (Anonymous 1979). The period of development from egg to adult is 1one month. There are 2–3 summer generations before winter generations develop in the fall. The adults of the last generation leave the pear trees in October–November and hide under the bark, under litter on the orchard floor,

Fig. 12.19 Damage of Pear psylla, Psylla pyri (Linnaeus)

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Fig. 12.20 Nymphs of Pear psylla, Psylla pyri (Linnaeus) feeding on leaf

Fig. 12.21 Nymph of Pear psylla, Psylla pyri (Linnaeus)

or in sites outsides the orchard, and return about 6 weeks before bloom (Alston 2007). Damage and Symptoms Pear psylla is a very small sap feeding hemipteran insect. It is considered the most serious insect pest of pear in the USA. Nymphs and adults suck the phloem sap from leaves and succulent tissues (Fig. 12.19). That results in production of a large amount of honeydew, which makes the tree sticky and promotes the growth of

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Fig. 12.22 Harmonia eucharis (Mulsant) predator of Pear psylla, Psylla pyri (Linnaeus)

sooty mould. The honeydew can run onto the fruit that can cause fruit russetting, thereby reducing the marketability of the fruit. The pest also transmits a phytoplasma that causes “pear decline”. This disease prevents the movement of nutrients towards the root side, results in root starvation. Besides this other symptoms on this are stunted shoots, small or curled leaves, reduced fruit size, twig dieback, and premature leaf drop. Pear psylla also injects toxin saliva as it feeds, and under heavy infestation, psylla shock can occur. The symptoms of psylla shock include stunted growth, wilting, defoliation and fruit drop. Management Monitoring should be done during summer season in order to keep the pest populations below damaging levels. Insecticides are more effective if applied when most of the nymphs are in the first three instars rather than later instars because fourth instar nymphs and adults develop a hard shell. For best results dormant oil alone or mixed with other insecticides, reduces the pear psylla population and delays the egg laying. It will synchronise egg hatch with subsequent sprays. A balanced fertiliser program and avoidance of excessive nitrogen greatly reduces flush vegetative growth that attracts psylla. Summer pruning should be avoided which encourages shoot growth. Predacious insects like anthocorid bugs, predaceous plant bugs, ladybird beetles, Harmonia eucharis (Mulsant) predator of Pear psylla (Fig. 12.22) and spiders could be used for biocontrol programme. Two parasitoid wasps—Trechnites psyllae and Prionomitus mitratus—lay eggs inside the bodies of psylla nymphs where the wasp larvae consume the psylla host as they develop. Commercial production often requires applications of broad-spectrum insecticides, highly refined summer oil, kaolin clay or insecticidal soap.

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Fig. 12.23 Blossom thrips (Frankliniella occidentalis)

12.2.5 Thrips 12.2.5.1 Blossom Thrips (Frankliniella occidentalis) Thrips are small (1–2 mm), slender insects, just visible to the naked eye which are an economic pest of pome and stone fruit. Damage can occur at flowering (early season) and/or when the crop matures (late season) (Fig. 12.23). Thrips have varied diets reflecting their host preferences. Many species feed on the cell contents of the tissue of green plants (e.g., leaves, flowers, fruit, and young shoots) as well as pollen, nectar, and other exposed plant liquids, small arthropods and fungal hyphae and spores (Kirk 1984). The Thysanoptera are the only insect group that sucks the contents of pollen (Grinfel'd 1959; Kirk 1984). Thrips have unique asymmetrical mouthparts comprising a single mandibular stylet and paired maxillary stylets, protected and supported within a mouth cone projecting downwards from the ventral surface of the head (Chisholm and Lewis 1984). The method of feeding in the Thysanoptera is incompletely understood (Lewis 1973). Unlike most piercing/sucking insects, thrips do not feed from vascular tissues. Rather, thrips are known to pierce or rupture plant cells with their single mandible and drain or lap up cell contents with the maxillary stylets. Some species also “drink” nectar from flowers (Lewis 1973). Life Cycle The thrips overwinter as a sexually mature female in the soil, curved leaves, and in evergreen plants. These emerge in spring season, and locate the host and begin to feed and oviposit. The eggs are deposited within the plant tissues singly. The eggs hatch approximately within 4 days. The emerging nymphs feed on plant tissues. The nymphs have two nymphal stages, the second-instar nymph when matures, then either drop to the ground or to a protected crevice of the plant. The thrips then moult into quiescent, pseudo-pupal stage that lasts 4–5 days during summer and then pupates in a loosely constructed cell in soil, or in crevices on the host plant. After

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emergence the adults move to the growing parts of the plant, where they feed and lay eggs, about 200 eggs per female. Adults are usually found on young leaves, while larvae are found on lower or older leaves. The life cycle is completed in approximately 17 days. They produce many generations in a year. Heaviest damage occurs in spring. Damage and Symptoms Thrips can cause commercially significant damage to fruits. Thrips cause damage to plant trees by various ways: 1. It forms pansy spots (also called ghost spotting) on the apples, cherries, pears, and plum fruits that is caused by the physiological reaction to the presence of thrips egg in the tissue of young fruit. Oviposition in the newly pollinated apples destroys plant cells that are not regenerated, resulting in the typical pansy spot (Newcomer 1921). 2. Adults and nymphs suck the sap from leaves, flower parts, as the thrips have highly modified asymmetrical mouth parts that allow the thrips to puncture the epidermis of the plant and then suck out the contents of the ruptured cells, as adult insects feed on flower stamens and styles and can cause severe flower abortion. Often, if populations are low, this damage has a similar effect to flower thinning. However, heavy infestations can cause a large reduction in fruit set. 3. In plums, apples, and cherries, dimples form as a result of differential growth between injured and uninjured tissue surrounding the oviposition site. 4. In pears, egg punctures result in a slightly depressed, russeted spot. At harvest this damage is characterised by russetting, scarring, and pitting and/or scarred blotches. 5. Thrips can transmit various types of diseases to the host plant (Bailey 1935). Management Monitoring with yellow sticky traps is useful for determining the presence or absence of thrips within an orchard. Orchards in which thrips are detected can be sprayed, minimising damage. The population of natural enemies should be encouraged by habitat manipulation. Many weed species (broad-leaved weeds) and ground debris act as reservoirs for thrips; therefore their management is essential. Clover is particularly favoured than other broadleaved weeds, such as cape weed, dock, and sorrel. Flowering plants are particularly attractive to thrips, with clover (particularly white clover) and lucerne hosting the highest thrips densities. Thrips feeds on the pollen of these plants; therefore, preventing flowering can be an effective management strategy. Total removal of clovers and other broad-leaved weeds is one option, but if this is not possible, then keep them down short throughout the year to prevent flowering. Managing your ground cover and weeds is critical to reducing thrips populations and preventing population carry over. As thrips does not feed on grasses, replace broad-leaved ground cover with grasses. Be careful if you spray weeds with herbicide. As the weeds die off, thrips can move off the weeds onto crop. Weeds may be treated with insecticide at the same time.

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Both eggs and pupal stages are protected from sprays, and applying insecticides is likely to be futile unless the timing is correct. For this reason, the sprays should be recommended to cover the time taken for eggs to hatch into larvae and for pupae to develop into adults. During blossom period the insecticide should be applied in such a manner so as to reduce the risk of bee poisoning. Some of the precautionary measures are as: (1) remove all hives from the orchards prior to spraying and return them when the risk period is over, (2) close the bee hives during the risk period, and (3) Pesticides, like endosulfan @ 140 mL/ 100 L of water, should be used for thrip management. It should be applied only at times when bees are not actively working the blossoms, such as in the evening or early morning.

12.2.6 Mites 12.2.6.1 European Red Mite, Panonychus ulmi (Koch) European red mite (ERM) is also known as the fruit tree red spider mite. It was first observed in North America in 1911. It was reported as pest of apple in New South Wales and Australia in 1975 and by 1986 it had become a major pest in the entire apple-growing areas (Bower and Thwaite 1986). In India, it was reported for the first time from North-Western Himalayan region of Jammu and Kashmir, Himachal Pradesh, and Uttar Pradesh (now Uttaranchal) on apple, plum, peach, walnut, apricot, quince, jackfruit, Hibiscus, rose, and wheat (Prasad 1974). Life Cycle ERM overwinter as eggs on rough bark. The eggs are most commonly found near the buds, fruit spurs, on small limbs, twigs and in fork of two branches and leaves (Fig. 12.24). These are deposited during August and September and hatch the following spring between tight cluster and bloom. Summer eggs are deposited along the leaf veins predominantly on the underside of leaves and sometimes on Fig. 12.24 Eggs of European Red mite, Panonychus ulmi (Koch)

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Fig. 12.25 Nymphs of European Red mite, Panonychus ulmi (Koch)

Fig. 12.26 Adult of European Red mite, Panonychus ulmi (Koch)

fruit if populations are high. After hatching the mite passes through three stages, that is larva, protonymph, and deutonymph (Fig. 12.25). The larva is slightly larger than egg, orange in colour with three pairs of legs, feeds on underside of the leaves. While as other two stages are gradually larger, red, and have four pairs of legs. Sex is distinguished at the deutonymph stage as females grow larger and more oval than males. The female ERM is red or brownish-red with conspicuous white spots at the bases of their white bristles. The adult male is smaller than the female, has a tapered abdomen, and is reddish-yellow (Fig. 12.26). These first appear around petal fall. Adults feed mainly on the underside of leaves; moves to upper leaf and fruit surface when populations are high. Female starts laying eggs after 2 days and live up to 15–20 days. Each female on an average lays about 30–35 eggs during its entire life time. One generation can be completed in 10–25 days depending upon the temperature. Typically there are five to eight overlapping generations per year.

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Damage and Symptoms All active stages of the European red mite injure the foliage by feeding with piercing and sucking mouthparts and removing cell contents, including chlorophyll. Both sides of the foliage are colonised but the lower surface is preferred. In case of severe infestation, the leaves may turn on a bronze appearance. At high ERM populations (100 per leaf), premature leaf drop may occur, as early as late July or early August. Serious feeding injury can result in poor fruit colour and reduced size and quality in that year’s crop, as well as reducing the number of fruit buds and size of fruits for the next season’s crop. Additionally mite infested leaves do not respond to growth regulators applied to delay harvest crop. Management The most effective and easiest time to control the ERM on all types of fruit trees is during the delayed dormant period (half-inch green to tight cluster) before overwintering eggs. Eggs are more vulnerable to control before hatching. Good coverage with a superior oil spray prior to bloom is an important ERM management strategy that all growers should use. The oil application coats the eggs closer to hatch, preventing the exchange of gasses through the egg shell, suffocating the embryo within the egg shell. The better control occurs during egg stage because the embryos are respiring more rapidly as they prepare to hatch from the eggs. Superior oil applications also control some aphids and scales. Oil should be applied only when temperatures are above 45
F and never just before or after freezing weather. If a delayed dormant spray is missed, later sprays may be required to keep ERM populations below damaging levels. Highly refined oils can be applied during the growing season that controls ERM. These oils function in the same way as superior oils, suffocating the mites. Therefore good coverage is a must. These oils have no residual activity. Oil has a physical effect on mite eggs and avoids to resistance development. A well-applied oil spray can keep ERM at a low level until mid-summer. If they do not land on the mite, they will have no effect. Predatory mites have the ability to climb out of the oil on the leaves and they are not as adversely affected by oil applications. Predatory mites, ladybird beetles, and the six-spotted thrips help to maintain this European red mite at non-damaging levels. Minimising insecticide usage and selecting insecticides that are least toxic to beneficial organisms may help to minimise problems with this mite. Conserving natural enemies is one of the important strategies for the management of ERM. If the insecticides are applied wisely, outbreaks of ERM will be infrequent and can usually be cleared up fairly easily with the miticides. However, if most of the predator mites are killed applying highly toxic insecticides, it will be very difficult to get ERM under control, requiring multiple applications of expensive miticides. Organic growers rarely have problems with ERM because the pest control materials they use tend to be relatively non-toxic to predatory mites. In India the primary biological control agent on ERM is the predatory mite, Amblyseius fallacis. Other predatory mites like Agistemis fleschneri and Zetzellia mali may be present in lower numbers. In addition, a small, black ladybird beetle, Stethorus punctum feed on mites, especially late in the season.

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In case of heavy summer season infestations occurs, the treatments with acaricides like abamectin, hexythiazox, clofentizine, HMO, etc. should be given. HMO should not be mixed with other sulphur containing products as these are phytotoxic at higher temperatures and may affect fruit finish. ERM have shown the ability to develop resistance to a number of miticides. Resistance usually occurs when a single miticide or related group of miticides are used repeatedly. Each miticide has a particular mode of action. One way of avoiding resistance is to rotate between products having different mode of action.

12.2.6.2 Two-Spotted Spider Mite (Tetranychus urticae Koch) It is one of the most polyphagous pests that feed on tree fruits. It is distributed worldwide and is of economic importance. This species is characterised by the presence of two dark spots, appearing throughout the transparent body wall. Since the spots are accumulation of body wastes, newly moulted mites may lack the spots. Among the tree fruits, apple, pear, peach, nectarine, apricot, cherry (sweet and sour), plum, and prune are suitable hosts of this mite. Life Cycle The life cycle is composed with the egg, the larva, two nymphal stages (protonymph and deutonymph), and the adult. The length of time from egg to adult varies greatly depending on temperature. It completes their development period in 5–20 days. The two-spotted spider mite (Fig. 12.27) overwinters as yellowish orange coloured adult female under the bark or ground cover. Only females are known to overwinter. They Fig. 12.27 Two spotted spider mite (Tetranychus urticae Koch)

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emerge from their overwintering sites at about the half-inch green stage of apple development. They gradually lose their orange colour and gain their normal greenish and dorsal spots. After about 2–5 days, egg laying begins, primarily on the underside of newly expanded leaves. Later the fruitlets develop at the rate of 2–6 a day. Overwintering females lay on an average of 39 eggs during her life time; Summerform females can lay about 100 eggs. These eggs may take 2–6 days to hatch, depending upon the temperature. Egg hatch takes only 1 or 2 days during the summer, and the entire generation time (oviposition to adult) may take only 10 days. After hatching the six legged larva immediately starts feeding on leaves. Later it changes into eight legged nymphal stage, and then into an adult stage. When leaf quality begins to decline (e.g. from excessive mite feeding), or when cooler temperatures and shorter day lengths occur during the fall, the orange overwintering forms are again produced. Under optimum conditions of high temperature and low humidity, the life cycle may be completed in 7 days. Females can lay 200 eggs. There may be 7–10 generations per season or even more during hot dry weather persists. Damage Because of their small size and habit of feeding on the underside of foliage, this species may go undetected until a population has caused serious damage to a plant. Hot dry weather favours the development of severe infestations of the pest. This species cause damage by sucking the plant sap from the foliage. In case of heavy infestations the leaves look chlorotic, mottles or stippled in appearance. On close investigation the infested plants reveal fine, silken threads over the foliage, twigs, and branches. These silken threads are produced by the mites as they move across the leaf surface. Hot dry weather favours the development of severe infestations of this pest. Management The population of overwintering mites can be reduced by orchard sanitation and destruction of weeds. Destruction of weeds adjacent to and in fields should be done in the fall or early spring. Growers should manage weeds around fields carefully during the season. Grass should be mowed regularly. Use of overhead-sprinkler irrigation may provide some short-term relief of mite infestations. Adequate watering of the plants during dry conditions can limit the population of spider mites. The natural enemies also play an important role in managing the mites. Predatory mites (Phytoseiidae) and ladybird beetle Stethorus are the primary arthropod natural enemy of two-spotted spider mites and could be used through conservation and augmentation techniques. The severe infestation of mite can be managed by applying registered miticides and should be repeated as needed at 7–10 days interval. Mite populations may develop resistance to any chemical used against them. Therefore, chemicals of different mode of action should be used if more than one treatment is required within a season, rather than applying a chemical with similar mode of action. Horticulture mineral oil may be applied when mites first appear in early spring. The miticide treatment may be delayed if rain with cooler temperature and high humidity are expected.

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12.2.6.3 Walnut Blister Mite Aceria erinea (Nalepa) The walnut blister mite occasionally occurs in walnut orchards. Adult mites are very small and cannot be seen without a 14–20 hand lens. They have a white, slender, striated body with a few long hairs and two pairs of legs. Immature forms resemble adults but are smaller in size. Eggs are spherical and pearly white. Life Cycle This mite overwinters beneath bud scales. When the weather warms up, the mites feed beneath the leaves among the leaf hairs. Fertilisation occurs when the females come in contact with sperm sacs left on the host by males. Females lay as many as 80 eggs in 1 month under favourable conditions. Several generations occur during the summer, which attack new foliage as soon as it unfurls. Alternating generation is more common in eriophyids that feed on deciduous, woody plants and appears to be an adaptation based on the seasonal changes of the hosts. Damage and Symptoms The mites cause raised yellowish-green blisters on the upper leaf surfaces (Fig. 12.28). Yellowish or brown concave pockets are formed on underside of leaf, where the mites can be found. These mites cause no appreciable harm to the tree, and do not inhibit the development of nuts or fruits, leave no control measures are necessary (the leaves just look bad). Management 1. Broad leaf weeds like mallow, bindweed, white clover, and knotweed enhance mite numbers. Avoid excessive nitrogen applications, as this encourages mites. Fig. 12.28 Walnut blister mite Aceria erinea (Nalepa) cause yellowish-green blisters on the upper leaf surfaces

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2. Pruning off and disposing of infested portions may offer some control. 3. Phytoseiid predator mites always keep mites under control if broad-spectrum insecticide applications are avoided. Heavy rain and cold weather also suppress mite numbers. 4. Heavy infestations can be controlled with miticides, but spraying will not get rid of the galls of erinea once produced. Apply miticides just after bud break in early spring. Dormant oil, horticultural mineral oils, and insecticidal soaps may be effective. Exposed mites are easily controlled, but most pesticides do not kill the mites living within galls.

12.3

Conclusions

Sucking pests pierce plants with slender, sharp-pointed mouthparts and suck the plant sap. Withdrawal of the sap results in minute white, brown, or red spotting on the leaves, fruits, or stems of the plant. They may cause curling of leaves, deformed fruit, general wilting, browning, and drying of the entire plant. Some of these pests are important virus vectors, transmitting a range of plant virus. Virus-infected plants cannot be cured. However, control measures can be used to prevent or reduce the levels of the disease in crops by removing, or avoiding the sources of virus infection, and minimising spread by these sucking pests. For sustainable management sucking insect pests and mites, it is important to know how to identify and monitor the pests and also identify the symptoms of the diseases they cause. Many of these pests are also resistant to pesticides. In addition, knowledge of management options is needed. Use an Integrated Pest Management (IPM) approach to monitor not just the pests but also the beneficial that are naturally occurring onto the crops. Experienced Integrated Crop Protection (ICP) researchers and consultants have provided valuable guidance and insights to assist the growers to implement IPM on their farms.

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Sucking Pests of Mango

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P. Venkata Rami Reddy, M. A. Rashmi, K. Sreedevi, and Sandeep Singh

Contents 13.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13.2 Leafhoppers (Hemiptera: Cicadellidae) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13.3 Mealybugs (Hemiptera: Pseudococcidae) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13.4 Thrips (Thysanoptera: Thripidae) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13.5 Shoot Gall Psylla (Hemiptera: Psyllidae) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13.6 Midges (Diptera: Cecidomyiidae) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13.7 Scales (Hemiptera: Coccidae) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13.8 Mites (Acari:Tetranychidae and Eriophyidae) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13.9 Minor Sucking Pests . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13.10 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract

Mango, Mangifera indica L., is grown throughout the subtropics and tropics and is one of the world’s most important fruit crops. Mango is vulnerable to a variety of pests including insects, mites, pathogens and vertebrates. However insect pests often pose serious threat to the profitable cultivation. Insect pests considered economically important to mango are leafhoppers, mealybugs, thrips, fruit flies, stone weevil, stem borer, etc. The changing climate, growing number of P. V. R. Reddy (*) Division of Entomology and Nematology, ICAR-Indian Institute of Horticultural Research, Bengaluru, India M. A. Rashmi Regional Plant Quarantine Station, Bengaluru, India K. Sreedevi ICAR-National Bureau of Agricultural Insect Resources, Bengaluru, India S. Singh Department of Fruit Science, Punjab Agricultural University, Ludhiana, India # Springer Nature Singapore Pte Ltd. 2020 Omkar (ed.), Sucking Pests of Crops, https://doi.org/10.1007/978-981-15-6149-8_13

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monovarietal orchards, clean cultivation and chemical intensive plant protection measures have contributed to significant shifts in pest community structures. Those pests considered to be of minor importance sometime back have become major, like thrips, and vice versa. Hence there is a need to constantly update the information on pest status to enhance our preparedness to tackle them effectively. This chapter deals exclusively with the sucking pests, including insects and mites, occurring on mango, their biology, damage potential and management strategies.

13.1

Introduction

Mango, Mangifera indica L., widely acclaimed as ‘King of Fruits’ is a major fruit crop of tropical and subtropical regions. A native of Indo-Burma region, mango is cultivated in India, China, Thailand, Mexico, Pakistan, Philippines, Indonesia, Brazil, Nigeria and Egypt. India contributes about 50% of the world’s mango production with 2.5 million hectares with an annual production of 18.0 million tons (Reddy et al. 2018). Like any other cultivated crops, mango is vulnerable to biotic stresses induced by several kinds of pests like insects, mites, pathogens, vertebrates, etc. Among these, insect pests are a major constraint in sustainable cultivation of mangoes. Exhaustive lists of insect pests attacking mango compiled by different workers indicate that about 400 species of insect pests infest mango in different parts of the world (de Laroussilhe 1980; Tandon and Verghese 1985; Veeresh 1989; Pena et al. 1998). They include sap feeders, borers, defoliators and fruit pulp feeders. Of them, sucking pests comprising mainly those from Orders Hemiptera (leafhoppers, mealybugs, scales), Thysanoptera (thrips) and non-insect group, Acarina (mites), form a major chunk. They cause both direct and indirect losses. Sucking insects, characterized by shorter life cycles and ability to produce asexually, pose serious challenge to plant protection experts. They are also more sensitive and responsive to climatic variations (Jayanthi et al. 2014), which leads to frequent outbreaks. This chapter deals with the species diversity, distribution, ecology and management of major sucking pests of mango (Fig. 13.1).

13.2

Leafhoppers (Hemiptera: Cicadellidae)

Species and Distribution Leafhoppers are the major sucking pests of mango. About 15 species are reported to occur on mango in Asia, of which four species viz. Amritodus atkinsoni (Lethier), Idioscopus nitidulus (Walker), I. clypealis (Lethierry) and I. nagpurensis (Pruthi) are of economic importance. They are distributed across mango growing regions of the country (Veeresh 1989; Waite 2002), though the geographical distribution of different species is not uniform across the country. For instance A. atkinsoni and I. clypealis are more severe in northern states while I. nitidulus and I. nagpurensis are predominant in the southern part of India (Veeresh 1989).

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Mango fruit infested with hoppers

Adult hoppers resting on tree bark

Sootymold growth on hopper infested leaf

Mealybug on mango fruits

Gall infested mango fruits

Maggot of gall midge inside fruit gall

Mango leaf galls

Mango blossom midge damage

Mites on mango leaf

Thrips damage on tender fruits

Fig. 13.1 Major sucking pests of mango

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Biology Major hopper species are easily distinguishable with naked eye based on certain morphological characters. A. atkinsoni is relatively bigger (4.2–5 mm) of all three species and is dark grey in colour. It has two prominent spots on the abdomen and scutellum. In case of I. nitidulus there are three spots on the scutellum with a prominent white band across wings. It is light brown in colour and is slightly smaller than A. atkinsoni and bigger than I. clypealis, the smallest of three (3.5 mm). It also bears two spots on the scutellum (Butani 1979). Eggs are laid singly by female adults on tender shoots, flower buds and new foliage. One adult female can lay up to 200 eggs. The duration of egg stage ranges from 4 to 7 days. Nymphal period lasts for 8–13 days with three to four instars. It takes 15–19 days to complete one generation and there are 2–3 generations in a year depending on the geographical location. The insect overwinters as adult. Hoppers are reported to rest in the cracks and crevices under the bark of main trunk during hot noon and rainy days (Patel et al. 1973). The population reaches a peak during March–April and maximum and minimum temperature and relative humidity are major abiotic factors contributing to population fluctuations (Tandon et al. 1983). The spacing of mango trees in orchards also plays an important role in breeding of the hoppers. I. nitidulus occurs during both vegetative and flowering stage while I. clypealis and I. nagpurensis survive only on inflorescence (Verghese and Devi Thangam 2011). The hopper incidence is more severe in closely planted orchards and on those varieties with dense inflorescence (Srivastava 1997; Reddy and Dinesh 2005). Nature of Damage Nymphs and adults of leafhoppers cause damage by sucking sap from flower buds, flowers, shoots, tender foliage and young fruits. The extent of damage due to leafhoppers, if unchecked, may be as high as 100% (Verghese and Devi Thangam 2011). Besides sucking sap, they excrete honey dew which attracts sooty mould. This makes affected plant parts turn black and adversely affects photosynthetic efficiency of leaves. Severe infestation causes withering and dropping of florets, thus leading to failure of fruit setting (Butani 1979). The affected trees exhibit leaves and fruits shining with honey dew. Another common symptom of hopper damage is congregation of honey bees on leaves to collect honey dew and also one would experience droppings of honey dew drops while walking under affected trees. Management 1. Full grown trees, especially centre branches, have to be pruned to facilitate adequate light penetrance. 2. Spray the botanical pesticides, like azadirachtin 1% @ 3 mL/L if the hopper population is low (