Microbes for Sustainable Insect Pest Management : An Eco-friendly Approach - Volume 1 [1st ed. 2019] 978-3-030-23044-9, 978-3-030-23045-6

Table of contents :
Front Matter ....Pages i-x
Synthetic Chemical Insecticides: Environmental and Agro Contaminants (Md. Aslam Khan, Wasim Ahmad)....Pages 1-22
Soil-Borne Entomopathogenic Bacteria and Fungi (Tan Li Peng, Samsuddin Ahmad Syazwan, Lee Seng Hua)....Pages 23-41
Molecular Phylogeny of Entomopathogens (Mudasir Gani, Taskeena Hassan, Pawan Saini, Rakesh Kumar Gupta, Kamlesh Bali)....Pages 43-113
Potential of Entomopathogenic Bacteria and Fungi (Lav Sharma, Nitin Bohra, Rupesh Kumar Singh, Guilhermina Marques)....Pages 115-149
Ascomycota and Integrated Pest Management (Tariq Ahmad, Ajaz Rasool, Shaziya Gull, Dietrich Stephan, Shabnum Nabi)....Pages 151-183
Thermotolerance of Fungal Conidia (Flávia R. S. Paixão, Éverton K. K. Fernandes, Nicolás Pedrini)....Pages 185-196
Oxidative Stress in Entomopathogenic Fungi and Its Potential Role on Mycoinsecticide Enhancement (Carla Huarte-Bonnet, M. Constanza Mannino, Nicolás Pedrini)....Pages 197-205
Effects of Cytotoxic Factors Produced by Entomopathogenic Bacteria on Insect Haemocytes (Carlos Ribeiro, Amélia Vaz)....Pages 207-245
Effects of Entomopathogenic Nematodes and Symbiotic Bacteria on Non-target Arthropods (Ramandeep Kaur Sandhi, Gadi V. P. Reddy)....Pages 247-273
Granuloviruses in Insect Pest Management (Pankaj Sood, Amit Choudhary, Chandra Shekhar Prabhakar)....Pages 275-298
Interactions of Entomopathogens with Other Pest Management Options (Surendra K. Dara)....Pages 299-316
Toxicological Prospects on Joint Action of Microbial Insecticides and Chemical Pesticides (A. R. N. S. Subbanna, J. Stanley, V. Venkateswarlu, V. Chinna Babu Naik, M. S. Khan)....Pages 317-340
Entomopathogen and Synthetic Chemical Insecticide: Synergist and Antagonist (Arash Zibaee)....Pages 341-363
Current State of Fungal Antagonists with Special Emphasis on Indian Scenario (Purnima Das, Lakshmi Kanta Hazarika, Surajit Kalita, Somnath Roy)....Pages 365-385
Back Matter ....Pages 387-396

Citation preview

Sustainability in Plant and Crop Protection

Md. Aslam Khan Wasim Ahmad Editors

Microbes for Sustainable lnsect Pest Management An Eco-friendly Approach - Volume 1

Sustainability in Plant and Crop Protection Series Editor Aurelio Ciancio, Sezione di Bari, Consiglio Nazionale delle Ricerche Istituto per la Protezione delle Piante, Bari, Italy

More information about this series at http://www.springer.com/series/13031

Md. Aslam Khan  •  Wasim Ahmad Editors

Microbes for Sustainable Insect Pest Management An Eco-friendly Approach - Volume 1

Editors Md. Aslam Khan Department of Biology, Faculty of Science Jazan University Jazan, Saudi Arabia

Wasim Ahmad Department of Zoology, Section of Nematology Aligarh Muslim University Aligarh, Uttar Pradesh, India

ISSN 2567-9805     ISSN 2567-9821 (electronic) Sustainability in Plant and Crop Protection ISBN 978-3-030-23044-9    ISBN 978-3-030-23045-6 (eBook) https://doi.org/10.1007/978-3-030-23045-6 © Springer Nature Switzerland AG 2019 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, express or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Springer imprint is published by the registered company Springer Nature Switzerland AG. The registered company address is: Gewerbestrasse 11, 6330 Cham, Switzerland

Foreword

Insect pests are responsible for major losses to agricultural products globally. Synthetic chemical insecticides have been the primary control agent of insect pests for decades. Hazards and side effects associated with the extensive and indiscriminate use of pesticides are well documented and are of great concern. Resistance to synthetic insecticides is also a critical problem in several parts of the world. Development of integrated pest management strategies with little or no reliance on chemical pesticides has become an important goal for modern agriculture. Biological control is a component of IPM strategies that minimize insecticide spray applications and move towards ecofriendly systems of pest management. Due to high specificity, effectiveness, and safety to nontargeted organisms, biological control agents are considered suitable alternatives to the use of chemical pesticides. In recent years, biological control using microorganisms like bacteria, fungi, viruses, and nematodes has emerged as a valuable tool. In the present scenario, it is praiseworthy to have a book comprising potentials of different entomopathogens in the sustainable management of insect pests. The current volume should prove a very timely action in this direction. The different chapters in this book provide valuable information in this regard. The editors of this volume together with the authors of the individual chapters have made a remarkable contribution in collating the up-to-date information on sustainable and ecofriendly management of pernicious insect pests.

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This information could be useful for researchers, educators, students, and industry persons for understanding and developing ecofriendly and sustainable pest management strategies. This book comprehensively addresses various methods related to sustainable management of insect pests through the expertise of the leading authors worldwide. Finally, this book in the series Sustainability in Plant and Crop Protection is highly innovative in covering both basic information and effective management of insect pests. Director, Zoological Survey of India Kolkata, India

Kailash Chandra

Preface

It is generally acknowledged that agricultural yields must be increased in the coming years in order to feed the projected growth of the global human population. An enormous amount of resources is spent each year, worldwide, to manage and control insect pests. A great proportion of the agricultural yields is, however, still lost to insect damage, predominantly in developing countries. Therefore, the need to keep insect pests away from destroying food crops has become even more urgent. The negative impacts of synthetic organic pesticides in crop pest management programs caused tremendous damages to the environment, including the development of resistance among target pests, and other detrimental effects on nontarget organisms, underlying the need for alternative and eco-friendly control methods. Biological control-based approaches provide a more environment-friendly and acceptable alternative to traditional chemical control measures, owing to high host specificity and safety for the environment and mankind. Microorganisms have been regularly isolated from natural sources around the world for pest management purposes. Some of them, known as entomopathogens, may safeguard crops and plants by causing epidemic diseases of insect populations. They have been widely tested and proved to be very effective against pernicious pests. Almost all the major insect groups are susceptible to entomopathogens. In certain developing and developed countries, a number of species and isolates have been registered for field application on various vegetables, fruits, and other crops of agricultural, horticultural, and forest importance. It is therefore imperative to study the entomopathogens and their potential role in achieving a sustainable pest management goal. This volume comprises 14 chapters in an attempt to bring available information on safe use of entomopathogens. The soil constitutes an important reservoir for harvesting various types of beneficial organisms. Chapters dealing with soil-borne entomopathogens and their phylogeny provide a review and most updated information of their isolation and molecular identification. Fungal pathogen applications play a key role in biological control programs. In other chapters, thermotolerance and oxidative stress are examined and explored. A number of entomopathogenic

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bacteria are able to kill their host quickly. The information provided upon cytotoxic factors for insect hemocytes constitutes an important contribution. Nematodes as biological control agents are safer alternatives for the environment, as shown by the information provided on their direct and indirect effects on nontarget organisms. Being highly specific, virulent, and safer to nontarget species, viruses are potential candidates for use as biological insecticides. A separate chapter on the role of granuloviruses in IPM contributes a wealth of information on this topic. Biopesticides in combination with synthetic insecticides are effective, economic, and eco-friendly. Understanding their interactions will certainly promote their use, as shown in reviews of synergistic and antagonistic interactions of microbial and chemical pesticides, provided in further chapters. We hope that this volume will be helpful to students, teachers, researchers, and industry technicians. We are highly grateful to all the contributors for providing their expertise in the form of stimulating contributions. Thanks are due to the Head, Biology Department, and Dean, Faculty of Science, Jazan University, Jazan, for their moral support. We are grateful to Dr. Aurelio Ciancio, CNR, Bari, Italy, for including this volume in the Springer series “Sustainability in Plant and Crop Protection.” We extend our thanks to Springer International team for their generous cooperation at every stage of the book production. Jazan, Saudi Arabia Aligarh, India 

Md. Aslam Khan Wasim Ahmad

Contents

1 Synthetic Chemical Insecticides: Environmental and Agro Contaminants........................................................................................... 1 Md. Aslam Khan and Wasim Ahmad 2 Soil-Borne Entomopathogenic Bacteria and Fungi.............................. 23 Tan Li Peng, Samsuddin Ahmad Syazwan, and Seng Hua Lee 3 Molecular Phylogeny of Entomopathogens........................................... 43 Mudasir Gani, Taskeena Hassan, Pawan Saini, Rakesh Kumar Gupta, and Kamlesh Bali 4 Potential of Entomopathogenic Bacteria and Fungi............................. 115 Lav Sharma, Nitin Bohra, Rupesh Kumar Singh, and Guilhermina Marques 5 Ascomycota and Integrated Pest Management..................................... 151 Tariq Ahmad, Ajaz Rasool, Shaziya Gull, Dietrich Stephan, and Shabnum Nabi 6 Thermotolerance of Fungal Conidia...................................................... 185 Flávia R. S. Paixão, Éverton K. K. Fernandes, and Nicolás Pedrini 7 Oxidative Stress in Entomopathogenic Fungi and Its Potential Role on Mycoinsecticide Enhancement.................................................. 197 Carla Huarte-Bonnet, M. Constanza Mannino, and Nicolás Pedrini 8 Effects of Cytotoxic Factors Produced by Entomopathogenic Bacteria on Insect Haemocytes............................................................... 207 Carlos Ribeiro and Amélia Vaz 9 Effects of Entomopathogenic Nematodes and Symbiotic Bacteria on Non-target Arthropods........................................................ 247 Ramandeep Kaur Sandhi and Gadi V. P. Reddy

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10 Granuloviruses in Insect Pest Management.......................................... 275 Pankaj Sood, Amit Choudhary, and Chandra Shekhar Prabhakar 11 Interactions of Entomopathogens with Other Pest Management Options............................................................................... 299 Surendra K. Dara 12 Toxicological Prospects on Joint Action of Microbial Insecticides and Chemical Pesticides..................................................... 317 A. R. N. S. Subbanna, J. Stanley, V. Venkateswarlu, V. Chinna Babu Naik, and M. S. Khan 13 Entomopathogen and Synthetic Chemical Insecticide: Synergist and Antagonist......................................................................... 341 Arash Zibaee 14 Current State of Fungal Antagonists with Special Emphasis on Indian Scenario................................................................. 365 Purnima Das, Lakshmi Kanta Hazarika, Surajit Kalita, and Somnath Roy Index ................................................................................................................. 387

Chapter 1

Synthetic Chemical Insecticides: Environmental and Agro Contaminants Md. Aslam Khan and Wasim Ahmad

Abstract  Synthetic pesticides are indispensable in intensive agricultural productions. For decades these compounds served as backbone in insect pest management. Due to persistence and pervasiveness of millions of tonnes of synthetic chemical pesticides applied, almost every ecosystem has received a negative impact. In the present chapter an effort has been made to highlight the environmental contaminations caused by synthetic chemical pesticides, their adverse effects on human health and other non target organisms, the development of resistance in target insect pests, along with the degradation of synthetic pesticides. Keywords  Synthetic pesticides · Environment · Contaminant · Non target organism · Human health

1.1  Introduction Insects, plants, bacteria, fungi and other organisms occur naturally in the environment but in some situations they can have environmental, health and economic impacts, and become pests that must be controlled. For many generations, natural pest control in agricultural practices relied heavily on crop rotation or mixed crop planting (Dayan et al. 2009). In view of the world rapid human population growth and limited croplands, it is needed to apply all available measures to increase crop production in order to ensure food security (Zhang 2009) and optimize provision of food at low cost. In this view, a worldwide agricultural movement that originated in Mexico in 1944, with the primary goal of boosting grain yields, was named as “Green revoluM. A. Khan (*) Department of Biology, Faculty of Science, Jazan University, Jazan, Saudi Arabia e-mail: [email protected] W. Ahmad Department of Zoology, Section of Nematology, Aligarh Muslim University, Aligarh, Uttar Pradesh, India © Springer Nature Switzerland AG 2019 M. A. Khan, W. Ahmad (eds.), Microbes for Sustainable Insect Pest Management, Sustainability in Plant and Crop Protection, https://doi.org/10.1007/978-3-030-23045-6_1

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tion”. Based on higher yields new cultivars were selected but most of those varieties were not widely resistant to pests and diseases. Therefore the pest control issue was mostly addressed by the use of synthetic chemical pesticides, as an integral part of the green revolution. More pesticide inputs were, however, needed for the green revolution, more than in the traditional agricultural systems. Following its success in Mexico, the green revolution concepts spread all over the world. As high temperature and humidity are highly conductive to rapid multiplication of pests, crop losses remain more severe especially in tropical countries. Insect pests, plant pathogens and weeds cause an estimated 14%, 13% and 13% of loss, respectively (Pimentel 2009). Without pesticides application a loss of 78% fruits, 54% vegetables, and 32% cereal crops has been reported, due to pest injury (Cai 2008). Therefore, pesticides appeared indispensable in agricultural productions as it was conceived that they were not a technical luxury rather a necessity for the well being of mankind. According to the Environmental Protection Agency (EPA 2009) a pesticide is any substance or mixture of substances intended for preventing, destroying, repelling, or mitigating any pest like insects, mites, nematodes, weeds and rats etc. Major categories of pesticides include insecticides, herbicides and fungicides/bactericides, but several other types of biocides, such as nematicides and rodenticides etc., are also included. Due to longer residual action and a wide toxicity spectrum synthetic chemical insecticides became more popular. Dichloro-diphenyl-trichloroethane (DDT) was the first important synthetic organic insecticide, synthesized by German scientist Ziedler in 1873 (Othmer 1996). Its insecticidal effect was discovered by Swiss chemist Paul Muller in 1939. DDT hailed as miracle in its early days because of a broad-spectrum activity, persistence, insolubility, low cost and ease to apply (Keneth 1992). For decades these compounds served as backbone in insect pest management and farmers relied heavily upon the conventional groups of synthetic insecticides such as organochlorines (DDT, BHC), cyclodienes (aldrin, dieldrin, endosulfan), organophosphates (monocrotophos, quinalphos, chlorpyriphos, profenophos, dimethoate, phosalone, metasystox, acephate, phorate, methyl parathion), carbamates (carbosulfan, carbaryl, thiodicarb, methomyl), pyrethroids (cypermethrin, deltamethrin, fenvalerate, λ-cyhalothrin), and formamidines (chlordimeform and amitraz) (Kranthi 2007). Many of the available pesticides are often a mixture of several chemicals ingredients mixed together in desired proportions, suspended in appropriate carriers or diluent materials. Different forms of pesticides include emulsifiable concentrate, wettable powder, granule, bait, dust and fumigant. A comprehensive review on identity, physical and chemical properties of pesticides was produced by Tano (2011). Application of million tonnes and hundred types of synthetic pesticides, however, reduced crop losses but due to their persistence and pervasiveness, almost every ecosystem on earth has receive one or more negative impacts (Reuter and Neumeister 2015), with longterm affect on society. It is now clear that adverse effects of pesticides are wide and varied. The public has expressed today many concern about the potential health and environmental impacts of these pesticides. This chapter collates the adverse effects of synthetic chemical insecticides from the growing body of related literature.

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1.2  Background and History Before 1870s natural pesticides, for instance sulfur in ancient Greece, were used to control pests. From 1870 to 1945 natural materials and inorganic compounds were mainly used as pesticides. Since 1945, man-made organic synthetic compounds terminated the era of inorganic and natural pesticides (Zhang et al. 2011). After that time most pesticides have been synthesized by man, and named as chemical pesticides. Based on chemical classification, insecticides are broadly grouped as: organochlorines, organophosphorous, carbamates and pyrethroids. For insect pests control most spectacular episodes began in 1945–1946 with the commercial introduction of organochlorines such as DDT, followed by organophosphate and carbamate introduced in the ‘60s (Nicholson 2007). Organochlorines, the first synthetic organic pesticides to be used in agriculture and in public health, are contact poisons that apparently act as nervous system disruptors leading to convulsions and paralysis of the insect and its eventual death. They have a long-term residual effect in the environment as they are resistant to most chemical and microbial degradations. Organophosphorous insecticides are cholinesterase inhibitors in target pests. As a result, nervous impulses fail to move across the synapses, causing a rapid twitching of voluntary muscles and hence paralysis and death. Unlike organochlorines, organophosphorous insecticides are easily decomposed in the environment by various chemical and biological reactions. Carbamates are organic pesticides derived from carbamic acid. They show high insect toxicity as cholinesterase inhibitors. The cholinesterase inhibition of carbamates differs from that of organophosphorous because it is species-specific and reversible (Drum 1980). Pyrethroids are synthetic analogues of the naturally occurring pyrethrins. They are known for the fast knocking down effect active against insect pests, with low mammalian toxicity and easy biodegradation. As they are particularly susceptible to photolysis, their uses as agricultural insecticides is relatively impractical. In each country, regulatory insect risk assessment related to agrochemicals use and registration follows specific guidelines such as the European Council Directive 91/414 in Europe, and the Federal Insecticide Fungicide and Rodenticide Act in the United States. In India the pesticides import, manufacture, sale, transport and use are regulated under a comprehensive statute, the Insecticides act of 1968. The worldwide consumption of pesticides is around 2–3 million tonnes per year (United States Environment Protection Agency 2011), of which 45% is consumed in Europe, 25% in the United States, and 30% in the rest of the world (De et al. 2014). The usage of pesticides in India is only 0.5 kg ha−1, while in Korea and Japan it is 6.6 and 12.0 kg ha−1, respectively (Abhilash and Singh 2009). Annual pesticide consumption in different countries is represented in Table 1.1. Total expenditure on pesticides is about US$ 40 billion per year (Popp et al. 2013). Ssynthetic insecticides entail several types of costs, including internal costs due to the purchase and application, and various other additional hidden costs due to the impact of treatments on the environment and human health (Bourguet and Guillemaud 2016). Zhang et al. (2011) reviewed the consumption, pollution and developmental trend

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Table 1.1  Annual pesticide consumption by different Asian countries (Abhilash and Singh 2009) Country Bangladesh Cambodia China DPR Korea India Rep. of Korea Lao PDR Malaysia Myanmar Nepal Pakistan Philippines Sri Lanka Thailand Vietnam

Ton a.i. 3635 42 258000 3000 41020 26610 10 51065 758 145 32500 7934 1696 49108 24473

Ton product 22100 198 1000000 12000 164080 100000 40 204260 3030 580 129589 31735 6329 132509 50000

US $ value (000) 75000 226 5670000 60000 820400 842638 200 85020 15095 2100 172300 158675 49000 253537 159000

of pesticide varieties and reported that the overall consumption structure has undergone significant changes since 1960s. The proportion of herbicides in pesticide consumption increased rapidly, and the consumption of insecticides and fungicides/ bactericides declined. Dayan et al. (2009) also reported that herbicides account for more than half of the volume of all agricultural pesticides applied in the developed world (Fig.  1.1). Pesticide production in India is dominated by insecticides and fungicides, followed by herbicides and rodenticides (Fig. 1.2). The major markets for pesticides are the USA, Western Europe and Japan (Dinham 2005). Europe is the largest pesticide consumer in the world. United States, Brazil, Germany, France, China and Japan are the largest pesticide producers, consumers or traders in the world (Zhang et al. 2011). India ranks twelfth in the world for the use of pesticides. The Indian pesticides industry started in 1952 with the BHC technical plant, near Kolkata. Thereafter Hindustan insecticides Ltd. set up two units to manufacture DDT. Union Carbide then set up a plant in 1969 named as Union Carbide India Ltd., at Bhopal city. The industry produced various pesticides for the Indian market till the 1984 Bhopal disaster. Now the Indian pesticide industry comprises of more than 125 basic producers of large and medium scale (Abhilash and Singh 2009). Although the Indian average pesticide consumption is far lower than many other developed economies, the problem posed by the pesticide residues is very severe (Abhilash and Singh 2009). Inappropriate application of pesticides affects the whole ecosystem by entering the residues in food chains and polluting soil, air, ground and surface water (Agnihotri 1999; UN/DESA 2002). For this reason Rachel Carson (1962) in her book “The Silent Spring” described this era as that of the “rain of chemicals”.

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Fig. 1.1  Comparison of pesticide use in India and worldwide. (Adapted from Abhilash and Singh 2009)

Fig. 1.2  Share of pesticide groups in total pesticide production (technical grade), in India. (Adapted from Subash et al. 2017)

Monocrotophos, phorate, phosphamidon, methyl parathion and dimethoate are some of highly hazardous pesticides that are continually and indiscriminately used in India. As a consequence, the country is now battling against the residual effects of extensively used pesticides (Rekha and Prasad 2006; Agoramoorthy 2008). The Supreme Court of India appointed an expert committee to examine all aspects of the ban on endosulfan and the disposal of the existing quantity of this pesticide. On May 2011 the Supreme Court of India passed an interim order to ban the production, distribution and use of endosulfan in the country. Moreover, the High Court of

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Karnataka state directed the State government machinery to provide medical cover to all persons affected by the use of endosulfan, in certain coastal districts. As early as 1972, DDT, one of the most noxious pesticides ever used, and related organochlorinated insecticides, were banned in the United States, by the Environmental Protection Agency, and then in most other countries because of its potential harmful effects on the environment, wildlife and humans. During the late twentieth century, pesticide consumption in the United States declined by 35% without reducing crop production (SDNX 2005). In Europe, as in the United States, older pesticides are being re-assessed one-by-one, to ensure that they meet the new regulatory standards (Damalas and Eleftherohorinos 2011). Overall pesticide consumption in Europe declined in last decades. Since 2001 to 2008  in Europe 704 pesticides were banned, 26% of which were insecticides, 23% herbicides and 17% fungicides (Karabelas et al. 2009). China has banned the application of high-­residual DDT, HCH and other organochlorined pesticides since 1983. Since 2007, the highly poisonous organophosphorus pesticides parathionmethyl, parathion, methamidophos, and phosphamidon have been banned for using and selling in China. Recent reports indicate that the use of persistent organic pollutants (POPs) is declining. Wang et al. (2008) noted that since the use of organochlorined pesticides (OCPs) has been banned long, the residual OCPs are declining but in some regions they are still abundant. Covacia et al. (2005) also suggested a slight decline in the concentrations of DDT from Belgium sediments, due to the restrictions in their usage.

1.3  Environmental Contaminants Pesticides hold a unique position among environmental contaminants due to their high biological activity and toxicity. Indiscriminate, inadequate, and improper use of these synthetic organic inputs in crop pest management programs around the world caused tremendous damage to the environment, development of resistance in target pests, pest resurgence, detrimental effects on non-target organisms, and impact on human health (Casida and Quistad 1998; Shen and Zhang 2000; Niyaki 2010; Al-Zaidi et al. 2011). According to the Food and Agriculture Organization (FAO) inventory (FAO/UNEP/OECD/SIB 2001), more than 500,000 tonnes of unused and obsolete pesticides are threatening the environment and public health, in many countries. A report published in proceedings of the international academy of ecology and environmental sciences by Zhang et al. (2011) shows that only 1% of the sprayed pesticides are effective, the remaining 99% being released to non-target soils, water bodies and atmosphere, and finally absorbed by almost every organism. Some synthetic pesticides are extremely persistent in the environment because of their resistance to natural breakdown processes and are routinely found in the environment today. POPs and related degraded products flow into the atmosphere, soils and riv-

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ers, resulting in the accumulation of toxic substances, thus threatening the environment. High-residual pesticides such as DDT have been detected in the Greenland ice sheet and the bodies of Antarctic penguins, as the result of atmospheric circulation, ocean currents and biological enrichment along food chains. Environmental costs are those resulting both from pesticide damage to animals, plants, algae and microorganisms and from induced pest resistance. These costs may be incurred by farmers or by the society as a whole. In agricultural areas in which pesticides are used, these substances drift in the air, pollute the soil and waterways, and are sometimes systemically absorbed by non-target plant species. Pesticide pollution to the local environment also affects the lives of birds, wildlife, domestic animals, fish and livestock. The environmental pollution caused by pesticides in Asia, Africa, Latin America, the Middle East and Eastern Europe is now serious. Even in earlier years in India the residuals of DDT, lindane and dieldrin have been much beyond the safety threshold for fish, eggs and vegetables (Wu 1986). Willet et al. (1998) reported that although environmental levels of some organochlorines have fallen over time, many can still be found as contaminants in soils, river or coastal marine sediments, reaching as far as the deep oceans and poles. The treatise dealing with environmental and pollution science by Pepper et al. (2006) is worthy to be mentioned here. Pesticide pollution of environmental waters is a pervasive problem with widespread ecological consequences. During applications, pesticides drift away in the air and seep into the soil (Pimentel 1995; Gil and Sinfort 2005). Once in the soil, some soluble compounds may be washed out in runoff water and during soil erosion, resulting in leaching into rivers and lakes (Chopra et al. 2011). Hence the pesticide concentration of water bodies can reach the magnitude of several dozen mg/L. Excess concentration of pesticides into runoff water should be removed to protect water resources or to achieve drinking water quality. Under Council Directive 98/83/EC in Europe, the legally permitted limit for an individual pesticide in drinking water is 0.1 μg/L, whilst the total of all pesticides must not exceed 0.5 μg/L (Boobis et al. 2008). Wastewaters from agricultural or industrial activities contain high levels of pesticide contamination. Agriculture, which accounts for 70% of water abstractions worldwide, plays a major role in water pollution. Farms discharge large quantities of agrochemicals, organic matter, drug residues, sediments and saline drainage into water bodies. The resultant water pollution poses demonstrated risks to aquatic ecosystems, human health and productive activities (UNEP 2016). Chiron et al. (2000) reported that among the different approaches to pesticide elimination from wastewater, photochemical and ozonation methods appear to be especially suitable for industrial applications, whereas ozonation is more easily controlled and easier to adapt to industrial applications than photochemical.

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1.4  Impact on Human Health Despite strict regulations on the registration and use of pesticides, there are major concerns about their direct impact on human health. There is a growing body of evidence showing that health hazards of pesticides are serious. People are inevitably exposed to a cocktail of pesticides through environmental contamination or occupational use. There are no groups in the human population that are completely unexposed to pesticides, and most diseases are multi-causal giving considerable complexity to public health assessments (Meyer-Baron et al. 2015). Many pesticides can damage human health (Damalas and Eleftherohorinos 2011). Indiscriminate use and improper handling of synthetic pesticides in agriculture have caused acute (high doses over short periods) and chronic (lower doses over longer periods of time) health problems in human. Pesticide poisonings are also divided into productive poisonings, generated in the process of agricultural production, and living poisonings, i.e., suicide, ingestion, and food intake with high-residuals etc. (Liu et al. 2008). Exposure to pesticides can occur via a number of pathways like food, drinking water, residential, occupational (Fig. 1.3), and different routes like oral, inhalation and dermal. Million cases of pesticide poisonings have been documented every year around the world. In the USA, the Environmental Protection Agency reports an estimated 300,000 human pesticide poisonings as a part of the cost of their application (Pimentel and Burgess 2012). Worldwide, the number of severe pesticide

Fig. 1.3  Different pathways of pesticides exposure

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poisonings is much higher, as Richter (2002) reports 26 million human pesticide ­poisonings with 220,000 deaths occuring each year, the majority in developing countries. While these countries use only around 25% of the world pesticides, they experience 99% of deaths linked to pesticides (UNICEF 2018). In developing countries agricultural pesticides are among the most commonly used substances for self-­ poisoning. Gunnell et  al. (2007) conservatively estimated that there are 258,234 (plausible range 233,997 to 325,907) deaths from pesticide self-poisoning worldwide each year, accounting for 30% (range 27–37%) of suicides, globally. Pesticides have long been proposed as a possible cause of human health, but information on their health impacts is quite limited in many developing countries. However, the true extent of the problem is hard to determine for a variety of reasons. Health-care professionals in rural areas often fail to correctly diagnose poisoning, as many of the related symptoms are quite general in nature or mimic other common health problems (FAO 2001). The majority of the pesticide poisonings and deaths in the developing countries are the result of farmers’ poor handling practices and less awareness of the relative toxicity of the product they are using. Often they use pesticides at rates more intensive than those recommended by the product labels, dispose residual pesticides into canals or ditches, re-apply them to the same crops, or spray crops that were not identified for initial use (Huan and Thiet 2000). Agricultural workers are responsible for mixing and loading pesticide preparations, spraying, sowing pesticide-treated seeds, harvesting sprayed crops, and cleaning and disposing of pesticide containers. Moreover, they are often occupationally exposed to pesticides. Workers in the pesticide industry are also likely to experience occupational exposures. The families living in rural areas in which pesticides are intensively used may also be indirectly exposed to these chemicals, through off-­ target pesticide drift (Lee et al. 2011). Generally, each exposure pathway is the responsibility of a different department or agency within national governments or international bodies. Hence, assessment by each route is generally undertaken independently. The risk assessment of pesticide residues in food is currently performed on a compound-by-compound basis. However, it is the totality of exposure i.e. through multiple routes and multiple pathways, that determines the actual risk (WHO 2007). The information provided by Allsop et al. (2015) in the form of “pesticides and our health: a growing concern” is worthy to be mentioned here. Unregulated pesticide use and lack of enforcement mechanisms has resulted in thousand acute and chronic poisoning cases, with effects of varying severity on human health, from mild ones to death. Exposure to pesticides has also been the subject of great concern in view of its possible role in the induction of congenital malformations. Abell et  al. (2000) reported that exposure to pesticides to female workers in flower greenhouses may have reduced fertility, and that exposure may be part of the causal chain. In a study by Bosma et al. (2000) in the Netherlands, farmers and gardeners showed a higher risk of developing mild cognitive dysfunctions. Long-term cognitive effects due to low-level exposure to pesticides in occupational conditions have been reported by Baldi et al. (2001). Clinical reports have shown

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that acute intoxication by organophosphates may be responsible for chronic impairment of cognitive functions (Rosenstock et al. 1990). Acute poisoning, leading to respiratory, gastrointestinal, allergic, and neurologic disorders, is commonly reported by farmers, and particularly by those carrying out pesticide applications (Kishi et al. 1995; Hudson et al. 2014). Increasing incidence of cancer, chronic kidney diseases, suppression of the immune system, among male and female sterility, endocrine disorders, neurological and behavioral disorders, especially among children, have been attributed to chronic pesticide poisoning (Agnihotri 1999). Besides cancers and reproductive effects, nervous system damage has been reported in terms of peripheral neuropathy and central nervous degenerative disease, with special emphasis on Parkinson’s disease (Checkoway and Nelson 1999). In children organophosphates have been linked to aplastic anemia, the failure of the bone marrow to produce blood cells, and leukemia. In particular, children with asthma may have severe reactions to organophosphates (Zahm et al. 1997). Pesticide exposure has also been associated with elevated cancer risks and reproductive dysfunctions in agricultural workers (Raschke and Burger 1997; Horrigan et al. 2002). Potential carcinogenicity of a wide range of insecticides, fungicides and herbicides has been reviewed by the International Agency for Cancer Research (IARC) and fifty-six pesticides have been classified as carcinogenic to laboratory animals. It is estimated that cancer patients resulted from pesticide poisoning account for nearly 10% of total cancer patients (Gu and Tian 2005). Review on environmental chemicals and breast cancer by Rodgers et al. (2018) is worthy to be mentioned here. The United Nations Environmental Program (UNEP) has cautiously taken action to protect human health, the environment and the earth from further destruction by persistent organic pollutants such as DDT and its principal metabolite, dichloro-diphenyl-trichloro-ethylene (DDE). To a large extent DDT, the first major synthetic insecticide, replaced lead arsenate, a major carcinogenic pesticide used before the modern era (Ames et al. 1990). Now DDT has emerged as a potential human carcinogen. Both DDT and DDE are highly lipid soluble, accumulate in fat-containing foods, and travel through the food chain (Leber and Benya 1994; Spear 1999). DDT exposure in young women during the peak period of DDT use in the USA predicts breast cancer later in their life time (Cohn et al. 2007). In India the DDT content in the human body was ever the highest in the world (Zhang et  al. 2011). Anti-androgenic activity has been reported for DDE, that bioaccumulates in meat and dairy products (Kelce et al. 1995). Compared with adults, children may be more vulnerable to particular risks due to exposure to pesticides and related toxic effects. This vulnerability can be due to a number of factors, including differences in physiology, behavior, and environmental conditions (Goldman 1995; Reigart and Roberts 2001). Organochlorine pesticides display diverse endocrine activity in vitro and in animal models. Cholinesterase-inhibiting pesticides are intended for insects but they can also be poisonous to humans, in some situations. Human exposure to cholinesterase-­inhibiting pesticides can result from inhalation, ingestion, or eye or skin contact during the manufacture, mixing, or applications of these chemicals. Insecticide toxicity test includes study with blood tests for cholinesterase inhibition

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(reductions in acetylcholinesterase enzyme [AChE] due to contamination by chemical insecticides use). To test the acute and chronic pesticide poisoning Dasgupta et al. (2007) conducted an acetylcholinesterase enzyme blood test for 190 rice farmers in the Mekong Delta (Vietnam) and reported high prevalence of pesticide poisoning. Pesticides have been also considered as potential chemical mutagens. Various agrochemical ingredients possess mutagenic properties inducing mutations, chromosomal alterations or DNA damage. Genetic damage associated with high exposure levels of pesticides in human populations is reported by Bolognesi (2003).

1.5  Resistance Among Insect Pests Melander (1914) first reported insecticide resistance. Since then the subject has received ever-growing attention. A population is considered resistant if its response to an insecticide in detection tests drops significantly, below its normal response (Georghiou and Mellon 1983). This is particularly the case in warmer climates where insect infestation pressure is high. For all major insecticide classes, resistance has been reported in one or more key pest species (Georghiou 1986), including stored product insects. This resistance has steadily increased over the last 30 years (Rex Consortium 2013). The doses of pesticides applied to many crops are, therefore, almost certainly higher than used in the past, resulting in a greater impact on the environment. In 1979, the United Nations Environmental Programme declared pesticide resistance “one of the world’s most serious environmental problems.” In 2009 the European Union brought in a new framework directive requiring that all member states should achieve a level of sustainable use of pesticides (European Union 2009). In view of all these problems, the availability of many broad-spectrum chemical pesticides is declining, as a result of the evolution of resistance (Ishtiaq et al. 2012), and legislation (Chandler et al. 2011). In addition to direct mortality, toxic substances can cause physiological responses, such as changes in biochemical contents of the exposed insects. Broad-­spectrum chemical insecticides have been the primary control agent for agricultural pests, with about 40% targeted to the control of lepidopteran insects (Brooke and Hines 1999). In the Indian subcontinent, Australia, China and Africa, cotton bollworm, Helicoverpa armigera (Hubner) (Lepidoptera: Noctuidae) is arguably the most important agricultural pest. It has a long history of resistance to almost all the insecticide types used for its control (Gunning et al. 1999; Kranthi et al. 2002; Srinivas et al. 2004). Nair (1981) reported H. armigera “not a serious pest” of cotton in India. A few years later the pest was noticed to cause heavy economic losses to cotton and was found to withstand a sustained insecticide pressure. Subsequently high levels of resistance to synthetic pyrethroids in H. armigera were confirmed by Dhingra et al. (1988) and McCaffery et al. (1989). By 1992, H. armigera resistance to insecticides had emerged as a great challenge to pest management in Asia and Australia. Unlike other lepidopteran species, H. armigera larvae don’t migrate far from their original

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host plant, consequently their populations in agricultural areas are exposed to consistent selective pressures, leading to greater resistance to i­nsecticides (Fitt 1994). Phokela and Mehrotra (1989) stated that pyrethroid resistance in different strains of H. armigera appears to be mainly due to the high rate of metabolism in resistant lines. Aurade et al. (2010) however reported that the presence of a P-glycoprotein could be one of the reasons for insecticide resistance in this pest. During the late 1970s, Spodoptera litura (Fabricius) (Lepidoptera: Noctuidae), another major pest of subtropical and tropical agricultural crops, was found to exhibit high resistance to several conventional insecticides recommended for its control (Ramakrishnan et al. 1984; Tong et al. 2013). Among sucking pests, aphids were reported to have developed resistance against organophosphates, carbamates and pyrethroids (Herron et al. 2001; Ahmad and Akhtar 2013). The whitefly Bemisia tabaci was found resistant to BHC, endosulfan, diamethoate, phosalone, acephate, monocrotophos, quinalphos, and carbaryl (Prasad et al. 1993). Because of health, safety, environmental and economic considerations, only a very limited number of chemicals are available for application to stored grain. A serious threat to the continued availability of these materials is the development of resistance in target pests. Stored-grain pests such as lesser grain borer, Rhyzopertha dominica (F.), red flour beetle, Tribolium castaneum (Herbst), and rice weevil, Sitophilus oryzae (L.) developed resistance against the fumigant phosphine, a most important insect control treatment for stored grain, in many regions (Sartori et al. 1990). The last quarter of the twentieth century has seen the withdrawal of many compounds formerly used as fumigants. Methyl bromide, the fumigant with the widest range of applications, is scheduled to be completely phased out in the second decade of the current century under the directive of the Montreal Protocol, an United Nations agreement on ozone depleting substances. Phosphine is also under regulatory review in several developed countries, with some unpredictability in the outcome (Bell 2000). Commercial development of insecticidal genes has focused on the Bacillus thuringiensis (Bt) toxins (Bravo et al. 2007; Pigott and Ellar 2007). More recently there have been reports of field resistance to Bt crops in cotton bollworm, Helicoverpa spp. (Luttrell et al. 1999; Ali et al. 2006; Ali and Luttrell 2007; Carriere et al. 2010), armyworm, Spodoptera frugiperda (Lepidoptera: Noctuidae) (Storer et  al. 2010), pink bollworm, Pectinophore gosspiella Saunders (Lepidoptera: Gelichiidae) (Bagla 2010; Dhurua and Gujar 2011), and western corn rootworm, Diabrotica virgifera LeConte (Lepidoptera: Chrysomelidae) (Gassmann et  al. 2011). The most common mechanism for Bt resistance is the disruption of the Bt toxin binding to the receptors in the host midgut membrane. Insects that show resistance to one insecticide generally develop resistance to other classes of insecticides, a phenomenon often referred to as cross-resistance. This phenomenon resembles multidrug resistance whereby resistance to one drug is accompanied by simultaneous resistance to a variety of structurally unrelated compounds (Lanning et  al. 1996). The complex patterns of cross resistance between chemical groups and within groups further complicate the use of this strategy. In some regions, the situation is precarious with insect populations containing multiple

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resistances, leaving no effective protective options available. The intensive use of insecticides to control peach potato aphid, Myzus persicae Sulzer (Hemiptera: Aphididae) over many years has led to populations that are now resistant to several classes of insecticide. Myzus persicae has a remarkable ability to evolve mechanisms that avoid or overcome the toxic effect of insecticides, with at least seven independent selective processes for resistance (Bass et al. 2014). Biochemical and molecular mechanisms underlying resistance in M. persicae has been reviewed by Bass et  al. (2014). A comprehensive database (Arthropod Pesticide Resistance Database) by the Insecticide Resistance Action Committee, Michigan State University, provides thorough information for insect species resistant to different pesticides, along with the locations where resistance is reported (www.pesticideresistance.com). In the situation outlined above it can be easily concluded that insecticide-­based pest management desperately needs new chemistries with different resistance profiles.

1.6  Effects on Non -Target Organisms Many studies have documented direct and indirect effects of both high and sublethal doses of pesticides on several wild vertebrates. As stated above, only 1% of the sprayed pesticides are effective whereas the remaining 99% are released to non-­ target soils, water bodies and atmosphere, and finally absorbed by almost every organism. A possible decline of amphibian population caused by synthetic pesticides has long been suggested (Carey and Bryant 1995; Lips 1998). Davidson (2004) reported association between the spatial patterns of declines for five amphibian species in California and cholinesterase-inhibiting pesticides, mostly organophosphates and carbamates. Pesticides have a particularly strong impact on birds (Mitra et al. 2011), through direct deaths and the reduction or elimination of habitats and food sources, including decreased levels of cereal grains, weed seeds, arthropods etc. Birds are particularly susceptible to cholinesterase-inhibiting pesticides mostly because, unlike mammals, they have low levels of anticholinesterase detoxifying enzymes (Walker 1983). The effects of pesticides on earthworms (Yasmin and D’Souza 2010), microarthropods (Adamski et  al. 2009), nematodes (Zhao et  al. 2013), fungi (Morjan et al. 2002) and microorganisms (viruses, protozoa and bacteria) (Lo 2010; Imfeld and Vuilleumier 2012) within the soil may also have major environmental consequences.

1.7  Effects on Beneficial Arthropods For the past 30 years, the effects of pesticides on beneficial arthropods have been the subject of an increasing number of studies. Insecticide treatments aiming at pest control also have damaging effects on many non-target terrestrial arthropods

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present in agroecosystems, including the natural enemies of agricultural insect pests (Croft and Brown 1975). Damage to these species may be greater than initially thought, because such damage can occur even at sublethal insecticide doses (Desneux et al. 2007). A sublethal dose or concentration is defined as inducing no apparent mortality in the experimental population. Experiments on bee physiology have been done by measuring the activity of enzymes after or during exposure to pesticides. Na+/K+ ATPase is a transmembrane enzyme that releases energy necessary for cell metabolism. Bendahou et al. (1999) reported that organophosphorus and pyrethroid led to a decrease in Na+/K+ ATPase and AChE activities. Thus, the inhibition of Na+/K+ exchange provoked by insecticides might affect a wide range of cellular functions. Malformations also occur in natural insect enemies after exposure to pesticides and may lead to reduction in predator or parasitoid efficiency and fitness. Deltamethrin, a pyrethroid, causes marked dysfunctions in myocardial bee cells (Papaefthimiou and Theophilidis 2001). Insect growth regulators (IGRs) that disrupt molting and, more generally, act on endocrine systems (Dhadialla et al. 1998) are also likely to perturb the development of beneficial arthropods. Malformation of ovaries in the parasitoid Hyposoter didymator Thunberg (Hymenoptera: Ichneumonidae) exposed to IGRs was reported by Schneider et  al. (2004). Hymenopteran social pollinators oral exposure with diflubenzuron, an IGR, reduced brood surface area (Chandel and Gupta 1992), weight gain and suppressed development of hypopharyngeal glands (Gupta and Chandel 1995). Insecticides can also interact with the immune capacity of insects. Depending on the type of insecticide, they can decrease or increase this capacity. Monocrotophos and methyl parathion applied at 10% of the LC50 decreased the number of plasmatocytes in the hemolymph of the predator Rhynocoris kumarii Ambrose and Livingstone (Hemiptera: Reduviidae) by 16% and 13%, respectively, whereas endosulfan increased these cells by 15% (George and Ambrose 2004). Therefore, insecticides may have an impact on both the immune capacity of a host and the capacity of parasitoids to evade the host immune reaction. Studies reporting pesticide impacts on the developmental period of natural enemies typically differ with the biology of the experimental subject and also by gender. Zanuncio et al. (2003) reported that exposure of the pentatomid predator Supputius cincticeps Stål (Heteroptera: Pentatomidae) to permethrin decreased development time for females, whereas this time increased for males. Reductions in fecundity associated with pesticides may be due to both physiological and behavioral effects.

1.8  Pesticides Degradation Microbial degradation of pesticides applied to soil is the main mechanism which prevents the accumulation of these chemicals in the environment. Microbial adaptation to pesticide degradation was recognized soon after their introduction into

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markets, with the pioneer research on biodegradation of 2,4-D (Audus 1949). In a recent study, Paul et  al. (2013) noticed that Azotobacter chroococcum strains degraded more than 90% of lindane, an organochlorine, under laboratory conditions. When these degraded compounds were tested against lepidopteran larvae, a lower mortality was recorded against the very high mortality induced at all concentrations by non-degraded lindane. One key factor associated to increased rates of microbial pesticide degradation in soil is the history (number) of previous applications of the same pesticide, or of other compounds, having a similar molecular structure. This phenomenon is known as “accelerated” or “enhanced degradation” (Racke 1990). The list of pesticides affected by accelerated degradation is long and constantly growing. In a review, Arbeli and Fuentes (2007) summarized the accelerated degradation processes, providing a list of important susceptible pesticides. One way to cope with the problem of reduced pesticide efficacy due to enhanced microbial degradation would be to increase the amount and/or frequency of applications, which may further generate other ecosystem problems. Thus, the increased difficulty of pest control clearly indicates the need for more sophisticated pest management, as the paradigm of relying almost exclusively on chemicals for pest control may need to be reconsidered. Insecticides and nematicides known to undergo accelerated degradation / loss of efficacy are listed as: Aldicarb, Bendiocarb, Benfuracarb, Cadusafos, Carbaryl, Carbofuran, Carbosulfan, Chlorfenvinphos, Chlorpropham, Chlorpyrifos, Cloethocarb, Diazinon, Ethoprophos, Fenamiphos, Fensulfothion, Fonofos, Furathiocarb, Isazofos, Isofenphos, Malathion, Mephosfolan, Methylparathion, Oxamyl, Parathion, Phorate, Terbufos, Trimethacarb (Arbeli and Fuentes 2007).

1.9  Conclusion Adverse effects of pesticides are wide and varied, as they hold a unique position among environmental contaminants. Due to unregulated use and lack of enforcement mechanisms, health hazards caused by pesticides are serious, especially in developing countries, and no groups in the human population are completely unexposed to pesticides or related residues. For almost all major insecticide classes, resistance has been reported in one or more key insect pest species. The complex patterns of cross resistance between chemical groups and within groups further complicate pest management strategies with chemicals. Direct and indirect adverse effects of pesticides on several non-­ target wild vertebrates are also widely documented. To cope with the problem, instead of relying almost exclusively on chemicals, eco-friendly alternative methods for insect pest management must be considered.

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Chapter 2

Soil-Borne Entomopathogenic Bacteria and Fungi Tan Li Peng, Samsuddin Ahmad Syazwan, and Seng Hua Lee

Abstract  Being rich in microorganisms, the soil is an ideal environment and important reservoir for harvesting various types of beneficial microorganisms. Soil-­borne entomopathogenic bacteria and fungi have been regularly isolated around the world to support crop producer in the never-ending arms race of pest management. Among these microorganisms, entomopathogenic bacteria and their toxins are the most successful microbial insecticides also from the commercial point of view. They grouped into spore- and non-spore-forming entomopathogens, in which the infection process starts upon ingestion by the susceptible insect hosts. Fungi, on the other hand, remain relatively underutilized as natural enemies despite their many advantages over other biological and chemical products. They mainly classified under the class of Entomophthoromycetes and Sordariomycetes in the larger Ascomycota division, which consists around 65,000 described species. In comparison to bacteria, fungi have a wider host range and are especially suitable for controlling pests with piercing and sucking mouthparts. Entomopathogenic bacteria and fungi can be released through inundative application methods and therefore play a critical role in integrated pest management (IPM) against several pests. This chapter provides a selective review on the different types of soil-borne entomopathogenic bacteria and fungi, including their distribution, infection mechanisms and host ranges.

T. L. Peng (*) Department of Paraclinical, Faculty of Veterinary Medicine, Universiti Malaysia Kelantan, Kota Bharu, Kelantan, Malaysia e-mail: [email protected] S. A. Syazwan Department of Forest Management, Faculty of Forestry, Universiti Putra Malaysia, Serdang, Selangor, Malaysia Mycology & Pathology Branch, Forest Biodiversity Division, Forest Research Institute Malaysia, Kepong, Selangor, Malaysia S. H. Lee Institute of Tropical Forestry and Forest Products, Universiti Putra Malaysia, Serdang, Selangor, Malaysia © Springer Nature Switzerland AG 2019 M. A. Khan, W. Ahmad (eds.), Microbes for Sustainable Insect Pest Management, Sustainability in Plant and Crop Protection, https://doi.org/10.1007/978-3-030-23045-6_2

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Keywords  Soil-borne · Entomopathogens · Bacteria · Fungi · Bio-insecticides

2.1  Introduction Soil, the unconsolidated mineral or organic material on the immediate surface of Earth, is the natural medium for the growth of land plants. It is also a natural body comprised of solids, liquid, and gases that harbours a complex, living and unique community on its own. This soil microbial community plays several important ecological and physiological functions (Sofo et al. 2014). In agroecosystems, the soil microbial communities are essential for plant nutrition and health (Gattinger et al. 2008). Soil microbial community in a narrow sense consists of viruses and organisms such bacteria, fungi, algae and protozoa (Sims 1990), that support a food web formed by nematodes and other micro-arthropods (Jones et al. 2010). Soil microorganisms in general maintain balances within soil through various roles, acting as e.g. decomposers, nitrogen binders, pathogens, etc. (Waldrop et  al. 2000; Glick 2010). They contribute to ensure soil health by sustaining its structure and composition, in turn sustaining plants’ growth (Jacoby et al. 2017). Even more interesting, some of these organisms are able to protect crops and plants by regulating harmful pests such insects and other arthropods (AGP – FAO 2018). Microorganisms that are able to cause a disease on an insect pest and, in some extent on other arthropods, are known as entomopathogens (Tanzini et al. 2001). They are natural enemies that kill or debilitate pests, keeping their populations under control (Lacey et al. 2015). Entomopathogens also represent a large component of the world’s biodiversity. There are for example about 750–1000 species of fungi being causal agents of insect diseases (St. Leger and Wang 2010). Although extensive studies and researches have been performed on entomopathogens, either as causal agents in a natural condition or in exploitation for biological control (Davidson 2012), only a small proportion of available species has been studied in depth for crop protection, and subsequently commercialized (Lacey et al. 2015). Among the mentioned soil microorganisms, bacteria and soil fungi are the most abundant (Beed 2011). Therefore, much interests and efforts have been invested in these two major groups in biological pest control. As for bacteria, also the proportion of fungi known to infect insects and several other arthropods is large. Entomopathogenic fungi can be taxonomically divided into 6 phyla: Chytridiomycota, Blastocladiomycota, Kickxellomycotina, Basidiomycota, Ascomycota and Entomophthoromycota (previously a subphylum under Zygomycota). Among these 6 phyla, substantial numbers of insect pathogenic species are found in Ascomycota and Entomopthoromycota (Mora et al. 2017). The two most important orders are the Entomophthorales (Entomopthoromycotina: Entomophthoromycetes) and the Hypocreales (Pezizomycotina: Sordariomycetes).

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Detailed life-cycles of entomopathogenic fungus belonging to Entomophthorales and Hypocreales have been described (Augustyniuk-Kram and Kram 2012). Slight differences are present between these orders, mostly considering two stages: (i) proliferation, whereby Hypocreales species proliferate inside the host body in the form of yeast-like hyphal bodies, multiplying by budding (Prasertphon and Tanada 1968), whereas members of Entomophthorales produce protoplasts (Butt et  al. 1996); and (ii) sporulation, whereby Hypocreales produce only asexual spores or sexual spores after host death (known as hemibiotrophic), whereas Entomophthorales can produce both asexual and sexual spores before host death (termed biotrophic) (Roy et al. 2006). Entomopathogenic fungi are widespread throughout the world, ranging from Antarctic to the Arctic Circle (Bridge et al. 2005; Eilenberg et al. 2007), reaching highest abundance at the equator level (Aung et al. 2008)(. Different habitats are known to be occupied by different group of entomopathogenic fungi, the distribution of these fungi being very much influenced by both biotic and abiotic factors. For instance, Hypocreales dominate soil layers, whereas Entomophthorales are mostly found at the arboreal level (Vandenberg and Soper 1978; Sosnowska et al. 2004). These differences could be mainly due to the presence of hosts at different habitat niches and to abiotic factors that affect the transmission of fungi. Various aspects collectively contribute to the “success” of an entomopathogenic fungus. An advantage, however, does not necessarily imply that a specific fungus is the best candidate for biological control. Hypocreales, for example, have a vast spectrum of potential victims but may not be suitable when beneficial insects (i.e., parasitoids) occur, that might also be infected by these fungi (Augustyniuk-Kram and Kram 2012). The relatively host-specific and obligate Entomophthorales, on the other hand, might not be sustainable in the environment after their host has been being wiped out and a continuous augmentation effort might be needed, is not cost-­ effective (Pell et al. 2001). Regardless of their systematics and even biology, these fungi and bacteria are all pathogenic, and their effectiveness in infecting their hosts is so significant that they can become a crucial factor regulating the population of many insects. Nevertheless, the success of their use requires a thorough knowledge about the biology and ecology of both fungi and hosts. It also requires recognition of abiotic and biotic factors that may interfere with their effectiveness. Biological control, rather than entirely replacing chemical products, is intended to reduce their use through their integration with other controlling measures to keep pest populations at a suitable, low damage level. Soil-borne entomopathogenic bacteria and fungi regularly isolated around the world for pest management (Bruck 2004). The use of entomopathogenic bacteria and fungi in pest management systems has a long and rich history. Despite a variety of obstructions, there are plenty of opportunities and benefits for the use of natural enemies in insect pest management. In general, both entomopathogenic bacteria and fungi play a critical role in integrated pest management (IPM) due to environmental and human safety concerns, development in pests of resistance to insecticides, increases in pesticides costs, etc. (Gangwar 2017). This chapter provides a selective

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review on the different types of soil-borne entomopathogenic bacteria and fungi, including their distribution, the primary infection mechanisms and their host ranges.

2.2  Soil-Borne Entomopathogenic Bacteria Most of the discovered entomopathogenic bacteria are soil-borne aoecies such as Brevibacillus laterosporus (Oliveira et  al. 2004; Ruiu et  al. 2007), Lysinibacillus sphaericus (Persinoti et  al. 2018), Clostridium bifermentans (Leja et  al. 2011), Serratia entomophila (Villalobos et  al. 1997), and numerous other entomopathogens. Bacteria are one of the most diversified microbiota existing in soil (Riesenfeld et al. 2004). Soil biota refers to organisms living in the ground, interacting within its environment, and involved in many ecological processes (Ritz et  al. 2004). Entomopathogenic bacteria provided the most common biological control agents for the management of insect pest populations in plantations. There are two main groups of entomopathogenic bacteria classified by their abilities to sporulate, either obligately or facultatively. Meanwhile, the facultative-sporulating group also divided into two groups known as crystalliferous, with the ability to form crystalline toxins, or non-crystalliferous species. More than a century ago, the discovery of entomopathogenic bacteria began with the first isolation from diseased silkworm larvae, Bombyx mori (Ishiwata 1901). The curiosity to examine a diseased silkworm and describing it as the sotto disease led to the first discovery of entomopathogenic bacteria in the world (Ishiwata 1901). The study continued by describing the morphology of isolated Bacillus spp. from Mediterranean flour moth, Anagasta kuehniella, named as Bacillus thuringiensis; in conjunction with the name of the city in which the moth found, Thuringia, in Germany (Berliner 1915). The discovery of Bacillus thuringiensis, also known as Bt, kick-started the biological pesticide discovery and subsequent developments, with applications depending on the technology availability, the industry demand and the raising awareness towards the impact of pesticides on environment and climate.

2.2.1  Isolation Even though almost all of the studied entomopathogenic bacteria are soil-borne species, few new species of entomopathogenic bacteria directly discovered from soil samples. Most findings proceeded either from diseased insects or insects’ cadavers. Despite this issue, several studies involved the isolation of soil-borne entomopathogenic bacteria, from various sampled places and soil conditions. Bacillus thuringiensis, Paenibacillus popilliae and Paenibacillus lentimorbus discovered from diseased insect larvae (Ishiwata 1901; Berliner 1915; Dutky 1940). Chromobacterium substugae, capable of  killing the Colorado potato beetle larvae, Leptinotarsa

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decemlineata was directly obtained from forest soil samples in central Maryland, USA (Martin et al. 2007). Several protocols were applied, which resulted in many soil-borne entomopathogenic bacteria isolated from soil samples. By using 1 g of soil, different dilution techniques, and treatments applied before streaking on to media. Based on Ernandes and Da Rosa experiment, the soil sample was dispersed into 10 mL saline solution, serial diluted, heated for 12 min at 80 °C and kept on ice for 5 min before streaked on nutrient agar medium (Ernandes and Da Rosa 2014). Meanwhile, González cultured 1 g of soil in nutrient broth, incubated for 24 h at 30 °C, shook at 150 rpm before seeding on nutrient agar (González et  al. 2013). Several techniques can also be applied to confirm the isolated bacteria species, including traditional Gram staining, phenotypic description, determination of respiratory type, colony and culture morphology and also biochemical tests, that may aid in the validation of the isolates species.

2.2.2  Mechanism of Action The insecticidal mechanism of Bacillus thuringiensis is well known by the scientific community. Over a century of works and studies focusing on this remarkable species yielded fruitful informations on each stage infecting the insects host. By noticing  the occurrence of a second body in the sporangium from the Steinhaus publication in 1951, also present in the Berliner previously published  the image, Hanney hypothesized that the body inidcated as a “parasporal crystal” had a major role in pathogenesis (Berliner 1915; Steinhaus 1951; Hanney 1953). The production of parasporal crystals, also known as δ-endotoxin (Cry proteins), within the Bt sporangium followed the sporogenesis time course, being approximately completed within 24 h in typical bacterial media (Federici et al. 1990). Brevibacillus laterosporus also produced similar parasporal crystals which also have an insecticidal activity on the infected hosts (Zubasheva et al. 2010). Variation in parasporal crystals produced by different B. laterosporus strains in different hosts were also discovered. Different insecticidal roles of excreted toxins were shown in the infection of various species of Coleopteran hosts. The insecticidal crystal is activated once ingested into the insect digestive system, harming its gut membranes by a pore-forming mechanism (Pigott and Ellar 2007). The alkaline condition of insect midgut solubilizes the ingested crystal proteins that are processed into the active form, due to reactions with midgut-excreted proteases (Lee et  al. 2003). Subsequently, activated parasporal proteins(δ-­ endotoxins) attach to the receptor sites on the gut epithelium causing the formation of pores (Martin et  al. 2007; Chilcott et  al. 1983). This process will imbalance osmotic regulations thus leading to a midgut paralysis and finally cell lysis. Rupture of insects midgut will produce a solution leakeage within the insects cavity, imbalancing the hemocoel pH level and causing a septicaemia, leading the infected insects to death.

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The specificity of entomopathogenic bacteria towards hosts depends on the presence of a corresponding cellular receptor of the crystal proteins in the host of interest. The suitable cellular receptors for the δ-endotoxin to bind and activate are vary and show a high affinity binding site, affecting host specificity and selection of entomopathogenic bacteria species (Honée and Visser 1993). In conclusion, several criteria should be taken to ensure the success of entomopathogenic bacteria application on insects pests, which are: (1) optimal condition of insects midgut (pH of midgut juice), (2) presence of suitable cellular binding receptors for the δ-endotoxin activation on infected gut epithelium, and (3) production of effective proteases digesting the activated δ-endotoxin.

2.3  Soil-Borne Entomopathogenic Fungi Back to 1888, the Russian microbiologist Elie Metchnikoff was the first to identify an entomopathogenic fungus from a wheat cockchafers. Krassilstick  then mass-­ produced the same fungus and used in the field against the sugar-beet weevil (Vega et al. 2009). His work after that ignited curiosity around the world for experimentation with “friendly fungi” against insect pests (Lord 2005), and searching for potential natural enemies towards insect pests was springing since then, until now. There are different groups of entomopathogenic fungi in various habitats, among which soil is the environment where these fungi may be most commonly found (Keller and Zimmerman 1989). Entomopathogenic fungi are a group of phylogenetically diverse and heterotrophic microorganisms. They are taxonomically classified into the six previously mentioned phyla (Mora et al. 2017) although some researchers group them into five phyla namely Chytridiomycota, Mycosporidia, Basidiomycota, Ascomycota and Entomophthoromycota (Araújo and Hughes 2016). Ascomycota and Entomophthoromycota are the two phyla with the most important groups of entomopathogenic fungi due to their substantial numbers (Maina et al. 2018) and thus focused on in this chapter. The occurrence and distribution of entomopathogenic fungi vary and mainly depend on the behavioural patterns of their hosts. Although the emphasis will be given to soil-borne Entomophthorales and Hypocreales, the occurrence of these fungi in aquatic and arboreal habitats are equally crucial as within these habitats they contribute to the regulation of arthropod populations. Entomophthorales, previously under Zygomycetes, are one of the lower fungi predominantly known as associated with aquatic habitats. This clade shows remarkable adaptations in which up to four spore types can be produced by the same species to facilitate dispersal and infection, both aerially and aquatically (Descals and Webster 1984). Both Entomophthorales and Hypocreales are abundant in forest habitats, including untouched humid, tropical forests or man-managed coniferous forests. They are especially ubiquitous at the understorey and below (Fig. 2.1), where the forest canopy buffers these layers from extremes of temperature and humidity, creating a

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Fig. 2.1  Diagram of the forest layer showing entomopathogenic fungi evolutionary diversity in forest habitats (Samson et  al. 1988). Although both Entomophthorales and Hypocreales found under the canopy layer, they dominate and develop succession at different layers, where forest soil litter and surface are dominated by Hypocreales, whereas understory trees and canopy mostly show Entomophthorales. (Sosnowska et al. 2004)

stable microclimate conducive to continual fungal activity, even during extreme seasons (Samson et al. 1988).

2.3.1  Entomophthorales (Entomopthoromycotina: Entomophthoromycetes) Entomophthorales (Schröter, 1893) is an order now treated under Entomophthoromycota, containing soil saprotrophic, opportunistic, broad-­ spectrum, or specialized insect pathogenic genera (Humber 2012). There are approximately 280 species in the phylum Entomophthoromycota, classified within three classes and six families (Humber 2012; Gryganskyi et  al. 2013). Entomophthorales is the only order in class Entomophthoromycetes and consists of approximately 17 genera classified under four families (Humber 2012). This order is notable for the epizootics they induce in populations of many insect orders, including Hemiptera, Homoptera, Lepidoptera, Coleoptera, Orthoptera  and Diptera.

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The modes of formation and germination of resting spores and the nature of vegetative growth and development are used to separate families (Humber 1989). The classification of genera in Entomophthorales, at present, follows the three systems proposed by Humber (1989), Keller (1991) and Balazy (1993). They classified according to: (1) morphology of primary and secondary conidia, conidiophores  and/or conidiogenous cells; (2) number of nuclei per conidium; (3) mode of discharge and formation; (4) pathobiology (e.g., host symptoms). Further down, the details of the primary conidia (size and shape); the number of nuclei per conidium and the basis of host insect species in the species identification (Balazy 1993). Entomophthorales have conidia and resting spores at different stages of life-­ cycle, divided into: (1) conidial cycle and (2) resting spore cycle (Fig.  2.2). The former allows the propagation and spread of the disease, the latter provides to the

Fig. 2.2  General scheme of the Entomophthorales life-cycle. (a): conidiophores within infected host grow and emerge out from the cadaver; finger shaped conidiophores show one apical spore, which is then forcibly ejected to drift on winds until they attach to another fly and start the cycle again. (b): infected host is compelled to find soil to land on; its abdomen becomes brittle and eventually falling apart to release the resting spores into the soil; the spores remain dormant throughout unfavourable conditions (winter) and attach to emerging hosts in favourable conditions (early spring). (c): host are infected by conidia drifted on winds, or by direct contact with cadavers (asexual), or by spores when in contact with soil where the spores resided. (d): infection starts with the conidia, asexual, non-motile fungal spores, attaching to a fly; the primary conidia then pierce through the exoskeleton with germ tubes; one long germ tube arises from the secondary conidium to branch out inside the insect hemocoel; hyphae then grow and start consuming the hemolymph and internal organs. (e): resting spores attached to the host may start an infection in resting spores cycle, as in the conidial cycle. (Roy et al. 2006)

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survival during unfavourable conditions. Both cycles are supposed to start in the same way, in which the conidia or spores adhere to the host cuticle and form a penetration tube. Unlike conidia, it is unknown at which stage resting spore formation is induced and if there are changes at the nuclear level. Resting spores are described as azygospores, however, their mode of formation has never been observed in detail. The infection mechanism of entomopathogenic fungi is generally the same. For a successful infection, the entomopathogenic fungus must first able to penetrate the insect cuticle. In Entomophthorales, the conidia can easily achieve their adhesion process by having A larger size and extra cell wall layers or mucus, that enable them to attach firmly to the host cuticle (Eilenberg et al. 1986). Conidia then germinate and produce germ tubes that penetrate through the pores or the layers of the epicuticle, procuticle and epidermal via mechanical pressure and production of cuticle-­ degrading enzymes (Vega et al. 2012). After reaching the hemocoel, the multiplication takes place either by protoplasts or by hyphae. These structures colonise the abdomen or, more commonly, the whole body of the host. Most host species die at this stage, as a result of a combination of effects that comprise physical damage of tissues, toxicity, cells dehydration due to loss of fluids, and consumption of nutrients. Many genera in this order contain obligate insect-pathogenic species, characteristically biotrophic with a narrow host range, capable of producing epizootics in natural insect populations and common among foliar insects (Pell et al. 2001). For instance, if conditions are right, one Entomophthorales species can cause epizootics of flies, with a 70–90% prevalence, killing the majority of flies. This makes members of this lineage as good candidates for biological control of multiple fly populations. Some important genera and their corresponding host range are summarized in Table 2.1.

2.3.2  Hypocreales (Pezizomycotina: Sordariomycetes) The order Hypocreales (Lindau, 1897) classified in Sordariomycetes, one of the largest classes in the Ascomycota with more than 600 genera and 3000 known species (Kirk et al. 2008). This order consists of biotrophic  fungi: they are mainly plant and insect pathogens, mycoparasites, and endophytes, as well as saprotrophic species (Sung et al. 2008; Zhang et al. 2006). Many important entomopathogenic fungi that have been developed into commercial mycoinsecticides belong to this order, including species of genera Lecanicillium, Isaria, Beauveria, Metarhizium, Pochonia, Cordyceps and others. Most of these species may be cultured artificially, thus are relatively easy for mass production. Although efforts in developing them as inundative biological control agents cannot be obliterated, in biocontrol strategies understanding of the ecology of entomopathogenic fungi are equally essential to achieve sustainable effects (Rai et al. 2014). Pezizomycotina is the most numerous, morphologically, and ecologically complex ascomycetes (Schoch et  al. 2009). Members of the Hypocreales, without exemption, are morphologically diverse with complex life cycles often with two

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Table 2.1 Most studied Entomophthorales (including main lineages in the family Entomophthoraceae) and associated hosts Genus Batkoa

Conidiobolusa

Entomophaga Entomophthora

Host Homoptera (aphids) Hemiptera Coleoptera Diptera Lepidoptera Aphids, other Homoptera Other insects (but weak pathogen) Lepidoptera Othoptera Muscoid flies,other Diptera

Aphids, other Homoptera Aphids, other Homoptera Diptera Lepidoptera Trichoptera Eryniopsis Coleoptera (Cantharidae) Diptera (Nematocera) Furia Diptera (Calliphoridae; Rhagionidae) Lepidoptera Macrobiotophthoraa Nematodes Massospora Cicadidae (Magicicada spp.) Erynia

Meristacruma Orthomyces Pandora

Strongwellsea Zoophthora

Nematodes Homoptera (Aleyrodidae) Homoptera Sacrophagid flies, other Diptera Plutella xylostella, other Lepidoptera Umbonia sp. Acraea sp. Aphids, other Homoptera Weevils, other Coeloptera

References Humber (2005)

Humber (2005)

Humber (2005) Humber (2005)

Li and Humber (1984), Steinkraus and Kramer (1989), and Capinera (2008)

Prischepa et al. (2011) Steinkraus et al. (2017) Samson and Nigg (1992) Prischepa et al. (2011) Bernard and Arroyo (1990) Cooley et al. (2018) Gryganskyi et al. (2013) Steinkraus et al. (1998) Humber (2005)

Samson et al. (1988) and Chien and Hwang (1997) Humber (2005) Capinera (2008)

a Taxa under other families with biocontrol importance: Macrobiotophthora, Conidiobolus = Ancylistaceae; Meristacrum = Meristacraceae (Humber 2012)

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stages, anamorph (asexual stage) and teleomorph (sexual stage), often leading to the identification of the same species with two different taxonomic names, one per stage. However, in several cases, the sexual stage of some Hypocreales may never or only rarely be produced, and anamorphs are used for taxonomic purposes (Taylor 2011; Mora et  al. 2017). Besides, sequences from the nuclear small subunit ­ribosomal RNA gene (nSSU rDNA) or other genes are also employed to solve the complexity in classifying this order, to  adopt the principle of “one fungus, one name” (Zhang et al. 2006; Hawksworth 2011). The Hypocreales once contained more than 70 genera in four major families (Rossman et al. 1999). Subsequent estimates showed approximately 237 genera and 2647 species, in seven families (Kirk et  al. 2008). Among this order, the former unique family Clavicipitaceae has split into three monophyletic families: Clavicipitaceae, Cordycipitaceae, and Ophiocordycipitaceae, including the most well-known entomopathogenic fungi. These three families are generally distinguished based on the host infected. For instance, spider pathogens are mostly found within the Cordycipitaceae, while Clavicipitaceae are common in scale. Pathogens of ants, termites, or dipterans are often found in the Ophiocordycipitaceae (Sung et  al. 2007). Other insect orders such as Hemiptera, Coleoptera, Lepidoptera, Thysanoptera, and Orthoptera also comprise most of the targets. The sexual (teleomorphic) stages of Hypocreales appear to be more host specialized, while their asexual (anamorphic) counterparts appear as more generalist. Among these entomopathogenic fungi, asexual stages often precede the production of sexual stages (Sung et al. 2007). Many Hypocreales have a well-developed parasitic phase that infects the host’s body. Taking the most commercialized species Beauveria bassiana as an example, it is the anamorph of Cordyceps bassiana (Li et al. 2001). The infection pathway of this species begins with attachment of conidia to the insect cuticle, followed by germination and penetration through the cuticle and proliferation within the host. This fungus is then switching to a saprotrophic nutrition (hemibiotrophic), by colonizing the cadaver to maintain hyphal growth and produce new conidia, even after the host’s death (Evans 1988). Beauveria bassiana is well characterized for infecting several insects. It is also recorded with variable strains which differ substantially in their ability to produce pathogenesis. Cordyceps bassiana, on the other hand, shows a more specialized behaviour (Bhushan et al. 2016). It is, perhaps, the most studied sexual stage in association with arthropods. The host range of Cordyceps, in a classical sense, is very broad and includes several orders such as Hymenoptera, Hemiptera, Orthoptera, Diptera, Blattodea, Mantodea, Dermaptera, Odonata, Phasmatodea etc. Coleoptera and Lepidoptera are the major host orders in which nearly 60% of Cordyceps species are recorded. Cordyceps are not monophyletic, as teleomorph stages occur in all the three families mentioned above. It is an intriguing genus considering the same fungal species differ in their ecology, depending on the reproduction stage (Hesketh et al. 2009). Entomophthorales and Hypocreales are the two main orders of entomopathogenic fungi distributed in wide terrestrial ecosystems. They are playing vital roles in regulating the insect density, balancing the ecosystem in natural conditions, and

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Table 2.2  Differences between Entomophthorales and Hypocreales Entomophthorales Narrow host range (Burges 1981; Pell et al. 2001) Dominantly distributed in temperate forests (Evans 1982) Biotrophic (Charnley 2003; Charnley and Collins 2007) Penetrate directly using germ tubes (Hajek and Delalibera 2010) Proliferation trough protoplasts (Boomsma et al. 2014) Toxin production rarely found / not known (Boguś and Scheller 2002; Freimoser et al. 2003a) Transmission Forcibly discharged: asexual conidia Passively released: sexual resting spores (Boomsma et al. 2014)

Hypocreales Narrow host range (Teleomorph) to Broad host range (Anamorph) (Vega et al. 2012) Dominantly distributed in tropical forests (Evans 1982) Biotrophic Hemibiotrophic (biotrophic + saprotrophic) (Charnley and Collins 2007) Penetrate using appressoria (Shah and Pell 2003; Sandhu et al. 2012) Proliferation trough hyphae (Boomsma et al. 2014) The toxin produced (Freimoser et al. 2003b; Sandhu et al. 2012) Transmission Forcibly discharged: sexual ascospores Passively released: asexual conidia (Boomsma et al. 2014)

reducing crop damage in many crops. They share some characteristics in common, yet showing uniqueness in many other ways (Table 2.2).

2.4  Current Status as Biopesticides Controlling the insect pest population is always a worldwide priority, with many industries and areas of study directly or indirectly connected and responding to this need. The increasing of the insect pest population is often due to poor agricultural practices, affecting crop yields, with severe impacts on the economy and food security issues.  Also, awareness towards environmental conservation is rising worldwide, and the application of highly toxic pesticides forbidden in many countries. Therefore, biopesticides emerge as the times required. Sporleder and Lacey (2013) stated that differing from conventional chemical pesticides; biopesticides are biocontrol agents-containing products used to control pest through specific biological effects. Biopesticide products potentially can be categorized into microbial insecticides, Plant-Incorporated Protectants (PIPs), and biochemical pesticides. Among these three classes, microbial pesticides contain microorganisms such as bacteria, fungi, or virus that kill insects and weeds. This section focuses on the application of soil-borne entomopathogenic bacteria and fungi as biopesticides. Using entomopathogenic bacteria and fungi as biopesticides offer some advantages such as: (1) they are more environment-friendly compared to chemical pesticide; (2) they are unharmful to mammals and other non-target organisms; and (3) they allow a prolonged pest control owing to their higher persistence (Singh et al. 2017). Nevertheless, some drawbacks have also identified regarding biopesticides

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Table 2.3  Example of commercially available biopesticides derived from entomopathogenic fungi and bacteria (Chandler et al. 2011; Mishra et al. 2015; Singh et al. 2017) Microorganism Fungi Beauveria bassiana

Product name

Target

Mycotrol®

Culicinomyces clavisporus Coniothyrium minitans Chondrostereum purpureum Paecilomyces lilacinus Aureobasidium pullulans Bacteria Bacillus thuringiensis var kurstaki Bacillus subtilis QST713 Pasteuria usgae

Mycar Contans WG Chontrol MeloCon WG Blossom Protect

Coding moth, pine caterpillar, European corn borer Mosquito larvae Sclerotinia spp. Cut stumps of hardwood trees and shrubs Plant parasitic nematodes in the soil Fire blight, postharvest diseases

Dipel DF

Caterpillars

Serenade ASO Pasteuria usgae BL1 Galltrol – A

Botrytis spp. Sting nematode (Belonolaimus longicaudatus) Crown gall disease

BlightBan C9-1

Fire blight

Agrobacterium radiobacter k84 Pantoea agglomerans C9-1

including: (1) longer time is required to observe their effects on targeted hosts, compared to chemical pesticides; (2) higher costs; (3) different control measures are needed, due to their high degree of host specificity; (4) a frequent treatments are necessary to maintain a long-term impact (Singh et al. 2017). Many biopesticides, mostly based on bacteria and fungi, have already been formulated and commercially manufactured to tend the global market. According to Kabaluk et  al. (2010), the whole biopesticide market accounts for 60% bacterial biopesticides, followed by fungal (27%), viral (10%) with a remaining 3% as “other” biopesticides. The most common bacterial insecticides include Bacillus species, with B. thuringiensis (Bt) as the most widely used (Mishra et  al. 2015; Khan et al. 2016). Table 2.3 lists some examples of commercially available biopesticides derived from entomopathogenic fungi and bacteria.

2.5  Conclusion Soil is rich in microorganisms and serves as an ideal reservoir for harvesting various types of beneficial species of entomopathogenic bacteria and fungi. Many biopesticides derived from these microorganisms are used in pest management. Many cutting-­edge innovations and developments in the industry enhanced the effectiveness and applicability of bacteria and fungi as a biological control agent. They may complement chemical pesticide applications to build up optimal protocols for insect pest management. Discovery of most adapted biological control agents such as

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entomopathogenic bacteria and fungi is one of the steps required for sustainable IPM practices. They may reduce the environmental impact of pests by decreasing the amount of chemical pesticide usage in the plantation industry. This concept indirectly will reduce carbon footprint and thus helps in combating climate change. Unfortunately, despite the wide variety of commercialized products, the usage of biopesticides worldwide is still not encouraging. The lack of awareness among farmers is one of the main constraints that limits the prevalence of biopesticides. Application of chemicals is highly price-dependent, as conventional chemical pesticides lower price and longer shelter life appear more attractive. Practically, being often highly specific, biopesticides are not as effective or generalist as chemical pesticides and occasionally, they may perform inconsistently in the field, inducing farmer to lose trust in their efficacy. Therefore, the intervention of government is of utmost important to achieve a balance between the two approaches, through the definition of more stringent rules on the use and impact of conventional chemical pesticides. Research and development of more effective and cheaper biopesticides is also vital in order to promote and encourage people to adopt safer IPM approaches.

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Chapter 3

Molecular Phylogeny of Entomopathogens Mudasir Gani, Taskeena Hassan, Pawan Saini, Rakesh Kumar Gupta, and Kamlesh Bali

Abstract  Insects, like other organisms, are susceptible to a variety of diseases caused by bacteria, viruses, fungi, protozoan and nematodes. Insect pathogens show a wide variety of interactions with their hosts that facilitate their replication and transmission, including strategies for evading the host’s defences towards invasion of microorganisms, and for manipulating their hosts physiology and behaviour. By applying a wide range of molecular techniques and approaches, better understandings of these interactions and of the roles played by both host and virulent genes have been understood. The control of insect pests with entomopathogens is an unique approach, in that naturally occurring host-pathogen relations are manipulated to the benefit of man, protecting agricultural crops and forests or controlling insect vectors of diseases. The isolation and identification of a pathogen followed by the phylogenetic classification of entomopathogens are the basic principles in insect pathology. Full genomic DNA sequencing techniques are used to assess the genetic diversity and phylogenetic analyses of entomopathogens. Alternatively, specific genes of interest can be targeted for sequencing. Sequences of single gene have been extensively used to assess phylogenetic relationships of known and novel isolates. However, the lack of sufficient resolution and disagreement with other gene phylogenies has prompted investigation of other genes and methods to further explore evolutionary relationships of entomopathogens. Therefore, phylogenies based on combined sequences of shared genes or the complete genome sequencing have been found to be more robust, providing more phylogenetic information and increasing robustness of evoM. Gani (*) · P. Saini Central Sericultural Research & Training Institute, Central Silk Board, Ministry of Textiles, Govt. of India, Kashmir, Jammu and Kashmir, India e-mail: [email protected] T. Hassan Department of Zoology, Aligarh Muslim University, Aligarh, Uttar Pradesh, India R. K. Gupta · K. Bali Division of Entomology, Sher-e-Kashmir University of Agricultural Sciences and Technology, Jammu, Jammu and Kashmir, India © Springer Nature Switzerland AG 2019 M. A. Khan, W. Ahmad (eds.), Microbes for Sustainable Insect Pest Management, Sustainability in Plant and Crop Protection, https://doi.org/10.1007/978-3-030-23045-6_3

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lutionary hypotheses. With the increasing efficiency and lower cost of whole genome sequencing, whole-genome studies of entomopathogens will refine knowledge about their evolutionary history and enable direct insight into the biology of entomopathogenesis. Keywords  Entomopathogen · Evolution · Genomics · Phylogeny · Taxonomy

3.1  Introduction Insects constitute the most diverse group of living organisms in the world, with more than one million species. They live almost everywhere, from steamy tropical jungles to cold polar regions. The microorganisms that are capable of causing diseases in insects are known as Entomopathogens (Greek entoma = insect, pathos = suffering, gennaein = to produce). The disease causing agents are ubiquitous in nature and have the capability to invade and reproduce in an insect and spread to infect other ones. Several species of naturally occurring noncellular agents (viruses), prokaryotes (bacteria), eukaryotes (fungi and protists) and multicellular animals (nematodes) infect a variety of insects and cause diseases (Table 3.1). Insect diseases include: (a) diseases of productive insects (honey bees, silkworm, lac insect); (b) diseases of pollinators (e.g. honey bees, bumble bees, Syrphid flies etc.); (c) diseases of biological control agents (Coccinellids, predatory stink bugs, praying mantis etc.); (d) diseases in natural insect pest populations (gypsy moth, codling moth, Alfa alfa looper etc.) and (e) diseases of edible insects (grub, caterpillar etc). These infectious microorganisms can be separated into four broad categories of opportunistic, potential, facultative and obligate pathogens based on their capability to infect hosts (Onstad et al. 2006). One the basis of the route of infection, the entomopathogens are classified as ingested (viruses, bacteria, protozoa) and penetrating microorganisms (fungi and nematodes). All of them effectively suppress pests when applied artificially as microbial pesticides or biopesticides (Saxena 2008) (Table 3.2). The use of microorganisms for biological control is commonly referred to as microbial control, an approach that includes introduction, augmentation (inoculation and inundation) and conservation (Eilenberg et al. 2001). The branch of entomology that deals with the study of insect diseases is known as Insect Pathology. In a broad sense, it refers to observations concerning the cause, symptomatology and epizootiology of insects diseases and the study of the resulting structural, chemical and functional body alterations. Insect diseases are generally recognised by obvious symptoms of infection such as movement and irritability, discolouration, changes in body size and shape, and physiological disturbance. The scope of insect pathology encompasses many subdisciplines in entomology. The basic concepts include the cause of the diseases and the classification and phylogeny of entomopathogens. Phylogenies provide a fundamental framework within which we can fit the knowledge produced on all aspects of biology (Lecointre and Le Guyader 2006). It also tells us how closely related the involved organisms are to

Usually cessation of feeding a few hours before death. Swoolen intersegmental regions, Larvae typically turn dark brown or black upon death and are extremely fragile, breaking open and spilling the liquefied internal contents at the slightest touch. Infected cells, primarily fat body, become hypertrophied and tissues disintegrate. Lepidoptera show external symptoms of whitening and softening. Diagnosis include spheroids and spindles (protein inclusions with no virions occluded) in host cell cytoplasm. Injected along with the wasp egg into the body cavity of a lepidopteran host caterpillar and infects cells of the caterpillar. Infection does not lead to replication of new viruses, rather it affects the caterpillar's immune system. Without the virus infection, phagocytic hemocytes (blood cells) will encapsulate and kill the wasp egg but the immune suppression caused by the virus allows for survival of the wasp egg, leading to hatching and complete development of the immature wasp in the caterpillar. Infection causes cell fusion, nuclear alterations, cell contraction and formation of masses in the cells. Infected cells remain "larval", even in adult insects. Most infections are systemic, but the fat body is particularly affected. Infections are usually lethal but low dosages may allow maturation of the host; however, infected pupa and adults are often abnormal. Adult honey bees cluster and crawl abnormally. White spots occur under the integument of some lepidopteran larvae (e.g. silk worms).

Pathogenic trait

(continued)

Common RNA Viruses Cytoplasmic Polyhedrosis Virus (CPV) or cypoviruses Strong effects include developmental lags, lower weight gain, etc. Heavily infected larvae may fail to pupate and usually die. Usually the nuclei of infected tissues are normal, but in heavy infections the cells can become hypertrophied. In fruit flies, abdomens are swollen and discoloured before the flies die. Birnaviruses This family of viruses has only been detected in natural field populations of Culicoides (biting midges). It replicates in the cytoplasm of cells and is somewhat systemic but does not infect fat body tissues.

Iridoviruses

Polydnaviruses (PDV)

Entomopoxviruses (EPV)

Pathogen 1. Viral diseases Baculoviruses

Table 3.1  List of common entomopathogens along with their virulence traits

3  Molecular Phylogeny of Entomopathogens 45

Pathogenic trait Develops in the cytoplasm of fat body cells in honey bee larvae. The virus kills larvae but is relatively benign in adults. The internal organs liquify and the integument remins intact assuming the typical saccular aspect. Brood cells holding infected larvae are recognizable because they are not capped or are only partially capped. Heads of infected larvae darken and turn upward. Dicistroviridae, including: Immature queens die and turn black. Paralyses adults. The disease can be transient, or can weaken Israel acute paralysis virus (IAPV), Kashmir bee virus colonies. The virus infects the cytoplasm of hindgut cells and neural ganglia. Adults are symptomatic: (KBV), Black Queen Cell Virus (BQCV), Acute bee bloated, crawl instead of fly, lose their hairs, tremble and eventually die. paralysis (APV or ABPV), Chronic bee paralysis, Drosophila C virus, Cricket paralysis virus, Aphid lethal paralysis virus, Viral Flacherie of the silkworm (Bombyx mori), Triatoma virus of Triatoma infestans, a hemipteran vector of Chagas disease 2. Bacterial diseases a. Spore-forming bacteria Bacillus thuringiensis Bt possesses 170 or more genes on chromosomes and plasmids for delta endotoxin production (crystal or “Cry” toxins) with specificity to insect gut tissues. Different combinations of these genes produce toxicity to particular insect orders. The three best known subspecies are:   Bt kurstaki with lepidopteran specific toxicity   Bt israeliensis with dipteran specific toxicity (Aedes spp. are most susceptible)   Bt tenebrionis with coleopteran specific toxicity Protoxins of Bt δ-endotoxin are released from the crystal inclusions and, in a susceptible host, are activated. The toxin binds to receptor sites on the brush borders of the midgut columnar cells and creates pores that interfere with ion transport across the cell membranes. Feeding cessation and paralysis occur quickly.

Pathogen Sacbrood virus (SBV)

Table 3.1 (continued)

46 M. Gani et al.

P. lentimorbus - Type B Milky Disease

Paenibacillus popilliae – Type A Milky Disease

Paenibacillus larvae

Pathogen Bacillus sphaericus

Table 3.1 (continued)

(continued)

Pathogenic trait B. sphaericus, a strict aerobe, is a pathogen of mosquitoes. Culex spp. are most susceptible, followed by Anopheles and Aedes. The most virulent species have parasporal crystals and produce two toxins, both required for pathogenesis. Unlike Bt, the endospore (reproductive spore) and the crystal are contained in an ‘exosporium’ that binds them together. B. sphaericus invades the posterior midgut cells and gastric caecae. Some cells swell and vacuolate, and midgut cells separate from each other. Peristalsis ceases, paralysis occurs, and larvae die within two days. P. larvae is the aetiological agent of American foulbrood in honey bees. A facultative anaerobe, this species does not produce a toxin crystal. Rods enter the midgut cells by phagocytosis. Those that survive gain access to the haemolymph where they reproduce. Adult bees are not affected but may harbor bacteria in the gut and serve to transmit the pathogen to larvae. Diagnosis: Larvae typically die late in the development period, turn brown and are of a sticky or ‘ropey’ consistency. Eventually, the carcasses dry into persistent scales at the bottom of brood cells, maintaining the bacterium in uncleaned hives. P. popilliae vegetative forms have a ‘footprint’ appearance with both an endospore and a crystal. The crystal appears to confer some toxicity (related to delta endotoxin of Bt) but toxicity doesn’t affect the course of the disease. Ingested spores multiply in the digestive tract where they penetrate midgut cells. The cells produced in the midgut are arrow-shaped, perhaps aiding penetration into the haemolymph. Spores are sometimes encapsulated by the haemocytes but can continue to reproduce inside the capsules. The host dies of septicaemia. All larvae die, but some survive the first phases or waves of sporulation. This species has no crystal and is less pathogenic. Infected larvae continue to grow and moult for some time. Larvae turn milky brown and haemolymph is clotted; circulation becomes blocked.

3  Molecular Phylogeny of Entomopathogens 47

Entomophthoralean Fungi (Entomophthora, Entomophaga, Massospora spp.)

Pseudomonas entomophila 3. Fungal diseases Ascomycota (Beauveria bassiana and Metarhizium anisopliae)

Xenorhabdus sp.

Photorhabdus sp.

Serratia entomophila Melissococcus pluton - European foulbrood

Lysinibacillus sphaericus Clostridium bifermentans b. Non-spore forming bacteria Serratia marcescens and S. liquifasciens

Pathogen Brevibacillus laterosporus

Table 3.1 (continued)

Smooth skinned larvae sometimes change colour shortly before death. Dead insects appear mummified and feel “bready” to touch. Host bodies may be covered with powdery conidia - the colour of conidia varies and is species specific. B. bassiana turns insects snowy white to yellowish; M. anisopliae whitish to dark green. They produce both conidia on the external surface of the dead host and either sexual, asexual resting spores (or both) internally. For these genera, disease in living insects is difficult to detect and it becomes evident just before or at death.

Ubiquitous in nature and common in the gut contents of insects but highly proteolytic and virulent if the haemocoel is breached. Some strains of S. marcescens are red-pigmented, rendering the dead host with a bright red or reddish tinge. Cessation of feeding and gut clearance. Amber coloration of the grub. Invasion of the haemocoel M. pluton is a seasonal (spring) pathogen of honey bees. The pathogen multiplies in the alimentary tract and is confined in the peritrophic membrane, a chitin and protein microfiber lining of the gut of insect (and other organisms). M. pluton produces a toxin that diffuses across the gut to other tissues. Released of symbiotic bacteria in the haemocoel of infected insects by nematodes of the family Heterorhabditidae. Invasion of the haemocoel by the bacteria, provoking toxaemia and septicaemia Release of symbiotic bacteria in the haemocoel of infected insects by nematodes of the family Steinernematidae. Invasion of the haemocoel by the bacteria provoking toxaemia and septicaemia Strong perturbation of the midgut epithelium. Food uptake blockage

Pathogenic trait Toxin produced during the vegetative phase of the bacterial growth cycle. Toxin maintained during sporulation Mode of action likely similar to that of B. thuringiensis Mode of action likely similar to that of B. thuringiensis

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7. Rickettsiosis and Mycoplasma spp. Wolbachia

Steinernema and Heterorhabditis nematodes

6. Nematodes Mermithids

5. Protozoan diseases –

Pathogen 4. Microsporidia Nosema, Vairimorpha, Octosporea spp.

(continued)

Effects are host species-specific and notably include: cytoplasmic incompatibility that kills offspring (often in the egg stage); male killing (males die during development); feminisation (infected males develop as females); and parthenogenesis.

Matures in the living host; the host dies upon egress. a well-known behaviour of mermithid- and nematomorph-infested insects is water seeking. This behaviour, induced by parasitism, allows emergence of the nematode/ nematomorph in water, which these species require to survive and lay eggs. Rhabditid nematodes in genera Steinernema and Heterorhabditus inject mutualistic bacteria (species specific - typically Xenorhabdus spp. in Steinernema spp. and Photorhabdus spp. in Heterorhabditis spp.) that kill the host via septicaemia and prevent saprophytes from reproducing. Tissues disintegrate, broken down by the bacteria. The invading nematodes mature, mate (if necessary) and produce offspring. When offsprings are in the 3rd stadium (often still protected by the second instar cuticle), they are released from the host via mouth, anus and other body openings. These infective juveniles (IJs, also called Dauer larvae) enter the environment to seek or ambush a new host.

Effects tend to be chronic and transmission ranges from oral to transovarial. Many species are commensal or opportunistic pathogens that only cause pathology when numbers build to very high levels in the gut (e.g. Trypanosoma, Crithidia)

Microsporidia are typically orally transmitted or transovarially transmitted from infected female to the offspring. Heavily infected larvae develop more slowly and may become sluggish in the last stages of disease. Failure to pupate successfully is frequent for heavily infected immature hosts. Some fat body microsporidia may completely fill the cells and cause hypertrophy, resulting in a swollen appearance and pale colour. Infected adults usually have shorter life spans and fecundity often declines.

Pathogenic trait 3  Molecular Phylogeny of Entomopathogens 49

b. Nutritional diseases

Pathogen 8. Degenerative diseases & Neoplastic diseases a. Genetic diseases

Table 3.1 (continued)

Lethal factors (mutants or deficiencies) - most genetic mutations are harmful. Malformations can occur in every body tissue and structure. Sterility factors - Many kinds of sterility factors are possible including malformation of organs and inability to produce eggs or sperm. In biological control programs, chemical or irradiation mutations in males cause sterility in the treated insects. The sterile males are released to compete with wild males for females, their mating producing no offspring. Structural alterations - Any imaginable malformations - missing or deformed wings, supernumerary appendages, deformed body parts. Tumours - Both malignant and non-malignant tumours occur, although non-malignant are most common. Tumour types include benign ovarian tumours, melanotic tumours, neoplasms, imaginal disk (wing formation), and invasive neuroblastomas. Gynandromorphs - or intersex mutants are usually sterile. Nutritional disease can occur in natural insect populations when the host is an outbreak species and population density increases until the normal food supply is depleted, or when the total food source is in short supply, independently of the population density (e.g. stores of honey for overwintering bees dissipate if pollen is in short supply or bees cannot fly in persistent poor weather conditions). In laboratory colonies, nutritional issues are a perpetual problem, particularly with meridic diets where all requirements may not be met despite years of nutritional studies. Symptoms include failure to grow and develop and deformed wings. Starvation mortality is also possible.

Pathogenic trait

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Table 3.2  Entomopathogens as biopesticides with their trade name and target pest Microbial Control Agent Bacteria Bacillus thuringiensis subsp. aizawai B. thuringiensis subsp. israelensis B. thuringiensis subsp. kurstaki B. thuringiensis subsp. tenebrionis Paenibacillus popilliae Fungi Beauveria bassiana

Tradenames of Biopesticides Agree WG and XenTari DF Mosquito Beater WSP

Target Pests Lepidoptera Diptera Lepidoptera

CoStar, DiPel ES, Monterey B.t., and Thuricide Novodor FC

Coleoptera

Milky Spore Powder

Japanese beetle, Popillia japonica

BotaniGard ES, Mycotrol-ESO, Myco-Jaal and Naturalis-LL Hirsutella thompsonii ABTEC Hirsutella Isaria fumosorosea NoFly WP and Pfr-97 WDG Lecanicillium lecanii Phule Bugicide L. longisporum Vertalec Metarhizium anisopliae BioCane, Metarril and Ory-X M. brunneum Met52 EC Paecilomyces lilacinus MeloCon WG Pochonia chlamydosporia Poch_ar Nematodes Heterorhabditis Nemasys and Terranem bacteriophora Steinernema carpocapsae Ecomask and NemAttack S. fatiae Entonem, Fungus Gnat & Rootknot Exterminator and Scanmask H. heliothidis and S. Double-Death carpocapsae Viruses Granulovirus (GV) Cyadia pomonella GV CYD-X and MADEX HP Nucleopolyhedrovirus (NPV) Helicoverpa zea NPV Gemstar LC Spodoptera exigua NPV Spod-X LC Lymantria dispar NPV Gypchek

One or more pests of Acarina, Coleoptera, Diptera, Hemiptera, Hymenptera, Lepidoptera, Orthoptera, Thysanoptera and others

Plant-parasitic nematodes

Several orders of soilborne pests

Lepidoptera

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each other (e.g., all animals are more closely related to each other than they are to plants) and how we can recognize groups of organisms (e.g., animals form a group). Pathogens are often grouped on the basis of phenotypic similarity (e.g., hosts, predilection sites, infection route, microscopy) or similarities concerning the ­ induced disease (e.g., symptoms, diagnostic procedures). There are many other important uses of phylogenies (Harvey et al. 1996), including the study of co-phylogeny of hosts and pathogens (e.g., understanding the role of hosts in a pathogen evolution) and biogeography (e.g., understanding the spread of pathogens). Different pathogens have varying distributions, different patterns of spread and rates of evolution. These differences result in a broad range of variations at the genetic and geographic levels. For example, a phylogenetic tree can be produced to reflect the geographic locations of the samples in order to investigate the spread of a disease, whereas the so-called molecular clocks can be applied to estimate the age of important events in the origin and spread of new pathogens. Most pathogens are unicellular or multicellular without specialized tissues, which severely limits the number and range of available characters. Traditionally, the characters used for phylogenetic and taxonomic analyses have been based mainly on life cycle features, disease characteristics and ultrastructure. It may be rather difficult to determine homologies among such characters (i.e., their evolutionary comparability), so that related characters are being compared, although the data are also regrettably incomplete for most species. Consequently, phylogenies based solely on these characters are rare and not particularly robust. For this reason, molecular data have now become the predominant character information source for phylogenetic studies. As DNA mutates, the sequences change and, as pathogens spread, they bring these changes with them. Molecular data are being generated using DNA-DNA hybridization, randomly amplified polymorphic DNA (RAPD), restriction fragment length polymorphism (RFLP), amplified fragment length polymorphism (AFLP), protein and nucleotide sequences. Of these, nucleotide sequences are by far the most common, used in general for all taxa, not only pathogens. Despite the growing understanding of the phylogeny of entomopathogens, our knowledge of their evolutionary processes is still limited. This chapter provides a comprehensive review on the taxonomy and evolution of entomopathogens and the contribution of genomics in deciphering their phylogeny.

3.2  Molecular Phylogenetics and its Importance Molecular phylogenetics is the branch of phylogeny that analyzes genetic, hereditary molecular differences, predominately in DNA sequences, to gain informations on an organism's evolutionary relationships. Conserved sequences, such as mitochondrial DNA, are expected to accumulate mutations over time. Assuming a constant rate of mutation, they provide a sort of molecular clock for dating divergence events. Molecular phylogeny uses such data to build a "relationship tree" that shows the probable evolution of various organisms. There are several methods available to

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perform a molecular phylogenetic analysis. Phylogenetic analysis of molecular sequences (the actual predominant type of data) usually consists of three distinct procedures: (i) sequence alignment, (ii) character coding and (iii) tree building. A tree from a single molecular sequence represents only the phylogeny of that specific gene the sequence belongs to, which does not necessarily reflect the phylogeny of the whole organism (Doyle 1992). Just how many genes might be required to reconstruct the entire organismal phylogeny is still an open question (Gatesy et al. 2007). Reconstructing a phylogenetic history is conceptually straightforward, although it may take a long time to explicate the most appropriate approach (Hennig 1966). The objective is to infer the ancestors of the contemporary organisms and their ancestors too, all the way back to the most recent common ancestor of the group being studied. Ancestors can be inferred because the organisms share unique characteristics. That is, they have features that they hold in common and that are not possessed by any other organisms. Phylogenesis scientists therefore feel confident that these methods also apply to situations where direct knowledge of the past is absent. Even the Insecta, which is usually considered to be the prime example of a poorly known group, has ~ 1 million species known thus far, out of an estimated total of 4.5–30 million. To date, most pathogen phylogenies have been based on the sequence of a single gene, usually the nucleotide sequence of the small-subunit (16S or 18S) ribosomal RNA gene. Indeed, most of the reclassification of bacteria since the late 1970s has been based principally on this gene (Sapp 2009). Other genes sequenced include those for host recognition or for dealing with the host immune system, often sequenced as part of projects producing new drugs or vaccines. These genes are often unique to each taxonomic group or are subjected to strong selective pressures, thus resulting not necessarily useful for phylogeny. In particular, bacterial genomes often show clusters of functionally related genes such as those for antibiotic resistance (Hedges 1972), which can affect phylogenetic analysis. Consequently, the data are rather fragmentary for many taxonomic groups. A multigene phylogeny is therefore unlikely to be produced from these current data sets. Several “analysis pipelines” have appeared recently, mostly aimed at microbiologists, which do indeed combine several computer programs together to perform a fast, single phylogenetic analysis. These include BIBI (Devulder et  al. 2003), PhyloGena (Hanekamp et al. 2007), WASABI (Kauff et al. 2007), AMPHORA (Wu and Eisen 2008), and ASAP (Sarkar et al. 2008). Probably a better approach is provided by services that allow the mixing and matching of various programs (e.g., Dereeper et al. 2008; Gouy et al. 2010). Attempts have even been made to provide descriptions of “standard procedures” for phylogenetic analysis (Peplies et  al. 2008). MEGA (molecular evolutionary genetics analysis, https://www.megasoftware.net/) is an analysis software that is user-friendly and free to download and use. It is capable of analyzing both distance-based and character-based tree methodologies. MEGA also contains several options one may choose to utilize, such as heuristic approaches and bootstrapping. Bootstrapping is an approach that is commonly used to measure the robustness of a topology in a phylogenetic tree, which demonstrate

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the percentage at which each clade is supported after numerous replicates. In general, a value greater than 70% is considered significant. Phylogenetics is important because it enriches our understanding of how genes, genomes, species (and molecular sequences more generally) evolve. The primary objective of molecular phylogenetic studies is to recover the order of evolutionary events and to represent them in evolutionary trees that graphically depict relationships among species or genes over time.

3.3  Entomopathogenic Viruses 3.3.1  Origin, Natural History and Geographical Distribution The term entomopathogenic refers to those microorganisms that are capable of attacking insects using them as hosts to develop part of their life cycle (Delgado and Murcia 2011). From an applied point of view, they reduce insect pest populations to levels that do not cause economic damage to crops (Tanzini et al. 2001) or regulate disease vectors (Scholte et al. 2004). They are also defined as facultative or obligate parasites of insects, with a high capacity for sporulation and survival (Delgado and Murcia 2011). The viruses that attack and kill insects are called “entomopathogenic viruses” (EV) and are reported from virtually every insect order. The diseases caused by EV are known since the 16th century. The grasserie disease of silkworm (Bombyx mori L.; Lepidoptera: Bombycidae) caused by B. mori nucleopolyhedrovirus (BmNPV) was described since 1524 by Italian bishop Marco Vida da Cremona in the poem “De Bombyce”. Later on another viral disease, Sacbrood was described in the honeybee (Apis mellifera L.; Hymenoptera: Apidae) in 1913. Two Italian scientists, Maestri (1856) and Cornalia (1856), first described the occlusion bodies (OBs) of the BmNPV and Paillot (1926) described the granulovirus (GVs) for the first time in the larvae of cabbage butterfly (Pieris brassicae; Lepidoptera: Pyralidae). Ishimori (1934) identified the cypovirus infection in B. mori by the abnormal presence of dense OB in the insect midgut. The introduction of electron microscopy greatly advanced the knowledge of EV, allowing Bergold to observe the rod-shaped nucleocapsids within the OBs of baculoviruses. The current view on the evolution of baculoviruses states that they evolved from non-occluded viruses infecting midgut tissue to occluded viruses infecting midguts (γ- and δ-baculoviruses) and finally to occluded viruses with the ability to spread the infection to other tissues (α- and β-baculoviruses) (Herniou and Jehle 2007). The most likely scenario is that baculoviruses have gained over time new features to infect more cell types, becoming more independent from the host cell machinery. Steinhaus and his collaborators (1950–1970) tested baculoviruses as biological control agents in the field by applying a nucleopolyhedrovirus (NPV) to control the alfalfa caterpillar (Colias eurytheme Boisduval; Lepidoptera: Pieridae). The first introduction of a baculovirus resulting in successful regulation of a pest insect,

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European spruce sawfly Gilpinia hercyniae occurred accidentally in 1930 in Canada (Bird and Elgee 1957). The first commercial viral insecticide, Elcar for the control of Heliothis/Helicoverpa complex (Lepidoptera: Noctuidae) was marketed in 1975 by Sandoz Agro, Inc. Some key examples of their use in biocontrol are Anticarsia gemmatalis (Ag) MNPV used on over two million Ha of soybean in Brazil (Moscardi 1999; Moscardi and Santos 2005) and H. armigera SNPV (HearNPV) used to control cotton bollworm in China (Zhang et al. 1995). To date, many commercial products based on baculoviruses are available. SPOD-X™, based on Spodoptera exigua (Se) MNPV is used in Europe and USA to control S. exigua larvae, mainly in greenhouses (Kolodny-Hirsch et al. 1997). Mamestrin™ based on Mamestra brassicae (Mb) MNPV is used on cabbage, tomatoes and cotton to control cabbage moth (M. brassicae), American bollworm (H. armigera), diamondback moth (Plutella xylostella), potato tuber moth (Phthorimaea operculella) and grape berry moth (Paralobesia viteana). GemStar™ based on H. zea (Hz) SNPV provides control of pests belonging to the genera Helicoverpa on a wide variety of crops, for these pests are polyphagous. The best commercial example of a GV in biocontrol is Cydia pomonella (Cp) GV being sold under at least five commercial names, Carpovirusine™, Madex™, Granupom™, Granusal™ and VirinCyAP. Also forest insects are being controlled with baculoviruses and several formulations of Lymantria dispar (Ld) MNPV (Gypchek™, Virin-ENSH™), Orgyia pseudotsugata (Op) MNPV (TM-BioControl-1™ and Virtuss™) and Gylpinia herciniae NPV (Abietiv™) are currently available in the market. Natural population of insects in any ecosystem are regulated in time by the action of both biotic and abiotic factors. The biotic factors include entomophagous insects such as parasitoids and predators and entomopathogens such as viruses, bacteria, fungi and nematodes. Abiotic factors include climate, soil, air, space and light (Hostetter and Bell 1985). Insect viruses have been studied for many years due to an intrinsic interest in the general study of invertebrate diseases and, more particularly, because of their potential as environmentally benign pest management agents (Evans 1986). The study of EV ecology and of their potential use for pest management began with the pioneering work of Steinhaus (1956a). Early in the 20th century the disease observed in silkworms was attributed to a virus infection, and in 1947 the visualization of rod-shaped virions, known to be characteristic of the virus family Baculoviridae, was reported. Out of 12 families, baculoviruses have been studied in detail because of their potential as pest control agents (Black et al. 1997; Van Beek and Hughes 1998) and, more recently, for their prominence as expression vectors for a wide range of heterologous genes (Miller 1988; Choi et al. 1999). The advantages of baculoviruses for pest control include their restricted host range (Gröner 1986), non-target effects on useful insects and lack of toxic residues, allowing growers to treat their crops even shortly before harvest, with low probability to develop stable resistance (Monobrullah 2003). Large quantities of baculoviruses are released into the environment during natural epizootics, which are common, widespread, and often important in regulating insect population levels (Federici 1978). There is evidence that the amount of virus

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particles which is artificially spread into the environment for insecticidal purposes is minimal compared with that produced during such epizootics. The baculoviruses have been reported from orders Lepidoptera, Diptera and Hymneoptera. Among them the nucleopolyhedroviruses (NPVs) attracted the attention of pest control scientists looking for an alternative to pesticides, because they cause a highly infectious disease that kills the host in 5-7 days. These viruses attack some of the most important Lepidopteran crop pests including species of Helicoverpa and Spodoptera. Some related GV species are also highly infectious, e.g. the C. pomonella (apple codling moth) GV and P. xylostella (diamond back moth) GV. However not all GVs are as fast acting as NPVs, because morphologically they have a single envelop with a single nucleocapsid per OB (Winstanley and O’Reilly 1999). In general, the host range of most NPVs is restricted to one or a few species of the genus or family of the host from which they were originally isolated. This factor also represents an important commercial draw back, restricting the use of these products to specific key pests or closely related pest complexes, such as Helicoverpa species (Chakraborty et al. 1999). Some of the few exceptions, having a broader host range are: (i) the Alfalfa looper, Autographa californica MNPV, infecting more than 30 species from about 10 insect families, all within the order Lepidoptera; (ii) the Celery looper, Anagrapha falcifera NPV, infecting more than 31 species of Lepidoptera from 10 insect families and (iii) the Cabbage moth, Mamestra brassicae MNPV, which was found to infect 32 out of 66 tested Lepidopteran species from 4 different families (Doyle et al. 1990; Hostetter and Puttler 1991). The maintenance of baculoviruses in insect populations requires transmission of the virus particles from infected to healthy individuals. Transmission in baculoviruses is thought to be primarily horizontal, via susceptible larvae ingesting OBs persisting in the environment. The OBs may be further distributed by excrements of infected larvae (Vasconcelos et al. 1996), rain (Kaupp 1981) and predators, such as birds (Entwistle et al. 1993). Baculoviruses can also be released in the environment by human activity, for example by application of sprays. Baculoviruses can also be transmitted vertically from adults to their young (Easwaramoorthy and Jayaraj 1989; Fuxa et al. 2002). Vertical transmission occurs by surface contamination of eggs (transovum transmission) or virus passing within the egg (transovarian transmission), including transfer of latent infections. A latent virus is a non-replicating form that can be reactivated to an infective state by some stressors (Fuxa and Richter 1992) such as host crowding, fluctuations in temperature or relative humidity, irradiation, dietary changes, chemical stress, parasitization and inoculation by a second pathogen (Burden et al. 2003; Cooper et al. 2003). Epizootics of baculoviruses irregularly occur in agricultural, forest and horticultural pests and have the potential to influence host population dynamics (Weiser 1987). Mathematical models have been used to study and gain insights in these complex systems. The theoretical relationships of host-pathogen dynamics of insects have been widely explored in mathematical models starting with those of Anderson and May (1981). More recent models have incorporated modifications such as variation in transmission parameters (Getz and Pickering 1983), vertical

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transmission (Régnière 1984), both density dependence and vertical transmission (Vezina and Peterman 1985), nonlinear transmission (Hochberg 1991), density dependence (Bowers et al. 1993; Bonsall et al. 1999), host stage structure (Briggs and Godfray 1995), heterogeneity in susceptibility (Dwyer et  al. 1997), discrete generations and seasonal host reproduction (Dwyer et al. 2000), and sublethal infection (Boots and Norman 2000). The EV are naturally occurring and widely distributed throughout the world. Genetic variations appear to be an essential component of natural baculovirus populations. Some variations may be kept in part by the inclusion of different genotypes in the same infectious particles, but also because they can confer a greater fitness to a virus populations. This is the basis on which natural selection drives the evolution of viral lineages adapted to new environmental conditions, such as new host species. Agrotis segetum NPV (AgseNPV) have been isolated only in Europe (Lipa et al. 1971; Allaway and Payne 1983) as its host, A. segetum is found only in Europe, while Agrotis ipsilon NPV (AgipNPV) has been identified only in the United States (Boughton et al. 1999; Prater et al. 2006), although the host, A. ipsilon is found in both, the United States and Europe. This distribution of hosts and viruses provokes the hypothesis that both viruses are descendants from a common ancestor, which is supported by their phylogenies, and then separated into two species, by adapting to different hosts, A. segetum and A. ipsilon, found in the same ecological niches. Since AgseNPV performs well in both hosts, AgseNPV may be the NPV naturally occurring in European populations of both insect species. However, it must be considered that the European AgipNPV has not been discovered yet. Interestingly it was reported that AgipNPV is equally effective against both Agrotis spp. while AgseNPV is about 40 times more effective on A. segetum than on A. ipsilon (El-Salamouny et  al. 2003). It is hard to find an explanation at this point for the equal AgipNPV performance in both insect species, given the fact that in natural conditions it never encounters A. segetum larvae. With the genetic information of both AgseNPV and AgipNPV, and the observation that both viruses replicate in a cell culture derived from the tissues of A. ipsilon the possibility to study this aspect further is now open. Geographical heterogeneity in host-pathogen interactions is thought to play indeed a major role in evolutionary processes (Cory and Myers 2003). Pathogens likely adapt to local host populations, likely due to genetic variability in host and virus (Ebert 1994). Leucoma salicis is a species native for Europe, introduced into North America at the beginning of 20th century (Langor 1995). Orygia pseudotsugata is a native species for North America, and is believed not to occur in Europe. Sequence analysis of a few core genes of two viruses isolated from these hosts, Leucoma salicis NPV (LesaNPV) and Orygia pseudotsugata NPV (OpMNPV), showed their very close relationship (98% nucleotide identity). It was then hypothesized that these viruses originated from one species that evolved by adaptation to different hosts and environmental conditions, and hence showed considerable variation at the genetic level. The Kimura-2 parameter proposed as a criterion for virus species demarcation was 0.010 for polyhedrin gene sequence of LesaNPV and OpMNPV, indicating that these two viruses represent actually the same species.

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Geographical separation might have led to speciation resulting in two viruses, from which one still is able to infect both insect species (LesaNPV), while the other is not (OpMNPV). LesaNPV and OpMNPV thus exemplify evolutional phenomena different from those of AgseNPV and AgipNPV. Many forest insects belonging to Lepidoptera (often from the family Lymantriidae) and Hymenoptera are known for their population cycles (Myers 1988). Populations of these species rise and fall over a predictable period of time. Baculoviruses isolated from these species appear to be specialists. Specialization may result from constant and predictable host availability, in annual cycles. Selection pressure is then switched to improved infectivity against one insect host. Leucoma salicis and Orgyia pseudotsugata are examples of forest lepidopterans showing cyclic population outbreaks. Intuitively there should be a trade-off associated with a broad host range. Generalist viruses, which obviously are less specialized, would be expected to be also less infective against their multiple hosts, than the specialist viruses. However, generalist viruses, in most susceptible hosts, are instead highly infectious (Goulson 2003). The use of molecular techniques to distinguish virus genotypes revealed that the host is usually infected with a mixture of genotypes and that maintaining this mixture is evolutionary favorable because it enables fast adaptation to changing host and environment conditions (Simón et al. 2004). In a given condition, especially in cyclic insect populations with regular outbreaks in a predictable time frame, a pathogen adapts to its host and some genotypes are more prevalent than others. A drastic change in genotype ratios can be seen when the virus is replicating in cell culture, in which the number of defective genotypes almost immediately increases after just one passage (Heldens et  al. 1996, Pijlman et  al. 2001). Defective genotypes are also present in natural baculovirus populations and are critical for high infectivity (Muñoz et al. 1998). Expanding the host range will be reflected in a shift in virus genotype ratios and may even lead to disappearance of some genotypes. Other mechanisms, such as mutations (Pijlman et al. 2001), homologous recombination (Arends and Jehle 2002), transposon insertions (Jehle et al. 1995) and horizontal gene transfer, including the introduction of host genes (Hughes and Friedman 2003), allow optimal adaptation of baculoviruses to local host and environmental conditions, which eventually may result in the evolution of new virus species. Successful (fatal) infection of an insect usually involves a number of stages. However, the ability/inability of the virus to establish infection may depend only on a single gene mutation. Croizier et al. (1994) reported that a change in only two amino acids of the AcMNPV helicase gene extended its host range to B. mori larvae. Thus we can expect that the susceptibility of the host may often be a matter of chance of a single mutation. Spodoptera exigua MNPV (SeMNPV) is believed to be monospecific, infecting only S. exigua. However, it should be mentioned that host range studies for this virus are very limited and so far cover only few insect species (Gelernter and Federici 1986; Smits and Vlak 1988; Simón et al. 2004). It has been confirmed that five proteins (P74, PIF1, PIF2, PIF-3 and PIF4) associated with the ODV envelope are indispensable for oral infectivity and that a deletion of any of them eradicates

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oral infectivity (Song et al. 2008; Fang et al. 2009). Thus per os (oral) infectivity factors (PIFs) are believed to be the main determinants of infectivity. The SeMNPV was non-infective for A. segetum larvae. However, transcription of the delayed early genes was detected in the midgut cells. This brings into question how specific PIFs are and suggesting at least that PIFs may be not the only determinants of oral infection. Apparently, the virus infection can be blocked at any following stage and the entry to midgut cells does not necessarily result in a successful infection. Some events in the midgut cells obviously prevent development of secondary infection. If this occurs at the level of virus gene expression, DNA replication, assembly of new nucelocapsids (NCs) or re-packaging of NCs into BVs, still remains unknown for SeMNPV infection in A. segetum.

3.3.2  Taxonomy and Evolution EV classification, just as for any other type of viruses, follows the indications of the International Committee on Taxonomy of Viruses (ICTV) (Van Regenmortel et al. 2000). The taxonomic classification of viruses is outlined in the Report of the ICTV (Fauquet et  al. 2005), the last being the 10th report (http://ictv.global/report/), as well as the ICTV Taxonomy and Index to Virus Classification and the Nomenclature Taxonomic Lists and Catalogue of Viruses that can be found on the National Centre for Biotechnology Information (NCBI) website (http://www.ncbi.nlm.nih.gov/ ICTVdb/Ictv/index.htm). Therefore, like other viruses, the criteria being followed for their classification include, among many others: type of genetic material (i.e. singe- or double-stranded DNA, singe- or double-stranded RNA, positive or negative strand), virion morphology and size (i.e. icosahedral, rod-shaped, etc.), presence of an envelope surrounding the virion, presence of an OB engulfing the virions, host and host range. All viruses falling into one of these nucleic acid classifications are further subdivided on the basis of whether the nucleocapsid (protein coat and enclosed nucleic acid) assumes a rod like or a polygonal (usually icosahedral) shape. The icosahedral shape viruses are further subdivided into families on the basis of the number of capsomeres making up the capsids. Finally, all viruses fall into two classes depending on whether the nucleocapsid is surrounded by a lipoprotein envelope. However, the ultimate criterion is the sequencing of the genetic material which determines not only the discrimination between viral species, but also the evolutionary relationship among viruses within the same group. The diversity of insect viruses is extensive. They are classified into different families including DNA and RNA viruses (Van Regenmortel et al. 2000) (Table 3.3). Most of the non-occluded (virions not occluded in a protein matrix) or non-­ aggregated viruses are not visible under light microscopy. The occluded viruses — Granuloviruses (GV), Nucleopolyhedrovirus (NPV), Entomopoxvirus (EPV) and Cytoplasmic Polyhedrosis virus (CPV) — are the most commonly observed due to the incorporation of the particles into a protein matrix which is large enough to be visible under light microscopy.

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Table 3.3  Groups of entomopathogenic viruses Nucleic acid dsDNA, circular

Nucleocapsid symmetry Baciliform

Reoviridae

dsRNA linear

Isometric

+

Poxviridae

dsDNA, linear

Ovoid

+

Iridoviridae

dsDNA linear ssDNA

Isometric

–

Isometric

–

Picornaviridae ssRNA

Isometric

–

Ascoviridae

dsDNA circular Polydnaviridae dsDNA circular Rhabdoviridae ssRNA Nodaviridae ssRNA linear Birnaviridae dsRNA linear Bunyaviridae ssRNA Iflaviridae ssRNA

Rod-ovoid

–

Host stage usually Recorded host orders infected Lepidoptera , Larvae, Hymenoptera, Diptera sometimes pupae and adults Lepidoptera, Diptera, Larvae, pupae, adults Hymenoptera, Coleoptera Larvae, pupae, Lepidoptera, adults Coleoptera, Diptera, Hymenoptera, Orthoptera Diptera, Coleoptera, larvae Lepidoptera Lepidoptera, Diptera, Larvae, pupae, adults Blattoidea, Odonata, Orthoptera larvae Diptera, Hemiptera, Hymenoptera, Lepidoptera, Orthoptera Lepidoptera Larvae

Rod, Fusiform

–

Parasitic Hymenoptera Adults

Bullet shaped Isometric

– –

larvae Larvae, adults

icosahedral

–

Diptera, Lepidoptera, Diptera, Coleoptera Diptera

circular Spherical

– –

adult Larvae, adults

Tetraviridae

Isometric

–

Diptera, Hemiptera Lepidoptera, Hymenoptera, Hemiptera, bee parasitic mites (Acarina) Lepidoptera

Isometric

–

Diptera, Hemiptera, Hymenoptera, Lepidoptera, Orthoptera

Larvae, adults

Family Baculoviridae

Parvoviridae

ssRNA linear Dicistroviridae ssRNA linear

Occlusion body +

adult

Larvae

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The vast majority of studies are focused on members of Baculoviridae, as a good number of these viruses are used as biological control agents for the management of insect pests (Copping and Menn 2000; Lacey et al. 2002; Szewczyk et al. 2008). In the past the classification of Baculoviridae was based on morphology, and was divided into the two genera NPVs and GVs (Theilmann 2005). NPVs form large (0.15-15 μm), polyhedral-shaped OBs called polyhedra, which contain many virions that can harbour a single nucleocapsid (SNPV) or multiple nucleocapsids (MNPV). The GVs form smaller (0.3–0.5μm), cylindrical (granule-like) OBs called granules, which contain a single virion with a single nucleocapsid (Fig. 3.1). Virion dimensions are in the size range of 30-60 nm × 260-360 nm for GVs, and 40-140 nm × 250-400 nm for NPVs. The major component of the OBs is a single, viral encoded protein called polyhedrin in NPV, and granulin for GV. The latter is genetically and serologically closely related to the NPV polyhedrin. However, differing from NPV, the host range of GV is narrower and mostly restricted to a single species. NPVs have been isolated from lepidopteran and non-lepidopteran hosts, whereas GVs were only found till now in lepidopterans. A new division was recently proposed on the basis of the comparison of genomic sequences which indicated that virus phylogeny followed more closely the classification of the hosts than the virion morphological traits (Jehle et al. 2006). In this

Fig. 3.1  On the basis of the OB morphology, baculovirus were originally divided in two major groups: the Nucleopolyhedrovirus (NPVs) and the Granulovirus (GVs). NPVs occlusion bodies are called polyhedra and their major occlusion protein is called polyhedrin. The GV occlusion bodies are called granules or capsules and their major protein is granulin. (Source: Haase et al. 2013)

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classification the family Baculoviridae contains four genera: Alpha baculoviruses (lepidopteran specific NPVs), Beta baculoviruses (lepidopteran specific GVs), Gamma baculoviruses (hymenopteran specific NPVs) and Delta baculoviruses (dipteran-­specific NPVs). These four major groups of baculoviruses were identified already in the ‘80s by N-terminal sequencing of major OB proteins (Rohrmann et  al. 1981). At that time hymenopteran baculoviruses were represented only by European pine sawfly, Neodiprion sertifer nucleopolyhedrovirus (NeseNPV). The European crane fly, Tipula paludosa (Tipa) NPV was the dipteran NPV. It has been shown that concatenated sequences of three conserved genes [polyhedrin (polh), late expression factor-8 (lef-8) and late expression factor-9 (lef-9)] or a combined phylogeny of the separate analyses of each of these, yield the same tree topologies as the analysis of complete genome sequences (Herniou and Jehle 2007). This indicated that the multiple conserved genes approach is strongly advised for identification of new baculovirus isolates. Insect viruses are named in acronyms, according to their host(s) and the viral group to which they belong to. For example, the entomopoxviruses are named EPV, the iridoviruses are IV, and the cytoplasmic polyhedrosis viruses (cypoviruses) are CPV. The Autographa californica multiple nucleopolyhedrovirus is named AcMNPV. Therefore, all nucleopolyhedroviruses are named NPV, just as the granuloviruses are named GV.  Still, lepidopteran baculoviruses was divided in NPVs (group I and II) and GVs by genomic sequence data (Van Oers and Vlak 2007). When genome sequences became available from NPVs infecting sawfly and mosquito species, it was clear that lepidopteran NPVs and GVs were closer to each other, than to NPVs from dipterans and hymenopteran hosts. There is a relatively large group of unassigned viruses that infect invertebrates (Fauquet et al. 2005). These are viruses whose key characteristics do not readily fit those of existing genera. Many of them are small ribonucleic acid (RNA) viruses (SRV) from Drosophila or honeybees, while others are well-characterized viruses such as Oryctes rhinoceros virus and Heliothis virus 1, which were once thought to be baculoviruses (Fauquet et al. 2005). The evolution of baculoviruses is governed by the same rules as that of any organism as can be seen by the changes in genotypes in viral populations. For evolution to happen there must first a genetic variation on which natural selection or genetic drift can act. It also depends on genetic variations in the insect hosts populations. Viruses in this co-evolutionary duet have of course more chances to adapt than the hosts, due to a shorter replication time, higher offspring numbers and a high natural heterogeneity. Since baculoviruses can persist outside their host for long times, also the environment exerts a selective pressure. The first hypothesis for evolution of baculoviruses in relation to their host stated that baculoviruses have evolved first in one insect order and then colonized other groups (Rohrmann 1986). The second postulated that the association between baculoviruses and their hosts dates back to the origin of insects and that they coevolved with their host during evolutionary time (Federici 1997). Recently, these two hypotheses have been tested and the results lead to a new hypothesis, postulating that ancestral baculoviruses horizontally infected hosts of different orders and that a progressive specialization

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into different baculovirus lineages later took place (Herniou et al. 2004). Support for this latter hypothesis proceeds from the fact that the phylogeny of baculoviruses follows that of different hosts within an order, hence reflects the pattern of insect families, but does not clearly reflect the evolution of insect orders. Based on sequencing studies, a criterion for demarcating baculovirus species has been depicted. The evolutionary distance between a pair of sequences usually is measured by the number of mutual nucleotide (or amino acid) substitutions. One of the models used to estimate the evolutionary distance between sequences is the Kimura 2-parameter. This method corrects for differences in the rates of transition (conserving the nucleotide ring number, i.e. A → G or C → T and viceversa) and transversion (changing the nucleotide ring number, i.e. A → T or C and C or T → G and viceversa). It allows weighing a quality of difference between transition and transversion (Kimura 1980). The proposed criterion suggests that when the Kimura 2-parameter between single or concatenated genes is larger than 0.050, two viruses are distant enough to be considered as different species. As a consequence, the proposed rules to discriminate baculovirus species are as follows: two (or more) baculovirus isolates belong to the same species if the Kimura-2-parameter between single and/or concatenated polh, lef-8 and lef-9 nucleotide sequence is smaller than 0.015. Two viruses should be considered as different virus species if that distance is bigger than 0.050. For the pair of viruses with the distance between 0.015 and 0.050 complementary information i.e. biological characteristics such as host range should be provided for species demarcation (Jehle et al. 2006). Viral species are defined as “a monophyletic group of viruses whose properties can be distinguished from those of other species by multiple criteria” (Adams et al. 2013).

3.3.3  Genomics and Phylogeny Over the last decade, there has been an explosion of genomic data on insect viruses, primarily because sequencing technologies have improved drastically in accuracy and speed, and whole-genome sequencing has become an affordable procedure. As a result, knowledge on the EV genomics has expanded significantly, yielding more accurate conclusions on the virus-hosts co-evolution (Herniou et al. 2004). The EV genetic is very intricate and complex, because most virus genomes may be as large as 300 kilobase pairs (kb). By far, the most studied group at the genomics level is the Baculoviruses, with 172 genomes belonging to viruses from Lepidoptera, Hymenoptera and Diptera totally sequenced (Wennmann et al. 2018) (Table 3.4). Five genomes of CPVs have been sequenced so far, Bombyx mori (BmCPV-1), Lymantria dispar (LdCPV-1), Dendrolimus punctatus (DpCPV-1), Lymantria dispar cypovirus 14 (LdCPV-14), Trichoplusia ni CPV 15 T (nCPV-15) (Mertens and Bamford 2009). Length and size of fragments vary according to each CPV species. That is, fragments are separated and visualized by agarose electrophoresis producing particular banding patterns in the gel, called electropherotypes. The CPVs genomes are com-

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Table 3.4  Entirely sequenced genomes of baculoviruses Group Virus Name Lepidoptera NPV Antheraea pernyi Group I MNPV – L2 Antheraea pernyi MNPV Antheraea pernyi – NPV Anticarsia gemmatalis MNPV Anticarsia gemmatalis MNPV – 43 Anticarsia gemmatalis MNPV – 26 Anticarsia gemmatalis MNPV – 35 Autographa californica MNPV – C6 Autographa californica MNPV - WP10 Autographa californica MNPV - E2 Bombyx mori NPV – T3 Bombyx mori NPV – India Bombyx mori NPV – C1 Bombyx mori NPV – C6 Bombyx mori NPV – C2 Bombyx mori NPV – Brazil Bombyx mori NPV – Zhejiang Bombyx mori NPV – Guangxi Bombyx mori NPV – Cubic Bombyx mandarina NPV – S1 Bombyx mandarina NPV – S2 Catopsilia pomma NPV – 416 Choristoneura fumiferana MNPV C. fumiferana DEF MNPV Choristoneura murinana NPV – Darmstadt C. occidentalis NPV C. rosaceana NPV

Abbreviation AnpeMNPV

Genome length (kb) 126.246

Genebank accession no. EF207986

AnpeMNPV AnpeNPV AgMNPV

126.629 126.593 132.239

DQ486030 LC194889 DQ813662

AgMNVP

132.077

KR815471

AgMNVP

131.678

KR815455

AgMNVP

132.176

KR815464

AcMNPV

133.894

L22858

AcMNVP

133.926

KM609482

AcMNVP

133.966

KM667940

BmNPV BmNPV BmNPV BmNPV BmNPV BmNPV BmNPV

128.413 126.879 127.901 125.437 126.406 126.861 126.125

L33180 JQ991010 KF306215 KF306217 KF306216 KJ186100 JQ991008

BmNPV

126.843

JQ991011

BmNPV BomaNPV

127.465 126.770

JQ991009 FJ882854

BomaNPV

129.646

JQ071499

CapoNPV

128.058

KU565883

CfMNPV

129.593

AF512031

CfDEFMNPV 131.160 ChmuNPV 124.688

AY327402 KF894742

ChocNPV ChroNPV

KC961303 KC961304

128.446 129.052

(continued)

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3  Molecular Phylogeny of Entomopathogens Table 3.4 (continued) Group

Virus Name Condylorrhiza vestigialis NPV Dashchira pudibunda NPV – ML1 Dendrolimus kikuchii NPV – YN Epiphyas postvittana NPV Hyphantria cunea NPV Lonomia oblique MNVP – SP2000 Maruca vitrata NPV Orgyia pseudotsugata MNPV Philosamia Cynthia ricini NPV Plutella xylostella NPV – CL3 Rachiplusia ou NPV Thysanoplusia orichalcea NPV – p2 Lepidoptera NPV Adoxophyes honmai NPV Group II Adoxophyes orana NPV Agrotis ipsilon NPV – Illinois Agrotis segetum NPV-A – Polish Agrotis segatum NPV-B – English Apocheima cinerarium NPV Buzura suppressaria NPV – Guangxi Buzura suppressaria NPV –Hubei Clanis bilineata NPV – DZ1 Chrysodeixis chalcites NPV – TF1H Chrysodeixis chalcites NPV – TF1C Chrysodeixis chalcites NPV – TF1B

Abbreviation CoveNPV

Genome length (kb) 125.767

Genebank accession no. KJ631623

DapuNPV

136.761

KP47440

DekiNPV

141.454

JX193905

EppoNPV HycuNPV LoobNPV

118.584 132.959 120.023

AY043265 AP009046 KP763670

MaviNPV OpMNPV

111.953 131.990

EF125867 U75930

PhcyNPV

125.376

JX404026

PlxyNPV

134.417

DQ457003

RoNPV ThorNPV

131.526 132.978

AY145471 JX467702

AdhoNPV

113.220

AP006270

AdorNPV AgipNPV

111.724 155.122

EU591746 EU839994

AgseNPV-A

147.544

DQ123841

AgseNPV-B

148.981

KM102981

ApciNPV

123.876

FJ914221

BusuNPV

121.268

KM986882

BusuNPV

120.420

KF611977

ClbiNPV

135.545

DQ504428

ChchNPV

149.624

JX560542

ChchNPV

150.079

JX560539

ChchNPV

149.080

JX560540 (continued)

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Table 3.4 (continued) Group

Virus Name Chrysodeixis chalcites NPV – TF1A Chrysodeixis chalcites NPV – TF1G Chrysodeixis chalcites NPV Chrysodeixis includens NPV – 1C Chrysodeixis includens NPV – 1B Chrysodeixis includens NPV – 1D Chrysodeixis includens NPV – 1A Chrysodeixis includens NPV – 1G Chrysodeixis includens NPV – 1F Ectropis obliqua NPV – A1 Ectropis obliqua NPV – unioasis Euproctis pseudoconspersa NPV – Hangzhou Helicoverpa armigera SNPV – G4 Helicoverpa armigera SNPV – C1 Helicoverpa armigera SNPV Helicoverpa armigera SNPV NNg1 Helicoverpa zea SNPV Helicoverpa armigera SNPV – Australia Helicoverpa armigera SNPV – AC53 Helicoverpa armigera SNPV – AC53C1 Helicoverpa armigera SNPV – AC53C6 Helicoverpa armigera SNPV – AC53T5 Helicoverpa armigera SNPV – H25EA1

Abbreviation ChchNPV

Genome length (kb) 149.684

Genebank accession no. JX535500

ChchNPV

149.039

JX560541

ChchNPV

149.622

AY864330

ChinNPV

140.859

KU669291

ChinNPV

138.869

KU669290

ChinNPV

140.787

KU669292

ChinNPV

140.808

KU669289

ChinNPV

139.116

KU669294

ChinNPV

139181

KU669293

EcobNPV EcobNPV

131.204 130.145

DQ837165 KC960018

EupsNPV

141.291

FJ227128

HearSNPV

131.403

AF271059

HearSNPV

130.759

AF303045

HearSNPV

154.196

EU730893

HearSNPV

132.425

AP010907

HzSNPV HearSNPV

130.869 130.992

AF334030 JN584482

HearSNPV

130.442

KJ909666

HearSNPV

130.460

KU738896

HearSNPV

130.435

KU738899

HearSNPV

130.439

KU738904

HearSNPV

130.440

KJ922128 (continued)

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3  Molecular Phylogeny of Entomopathogens Table 3.4 (continued) Group

Virus Name Helicoverpa armigera SNPV – LB3 Helicoverpa armigera SNPV – LB6 Helicoverpa armigera SNPV SP1B Helicoverpa armigera SNPV –SP1A Helicoverpa armigera SNPV – LB1 Helicoverpa armigera SNPV – L1 Helicoverpa zea SNPV – HS18 Helicoverpa zea SNPV – Brasil/South Heimileuca sp. NPV Lambdina fiscellaria NPV – GR15 Leucania separata NPV Lymantria dispar MNPV Lymantria dispar MNPV –BNP Lymantria dispar MNPV – Japan-3041 Lymantria dispar MNPV – Korea-2161 Lymantria dispar MNPV – Spain-3054 Lymantria dispar MNPV – 27/2 Lymantria dispar MNPV – Russia-27/0 Lymantria dispar MNPV – Russia-3029 Lymantria dispar MNPV – RR01 Lymantria dispar MNPV – USA-45/0 Lymantria dispar MNPV – USA-Ab-a624 Lymantria xylina NPV- 5 Mamestra brassicae MNVP – CHb1

Abbreviation HearSNPV

Genome length (kb) 130.949

Genebank accession no. KJ701030

HearSNPV

130.992

KJ701031

HearSNPV

132.265

KJ701033

HearSNPV

132.481

KJ701032

HearSNPV

131.966

KJ701029

HearSNPV

136.760

KT013224

HzSNP

130.890

KJ004000

HzSNP

129.694

KM596835

HespNPV LafiNPV

140.633 157.977

KF158713 KP752043

LeseNPV LdMNPV LdMNPV

168.041 161.046 157.270

AY394490 AF081810 KU377538

LdMNPV

162.658

KT626571

LdMNPV

163.138

KF695050

LdMNPV

164.478

KT626570

LdMNPV

164.158

KP027546

LdMNPV

161.727

KU249580

LdMNPV

161.712

KM386655

LdMNPV

159.729

KX618634

LdMNPV

161.006

KU862282

LdMNPV

161.321

KT626572

LyxyNPV MbMNVP

156.344 154.451

GQ202541 JX138237 (continued)

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Table 3.4 (continued) Group

Virus Name Mamestra brassicae MNVP – Cta Mamestra brassicae MNVP – K1 Mamestra configurata NPV-A 90/2 Mamestra configurata NPV-A 90/4 Mamestra configurata NPV-B Operophtera brumata NPV – MA Orgyia leucostigma NPV CFS-77 Peridroma sp. NPV – GR-167 Perigonia lusca NPV Pseudoplusia includens NPV – IE Spodoptera exigua multiple NPV – HT-SeSP2A Spodoptera exigua multiple NPV – HT-SeG26 Spodoptera exigua multiple NPV – HT-SeG24 Spodoptera exigua multiple NPV – HT-SeG25 Spodoptera exigua multiple NPV – VT-SeAl1 Spodoptera exigua multiple NPV – VT-SeAl2 Spodoptera exigua multiple NPV – VT-SeOx4 Spodoptera exigua multiple NPV Spodoptera frugiperda MNPV Spodoptera frugiperda MNVP– 3AP2 Spodoptera frugiperda MNPV –Colombian Spodoptera frugiperda MNPV – 19

Abbreviation MbMNVP

Genome length (kb) 153.890

Genebank accession no. KJ871680

MbMNVP

152.710

JQ798165

MacoNPV-A

155.060

U59461

MacoNPV-A

153.656

AF539999

MacoNPV-B

158.482

AY126275

OpbuNPV

119.054

MF614691

OrleNPV

156.179

EU309041

PespNPV

151.109

KM009991

PeluNPV PsinNPV

132.831 139.132

KM596836 KJ631622

SeMNPV

135.395

HG425349

SeMNPV

135.718

HG425348

SeMNPV

135.292

HG425346

SeMNPV

135.556

HG425347

SeMNPV

135.653

HG425343

SeMNPV

134.972

HG425344

SeMNPV

142.709

HG425345

SeMNPV

135.611

AF169823

SfMNPV

135.611

AF169823

SfMNPV

131.330

EF035042

SfMNPV

134.239

KF891883

SfMNPV

132.565

EU258200 (continued)

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Table 3.4 (continued) Group

Lepidoptera GV

Virus Name Spodoptera frugiperda MNPV – G(def) Spodoptera frugiperda MNPV – Nicaraguan Spodoptera littoralis NPV – AN1956 Spodoptera litura MNPV – G2 Spodoptera litura NPV – II Sucra jujube NPV - 473 Trichoplusia ni SNPV Urbanus proteus NPV – Br/South Adoxophyes orana GV – En Adoxophyes orana GV – Mi Agrotis segetum GV Agrotis segetum GV – L1 Agrotis segetum GV - DA Choristoneura occidentalis GV Clostera anachoreta GV – HBHN Clostera anastomosis GV-A Henan Clostera anastomosis GV-B Cnaphalocrocis medinalis GV - Enping Cnaphalocrocis medinalis GV Cryptophlebia leucotreta GV – CV3 Cydia pomonella GV – M1 Cydia pomonella GV – S Cydia pomonella GV – M Cydia pomonella GV – 112 Cydia pomonella GV – E2 Cydia pomonella GV – I07 Diatraea saccharalis GV Epinotia aporema GV Erimmyis ello GV

Abbreviation SfMNPV

Genome length (kb) 128.034

Genebank accession no. JF899325

SfMNPV

132.954

HM595733

SpliNPV

137.998

JX454574

SpltMNPV

139.342

AF325155

SpltNPV II SujuNPV TnSNPV UrprNPV

148.634 135.952 134.394 105555

EU780426 KJ676450 DQ017380 KR011717

AdorGV

99.657

AF547984

AdorGV

99.507

KM226332

AgseGV AgseGV AgseGV ChocGV

131.680 131.442 131557 104.710

AY522332 KC994902 KR584663 DQ333351

ClanGV

101.487

HQ116624

ClanGV-A

101.818

KC179784

ClanGV-B

107.439

KR091910

CnmeGV

111.246

KU593505

CnmeGV

112.060

KP658210

CrleGV

110.907

AY229987

CpGV CpGV CpGV CpGV CpGV CpGV DisaGV EpapGV ErelGV

123.500 123.193 123.529 124.269 123.858 120.816 98.392 119.082 102.759

U53466 KM217573 KM217575 KM217576 KM217577 KM217574 KP296186 JN408834 KJ406702 (continued)

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Table 3.4 (continued) Group

Hymenoptera NPV

Diptera NPV

Virus Name Helicoverpa armigera GV Mocis sp. GV Mythina unipuncta GV Phthorimaea operculella GV – Tu Phthorimaea operculella GV – Sa Pieris rapae GV – (1) Pieris rapae GV –E3 Pieris rapae GV – (2) Plodia interpunctella GV – Cambridge Plutella xylostella GV – K1 Plutella xylostella GV – T Plutella xylostella GV – M Plutella xylostella GV – C Plutella xylostella GV – K Plutella xylostella GV – SA Pseudaletia unipuncta GV – Hawaii Spodoptera frugiperda GV – VG008 Spodoptera litura GV – K1 Trichoplusia ni GV – LBIV12 Xestia c-nigrum GV Neodiprion abietis NPV Neodiprion lecontei NPV Neodiprion sertifer NPV Culex nigripalpus NPV

Abbreviation HearGV MospGV MyunGV PhopGV

Genome length (kb) 169.794 134.272 144.673 119.217

Genebank accession no. EU255577 KR011718 KX855660 AF499596

PhopGV

119.004

KU666536

PiraGV PiraGV PiraGV PlinGV

108.592 108.476 108.658 112.536

GQ884143 GU111736 JX968491 KX151395

PlxyGV

100.999

AF270937

PlxyGV PlxyGV PlxyGV PlxyGV PlxyGV

100.978 100.986 100.980 101.004 100.941

KU529794 KU529793 KU529791 KU529792 KU666537

PsunGV

176.677

EU678671

SfGV

140.913

KM371112

SpltGV TnGV

124.121 175.360

DQ288858 KU752557

XecnGV NeabNPV

178.733 84.264

AF162221 NC008252

NeleNPV NeseNPV CuniNPV

81.755 86.462 108.252

NC005906 NC005905 AF403738

Source: Wennmann et al. (2018)

prised in average by 10 dsRNA fragments with a varying length of 0.4–4 kb. Different electropherotypes from cypovirus were initially identified on the basis of differences in the migration patterns of their genome segments during gel electrophoresis, which was used to identify them. The sum of all these fragments adds up to a total genome length of 19–25 Kb. So far, 14 different electropherotypes have been identified (Martens et  al. 1989). Due to these studies, it is known that the smallest fragment contains the coding gene for the CPV’s polyhedrin, which is the major OB component.

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The molecular biology of Entomopoxviruses has been limited by the lack of genomic information. EPV genomic organization and molecular mechanisms of replication, pathogenesis and host range are largely unknown. Known EPV genomes are constituted of a single dsDNA molecule of 130 to 375 kb in length. Such an extensive genome may have the potential to code for as many as 150 to 300 genes (Arif 1984). A peculiarity of these genomes is the presence of isometric DNA sequences at the terminal ends of the molecule, constituted by inverted palindromic motifs. Two EPVs genomes have been sequenced so far, from migratory grasshopper, Melanoplus sanguinipes (MsEPV) (Afonso et al. 1999) and from red hairy caterpillar, Amsacta moorei (AmEPV) (Bawden et al. 2000). The 236-kb MsEPV genome contains a subset of genes shared between all sequenced poxviruses, and allowed the concept of a common, universally shared genetic core of poxvirus genes. Poxvirus core genes included many of those associated with RNA transcription, post-transcriptional modification, DNA replication and core structural proteins. The EPV genes are classified according to phases of expression as: (a) early (b) intermediate and (c) late genes. Early genes are expressed before the genomic DNA is replicated. Even, some genes are expressed before the DNA is totally released from the capsid. It is known that proteins expressed at the early events of infection inhibit the synthesis of macromolecules by the host cell. These early genes code for non-structural proteins, including enzymes involved in DNA replication, and modification of DNA and RNA, as well as those implicated in the inactivation of host defence mechanisms. Additionally, these genes code for transcriptional factors used by intermediate genes. During the intermediate phase, viral DNA replicates and intermediate genes code for transcriptional factors used by the late genes. Finally, late genes are expressed once the viral DNA has been replicated and code mostly for structural proteins, both for the assembling of the capsid and the formation of spheroids. Interestingly, some of these genes also code for transcriptional factors used by early genes. Only two entomopathogenic Invertebrate Iridescent Virus (IV) genomes have been sequenced to date, IIV-6 or Chilo iridescent virus (Jakob et al. 2001) and iridescent virus type 3 (IIV-3) or mosquito iridescent virus (Delhon et al. 2006). The IV genome is packed as a circularly permuted and terminally redundant ds-DNA molecule of 140–303 kb (Goorha and Murti 1982). IV genomes range in size from 105 to 212 kb and contain 96 to 234 largely non-overlapping open reading frames (ORFs), a G-C content ranging from 27% to 55% and complex repeat sequences mostly located between coding regions. Genomes exhibit little to no co-linearity among genera (Delhon et al. 2006). The replication process of IV genome is highly complex. Replication starts early after infection, as DNA replicates in the nucleus, producing similar or shorter copies. Replication continues in the nucleus but some copies are transported to the cytoplasm, where replication goes on and recombination occurs among the copies constructing a complex of concatamers (Ward and Kalmakoff 1991). So far, it is known that a virus encoding enzyme called DNA integrase-recombinase may be involved in the recombination of small pieces of DNA as well as in the resolution of

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the concatamer, just before the DNA is packed within the capsids. Transcription of the early genes starts with the RNA polymerase II from the host, which has been modified by one of the viral proteins (Jakob and Darai 2002). Only three genomes of ascoviruses have been fully sequenced: Trichoplusia ni ascovirus 2c (TnAV-2c), Spodoptera frugiperda ascovirus1a (SfAV-1a) and Heliothis virescens ascovirus 3e (HvAV-3e) (Cui et al. 2007). The study of gene content has the potential to show the extent of variation between baculovirus genomes whereas the comparison of genomes may provide valuable insights into their evolution and biology (Herniou et al. 2003). Baculovirus genomes range in size from approximately 80–180 kb (Theilmann 2005). Currently, the baculovirus with the largest genome is Xestia c-nigrum NPV (XecnGV) with 178,733 bp (Hayakawa et al. 1999), while the smallest belongs to Neodiprion lecontei NPV (NeleNPV) with 81,755 bp (Lauzon et al. 2004). The recently determined complete genome of Antheraea proylei nucleopolyhedrovirus (AnprNPV) is 126,930 bp in lenth and encodes 147 ORFs (Shantibala et al. 2018). The number of predicted ORFs found in sequenced baculoviruses encoding 50 or more amino acids range from approximately around 89–181 ORFs. The average G + C content is quite variable in baculoviruses, ranging from around 30 to 60%. A distinctive feature of most sequenced baculoviruses is the presence of repeat regions or homologous regions (hrs) dispersed throughout the genome, ranging from approximately 3 to 17 hr/repeat regions. In NPVs, most hrs contain 30 bp palindromes within direct repeats and are similar to other NPV hrs, whereas GV repeat regions are more variable and often lack palindromes (Wormleaton et  al. 2003). Hymenopteran baculovirus genomes contain repeated regions that do not conform to the typical structure of hrs in lepidopteran NPVs. Comparative analysis provided insight into their evolutionary history in that differences and similarities in amino acid sequences and gene order aided in the division of baculoviruses into groups, sharing gene characteristics and overall genome relatedness. More closely related viruses share a higher degree of gene co-linearity. Hu et al. (1998) developed a method called “gene parity plots” that compared the positions of homologous genes in different genomes and is used to show conservation between baculovirus genomes. Another method for comparing and classifying genomes is the use of phylogenetic trees (Herniou et al. 2003). Early attempts to infer relationships between baculoviruses and to study their evolution were approached by phylogenetic analyses of the amino acid and nucleotide sequences of the polh and granulin (gran) gene, which constitute the OB matrix of NPVs and GVs, respectively. The phylogenetic analyses based on polyhedrin gene sequences further subdivided lepidopteran NPVs into group I and group II NPVs (Zanotto et al. 1993). This division moreover appeared to correlate with the utilization of two different budded virus (BV) envelope fusion proteins. Group I NPVs contain the major envelope glycoprotein GP64, which mediates membrane fusion (Blissard and Wenz 1992). Group II NPVs as well as GVs lack GP64 protein, but contain a functional homolog designated as F protein (Westenberg et al. 2002). The major OB proteins polyhedrin and granulin are made in large amounts and for this reason large quantities can be obtained and sequenced from the N-terminus

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(Rohrmann et al. 1981). Moreover, the genes coding for polyhedrin and granulin are highly conserved and thus easily identified in new baculovirus isolates, by DNA hybridization or more commonly by PCR analysis with degenerate primers (Gani et al. 2017). Single gene phylogenies, however, may occasionally lead to misinterpretation as they not always reflect authentic relations between viruses. This was for example the case for the AcMNPV polh gene, which appeared to have a mosaic structure resulting most probably from recombination events (Lange et al. 2004). While biological data as well as sequence information for other AcMNPV genes clearly show that AcMNPV belongs to group I NPVs, its polh gene is most closely related to the homologue in Trichoplusia ni SNPV, a group II NPV. Nevertheless, polh gene phylogenies as well as phylogenies of other conserved genes usually reflect baculovirus relationships and evolution. In fact, polh was until recently the only choice for analysis of large numbers of baculoviruses due to the limited sequences available for other genes. The use of a concatenation of shared genes has been shown to produce more reliable trees (Herniou et al. 2003). This is done by producing concatamers of the core proteins found in all baculoviruses, aligning their amino acid sequences and generating trees that reflect different taxonomic divisions. Gene order and phylogeny provides essential information on the evolution and relatedness of baculoviruses. Comparative analysis of all completely sequenced baculoviruses revealed a set of core genes conserved in all genomes that have essential roles in the baculovirus life cycle (Garcia-Maruniak et al. 2004). There are 38 conserved core genes found in all baculoviruses (Wennmann et al. 2018). Most of them have a known function within the genome, either required for RNA transcription, DNA replication or as structural and auxiliary proteins. Identification of genes that are essential or that stimulate DNA replication in baculoviruses has provided a basis for elucidating the process by which they replicate their genomes (Kool et al. 1995). Baculovirus DNA replicates in the nucleus and they carry their own complement of genes encoding DNA replication proteins. Four of these genes are found in all sequenced baculoviruses to date: DNA polymerase, DNA helicase (p143), lef-1 and lef-2 (Herniou et al. 2003). A list of additional genes involved in DNA replication found in lepidopteran baculoviruses also includes: lef-3, ie-1 and me53 (Herniou et al. 2003; Lange and Jehle 2003). An additional gene, dbp (DNA binding protein), is found in lepidopteran and dipteran baculoviruses (Lauzon et al. 2004). Four major groups of baculovirus genotypes were identified by sequencing the N-terminus of purified OB proteins (Rohrmann 1986). (i) the dipteran group, whose OB proteins did not show any relatedness to those of other baculoviruses, (ii) the hymenopteran group, with limited relatedness to polyhedrins of the lepidopteran viruses, (iii) the nucleopolyhedroviruses and (iv) the granuloviruses. These main clusters were later corroborated using DNA sequence analyses of other genes (lef-8, ac22-pif2) (Herniou et  al. 2004) and by genome sequence comparisons, which revealed that a NPV isolated from the mosquito Culex nigripalpus (CuniNPV) did not have a homologue of the polyhedrin gene, but used a different gene to encode its OB protein (Afonso et al. 2001). Whether any of the single gene trees indeed repre-

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sented the phylogeny of the baculoviruses had not been evaluated until whole genome sequence comparisons were performed. Herniou et  al. (2001) was the first study to use the information derived from whole genomes to infer a baculovirus phylogeny and compare the trees based on complete genomes to those based on single genes. The inference of phylogeny based on complete genomes followed three complementary approaches using: (i) the concatenated sequences of the core genes present in all baculoviruses, (ii) gene order and (iii) the gene content of the genomes. These approaches had separately been successfully applied to the phylogenetic analyzes of herpesviruses (Montague and Hutchison 2000). Applying these three methods on 9 and 13 complete genomes, the phylogeny of baculoviruses was clarified, permitting the comparison of single gene trees (Herniou et al. 2003). Of all the core genes, comparisons of only seven genes (ac22/pif-2, ac81, ac119, ac142, ac145, lef-8 and lef-9) produced the same phylogenetic tree comparisons based upon complete genomes (Herniou et al. 2001). Despite the limitations of single gene trees, their use in phylogenetic analysis is still needed when less characterized viruses are being investigated. In order to overcome the limitation of single gene trees, a small number of phylogenetically informative genes may be used. Herniou et al. (2004) developed degenerate PCR primers that are suited to amplify highly conserved gene fragments within pif-2 (ac22) and lef-8 (ac50), which can be directly subjected to DNA sequencing and molecular phylogenetics (Herniou et al. 2004). By using this approach the separation of the baculovirus tree into four main groups as described above was validated for an unprecedented number of viruses. Lange et  al. (2004) also succeeded in developing degenerate primers for conserved sequences within lef-8, lef-9 and polh/gran of Lepidoptera-specific NPVs and GVs (Lange et al. 2004). Using this method, a comprehensive phylogenetic tree of 117 baculoviruses was inferred by Jehle et  al. (2006). Baculovirus phylogeny based on the maximum-likelihood (ML) method [using RAxML (randomized accelerated maximum likelihood)] software was recently inferred by Wennmann et  al. (2018), using concatenated 38 core-gene amino acid sequences from 172 entirely sequenced baculovirus genomes. It was found that the 38 core-gene data provided a more accurate method to define distinctions between baculovirus species than the original 3-gene concept. A baculovirus phylogenetic tree using 37 core genes from 81 baculovirus complete genomes was constructed by Nguyen et al. (2018) (Fig. 3.2). On the basis of complete genome analyses (Herniou et al. 2001; Herniou et al. 2003; Lauzon et al. 2004) and gene phylogenies (Herniou et al. 2004; Lange et al. 2004; Jehle et al. 2006), the following conclusions can be derived: (i) Baculoviruses comprise at least four distinct clades, which should be considered as different genera. These are (a) lepidopteran-specific NPVs, (b) lepidopteran-­specific GVs, (c) hymenopteran-specific NPVs and (d) dipteran-­ specific NPVs; (ii) the lepidopteran-specific NPVs fall into two most likely monophyletic clades, the so-called group I and group II NPVs, utilizing different envelope fusion proteins for cell-to-cell spread;

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Fig. 3.2  Molecular phylogenetic analysis by Maximum Likelihood method. The tree was constructed using 37 core genes from 81 baculovirus complete genomes. Bootstrap value resulted from 1000 replications is shown in each node. Red arrow – HytaNPV. (Source: Nguyen et al. 2018)

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(iii) within group I NPVs, two monophyletic clades (Ia and Ib) have been defined (Jehle et al. 2006). Clade Ia includes Autographa californica NPV (AcMNPV) and variants of this virus isolated from other insects, as well as Rachiplusia ou MNPV, Bombyx mori NPV, and Thysanoplusia orichalcea NPV. These viruses infect members of the families Noctuidae, Bombycidae, Pyralidae and Plutellidae. Clade Ib comprises hosts from the Lepidoptera families Arctiidae, Geometridae, Lymantriidae, Notodontidae, Nymphalidae, Saturniidae, and Tortricidae. There are further group I NPVs, which do not belong to clade Ia and Ib; (iv) group II NPV relationships are characterized by long branches and weak resolution at the basal nodes, which suggests that this group is more ancient than group I NPVs (Jehle et  al. 2006). However, their host range is apparently restricted to fewer lepidopteran families, as group II NPVs have mostly been isolated from higher Macrolepidoptera families, such as Noctuidae, Lymantriidae, Lasiocampidae and Geometridae. Thus far, the only Microlepidoptera-­specific group II NPVs have been identified from Adoxophyes honmai (Tortricidae) and Wiseana cervinata (Hepialidae). Improved virus sampling might show an increase in the number of lepidopteran host families infected by group II NPVs; (v) GVs have so far been identified from 9 Lepidoptera families including Noctuidae, Arctiidae, Plutellidae, Nodontidae, Pieridae, Tortricidae, Bombycidae Sphingidae, and Gelechiidae. Similar to group II NPVs basal resolution is weak and the branches are generally long, suggesting an ancient radiation of these viruses. Midgut infecting GVs (Harrsinia billions GV) and slow killing GVs (Adoxophyes orana GV, Xestia c-nigrum GV and Trichoplusia ni GV) are scattered within the vast majority of fast GVs causing systemic infection (Federici 1997; Wormleaton and Winstanley 2001). This clearly indicates that these traits are not phylogenetically informative and therefore not suitable for a natural classification of GVs. GVs isolated from Tortricidae appear to have a monophyletic origin irrespective of their pathogenesis and replication speed (Lange et al. 2004; Jehle et al. 2006).

3.4  Entomopathogenic Bacteria (EB) 3.4.1  Origin, Natural History and Geographical Distribution Bacteria may be generally defined as unicellular and ubiquitous microorganisms possessing a single chromosome deprived of a nucleus membrane and having ribosomes of the 70S type. These organisms proliferate through binary fission, a process resulting in daughter cells that are essentially identical copies of the mother cell. Bacteria constitute a large domain of prokaryotic microorganisms. Typically a few micrometres in length, they have varying shapes which include spherical, rod-like

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and spiral cells. Bacteria were among the first life forms to appear on Earth and inhabit all environments, including soil, oceans and land waters, acidic hot springs, radioactive waste and the deep portions of the Earth's crust. They also live in symbiotic and parasitic relationships with or within plants and animals. The ancestors of modern bacteria were unicellular microorganisms, the first forms of life to appear on Earth, about four billion years ago. Although bacterial fossils exist, such as stromatolites, their lack of distinctive morphology prevents them from being used to examine their evolutionary history or to date the time of origin of a given species. The most recent common ancestor of bacteria and archaea (the other branch of Prokaryotes) was probably a hyperthermophile that lived about 2.5–3.2 billion years ago (Di Giulio 2003; Battistuzzi et al. 2004). As with other living organisms, insects are intimately associated with bacteria at all stages of their lives. EB infecting insects have sizes ranging from less than 1 μm to several μm in length. Insects inhabit diverse niches and interact with various bacteria to form relationships that range from mutualistic symbiosis to pathogenesis. Depending on the relationship, symbiotic associations can be divided into commensalism, mutualism and parasitism (Moya et al. 2008). The mechanisms leading to these interactions are presumed to have ancient origins and to have developed throughout a long co-evolutionary process (Vilcinskas 2010). Pathogenic interactions require a high degree of specialisation and intimate contact with the host. In this respect, many of the molecular mechanisms used by bacterial pathogens and mutualists are similar. Furthermore, the same microorganism can behave differently, depending on the fitness of the host and the environmental circumstances, turning from beneficial to detrimental, thus complicating the arbitrary definition of bacterial behaviour. Therefore, the concepts of mutualism and pathogenesis are not clearly differentiated, but rather a matter of balance between the bacteria and the insect host in terms of fitness, reproductive success, feeding and influence of other symbionts. Given that insects and bacteria are some of the most numerous organisms on our planet, their interactions hold significance in many areas. Most bacteria currently are spore forming members of the bacterial family Bacillaceae and Enterobacteraceae. In Bacillaceae the genus Bacillus is the most important. The insect pathogenic Bacilli occur in healthy and diseased insects and can also be isolated from many other habitats including insect frays, soil, plants, granaries and aquatic environments. Over 90 species of naturally occurring insect-specific (entomopathogenic) bacteria have been isolated from insects, plants and soil, but only a few have been studied intensively. Much attention has been given to Bacillus thuringiensis (Bt) (Firmicutes: Bacillaceae) a species that has been developed as a microbial insecticide and safe alternative for the management of insect pests (Lacey et  al. 2001; Lacey and Kaya 2007). Indeed, Bt originally discovered in 1901 by Ishiwata and later rediscovered and isolated by Berliner in 1915 in Germany from the diseased larvae of the Mediterranean flour moth, Ephestia kuhniella has been the most studied and broadly used in microbial control. Screening programs have identified thousands of different strains of Bt all of which have a limited host range but together span a wide range of insect orders (Lepidoptera, Diptera, Coleoptera, Hymenoptera,

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Homoptera, Orthoptera and Mallophaga) and even other organisms such as nematodes, mites and protozoa (Feitelson et al. 1992). Manonmani and Balaraman (2001) isolated several strains of B. thuringiensis from various sources, i.e. soil, water, larvae and roots of aquatic weeds representing diverse habitats. These strains were examined for their flagellar antigenicity, mosquito larvicidal activity and protein composition. Among these, one strain belonging to the serotype B. thuringiensis thompsoni (H-12) was found to be highly toxic to different species of mosquitoes. Nine different indigenous B. thuringiensis isolates were recovered from the soil of cotton fields in different Egyptian governorates namely, Minofiya, Sharkiya, Gharbiya, Kalyoubiya, Dakahliya and Kafr El-Sheikh (Saker et al. 2012). The discovery that Bt spore-associated toxins are extremely virulent and can persist in the environment with high potency, prompted the development of bacterial spray formulations (Wilcox et al. 1986) and transgenic plants expressing certain Bt toxins (Fischhoff et al. 1987). Different Bt varieties produce a crystal protein that is toxic to specific groups of insects. Sprays of sporulated B. thuringiensis have a long history of safe use for pest control in agriculture (Paul et  al. 2017). Sprays of B. thuringiensis subsp. israelensis (Bti) and of Bacillus sphaericus (Bsp) have been used to control disease-carrying mosquitoes and blackflies (Land and Miljand 2014). Since 1996, transgenic crop plants expressing entomocidal Cry proteins from B. thuringiensis have been commercialized, resistant to several insect pests (de Maagd et al. 1999). The most common species applied are B. thuringiensis var. kurstaki, var. israeliensis, var. tenebrionis, B. popilliae and B. sphaericus (Lysinibacillus sphaericus). These biological control agents being persistent, cheap, readymade, highly specific and safe for non-target flora and fauna, certainly contributed to the development of management practices for agricultural and forest insect pests. Advancements in the characterization of bacterial pathogens including purification and culturing methods, molecular identification of virulence factors and whole genome characterization and comparisons have prompted the discovery of novel pest management tools. In this view, insecticidal molecules expressed and secreted by various EB have been targeted for genetic manipulation to enhance toxicity (Gatehouse 2008). Recently, other insect pathogenic bacteria with modes of action similar to Bt have been hailed as agriculturally relevant (Chattopadhyay et al. 2004). Certainly, EB of diverse taxonomic groups and phylogenetic origins have been shown to have striking similarities in the virulence factors they produce (Priest et al. 2004). Bacterial virulence factors are often encoded on mobile genetic elements such as plasmids and can easily be spread through horizontal gene transfer by bacteriophages. The discovery that both Gram-negative and Gram-positive bacteria produce analogous insect-specific toxins infers history of gene transfer between them (Ochman et al. 2000). For example, Photorhabdus luminescens (Proteobacteria: Enterobacteriaceae), the bacterial symbiont of the entomopathogenic nematode Heterorhabditis bacteriophora (Rhabditida: Heterorhabditidae), has virulence factors similar to those of Bt (Nielsen-LeRoux et  al. 2012). The first commercial biopesticide, Sporeine, commercialized in 1938  in France, was based on Bt. Moreover, the first EB used in a major insect control program was Paenibacillus

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popilliae (Dutky, previously Bacillus popilliae; Klein and Jackson 1992), the causative agent of milky disease in Japanese beetle (Popillia japonica Newman). Despite successful use, problems related to the mass production of viable P. popilliae spores (Stahly and Klein 1992) reduced commercial interest in this bacterium. Although commercial products based on P. popilliae are currently available, their use is limited to the control of grubs, especially in organic agriculture (Johnson et al. 2001). Both B. thuringiensis and L. sphaericus are ubiquitous soil microorganisms and isolates have been obtained from multiple environments worldwide (Guerineau et al. 1991; Bernhardt et al. 1997). Photorhabdus and Xenorhabdus are the genera of EB that live in symbiosis with nematodes of the family Heterorhabditis or Steinernema, respectively, and together they infect insect larvae. As symbionts, the bacteria are food that supports the bacteriovorous nematode host development, but also produce toxic natural products (NP) and proteins that kill the insect prey, in which they actively multiply (Bode 2009).

3.4.2  Taxonomy and Evolution Bacterial classification began with taxonomic units based solely on the morphological characteristics of the organisms, and progressed to the use of a wide variety of characteristics including physiology, biochemistry and genetic data. Their classification was started by Muller (1773) with the discovery of different morphological forms. Cohn (1872) classified bacteria into 4 major types depending on their shapes as Cocci (spherical or elliptical), Bacilli (rod or cylindrical), Vibro (curved or comma) and Spirilla (spiral or spring like). Depending upon the staining reactions by the Gram stain, bacteria are classified into Gram positive and Gram negative. Today, the sheer number of species that have been identified and the wide diversity among them make classification of these organisms into discreet arrangements both difficult and necessary. Two main groups of prokaryotic microorganisms are recognized based on the 16S ribosomal RNA sequence: Archaea, containing microorganisms that share DNA replication, transcription and translation features with eukaryotes; and Eubacteria or true bacteria. EB are classified within Eubacteria. This group contains three major divisions, based on the presence or structure of the cell walls: bacteria with a Gram-negative type cell wall (Gracilicutes), those with a Gram-positive type cell wall (Firmicutes) and Eubacteria lacking a cell wall (Tenericutes). EB may be found in all three main Phyla and are mainly classified as spore producers and ­non-­spore producers (Fig.  3.3). However, when focusing on bacteria with a ­demonstrated use in insect control, species of interest occur in the following families: Bacillaceae, Paenibacillaceae, Enterobacteriaceae, Neisseriaceae, ­ Pseudomonadaceae, Lactobacillaceae and Micrococcaceae. Classification of EB into groups on the basis of their pathogenicity presents difficulties, as most bacteria are facultative pathogens and present virulence variations depending on the host environment and the specific strain. Bacterial populations,

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Fig. 3.3  Phenotypic classification of entomopathogenic bacteria

including pathogenic ones, are essentially clonal, implying that exchange of DNA in nature is infrequent. Many species displaying pathogenicity exist as a number of distinct lineages, or clones, of which only a few contain the genetic information that encodes pathogenicity (Selander et al. 1987). For example, genetic diversity among strains of B. sphaericus explained variation in toxicity against mosquito larvae (Krych et al. 1980). In some cases, non-pathogenic and disease-causing clonal lineages have been named and classified as diverse species based on their pathogenic niche (Lan and Reeves 2001). To resolve this issue, a bacterial species concept based on members sharing a core genome (housekeeping genes) was proposed (Dykhuizen and Green 1991). In this view, lineages incorrectly described as diverse are named as clones within a species, based on sequence similarity (Lan and Reeves 2000, 2001). Advances in DNA sequencing technology with the concomitant increase in available bacterial genomic data, together with increased characterization of bacterial population genetics, are expected to allow for a more accurate and useful definition of species that will increase EB understanding. Bergey’s Manual of Systematic Bacteriology defines the bacterial species as “a collection of strains that share many features in common and differ considerably from other strains.” It goes on to say that "a species consists of the type strain and all other strains that are considered to be sufficiently similar to it as to warrant inclusion with it in the species" . A more uniform definition of the bacterial species is desirable and can possibly be obtained through the use of genetic relatedness among bacteria. Established taxonomic techniques, including biochemical tests and serological techniques, allow ready qualitative identification. Bacterial strains are grouped into species based on sharing of certain distinguishing phenotypes of ecological importance and overall genomic similarity (Stackebrandt et al. 2002). A bacterial strain is defined as the descendants of a single isolation in pure culture and is usually derived from successive cultures, from an initial single colony. Phenotypic screening is usually considered the first step in the identification of bacterial strains as it allows a rough

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comparison and placement into similar or different groups. Most commonly considered phenotypic traits include: (i) colony morphology (size and shape); (ii) colony pigmentation; (iii) cell shape and size; (iv) motility and flagellar arrangement; (v) Gram stain reaction; and (vi) aerobic or/and anaerobic metabolisms. In practice, more than 70% DNA-DNA hybridization values are considered evidence of a single species (Wayne et al. 1987). The DNA hybridization method has been then integrated and partially replaced by sequencing methods. Within Eubacteria, current classification is mostly based on polyphasic (consensus) taxonomy, including analysis of the nucleotide sequence of the ribosomal small-subunit RNA (16S rRNA), DNA-DNA hybridization, as well as phenotypic, genotypic and phylogenetic data (Brenner et al. 2005). In the past 50 years, numerous bacteria such as B. sphaericus, B. thuringiensis and Serratia marcescens have been isolated, classified and demonstrated in the laboratory to be pathogenic to various insects (Bahar and Demirbag 2007; Kleespies et  al. 2008). Diverse methods have been used to classify B. thuringiensis and L. sphaericus isolates, with flagellar H-serotyping being the most widely present in the literature. For B. thuringiensis, 85 serotypes have been described (Jurat-Fuentes and Jackson 2012), although most commercial biopesticides are based on serovars (or subsp.) kurstaki (Btk), aizawai (Bta), israelensis (Bti), and tenebrionis (Btt). Each of these serovars includes isolates expressing crystal (Cry) parasporal proteins with entomotoxicity against specific insect taxonomic orders, including Lepidoptera, Coleoptera and Diptera. Additional noncrystal toxins and other virulence factors that enhance entomotoxicity may also be produced by diverse isolates. Important for entomotoxicity of Bti are the cytolytic (Cyt) toxins they produce, which synergize activity of Cry proteins against mosquito larvae (Wu and Chang 1985). On the other hand, potential secretion during the vegetative stage of some strains of thermostable insecticidal toxins called β-exotoxins is of concern during commercial production of Bt-based biopesticides. The Bacillus species commonly recognized as insect pathogens are B. thuringiensis, B. popilliae, B. lentimorbus, B. larvae, and certain strains of B. sphaericus (Stahly et al. 2006). Apart from Bacilli, there are further non-Bacillus EB such as Clostridium, Serratia, Photorhabdus, Xenorahbdus and Pseudomonas (Boemare and Tailliez 2009). The red-pigmented varieties of S. marcescens are most often reported as pathogens of insects. However, non-chromogenic strains of S. marcescens, for example S. marcescens Bizio, are also pathogenic. In the last few years, approximately 60 pathogenic bacterial species and their products have been developed as biopesticides, worldwide. These various pathogens are being used successfully in the biological control of insects (Betz et al. 2000). Bacteria and insects have co-evolved a wide range of complex relationships from commensalism to parasitism or pathogenesis over more than 250 million years. The specific host-bacteria relationships that have developed are the outcome of dynamic co-evolution underpinned by genetic diversity and driven by selection pressure. While opportunistic bacterial infections occur, often as a result of injury or stress, most consistent EB have obligate or facultative relationships with their hosts. EB occupy many niches, functioning as rhizosphere and phyllosphere colonizers, both

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obligate and facultative pathogens and as endophytes of plants. The main ecological niche of some species is primarily as pathogen, surviving between hosts as environmentally resistant spores. It has been proposed that Bt evolved in commensal relationships with plants to provide defence against herbivores’ attacks (Smith and Couche 1991). This “bodyguard theory” states that microbe–plant symbiosis has evolved so that microorganisms that are insect antagonistic are recruited and/or maintained by the plant for protection. It has been supported by recent ecological and molecular studies on Bt demonstrating rhizosphere competence (Vidal-Quist et al. 2013), endophytic ability (Tao et al. 2014) and even plant-mediated volatile attraction. Genetic variation in bacteria originates from mutation and selection or acquisition of genetic material from the environment (transformation), bacteriophages (transduction) or other bacteria (conjugation). These processes, combined with the rapid generation time of bacteria, result in a high level of variation and wide ranges of functionality among strains. Horizontal gene transfer of mobile genome fragments known as genomic islands also increases genetic variation among populations. Genetic information is also sometimes stored in a number of plasmids, small and usually circular DNA molecules that are self-replicating and that can be transferred between bacteria through a process known as conjugation. The plasmids usually contain genes that are crucial for specific functions, including pathogenicity. For instance, most crystal toxins from Bt are located in plasmids (Held et al. 1982), which can be exchanged between diverse strains and Bacillus spp. (González et al. 1982). The pathogenicity island of Serratia spp. containing tc genes is also located on a plasmid (Dodd et  al. 2006). Genetic information can also be stored in prophages, DNA from bacteriophages that is inserted into the bacterial chromosome or plasmid through transduction, which can also confer phenotypes conducive to pathogenicity. The ecology of B. thuringiensis and L. sphaericus and the evolutionary advantage of producing insecticidal crystalline protein inclusions are still matters of debate. Supporting the identity of these bacteria as bona fide insect pathogens is their commitment of relevant energy resources to production of high amounts of insecticidal parasporal proteins. However, although spores of B. thuringiensis are very persistent in the environment, the bacterium displays low rates of horizontal transmission between hosts, with rare epizootics. Horizontal transmission of B. thuringiensis was reported to be tightly related to the production of urease (Martin et al. 2009), but this is not observed in commercially used strains. Thus, secondary infections are uncommon after spray with biopesticides based on Btk and subsp. aizawai (Smith and Barry 1998), which are urease positive (Martin et al. 2010). In contrast, persistent activity was reported for applications of biopesticides containing the urease-negative Bti (Tilquin et al. 2008). Persistence of L. sphaericus biopesticides is related to the low settling of L. sphaericus spores in the water column (Nicolas et al. 1987) and recycling in the host (Charles and Nicolas 1986). The biological control paradigm changed when the potential of EB was discovered, especially associated to species belonging to the genus Bacillus (Glare and O’Callaghan 2000). Initially, the species Paenibacillus (former Bacillus) popilliae Dutky was introduced for the management of the

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Japanese Beetle Popillia japonica Newman (Steinhaus 1975), but more concrete results were achieved with the discovery of new Bt strains showing high toxicity against specific insects at a competitive level, compared to conventional insecticides, in terms of efficacy and costs of production. The strain HD-1, belonging to subsp. kurstaki (De Barjac and Lemille 1970), soon became the main commercial focus for the management of lepidopteran pests in agriculture and forestry. Beside it, other strains are actually commercially available, such as SA-11, SA-12, PB 54, ABTS-351 and EG2348, all isolated from insects or soil and expressing a range of different toxins mostly belonging to the Cry1 and Cry2 families. Subsequently, the discovery of a Bt strain belonging to the subsp. israelensis (Bti) was followed by its commercialization for the management of mosquitoes and simulids (Goldberg and Margalit 1977). Then, a particularly active strain of the subsp. tenebrionis was discovered and employed against Coleoptera (Krieg et  al. 1983). Homology among toxins from B. sphaericus and Bt or other EB has been shown, demonstrating their phylogenetic relationships and revealing a probable common co-evolution (de Maagd et al. 2003).

3.4.3  Genomics and Phylogeny Bacteria are characterized by one single chromosome within the nucleoid, whose size varies among species. For example, the bacterial genomes of B. thuringiensis (the widely studied insect pathogen), B. cereus and B. anthracis are around 5.4 Mb (Carlson and Kolstø 1993; Ivanova et al. 2003). Other Bacillus spp. such as B. subtilis and B. licheniformis have a smaller chromosome (4.2 Mb) (Kunst et al. 1997; Rey et al. 2004). While the phenotypes of these species are different, their intra and inter phylogenetic relationships are not clear. Several approaches have been used to classify B. thuringiensis strains, including rRNA gene sequences, amplified fragment length polymorphisms (AFLP), restriction fragment length polymorphisms (RFLPs) in small subunit (SSU) rRNA sequences, GryB (gyrase subunit B) and AroE (shikimate-5-dehydrogenase) gene sequences. The results suggest that there is a high level of sequence homology among strains of B. thuringiensis. Similarly, overall genetic studies have shown that B. thuringiensis and B. cereus are essentially identical (Helgason et al. 1998). Bacillus anthracis can only be distinguished from B. thuringiensis and B. cereus through microbiological and biochemical tests. Since these genetic methods are not able to easily distinguish different members of B. thuringiensis, B. anthracis and B. cereus, it becomes necessary to look for some more easily recognizable markers. With the advent and development of next generation sequencing technologies, a great deal of sequencing data has been generated in recent years, as the whole-genome-sequencing gives information on the organization of the bacterial genomes. Genome sequencing is informative about the chromosome, which can be linear or circular, the presence of plasmids and the genome size and variation, within a species (Casjens 1998). It also provides information on new operons, origin of replication and genome polarity. This increases with the growing

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number of sequenced genomes. Finally, genome sequencing allows the characterization of guanine-cytosine content (GC) and its variation, genomic islands and synteny with others organisms. All these data give information on DNA regions acquired by horizontal gene transfer and therefore on the evolutionary history of the bacterium. The rapid accumulation of whole genome data of Bacillus species in Genbank makes it possible the comparison of genomic differences over the entire genome that cannot be identified in analyses of specific, single gene sequences. However, the size of the whole genome data poses great challenges on alignmentbased algorithms, which are effective in dealing with closely related sequences but are unable to evaluate the recombination, shuffling and rearrangement events of the whole genomes. Thus, alignment-free sequence analysis approaches, such as Feature Frequency Profile (FFP), provide attractive alternatives over alignmentbased approaches. FFP is a new method used to study the whole genome phylogeny based on k–mers (Jun et al. 2010). In this method, the number of features of a particular length that occur in a particular genome is counted and assembled into a FFP vector. FFPs from different species are then compared using the Jensen–Shannon (JS) Divergence (Lin 1991). A neighbor-joining phylogenetic tree can thus be constructed based on the resulting distance matrix. Compared to the traditional multiple sequences alignment (MSA) based method, the alignment free FFP method can compare both genic and non-genic regions of the whole genome at higher speed. It can incorporate a wide variety of genomic features into each comparison including intron deletions, exon sequence indels, transposable element insertions, base transversions in coding sequences and some rare genomic changes such as short interspersed element/long interspersed element (SINE/LINE) insertions (Sims et  al. 2009). Wang and Ash (2015) reconstructed the whole-genome phylogeny of Bacillus species using the FFP approach, with an aim to better understand the phylogenetic relationships among them. Fifty complete Bacillus genome sequences and associated plasmids were compared using the FFP method (Fig. 3.4). The resulting wholegenome phylogeny supports the placement of three Bacillus species (B. thuringiensis, B. anthracis and B. cereus) as a single clade. These results clearly suggest the close relationship among B. thuringiensis, B. anthracis and B. cereus species, are in agreement with earlier results from DNA-DNA hybridization analysis and Multi Locus Enzyme Electrophoresis (MEE), which showed high identity among them (Priest et  al. 1994). These three species have been grouped under the name of Bacillus cereus sensu lato (Rasko et al. 2005) despite their obvious difference in phenotype and pathological effects, which result from the genetic difference in plasmid rather than in chromosome. The B. weihenstephanensis strain KBAB4 was found to be very closely grouped with the major cluster I-d consisting of all B. thuringiensis isolates and proximal to cluster I-e (B. cereus) and cluster I-c (a cluster containing both B. thuringiensis and B. cereus strains). FFP proved to be more effective in inferring the phylogeny of Bacillus than methods based on single gene sequences [16s rRNA gene, GryB (gyrase subunit B) and AroE (shikimate-5-dehydrogenase)] analyses. Furthermore, the availability and reduced cost of whole genome sequencing can be used without extensive gene annotation to provide robust phylogenetic analysis of new isolates as they become

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Fig. 3.4  Phylogenetic tree of 50 Bacillus strains, constructed using the NJ algorithm based on the FFP features of the Whole Genome Data. Escherichia coli Bl21 (DE3) (AM946981.2) was used as an outgroup in the analysis. The bootstrap confidence values were generated using 1000 permutations. Different symbols were allocated to represent different species: Blue triangle for B. thuringiensis; Pink diamond for Bacillus cereus; Red circle for Bacillus anthracis; Green Square for Bacillus subtilis. (Source: Wang and Ash 2015)

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available. Entomopathogenic nematode symbiotic bacteria Photorhabdus luminescens has a genome size of 5.7  Mb (Duchaud et  al. 2003), whereas Xenorhabdus bovienii and X. nematophila, other nematode symbiotic bacteria, are estimated to have a genome size of approximately 4.3 Mb (http://www.xenorhabdus.org/). Many bacteria also contain extrachromosomal elements (plasmids), which are smaller double-stranded DNA (dsDNA) molecules that replicate independent of the chromosomal DNA. Genes located in bacterial plasmids usually code for proteins that determine specific phenotypes, but do not code for products needed for bacterial survival and growth. Bacterial genes are organized in operons or cassettes that consist of a promoter, a series of genes and a transcription terminator. The most important genes studied for EB are the Cry genes that code for insecticidal crystal proteins. They are usually found in large transmissible plasmids or more rarely in the chromosome. These proteins show entomopathogenic properties to insects from orders Lepidoptera, Diptera and Coleoptera (WHO 1999). Different combinations of Cry genes are found in various Bt strains including those with one, two or even four different genes (Lereclus et al. 1993). More than 200 toxin genes with pesticidal activity have been cloned from a wide range of B. thuringiensis strains (Table 3.5), grouped into 80 classes (Crickmore et al. 1998). Sequence analysis of these genes is currently considered not only relevant to the classification of bacteria but also for interpreting evolutionary relationships among taxa (Crickmore 2000). Bacterial genome studies have provided

Table 3.5 Some B. thuringiensis toxins active against different insect pest orders Order and Insects Lepidoptera Cydia pomonella Trichoplusia ni Spodoptera exigua Ariogeia rapae Bombyx mori Hyphantria cunea Plutella xylostella Manduca sexta Spodoptera frugiperda Spodoptera littoralis Helicoverpa zea Heliothis virescens Ostrinia nubilalis Spodoptera litura Lymantria disper Galleria mellonella

Active Toxin Cry1Aa, Cry1Ab, Cry2Aa2, Cry15Aa1 Cry1Aa, Cry1Ab, Cry1Ac, Cry1Ba, Cry1Ca, Cry1Da, Cry1Ea, Cry1Fa, Cry1Ja, Cry2Aa1, Cry2Ab2, Cry2Ac1 Cry1Ab1, Cry1Ba1, Cry1Bd1, Cry1C, Cry1Ca1, Cry1Fa1, Cry1Gb1, Cry1Ka1 Cry1Ba1, Cry1Ka1 Cry1Ba1, Cry1Ia1, Cry1Ka1 Cry1Ba1, Cry1Ka1 Cry1Ba1, Cry1Bd1, Cry1Ea2, Cry1Gb1, Cry1Ia1, Cry1Ka1, Cry2Ab1 Cry1C, Cry1Ea1, Cry1Ea4, Cry2Ac1, Cyt1Aa2, Cyt2Aa1 Cry1C Cry1C, Cry1Ca2, Cry1D, Cry1Ea1, Cry1Fa1 Cry1Fa1, Cry2Aa1, Cry2Ab2 Cry1Fa1, Cry2Aa1, Cry2Ab2, Cry2Ac1 Cry1Fa1, Cry2Aa1, Cry2Ab2 Cry1Ia1 Cry2Aa1, Cry2Ab, Cry2Ab2 Cry9Aa1 (continued)

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Table 3.5 (continued) Order and Insects Coleoptera Diabrotica undecimpunctata Leptinotarsa decemlineata Cotinis spp. Cyclocephala sp. Popillia japonica Chrysomela scripta Diptera Musca domestica Aedes aegypti

Active Toxin Cry3Ba1, Cry3Bb1 Cry3Ba1, Cry3Bb1, Cry8Aa1 Cry8Ba1 Cry8Ba1 Cry8Ba1 Cyt1Aa4

Cry1Bd1, Cry1Gb1 Cry1C, Cry2Aa1, Cry2Ab, Cry2Ab2, Cry2Ac1, Cry4Aa, Cry4Aa1, Cry4Ba, Cry4Ba1, Cry10Aa1, Cry11Aa, Cry11Aa1, Cry17Aa1, Cry19A, Cry20Aa1, Cry27Aa1, Cyt1Aa2, Cyt1Aa4, Cyt1Ab1, Cyt2Aa1 Anopheles stephensi Cry4Aa, Cry11Aa, Cry17Aa1, Cry19A, Cry27Aa1, Cyt1Ab1 Culex pipiens Cry4Aa, Cry4Ba, Cry11Aa, Cry17Aa1, Cry19A, Cry27Aa1, Cyt1Aa2, Cyt1Ab1, Cyt2Aa1 Anopheles gambiae Cry4Aa1, Cry4Ba, Cyt1Aa2, Cyt2Aa1 Culex Cry4Aa1, Cry4Ba, Cry11Aa, Cry11Ba1, Cyt1Aa4 quinquefasciatus Callíphora stygia Cyt1Aa2, Cyt2Aa Lucilia cuprina Cyt1Aa2, Cyt2Aa Lucilia sericata Cyt1Aa2, Cyt2Aa Hymenoptera Acromyrmex Cry1, Cry9 Diprion pini Cry5A, Cry5B Cephacia abietis Cry5A, Cry5B Nematodes Caenorhabditis Cry5Aa1, Cry5Ab1 elegans Pratylenchus spp. Cry5Aa1, Cry5Ab1, Cry6Ba1, Cry12Aa1 Fasciola hepatica Cry5Ab1 Panagrellus Cry6Aa1 redivivus Pratylenchus Cry6Aa1 scribneri Source: Hernandez-Fernandez and Lopez-Pazos (2011)

extremely valuable information regarding the genetic diversity of species. Not only have they assisted in resolving taxonomies at various levels, but also contributed in assessing evolutionary relationships. EB are distributed in phylogenetically diverse groups of prokaryotes. Gram-­ negative EB include: (i) members of the Enterorbacteriaceae such as Serratia entomophila, S. marcescens, S. proteamaculans and the nematophilic genera

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Photorhabdus and Xenorhabdus; and (ii) members of the Pseudomonadaceae such as Pseudomonas entomophila. Gram positive EB comprise the most studied group of bacteria in insect pathology and include members of Bacillus (Bacillaceae). This genus has recently been submitted to several taxonomic revisions and a number of new genera such as Brevibacillus, Lysinibacillus and Paenibacillus have been recognized from species previously described in Bacillus (Ash et al. 1993; Shida et al. 1996; Ahmed et al. 2007). Since the 1970s, ribosomal RNA (rRNA) molecules have been considered for studying bacterial molecular genealogies. These molecules are universally distributed, easily sequenced and carry generally useful phylogenetic information. Comparison of rRNA sequences enabled notably the discovery of the domain Archaea (Balch et al. 1977), a branch of prokaryotes more closely related to Eukarya than to the other prokaryotes (Eubacteria) (Iwabe et al. 1989). Sequences of 16S rRNA gene have been considered to assess evolutionary relationship of Serratia spp. (Dauga et al. 1990; Ashelford et al. 2002) and other lineages. They have also helped to resolve taxonomic conflicts, particularly for identification of new isolates, previously classified in this genus using basic phenotypic characters. Pseudomonas entomophila shares a common ancestor with P. putida biovar A (bootstrap value of 94% indicating a high robustness of the node). The genomes of these two species are also very similar (Vodovar et al. 2006), as P. entomophila may have also acquired insect virulence factors through lateral gene transfer. Furthermore, analysis of the P. entomophila genome confirmed the presence of genes that encode insecticidal toxin complexes also found in entomopathogenic enterobacteria such as Photorhabdus luminescens, S. entomophila, Xenorhabdus nematophila and Yersinia spp. (Bowen et al. 1998; Waterfield et al. 2001). Protein-coding genes have been suggested as an alternative option to avoid some of the problems encountered in phylogenetic studies with prokaryotes (Hedegaard et  al. 1999; Lerat et  al. 2003). Recently, two protein-coding gene sequences, gyrB and recA, have been considered for assessing evolutionary relationships in various groups of bacteria, including the entomopathogenic types (Eisen 1995; Dauga 2002). Akhurst et al. (2004) considered gyrB gene sequences to assess evolutionary relationships among Photorhabdus spp. Three Photorhabdus groups can be recognized based on the phylogenetic trees of gyrB gene sequences: the P. luminescens group, represented by subspecies P. luminescens ssp. laumondii, P. luminescens ssp. kayaii, P. luminescens ssp. luminescens and P. luminescens ssp. akhurstii, together with an unidentified strain C8404. The P. asymbiotica group comprises two subspecies: P. asymbiotica ssp. asymbiotica, P. asymbiotica ssp. australis, Photorhabdus strain Q614 isolated from an uncharacterized Heterorhabditis spp. from Queensland, Australia (Akhurst and Boemare 1986) and strains Cbkj163 and Onlr40 isolated from Heterorhabditis indica isolates from Japan. The P. temperata group includes two subspecies: P. temperata ssp. temperata and P. luminescens ssp. thracensis (strain DSM15199T). The phylogenetic position of this latter strain, whatever the method of reconstruction used, is not congruent with its taxonomic classification as P. luminescens based on 16S rRNA gene sequences comparison (Hazir et al. 2004). The distance and maximum likelihood analyses suggest a com-

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mon ancestor for P. asymbiotica and the P. luminescens groups; whereas maximum parsimony analysis suggests a common ancestor for P. asymbiotica and the P. temperata groups.

3.5  Entomopathogenic Fungi (EF) 3.5.1  Origin, Natural History and Geographical Distribution EF are a group of phylogenetically diverse, heterotrophic, eukaryotic, unicellular or multicellular (filaments) microorganisms that reproduce via sexual or asexual spores, or both. They have chitinized cells and are generally non-mobile (Badii and Abreu 2006). Approximately 80% of the diseases that occur in insects have a fungus as the causative agent. Practically all insects are susceptible to some diseases caused by fungi, which can lead to death (Batista 1989). The kingdom Fungi is one of the major groups of eukaryotic microorganisms in terrestrial and aquatic ecosystems (Mueller and Schmit 2007), with approximately 100,000 described species (Kirk et al. 2008a), that represent only a fraction of the estimated diversity, considered to range between 1.5 and 5 million species (Blackwell 2011). The evolution of fungi has been going on since they diverged from other ancestors around 1.5 billion years ago (Wang et al. 1999). Fungi probably colonized land during the Cambrian, over 500 million years ago, but terrestrial fossils only become uncontroversial and common during the Devonian, 400 million years ago (Lucking et  al. 2009). It is considered that animals and fungi evolved separately from a common ancestor long ago. Although it is believed that fungi first evolved in aquatic environments, the best fossil evidence suggests that they arose on land. Analyses with molecular phylogenetics support a monophyletic origin of fungi (Hibbett et al. 2007). Insects are known to form intimate relationships with many fungal groups that include: mutualistic endosymbionts that assist in nutrition (Suh et al. 2005), fungi as food sources that are farmed as crops by leaf cutter ants (Currie et al. 2003), vertically transmitted parasites (Lucarotti and Klein 1988), commensals (De Kesel 1996) and pathogens with pronounced effects on host populations (Evans and Samson 1982, 1984). Fungal associations involve members of Coleoptera, Diptera, Homoptera, Hymenoptera and Isoptera as well as others. The fungi may be clustered taxonomically, as is the case of Ascomycetes in the Hypocreales (e.g., Beauveria, Metarhizium, Fusarium), ambrosia fungi in genera Ophiostoma and Ceratocystis and their asexual relatives, Laboulbeniomycetes, Saccharomycetes and the basal Microsporidia. Other groups, however, only have occasional members (e.g., mushrooms cultivated by attine ants and termites) in such associations. These fungi usually attach to the external body surface of insects in the form of microscopic spores (usually asexual, mitosporic spores also called conidia). Under the right conditions of temperature and humidity (usually high), these spores germinate,

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grow as hyphae and colonize the insects' cuticle penetrating through the cuticle or through body openings, by enzymatic hydrolysis, thus reaching the host body cavity (hemocoel) ((Tanada and Kaya 1993; Fernandes et al. 2012). Then, the cells proliferate in the host, usually as walled hyphae or in the form of wall-less protoplasts (depending on the EF species involved). After some time the insect is usually killed (sometimes by fungal toxins) and new propagules (spores) are formed inside or on the insect body, if the environmental conditions are again favourable. High humidity is usually required for sporulation. EF have evolved specialized mechanisms for the enzymatic degradation of the integument and for overcoming insect defence compounds. The relationships by which different fungal species obtain energy from their hosts (i.e., their eco-­ nutritional mode) include biotrophy (nutrition derived only from living cells, which ceases once the cells die), necrotrophy (killing and utilization of dead tissues) and hemibiotrophy (initially biotrophic and then becoming necrotrophic). Following entry, some groups (i.e., Metarhizium and Beauveria in the order Hypocreales, phylum Ascomycota) are known to grow inside the host as yeast-like hyphal bodies, multiplying by budding (Prasertphon and Tanada 1968). Others, for example, some species within the Entomophthoromycota, produce protoplasts (cells without cell walls) instead (Butt et  al. 1996). A third group encompassing some species within Oomycota, Chytridiomycota and the genus Entomophthora, that infect aphids, are known to grow directly as hyphal filaments inside the host (Lucarotti and Shoulkamy 2000). Most EF kill their hosts before the spore production starts (as such they are termed hemibiotrophic). A few of them, especially some in the phylum Entomophthoromycota, sporulate from the living body of their hosts (and as such are termed biotrophic) (Roy et al. 2006). All entomopathogenic oomycetes kill the host before transmission. There are about 750 species of fungi that cause infections in insects or mites. As a group, they attack a wide range of insect and mite species, but individual species and strains of fungus are very specific. Host death requires between 4 and 10 days, depending on the type of fungus and the number of infecting spores. After host death, the fungus produces thousand new spores on the dead body, which disperse and continue their life cycle on new hosts. Oomycete infections have been recorded from mosquito larvae in freshwater, primarily in well-aerated streams, rivers, ponds, lakes (Alexopoulos et al. 1996), and even treeholes (Saunders et al. 1988) or water collected on leaf axils (Frances et al. 1989). A single example of oomycetes infecting a nondipteran was Crypticola entomophaga, which was described attacking caddis flies (Trichoptera), which are also aquatic. The earliest known fossil of an insect pathogenic fungus, an Ophiocordyceps-­ like fungus infecting a scale insect (Hemiptera) is from Myanmar amber (100– 110 million years ago) (Sung et al. 2008). Later fossils include an Entomophthora-like fungus infecting a termite (Isoptera) and a Beauveria infecting an ant (Hymenoptera), both from Dominican amber (20–30 million years ago) (Poinar and Thomas 1982). Agostino Bassi described Beauveria bassiana in 1835 as the cause of the devastating muscardine disease of silkworms and it was instrumental in his development of

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the germ theory of disease (Steinhaus 1956b). In 1880, Elie Metchnikoff was among the first to propose practical methods of microbial control of an insect crop pest, initiating trials of the fungus Metarhizium anisopliae against grain beetles (Lord 2005). The approximate number of described insect pathogenic species varies from just a few in the Chytridiomycota, Blastocladiomycota, Kickxellomycotina and Basidiomycota to a substantial number in the Ascomycota and almost complete dominance in Entomophthoromycota. The phylogenetic groups of Ascomycota that harbor most of the known insect pathogens are the Hypocreales and the Onygenales. Insect pathogenic life forms evolved repeatedly in the Ascomycota (Spatafora et al. 2007). Molecular phylogenies have demonstrated that Fungi, which traditionally have been grouped with plants, are more closely related to the Metazoa or animals (Baldauf et  al. 2000; Berbee and Taylor 2001; Lang et  al. 2002). Fungi are thus likely to have evolved from a flagellated ancestor (Cavalier-Smith 2001), which bolsters the long-held assumption that Chytrids, which produce flagellated zoospores, represent the earliest surviving lineages of fungi and point to an aquatic origin for the fungi. Clades of insect pathogens have also originated in both phyla of the subkingdom Dikarya, the Basidiomycota and Ascomycota. The Basidiomycota includes only a single group of insect pathogens, the monophyletic Septobasidiobasidiaceae, which are classified in the subphylum Pucciniomycotina or rust clade (Aime et al. 2006). Three notable radiations of insect pathogens have originated within Ascomycota (James et al. 2006), within the Eurotiomycetes and Sordariomycetes and in the singular class Laboulbeniomycetes. The Eurotiomycete genus Ascosphaera is an obligate pathogen of bees and A. aphis in the well-known causal agent of chalkbrood disease of honeybee larvae (Apis mellifera). Interestingly, honeybee defences towards A. apis appear to have a ‘lock and- key’ dynamic, leading to suggestions that this pathogen has pushed bees, over evolutionary time frames, towards lower nest-level relatedness (Tarpy and Seeley 2006; Seeley and Tarpy 2007). There are 29 described species of Ascosphaera that infect various bee species, each of which generates pathologies similar to that of A. apis. A ribosomal phylogeny of selected species of Ascosphaera supports the monophyly of the genus and provides support for traditional species concepts (Anderson et  al. 1998). A whole genome sequence, the first for any fungal entomopathogen, is available (Qin et al. 2006) providing an unique opportunity to analyse the genetic architecture of a specialized entomopathogen. EF are not a monophyletic group, but rather a heterogeneous assemblage of fungi independently derived from Ascomycota, Basidiomycota, Entomophthoromycotina, Blastocladiales, Kickxellomycotina, Neocallimastigomycota and Microsporidia. There are only a few exceptions, such as the Glomeromycota, which contain only mycorrhizal clades. Nutritional modes of the earliest fungi remain poorly understood, but the ability to infect insects is clearly represented in several basally diverging clades, e.g., Blastocladiomycota, Chytridiomycota, and Entomophthoromycota (Boomsma et al. 2014).

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EF are particularly well suited for development as biopesticides because, unlike bacteria and viruses that have to be ingested to elicit disease, fungi are contact insecticides and typically infect insects by directly penetrating their surface (cuticle) and multiplying in the hemocoel. Approximately 170 pest control products have been developed based on at least 12 species of fungal entomopathogens (De Faria and Wraight 2007). Anamorphic EF, such as B. bassiana and M. anisopliae, are usually developed as inundative control agents which are applied en masse to a pest population and there is little expectation that they will persist and reproduce within the biotic environment. As a consequence, research on these fungi has tended to concentrate on technical aspects of biopesticide development (mass production, formulation, application, response to abiotic variables etc) with significantly less work done on understanding their basic ecology. Anamorphic EF are naturally widespread, particularly in soil and yet little is known about the factors that influence their distribution, population structure, econutritional behaviour and the evolution of virulence related characteristics. In contrast, researchers studying entomophthoralean fungi, which are not easy to mass produce, have focused on the ecology of these organisms and their role as causative agents of natural epizootics. Identifying fungal fitness traits and the selective forces that act upon them will improve our understanding of how and why EF interact with their hosts. Current theories on the evolution of virulence in micro-parasites, which at present are not being applied to EF, could provide fresh insights into their ecology and exploitation for biocontrol. At the same time, the wide arrays of experiments that can be done with EF are ideal for answering basic questions in parasitology and entomology.

3.5.2  Taxonomy and Evolution Fungi are more closely related to animals than to plants and are placed with the animals in the monophyletic group of Opisthokonts (Shalchian-Tabrizi et al. 2008). Fungal species can have multiple scientific names depending on their life cycle and mode of reproduction (sexual or asexual). There are fungi that invade dead insects, called saprotrophs and fungi that infect living insects called entomophagous (Butt et al. 2006). Of the estimated 1.5 to 5.1 million species of fungi in the world (Hibbett et al. 2007), approximately 750 to 1,000 are fungal entomopathogens placed in over 100 genera (St. Leger and Wang 2010). Fungal entomopathogens, thus, constitute the largest number of taxa that are insect pathogens (Ignoffo 1973). De Faria and Wraight (2007) identified 171 fungal-based products used as biocontrol agents since the 1960s, most of them based on B. bassiana, B. brongniartii, M. anisopliae and Isaria fumosorosea. The 2007 classification of Kingdom Fungi is the result of large-scale collaborative research efforts involving dozens of mycologists and other scientists working on fungal taxonomy (Hibbett et  al. 2007). It recognizes seven phyla which include Microsporidia, Chytridiomycota, Blastocladiomycota, Neocallimastigomycota, Glomeromycota, Ascomycota and Basidiomycota (Hibbett et  al. 2007) (Fig.  3.5). Out of these, two phyla  - the Ascomycota and the

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Fig. 3.5  Phyla of fungi (based on Hibbett et al. 2007) indicate that fungi are more diverse than previously appreciated. Major changes include separation of groups with flagellated cells (Chytrids) in three phyla and separation of zygosporic fungi (Zygomycetes) in at least three lineages. Numbers of described fungal phyla are from Kirk et al. (2008b) and for the outgroup from The IUCN Red List of Threatened Species (http://www.iucnredlist.org/static/stats). (Source: Vega et al. 2009)

Basidiomycota are contained within a branch representing subkingdom Dikarya, the most abundant in species and familiar group. Fungal species of different phyla like Microsporidia, Chytridiomycota, Entomophthoromycota, Basidiomycota and Ascomycota are known to infect and kill insects (Sung et  al. 2007; Shang et  al. 2015). The two best-studied groups are the ascomycete entomopathogens and the Entomophthoromycota. The most well-studied insect ascomycete pathogens fall into three families within the order Hypocreales: Cordycipitaceae, Clavicipitaceae and Ophiocordycipitaceae. Many common and important EF is in order Hypocreales

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of Ascomycota: the asexual (anamorph) phases Beauveria, Isaria (Paecilomyces), Hirsutella, Metarhizium, Nomuraea and the sexual (teleomorph) Cordyceps; others such as Entomophthora, Zoophthora, Pandora, Entomophaga belong to the order Entomophthorales (Zygomycota). Over the past 400 million years fungi and insects have coevolved a wide array of intimate interactions (Araujo and Hughes 2016), that include mutualistic endosymbiosis (Suh et  al. 2001), fungi as obligate food sources, such as those found in fungus-­gardening ants (Mueller et al. 2005), sexually and behaviourally transmitted parasites, such as Laboulbeniales (De Kesel 1996) and the most common disease-­ causing agents of insects (Roberts and Leger 2004). Entomopathogenicity evolved independently and repeatedly in all the major phyla of the Kingdom Fungi (Araujo and Hughes 2016). The EF have been traditionally considered as important mortality factors for insects. There are two main taxonomic orders of entomopathogenic fungi. The Entomophthorales from phylum Zygomycota include important genera such as Pandora, Entomophthora and Conidiobolus. Many of their species are co-evolved, obligate pathogens that show specific eco-morphological adaptations to the life cycles of their hosts, such as the production of forcibly-ejected infective spores that are produced on insect cadavers during the night, when environmental conditions are most conducive to infection. These fungi often cause natural epizootics in insect and mite populations. Recently, molecular methods have revealed the telemorph connections of these fungi. Molecular studies are also shedding light on the phylogenetic relationships with other fungi. Data showed that entomopathogenicity has evolved multiple times in the ascomycete fungi, characterised by host shifting from an intimate association with plants to insects and vice versa. Recently, molecular tools such as DNA sequence analysis have led to a new phylogenetic classification of fungi that challenged many assumptions about their relationships with other fungi. This new phylogeny is already leading to significant new insights that should allow us to better understand the EF ecology. In addition, it has been discovered recently that many EF play additional roles in nature, as plant endophytes, antagonists of plant pathogens, beneficial rhizosphere-associates and possibly even plant growth promoters. Comparative genomics utilizing the Ophiocordyceps sinensis genome provided an unparalleled opportunity to develop a deeper understanding of how this unique pathogen interacts with insects within its ecosystem. It is clear that host-pathogen interactions are a major driving force for diversification, but the genomic basis for speciation and host shifting remains still unclear. The genus Metarhizium has been subdivided into 12 different species according to the sequences of several genes (Bischoff et al. 2009). Some of these species have a wide host range, whereas others show specificity for certain insect families and can be used to test hypotheses on speciation and host specificity. Comparative genomic analyses of seven species revealed a directional speciation continuum from specialists with narrow host ranges (i.e. M. album and M. acridum specific to hemipterans and acridids, respectively), to transitional species with intermediate host ranges (Metarhizium majus and M. guizhouense both have host ranges limited to two insect orders), and then to

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generalists (i.e., M. anisopliae, M. robertsii and M. brunneum) (Hu et  al. 2014). Besides host range, generalist and specialist Metarhizium spp. differ in the way they colonize hosts (Kershaw et al. 1999). Generalists, like M. robertsii, typically kill hosts quickly via toxins and grow saprophytically in the cadaver. In contrast, the specialist M. acridum causes a systemic infection of host tissues before the host dies. This may reflect greater adaptation by the specialists to subverting or evading the immune systems of their particular hosts so they do not need to kill quickly. Generalists also have mechanisms for evading host immunity, but appear less able to subvert immune responses specific to certain hosts. Lack of specific adaptations could have selected for rapid killing of hosts before the host can mount an enfeebling immune response. Sequencing related entomopathogen species that have evolved specialist or generalist lifestyles has increased their utility as models and provided insights into the evolution of pathogenicity. Thus, cross-species comparative analysis identified novel and specialized virulence mechanisms and, compared to experimental methods, has allowed for more rapid identification of genes encoding biologically active molecules and genes responsible for interactions between fungi, plants and insects. Undoubtedly, the information from comparative genomics will benefit future functional studies of insect-fungus interactions.

3.5.3  Genomics and Phylogeny Fungi are the most common pathogens of insects and crucially regulate insect populations. The rapid advance of genome technologies has revolutionized our understanding of entomopathogenic fungi with multiple Metarhizium spp. sequenced, as well as Beauveria bassiana, Cordyceps militaris and Ophiocordyceps sinensis among others. Comparative genomics offers a way forward by disentangling common themes of fungal biology, from specific components involved in insect pathology and allowing broad host range pathogens to be studied in the context of narrow host range pathogens (Wang and St. Leger 2014). It is also extremely valuable for assessing poorly characterized species such as Ophiocordyceps sinensis. Comparative genomics has facilitated identifying fungal fitness traits and the selective forces that act upon them to improve our understanding of how and why EF interact with insects and other components in their environments. Thus, sequence data provide crucial information on the poorly understood ways that these organisms reproduce and persist in different environments. Alongside the recent availability of genomic resources, the wide array of experiments that can be performed with EF make them ideal models for answering basic questions on the genetic and genomic processes behind adaptive phenotypes. Key challenges for fungi as models for other eukaryotes include identifying the genes involved in ecologically relevant traits and understanding the nature, timing and architecture of the genomic changes governing the origin and processes of local adaptation (Gladieux et al. 2014). As of February 2016, one published Entomophthoromycota genome (Conidiobolus

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coronatus (Chang et al. 2015) and nine incomplete Entomophthoromycota genome sequencing projects were listed in the Genomes OnLine Database (GOLD) (Licht et al. 2016). To date, there is much more genomic information on ascomycete insect pathogens, as sequences are available from nine Metarhizium strains (Gao et  al. 2011; Hu et al. 2014; Pattemore et al. 2014; Staats et al. 2014), Beauveria bassiana (Xiao et al. 2012), Cordyceps militaris (Zheng et al. 2011), Ophiocordyceps sinensis (anamorph, Hirsutella sinensis) (Hu et al. 2013), Ophiocordyceps unilateralis (de Bekker et al. 2015), Tolypocladium inflatum (Bushley et al. 2013) and Hirsutella thompsonii (Agrawal et al. 2015). Entomophthoromycota boasts one of the largest fungal genomes ever measured, at 8000  Mb in E. aulicae, which correlates with microscopic observations of extensive condensed chromatin in the nuclei (Murrin et al. 1986). Also, Basidiobolus ranarum has a large haploid genome of 350 Mb (Henk and Fisher 2012), compared to an average genome size of around 40 Mb in the kingdom Fungi (Gregory et  al. 2007; Henk and Fisher 2012). Earlier microscopic analyses have found numerous chromosomes in B. ranarum, ranging from 60 to more than 500 (Sun and Bowen 1972), but it is unclear whether ploidy level or genome duplication governs genome size in B. ranarum (Henk and Fisher 2012). In general, entomophthoromycotan genomes are considered to be haploid (Humber 2012; Gryganskyi and Muszewska 2014) and the basal chromosome count in Entomophthoromycota appears to be 8, whereas 12, 16 and 32 have also been estimated (Riddle 1906; Sawyer 1933; Humber 1982). The large nuclei with numerous chromosomes and condensed chromatin seen in several species suggest that large genomes may be an ancestral trait within Entomophthoromycota, but less than 10 out of 280 species have been analyzed for either chromosome count or genome size, making it unclear whether having a large genome is unusual or the norm within Entomophthoromycota. However, the first genome assembly (ver 1.0) within the Entomophthoromycota was the phylogenetically basal Conidiobolus coronatus, which was sequenced with 454-technology by the US Department of Energy Joint Genome Initiative (JGI). C. coronatus has 10,635 predicted genes and a genome size of 39.9 Mb (Chang et al. 2015), which is similar to the average fungal genome size of 40  Mb. The genus Conidiobolus is paraphyletic, consisting of one clade, exemplified by C. coronatus, including soil-living saprotrophs that appear to be facultative insect pathogens and another clade, exemplified by C. thromboides, which includes primarily insect pathogens (Gryganskyi et  al. 2012; Gryganskyi et  al. 2013). This may suggest that the larger genomes reported for E. aulicae (Murrin et al. 1986) and B. ranarum (Henk and Fisher 2012) could be associated with specialization to obligate insect pathogenicity. This process has been also observed in some plant pathogenic fungi, where the expansion of specific gene families and/or whole-genome duplication has been found to be associated with host specialization (Raffaele and Kamoun 2012). If these inferences are correct it implies that the evolution of the large haploid genome of 350 Mb in B. ranarum is independent from potential genome expansion events within Entomophthorales, and might be driven by adaptation to amphibian and reptile guts. This is supported by recent phylogenetic analyses where the genus Basidiobolus is the sister group to all other Entomophthoromycota and is placed somewhere between Entomophthoromycota

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and chytrid fungi (Henk and Fisher 2012; Gryganskyi et al. 2013). From the reconstructed phylogeny it is evident that entomopathogenicity evolved independently in these families and that genera of hypocrealean entomopathogens cluster among closely related phytopathogens, endophytes and mycoparasites. These ancestral associations are consistent with repeated transitions (host switching) between plant, fungi and insect hosts, as suggested by Suh et al. (2001) in their study on Cordyceps spp. A comparative genome analysis of seven Metarhizium (Clavicipitaceae) genomes confirmed the genus as a monophyletic lineage that diverged from clavicipetacean plant pathogens and endophytes about 231 million years ago and placed the hemipteran-­specific M. album as basal to the Metarhizium clade, with an estimated divergence time about 117 million years ago (Hu et al. 2014). It was suggested that the close physical proximity of the plant-associated ancestor of M. album to plant-­ sap sucking hemipteran bugs may have facilitated this particular host switch to entomopathogenicity (Hu et  al. 2014). As part of a wider study on the biocontrol potential of EF from Central Asia, the phylogenetic characterization of a number of insect-derived isolates from Uzbekistan was assessed (Ergashev et al. 2009). Specialization in Metarhizium is associated with retention of sexuality and rapid evolution of existing protein sequences, whereas generalization is associated with protein family expansion, loss of genome-defence mechanisms, genome restructuring, horizontal gene transfer and loss of sexuality (Hu et al. 2014). Fungi that are able to infect insects are not just comprised by a single monophyletic group. Different groups have arisen independently and repeatedly in many different lineages through fungal evolution (Humber 2008). Recent phylogenetic studies indicate that the ability to utilize insects as a source of nutrition has arisen more than once among fungi (Spatafora et al. 2007). Scale insects, particularly Coccidae and Aleyrodidae have the greatest diversity of fungal pathogens documented (Humber 2008). These insects occur in dense and mainly immobile populations feeding on plants. The sustained proximity between these insects, fungi and other potential hosts may provide pathogenic fungi with the opportunity to move from plant to insect and beyond.

3.6  Conclusion Molecular identification and the phylogenetic analyses of entomopathogens are basic principles in insect pathology. Molecular phylogenetics applies a combination of molecular and statistical techniques to recover the order of evolutionary events and represent them in evolutionary trees that graphically depict relationships among species or genes over time. Phylogenetics based on sequence data provide us with more accurate descriptions of patterns of relatedness than was available before the advent of molecular sequencing. Genome wide studies on gene content, conserved gene order, gene expression, regulatory networks, metabolic pathways, functional genome annotation can all be enriched by evolutionary studies based on

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phylogenetics. Thus, the molecular criteria and bioinformatic tools for pathogen discrimination and species demarcation is obvious and need of the hour. However, molecular phylogenies are sensitive to the assumptions and models used for trees constructing. They face issues such as long-branch attraction, saturation and taxon sampling problems. This means that strikingly different results can be obtained by applying different models to the same dataset. Therefore, it is concluded that developing new experimental approaches to discover the links between these molecular mechanisms and ecological processes can substantially improve inference in evolutionary biology, in a significant and powerful way.

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Chapter 4

Potential of Entomopathogenic Bacteria and Fungi Lav Sharma, Nitin Bohra, Rupesh Kumar Singh, and Guilhermina Marques

Abstract  Soil is a reservoir of numerous microorganisms critical for the sustainable functioning of natural and managed ecosystems. Entomopathogenic bacteria and fungi are natural enemies of pest-insects, whose utility in agroecosystems has been studied since decades. These entomopathogens spend significant time period in soil, either as saprotrophs, active conidia, resting spores or dormant endospores. In this chapter, we focus on: (a) the different bacterial and fungal species exhibiting entomopathogenicity; (b) insect-hosts and pathology; and (c) their survival in soil. Studying these aspects is of the utmost importance in fully exploiting the potential of these microorganisms. The bacterium Bacillus thuringiensis and fungi from the orders Entomophthorales and Hypocreales are discussed in more details, pertaining to the amount of literature and their dominance in the microbial biopesticide industry. Keywords  Hypocreales · Entomophthorales · Beauveria · Metarhizium · Paenibacillus

L. Sharma (*) · G. Marques CITAB – Centre for the Research and Technology of Agro-Environmental and Biological Sciences, University of Trás-os-Montes and Alto Douro, UTAD, Vila Real, Portugal e-mail: [email protected] N. Bohra School of Genetics and Biotechnology, University of Trás-os-Montes and Alto Douro, UTAD, Vila Real, Portugal Department of Biotechnology, National Institute of Technology Warangal, Warangal, Telangana, India R. K. Singh Centro de Química de Vila Real (CQ-VR), Universidade de Trás-os-Montes e Alto Douro, Vila Real, Portugal © Springer Nature Switzerland AG 2019 M. A. Khan, W. Ahmad (eds.), Microbes for Sustainable Insect Pest Management, Sustainability in Plant and Crop Protection, https://doi.org/10.1007/978-3-030-23045-6_4

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4.1  Introduction Bacteria exhibit different levels of mutualistic relationships with insects, that arose during the last 250 million years of evolution. Only a few lineages, however, evolved strategies to invade a host-insects and kill them by overcoming their immune response, and are known as bacterial entomopathogens or entomopathogenic bacteria (EB). The mechanisms of entomopathogenicity developed through a long co-­ evolutionary process (Vilcinskas 2010). Considering the life strategies, EB which have been commercialised are either obligate (those completing their life cycle inside the host insect), or facultative (which can grow outside the host). Some bacteria, such as Bacillus spp., produce endospores which are resistant/tolerant to adverse conditions. These endospores prove remarkably beneficial for the bacterium, as they overcome limited temporal availability of the host insect (Jurat-­ Fuentes and Jackson 2012). Moreover, bacteria such as Chromobacterium subtsugae and Bacillus thuringiensis produce toxins which can be exploited even in absence of the live organisms (Glare et al. 2017). Such organism-free approaches for insect biological control facilitate the production of stable and non-infectious biopesticides. Fungi are commonly found in the terrestrial and aquatic environment, and those dwelling on land can have various ecological roles ranging from being parasites, pathogens, endophytes, or symbionts of plants or animals. In a broader sense, those fungi which cause infections (mycosis) in insects or other arthropods such as mites, ticks and spiders are known as fungal entomopathogens or entomopathogenic fungi (EPF). A common difference between EB and EPF is that the former must be ingested by the insect, whereas the latter generally penetrates their hosts’ cuticle (Hajek and Meyling 2018). Two fungal orders, Hypocreales and Entomophthorales, deserve attention due to their applications in various strategies for the biological control of arthropod pests.

4.2  Bacterial History and Diversity During the late nineteenth century, the silk industry in Japan observed sudden deaths of silkworm, Bombyx mori (Beegle and Yamamoto 1992; Milner 1994; Davidson 2012). In 1898, a spore forming bacterium was observed in these caterpillars by the Japanese scientist Sigetane Ishiwata. This bacterium was lethal enough to kill the silkworm caterpillars, hours after its ingestion.. The disease was termed as “sotto-­ byo-­kin” which translates as “collapse-disease-microorganism” (Ishiwata 1901; De Barjac and Bonnefoi 1968). That led to the discovery of the bacterium Bacillus sotto. In 1915, Aoki and Chigasaki confirmed the presence of a toxin within the bacterium (Angus 1954). In another independent study, a similar bacterium was found in the Mediterranean flour moth larvae by a German scientist Ernst Berliner in Thuringia, Germany, in 1909, that he named Bacillus thuringiensis (Bt) (Berliner

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1915). He also proposed the use of Bt for insect biological control, however, the strains were lost. Luckily, German scientist Mattes isolated Bt from flour moth and successfully tested it against European corn borer (Angus 1954). This study later led to the first commercial product based on B. thuringiensis in 1938 (Beegle and Yamamoto 1992; Milner 1994). Bacterial  Entomopathogens belong to four phyla: Firmicutes, Actinobacteria, Proteobacteria and Tenericutes. However, most of the known EB are from families Neisseriaceae, Enterobacteriaceae, Paenibacillaceae and Bacillaceae. Bacteria belonging to the genus Bacillus are the most widely studied and used as microbial insect pathogens. Commonly known EB are considered in this work, and are presented in Fig.  4.1. Other insect-pathogenic bacteria such as Xenorhabdus and Photorhabdus from the family Enterobacteriaceae are symbionts of entomopathogenic nematodes, and are out of the scope of this chapter.

4.2.1  Gram Positive: Firmicutes and Actinobacteria Bacteria that exhibit entomopathogenicity in phylum Firmicutes belong to the orders Bacillales, Lactobacillales, and Clostridiales. Entomopathogenic bacterial families from these orders are presented in Fig.  4.1. Bacteria from the phylum Actinobacteria can be pathogenic to plants, animals, and humans. Entomopathogens from this group belong to the families Streptomycetaceae and Pseudonocardiaceae, of order Actinomycetales. 4.2.1.1  Bacillaceae This family consists of some of the bacteria which have been dominating the market of bacterial entomopathogens for some decades. These bacteria are endospore-­ forming, as they undergo sporulation and form one oval-shaped spore per cell during adverse conditions. Bacillus thuringiensis Bacillus thuringiensis is commonly found in soil, plants, water, dead insects and stored cereals. It is a gram-positive, facultative anaerobe which produces crystalline proteinaceous inclusions or “parasporal crystals” during its vegetative and sporulating growth phases (Argôlo-Filho and Loguercio 2014). Approximately 98% of the sprayable bacterial biopesticide formulations are from Bt and its subspecies. These EB collectively cause mortality among six taxonomic orders of insects (van Frankenhuyzen 2009). Moreover, Bt also infects root-knot nematodes (Jurat-­ Fuentes and Jackson 2012). Bacillus thuringiensis generally differs from other bacteria as it contains parasporal crystals. The insect specificity of a Bt strain is determined by the toxins it produces. Crystal (Cry) and cytolytic (Cyt) toxins are the two proteins present in the parasporal crystals, in general. Besides, Vegetative

Fig. 4.1  Classification of entomopathogenic bacteria. (Modified from Jurat-Fuentes and Jackson 2012)

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Insecticidal protein (VIP) toxins are synthesized and secreted by vegetative Bt cells. At least 732 Cry toxins have been identified from 73 families of crystal toxins. Moreover, 38 Cyt toxins belonging to three cytolytic protein families, and 125 VIP toxins belonging to four families, have also been reported (Crickmore et al. 2014). Toxins from Bt target the receptors of the midgut epithelial cells and disrupt the natural cell membrane permeability through a pore-forming mechanism. This leads to the cell lysis, followed by gut paralysis and insect death. The bacteria continue to grow until nutrition depletion, and ultimately sporulate (Raymond et  al. 2010). Most of the commercially available Bt formulations are spore-crystal mixtures that are effective against different insects. Bacillus thuringiensis var. kurstaki and B. thuringiensis var. aizawai are used against lepidopteran larvae. Other strains, such as B. thuringiensis var. san diego and B. thuringiensis var. tenebrionis are used against beetle pests, whereas B. thuringiensis var. israelensis is a mosquito pathogen. Strategies for integrating cry genes into the crop plants, such as maize and cotton, to create genetically modified plants, have also been successful (Ruiu 2015). Lysinibacillus sphaericus Previously known as Bacillus sphaericus, Lysinibacillus sphaericus is commonly isolated from soils and aquatic habitats. Based on DNA homology analyses, entomopathogenicity is exhibited only by the IIA sub-group of the species (Krych et al. 1980; Charles et al. 1996). The insecticidal activity varies with the kind of toxin produced, i.e., Bin (Binary) and/or the Mtx (Mosquitocidal toxin) (Silva Filha et al. 2014). Some bacterial strains also produce other binary toxins which are related to the crystal toxins of Bt, such as Cry48 and Cry49. These binary toxins are synthesized and stored as parasporal bodies, however, the Mtx toxins are synthesized during bacterial vegetative stage (Wirth et al. 2014). Among different mosquito species, Lysinibacillus infections are more common in Culex, followed by Anopheles and Mansonia (Berry 2012). L. sphaericus is also toxic to some species of Aedes. However, several Aedes spp., such as Aedes aegypti, are resistant to L. sphaericus (Lacey et al. 2015). The bacterium is also applied to control non-biting midges and black flies (Glare et al. 2017). After ingestion by the host, Bin and Cry toxins target the midgut cells and induce hypertrophy. This is followed by an increase in the presences of lysosomes, induction of apocrine secretion, and damage to the epithelium. Parasitized insects show the onset of paralysis following damages to neural and skeletal cells. Continual growth of vegetative cells and sporulation inside the hemocoel sustain the mosquitocidal activity, probably through a synergistic action mediated by the Mtx and Bin toxins (Berry 2012). Lysinibacillus sphaericus has some specific advantages over B. thuringiensis var. israelensis, as the former is quite specific to only some species of mosquitoes, as mentioned earlier. Furthermore, the bacterium persists longer than Bti in polluted habitats. However, populations of the mosquito Culex quinquefasciatus from Brazil, Tunisia, France, Thailand, India, and China exhibit low to very high resistance against L. sphaericus (Lacey 2007). Approaches such as adding B. thuringiensis var. israelensis genes to the L. sphaericus genome can overcome insect resistance, and increase its infectivity spectrum (Federici et al. 2007).

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4.2.1.2  Paenibacillaceae The bacteria from Paenibacillaceae are characterized by a larger spore which can be oval or ellipsoidal. These bacteria are catalase negative and are difficult to produce in-vitro, unlike other members of Bacillaceae. Paenibacillus spp. Paenibacillus spp. are spore-forming and obligate pathogens attacking larvae of species within the family Scarabaeidae (order Coleoptera). Paenibacillus spp. exhibit a limited growth on nutrient media, a characteristic which distinguishes them from Bacillus spp. These bacteria cause the “milky disease”, named after the milky hemolymph aspect of infected larvae. Paenibacillus spores can persist in soils for many years. However, the infection process is induced only when the spores are ingested (Jackson et al. 2018). Following intake, the spores germinate, and the vegetative cells reach the luminal side of the basal membrane, after penetrating the midgut epithelium, undergoing a primary multiplication cycle (Splittstoesser et al. 1973). The toxins from the parasporal body supposedly disrupt the gut epithelial barrier and facilitate the invasion of the hemocoel. Vegetative multiplication is followed by sporulation, which occur until the refractive spores dominate and confer the characteristic milky appearance (Zhang et al. 1997). An exception is the infection by P. lentimorbus in scarab larvae, whichinduces a clotting of the hemolymph, leading to a brown discoloration of the larvae (Sharpe and Detroy 1979; Glare et al. 2017). Paenibacillus popilliae was the first microbial insecticide registered in North America, for the biological control of the Japanese beetle Popillia japonica (Klein 1988). However, in some reports, epizootics caused on P. japonica showed variable results with limited or no success (Lacey et al. 2015). Its use as a microbial pesticide was not fruitful as the bacterium required in-vivo culturing, which hampered its large-scale production. The very narrow host range within Scarabaeidae is also a limiting factor for commercial exploitation of P. popilliae (Lacey et al. 2015). Brevibacillus laterosporus Brevibacillus laterosporus is commonly found in soil and water (Ruiu et al. 2013). It is aerobic, but can also be a facultative anaerobe. It is reported to infect insects from at least three orders. Among lepidopterans, B. laterosporus is pathogenic to the diamondback moth Plutella xylostella and the velvet bean caterpillar Anticarsia gemmatalis (De Oliveira et al. 2004). Within Coleoptera, B. laterosporus infects the Mexican cotton boll weevil Anthonomus grandis and the corn rootworm Diabrotica virgifera virgifera (De Oliveira et al. 2004; Ruiu et al. 2006). Among Diptera, B. laterosporus has been applied against the larvae of the housefly Musca domestica, of mosquitoes such as C. quinquefasciatus and A. aegypti, and against the black fly Simulium vittatum (Favret and Yousten 1985; Ruiu et al. 2007). This bacterium also infects other invertebrates, such as nematodes and freshwater snails (Ruiu et  al. 2013). Insecticidal toxins of infectious B. laterosporus have been a topic of debate. The bacterium produces a typical “canoe-shaped” parasporal body (CSPB), and these crystals were supposed to be required for infections against mosquitoes, e.g. A.

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aegypti (Ruiu et al. 2013). However, strains lacking CSPB were also found to be toxic (Ruiu et al. 2007). Insecticidal secreted proteins (ISPs), with a resemblance to the Bt VIP proteins, have also been reported from B. laterosporus. Besides, recent studies have reported the presence of the genes for Cry proteins in B. laterosporus (Sharma et al. 2012). The progressive symptomatology and the changes that occur within the midgut are similar to those caused by Bt (Ruiu et al. 2012). 4.2.1.3  Clostridiaceae These are gram-positive, anaerobic and spore-forming rod-shaped bacteria. Entomopathogenicity amongst the bacteria from the Clostridiaceae is rarely noticed. Clostridium bifermentans Clostridium bifermentans is occasionally isolated as a human pathogen. For insects, C. bifermentans var. malaysia is a highly toxic entomopathogen of black flies and mosquitoes. However, C. bifermentans var. malaysia does not exhibit short-term toxicity against humans, mammals and goldfish (Thiéry et al. 1992; Qureshi et al. 2014). A range of toxins are required for the bacterial infections in mosquitoes. For example, toxins encoded by Cry operon, Cry16A, Cry17A, Cbm17.1, and Cbm17.2, are only insecticidal to Aedes spp., however, a different set of proteins are supposed to cause infection in Anopheles spp. (Qureshi et al. 2014). 4.2.1.4  Streptomycetaceae Representatives of this family produce extensive mycelium which does not fragment, in general. After maturity, the mycelium forms a typical chain of spores (Kämpfer et al. 2014). Streptomyces spp. Some Streptomyces spp. produce a variety of metabolites that are toxic to insect pests, such as Aphis gossypii, Chilo partellus, Helicoverpa armigera and S. litura. These metabolites include antimycin A, avermectin, flavensomycin, macrotetralides, piericidins and prasinons (Ruiu et al. 2013). Metabolites like avermectin inhibit the neurotransmission in insects by binding to their γ-aminobutyric acid (GABA) receptor, which eventually leads to the paralysis of the insect neuromuscular systems. Species exhibiting entomopathogenicity include S. albus and S. avermitilis (Glare et al. 2017). 4.2.1.5  Pseudonocardiaceae Like Streptomycetaceae, the family Pseudonocardiaceae also belongs to the order Actinomycetales.

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Saccharopolyspora spinosa Saccharopolyspora spinosa produces insecticidal metabolites such as “spinosyns” during aerobic fermentation. Spinosyns are active towards a wide range of insects. However, they also exhibit toxicity towards non-target insects, aquatic organisms, and mammals (Ruiu et al. 2015).

4.2.2  Gram-Negative: Proteobacteria The Proteobacteria form a phylum which consists of many pathogenic bacteria. Entomopathogenic species belong to the families Enterobacteriaceae, Pseudomonadaceae, Coxiellaceae, Neisseriaceae, and Burkholderiaceae. 4.2.2.1  Enterobacteriaceae Members of the Enterobacteriaceae are rod-shaped facultative anaerobes. Most of them are pathogenic in nature and do not form spores. Serratia spp. Serratia spp. are ubiquitous in the environment, and some are reported from dead or diseased insects. Serratia marcescens supposedly causes disease in the May beetles Melolontha melolontha, the tsetse flies Glossina spp. (Poinar et al. 1979), and the blowfly Lucilia sericata (O’Callaghan et al. 1996; Jurat-Fuentes and Jackson 2012). Moreover, S. marcescens was reported as toxic against the diamondback moth (Jeong et al. 2010). Association of Serratia spp. like S. nematodiphila with entomopathogenic nematodes, such as Heterorhabditinoides sp., suggests potential for its use in the biological control of insects pests (Zhang et al. 2009). Serratia entomophila, the causal agent of the “Amber disease” in the New Zealand grass grub Costelytra zealandica, is commonly found in soils (Jackson et al. 1993). Recently, some S. entomophila strains were found to be pathogenic to several Phyllophaga and Anomala spp. (Nuñez-Valdez et al. 2008). In general, C. zealandica larvae stop feeding in a few days after bacterial ingestion. The bacterium facilitates gut clearance, which is followed by the appearance of characteristic amber colouration. The diseased larva stays active for 1–3 months in the soil without feeding, and shrinks while consuming its own body fat. This leads to the weakening of the structural tissues, eventually allowing the bacteria to enter the hemocoel and cause septicaemia leading to the host death (Jackson et al. 1993, 2001). The entomopathogenicity of S. entomophila and S. proteamaculans is attributed to two different gene clusters: sep (Serratia entomophila pathogenicity) and afp (anti-feeding prophage). The sep cluster is composed of three genes, sepA, sepB, and sepC, and is responsible for the clearance of the gut, and the development of amber colouration of larvae (Hurst et al. 2007a). The Sep proteins are similar to the Tc (toxin complex) proteins found in Photorhabdus luminescens. Another gene cluster, afp, codes for the novel “anti-feeding prophage” which is a toxin delivery apparatus that makes larvae cease feeding (Hurst et al. 2007b).

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Yersinia spp. The bacterial family Enterobacteriaceae includes some well-known human pathogens. Entomopathogenicity among Yersinia spp. have been described only recently. It was found that the Tc proteins were necessary for the insect pathogenicity of Yersinia enterocolitica (Bresolin et  al. 2006). Later, a few other species, such as Yersinia mollaretii, Y. pestis, and Y. pseudotuberculosis are presumed entomopathogenic due to the presence of Tc genes in their genomes (Fuchs et al. 2008). Recently, Y. entomophaga was isolated from a diseased C. zealandica larva in New Zealand, and was found highly pathogenic to the insects from the orders Coleoptera, Lepidoptera, and Orthoptera (Hurst et al. 2011a). After bacterial ingestion, the larva stops feeding, this is followed by the regurgitation and clearance of the gut content, leading to a similar appearance to the amber disease, however for a short duration. The gut epithelial membrane degrades, allowing the invasion of hemocoel, which leads to septicemia and death in 2–5 days after bacterial ingestion (Marshall et al. 2012). The pathogenicity of the bacteria is governed by a “Tc protein complex” which is composed of TcA, TcB and TcC toxins, and two chitinase proteins, collectively constituting the Tc molecule (Hurst et al. 2011b). 4.2.2.2  Pseudomonadaceae Bacteria belonging to the Pseudomonadaceae are widely distributed in the environment and include some well-known species, such as Pseudomonas aeruginosa, which are pathogenic to plants, and animals including humans. Some pseudomonads can also be isolated from diseased and dead insects. Pseudomonas spp. A soil isolate of P. entomophila showed high insect pathogenicity against the common vinegar fly Drosophila melanogaster. With time, additional reports included the pathogenicity of P. entomophila against the insect orders Diptera, Lepidoptera and Coleoptera, nematodes, and amoebae (Dieppois et al. 2015). Genome analysis identified a two-component system, GacS/GacA, that regulates potential virulence factors, such as toxins, putative hemolysins, hydrogen cyanide, and proteases, and supposedly assists the entomopathogenicity in P. entomophila. Its genome also consists of genes encoding TcB and TcC toxins, as in the case of Yersinia spp. However, P. entomophila lacks the genes for TcA component (ffrench-Constant and Waterfield 2005). Another species, P. putida, was reported pathogenic to the Colorado potato beetle Leptinotarsa decemlineata (Muratoglu et al. 2011). 4.2.2.3  Coxiellaceae These are gram-negative, obligate intercellular pathogens, and those exhibiting entomopathogenicity have been reclassified from the order Rickettsiales (α-proteobacteria) to the Legionellales (γ-proteobacteria) (Leclerque 2008).

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Rickettsiella spp. Rickettsiella popilliae, R. chironomi and R. grylli are the three most widely known entomopathogens. R. popilliae typically targets the hemolymph cells and the fat body of the host-insect. The diseased larva is characterised by the development of white to blue colouration although, in the case of infection in Japanese beetle, the colonisation causes a greenish-blue color (Jurat-Fuentes and Jackson 2012). In general, the infected larvae act normal for over a month but with passing time they loose vigour, become sluggish, and stop feeding before death. In the cases of infections in crickets by R. grylli, a “behavioural fever” is observed. The diseased insect tries to reach an environment with a higher temperature, to reduce bacterial growth. Some concerns, such as the ability of the entomopathogenic Rickettsiella spp. to cause inflammation and infections in vertebrates, and a very slow insect death post bacterial ingestion, have raised doubts on their use as microbial agents for insect pest control (Jurat-Fuentes and Jackson 2012). 4.2.2.4  Neisseriaceae Most of the bacteria from the Neisseriaceae are animal commensals, although, a few are human pathogens. Generally, they do not exhibit entomopathogenicity but there are some exceptions as discussed below. Chromobacterium subtsugae A soil isolate of C. subtsugae from Maryland demonstrated high insecticidal activity against insects from different orders, including Lepidoptera, Hemiptera, and Coleoptera. The strain produces a chemical “violacein” which confers a violet colour to its colonies. Pathogenicity of the bacterium is attributed to the various molecules produced during the infection process, including heat stable insecticidal toxins (Martin et al. 2007). Recently, another Chromobacterium strain, Csp_P, was isolated from the midgut of the mosquito A. aegypti, which reduced the larval and adult survival rates. Furthermore, an anti-pathogen activity of the Csp_P strain against the malaria-causing Plasmodium falciparum and the dengue virus, was observed in-vitro. The strain compromised the vector competence of the mosquito (Ramirez et al. 2014). 4.2.2.5  Burkholderiaceae Species from the Burkholderiaceae have wider biological roles including human, animal and plant pathogens. Burkholderia spp. Burkholderia spp. have been isolated from insect gut, and can negatively affect the oviposition time and the number of eggs in the bean bug Riptortus pedestris (Kil et al. 2014). Recent studies have reported the entomopathogenicity of B. rinojensis

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against the two-spotted spider mite Tetranychus urticae and the beet armyworm Spodoptera exigua (Cordova-Kreylos et al. 2013).

4.3  Soil Habitat and Pathogenesis of Bacteria Bacillus thuringiensis is ubiquitous, but the numbers of cases of natural insect infections are rare. Such contradiction has led to an on-going debate: is Bt an opportunistic pathogen with soil being the primary environment for bacterial reproduction? It is hypothesized that Bt survives on the decaying organic matter or the root exudates. The bacterium reaches to the aerial parts of the plants as they grow (Argôlo-Filho and Loguercio 2014). The ability to grow on a wide range of substrates and a high persistence in soil samples are some of the arguments supporting this hypothesis (Glare and O’Callaghan 2000, Raymond et  al. 2010). However, other works disagree with Bt growth, and survival of parasporal protein crystals, in natural soil conditions (West et al. 1985). Some researchers convincingly argue that Bt is a “bona fide insect pathogen” whose primary reproduction occurs within a host-insect, and only after multiplication, the bacterium is deposited on plants and soils, eventually making them its natural reservoirs (Raymond et al. 2010; Argôlo-­ Filho and Loguercio 2014). The scarcity of Bt natural insect infections is attributed to a rare combination of susceptible insects, optimal temperatures, and nutrient availability. Although, a successful horizontal transfer of Bt spores into the host-­ insect after ingesting them from the soil, is quite difficult, as the spores will have to germinate and undergo at least one cycle of vegetative growth to synthesize Cry toxins. Therefore, bacterial transmission through plants seems to be more feasible (Raymond et al. 2010). The strategy of host-seeking by a resistant bacterial spore is also adopted by the milky disease-causing P. popilliae, as the bacterium produces ultra-dormant spores which can survive many years. The bacterium gets activated after the ingestion of the soil organic matter and the plant root by the scarab larvae (Jackson et al. 2018). Nonetheless, the soil is an excellent reservoir for Serratia spp., and many methods have been developed to study the ecology of facultative insect pathogens, S. entomophila and S. proteamaculans, that cause the amber disease in C. zealandica, as discussed earlier. Studies within the pasture soils from New Zealand revealed that both pathogenic as well as non-pathogenic strains of Serratia spp. can co-exist. Pathogenic strains, however, are generally localised to the areas with grass grub populations (Jackson et al. 2018). Some bacterial species, such as B. laterosporus, Bacillus spp. and P. putida may also act as plant endophytes enhancing plant growth, in the absence of insects. Different EB undertake distinct mechanisms to infect their host-insects, as already briefed earlier. A brief pictorial description is presented in Fig. 4.2. There are two distinct phases in the life cycle of Bt, i.e., vegetative cell development and spore development. Bacterial vegetative cells measure 2–5 μm long and 1 μm wide, with short hair-like flagella (Bulla et  al. 1980). These cells are divided into two

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Fig. 4.2  Mechanism of infection by Bacillus thuringiensis. (Modified from Jean-Michel Vassal, CIRAD, France)

uniform daughter cells due to the formation of division septa initiated midway alongside the plasma membrane. Spore development is a typical seven stages process that spans from (1) the formation of the axial filament; (2) forespore septum formation; (3) cell engulfment (4–6) the formation of the exosporium, primordial cell wall, cortex and spore coats, and subsequent transformation of the spore nucleoid; and, lastly, (7) spore maturation and sporangial lysis (Ibrahim et al. 2010). Bacillus thuringiensis crystals get solubilised in the alkaline environment of the host gut upon ingestion. These proteins get processed by mid-gut proteases (for e.g., tripsin like, or chemotripsin like in case of Lepidoptera) and change from protoxins to an active toxin core. The processed toxin binds to the cadherin receptors present on the host epithelial mid-gut cell membrane. Here onwards, two models have been reported to define the possible mode of action of a Cry toxin. One mode of action is the “pore formation model” where cadherin initiates the cleavage of helix ɑ1 resulting in the oligomerisation of the Cry toxin, which in turn binds to the GPI-anchored receptors, assisting the toxin to penetrate into the membrane, leading to the formation of pores and cause colloid osmotic lysis (Bravo et al. 2004). Another mode of action is the “signal transduction model” where interaction of the Cry toxin with cadherin activates a guanine nucleotide binding protein (G-protein), which initiates an adenylyl cyclase activity resulting in the production of the intracellular cyclic adenosine monophosphate (cAMP). This causes the activation of the protein kinase A, and the induction of an intracellular pathway triggering cell death (Zhang et al. 2006). Where and when one or the other mechanisms lead to the larval death is still controversial (Soberón et al. 2009). Crickmore et  al. (1998) defined Cry protein as “A parasporal crystal protein from B. thuringiensis that shows some experimentally verifiable toxicity against a

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target pest”; whereas, Cyt represents “a parasporal crystal protein from B. thuringiensis that exhibits haemolytic activity or any protein which has sequence homology with a known Cyt protein”. The difference between the Cry and the Cyt proteins mode of action is that the latter bind to the non-glycosylated lipid portion of the microvillar bilayer, and therefore do not need binding to a protein receptor (Federici and Siegel 2007). Since the first cloning of Cry protein was reported by Schnepf and Whiteley (1981), approximately 200 Bt-proteins have been characterised for their action against many insects from orders such as Lepidoptera, Coleoptera, Diptera, Homoptera, Orthoptera, Hymenoptera, and suborder Mallophaga (Schnepf et  al. 1998; Baxter et  al. 2011). Besides, Bt-proteins have also been reported for their toxicity against protozoa, mites, and nematodes. In general, the insecticidal property is attributed to the crystal proteins. However, recent studies have shown the utility of the VIP toxins against a wide range of lepidopteran larvae (Estruch et al. 1996). To summarise, some of the criteria which define the insecticidal range and the efficacy of a particular Cry protein are: (a) a sufficient ingestion by the pest-­ insect and an effective solubilisation in the gut media; (b) the efficiency of the protoxin-­toxin conversion, and the specificity of binding to a membrane receptor (such as Cadherin); and, (c) the membrane pore formation, i.e. membrane insertion and ion-channel activity (Schnepf et al. 1998).

4.4  Fungal History and Diversity The earliest evidence of a fungal entomopathogen to date is Paleoophiocordyceps coccophagus, an Ophiocordyceps-like anamorph, attacking an early Cretaceous Burmese male scale insect dating 100–110 million years ago (Sung et  al. 2008). Nonetheless, during human civilization, the earlier reports on EPF are also credited, like for bacteria, to the silk industry. In China, Cordyceps fungi had been observed on caterpillars and were used in traditional Chinese medicine. However, the first published report on Cordyceps was provided by a French scientist, René Antoine Ferchault de Réaumur, as “vegetable growths” (Réaumur 1734–1742, Davidson 2012). In the early nineteenth century, a “white muscardine” disease of silkworms became a big problem for the silk industry in France. The disease was called “calcinacio” because of the white powdery deposit on the body of caterpillars. Agostino Bassi, who was later renowned as the “Father of Insect Pathology”, investigated the causal agent, and reported a fungus or a “vegetable parasite” (Bassi 1835–1836). Giuseppe Gabriel Balsamo-Crivelli then named the fungus Botrytis bassiana in honour of Bassi. The fungus is now renowned as Beauveria bassiana. These discoveries led to the idea of using EPF against other insects, and a few reports emerged in this direction. In 1878, a similar outbreak of “green muscardine” disease was analysed near Odessa (Ukraine) by Élie Metchnikoff, on the wheat cockchafers Anisoplia austriaca. The pathogen was identified as Entomopthora anisopliae (Metchnikoff 1879). Sorokin (1883) named it as “Metarrhizium” anisopliae, presently spelt as Metarhizium. Years later, in 1888, the same fungus was used against

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the sugar-beet weevil Bothynoderes (Cleonus) punctiventris by Isaak Krassiltstchik (Krassiltstchik 1888). That was the first large-scale production for a field trial of any microbial biopesticide. Whilst the genus Beauveria started gaining importance, Steinhaus (1949) briefed the process of infection, disease development, and practical usage of B. bassiana against some pest-insects, such as the chinch bug Blissus leucopterus, the codling moth Carpocapsa (Cydia) pomonella, and the European corn borer Pyrausta (Ostrinia) nubilalis (Zimmermann 2007). Similar attention was seen in the usage of M. anisopliae against numerous pest-insects, such as the sugarcane froghopper Aeneolamia flavilatera, the wireworms Agriotes obscurus and Agriotes sputator, the turnip moth Agrotis segetum, the sugarcane white grub Alissonotum impressicolle, the sugar-beet weevil, moths from Euxoa spp., the European corn borer, the coconut rhinoceros beetle Oryctes rhinoceros, P. japonica and the black rice bug Scotinophara lurida (Müller-Kögler 1965). However, the continual development of chemical insecticides hampered the studies on the ecology of the natural biocontrol agents. Besides, the efficacies of microbial insect-pathogens were compared with chemical pesticides, without taking the ecological requirements of a microbial biopesticide in consideration. This biased comparison resulted in a disinclination towards the fungal microbial pesticide agents (Vega et al. 2009). Finally in 1981, after a long delay of a few decades, the fungus Hirsutella thompsonii was registered as the first fungal insecticide for its use against the citrus rust mite Phyllocoptruta oleivora, in the USA (Tanada and Kaya 1993). Nonetheless, in terms of biology, entomopathogenicity in fungi is restricted to a few orders, as showed in the Fig. 4.3. However, occasional insecticidal activities can be noticed in most fungal phyla.

4.4.1  Oomycota (Kingdom: Chromista) The oomycetes are eukaryotic microorganisms with filamentous features, and were thought to be fungi while they are not. They are aquatic microorganisms with a coenocytic hypha which is mainly aseptate, whose cell walls are mainly composed of glucan-cellulose. Recent classifications have placed them closer to the diatoms and the brown algae, in the kingdom Chromista. Asexual reproduction takes place by “zoospores” exhibiting two flagella of varying lengths. Sexual spores are termed “oospores” (Wraight et  al. 2007). They are able to infect algae, protists, plants, fungi, vertebrate animals, and arthropods. Although plant pathogens such as Phytophthora infestans, the potato late blight causal agent, are well known, the information on entomopathogenic oomycetes is indeed limited. Most of insect pathogenic oomycetes belong to three different orders: Myzocytiopsidales, Pythiales, and Saprolegniales. Previous reports showed that a few species from these groups are facultative pathogens of mosquitoes. In terms of insect pathology, the species Aphanomyces laevis and Lagenidium giganteum are among the best studied oomycetes. However, species from genera Leptolegnia, Pythium, Couchia

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Fig. 4.3  Classification of entomopathogenic fungi. (Modified from Kirk et al. (2008), Vega et al. (2009), Gryganskyi et al. (2012) and Boomsma et al. (2014))

and Crypticola have also been spotted occasionally (Scholte et al. 2004). Oomycete infections on mosquito larvae have been observed on the waters from rivers, ponds, well-aerated streams, lakes, tree holes, and on the leaf axils (Araújo and Hughes 2016).

4.4.2  Microsporidia The Microsporidia are among the first diverging lineages of fungi, but in earlier times they were considered as protozoans (James et al. 2006; Hibbett et al. 2007). They are ubiquitous, obligate intracellular parasites of animals including invertebrates (Wittner and Weiss 1999). They inject spores into the cytoplasm of the host rapidly, through a thin polar tube (James et al. 2006). Sixty-nine genera have been reported for entomopathogenicity, and 42 of them infect dipteran insects only. Amblyospora is the most abundant and the largest genus that alone kills 79 dipteran species belonging to eight genera (Araújo and Hughes 2016). Amblyospora exhibits a complex life cycle and requires an intermediate copepod host, and two generations of insect (mosquito) host for completion of its full lifespan. Nosema is also an important and widely distributed microsporidian genus. Nosema apis and N. ceranae target honey bees and are hence considered quite devastating for apiculture. The infection occurs as N. apis invades the epithelial layer of the ventriculus and the

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midgut of Apis mellifera adult bees. This causes digestive disorders leading to a shortened bee lifespan, resulting in a decrease in the bee population after winter conditions (Higes et al. 2006). Studies in the past have also reported microsporidian pathogenicity against caterpillars, such as the European corn borer, locusts, mosquitoes and grasshoppers. However, their utility as biological control agents is marred by the difficulties in their mass productions (Corradi 2015).

4.4.3  Chytridiomycota The Chytridiomycota are supposed to be the most primitive lineage of fungi, dating as back as 400 million years (Taylor et al. 1992; James et al. 2006). They produce motile zoospores and gametes with smooth whiplash flagellum. The majority of them are wet soils and freshwater saprotrophs (Wraight et al. 2007). Many chytrid species are parasitic on protists, tardigrades, fungi, rotifers, plants and animals. Fewer species, belonging to Myiophagus, have been reported for entomopathogenicity, but such cases are rare (Araújo and Hughes 2016).

4.4.4  Blastocladiomycota Fungi belonging to the phylum Blastocladiomycota can be saprotrophs, plant and invertebrate pathogens (Longcore and Simmons 2012). They are distinct from the other fungi as they include an alternation between haploid and diploid generations, and for the fact that meiosis occurs during the formation of spores within a thick-­ walled “meiosporangia”. Entomopathogenic blastocladiomycetous fungi belong to the genera Catenaria, Coelomycidium and Coelomomyces of the order Blastocladiales. Catenaria spp. are pathogenic to nematodes, although, some of them may also infect flies. Coelomycidium spp. are toxic to beetles, scales and some Diptera (Tanada and Kaya 1993). For example, Coelomycidium simulii is a pathogen that occasionally attacks larvae of the black fly Simulium japonicum larvae (Kim 2011). The genus Coelomomyces includes over 70 species that are pathogenic to different families of Diptera, such as the Chironomidae, Culicidae, and Psychodidae (Scholte et  al. 2004). Coelomomyces are obligate pathogens, and species like C. psophorae require an intermediate copepod host and a mosquito larva at different stages of their life cycles (Vega et al. 2012). Some species, such as C. stegomyiae, do not kill the larva but, on the contrary, reside inside the mosquito, passing through the larval and pupal stages, and maturing within the ovaries of the adult female. After the first blood meal, a fungal hypha matures in the zoospore producing “sporangia”. Therefore, as the mosquito reaches any breeding site, instead of eggs, it lays sporangia full of zoospores, discharging the fungus for a new host (Lucarotti and Shoulkamy 2000). Despite such infection-causing strategies, problems in the

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mass production have rendered these species ineffective for their use on a larger scale (Chandler 2017).

4.4.5  Zygomycota The phylum Zygomycota is constituted by two main classes, the Zygomycetes and the Trichomycetes. Their members are characterised by coenocytic mycelia, asexual reproduction by “sporangiospores”, and absence of flagellate cells and centrioles (Alexopoulos et al. 1996; Araújo and Hughes 2016). The Zygomycota differ from other fungal groups as they produce “zygospores” (thick-walled resting spores) within the “zygosporangium”, a structure formed when two specialized hyphae (gametangia) fuse (White et al. 2006). The Zygomycota consist of a wide group of species which are quite ecologically diverse. These species dwell as saprotrophs in soil and dung, and may colonize bread, vegetables, and fruit (Alexopoulos et  al. 1996). Based on their phylogenetic characterisation, the Zygomycota were classified in five monophyletic taxa, i.e., four subphyla, the Zoopagomycotina, the Mucoromycotina, the Kickxellomycotina, and the Entomophthoromycotina, and one phylum, the Glomeromycota, belonging to the arbuscular mycorrhizal fungi (Hibbett et  al. 2007). Later, Humber (2012) described the phylum Entomophthoromycota, for the Entomophthoromycotina. Subphyla Zoopagomycotina, Mucoromycotina and Kickxellomycotina are mainly constituted of saprotrophs. Species belonging to the Zoopagomycotina are also mycoparasites and nematophagous (Zhang and Hyde 2014). The Mucoromycotina form the most morphologically diverse and largest subphylum within zygomycetous fungi. Some orders of Mucoromycotina, such as Mortierellales and Mucorales, are weak parasites of animals and plants. Just one species i.e., Sporodiniella umbellate can be classified as entomopathogenic, as it causes mortalities amongst the lepidopteran genus Acraea in Taiwan, and the hemipteran genus Umbonia in Ecuador (Araújo and Hughes 2016). The members of the Kickxellomycotina, from the orders Asellariales and Harpellales, are considered as gut commensals of arthropods, however, some species, such as Smittium morbosum, are pathogenic to mosquitoes (Vega et al. 2012).

4.4.6  Entomophthoromycota The Entomophthoromycota are the most important group of entomopathogens amongst the basal lineages of fungi. They produce coenocytic hyphae with some septation, especially in the older mycelial parts. Asexual “azygospores” or sexual zygospores formed by the Entomophthorales are termed “resting spores”, and are capable of surviving harsh environmental conditions. The Entomophthoromycota also produce uninucleate or multinucleate asexual conidiawhich, upon germination

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may produce secondary or tertiary conidia (Wraight et al. 2007). This phylum consists of approximately 280 species, and the majority of them are obligate insect or mite pathogens, with a relatively narrower host range (De Fine Licht et al. 2016). Other species are pathogens of fern gametophytes, desmid algae, vertebrates, reptiles and macromycetes (Gryganskyi et  al. 2012). Some members of families Basidiobolaceae and Anyclistaceae dwell as saprotrophs (Humber 2008). Most entomopathogens within the zygosporic fungi belong to the families Entomophthoraceae and Neozygitaceae. Entomophthora, Entomophaga, Pandora and Zoophthora are some of the widely known insect pathogenic genera of Entomophthoraceae. The Neozygitaceae consists of a few genera, such as Neozygites, which are specialist entomopathogens. Other families, such as Ancylistaceae and the Basidiobolaceae, also represent fewer insect pathogens, such as Conidiobolus spp. and Basidiobolus spp., respectively (Vega et al. 2012). Entomophthoroid fungi generally attack adult insects, although, three species, Entomophthora conglomerata, Entomophthora aquatica and Erynia aquatica infect mosquito larvae (Scholte et  al. 2004). The Entomophthoromycota target a large number of insects, however, in small patches within agroecosystems or forests. Fungal transmission occurs via forcible discharge of spores into the environments, with an exception of the genus Massospora (Lovett and St. Leger 2016). Moreover, species from Massospora, Strongwellsea, and a few from the genera Entomophaga, Entomophthora and Erynia, produce spores while the host is still alive. For example, in case of the infections in dipteran insects Hylemya brassicae and Hylemya platura by Strongwellsea castrans, the flies are seen with a large circular hole on the lateral side of the abdomen, filled with “conidiophores” (spore-producing cells) and fungal tissues (Araújo and Hughes 2016). However, the insect can still be seen as behaving normally. Therefore, these fungi are considered as biotrophs which consume the host while it’s still alive and exhibit no somatic growth after the host’s death (Roy et al. 2006).

4.4.7  Basidiomycota The basidiomycetous fungi are quite diverse and exhibit various ecological traits. They can be saprophytes, and plant pathogens such as the smut and the rust fungi. Animal pathogens include those attacking nematodes. A few genera are also recorded as insect parasites. For example, those from order Septobasidiales, i.e. Auriculoscypha, Coccidiodictyon, Ordonia, Septobasidium and Uredinella, that infect scale insects. Fibularhizoctonia, a member of order Atheliales, attack termite eggs (Henk and Vilgalys 2007). The Septobasidiales are the specialist parasites of scale insects, and use them for their nutrition. However, these fungi seldom kill their host-insects (Humber 2008).

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4.4.8  Ascomycota Ascomycota is the largest fungal phylum with at least 64,163 species (Kirk et al. 2008). The majority of the ascomycetous fungi are filamentous and produce septate hyphae. Sexual spores or “ascospores” are packaged in a sac-like structure called “ascus”. These fungi exhibit very diverse ecology ranging from decomposers to pathogens of animals, humans, and plants. Lichens are also a part of this phylum (Money 2016). The ascomycetous fungi are classified into three subphyla, the Taphrinomycotina, the Saccharomycotina, and the Pezizomycotina. The Pezizomycotina are the most numerous and the most complex subphylum in terms of ecology and morphology, and entomopathogenicity is demonstrated by the members of this group only (Schoch et al. 2009). These fungi can be “anamorphs” (exhibiting an asexual state); “teleomorphs” (exhibiting sexual stage); or “synanamorph” (presences of more than one morphologically distinct asexual stage) (Vega et al. 2012). Some species belonging to the genera Aspergillus and Penicillium are pathogenic to insects, besides other members of the order Eurotiales (class Eurotiomycetes) (Sharma et al. 2018a). Besides, genus Ascosphaera, in the order Ascosphaerales, are the obligate parasites of the bee larvae, and species like Ascosphaera apis cause the honeybee “chalkbrood disease” (Maxfield-Taylor et al. 2015). The spores of A. apis must be ingested for infection. Amongst the members of the Dothideomycetes incertae sedis, species like Podonectria (order Pleosporales) are pathogenic to scale insects and cover the body of the insect with a crust resembling cotton (Roberts and Humber 1981, Kodsueb et al. 2006). Related anamorphs are from the genera Tetranacrium and Tetracrium. Around 2000 species of the Laboulbeniomycetes (order Laboulbeniales), are the obligate haustorial ectoparasites of insects, especially beetles and flies (Weir and Blackwell 2005). Species from the order Myriangiales, within the class Dothideomycetes, can also be found on scale insects (Alexopoulos et  al. 1996). Entomopathogenic Myriangiales exhibit perennial growth and the reproduction is only sexual (Vega et al. 2012). The hypocrealean fungi from the class Sordariomycetes are probably the most well-known and the most studied fungal entomopathogens. Most of the entomopathogens were earlier classified within the family Clavicipitaceae, however, some studies classified the entomopathogens in other families, especially, the Bionectriaceae, the Hypocreaceae, and the Nectriaceae (Vega et  al. 2009; Sosa-­ Gómez et al. 2010; Oliveira et al. 2012; Gouli et al. 2013; Sharma et al. 2018a, b, c; Sharma and Marques 2018). The classical studies on the hypocrealean phylogeny by Sung et al. (2007a) and Sung et al. (2007b) suggested three families within the Clavicipitaceae, i.e., the Cordycipitaceae, the Clavicipitaceae sensu stricto (s.s.), and the Ophiocordycipitaceae. The most specious genus amongst the entomopathogenic hypocrealean fungi is Cordyceps sensu lato (s.l.). It comprises at least 400 species (Chandler 2017). Spatafora et al. (2007) provided evidence of host switching within the hypocrealean families.

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The Clavicipitaceae can be characterised by either lilac, pale, or strongly pigmented green, yellow or occasionally red stromata (Sung et al. 2007a). Cordyceps from the family Cordycipitaceae are characterised by brightly coloured and fleshy stromata, and their anamorphs include species, such as Beauveria, Isaria and Lecanicillium. Members of the Ophiocordycipitaceae exhibit darker stromata and mature ascospores. Some of the widely known entomopathogens from this family include Purpureocillium lilacinum and Ophiocordyceps unilateralis s.l. (Sung et al. 2007a). Recently, Purpureocillium lavendulum was also reported as an EPF from cultivated soils (Sharma et al. 2018b). Whilst the fungi from the Cordycipitaceae are mostly entomopathogens, those belonging to the Clavicipitaceae and the Ophiocordycipitaceae obtain nutrition from animals, plants and fungi (Chandler 2017). Although not specified, it seems that also spider pathogens belong to the Cordycipitaceae; pathogens of dipteran insects, ants and termites are from Ophiocordycipitaceae; and scale insect pathogens are from the Clavicipitaceae (Vega et al. 2012). Ant infections caused by the members of Ophiocordycipitaceae, such as O. unilateralis, present a spectacular case of a host behaviour manipulation by the parasite. The ants leave their nests and climb up the plant, and die sticking themselves onto the leaves. This facilitates fungal dispersal to a longer distance (Araújo et al. 2018), and can be correlated with “walking zombies”, as the fungus manipulates the behaviour of the ants to increase its own fitness. The most important EPF in terms of the usage in agriculture are arguably from the genera Beauveria and Metarhizium. It is believed that B. bassiana alone can kill at least 750 different pest-insects species (Ghikas et al. 2010). Besides, there has been an increase in the literature suggesting the use of EPF, such as B. bassiana, as endophytes and mycoparasites (Vega et  al. 2009, Quesada-Moraga et  al. 2014). Recent multi-gene phylogenies have resolved Beauveria and Metarhizium into many new species (Rehner and Buckley 2005; Bischoff et al. 2009; Rehner et al. 2011). Moreover, habitat-specific preferences have been noticed for these genera within different agroecosystems (Vänninen et  al. 2000; Meyling and Eilenberg 2006, 2007; Quesada-Moraga et  al. 2007; Sun et  al. 2008; Meyling et  al. 2009; Goble et al. 2010; Medo and Cagáň 2011; Sánchez-Peña et al. 2011; Schneider et al. 2012; Clifton et al. 2015; Sharma et al. 2018b).

4.5  S  oil Habitat and Pathogenesis of Entomophthorales and Hypocreales The soil is a reservoir of many microorganisms, and EPF such as Beauveria, Metarhizium and Isaria are considered as weak saprophytes in the competitive soil environment. These fungi spend a considerable part of their life cycle within the soils, and shown in related literature (Hughes et al. 2004; Sevim et al. 2009; Jaronski 2010; Fisher et al. 2011; Scheepmaker and Butt 2010; Meyling et al. 2011, 2012; Muñiz-Reyes et al. 2014; Pérez-González et al. 2014; Kepler et al. 2015; Keyser

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et al. 2015; Aguilera Sammaritano et al. 2016; Hernández-Domínguez and Guzmán-­ Franco 2017; Kirubakaran et al. 2018; Sharma et al. 2018b). Optimum temperature requirement for Entomophthorales is 15–25 °C, whereas for Hypocreales it is 20–30 °C. Soil protects EPF from solar radiation and acts as a buffer against variations in temperatures as well as water availability. It also provides a habitat to numerous soil-dwelling insects, which serve as potential hosts for EPF (Quesada-Moraga et al. 2007). Metabolites secreted by other soil microorganisms may hinder the pathogenicity of EPF, as they may affect the fungal germination and growth adversely and/or can be toxic (Sinha et al. 2016). Therefore, it is not surprising that the EPF efficacy and survival is superior in sterilised soils, when compared with non-sterilised conditions (Jaronski 2007). However, soil amendments with nutrients help in overcoming this apparent fungistasis. Nonetheless, EPF continue to germinate and infect the potential host-insects indicating that, in addition to a reduction in fungistasis, EPF also rely on host or nutrient derived cues for development (Lacey et al. 2015). Soil habitat-types can have some significant effects on the occurrences of EPF. Although there are exceptions, it has been noticed that M. anisopliae is associated with tilled agricultural soils, and B. bassiana is more frequent in uncultivated soils, such as hedgerows (Sharma et al. 2018b). This can be explained as B. bassiana is thought to rely more on host-insect, whereas M. anisopliae can also sustain itself on the plant exudates in the rhizosphere. In return, fungi such as M. anisopliae translocate nitrogen from the insects directly into the plants (Behie et al. 2012). Other factors, such as the soil moisture, pH, organic matter, and chemical properties are also important in determining the abundance and infectivity of EPF (Rath et al. 1992; Jabbour and Barbercheck 2009; Garrido-Jurado et al. 2011a, b; Sharma and Marques 2018). Herbicide usage can also have distinct effects of the EPF persistence (Yousef et  al. 2015). Nonetheless, numerous abiotic and biotic factors shape the dynamics of EPF community in the soil, and it is difficult to access the exact effect of each of these. Main biotic components which influence EPF persistence and efficacy the most are: plants, soil microorganisms, and invertebrates (Sinha et al. 2016). Entomopathogenic fungi exhibit diverse lifestyle and mode of action. In presence of suitable conditions, EPF invade percutaneously into the host-insect body and cause mycosis. A brief pictorial description of the mechanism of infection of EPF is provided in Fig.  4.4. In the majority of cases, the entire infection cycle requires these following steps: (1) spore attachment to the cuticle; (2) germination of the spore, which initiates cascades of many reactions related to recognition and enzyme activation, both by the EPF and by the host-insect; (3) the penetration of host’s integument through hyphal tubes; (4) the suppression of host immune defence mechanisms by the fungus; (5) the alteration of the fungal morphology into a yeast-­ like phase, leading to the formation of hyphal bodies or “blastospores”, which circulate within the insect’s hemolymph leading to the insect death; and, (6) the attainment of the previous saprophytic phase, the typical hyphal form, which comes out of the dead body to produce new conidia (Vega et al. 2009; Vega et al. 2012; Chandler 2017).

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Fig. 4.4  Life cycle and mode of action of entomopathogenic fungi (saprotrophic and parasitic cycle). (Modified from Ortiz-Urquiza et al. (2015) and Lu and St. Leger (2016))

As the majority of the research has focussed on the entomophthoralean and the hypocrealean fungi, we will only discuss these two fungal groups in more detail. Entomophthoralean fungi are obligate insect pathogens and do not grow outside their hosts. Therefore, they are quite host-specific and restricted to only one family or genus. For the same reason, they are difficult to mass produce in an artificial culture medium. These fungi eject spores actively and rapidly upon the realisation of favourable conditions. This property makes them different from their hypocrealean counterparts (Steinkraus 2007). The life cycle of the entomophthoralean fungus is indeed quite complex and often involves at least two different types of spore-forms: resting and conidia. Condia spore-forms are responsible for infecting insect-hosts, especially when the hosts are active. Conidia are actively discharged from the dead host, although there are few exceptions, for e.g., Entomophthora thripidum is discharged by living thrips (Pell et al. 2001). Conidia are usually coated with preformed mucous, and they adhere to the integument upon landing on the insect cuticle. After conidial attachment, the formation of an “appressorium” (a specialised attachment cells below which the EPF penetrates the cuticle) facilitates fungal penetration. However,

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some entomophthoralean fungi such as Conidiobolus obscurus, Entomophthora planchoniana, and Pandora neoaphidis do not form appressoria, and penetrate the insect cuticle directly through the germ-tube (Vega et  al. 2012). The conidia are fragile and short-lived but germinate quite rapidly. In the cases when the primary conidia do not land on the host-insect surface, they can give rise to the secondary conidia, which can further produce the tertiary conidia. These supernumerary conidia increase the chances of fungal infection, although, after every passing generation, they become smaller in size. Interestingly, only secondary conidia are infective in Neozygites spp. (Pell et al. 2001). Mechanical pressure and cuticle-degrading enzymes are indispensable for cuticle penetration. Thereafter, some Entomophthorales initially grow as wall-less protoplasts, to avoid the insect immune response, and later turn into hyphal bodies. Others grow either as hyphal bodies or protoplasts within the insect host. Host ceases eating as the infection progresses, and several behavioural changes are noticed prior to death. These behavioural changes can be noticed in Entomophaga infecting acridid hosts, Pandora infecting the ant genus Formica, and those infecting flies such as M. domestica and Scatophaga stercoraria. The fungus multiplies within the host, and manipulates its behaviour by making the host climb to an elevated position (referred as “summit disease”) and kills it. In other cases, for e.g., infections by Erynia spp., some fungal structures (rhizoids) are formed to fasten the hosts more securely to the substrate (Małagocka et al. 2015). Some physiological changes, such as behavioural fever (an increase in the body temperature of infected hosts) generally originated when reaching warmer locations, is noticed in the house flies M. domestica infected by Entomophthora schizophorae or E. muscae. Furthermore, changes in the mate-seeking behaviour for healthy males is also noticed, which is characterised by an increased attraction towards the infected females. The fungi attack the tissues once the hemolymph has significant numbers of fungal cells, in order to fill the host body cavity with cells by the time the insect dies (Hajek and Meyling 2018). The Entomophthorales kill the host-insect by depleting most of the resources inside the hemolymph, probably trehalose sugar, and sporulate. This leaves host with no time for toxin production (De Fine Licht et al. 2016). As the host stays alive for some time before sporulation, it may even succeed to complete a reproductive cycle before dying (Gryganskyi et al. 2017). Most entomophthoralean species also produce resting spores for survival, when hosts are either not active or present. For example, during lower temperature periods, or at a late instar larval stage, or during a decreased photoperiod. The resting spores are produced inside the cadaver after host death and remain dormant. Resting spores can either be azygospores (formed when one hyphal cell rounds up at one end) or zygospore (when two hyphae unite) (Pell et al. 2001). These spores possess a thick wall, and are deposited in the soil when cadavers disintegrate. Resting spores may germinate if they find the hosts, however, many of them do not, at least in the first year of their genesis (Butt et  al. 2001). Moreover, in some cases, for e.g. Entomophaga maimaiga, only a part of the total resting spores germinate in a year. This is done to ensure fungal infectivity in the cases of prolonged seasons of host

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unavailability or inactivity. Resting spores may retain the capacity of germination for many years (Hajek and Meyling 2018). The hypocrealean fungi represent some of the well-known, easy to culture, anamorphic EPF, such as Metarhizium and Beauveria (Meyling and Eilenberg 2006, 2007; Quesada-Moraga et al. 2007; Garrido-Jurado et al. 2011a; Carlos et al. 2013; Rudeen et  al. 2013; Garrido-Jurado et  al. 2015; Steinwender et  al. 2015; Castro et  al. 2016; Fernández-Salas et  al. 2017; Gan and Wickings 2017; Sharma et  al. 2018b). The host range varies from being specialist pathogens, such as Hemiptera-­ specific M. album and white grubs specialist B. brongniartii, to generalists, such as M. robertsii and B. bassiana (Enkerli et al. 2004; Hu et al. 2014). Hypocrealean teleomorphs show a narrower host range (Boomsma et al. 2014). Infection occurs either through: (a) an actively discharged ascospore or (b) a passively transmitted conidia. Passive transmission is facilitated by rain splash or wind currents, contact with an infected individual, or dispersal by the bodies of arthropods (Boomsma et al. 2014). Conidia of some species are hydrophobic and have a proteinaceous rodlet layer, whereas, some species have hydrophilic conidia with a smoother surface (Hajek and Meyling 2018). Conidial adhesion of M. anisopliae is mediated by “Metarhizium adhesion-like proteins”, for e.g., MAD1 and MAD2, which are also responsible for conidial germination and blastospore formation. Carbon and nitrogen sources are required for spore germination and germ-tube formation in case of B. bassiana. Other EPF, such as Isaria farinosa and M. anisopliae, produce mucilage for fungal adhesion during the germ-tube formation. Cuticle penetration requires a set of enzymes, such as proteases, chitinases, and lipases (Vega et al. 2012). Hypocrealean anamorphs often grow inside the insect hemocoel as blastospores or hyphal bodies, by utilising available nutrients for growth and reproduction. Host death by EPF, such as M. anisopliae, is supposed to be facilitated by the fungi undergoing either a “toxin strategy” or a “growth strategy”. The toxin strategy suggests that the fungus employs the production of the secondary metabolites, such as destruxins, to kill the host, while maintaining very little vegetative growth. On the contrary, the growth strategy suggests that the fungus relies on a profuse growth inside the hemolymph, leading to the host starvation and death (Kershaw et  al. 1999). Nonetheless, a fungal hypha emerges from within the insect and sporulates to produce aerial conidia.

4.6  Conclusion With the growing concerns about the use of chemical pesticides, the use of microbial biopesticides does exhibit a great potential. The majority of the microbial pesticides either spend some part of their life cycle in soils, or somehow survive the harsher conditions in the soil environment as spores. Hence, understanding the diversity and ecology of the EB and EPF is of utmost importance. More studies on the ecology of entomopathogens will provide a better understanding of insect

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population cycles and pest outbreak. Strategies considering the bacterial and fungal ecologies within the agroecosystems may provide a better application design for the use of microbial entomopathogens in controlling pest-insects. Attributes such as higher efficacy, low production cost, desired specificity and enhanced persistence are always crucial in commercialising microbial agents for biological control. Moreover, the evolution of resistance in the target insect pest and the deaths of the non-target arthropods are some of the challenges needed to be addressed. Constant screening of toxins and virulence factors is always underway, and the discoveries of newer species, approaches for directed evolution, and genetic engineering provide enormous opportunities to be looked upon in the development of a biopesticide. Acknowledgement  This work is a part of L. Sharma’s PhD dissertation at the ‘University of Trás-­ os-­Montes and Alto Douro’, Vila Real, Portugal. The funding was provided by the ‘Centre for the Research and Technology of Agro-Environmental and Biological Sciences’ (CITAB) through the fellowship: BIM/UTAD/16/2018; and by the EcoVitis project. National Funds by FCT – Portuguese Foundation for Science and Technology, UID/AGR/04033/2013; and the European Investment Funds by FEDER/COMPETE/POCI Operacional Competitiveness and Internationalization Programme, under the Project POCI-01-0145-FEDER-006958, are also acknowledged. Due to the limited space, we could not cite many relevant articles. However, we deeply acknowledge those works.

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Chapter 5

Ascomycota and Integrated Pest Management Tariq Ahmad, Ajaz Rasool, Shaziya Gull, Dietrich Stephan, and Shabnum Nabi

Abstract Employing fungal pathogens to combat insect pests of agricultural importance has gained momentum due to its ecofriendly approach, availability and host specificity. Successful future prospects include efficiency and improvements in the research methods, proper selection of strains, mass production, genetic manipulations and other innovative techniques. Further aspects are the preparation of formulations that will increase persistence, longer shelf life and ease of application, pathogen virulence and spectrum of action. A description of Ascomycetes biology, taxonomic characters and mode of action is presented here with a focus on its role in Intergrated Pest Management strategies. Recent studies on genetic modifications for improving their virulence are also discussed. Keywords  Ascomycota · Fungi · Entomopathogens · Pest · IPM · Biological control

T. Ahmad (*) Federal Research Centre for Cultivated Plants, Institute for Biological Control, Julius Kühn Institute, Darmstadt, Germany Entomology Research Laboratory, Postgraduate Department of Zoology, University of Kashmir, Srinagar, Jammu and Kashmir, India e-mail: [email protected] A. Rasool · S. Gull Entomology Research Laboratory, Postgraduate Department of Zoology, University of Kashmir, Srinagar, Jammu and Kashmir, India D. Stephan Federal Research Centre for Cultivated Plants, Institute for Biological Control, Julius Kühn Institute, Darmstadt, Germany S. Nabi Interdisciplinary Brain Research Centre, J. N. Medical College, Aligarh Muslim University, Aligarh, Uttar Pradesh, India © Springer Nature Switzerland AG 2019 M. A. Khan, W. Ahmad (eds.), Microbes for Sustainable Insect Pest Management, Sustainability in Plant and Crop Protection, https://doi.org/10.1007/978-3-030-23045-6_5

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5.1  Introduction With the world population expected to reach 10–12 billion by 2100 (UN Report 2011) the agricultural sector is facing the biggest challenge of providing enough food to meet the pressing demands of nations. To meet this ever increasing demand, efforts have been done towards improving crop varieties for higher quality and yields, shorter durations of life cycle and resistance to insect pests and diseases, which are responsible for 20–40% crop losses, annually (FAO 2012). Integrated pest management (IPM) is a broad-based approach that integrates practices for economic control of pests and aims at suppressing pest populations below economic injury level (EIL). It emphasizes the growth of a healthy crop with least possible disturbance to agro-ecosystems, and encourages natural pest control mechanisms. IPM makes use of a range of actions including cultural controls, physical barriers, chemical control or pesticides, and finally biological control, including, adding and conserving natural pest predators and enemies. For conventional farms, IPM reduces human and environmental exposure to noxious chemicals, and potentially lowers overall costs. It extends the concept of integrated control to all classes of pests and includes all tactics. Other tactics, such as host-plant resistance and cultural manipulations also became part of the IPM framework while uniting Entomologists, Plant Pathologists, Nematologists and Weed Scientists at one platform to define organized and integrated strategies against a particular pest species. Entomopathogenic fungi (EF) trials were started by Russian Microbiologist, Elie Metchnikoff way back in 1888 (Samson et al. 1988; Glare and Milner 1991), by mass producing fungal conidia on sterilized brewer’s mash and combined cultures with sand granules for spreading on field crops. Although his results were inconsistent, the work of Metchnikoff (1888) ignited inquisitiveness around the world and led to programs in Europe and United States for experimentation with “friendly fungi” against insect pests (Lord 2005). Boverin, a Beauveria bassiana-based mycoinsecticide was developed in 1965 for the control of Colorado potato beetle and codling moth in the former USSR (Kendrick 2000). The first formal and published proposal of microsporidia to control grape phylloxera, Daktulosphaira vitifoliae (Fitch) was suggested by LeConte (1874). The study offered imaginative ideas for an effective and economic alternative method of pest control and paved the way for future scientists to study EF. Owing to greater environmental awareness, food safety concerns and insecticide resistant species, the application of EF is growing by leaps and bounds (Shahid et al. 2012). Ascomycota, the largest and main phylum of the Kingdom Fungi with over 64,000 species (Kirk et  al. 2008), is one of the most diverse eukaryotic phyla, encompassing some important species in biological control programmes throughout the world. Ascomycetes are very varied and can be identified from the fruiting bodies and the way in which the asci develop. They either act in nature per se or have been manipulated and used worldwide to control various pest species, with some level of success in Asia and South America (Fuxa 1987). The distribution of some species is variable and limited while some are cosmopolitan such as the white

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­truffle, Tuber magnatum found in isolated locations in Italy and Eastern Europe (Mello et al. 2006). Around 90% of microbial pesticides are used as insecticides, accounting for the 1.3% of the world’s total pesticide market (Hajek and Leger 1994), with few commercial formulations of ascomycetes developed for crop pest management. Among 171 products of EF developed, products based on B. bassiana and M. anisopliae represent 33.9%, while Isaria fumosorosea and B. brongniartii represent 5.8 and 4.1% respectively (Moorhouse et al. 1992; De Faria and Wraight 2007). Although mycoinsecticides have been found effective in controlling various insect pests of economic importance, successful marketing and utilization of these products have been somewhat slow, largely due to low production efficiency, high cost, low performance under testing environmental conditions and lack of awareness programs. Nonetheless, these mycoinsecticides have potential to play a key and defining role in IPM programs while focusing on formulations with increased persistence, longer shelf life and pathogen virulence, ease of application and spectrum of action.

5.2  Biology and Taxonomy Ascomycetes comprise 75% of all the described fungi speciesThey are heterotrophs and obtain nutrients from dead or living organisms (Carroll and Wicklow 1992) while as biotrophs they form symbioses with algae (lichens), the leaves and stems of plants (endophytes), and roots (mycorrhizae). Ascomycota are either single-­ celled (yeasts), filamentous (hyphal) or both (dimorphic). While most yeasts and filamentous Ascomycota are haploid, some species such as Saccharomyces cerevisiae can also be diploid. Ascomycetes are mostly terrestrial or parasitic, however, few have adapted to marine or freshwater environments whilst in most land-based ecosystems, and act as decomposers facilitating detrivores to obtain nutrients. The key character that defines Ascomycota is the ascus, a special, elongated cell or sac within which nuclear fusion and meiosis take place, and that give the group its name. Ascomycetes are ‘spore shooters’ and produce microscopic spores insidethe asci. Asexual reproduction is the dominant form of propagation, a part of reproducing by means of haploid conidiospores and through budding. The ascus, which is the result of the sexual reproduction, is a tube-shaped vessel, a meiosporangium, containing the sexual spores called ascospores. It starts with the development of special hyphae from either one of two types of mating strains. The “male” strain produces an antheridium while the “female” strain develops an ascogonium which combine in plasmogamy at fertilization. Later on, special ascogenous hyphae arise, in which pairs of nuclei migrate: one from the “male” strain and one from the “female” strain. In each ascus, two or more haploid ascospores fuse their nuclei in karyogamy, giving rise to diploid nuclei which, by the process of meiosis, give rise to haploid nuclei. The ascospores are then released, germinate forming hyphae and are disseminated in the environment to start new mycelia.

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Fungi and fungus-like organisms are usually defined by their heterotrophic, absorptive mode of nutrition and their apical hyphal growth (Deacon 1997). The classification of fungi has been traditionally defined upon morphology (e.g. Conidiogenesis) and ultrastructure (e.g. cell wall and septum structure). The Phylum Ascomycota, together with the Basidomycocetes, forms Sub-Kingdom Dikarya which includes members that form septate haploid hyphae and yeasts. More precise placement of EF into defined taxonomic groups has been achieved by approaches such as metabolisms comparison (e.g. nutrient utilization, enzyme and toxin production) and/or using genetic analyses (e.g. rRNA sequences, karyotyping, mtDNA restriction length polymorphism (Khachatourians 1992). Actually, true fungi are placed in four Phyla (Ascomycota, Basidiomycota, Chytridiomycota, and Zygomycota) plus one artificial form, the Phylum Deuteromycota, that contains filamentous fungi. The Deuteromycetes, also known as Fungi imperfecti, are actually thought as ascomycetes or basidiomycetes which have either lost their potential for sexual reproduction or comprise undescribed sexual forms (Barr 1992). There are only three subphyla in the phylum Ascomycota that have been described (Berbee and Taylor 1992): • The Pezizomycotina is the largest subphylum of Ascomycetes producing ascocarps (fruiting bodies), except for the genus Neolecta which includes most macroscopic “ascos” such as truffles, ascolichens, ergot, cup-fungi (discomycetes), pyrenomycetes, lorchels, powdery mildews, dermatophytic fungi, and Laboulbeniales. • The Saccharomycotina comprising “true” yeasts such as baker’s yeast and Candida, that reproduce vegetatively by budding. • The Taphrinomycotina originally named Archiascomycetes (or Archaeascomycetes), including a disparate and basal group within the Ascomycota, and recognized by molecular analyses. It includes hyphal fungi (Neolecta, Taphrina, and Archaeorhizomyces), mammalian lung parasite Pneumocystis and fission yeasts (Schizosaccharomyces). Supraordinal fungal classification in Fungal Tree of Life (AFTOL) listed 3 Subphyla, 14 Classes, and 60 Orders of Ascomycota (Hibbett et al. 2007). Of the five classes that contain EF, most species are found in Laboulbeniomycetes and Pyrenomycetes (Tanada and Kaya 1993). The class Hemiascomycetes, consisting of yeast like entomopathogens, includes species typically slow acting and cause chronic infections, such as infection of the biting midge (Dasyhelea obscura) by Monosporella unicuspidata. Members of the class Plectomycetes are responsible for chalkbrood disease in bee colonies. The class Loculoascomycetes is characterized by bitunicate asci, which are released in specialized stomatic compartments called locules. Two entomopathogenic orders of this class, Myriangiales and Pleosporales are considered important pathogens of scale insects. The Pyrenomycetes, a major class in Ascomycota, possess distinguishing unitunicate cylindroid asci and consist of all EF species in order Sphaeriales, most of which are described within the genus Cordyceps including over 250 species. Both

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Exopteygota and Endopterygota insect orders are infected by members of this genus, and some are even pathogens of spiders. Cordyceps spp., abundant in tropical forest ecosystems, is an important insect pathogen (Samson et al. 1988) infecting numerous soil dwelling pests. Laboulbeniomycetes is the other major class of Ascomycota composed of minute, ostensibly inconspicuous fungi that were originally thought to be external commensals of insects (Tanada and Kaya 1993). An estimated 115 genera of this class are known and only few entomopathogenic species have been found in a wide variety of habitats (Carruthers and Hural 1990). Although Coleoptera come into view to be the most common targets, these are known to be pathogenic to members of at least 11 insect orders with most species reported from Aspergillus, Lecanicillium (formerly Verticillium), Metarhizium, Beauveria, Nomuraea, Aschersonia, Hirsutella, Culicinomyces, Paecilomyces and Sorosporella. Most of these genera have been linked to one or several ascomycete genera. Such linkages can be demonstrated either biologically (an insect infected with an anamorph dies and a teleomorph is produced) or by using molecular tools showing the genetic relationship between anamorphs and teleomorphs (Huang and Erickson 2002; Liu et al. 2002).

5.3  Mode of Action Entomopathogens typically have unique mode of action and infection while coming in contact with their arthropod host. With ideal conditions of moderate temperature and relative humidity, their spores germinate and breach the host cuticle with the combined aid of enzymes and mechanical pressure. Later on the germ peg invades the tissues, multiply yielding a mycelium that produces more spores, killing the host by a variety of means such as starvation through multiplication or production of toxins. The mode of action of various ascomycete species is discussed as follows.

5.3.1  Beauveria spp. Beauveria is an EF that dwells in soil all over the world. Some strains are exceedingly adapted to specific host insects with high potential to be used as biological insecticides against pests. The conidiospores germinate on the superficial area of the cuticle, resulting in the formation of long hyphal tubes directly piercing the insect host integument. After crossing this layer, the fungus shows morphological changes, becomes yeast like and produces hyphal bodies which move in the haemolymph and start fast budding. This yeast-like phase is essential for the pathogenicity and subsequent propagation by conidia by emerging mycelia. Attachment of fungal spores to the host is mostly passive and represents the first event for mycosis development. Spores are randomly transferred by various agents like air and water.

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Dry spores of B. bassiana have an outer layer made of interwoven fascicles of hydrophobic rodlets. Each conidial stage has a unique rodlet layer which is never found on the vegetative cells. Rodlets exert non-specific hydrophobic forces which result in hold of dry spores to the cuticle (Boucias and Pendland 1988). Nevertheless, the mechanism, that is yet to be known in detail, results in yielding adhesive forces between conidia and the insect cuticle (Latge and Monsigny 1988). When the fungal spores reach the haemocoel, it shows rapid germination and growth which is enhanced by nutrient availability, water, oxygen and temperature. Beauveria enters through the cuticle into the insect body to acquire nutrients for reproduction and growth. It results in both an enzymatic activity as well as a mechanical pressure allowing entering the host, owing to physical separation by inserted hyphae. Different pathogenic enzymes such as chitinases, lipases, esterases and at least four different classes of proteases, are responsible for breaking of host cuticle, degrading its major components. Enzymes such as endoproteases (PR1 and PR2) and aminopeptidases are produced on the cuticle, coincident with formation of appressoria. Proteolytic enzymes are formed at faster pace in comparison to N-acetylglucosaminidase (St Leger and Butt 1989). The most important factor for of cuticle breaking is the activity of endoproteases, although other enzymes synergistically work together for penetration. Beauveria spp. also release lethal fungal toxins leading to cellular disturbances in the host. Many insects show changes in behaviour such as overall paralysis, sloth appearance and decreased petulance, due to neuromuscular toxins. Toxins extracted from the infected host body include beauvericin, beauverolides, bassianolide and isarolides (Hamill and Sullivan 1969; Elsworth and Grove 1977), lethal to tissues leading to cellular disturbances and death.

5.3.2  Isaria spp. Isaria (I. farinosa and I. fumosorosea), includes filamentous fungi attacking various insect pests, worldwide. Isaria spp. can be used as biocontrol agents for controlling predominantly white flies. They produce cuticle destroying enzymes viz. proteases (Pr1 and Pr2), chitinases, chitosanases and lipases. They act as a strong antifeedant decreasing the host feeding potential and body weight (Shaukat et al. 2010). They target the host cuticle through the enzymatic action of chitinases. The chitinase gene isolated from the genome of I. fumosorosea (Ifchit1) encodes a polypeptide of 423 amino acids. Ifchit1 chitinase has proved to be a virulent pathogenic factor towards the host (Zhen et al. 2016). When in contact with a specific host, Isaria becomes rapidly pathogenic when the conidia are inoculated on the hind wings. The fungus shows a higher response in comparison to the conidia attachment to other part of the body, due to a germination rate which is higher on wings.

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5.3.3  Metarhizium spp. Metarhizium, formerly known as Entomophthora anisopliae, is an EF having worldwide distribution, acting as an insect parasite. Metarhizium spp., like other entomopathogens, penetrate the cuticle by adhesion of asexual, dry spores and get attached to the upper waxy insect coating by hydrophobic, electrostatic forces and protein interactions between conidia and host integument. Different species have specific genes for pathogenicity, such as Mad 1 kinase gene of M. anisopliae. Asexual spores of Metarhizium spp. contain hydrophobins, that increase adhesion properties reinforcing attachment (Ment et al. 2010a). The adhesion is largely dependent on environmental factors and on the chemical composition of hosts (Santi et al. 2010). At the onset of germination, concentration of trehalase, that uses the sugar trehalose present in the haemolymph, increases. The process is initiated by the presence of exogenous carbon and nitrogen (Ment et  al. 2010b). Trehalase gives energy for spores development and swell up, germ tubes formation and subsequent development of the appressorium (Lovett and Leger 2015). Mad 1 and Mad 2 are formed by replacing hydrophobins so that conidia attach to the integument in a firm way. Besides, some fatty acid esters, amino acids, glucose and peptides are also required during germination. After germination, appressorium formation takes place with the help of two genes, ODC1 and Mpl1. Through the conidial germination, ODC1 translates for ornithine decarboxylase and appressorium is formed from the germ tube (Wang and St Leger 2007a, b; Pulido et al. 2011). Mechanical and turgor pressure is generated by conidia penetrating inside the insect cuticle, a process enhanced by a mucilage layer (Greenfield et al. 2014; Staats et al. 2014). In M. anisopliae, another enzyme MAPK (Mitogen-Activated Protein Kinase) is present that helps in regulation of the expression of Mad1 gene for adhesion, while appressorium development is highly enhanced by the gene MaMK1 which is an extracellular signal-­ regulated kinase. Tetraspanins determined by the MaPls1 gene also helps in turgor pressure and in the development of the appressorium in a specific pathway (Luo et al. 2013). Penetration of conidia is the next stage in M. anisopliae that results in the secretion of proteins such as chymotrypsins, trypsins, carboxypeptidases and subtilisins by which the protein rich procuticle of arthropods is digested (Wang et al. 2008). M. anisopliae produces many types of proteins which affect different hosts and are very specific. Enzymes such as subtilisin proteases (Prl) are found to be degrading cuticle by hydrolysing the integument, while other enzymes such as chitinases work with other proteases to pierce the cuticle (Butt et al. 2013). On the surface of conidia, lipases are also present that enhance adhesion of conidia by lipolytic activity and results in the release of free fatty acids (Beys da Silva et al. 2011). All these enzymes work synergistically to enhance penetration and are mostly dependent on the amount of nutrients present in the host haemocoel . The chi2 gene, which encrypts for an endochitinase, is also decisive for the infiltration of the host cuticle by plummeting the lethal time required to kill a specific insect host (Boldo et al. 2009). Certain substances, such as tetraspanins, help in regulation of apsA and kinesin, responsible for the structure and migration of the cytoskeleton. It uses

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nutrients proceeding from the host cuticle to start germination. All the cellular activities are under the control of GTPases and tetraspanins, that code MaPlsl while helping the fungus to thrive in the host environment. Colonization is the most important stage in the pathogenicity caused by M. ansiopliae in the haemolymph of host insect. Dextruxin A and dextruxin E, natural insecticidal molecules, act on the immune system of the host suppressing all the cellular activities and encapsulated spores are released. The gene dtxS1 is responsible for the synthesis of dextruxin and is mostly found in Metarhizium (Wang et al. 2012). However, some other fungal proteins elude the host immune system, such as the evasion proteins Mcl1 The glucose required for their activity is provided by trehalose from the trehalases released in the haemolymph (Ment et al. 2010b). When germinated conidia enter the host body, there is a fast multiplication and hyphal differentiation step, controlled by the Mad1 protein and in turn, these proteins regulate every activity in the cell cycle (Wang and St Leger 2007a). Cytochrome P504s genes are also found in M. acridium, which reduces the antimicrobial host activity. Finally during sporulation, hyphal growth is enhanced due to the action of a Hog 1 kinase, resulting in a thick layer of green spores formed on the dead body of infected insects (Ocampo and Caoili 2013).

5.3.4  Lecanicillium spp. Lecanicillium, earlier identified as Verticillium (i.e. V. lecanii), is one of the most important EF with a high potential to be used as commercial biopesticide. It relies on hydrolytic enzymes and mechanical forces to pierce into the integument of the host. They act as mycoparasites to plant pests (Goettel et  al. 2008) being highly virulent against aphids and white flies. Lecanicillium spp. have a wide host range and are known as white halo fungi as their mycelial growth is white in colour. Their potential and virulence have been reported on pests such as thrips, mealy bugs, scale insects, white flies (Horn 1915; Ekbom 1979a, b; Kanagaratnam 1982). These fungi can be grown on potato dextrose medium and are initially white in colour and later develop light yellow nuances in the medium (Naejrech 1973). Lecanicillium spp. act on the host while contacting its integument and get attached to the outer, waxy, epicuticle coating. Afterwards, they start germinating with hyphae formation that penetrate the cuticle directly (Hughes and Gillesipie 1985). However, at times they may grow on the external surface of the cuticle as well. Various extracellular enzymes are responsible for piercing the integument such as chitinase, protease, esterase, n-acetyl glucosamine, endoprotease, carboxypeptidase A, PR 1- chymoelastase serine protease and endoprotease (St Leger and Cooper 1986, 1987). All these enzymes work synergistically for breaking the cuticle wall, while PR-1 acts as the major cuticle breaking enzyme, by increasing its concentration at the point of penetration. Later on they start forming blastospores in the form of hyphal growth. Infested insects die due to combined effect of mycotoxins and outer mechanical pressure (Ferron 1981). Beauvericin, dipicolinic acid (Claydon and Grove 1982),

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bassianolide (Kanaoka 1978), vertilecanin-A1, decenedioic acid and 10-hydroxy 8-decenoic acid (Soman 2001) are some of the chemicals present in the process. When the nutrient concentration is reduced inside the host body, hyphae are protruded outside to form a mycelial mat on the external body surface, which rapidly leads to the host insect death (Yeo 2000).

5.3.5  Nomuraea rileyi Nomuraea, a potential dimorphic hyphomycete EF, is effectual on various pests, mostly Lepidoptera and Coleoptera (Ignoffo 1981). It is a versatile fungus, with a broad host range, widespread occurrence and infecting any developmental stage. Its mode of action starts with the attachment of conidia to the insect body (Srisukchayakul et al. 2005). On the host epicuticle, tube-like structures are formed, puncturing the endocuticle and producing hyphal bodies inside the haemocoel, leading to the insect death. Conidiophores are formed at the end of the infection cycle, when mycelia develop from the insect cuticle surface (Srisukchayakul et  al. 2005). The fungus penetrates the body wall by both mechanical pressure and enzymatic activity, as well as by secreting on the epicuticle various proteinaceous substances such as proteinases, lipases and chitinases. It starts budding in the insect haemocoel, when hyphal bodies are formed. It has been reported that N. rileyi produces one specific insecticidal toxin (Ye et al. 1993) which causes infection to sap sucking insects and several noctuid pests, with a potential to infect any developmental stage including pupae (Ferron 1978; Anand et al. 2009).

5.3.6  Hirsutella thompsonii Hirsutella thompsonii is cosmopolitan in distribution and considered a very important entomopathogen. It is highly effective on mites, penetrating through the legs and forms hyphal bodies inside the haemolymph. Spores which are formed inside the body show their emergence through openings such as mouth and genitalia, and then from all over the body. The insects which are infected by this fungus in in-vivo conditions show the same general mechanism, i.e. conidia gets adhered to the cuticle, followed by germination and body penetration (Liu et  al. 1996). Hirsutella thomopsonii is highly effective towards various arthropods such as mosquito larvae, aphids and mites (Krasnoff and Gupta 1994). It secretes an insecticidal polypeptide chain named Hirsutelin A (Ht A), showing ribosomal inhibiting activity on the pests when infected. Ht A has insecticidal properties (Liu et al. 1995; Mazet et al. 1995). Its toxicity was observed in vivo against adult citrus rust mite, Phyllocoptruta oleivora, which is its natural host (Omoto and McCoy 1998). Ht A has an ability to form a ribonucleolytic enzyme and other ribotoxins (Liu et al. 1995) and acts on insect cells at increasing concentration of phosphatidyl ethanolamine and

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phosphatidylinositol (Marheineke et al. 1998), increasing permeability due to the presence of thin membranes and ribotoxin targets. Ht A causes death of larvae upon infection, albeit infection rates depend on the ribotoxin concentration (Lacadena et al. 1995). In addition, H. thompsoni causes cellular damage by generating oxidative stress in the insects and by the action of enzymes such as CAT, GPx and SOD (Fornazier et al. 2002).

5.3.7  Aschersonia spp. Species of Aschersonia are predominantly effective biocontrol agents of whiteflies. Immature stages, especially the first, second and third nymphal instars of whiteflies are susceptible to infection by Aschersonia while all other substages, from fourth nymphal instar to adults, are usually less susceptible (Fransen 1987). Its action starts with the attachement of conidia to the host insect cuticle. Changing the attachment from passive to active induces an enzymatic process of secretion of various mucilaginous substances, by germinating conidia (Fargues 1984). There is a visible colour change in the host at early infection stages, as germ tubes are produced on the cuticle which start penetration or form an appressorium. All these events involve enzymatic and physical activities whilst hyphal bodies are produced inside the host. They circulate in the haemolymph before germinating to yield a mycelium on which masses of spores are produced. The fungus pervades all body organs while protruding outside, resulting in the host death (Roberts and Humber 1981). Under optimal conditions, the first symptoms occur within 24–48  h. A vigorous hyphal growth takes place in 4–6  days and production of conidia occurs usually from 7–9  days after initiation of infection (Fransen 1987). Relative humidity plays an important role in the infection cycle. Conidial germination is retarded below 98% RH and is impaired below 90% (Fransen 1987).

5.4  Ascomycetes in IPM Programmes EF are known to infect insects of almost all orders, and especially Hemiptera, Lepidoptera, Diptera, Coleoptera, Orthoptera and Hymenoptera (Ramanujam et al. 2014; Khan and Ahmad 2015). Nymphal and larval stages are more susceptible to EF attacks than adults, while in others the reverse may be true. Some fungi such as Aschersonia aleyrodis have restricted host ranges and infect only whiteflies, whereas N.rileyi infects only lepidopteran larvae. Others like B. bassiana and M. anisopliae infect more than 700 species in several insect orders and have been tested against aphids, whiteflies, thrips, and a few against coleopteran and lepidopteran pests. Ascomycota harbour most number of species, some of which play a key role in IPM and natural regulation. Various species in Ascomycota may provide effective

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long term and short term control including members of Beauveria, Verticillium, Isaria, Aschersonia, Hirsutella, Metarhizium and Nomuraea, that are integral components of IPM used in myco-biocontrol of insect pests.

5.4.1  Beauveria spp. Beauveria bassiana (family Cordycipitaceae), is one of the widely and popularly used entomopathogens. It is the anamorph (asexual form) of Cordyceps bassiana, the sexually reproducing form (teleomorph) collected only in eastern Asia (Li et al. 2001). In cultures, this fungus grows as a white mould producing dry, powdery distinctive conidia in white spore balls. Rehner and Buckley (2005) argued that B. bassiana consists of many diverse lineages that should be recognized as distinct phylogenetic species. For a long time B. bassiana has been known as the most common causal agent of the disease related to dead and moribund insects in nature (Mcleod 1954). It has also been scrutinized worldwide as a microbial control agent of hypogeous insect species (Ferron 1981). Many curculionid weevils with subterranean larval stages are highly susceptible to it (Beavers et al. 1983). Growing naturally in soils throughout the world, B. bassiana acts as a pathogen on various insect species, inflicting diseases such as the white muscardine (Sandhu and Vikrant 2004; Thakur et al. 2005; Jain et al. 2008). Some of the hosts of agricultural and forest importance of Beauveria include Codling moth, the Colorado potato beetle and several genera of termites, the American bollworm, Helicoverpa armigera, the teak defoliator, Hyblaeapara and teak skeletonizer, Eutectona machaeralis (Thakur and Sandhu 2010). With worldwide distribution, B. bassiana is also used as a biological insecticide to control a number of pests such as termites, whiteflies, and malaria-transmitting mosquitoes (Hamlen 1979). When used as an insecticide, spores are sprayed on affected crops as an emulsified suspension or wettable powder. While acting as a non-selective biological insecticide, it parasitizes a wide range of arthropod hosts. It has been effective against pine caterpillars, Dendrolimus spp., the European corn borer, Ostrinia nubilalis and green leafhoppers, Nephotettix spp. Pathogenity studies of two isolates of B. bassiana, M. anisopliae and Paecilomyces farinosus against soldier ants under laboratory conditions revealed high mortality rates around 80%, killing the ants in the first four days after inoculation (Loureiro and Monteiro 2005). In China, one of the major forest pests, the pine caterpillars Dendrolimus spp., have been successfully controlled through aerial applications of B. bassiana in oil, spraying up to 300,000  ha of forest cover over a 5-year period, resulting in mortality ranging from 43% and 93% (Pan and Zheng 1988). Our recent studies on walnut pests and their management by entomopathogens showed efficacy of Beauveria bassiana on Myllocerous fotedari. (Fig. 5.1a).

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Fig. 5.1 (a) Myllocerous fotedari infested with Beauveria bassiana (b) Verticillium lacanii infecting whitefly (c) Bagrada bug killed by the green fungus Metarhizium anisopliae (d) Spodoptera larvae infected with Nomuraea sp. (Elangbam et al. 2016)

5.4.2  Lecanicillium lecanii Lecanicillum lecanii (previously known as Verticillium lecanii) is another widely distributed EF with broad host ranges, affecting insects of the orders Homoptera (Milner and Lutton 1986; Etzell and petitt 1992), Coleoptera (Barson 1976) Lepidoptera (Gopalakrishnan 1989) and Orthoptera (Khachatourians 1992). Some plant disease pathogens such as cucumber powdery mildew (Verhaar et al. 1996) and Chrysanthemum rust fungi (Whipps 1993) may be also affected by this fungus. Kim et al. (2002) reported that L. lecanii was an effective biological control agent against the greenhouse whitefly, Trialeurodes vaporariorum in South Korean greenhouses (Fig. 5.1b). While attacking nymphs and adults, it sticks to the leaf underside by means of a filamentous mycelium (Nunez et al. 2008). In the ‘70s, L. lecanii was used to control whiteflies and several aphid species, including the green peach aphids (Myzus persicae) in greenhouse cultivated Chrysanthemum (Hamlen 1979). Aspergillus flavus and L. lecanii were applied to control Helopeltis spp. (Hemiptera: Miridae) under laboratory conditions (De Faria and Wraight 2007). Pathogenicity test revealed that the average mortality of Helopeltis spp. occurred from the second until the seventh day after inoculation, and that mortalities were 90% with Aspergillus sp., 80% with A. flavus and 77% with L. lecanii. The latter has been also reported to inflict natural epizootics in aphid and scale populations in

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tropical and sub-tropical regions. It was also the first fungus to be studied and ­developed for use as an inundative mycoinsecticide in glasshouses and available in the form of two products manufactured by Koppert Biological Systems in the Netherlands (De Faria and Wraight 2007). The product contains different isolates as active ingredients such as “Vertalec” against aphids and “Mycotal” against whiteflies and thrips and is registered in Denmark, Finland, Netherlands, Norway and UK.

5.4.3  Metarhizium spp. Metarhizium anisopliae is explored for IPM of various notorious insect pests (Sandhu and Mishra 1994). Several strains of M. anisopliae have been developed as biological control agents for grasshoppers, locusts, cockchafers, spittlebugs, grubs, bagrada bugs and borers (Fig. 5.1c). A complete bioactivity of M. anisopliae tested on teak skeletonizer, Eutectona machaeralis has confirmed its potential as a myco-­ biocontrol agent also for this pest (Sandhu et al. 2000). Between 1985 and 1989, due to outbreaks of the Desert locust, Schistocerca gregaria, a devastating and destructive pest of crops and pasture grasses in many parts of Africa, extensive chemical insecticides were applied for control. Later on, looking into the possibility of using EF as biological control agents, a collaborative research programme, prompted by international concern on adverse environmental impacts of insecticide applications, was initiated between research institutes in the UK, the Netherlands, and the Republics of Benin and Niger (Prior and Greathead 1989). During the course of the study, M. anisopliae var. acridum was found to be an important natural pathogen of locusts and grasshoppers (Shah et al. 1997; Driver et al. 2000). Currently, a Metarhizium-based mycoinsecticide is supplied in sachets containing dried conidia which can be mixed with diesel or kerosene oil before spraying for pest control (Bateman et al. 1998). Spraying this mycoinsecticide led to infection and death rates around 70–90% of treated locusts or grasshoppers within 14–20 days after application, without negative effects on non-target organisms (Lomer et al. 2001). “Green Muscle”, a patented product of M. anisopliae, is recommended for locust and grasshopper control by the Food and Agriculture Organisation of the United Nations (Lomer et al. 2001). A number of entomopathogenic Hyphomycetes, including M. anisopliae and B. bassiana, commonly isolated from termite colonies demonstrated considerable potential for controlling termites (Milner et al. 1998a). Application of M. anisopliae conidia to mound- and tree-nesting termites in Australia showed substantial mortality (Milner and Staples 1996; Milner et al. 1998b). In Brazil, a high prevalence of termite mortality (100%) was observed in 19 of 20 nests treated with M. anisopliae. Brazil is the single largest country where commercial biopesticides based on M. anisopliae are used against spittlebugs on sugarcane and grassland, annually (Li et al. 2010). Rangaswami et al. (1968) reported 100% mortality by Metarhizium sp. in Pyrilla purpusilla in field conditions, besides controlling spittle bugs (Mahanarva posticata) in sugarcane (Ferron 1978). Mweke et  al. (2018) evaluated the

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p­ athogenicity of 23 fungal isolates including M. anisopliae, B. bassiana and Isaria sp. against adults of Aphis craccivora in the laboratory. All the fungal isolates were pathogenic to A. craccivora with mortality rate between 34.5 and 90%. Adults of the coconut-palm beetle, Oryctes rhinoceros, one of the major pests of Asian- and Pacific-grown coconut and oil-palms, feed on palm fronds, boring into the axils and destroying plant tissues (Bedford 1980). Its larvae are naturally infected by M. anisopliae which considered as an indispensable natural mortality factor (Carruthers and Soper 1987).

5.4.4  Nomuraea sp. Nomuraea rileyi is responsible for epizootics in various insects populations. Many insect species belonging to Lepidoptera, including Spodoptera litura and some Coleoptera, are susceptible to N. rileyi (Ignoffo 1981) (Fig. 5.1d). The host specificity of N. rileyi and its ecofriendly behaviour encourages its use in insect pest management, as shown by epizootics in populations of several noctuid pests (Tang and Hou 1998). Its infection and development have been reported for several insect hosts such as Trichoplusia ni, Plathypena scabra, Heliothis zea, Bombyx mori, Pseudoplusia, Helicoverpa armigera and Anticarsia gemmatalis. Spilosoma moths are severely attacked by N. rileyi and various species were studied in detail for mycobiocontrol properties (Mathew et  al. 1998). In addition, an epizootic of N. rileyi observed on Hedge plant eater, Junonia orithya proved to be the best alternative for management (Rajak et al. 1991).

5.4.5  Isaria spp. Formerly known as Paecilomyces, Isaria spp. have the ability to grow extensively over leaf surface under humid conditions thus enhancing their spread through whitefly populations (Wraight et al. 2000). Natural epizootics of these fungi suppressed Bemisia tabaci populations (Seryczynska and Bajan 1975) while its closed taxon, Paecilomyces fumosoroseus (Wize), one of the key natural enemies of whiteflies worldwide, causes the disease known as “Yellow muscardine” (Nunez et al. 2008). It has strong epizootic potential against Bemisia and Trialeurodes spp. in greenhouse and open field environments. It has been reported to cause considerable reductions in B. tabaci populations during prolonged periods of cool and humid conditions in the field or greenhouse or immediately following rainy seasons (De Faria and Wraight 2007). Kim et al. (2002) reported that P. fumosoroseus is paramount for controlling the nymphs of whitefly, while it covers their body with mycelial threads and sticks them to the leaves underside (Nunez et al. 2008). Paecilomyces furiosus is also used to control Culex pipiens (Sandhu and Mishra 1994). Furthermore, isolates of Paecilomyces induce highest mortality rates in European pepper moth,

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Duponchelia fovealis (Lepidoptera: Crambidae), a greenhouse pest of cut flowers, vegetables, and aquatic plants in northern Europe and Canada (Matuzzia et al. 2016).

5.4.6  Aschersonia sp. Aschersonia is characterised by bright coloured stromata, filiform ascospores and pycnidial to acervular anamorphs. It was recognised as an effective biocontrol agents in Florida for control of whitefly, Aleyrodes citri (Fawcett 1908) and of citrus white fly, Dialeurodis citri in USSR. Aschersonia spp. were introduced from India, China, Japan, Vietnam, USA and Cuba. Protsenko (1967) reported 80% larval mortality of white flies with foliar spray of conidia. Experiments carried out by Uchida (1970) reported fall in D. citri effectiveness. Similar experiments carried out by Solovey and Koltsov (1976) found 83% mortality of orange white fly, Aleurocanthus spiniferus. The pathogenicity and effectiveness of Aschersonia spp. was carried out on a wide range of insects, indicating their potential role as biocontrol agents (Spassova et al. 1980; Ramakers 1983 & Ellis et al. 2002).

5.4.7  Hirsutella spp. The entomopathogenic Hirsutella species have a worldwide distribution and are known to infect larval, pupal, and adult stages of insects living in diverse habitats. The genus Hirsutella include members which represent mesothermic mycopathogenens of pest and non pest mites, insects and nematodes. These are known to be host specific and regulate host populations in a density dependent relationship. Many members within the genus Hirsutella are anamorphs (asexual state) of telomorphs (sexual state) within genera Cordyceps and Torrubiella (Ascomycota: Hypocreales) (Hywell-Jones 1995, 1997), or synanamorphs (second anamorph state) of genus Harposporium (Hodge et  al. 1997; Evans and Whitehead 2005; Li et  al. 2005). Presently, the genus Hirsutella includes more than ninety species attacking a wide range of mites, insects, and nematode hosts. Hemipteran vectors of plant diseases are the major group of economically-­ important insects that harbors Hirsutella infections. The highly polyphagous, xylem-feeding glassy winged sharpshooter, Homalodisca coagulata (Hemiptera: Cicadellidae) are one of the regulated pests. This insect harbours the phytopathogenic bacterium Xylella fastidiosa, the causal agent of Pierce’s disease of grape, almond leaf scorch, plum leafscald, oleander leafscorch, citrus variegated chlorosis and many other plant diseases (Purcell 1989; Hopkins and Purcell 2002). Hirsutella spp. have been also reported to be specific towards mites inhabiting foliar substrates. Species such as H. nodulosa, H. kirchneri, H. necatrix, H. gregis, and H. thompsonii are known to attack eriophyoid mites (McCoy et al. 1988; Maimala 2004). Likewise, Rossi-Zalaf and Alves (2006) reported that H. thompsonii isolated from the rubber

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tree mite Calacarus heveae was highly lethal (>90%) to the false spider mite, Brevipalpus phoenicis. Recent laboratory bioassays revealed that H. kirchneri, isolated from the cereal rust mite, Abacarus hystrix was infectious to prostigmatids, including various spider mites and an astigmatid parasitic mite (Sztejnberg et al. 1997). Moreover, well characterized H. thompsonii strains isolated from eriophyoid mites have been reported to infect and kill the mesostigmatid Varroa destructor, the devastating ectoparasitic mite of the honeybee, Apis mellifera (Muma 1958; Gerson et al. 1979; Shaw et al. 2002).

5.5  Ascomycetes as Mycoinsectides Thanks to the more than 750 species of fungi pathogenic to insects, offering great potential for IPM, about 171 products—based on at least 12 Ascomycete species— have been developed worldwide (De Faria and Wraight 2007; Ramanujam et  al. 2014; Hu et al. 2016). Some species. Pathogenic to insect pests of agricultural crops. Already used in formulated mycoinsecticides include: Beauveria bassiana, B. brogniartii, Lecanicillium spp., M. anisopliae, H. thompsonii and I. fumosorosea (Maina et  al. 2018) (Table  5.1). These Entomopathogenic fungi infect insects of almost all orders with most common targets being Hemiptera, Diptera, Lepidoptera, Coleoptera, Orthoptera and Hymenoptera (Ramanujam et  al. 2014). Some fungi have restricted host ranges viz., Aschersonia aleyrodis infects only whiteflies while others such as B. bassiana and M. anisopliae infect more than 700 species in several

Table 5.1  Fungi pathogenic to insect pests of agricultural crops (Maina et al. 2018) Fungus Beauveria bassiana BB-01

Beauveria bassiana PDRL1187 Beauvaria bassiana Beauvaria bassiana Verticillium lecanii V17, PDRL922 Metarhizium anisopliae L6, M440, PDRL711, PDRL526 Paecilomyces fumosoroseus n32, Verticillium lecanii Peacilomyces lilcinus PDRL812 Hirsutella thompsonii

Target pest Schizaphis graminum, Rhopalosiphum padi, Brevicoryne brassicae and Lipaphis erysimi Mustard Ahpid, Lipaphis erysimi, Aphis craccivora Koch Whiteflies Myzus percsicae Cabbage aphid, mustard Ahpid (Myzus persicae, Lipaphis erysimi) Cabbage aphid, mustard Ahpids, Lipaphis erysimi, Aphis gossypii, Aphis craccivora Koch Mustard aphids, diamondback moth Lipaphis erysimi, Plutella xylostella Myzus persicae, Aphis craccivora Koch Mustard Ahpids, Lipaphis erysimi Aphis craccivora Koch

Crop Laboratory

Canola (Brassica napus L.) Melon Cabbage Cabbage, canola (Brassica napus L.) Cabbage, canola (Brassica napus L Cabbage, canola (Brassica napus L.) Chili Canola (Brassica napus L.) Cowpea

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insect orders. Due to their insecticidal properties, many EF have been formulated into commercial products for controlling insects of economic importance even though temperature and humidity limit their efficacy in field applications. Some of the well known products include, Metarhizium 50, Biogreen and Green Guard, Cryptogram and Bb plus, BIO 1020 and Green Muscle (Bidochka and Small 2005). These mycoinsecticides have been tested and used to control a number of insect pests in glasshouse and field crops worldwide. A list of some commercially available mycoinsecticides, with brand names, target pests and country of production are presented in Table 5.2. Mycoinsecticides, being part of biopesticides, are produced and marketed for pest management strategies while representing only 3% of the global crop protection business. Nonetheless, owing to successes and ecofriendly approach, its growth rate is high, reaching 10% per year (Patrick and Kaskey 2012). Of the total global biopesticide market, mycoinsecticides are second (27%) to Bacillus thuringiensis products, mostly produced in America, Europe and Asia (Kabaluk et al. 2010). In China, at least 30 mycoinsecticides have been registered with B. bassiana being the most popular. Among them up to 14 products have been used for the control of locusts, pine moth and diamond back moth (Hu et al. 2016). Furthurmore, M. anisopliae and Paecilomyces lilacinus with eight seven products, respectively, are registered for application on grubs, corn borer, aphids and whiteflies. Commercially available B. bassiana is the most widely used species. Its products are available for a very wide range of insect pests, including pine caterpillars (Dendrolimus spp.) in China (Feng et al. 1994), banana weevils (Cosmopolites sordidus) in Brazil (Alves et al. 2003) and the European corn borer and greenhouse aphids in the Western world (Shah and Goettel 1999; Copping 2001). Formulations based on Beauveria consist of aerially produced conidia as wettable powders or in emulsifiable oil. Products based on B. brongniartii are available against a wide variety of Coleopteran, Lepidopteran, Homopteran and Dipteran pests in flowers, oil palms, vegetables, and other crops in Colombia (Alves et  al. 2003), against the European cockchafer (Melolontha melolontha L.) and other white grubs in Europe (Copping 2001) and against cerambicid beetles in Japan (Wraight et al. 2001). Clay granules or barley kernels of L. lecanii have been used to produce conidia in solid-state and marketed for control of greenhouse aphids, thrips and whiteflies (Copping 2001; Wraight et  al. 2001) as well as for the control of lepidopteran, homopteran, and dipteran pests of flowers, vegetables, and other crops (Shah and Goettel 1999; Alves et al. 2003). Products with blastospores produced in submerged fermentation or conidia in solid-state fermentation are commercially available. Products based on M. anisopliae are available for a wide range of pests, including red-headed cockchafer in Australia (Shah and Goettel 1999), sugarcane spittlebugs (Mahanarva spp.) in Brazil (Alves et al. 2003), termites in USA (Copping 2001), grasshoppers and locusts in Africa (Lomer et al. 2001) and Australia (Copping 2001). Formulations of P. fumosoroseus are primarily used in greenhouse applications and marketed against whiteflies, thrips, aphids and spider mites in Latin America, Europe, and North America (Copping 2001; Alves et al. 2003). In addition, numerous other products are currently available in many countries such as Entomophthora

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Table 5.2  Some familiar commercial mycoinsecticides with their brand names, target pests and country of production (De Faria and Wraight (2007), Kachhawa (2017), Mishra et al. (2015)) Fungus Beauveria bassiana

Brand name Mycotrol WP Myco-Jaal Conidia Naturalis L Boverol Ostrinil BioGuard rich

Metarhizium flavoviride Metarhizium anisopliae

Boverin Biogreen Bioblast Metaquino Metabiol  Bio-magic  BIO 1020 Metarhizium Andermatt  Fitosan-M

Metarhizium anisopliae (var. acridum) Isaria fumosoroseus

Baeuvaria brongniartii Nomuraea rileyi Hirsutella thompsonii Conidiobolus thromboides Lecanicillium longisporum L. Muscarium B. bassiana; M. anisopliae; I. fumosorosea Aschersonia aleyrodis B. bassiana; M. Anisopliae; N. rileyi I. fumosorosea; Bacillus thuringiensis

Target pest Whiteflies/aphids/Thrips Diamondback moth Coffee berry borer Whiteflies/thrips/aphids/ white grub Leaf beetles Moths Moths, Thrips, weeils, aphids, scarab beetles Whiteflies, Thrips,Mites Scarab larvae Termites Spittle bugs Froghoppers Weeils, scarab beetles, plant hoppers Weevils Scarab beetles

Country USA India Germany USA Czech Republic France India Russia Australia USA Brazil Venezuela India Germany Switzerland Mexico

Betel Numoraea 50 Mycohit Vektor 25SL Vertalec Mycotal Verticillin Tri-sin

Scrab beetles, locusts, grasshoppers Scrab beetles, grubs bugs Locust, grasshoppers Locusts Whitefly Whitely Aphids, Thrips, mealybugs whiteflies Scarab beetle larvae Lepidoptera Acari Aphids/Thrips/whiteflies Aphids Whiteflies/Thrips Whiteflies, Thrips, mites Psyllid

Aseronia

Whiteflies

Micobiol Completo

Mites, beetles, bugs, aphids, flies, locusts

Former USSR Colombia

 DeepGreen Green muscle Green muscle PFR-97 Pae-sin Fumosil

Colombia South Africa China USA Mexico Colombia France Colombia India South Africa Netherlands Netherlands Russia Mexico

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virulenta for control of whiteflies and N. rileyi for management of Lepidoptera in Colombia (Shah and Goettel 1999; Alves et al. 2003), P. lilacinus against plant parasitic nematodes in Australia (Copping 2001) and Hirsutella thompsonii for control of mite species (Kumar and Singh 2001; Copping 2004).

5.6  Genetic Modifications to Enhance Virulence 5.6.1  Augmenting Virulence Owing to their eco-friendly approach and efficiency against agricultural pests, EF have been developed as an alternative to chemical insecticides and a vital component in biocontrol programmes around the globe. However, use has been marred by low virulence and inconsistencies in performance against various pests, which in turn has led to small market shares (Fang et al. 2012) being the prime reason the environmental stresses (Lovett and St. Leger 2015). Traditional approaches to improve pathogen efficacy are based on physiological manipulation or development of better formulation and application strategies (Butt et al. 2016). On the other hand, genetic engineering has given a new lease of life to these entomopathogens (Zhao et  al. 2016a), chiefly Ascomycetes, to significantly improve their virulence and adaptations in adverse conditions prevailing in various habitats. Genetic engineering, in concert with a better understanding of fungal pathogenesis and ecology, has provided a myriad of opportunities to improve the efficacy and thus cost-­effectiveness of mycoinsecticides by recuperating their tolerance to environmental stresses and enhancing their virulence. These methods have resulted in the development of strains displaying: (1) increased cuticular degradation and faster ingress into the host; (2) engineered delimited insect host range; (3) rapid cessation of feeding and paralysis via expression of insecticidal toxins; (4) increased resistance to abiotic stress; and (5) ability to block transmission of human disease-causing agents. Various fungi have been engineered to express insecticidal proteins or peptides or by over expressing the pathogen’s own genes for virulence enhancement. Functional domains from diverse genes of pathogenic fungi and other organisms have also been matched and engineered for producing insecticidal proteins with novel characteristics. Furthermore, fungal tolerance to abiotic stresses, especially UV radiation, has been improved by introducing a photoreactivation system from an Archaean and pigment synthesis pathways from non-entomopathogenic fungi (Zhao et al. 2016b). Genetic engineering to increase virulence has focused on reducing both lethal conidial dosage and time to kill, while improving infection rates. Studying molecular mechanisms of fungal pathogenesis (primarily Metarhizium spp. and B. bassiana), has opened new avenues while allowing many pathogenicity-related genes to be characterized, and used as a resource for enhancing EF performance. For example, notwithstanding insect cuticle that acts as a barrier, entomopathogens produce

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proteases and chitinases to overcome this obstacle and grow on insect pests. Under normal regulation, expression of most of these genes is under tight control (Fang et al. 2009a, b). Constitutively overexpressing the gene encoding the subtilisin-like protease Pr1A increased the virulence of Metarhizium anisopliae towards Manduca sexta the recombinant strain showed a higher performance in survival time towards the insect as compared to the parent wild-type (WT) strain (St. Leger et al. 1996). Similarly, constitutive overproduction of B. bassiana chitinase (CHIT1) also resulted in improvement of virulence by 23% (Fang et al. 2005). Expression of M. anisopliae Pr1A into B. bassiana also increased its killing efficiency (Gongora 2004). These examples illustrate that pathogenicity-related genes from one fungus can be used to improve the virulence of other fungi. Likewise, these fungi need carbohydrate sources for growth and proliferation in the haemolymph. As trehalose is the main sugar in the insect hemolymph, EF must utilize it by secreting the enzyme trehalase (Thompson and Borchardt 2003; Jin et al. 2015). Overexpression of the acid trehalase gene ATM1 accelerated the growth of M. acridum in the hemocoel of locusts, reducing the number of conidia and causing 50% mortality (Peng and Xia 2015). Therefore, modifying the way entomopathogens exploit hosts for nutrition is another reasonable way to improve virulence with many possible mechanisms. The range of endogenous genes suitable for genetic engineering is likely to be in high numbers, including species-specific toxin encoding genes, and gene required to evade the host immune response (Fang and St. Leger 2010; Gao et al. 2011; Lin et al. 2011; Wang and St. Leger 2006, 2007a, b). Prospectively, combining the available genomes from B. bassiana and the several Metarhizium species with robust genetic manipulation technologies, might open doors for characterization of the full range of pathogenicity and host-specificity-related genes (Fang et  al. 2004; Fang et al. 2006; Gao et al. 2011; Hu et al. 2014; Xiao et al. 2012; Xu et al. 2014). Wang et al. (2011) demonstrated that transfer of an esterase gene (Mest1) from the generalist Metarhizium robertsii to the locust specialist M. acridum enabled it to expand its range ad potential to infect caterpillars. Genome-wide analyses of horizontal gene transfer events in Fungi have revealed that Metarhizium species acquired diverse genes from bacteria, archaea or even arthropods, plants and vertebrates (Hu et al. 2014). For instance, sterol transporter (Mr-NPC2a) which allows fungus to compete with host for the growth-limiting sterols present in the haemolymph hace been acquired by horizontal gene transfer (Zhao et  al. 2014). This evolutionary event has been reproduced in B. bassiana which lacks an endogenous (Mr-NPC2a) homolog, thus improving its pathogenicity (Zhao et al. 2014). Inspired by this, virulence of B. bassiana was improved by transgenic expression of several insect molecules (Ortiz-Urquiza et al. 2015). Similarly, expression of the M. sexta diuretic hormone (MSDH) considerably increased the virulence of B. bassiana against various lepidopteran targets (e.g., M. sexta and Galleria mellonella) as well as mosquitoes (Anopheles aegypti) (Fan et al. 2012a). Expression of an inhibitory regulator of toll signalling pathway, a key immune-­ related signal pathway, also increased B. bassiana virulence against two taxonomically different insect species, G. mellonella (wax moth) and adult Myzus persicae

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(green peach aphid) (Yang et al. 2014). Hence, a conserved molecule among taxonomically distant insect species can add to the EF virulence against various target species. In addition, expression of the species-specific pyrokinin b-neuropeptide from fire ants (Solenopsis invicta) increased the virulence of B. bassiana against fire ants. However, it was ineffective against lepidopteran hosts such as G. mellonella and M. sexta (Fan et al. 2012b). To date, insect molecules that have been reported to increase EF virulence are involved in five types of biological processes: sterol hemostasis, food digestion, osmotic balance, immunity, and neural system. However, in theory, any biological process in insects could be a potential target for disruption by mixing and matching different insect molecules, thus making it possible to create more virulent fungal strains with higher specificity against target pests. Bacterial and viral pathogens of insects could benefit by additional toxins by means of novel modes of action. Besides crystal proteins, Bt vegetative insecticidal proteins having insecticidal activities have proven to kill a broad spectrum of lepidopteran insects by lysing the midgut epithelium. Agrobacterium-mediated transformation has progressed to a high state of efficiency in Metarhizium and Beauveria spp. (Xu et al. 2014) while genome-wide functional screening using Agrobacterium insertional mutagenesis in Metarhizium robertsii identified genes involved in sporulation (Fang et al. 2010) and enhancement of virulence (Zhao et al. 2014).

5.6.2  Tolerance to Abiotic Stresses – UV Radiation and Heat The efficacy of mycoinsecticides is often limited by their susceptibility to abiotic stresses viz., UV radiation and temperature (Lovett and St. Leger 2015; Ortiz-­ Urquiza and Keyhani 2015). Screening for increased growth has been effectively used to identify stress tolerance and hypervirulence in UV-induced mutants of M. anisopliae (Zhao et  al. 2016b). However, genetic engineering has been used to increase tolerance to environmental stresses such as UV radiations, producing lines that reliably perform for longer durations in variable conditions and diverse ecosystems. UV radiation is the most challenging environmental factor for solar-exposed mycopesticides that primarily damages DNA through the generation of chemical modifications, most of which are cyclobutane pyrimidine dimers (CPDs) (Sinha and Häder 2002). Expression of photolyases from highly UV-tolerant Halobacterium improved the survivability of M. robertsii and B. bassiana by more than 30-fold, maintaining virulence against Anopheles gambiae even after exposure to several hours of sunlight (McCready and Marcello 2003; Fang and Leger 2012). Besides directly damaging DNA, UV radiation elevates oxidative stress through the production of reactive oxidative species (ROS) (Lesser 1996). Superoxide ­dismutase (SOD) overexpression improves the ability of B. bassiana to detoxify ROS, enhancing UV tolerance (Ying and Feng 2011). Expression of a tyrosinase from Aspergilllus fumigatus activated the production of pigments in B. bassiana,

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thus increasing their conidial tolerance to UV radiation (Shang et  al. 2012). The dihydroxynaphthalene-­melanin (DHN-melanin) synthesis pathway of Alternaria alternata, that increases UV tolerance in many fungi, has been transferred to M. anisopliae, resulting in doubling its tolerance to UV radiation, with increased tolerance to thermal stress (35 °C) and low water activity (Tseng et al. 2011). Temperature extremes can also limit the effectiveness of pest control agents. As with UV radiation, heat stress produces ROS and small heat-shock proteins (HSPs) bestowing thermotolerance in many organisms. Overexpressing HSP25 in M. robertsii also increased thermotolerance and survival under heat stress (Liao et  al. 2014). Exposing M. robertsii to continuous culture with increasing heat stress, produced thermotolerant variants (De Crecy et al. 2009). Consequently, tools of genetic engineering and synthetic biology could offer ample scope to overcome the low tolerance to abiotic stresses and low virulence that constrained the development of biocontrol agents. This approach could generate an unlimited arsenal of anti-insect proteins at pace with the accelerating discoveries of virulence genes and insect vulnerabilities.

5.7  Conclusion and Future Prospects Agricultural pests including arthropods, plant pathogens, weeds and other invertebrates together with ever changing environmental conditions pose serious threat to crop production and dwindling crop yields, worldwide. Entomopathogenic fungi, being naturally available biological control agents, play a key role in population dynamics of pest populations while limiting their economic injury level. Various species of EF have been applied to control pests in different countries and in diverse environmental settings, while some commercial products acting as mycoinsecticides have also been developed and registered. Nonetheless, compared to other technologies and insect management techniques, use of entomopathogens and mycoinsecticides for regulating pest populations is still in its infancy. Need of the hour is to draw attention of Agricultural Scientists, Entomologists, Researchers, Farmers, NGO’s, Goverment agencies and other stakeholders, who are directly or indirectly involved, with a focus on extensive research, innovation, awareness, compatibility, availability and ecofriendly approaches. A range of entomopathogens particularly in the Phylum Ascomycota such as Metarhizium, Lecanicillium, Beauveria and Isaria are grown at large scale while playing a role in IPM strategies. Mycoinsecticides involving M. anisopliae and B. bassiana have been used to control a wide range of pests like mealy bugs, aphids and thrips. Nonetheless, their utilization has not yet reached its zenith in view of the fact that only 3% crop protection globally relies on entomopathogens, with an annual increase around 10% (Patrick and Kaskey 2012). Therefore, paying attention to ecofriendly entomopathogens principally in terms of formulations, strain selectivity, stability in long term use and efficacy under field conditions, is of paramount importance.

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Future prospects of entomopathogenic fungi depend on ecological niches where they dwell for better production and utility. Focus should be given to finding alternative methods for easy use, longer shelf-life and, most importantly, exploitation towards broad spectrum pests. On the contrary, inadequate production of mycotoxins in some species, carcinogenic mycotoxicosis in non-target organisms and slow effectiveness of conidia are some of the hindrances to EF application in IPM. Nonetheless, to overcome this issue, strategies based on synergistic approaches and genetic manipulation have been developed by researchers. Lethal chemical pesticides in low doses have been combined with fungi to kill the pests, synergy applied for example using imidacloprid and B. bassiana against the Colorado beetle (Furlong and Groden 2001) and Spilarctia obliqua caterpillars (Purwar and Sachan 2006). Likewise in genetic manipulation, improvements of pathogens have been attempted through parasexual crossing and protoplast fusion as well as by conventional mutagenesis. Even though great strides have been made in improving methods of production, formulation, and application, many promising innovations and improvements are still possible. Production of “cocktails” especially combining microbials from different classes of entomopathogens or even with chemical or botanical antagonists, is only just beginning. Whatever strategies are contemplated, they should be considered with an intention of not just replacing chemical pesticides by EF, but contributing to the overall development of sustainable agriculture, horticulture, and forestry as well as the preservation of biodiversity in the long run.

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Chapter 6

Thermotolerance of Fungal Conidia Flávia R. S. Paixão, Éverton K. K. Fernandes, and Nicolás Pedrini

Abstract  Conidia of entomopathogenic fungi (EF) are the propagules most frequently used in arthropod biocontrol programs. This anamorphic form is essential for the infection process, including spore germination, penetration, vegetative growth, conidiogenesis and dissemination. Most EF are mesophilic and can develop between 10 and 40 °C, but optimal growth is between 25 and 35 °C. Abiotic factors, especially temperature (high or low) can determine their viability, virulence and success or failure of infection process. Temperature has the highest impact on conidial stress inhibiting metabolic processes, such as decreased morphogenesis during germination, protein denaturation and membrane disorganization. Several studies show that some strains of Beauveria spp., Metarhizium spp., and Isaria spp. exhibit conidial survival even when grown at high temperatures, indicating a relationship between conidial thermotolerance and their geographical isolation origin. Moreover, the high variability in fungal thermotolerance is also dependent of the culture media composition and growth condition. EF that grow at high temperatures do not grow at low temperatures and vice versa. Moreover, when growth conditions are not set at optimal temperatures, EF development is affected and their effectiveness in biological control programs of arthropods is reduced. Thermal stress directly impacts on fungal strains ability to target arthropods and their environmental activity performance. The screening for fungal strains with a higher thermotolerance and the improvement on conidial formulations may aid in optimizing the conditions for biocontrol agent application. Keywords  Temperature · Conidia · Germination · Biological control

F. R. S. Paixão · N. Pedrini (*) Instituto de Investigaciones Bioquímicas de La Plata (INIBIOLP), Consejo Nacional de Investigaciones Científicas y Técnicas (CONICET)-UNLP, Universidad Nacional de La Plata (UNLP), La Plata, Argentina e-mail: [email protected] É. K. K. Fernandes Instituto de Patologia Tropical e Saúde Pública (IPTSP), Universidade Federal de Goiás (UFG), Goiânia, GO, Brazil © Springer Nature Switzerland AG 2019 M. A. Khan, W. Ahmad (eds.), Microbes for Sustainable Insect Pest Management, Sustainability in Plant and Crop Protection, https://doi.org/10.1007/978-3-030-23045-6_6

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6.1  Introduction Entomopathogenic fungi (EF) are responsible for epizootics that often regulate insect pest populations. The genera Metarhizium (Hypocreales: Clavicipitaceae), Beauveria (Hypocreales: Cordycipitaceae), and Isaria (Hypocreales: Cordycipitaceae) are the fungi most frequently used in biological control programs. The asexual spore, named conidium, is an anamorphic and primary form essential in the life cycle of many filamentous fungi (Osherov and May 2001). Mostly, entomopathogenic fungal infection starts with the attachment of conidia to the insect cuticle, and then progresses to conidial germination, penetration, vegetative growth (as hyphal bodies), conidiogenesis and, finally, dissemination (Lacey et al. 2001; Pedrini 2018). Temperature is an abiotic factor that influences all fungal development stages, from primary processes such as biochemical and cellular reactions in conidia to overall infection and fungus-host interaction (Zimmermann 1982; Cabanillas and Jones 2009) (Fig. 6.1). Generally, three thermal conditions are considered for fungal development, i.e., low (below 11 °C), intermediate (11–28 °C) and high temperatures (above 28 °C) (Vidal et al. 1997). Under this classification, EF are considered mesophilic because they develop well between 10 and 40 °C, with optimal growth between 25 and 35 °C (Crisan 1973; Fargues et al. 1997; Vidal et al. 1997; Dimbi et al. 2004). Metarhizium rileyi (= Nomurea rileyi) presented optimum growth at 25 °C (Ignoffo et al. 1976). Similar studies with Metarhizium spp. strains other than M. rileyi showed optimum growth between 11 and 35 °C (Ouedraogo et al. 1997), whereas B. bassiana strains grew over a wider temperature range, from 8 to 35 °C, exhibiting optimal growth at temperatures as low as 20 °C and as high as 30 °C (Fargues et al. 1997).

Inhibiting metabolic processes

Decreasing: Inducing protein denaturation

Stressful conidia Temperature

 Viability  Morphogenesis  Germination

Inducing membrane disorganization Increasing

 Host interaction  Virulence  Sporulation

oxidative stress

Fig. 6.1  Effect of non-optimal temperatures for fungal growth in various processes of conidia life cycle

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Thermotolerance is defined as the ability to withstand relatively hot (or cold) conditions. Viability in thermotolerance above 40 °C was observed in several strains of Metarhizium spp. (Hedgecock et  al. 1995; Li and Feng 2009). However, the growth of some strains of Beauveria spp. was inhibited after 2 h at 45 °C. On the other hand, cold activity was reported for both Beauveria spp. and Metarhizium spp., B. bassiana being able to grow at temperatures as low as 5 °C and some M. anisopliae strains at 8 °C, due to their cold adaptation (McCammon and Rath 1994; Croos and Bidochka 2001; Fernandes et al. 2008; Santos et al. 2011). This physical factor is very important to regulate all development processes, since the beginning of germination up to conidial sporulation (Edelstein et al. 2005; Keyser et al. 2014) (Fig. 6.1). Thus, the screening of fungal strains thermotolerant to high and low temperatures, together with the molecular and genetic characterization, and the investigation on formulations to increase fungal performance are key factors to distinguish strains with high potential to be used in biological control programs of arthropods (Dillon and Charnley 1990; Fernandes et al. 2010; Oliveira et al. 2018).

6.2  Thermal Characteristics and Geographical Origin Interactions between different environmental abiotic factors, spore germination and other physiological traits in fungi were early reported by Gottlieb (1950). EF are ubiquitous in soils worldwide, from the Arctic to the tropics (Zimmermann 2007), thus different thermal behavior in fungi isolated from different geoclimatic origins and/or from diverse hosts might be expected. In fact, Isaria fumosorosea (= Paecilomyces fumosoroseus) strains from tropical or subtropical origin (Cuba, USA, India, Nepal, and Pakistan) demonstrated high tolerance to upper limits (optimal development at 32–35 °C) than European strains (optimal growth at 25–28 °C). Conversely, European strains isolated from temperate areas are able to grow at 8 °C and show to be more tolerant to low temperature than those fungal strains originated from tropical or subtropical regions (Vidal et al. 1997; Fargues and Bon 2004). Some studies reported no thermal variability between B. bassiana strains isolated from temperate or tropical areas from Europe, Africa, Asia and America (Fargues et  al. 1997, Devi et  al. 2005, Rangel et  al. 2005, Borisade and Magan 2014). Although M. anisopliae strains from the tropical region (Africa) were susceptible to germinate at 15  °C (Dimbi et  al. 2004), M. anisopliae isolates from Ontario (Canada) showed large variation in both growth rate and conidial production at temperatures between 8 and 22 °C. This latter study shows that M. anisopliae isolated from forested areas were cold-active, while the isolates from agricultural areas showed an ability for growth at high temperatures and resilience to UV exposure (Bidochka et al. 2001). Regarding cold activity, fungal strains isolated far from the equator presented higher relative germination under cold conditions than strains originated from near the equator. Beauveria bassiana isolated from higher latitudes were cold-active,

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however, there was not a similar correlation for heat (Fernandes et  al. 2008). Thermal characteristics and geographical origins coincide with conditions during natural epizootics between EF and hosts. Persistence in the environment indicates certain adaptation as it was reported for Isaria spp., which showed to be well adapted to semiarid region (Cabanillas and Jones 2009). Finally, the specific pathogen M. acridum, virulent against desert locusts, is able to tolerate temperatures up to 42 °C, and thus to avoid the behavioral fever developed by host in an attempt to stop the development of the fungus, i.e., a successful strategy to avoid infection by thermosensitive EF such as B. bassiana (Elliot et al. 2002).

6.3  Culture Conditions and Conidial Thermotolerance The production of conidia by EF is influenced mostly by culture conditions (viz., temperature, pH, water activity, aeration) and media composition (viz., carbon and nitrogen sources, metal ions) (Hallsworth and Magan 1996; Ibrahim and Jenkinson 2002; Ying and Feng 2006; Rangel et al. 2008). Elevated temperatures reduce spore viability, growth, germination and virulence (Anderson and Smith 1972; McCammon and Rath 1994; Inglis et al. 1996; Rangel et al. 2010; Tumuhaise et al. 2018). It is possible, however, to optimize both conidial production and thermotolerance by efficient culture conditions and/or supply of culture media (Ouedraogo et al. 1997; Cabanillas and Jones 2009; Kim et  al. 2010c). Accordingly, Isaria spp. showed greater tolerance (from 20 to 30  °C) after growing on Sabouraud Dextrose Agar Yeast extract (SDAY) than fungi grown on Sabouraud Maltose Agar (SMA) (Cabanillas and Jones 2009). Submerged cultures of I. fumosorosea grown in Sabouraud Dextrose Broth (SDB) were able to develop well from 20 to 34  °C, showing optimal growth at 28  °C.  In solid fermentations, however, these strains grew optimally at 25 °C (Esther et al. 2013). Although the excess of heat on conidia causes unviability in the derived fungal propagules, it is possible to improve thermotolerance from 26 to 30 °C by increasing the sucrose content in the culture media (McClatchie et al. 1994). Conidia of M. robertsii produced on potato, dextrose, agar and yeast extract (PDAY) medium, containing low concentrations of salicylic acid, demonstrated increased tolerance to heat (Rangel et al. 2012). Millet grain was used as a substrate to produce conidia by B. bassiana and M. anisopliae, potentially enhancing conidial thermotolerance of fungi grown on a massive production system (Kim et al. 2011). Metarhizium acridum grows on agar medium and produces both aerial and microcycle conidia. Tolerance of both propagules was compared at 40–45 °C, showing that microcycle conidia were more heat resistant than normal aerial conidia (Zhang et  al. 2010). Microcycle conidiation is defined as a process in which the germination of spores directly produces the formation of conidia, without the intervention of an intermediate mycelial growth. This microcycle conidiation can be induced by manipulation of environmental conditions, especially culture conditions

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that are stressful to fungi (Hanlin, 1994; Bosch and Yantorno 1999; Zhang et  al. 2010; Jung et al. 2014). A relationship between thermotolerance and hydrophobicity can be traced. Employing a siloxane-mediated conidial collection method based on hydrophobicity, it is possible to classify conidia from B. bassiana and M. anisopliae into two groups with different thermotolerance (Kim et  al. 2010b). Similar results were observed for species with hydrophobic conidia such as B. bassiana, M. brunneum, M. robertsii, M. anisopliae and I. fumosorosea, which were more thermotolerant than species with hydrophilic conidia such as Tolypocladium cylindrosporum, T. inflatum, Simplicillium lanosoniveum, Lecanicillium aphanocladii, Aschersonia placenta and A. aleyrodis (Souza et al. 2014). The sugar content (types and concentrations) used as carbon source in culture media for conidial production may affect both conidial thermotolerance and hydrophobin-like or formic-acid-extractable (FAE) protein content (Ying and Feng 2004), (see Sect. 6.5).

6.4  Effect of Abiotic Factors on Conidial Germination The interaction of abiotic factors (temperature, humidity, light, pH) are important to germination, dispersion, and development of fungal conidia (Glare et  al. 1986; Jaronski 2010; Osherov and May 2001; Oliveira et al. 2018). Water is fundamental to start conidial germination. However, the interaction between water and temperature can reduce the viability and/or limit conidial viability of some EF, e.g., B. bassiana, M. anisopliae, I. farinosa and I. fumosorosea require humidity and optimum temperature conditions to their development (Hallsworth and Magan 1999; Devi et al. 2005; Borisade and Magan 2014). Humidity and temperature are key factors in activation of metabolic pathways allowing the nutrients mobilization required for conidial germination, but are also important during storage periods, longevity and persistence of quiescent conidia (Daoust and Roberts 1983; Dillon and Charnley 1990, Bouamama et  al. 2010). These factors also affect physiological interactions between the host and pathogen (Walstad et  al. 1970; Luz and Fargues 1997; James et  al. 1998). The ultraviolet (UV) radiation (UV-A and UV-B) also affects conidial germination of M. acridum and M. robertsii, limiting entomopathogenic fungal development, however increasing thermotolerance by accumulating trehalose and mannitol (Braga et  al. 2001; Pereira-Junior et al. 2018; Rangel and Roberts 2018).

6.5  Thermal Effects and Metabolic Processes EF have mechanisms to overcome and circumvent thermal effects (Rangel et  al. 2010; Tseng et  al. 2011; Keyser et  al. 2014), triggering signal transduction and metabolic pathways that synthesize the molecules that will ultimately protect the

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fungal cells from damage caused by heat exposure (Farrell and Rose 1967; Ying and Feng 2004; Zhang et al. 2011; Liao et al. 2014; Wang et al. 2017, 2018). Among them, fungal proteins of hydrophobic nature associated to cell walls are often linked with thermal protecting functions. Hydrophobins are small proteins important for fungal growth and development (Wösten and Vocht 2000). Hydrophobin-like or formic-acid-extractable (FAE) proteins were studied in aerial conidia of B. bassiana and I. fumosorosea based on thermotolerance. FAE proteins provide hydrophobicity to conidia, exhibiting different composition between B. bassiana and I. fumosorosea. For both fungi, conidial viabilities decreased after exposure to heat stress (48 °C for up to 150 min), perhaps as a result of different conidial structure related to FAE proteins (Ying and Feng 2004). Zhang et al. (2011) characterized structurally the cell wall carbohydrates in B. bassiana, and demonstrated that targeted gene knockouts lacking β-1,3-glucanosyltransferase destabilize the cell wall and decreased germination after 1 to 4 h of heat shock at temperatures >40 °C. Trehalose is a disaccharide that accumulates in fungi during stress situations, such as adverse growth conditions, heat, and hyperoxidative shock. Thus, along with other polyols these molecules are known as stress metabolites (Van Laere 1989; Fillinger et al. 2001; Liu et al. 2009). Polyols accumulation is associated with thermotolerance by helping in the stabilization of structure (and function) of proteins and enzymes at high temperatures (Kim and Lee 1993). Accumulation of glycerol, erythritol, arabitol, mannitol, and trehalose in conidia of M. anisopliae, B. bassiana, and I. farinosa under different culture age (up to 120 days), temperature (5–35 °C) and pH (2.9–11.1) were reported by Hallsworth and Magan (1996). Also, high accumulation of both trehalose and mannitol were observed in abiotic stressed conidia of M. acridum, suggesting they are part of a mechanism that the fungus uses to attain its high tolerance to UV-B radiation and heat (Rangel and Roberts 2018). The heat stress also triggers the production of toxic reactive oxygen species (ROS), favoring oxidative stress in fungal propagules (Zhang and Feng 2018). Catalase is an antioxidant enzyme characterized in B. bassiana (Pedrini et al. 2006) that showed to be an important regulator of conidial thermotolerance (Wang et al. 2013). The relationship between oxidative stress and elevated culture temperature also was reported for Aspergillus niger (Bai et al. 2003). The heat shock proteins (HSPs) are also associated with tolerance to heat: overexpressing the gene encoding for HSP25  in M. robertsii increased fungal growth under heat stress either in nutrient-­rich medium or on insect wings, and also enhanced the tolerance of heat shock-treated conidia to osmotic stress (Liao et al. 2014).

6.6  Conidial Formulation and Thermotolerance Formulations preserve the viability of conidia exposed to environmental stresses, improving the efficiency of fungal propagules in microbial control (Faria and Wraight 2007). Conidial formulations based on oil or oil-in-water emulsions are investigated because the combination of conidia with oils improved their

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performance against heat stress (Malsam et al. 2002; Mendonça et al. 2007; Barreto et al. 2016; Paixao et al. 2017; Oliveira et al. 2018). Oil-based formulations of M. anisopliae s.s. were used to improve both germination and appressorium production in conidia used for tick control (Barreto et  al. 2016; Alves et al. 2017), and also to protect conidia against the effect of solar radiation (Alves et al. 1998) and high temperatures (McClatchie et al. 1994). Conidia of M. anisopliae s.l. and M. robertsii formulated on either vegetable or mineral oils were more tolerant to heat stress than those either unformulated or formulated on carboxymethylcellulose gel (Paixao et  al. 2017). M. anisopliae and B. bassiana viability also increases when fungi are formulated in emulsifiable oil (Oliveira et al. 2018), and vegetable oil improved both performance and thermotolerance of B. bassiana (Kim et al. 2010a). Thus, formulation is considering a very important tool to manage heat stress on conidia.

6.7  Conclusion Temperature is a key factor that limits survival of entomopathogenic fungal conidia used in biological control programs. As detailed in this chapter, most of the investigations in this area have concentrated in: (i) fungal screening for thermotolerance, based on geographical origin of the strains (McCammon and Rath 1994; Morley-­ Davies et al. 1996; Fargues et al. 1997; De Croos and Bidochka 1999; Devi et al. 2005), (ii) test of tolerance to low or high temperature (Fernandes et al. 2008; Paixão et al. 2017), (iii) appropriate culture medium for conidial production (Hallsworth and Magan 1999; Cabanillas and Jones 2009; Esther et al. 2013), (iv) formulations to increase conidia thermotolerance and protection (Hedgecock et al. 1995; Barreto et  al. 2016; Paixão et at. 2017), and (v) biological/molecular characteristics and mechanisms that mediate stress tolerance (Liu et al. 2009; Fernandes et al. 2010; Rangel et al. 2018). On the basis of the literature available, we can conclude that EF are promising tools against many arthropods (Zimmermann 2007; Faria and Wraight 2007). However, additional research is still needed mostly in both screening of thermotolerant strains and formulation types of fungal propagules, to circumvent the negative effects of abiotic factors that potentially limits their efficacy, thus improving the use of EF in biological control programs.

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Chapter 7

Oxidative Stress in Entomopathogenic Fungi and Its Potential Role on Mycoinsecticide Enhancement Carla Huarte-Bonnet, M. Constanza Mannino, and Nicolás Pedrini

Abstract  Entomopathogenic fungi (EF) are used worldwide as environmentally friendly mycoinsecticides. A successful invasion process depends on the fungal ability to cope with several stress factors, such as osmotic stress, temperature, UV radiation, and oxidative stress. Reactive oxygen species (ROS) can appear due to either previous environmental stresses or endogenous metabolic changes. Moreover, ROS may be either part of the host defense against fungi or the fungus itself can release ROS in the hemolymph to overcome insect defenses. Regardless of its source, fungi must mitigate ROS damage in their cells. Antioxidant response in fungi involves the action of enzymes as well as non-enzymatic compounds. Oxidative stress and antioxidant responses are known to have several direct and/or indirect consequences in fungal adaptation. Nutritive stress produced by non-­ preferred carbohydrate sources in conidia production can increase ROS scavengers consequently enhancing UV tolerance. Additionally, growth in long chain cuticular hydrocarbons triggers ROS production and antioxidant gene induction, leading to more virulent conidia. Also, ROS can act as signaling molecules for cell differentiation into new propagules such as microsclerotia and mycelial pellets that tolerate desiccation and produce new infective conidia in the field. In this chapter we will summarize ROS sources and antioxidant scavengers during conidial production and fungal invasion into their hosts, and the beneficial consequences for stress tolerance, virulence and cell differentiation that can arise from these initial drawbacks. Keywords  Beauveria bassiana · Metarhizium anisopliae · ROS · Stress tolerance · Cell differentiation

C. Huarte-Bonnet · M. C. Mannino · N. Pedrini (*) Instituto de Investigaciones Bioquímicas de La Plata (INIBIOLP), Consejo Nacional de Investigaciones Científicas y Técnicas (CCT La Plata CONICET-UNLP), Universidad Nacional de La Plata (UNLP), La Plata, Argentina e-mail: [email protected] © Springer Nature Switzerland AG 2019 M. A. Khan, W. Ahmad (eds.), Microbes for Sustainable Insect Pest Management, Sustainability in Plant and Crop Protection, https://doi.org/10.1007/978-3-030-23045-6_7

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7.1  Introduction Entomopathogenic fungi (EF) are currently used as biocontrol agents of a large number of pests in agricultural systems, including aphids, beetles, grasshoppers, moths, butterflies, termites, weevils, whiteflies (Lacey et al. 2015; Deshayes et al. 2017), as well as for arthropods of medical and veterinary importance, such as mosquitoes, kissing bugs, bed bugs, ticks and tsetse flies (Blanford 2005; Scholte et al. 2006; Pedrini et  al. 2009; Fang and St. Leger 2012; Maniania and Ekesi 2013; Barbarin et  al. 2017). Unlike any other insect pathogen, EF do not need to be ingested to invade their hosts since they can initiate the infection cycle by penetrating the insect cuticle. Infective fungal cells meet the host surface and subsequently start a series of physical, molecular and physiological changes allowing adhesion to the epicuticle, penetration through the cuticle into the insect haemocoel, colonization and sporulation throughout the cadaver (Pedrini 2018). In all those stages, fungi are exposed to both external and internal sources of stress that can increase reactive oxygen species (ROS), e.g. singlet oxygen, peroxide ion, hydrogen peroxide, and superoxide anion, hydroxyl and peroxide radicals. In these scenario, a hyperoxidant state in which ROS exceed the antioxidant capability of the cell can transiently appear, endangering the integrity of cellular components such as DNA, proteins and lipids (Georgiou et al. 2006; Gessler et al. 2007; Hernandez et al. 2010; Zhang and Feng 2018). Fungal antioxidant responses include antioxidant enzymes and other antioxidant compounds, which have been reviewed by Gessler et al. (2007). In this chapter we will summarize some ROS sources during fungal infection and in vitro fungal production and its potential role in further stress resistance, cell differentiation and virulence enhancement.

7.2  ROS During Fungal Infection During host colonization, fungi must mitigate the effect of several stresses to rapidly and efficiently kill the target insect. Before penetration into the host, fungal conidia are exposed to solar radiation and temperature fluctuations. Ultraviolet radiation causes several changes in the cell. UV-B radiation mainly provokes DNA modifications such as pyrimidin dimers and 6–4 photoproducts, whereas UV-A induces the formation of ROS in the cell (Griffiths et al. 1998). Temperature fluctuations can result in osmotic imbalance as well as free radicals generation (Ortiz-­ Urquiza and Keyhani 2015). Penetrating inside the host is also challenging. Fungi need to trespass the hydrophobic epicuticle and the protein-chitin procuticle to reach the hemolymph. The epicuticle is a thin layer mainly composed of long chain hydrocarbons (Pedrini et al. 2007). Alkane degradation by fungi involves ROS formation in peroxisomes through β-oxidation, as well as antioxidant stress response induction in Beauveria bassiana and other EF (Huarte-Bonnet et al. 2015). Once inside the hemocoel, fungi must struggle with the host immune response as well as

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osmotic changes due to hemolymph solutes. Cellular and humoral insect immune responses includes phagocytosis, pathogen encapsulation, induction of a variety of antimicrobial peptides, lectins, and the prophenoloxidase cascade. The latter induces the release of insect ROS into the cavity and melanin production, toxic to both insect and fungi. Insect can also subject themselves to a behavioral fever by selecting environments warmer than their normal preference, to delay or inhibit fungal pathogenicity (Lovett and St. Leger 2014; Ortiz-Urquiza and Keyhani 2015; Pedrini 2018). Subsequently, invading fungi use hemolymph substrates to produce hyphal bodies until the nutrient availability is exhausted. At this stage, they return to starvation, under osmotic and oxidative stress. Finally, the pathogenic fungi transit through the host in the opposite direction and proliferate on the cadaver surface, being once again vulnerable to environmental stress (Lovett and St. Leger 2014; Rangel et al. 2015). Several studies indicate that antioxidant stress enzymes, as well as the protectant compounds produced are crucial to the fungal fitness, virulence and tolerance to stress. The catalase (cat) gene family of B. bassiana consists of catA (spore-­ specific), catB (secreted), catP (peroxisomal), catC (cytoplasmic) and catD (secreted peroxidase/catalase). Single deletion knockouts of these genes shed light into their contribution to stress tolerance and virulence: BbcatP, BbcatA and BbcatD are influential virulence factors against insect pest whereas BbcatA and BbcatD are equally crucial in regulating conidial UV-B resistance, followed by BbcatB. Moreover, BbcatA also acts as the most important regulator of conidial thermotolerance and BbcatP is crucial to oxidative stress management (Wang et al. 2013a). Superoxide dismutase gene (sod) single knockouts and knockdowns were also constructed for five sod in B. bassiana (Xie et al. 2012; Li et al. 2015). Whereas all transgenic strains were more sensitive to UV stress and less virulent, only Bbsod1, Bbsod2, Bbsod3 and Bbsod5 knockout strains delayed germination, but none of them showed differences in thermotolerance. Interestingly, Bbsod5 is involved in oxidative stress response, more specifically in the fungal response to the host immune defense after penetrating the cuticle. Additionally, six thioredoxins (trx), four glutaredoxins (grx) and one glutathione reductase (glr) were disrupted in B. bassiana. The six ∆Bbtrx mutants, ∆BbGrx3 and ∆BbGlr displayed one or more phenotypic changes associated with the fungal biocontrol potential, i.e., conidiation and germination, thermotolerance, UV-B resistance and virulence of their conidia (Zhang et al. 2015, 2016). Melanins present in Metarhizium spp. strains are involved in heat stress and UV tolerance. In M. anisopliae, the laccase MLAC1 is involved in melanin biosynthesis and contributes to conidial pigmentation, tolerance to abiotic stresses and pathogenicity (Fang et al. 2010). Moreover, overexpression of antioxidant stress enzymes improved EF biocontrol potential: Bbsod2 overexpression led to a significant increase in superoxide dismutase activity, accompanied by increased oxidative and UV stress tolerance, as well as an increased virulence against larvae of Spodoptera litura (Xie et al. 2010). A transgenic strain of M. anisopliae overexpressing Macat1 and a B. bassiana transgenic strain overexpressing BbcatE7 showed increased resistance to hydrogen peroxide, decreased germination times and improved virulence

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against Plutella xylostella and S. exigua larvaes, respectively (Hernandez et  al. 2010; Chantasingh et al. 2013).

7.3  ROS During Conidial Production EF are produced and commercialized worldwide as mycoinsecticides. Production and formulation processes are crucial to obtain fungal propagules that are more virulent and more tolerant to stress (Faria and Wraight 2007; Jackson et al. 2010). In this regard, several studies have changed nutritional conditions to enhance germination rates, tolerance to abiotic and oxidative stress, and virulence. Conidia of M. robertsii produced in media without a carbon source, known as minimal media (MM), were found to be more tolerant to UV-B radiation. Additionally, fungi grown in MM supplemented with non-preferred carbon sources such as arabinose, fructose, galactose, inositol, and lactose, also increased UV-B tolerance, and germinated faster with superior cuticle adherence (Rangel et al. 2006, 2008). Fungal growth in absence of glucose is a nutritional stressor that triggers different cell responses, including apoptosis, macroautophagy, production of secondary metabolites and antioxidant stress responses alteration (Rúa et al. 2014). During nutritional stress (and other stresses) trehalose accumulation in the cell was shown to be increased (Rangel et al. 2015). Trehalose functions as a free radical scavenger and protects cellular constituents from oxidative stress. Therefore, trehalose can protect fungal cells from further oxidative stress, and its levels can be considered as stress indicators. Thus, trehalose and other stress responses can provoke “cross-­ protection” in the cell: some stress responses can be beneficial to overcome further stress, since response pathways are common to different stress sources (Hallsworth 2018). Since EF penetrate the wax-rich epicuticle, supplementation with insect-like hydrocarbons has also been tested. Napolitano and Juárez (1997) first described the ability of B. bassiana and M. anisopliae to degrade hydrocarbons. Isolates of B. bassiana grown on MM supplemented with hydrocarbons were more virulent than glucose-grown fungi (Crespo et al. 2002; Pedrini et al. 2009). Moreover, alkane-­ grown cells showed peroxisome proliferation (Crespo et al. 2000), and an oxidative stress scenario was triggered (Huarte-Bonnet et al. 2015). In Isaria fumosorosea, conidia produced with hydrocarbons showed higher antioxidazing enzymes activity, lower germination times, increased resistance to exogenous hydrogen peroxide and higher virulence against S. exigua (Ali et al. 2013). The effect of different oxygen concentrations during production of I. fumorosea conidia was described by Miranda-Hernández et al. (2014). Oxygen pulses during fungal growth provoked greater germination rates and resistance to thermal and osmotic stresses, as well as higher virulence against Galleria mellonera larvae. This was presumably due to the induction of an antioxidative stress response, as catalase activity in these conidia was increased. Furthermore, oxidative stress can also be generated by exposing conidia to UV-A radiation, hydrogen peroxide

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s­ upplementation in the culture media, as well as other oxidants such as menadione. Rangel et al. (2011) induced cross-protection in M. robertsii conidia to other stress conditions after exposure to visible light, including UV-A wavelengths. However, not all oxidative stress generators are capable to induce tolerance against other types of stress (Rangel et al. 2015).

7.4  ROS and Cell Differentiation A hyperoxidant state in the cell is a transient condition that needs to be battled to prevent harmful damage that could lead to the cell death. Cell differentiation in eukaryotic microorganisms has been proposed as a process triggered by hyperoxidant states, allowing cell isolation from molecular oxygen (Hansberg and Aguirre 1990). However, isolation from O2 also isolates the cell from water and other nutrients, forcing the exploitation of endogenous sources to survive and maintain such a condition (Georgiou et al. 2006). In some filamentous fungi, cell differentiation can induce sclerotial biogenesis. Sclerotia are macroscopic, pigmented aggregates that can survive long periods in adverse environmental conditions. They have ben previously reposted in several plant pathogenic fungi. In particular, oxidative stress has shown to be involved in cell differentiation into sclerotia in Sclerotium rolfsii, Sclerotinia minor, Sclerotinia sclerotiorum and Rhizoctonia solani (Sideri and Georgiou 2000; Georgiou et  al. 2006; Papapostolou and Georgiou 2010a,b). EF, however, produce smaller sclerotia known as microsclerotia (MS). Up to date MS formation was described in Metarhizium anisopliae, M. acridum, M. robertsii, M. brunneum, M. rileyi, Lecanicillium lecanii, Beauveria bassiana, B. brogniartii and B. pseudobassiana (Jaronski and Jackson 2008; Jackson and Jaronski 2012; Wang et  al. 2013b; Mascarin et al. 2014; Song et al. 2014; Villamizar et al. 2018). These propagules have been explored as potential propagules for biocontrol strategies since they can tolerate desiccation and germinate without any exogenous carbon source, producing new conidia in the field. These characteristics make MS promising propagules since they can be formulated as solid granules, stored for long periods of time for later use and produce great quantities of conidia once they are applied and spread (Jaronski and Jackson 2008). To date, oxidative stress involvement in MS differentiation was only studied in M. rileyi. First, upregulation of antioxidant enzymes during MS formation was found by transcriptome analysis (Song et al. 2013). During culture optimization, Fe2+ was found to promote MS formation (Song et al. 2014). Iron ions are known to increase ROS concentration via Fenton reaction (Georgiou et al. 2006; Song et al. 2013). Additionally, whereas exogenous menadione and peroxide hydrogen in the culture media induced MS biogenesis, ascorbic acid, a ROS scavenger, inhibited MS formation (Liu et al. 2014). It was recently described that alkane-cultured B. bassiana differentiated into mycelial pellets, compact spherical aggregates previously described only in non-­ entomopathogenic filamentous fungi (Huarte-Bonnet et  al. 2018). These hyphal

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aggregates tolerated desiccation, sporulated and produced viable conidia after rehydration. They were found to be pathogenic against larvae of Tenebrio molitor and Tribolium castaneum, making them also a promising propagule for mycoinsecticide production. Electron microscopy imaging showed that cells forming the pellets had high peroxidase activity, with peroxisomal proliferation. Several antioxidant response genes encoding for catalases and superoxide dismutases were induced in these pellets, compared to mycelia from glucose-grown fungi. These results suggested that oxidative stress previously found in alkane-grown conidia (Huarte-­ Bonnet et al. 2015) could have triggered B. bassiana differentiation into mycelial pellets. Although not yet tested, as conidia produced by hydrocarbon-grown fungi were more virulent than those produced with glucose, it is possible that conidia produced after pellet rehydration would result more virulent as well.

7.5  Conclusion Microbes are constantly under stress. Stress biology involves all microbial organizational levels. Therefore, the study of stress is crucial to better understand and successfully implement microorganism-based technologies (Hallsworth 2018). At this regard, EF do not represent an exception. In particular, oxidative stress can appear either as a direct or indirect consequence of ROS formation inside the cell, due to other stresses, or ROS can be produced as metabolic subproducts. If ROS accumulation is higher than the amount a fungal cell can handle, induction of antioxidant enzymes and production of antioxidant compounds are activated to defend the cell integrity. In some other cases, supposedly when antioxidant stress responses are not sufficient, cell differentiation can be triggered to isolate fungal cells from subsequent stress. If fungi can finally overcome stress, molecular and phenotypic changes transformed formerly stressed cells into better pathogens, improving their germination rates, tolerance to more harmful stress levels or to other stresses sources, enhancing virulence against insects, or even forming new propagules more tolerant to environmental stress and more suitable to be formulated and spread in the field. In conclusion, if sublethal, stress apears as an useful improvement stimulant.

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Chapter 8

Effects of Cytotoxic Factors Produced by Entomopathogenic Bacteria on Insect Haemocytes Carlos Ribeiro and Amélia Vaz

Abstract  Members of genera Xenorhabdus and Photorhabdus are entomopathogenic bacteria (EB) that associate with nematode hosts from the genera Steinernema spp. and Heterorhabditis spp., respectively, providing them with entomopathogenic services through exceptional physiological and metabolic mechanisms. Both symbiotic pairs infect and kill insects, with the bacteria contributing to host pathogenesis and death, supplying nutrition for the nematode from available insect-derived nutrients. In turn, the nematode provides the bacteria with protection from soil predators, access to nutrients, and a dispersal mechanism. These EB are able to produce and secrete a broad range of cytotoxic metabolites, aimed not only at circumventing and disabling the insect’s complex web of humoral and cellular defences, but also at killing it quickly, eliminating all potential competitors for the newly acquired food source. An overview of most studied compounds and of their impact on the immune response effectors is provided, with emphasis on the cellular components of the insect immune system. The different cell types found in the haemolymph of lepidopterans are described and characterised, in comparison to the cell types recognized in Drosophila. Keywords  Xenorhabdus · Photorhabdus · Immunity · Haemocytes · Cytotoxins · Cytolysins · Haemolysins

C. Ribeiro (*) · A. Vaz Departamento de Biologia, Faculdade de Ciências e Tecnologia, Universidade dos Açores, Ponta Delgada, Portugal e-mail: [email protected] © Springer Nature Switzerland AG 2019 M. A. Khan, W. Ahmad (eds.), Microbes for Sustainable Insect Pest Management, Sustainability in Plant and Crop Protection, https://doi.org/10.1007/978-3-030-23045-6_8

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8.1  Introduction Insects are ubiquitous organisms, and a small but significant part of them are considered pests, despite the subjectivity of that classification, because what qualifies as “pests” varies greatly from case to case. In general, this designation is reserved for organisms that cause potential damage in several domains, namely to animal and/or human health, to natural habitats and ecosystems, and agricultural crops, wherein they cause a series of well-documented socio-economic losses. The current review deals, primarily, with the effects of entomopathogenic bacteria (EB) on the cellular immune responses of insect pests, when challenged with two entomopathogenic nematodes (EPNs) living in soil, Steinernema carpocapsae (Nematoda: Steinernematidae) and Heterorhabditis bacteriophora (Nematoda: Heterorhabditidae), strictly associated with a monospecific natural symbiont from the bacterial genera Xenorhabdus (Akhurst and Boemare 1988; Akhurst 1993; Nishimura et al. 1994) and Photorhabdus (Boemare et al. 1992; Akhurst et al. 1996; Szállás et al. 1997; Fisher-Le-Saux et al. 1999), respectively. Both genera comprise Gram-negative motile rods from the Enterobacteriaceae family of γ-Proteobacteria. These bacteria are carried in the nematode gut, where they find shelter from soil stressors and protection from antagonists, namely telluric bacterial consortia and bacteria present in the insect gut. In fact, neither has ever been isolated from soil samples in the absence of their specific nematode host. These symbiotic pairs are pathogenic, and capable of parasitizing and killing the larval stages of their hosts, namely Lepidoptera, Diptera, Coleoptera, Orthoptera, Hymenoptera and Isoptera (Laumond et al. 1979; Poinar 1979; Akhurst and Boemare 1990; Boemare 2002; Khan et al. 2016). EPNs from genera Steinernema and Heterorhabditis are among the most widely known biological control agents, being often used in classical, conservative and augmentative biological control approaches (Bedding 2006). In this symbiosis, the eukaryote partner provides not only safe shelter, but also a mean of entering the insect body cavities where the bacteria are released, first in the insect gut and then in the haemocoel. Upon release, the bacteria start to produce and secrete (1) a large spectrum of antibiotics and/or antimicrobial metabolites, in order to begin dealing with the bacterial consortia in the insect gut, annihilating all the competitors, (2) several toxins, secondary metabolites and enzymes, that help in the destabilization of the intestinal epithelium and, once in the haemocoel, circumvent and inhibit the humoral and cellular defence reactions of the insect, ultimately killing it, and (3) bacterial enzymes, initiating the biotransformation of the macromolecular contents of the insect cadaver, preventing its putrefaction and making available the nutrients necessary for both nematodes and bacteria to develop and grow, completing their life cycle (Forst and Nealson 1996; Forst and Clarke 2002). This interaction between the nematobacterial complex and the insect is somewhere between host-parasite and predator-prey. The penetration of the insect, coupled with the rapid multiplication of the bacteria and nematodes once inside the larvae, are close to a parasitic behaviour, while the speed with which the complex kills the victim evokes the actions of a

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predator (Ribeiro 2002). In the end, it must be pointed out that the bacterial symbiont inhabits and influences the life cycles of two host animals, helping one to feed and reproduce optimally in a controlled environment (the nematode), while killing the other (the insect) (Herbert and Goodrich-Blair 2007). As far as the host insects are concerned, they defend themselves against the nematodes and their symbiotic bacteria with cellular and humoral innate immune responses. The cellular responses against bacteria include phagocytosis and/or nodule formation, whereas against EPNs these reactions are melanisation and encapsulation. The major humoral response factors in lepidopteran larvae include the synthesis of antimicrobial peptides (AMPs, e.g. cecropins, defensins and attacins), lysozyme and phospholipase A2, and the prophenoloxidase/melanisation and coagulation cascades.

8.2  Life Cycles of the Nematode-Bacteria Symbionts The life cycles of the two nematobacterial mutualistic symbionts, Steinernema: Xenorhabdus and Heterorhabditis: Photorhabdus are generally described as one, despite the fact that there are four living entities involved, that the nematodes have different life cycles and that the molecular components of the regulatory networks controlling pathogenicity and mutualism in Photorhabdus and Xenorhabdus are very different (Goodrich-Blair and Clarke 2007). Indeed, the life cycles of these entities appear to be quite similar at the macroscopic level, with only slight differences, namely for the foraging strategies and reproductive cycles (Poinar 1975; Johnigk and Ehlers 1999a, 1999b). However, at the cellular and molecular levels, there are significant differences that are important to consider and study, both related to the pecularities of the specific interactions of the symbiotic pairs, during mutualism (Herbert and Goodrich-Blair 2007). They include symbionts evolution, leading to two convergent life styles originated from two divergent genomes (Chaston et al. 2011), and the interactions with the insect preys during parasitism (Chapuis et al. 2009; Lee and Stock 2010; Stilwell et al. 2018). Furthermore, coevolution with the symbiotic partners has led to a high level of mutualism specialization, which effectively prevents horizontal transmission of bacterial symbionts between different nematode species (Sicard et al. 2004a). The free-living infective juvenile (IJ) nematode stages are soil-dwelling and not-­ feeding, and protected by a double cuticle. They rely upon their energy reserves for survival up to several months, while seeking their insect preys, using different foraging strategies that vary from ambush to cruise (Lortkipanidze et al. 2016). At this point, the IJs harbour their bacterial symbiont either disseminated in their intestinal lumen, in the case of Heterorhabditis: Photorhabdus (Boemare 2002) or, in the case of Steinernema: Xenorhabdus, specifically in a vesicle (Bird and Akhurst 1983). Once a suitable host is encountered, the nematode enters the insect via its natural openings, namely the mouth and/or anus and, during this process, sheds its outer cuticle (Poinar and Himsworth 1967; Sicard et  al. 2004b). Nematodes may also

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enter the body cavity via the tracheal system, provided that the aeropyles of the spiracles are wider than their body diameter. Direct penetration of the insect cuticle by Steinernema has never been described. However, Heterorhabditis is capable of puncturing the larval cuticle, entering the host’s body this way (Bedding and Molyneux 1982). Once in the insect gut, and as they begin to pass through the ectoperitrophic matrix and the intestinal epithelium on their way to the haemocoel, the nematodes begin releasing the bacteria. During infestation of the cotton leafworm Spodoptera littoralis larvae with EPNs harbouring GFP-labelled Xenorhabdus (Sicard et  al. 2004b), the authors described in detail all the sequential steps of the parasitic path. In particular, the visualization of Xenorhabdus in IJ nematodes provided new insights on the stimuli triggering the bacterial release in the insect midgut. Moreover, in the early stages of colonization, as the bacteria reach the haemocoel, living bacterial cells were found in two places: in the connective tissues of the hematopoietic organ, and in the haemolymph plasma. These results gave new insight into the behaviour of Xenorhabdus during the interaction with host tissues. It seems that these bacteria need to find a connective tissue to ensure stability for growing successfully in the early steps of haemocoel colonization. It would be interesting to investigate which putative molecular components present in the cellular matrix of the basal membranes (e. g. laminin, collagen, fibronectin or other proteinaceous and/or glycoproteinaceous materials) are the most suitable for bacteria to interact with. After the pair reaches the insect haemolymph and the nematodes release their symbionts, the bacteria begin the task of overcoming the insect immune response and killing it, by toxaemia and/or septicaemia, through the production and secretion of an array of toxic molecules (Akhurst and Boemare 1990; Forst et al. 1997). After the onset of generalized toxaemia and/or septicaemia, and the concomitant monoaxenic colonization by the nematobacterial complex, the insect cadaver undergoes substantial changes, starting with the digestion and biotransformation of the entire molecular and tissue content, providing a rich nutritional environment for nematodes and bacteria to feed and reproduce. Ultimately, the massive amount of bacterial biomass produced serves also as a source of nutrition for both organisms. In fact, the bacteria grow exponentially in a way that makes it impossible to fit not only in the cadaver, but also in the intestine of the next generation of nematodes and so, it is assumed that a significant part of their biomass will also serve as nutrient content, after the stationary and declining growth stages are reached. In fact, Ribeiro (1994, 2002) reported the presence of live Xenorhabdus rods alongside bacterial spheroplasts and debris from dead bacteria in microscopic observations of greater wax moth, Galleria mellonella, cadaver contents. Spheroplasts appear in the last third of exponential growth in artificial medium (Akhurst and Boemare 2015), as they do inside the insect cadaver. In this nutrient-rich environment, the nematodes begin several rounds of reproduction cycles, sexually in Steinernema and hermaphroditically in Heterorhabditis (Johnigk and Ehlers 1999a, 1999b). Concomitantly, with their growing numbers being limited by nutrient scarceness, they successively stop their reproductive cycle

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in the L3 instar larvae. At this stage, the nematode progeny receives uncharacterized environmental cues in response to a signal, possibly nutrient deprivation or space limitation (Popiel et al. 1989; Herbert and Goodrich-Blair 2007) which stimulate the development of a new generation of IJs, that are then colonized by their symbiont partner, before gaining the protection of a new double cuticle and emerging by the thousands from the insect cadaver. The success of the symbiotic pair entails a high degree of specialization, namely in the specificity of the re-association and the high efficiency in vertical transmission, that are potentially under a strong selective pressure. In fact, in Steinernema: Xenorhabdus, it has been found that an association with bacteria other than their natural symbiont is clearly detrimental for the nematode’s fitness and only symbionts that were cognate to the nematode were associated and transmitted (Sicard et al. 2004a). Therefore, an effectively axenic environment in the insect cadaver is paramount in order to exclude any bacteria other than the natural symbiont, since this is the only suitable biological reservoir for the entomopathogenic symbionts to complete their life cycles. Looking at the bacterial symbiont life cycle, it can be divided into three separate stages: infection, after the nematode enters the insect and the bacteria are expelled from its gut into the insect gut and/or haemocoel; reproduction, after the insect is killed and its biomass is converted by the bacteria into a nutrient-rich soup enabling the growth and development of both organisms; and re-association, as the nematodes stop reproducing, and the bacteria regain shelter de novo in the nematode gut before the pair emerges. To successfully complete these stages, the bacteria must accomplish five distinct tasks: (i) within the nematode gut, they must keep a stabilized sessile condition, in order to maintain a successful symbiosis, (ii) once in the haemocoel, they must overcome the insect’s immune response and kill it, (iii) establish an axenic environment, (iv) produce nutrients from the insect cadaver to facilitate the development of the nematode and their own growth, and (v) colonize the next generation of nematode IJs. In the specific case of X. nematophila, three regulons (Lrp, LrhA and CpxR) are thought to be responsible for the modulation of activities necessary for the fitness of both symbionts in response to the shifting environment throughout the bacteria’s life cycle. Their activation possibly responds to nutritional cues, expressing virulence factors, degradative enzymes or nematode colonization factors accordingly (Richards and Goodrich-Blair 2009). LrhA is a LysR-type regulator heavily involved in the modulation of virulence, flagellar motility and lipase activity through positive regulation of FlhDC. This is a transcription factor controlling flagellar synthesis (Givaudan and Lanois 2000), transcription and secretion through the flagellar apparatus of the XlpA lipase, supporting Steinernema reproduction (Park and Forst 2006; Richards and Goodrich-­ Blair 2010), and the expression of several virulence effectors such as the cell-associated haemolysin, encoded by xhlA, and the secreted haemolysin, encoded by xaxAB (Cowles and Goodrich-Blair 2005; Vigneux et al. 2007). Lrp, the leucine-responsive regulatory protein encoded by lpr, is involved in all stages of the X. nematophila life cycle. It regulates both pathogenic and mutualistic

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host interactions, and is thought to be responsible for coordinating the so called “feast or famine” adaptation to fluctuations in nutrient availability, found ubiquitously in both bacteria and archaea (Yokoyama et al. 2006; Cowles et al. 2007). Lrp positively regulates motility, antibiotic production, protease activity and the haemolysins encoded by xhlA and xaxAB, as well as the production of LrhA and, by extension, of XlpA (Richards and Goodrich-Blair 2009). It negatively regulates the expression of the nematode colonization genes nil, working synergistically with NilR in their repression (Cowles and Goodrich-Blair 2006). CpxR, the response regulator of the two-component system CpxRA, positively regulates motility and production of LrhA and XlpA, as well as the expression of the nil colonization genes, and is thought to have an opposite, negative effect on antibiotic, protease and haemolysin activities (Herbert et al. 2007). Like Lrp, it is implicated in mutualistic and pathogenic host interactions, with both of them appearing to regulate the transition between hosts, via their opposed influence on the nil genes associated with nematode colonization (Richards and Goodrich-Blair 2009). Lrp may aid in the repression of nil gene expression both in the early stages of infection and during the reproduction stage, when they are not needed, as the mutualistic relationship with the nematode host is interrupted and the bacteria are engaged in a pathogenic relationship with the insect host. Once the insect is depleted of nutrients, and transmission to a new host becomes beneficial, CpxR may then activate nil gene expression, triggering the re-association process and therefore inducing the shift from a pathogenic relationship to a mutualistic one.

8.3  Insect Immunity The insects defence mechanisms can be classified into two broad strategies based on their specificity. The first one relies on non-specific structural and passive barriers such as the cuticle, designed to either keep the pathogens from entering the insect body altogether or, if the pathogens are ingested and therefore already inside, to prevent them from accessing the haemocoel through the gut wall and ease their destruction. This action is carried out by means of the peritropic membrane that lines the gut, and its physicochemical properties, respectively. The second strategy, much more complex, is specific and comprises the immune system as a whole, with its concomitant cellular and humoral immune responses, that interact in a coordinated way to respond to the pathogen challenges. Insects lack the adaptive immune defence mechanisms that vertebrates have. They instead rely entirely on the so-called innate immune system. In fact, this is the most ancient and common system of defence against microbes and parasites, and its elements have been conserved throughout evolution in the animal kingdom. The innate immune system in vertebrates and invertebrates is mediated by genes that are homologous or very closely related (Hoffmann 2003; Lemaitre and Hoffmann 2007). Both utilize a similar set of receptors, signalling pathways, transcription ­factors, humoral factors and cell-mediated mechanisms, in the course of the immune response.

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Although the cellular and humoral defence reactions are usually studied and considered separately in immunology, this division is somewhat artificial. The detection of a pathogen elicits in vivo a large array of interconnected and synergistic defence reactions in the immune-responsive connective tissue (i.e. haemolymph). The two systems clearly overlap, as the immune response involves a combination of soluble and cell-derived factors that are activated in conjunction (Theopold et  al. 2002; Jiravanichpaisal et  al. 2006). In the case of Drosophila, these switches between humoral and cellular responses in the course of the immune reaction are mediated by cytokine growth-blocking peptides (Tsuzuki et al. 2014). Depending on the final effector that attacks the foreign body, the innate immune reactions are characterized as either humoral or cellular. The humoral factors include activation of i) the prophenoloxidase/melanisation cascade, leading to wound healing and opsonisation of invading pathogens, ii) the coagulation cascade that immobilize foreign bodies, and iii) the induction of immune proteins such as lysozymes, lectins and AMPs (anti-bacterial and anti-fungal proteins). The cellular mechanisms are the haemocyte-mediated immune responses, interconnected with and stimulated by the previously mentioned humoral factors, and include coagulation, melanisation, phagocytosis of microbes and apoptotic tissues, nodulation and encapsulation of larger particles, such as parasitoid eggs, protozoa and nematodes, and production of reactive intermediates of oxygen and nitrogen.

8.3.1  Insect Haemocytes Like Arabidopsis thaliana, Caenorhabditis elegans or Danio rerio, Drosophila melanogaster is a model system, one that is extensively used for studies on the development of metazoans. Most studies in immunology have been devoted to the production of AMPs, and the reactions of Drosophila humoral immunity are now well understood. Cellular immune reactions have not been explored so extensively, although there is a growing tendency in studies investigating them, mostly being performed on lepidopterans (Liu et al. 2013), making comparisons between Drosophila and other insects increasingly necessary. Unfortunately, while the bulk of haemocyte types (plasmatocytes, granulocytes, oenocytoids and spherulocytes) are recognized and accepted in most insect species (Lavine and Strand 2002; Ribeiro and Brehélin 2006), Drosophila emerges as a peculiar exception in the class Insecta, as far as the cellular components of its haemolymph are concerned. Among the three main Drosophila haemocyte types mentioned in the literature, two of them, lamellocytes and crystal cells (Meister and Lagueux 2003; Ribeiro and Brehélin 2006), seem to be present only in Drosophila species. Furthermore, the third Drosophila haemocyte type, called plasmatocyte, seems to be very different from its homonym observed in species belonging to other insect orders, especially in Lepidoptera. For these reasons, and in an effort to facilitate the comparison of results obtained on cellular immune reactions, a thorough comparison between the Drosophila hae-

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mocyte types and those described in Lepidoptera is in order, and changes in nomenclature are long overdue (Ribeiro and Brehélin 2006). In fact, the lack of normalization in nomenclature has long been a standing issue, noted by more than one author (Strand 2008; Hillyer 2016). Notwithstanding difficulties in reaching a general agreement, there is a growing consensus and consistency in the homology and nomenclature uniformity between the species of Lepidoptera and those belonging to Drosophila (Brehélin and Zachary 1986; Ribeiro and Brehélin 2006; Lemaitre and Hoffman 2007; Tsuzuki et al. 2014). In fact, there is actually a clear correspondence between the morphology and physiology of the above mentioned insect haemocytes and comparisons are possible with lepidopteran plasmatocytes, granulocytes and oenocytoids, being homologous to Drosophila lamellocytes, plasmatocytes and crystal cells, respectively (Ribeiro and Brehélin 2006).

8.3.2  S  tructural and Functional Analysis of Typical Haemocyte types in Lepidoptera and their Corresponding types in Drosophila Insect haemocytes originate from mesodermal tissues, namely the hematopoietic organs and the lymph glands, from where stem cells are derived. These differentiate into specific haemocyte lineages identified by morphology, function, and molecular markers (Lavine and Strand 2002). A summary of the accepted insect haemocyte types and their respective functions is provided in Table 8.1.

Table 8.1  Most accepted insect haemocyte types for Lepidoptera and D. melanogaster, and related specific functions Lepidoptera

Haemocyte Type Prohaemocyte Plasmatocyte Granular cell Granulocyte Oenocytoid Spherule cell Spherulocyte

Drosophila melanogaster

Prohaemocyte Lamellocyte Plasmatocyte Crystal cell

Main functions Haematopoietic stem cell Spreading and adhesion, nodulation and encapsulation Adhesion, phagocytosis, coagulation, nodulation and encapsulation Prophenoloxidase reservoir; wound cicatrisation, cuticular melanization and sclerotization Reservoir and transporter of components for cuticular renovation, pupae/chrysalis formation and silk production for cocoons Haematopoietic stem cell Spreading and adhesion; encapsulation Adhesion, phagocytosis, coagulation, nodulation and encapsulation Prophenoloxidase reservoir; wound cicatrisation

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8.3.2.1  Lepidopteran Haemocytes Five main circulating haemocyte types have been recovered in all the lepidopteran species studied to date: (i) prohaemocytes, (ii) plasmatocytes, (iii) granulocytes or granular cells, (iv) oenocytoids and (v) spherulocytes or spherule cells. (i) Prohaemocytes Maintenance of circulating haemocytes in larvae of Lepidoptera has been attributed to both mitosis of haemocytes already in circulation and release from haematopoietic organs (Gardiner and Strand 2000). In all studied species, rare small haemocytes, regular in shape and with a high nucleus-to-cytoplasmic ratio have been reported. They are described as prohaemocytes, hypothesized to serve as stem cells or immature haemocyte progenitors for one or more of the remaining types (Grigorian and Hartenstein 2013). They are believed to be precursors of differentiated haemocyte types. This interpretation, however, is supported mostly by cytological features and is controversial. Further work is needed before this hypothesis can be confirmed. Additionally, different species of insects will likely show differences in haematopoietic organs and even in haematopoiesis as well (Liu et al. 2013). (ii) Plasmatocytes In vivo, plasmatocytes are spherical or oval cells (up to 20 μm long) with a regular shape, although they can sometimes appear spindle-shaped. In transmission electron microscopy (TEM), they show few pseudopods, small clear vacuoles, numerous polyribosomes and a moderate amount of rough endoplasmic reticulum (RER). The Golgi apparatus is present, but is often little developed, and is devoid of granules in most lepidopteran species. These cells always present pinocytotic vesicles largely distributed in the cell membrane (Ribeiro et  al. 1996; Ribeiro and Brehélin 2006). In monolayers, after a few minutes incubation, plasmatocytes are easily recognizable as they spread rapidly on contact with the glass slides, presenting a characteristic fibroblast-like morphology. They develop numerous pseudopodia and long, wide lamellipodia. They show a large rounded nucleus, with small scattered chromatin masses. It is widely recognized that plasmatocytes form the bulk of capsules around foreign bodies too large to be phagocytised, or nodules around masses of bacteria and necrotic melanised material, in vivo. Capsule and nodule formations look identical at the cytological level (Ratcliffe and Gagen 1976, 1977; Lavine and Strand 2002), and in these formations plasmatocytes synthesize numerous desmosomes and contain large amounts of microtubules in their cytoplasm (Ribeiro and Brehélin 2006). The role of plasmatocytes in phagocytosis is, however, disputed. For some authors, they are phagocytes (Ratcliffe and Rowley 1975; Tojo et al. 2000; Ling and Yu 2006), but for others they are clearly not phagocytic cells (Akai and Sato 1973; Neuwirth 1974; Ribeiro et  al. 1996; Beaulaton 1979; Ribeiro 2002; Ribeiro and Brehélin 2006).

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(iii) Granulocytes or Granular Cells After fixation upon haemolymph removal, lepidopteran granulocytes appear as spherical (diameter from 5 to 8 μm), very refractive cells in phase contrast. In TEM, they show a developed RER with enlarged cisternae filled with flocculent material, numerous Golgi complexes and mitochondria, and sparse glycogen particles dispersed in the cytoplasm. Several thin and long pseudopodia/filopodia were often visible. Similarly to plasmatocytes, granulocytes also possess numerous pinocytotic vesicles in their cell membrane. Three different kinds of membrane-bound inclusions have been described in these cells (see Brehélin and Zachary 1986), namely vesicles containing dense granules or structured granules, both having a rather regular, rounded shape, and being exclusively filled with materials synthesized by the Golgi apparatus. A third type of vesicles resemble phagolysosomes, highly irregular in shape and filled with heterogeneous material, either originated from cytoplasmic areas in autolysis or from a fusion between phagosomes and lysosomes containing hydrolases, synthesized by the Golgi complex (Ribeiro et al. 1996). Inclusions of this type can become very numerous and of large size (Raina 1976), for instance, at the end of the larval development (Essawy et  al. 1985; Ribeiro et  al. 1996; Nardi et  al. 2001, 2003). Granulocytes, overloaded with large phagolysosomes, could be mistaken for spherule cells on light microscopy. One of the main functions of granulocytes is phagocytosis (Costa et al. 2005; Ribeiro and Brehélin 2006). Granulocytes also represent the first cells to come into contact, in small numbers, with a foreign body at the beginning of the capsule/nodule formation. When in contact with a foreign body, they release their granular content, another of their critical functions (Akai and Sato 1973; Ratcliffe and Gagen 1977; Schmit and Ratcliffe 1977; Ribeiro et al. 1996). According to most authors, this exocytosis of typical opsonin-like material serves to attract plasmatocytes (Gillespie et al. 1997), or at least to help plasmatocytes to build the capsule or nodule (Pech and Strand 1996). Granulocytes and plasmatocytes are recognized as defensive haemocytes in most lepidopterans (Ribeiro et al. 1996; Brillard et al. 2001; Ribeiro and Brehélin 2006; Costa et al. 2009). The former are involved in mediating phagocytosis and nodule formation, whereas the latter act in the encapsulation process. The pinocytic vesicles, clearly visible in the cell membranes of granulocytes but also found on plasmatocytes, cannot go unnoticed, and studies on pinocytosis in insect immunocompetent cells are needed. Pinocytosis is usually considered to be constitutive, but it can also be a highly regulated, receptor-mediated process. Depending on the molecular mechanism involved, pinocytosis can be divided into four categories: caveolae-mediated, clathrin-dependent, macropinocytosis, and dynamin- and clathrin-independent (Seto et al. 2002). All of the abovementioned categories are involved in key signalling phenomena in eukaryotic cells. It is reasonable to assume that membrane trafficking and signal transduction are as important in insects as they are in mammals. Signalling pathways, namely those related to the presence of lipid rafts associated with c­ aveolae/

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caveolin-dependent pinocytic vesicles, may interact and regulate components of the membrane trafficking machinery in functions pertaining to the immune system, such as recognition of foreign bodies and cellular and humoral responses in insects. For instance, a large fraction of the cholera toxin can be endocytosed by clathrin-­ dependent mechanisms, as well as by caveolae-mediated and clathrin-independent endocytosis, in different cell types (Torgersen et al. 2001). Some bacterial pathogens have also been shown to exploit caveolin-1-rich membranes for infectious entry into cells (Parton and Richards 2003). Pinocytic vesicles seen in lepidopteran haemocytes (Ribeiro 1994; Ribeiro et al. 1996; Ribeiro 2002; Ribeiro and Brehélin 2006) resemble the caveolae that are generally associated with lipid rafts and are described as being flask-shaped invaginations present in the plasma membrane of many cell types. Caveolae have long been implicated in endocytosis, transcytosis, and cell signalling. They have been confirmed as directly involved in the internalization of membrane components such as glycosphingolipids, extracellular ligands (e.g. folic acid or albumin), bacterial toxins (e.g. cholera and tetanus toxins), and non-enveloped viruses, namely polyomaviruses. Unlike clathrin-mediated endocytosis, internalization through caveolae is a triggered event that involves complex signalling (Pelkmans and Helenius 2002). (iv) Oenocytoids After fixation, oenocytoids appear as large cells (diameter up to 25 μm) rather regular in shape, with variable refractivity in phase contrast microscopy, a low nuclear to cytoplasmic ratio and an often eccentric nucleus. In TEM, the cytoplasm is filled with numerous free ribosomes. Other typical cytoplasmic organelles are poorly developed, especially the Golgi apparatus and the RER cisternae, which are rare and very short, but often enlarged (Ribeiro and Brehélin 2006). In Lepidoptera, prophenoloxidase, the precursor of the cascade responsible for the melanisation reaction, is synthesized and stored within these haemocytes, and is released into the haemolymph when these cells lyse (Neuwirth 1973; Essawy et al. 1985; Iwama and Ashida 1986; Ashida et  al. 1988; Ribeiro 1994, 2002; Ribeiro et al. 1996). This causes the characteristic darkening, due to the oxidation of phenols to quinones, leading to spontaneous polymerization and to the formation of insoluble melanin (Söderhäll and Cerenius 1998; Nappi and Ottaviani 2000; Cerenius and Söderhäll 2004; Christensen et al. 2005). Activated prophenoloxidase also plays an important role in cuticular melanization and sclerotization, during larvae development (Marmaras et al. 1996). (v) Spherulocytes or Spherule Cells These are round cells, relatively stable in monolayers, containing a small number of large inclusions (the spherules) that cause the cell to adopt an irregular shape. In TEM, these inclusions show an internal structure of either lamellate concentric layers or a crystal-like lattice of dense particles, depending on the section (Ribeiro et al. 1996). The exact functions of these cells aren’t currently known, but they are considered as reservoirs and transporters of cuticular components. In G. mellonella, spherule cells appear full of globular contents, becoming conspicuously empty of

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their initial granules when the insects initiate silk production and pupation (Ribeiro 1994). In armyworm, Mithymna unipuncta, differential haemocyte counts during the last instar larval stage showed a dramatic reduction of their numbers, with the few present also devoid of sphere content (Ribeiro et al. 1996; 2002). Additionally, cytochemical assays (Thiéry 1967) using the Periodic Acid-Thiosemicarbazide-­ Silver Proteinate technique (PATAg) showed presence of polysaccharides in the spherule inclusions (Ribeiro 1994). These results were in conformity with those reviewed in Gupta (1991), Gupta and Sutherland (1967) and Ashhurst (1982), which also pointed at the presence of neutral and acidic mucopolysaccharides, glycomucoproteins and glycosaminoglycan in the spherule content. Despite their eventually different ontogeny, spherule cells participate in important physiological tasks, namely those related to cuticle renovation and pupae/chrysalis formation and cocoon silk production (Nittono 1960; Gupta 1991). In insects, the cuticle, which is secreted by the epidermis, is a complex structure, consisting primarily of an unbranched polymer of high molecular weight  – an amino-sugar polysaccharide composed of β(1-4) linked units of N-acetyl-D-­ glucosamine – combined with lesser amounts of phenolics, lipids, waxes and minerals. The cuticle sclerotization is an irreversible process that darkens insect’s exoskeleton and transforms the cuticular proteins in a water-insoluble matrix, via the insertion of phenolic compounds and quinones and the deposition of melanin, in a phenomenon known as tanning/darkening (Gullan and Cranston 1994). Considering the underlying role of oenocytoids in the melanisation process involved in sclerotization and the fact that spherule cells are transporters of cuticular components, a close physiological and biochemical collaboration between these two types of cells during the development, moulting and ecdysis of the insect larvae, seem likely. 8.3.2.2  Drosophila Haemocytes Drosophila has long been established as one of the best invertebrate models to study haematopoiesis. As a result, the process of haemocyte formation in its embryonic and larval stages is well documented (Mandal et al. 2007; Krzemien et al. 2007, 2010a, 2010b; Grigorian et  al. 2011, 2013; Grigorian and Hartenstein 2013; Yu et  al. 2018). During the embryonic stage, haemocytes are produced by the head mesoderm, whereas in the larval stage the lymph gland takes over this task (Lanot et al. 2001; Holz et al. 2003). Since the circulating haemocytes in the adult flies have been demonstrated to be either of embryonic or larval origin, adults were, until recently, perceived as being incapable of haematopoiesis. However, Ghosh et  al. (2015) presented evidence that there are active haematopoietic centres in adult flies. They house haemocytes from the two previously known lineages, in a network of laminin A and pericardin. Moreover, they also harbour stem cells derived from the larval lymph gland, capable of originating both crystal cells and fully functional plasmatocytes. These clusters are akin to simplified versions of the vertebrate bone marrow. This further reinforces the importance of Drosophila as a model for the

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study of vertebrate haematopoiesis. There are in fact significant similarities between the two processes, with conservation of signalling molecules, namely the phosphoinositol 3 kinase (PI3K) and GTPase Rac1 (Stramer et  al. 2005; Wood et  al. 2006), and the components of Toll, Imd, Jak/Stat and JNK pathways (Irving et al. 2005; Wertheim et al. 2005; Strand 2008). Three main types of circulating haemocytes have been reported in Drosophila, with a fourth being mainly described in the embryonic stage: (i) prohaemocytes, (ii) lamellocytes, (iii) plasmatocytes and (iv) crystal cells. (i) Prohaemocytes Circulating cells, resembling lepidopteran prohaemocytes (and presumed to have similar functions), are rare but present in Drosophila larvae, in particular during the embryonic stage, likely as stem cells of the haematopoietic tissues (Brehélin 1982; Lanot et al. 2001). Prohaemocytes rapidly differentiate to lamellocytes when insects are immune-challenged by intruders, such as parasitoid wasps, and during metamorphosis (Lanot et  al. 2001). Furthermore, invertebrate lymph glands contain dividing stem cells or haemocyte progenitors called prohaemocytes (Grigorian et al. 2011, 2013; Grigorian and Hartenstein 2013). (ii) Lamellocytes These are large (25-40 μm), flat cells, that differentiate upon immune induction and are rarely seen in healthy larvae (Brehélin 1982; Lanot et al. 2001). These haemocytes exhibit the same characteristics as the lepidopteran plasmatocytes (Ribeiro and Brehélin 2006). They have a regular shape, with very rare pseudopodia, lacking cytoplasmic inclusions, and appearing as flat, thin and adhesive cells in monolayers. They are recognized by reporters related to the Jun kinase signalling and L1 antigen (Lanot et al. 2001; Asha et al. 2003). In their cytoplasm there are numerous polyribosomes, whereas the Golgi apparatus and RER are poorly developed, with narrow cisternae. Like lepidopteran plasmatocytes, Drosophila lamellocytes form the bulk of capsules and nodules, showing the formation of numerous desmosome-like junctions (Russo et al. 1996) and microtubules (Rizki and Rizki 1979; Ribeiro and Brehélin 2006). Similarly, injected particulate material is phagocytosed in very low amounts, if at all, by Drosophila lamellocytes (Brehélin 1982; Lanot et al. 2001), as is the case with lepidopteran plasmatocytes (Costa et al. 2005). (iii) Plasmatocytes To add to the confusion in the nomenclature, the haemocytes that are called plasmatocytes in Drosophila are not equivalent to the lepidopteran plasmatocytes, as stated above. They have different histological and cytological features, different behaviour in monolayers and different functions (Ribeiro and Brehélin 2006). The Drosophila plasmatocytes are round cells that make up > 90% of circulating haemocytes. They are strongly adhesive in vitro, and function as phagocytes that engulf pathogens, dead cells and other entities, also participating in encapsulation and in the production of AMPs (Brehélin 1982). Furthermore, they possess cell

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membrane molecular markers that include the extracellular matrix protein peroxidasin and an uncharacterized surface marker called the P1 antigen (Nelson et al. 1994; Asha et  al. 2003). Although there is no true equivalent of lepidopteran granular haemocytes in the haemolymph of Drosophila species, we must emphasize that the cells called plasmatocytes in Drosophila look more similar to lepidopteran granular haemocytes than to lepidopteran plasmatocytes. Like lepidopteran granular haemocytes, circulating Drosophila plasmatocytes are spherical cells (diameter 5-8 μm), with a developed Golgi apparatus and RER, enlarged cisternae filled with flocculent material, numerous pinocytotic vesicles, thin pseudopodia and presence of more or less numerous phagolysosome-like inclusions. (iv) Crystal Cells Crystal cells are non-adhesive haemocytes that comprise approximately 5% of total circulating cells. They are considered equivalent to oenocytoids, due to high similarities both in structure and function, but are distinguishable by the presence of crystal-like cytoplasmic inclusions (Rizki and Rizki 1959; Nappi et al. 1995). In phase contrast microscopy, D. melanogaster crystal cells are rather large with a regular shape, an eccentric nucleus and irregular, sharp contour crystalline cytoplasmic inclusions. In other studied species of this genus, the above mentioned inclusions are rounded and do not look like crystals, resembling the lepidopteran oenocytoids instead (Rizki and Rizki 1980; Brehélin 1982). Crystal cells, like oenocytoids express components of the prophenoloxidase cascade, the activation of which leads to the formation of melanin (Rizki and Rizki 1959; Nappi et al. 1995).

8.4  The Bacterial Symbiont Arsenal Xenorhabdus and Photorhabdus produce numerous secondary metabolites that play different roles in their life cycles. These include not only several insecticidal and antimicrobial compounds, but also other pathogenicity factors, namely cytotoxins and cytolysins/haemolysins, as well as exoenzymes such as proteases and lipases. These metabolites may be produced upon release into the host haemocoel in order to suppress its immune response or actively circumvent it. This allows them to quickly kill the insect during their growth phase, as well as to protect the insect cadaver from competitors, afterwards. These may be saprotrophic organisms present in soil or the insect own gut microbiota. Moreover, the metabolites convert the host tissues into readily available nutrients to sustain the growth of both the bacteria and the EPN.

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8.4.1  Interacting with the Insect Host Immune System There are several mechanisms by which both Xenorhabdus and Photorhabdus interact with the insect host immune system. Although recognized by the insect immune system, they suppress its humoral response by inhibiting the activation of its major signalling pathways and response cascades, directly engaging and destroying the cellular arm of the immune system (ffrench-Constant et  al. 2007; Eleftherianos et al. 2010a; Costa et al. 2010; Nielsen-LeRoux et al. 2012). 8.4.1.1  Recognition and Triggering of the Immune Reaction The insect immune system is triggered when microbes are recognized by the host cells, in a process mediated by pattern recognition receptors (PRRs), namely hemolin, immunolectin-2 and peptidoglycan-recognition protein. These detect invariant features of microorganisms called pathogen-associated molecular patterns (PAMPs) (Seufi et al. 2012). PAMPs are, essentially, structural components of the bacteria cell wall, such lipopolysaccharides (LPS), peptidoglycan, lipoteichoic acid or lipoproteins. Specific recognition of a wide array of PAMPs by the PRRs allows the host to react to the pathogen triggering signalling responses (Medzhitov and Janeway 2002). Such a mechanism constitutes a very effective defence network against any possible invading microorganisms. Hemolins are bacteria-inducible immunoglobulin-like proteins acting as multifunctional molecules involved in a diverse range of cell interactions. They are able to: (i) bind lipopolysaccharides (Ladendorff and Kanost 1991; Daffre and Faye 1997; Yu and Kanost 2002); (ii) be upregulated during metamorphosis (Yu and Kanost 1999; Lindquist et al. 2005); (iii) agglutinate bacterial cells (Schmidt et al. 2010); (iv) promote opsonin-like effects, such as increased cell adhesion and phagocytosis (Lanz-Mendoza et al. 1996; Bettencourt et al. 1997); (v) bind to haemocytes and inhibit their aggregation (Kanost et  al. 1994), and (vi) regulate embryonic development (Bettencourt et al. 2000, 2002). Lectins are carbohydrate-binding proteins, and imunolectin-2 is a C-type lectin that is present at a constitutively low level in the larval haemolymph, recognized as a potent protector from bacterial infection. Its synthesis was experimentally induced after injection of Gram-negative bacteria or LPS, stimulating prophenoloxidase in haemolymph (Yu and Kanost 2003). After pathogen recognition, insects produce a plethora of AMPs that include attacin, cecropin and moricin in Lepidoptera and metchnikowin, diptericin, drosomycin and attacin in Drosophila. The recognition of pathogenic Photorhabdus and Xenorhabdus bacteria by PRRs, and the subsequently produced antimicrobials, have been shown to slow down the otherwise rapid killing of the infected insects.

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The genes encoding PRRs and AMPs are known to be transcriptionally induced at higher levels after infection by EB. Accordingly, silencing by RNA interference of the same genes results in an increase of the larval sensitivity to the pathogens (Eleftherianos et  al. 2006a, 2006b, 2010b). Despite the fact that the insects are clearly able to deploy various immune responses towards pathogenic bacteria, these systems are, in most cases, simply not effective enough. The bacteria, in the end, almost always prevail as they have the necessary genetic and molecular strategies to deal with, and dismantle the host response and escaping death (Costa, 2008). 8.4.1.2  Suppression of the Humoral Response As far as the suppression of the insect immune system is concerned, the first step appears to be the inhibition of the biosynthesis of eicosanoids (Eom et  al. 2014; Sadekuzzaman and Kim 2017; Sadekuzzaman et al. 2017a, b). Eicosanoids are a group of oxygenated C20 polyunsaturated fatty acids involved in the activation of various cellular and humoral immune responses, both in vertebrates and invertebrates (Park and Kim 2000; Park et al. 2004a, b, 2005; Kim et al. 2005; Shrestha and Kim 2007; Stanley and Kim 2014). They are extremely important in mediating the insect immune responses, being responsible for: (i) the increase of circulatory haemocytes and their migration, (ii) the expression of AMPs and (iii) the release of prophenoloxidase, the zymogen of phenoloxidase involved in the melanisation cascade (Kim et al. 2018). These polyunsaturated fatty acids are synthetized from arachidonic acid, which is in turn produced as the result of membrane lipid hydrolysis by phospholipase A2 (Burke and Dennis 2009; Cao et al. 2013). By inhibiting the production of arachidonic acid, several immune response pathways are directly and indirectly impacted. The host immune system hence becomes severely compromised, making phospholipase A2 an obvious target for any pathogen. This inhibition is achieved, in the case of X. nematophila, by producing, at least, eight different metabolites, that are synthetized at different stages of bacterial growth and provide differential immunosuppressive activity (Eom et al. 2014). All of the tested compounds produced significant results to varying degrees, but benzylideneacetone (BZA), Pro-Tyr (PY) and oxindole stood out as the most effective. Oxindole prevented phenoloxidase activity and BZA, which had already been reported to inhibit aggregation, spreading and nodulation of haemocytes (Kwon and Kim 2008), was shown to significantly inhibit haemocyte nodulation and, along with PY, exhibited phospholipase A2 inhibitory activity and cytotoxicity against insect cells. Another target for immunosuppression is phenoloxidase, a metalloenzyme that has a crucial role in the invertebrate immune response, triggering the deposition of melanin onto the cell surface of the invading pathogens, opsonizing them in order to stimulate encapsulation, effectively isolating the intruder from the surrounding haemolymph (Gillespie et al. 1997). Xenorhabdus and Photorhabdus are able to circumvent this melanisation reaction by targeting different steps of its cascade, through different mechanisms. For instance, Photorhabdus produces a hydroxystilbene compound that serves a dual function as antibiotic, inhibiting the growth of

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microbial competitors in the insect cadaver, and as a potent inhibitor of activated phenoloxidase (Eleftherianos et  al. 2007). Rhabduscin, an isocyanide-containing tyrosine derivative produced by both bacterial genera, is a virulence factor located in the bacterium outermost layer. It acts as an inhibitor for phenoloxidase, mimicking its substrate (Crawford et  al. 2012). Another example are the rhabdopeptide/ xenortide peptides, a large family of compounds that are commonly produced by Xenorhabdus and Photorhabdus. Given their high similarity with protease inhibitors are likely responsible for disrupting the serine cascade that results in the activation of prophenoloxidase into phenoloxidase (Tobias et al. 2018). Other compounds target the proteasome, whose main task is to degrade proteins that are no longer needed for the normal cellular function, and may be implicated in the removal and disabling of toxins and inhibitors produced by the pathogens during infection. Photorhabdus produces glidobactin and cepafungin, some of the more potent proteasome inhibitors currently known (Stein et  al. 2012). These compounds are included, alongside syringolins, in the syrbactin natural product class (Groll et al. 2008). These bear a 12-membered dipeptide-macrolactam resulting from linkage of a vinylogous amino acid and a modified lysine residue, a unique structure that confers them particular chemical and biological properties (Krahn et  al. 2011). Xenorhabdins are dithiolopyrrolone derivatives, first isolated from Xenorhabdus, that  were described as having a significant antibacterial activity against Gram-­ positive bacteria, but little effect against Gram-negative bacteria (McInerney et al. 1991a). These compounds are members of the dithiolopyrrolone group of bicyclic antibiotics that have been isolated from Actinomycetes and Proteobacteria (Li et al. 2014). Thiolutin, one of the most well studied members of this group, has recently been reported as a zinc-chelator, capable of inhibiting several metalloproteases, including one involved in the proteasome (Lauinger et al. 2017). 8.4.1.3  Directly Engaging the Cell-Mediated Response Effectors In their interactions with the cellular arm of the insect immune system, both Xenorhabdus and Photorhabdus produce several different types of compounds that directly target haemocytes, namely cytotoxins, cytolysins, haemolysins and proteases, as well as other pathogenicity factors. For instance, Xenorhabdus also employ specific morphological features that have been previously characterized and include: i) a cytotoxic fimbrial structural subunit (MrxA), a pore-forming toxin that lyses haemocytes (Banerjee et al. 2006); ii) a pilin subunit, secreted through outer membrane vesicles (OMVs) with binding cytotoxicity (Khandelwal et  al. 2004a) and agglutinating properties against haemocytes (Khandelwal et  al. 2004b); iii) lipopolysaccharides, which confer adverse effects on haemocyte functions (Giannoulis et al. 2008), and iv) type 1 fimbriae, that exhibit agglutinating activity with sheep, rabbit, and human erythrocytes and with haemocytes of G. mellonella (Moureaux et al. 1995).

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Type 1 fimbriae, filamentous appendages that are expressed by a variety of Enterobacteriaceae, are mannose-sensitive haemagglutination (MSHA) factors comprised of filamentous surface proteins that have been shown to facilitate bacterial adherence to specific host tissues by carbohydrate binding to surface receptors and to mannose-containing residues. They mediate the agglutination of erythrocytes of many vertebrate species and contribute to the adherence of pathogenic bacteria to eukaryotic cells (Sharon and Ofek 1986; Clegg and Gerlach 1987; Paranchych and Frost 1988; Gerlach et al. 1989; Johnson 1991; Krogfelt 1991; Forst et al. 1997; Duncan et al. 2005; Pizarro-Cerdá and Cossart 2006; Chandra et al. 2008). Several other types of enterobacterial fimbriae exhibit mannose-resistant haemagglutination (MRHA) and cell adherence, by recognizing either a carbohydrate moiety of a membrane glycoprotein (or glycolipid) or a carbohydrate-conjugated mucosal protein (Graaf and Mooi 1986; Johnson 1991; Hultgren et al. 1993). Fimbriae are an important virulence factor in enterotoxigenic Escherichia coli and uropathogenic strains of E. coli and Proteus mirabilis, enhancing colonization of epithelial cells (Gaastra and Graaf 1982; Johnson 1991; Massad et al. 1994; Li et al. 1999). The global regulator Lrp has long been implicated in the regulation of fimbriae production, being reported as both a positive and a negative regulator of the pap operon in E. coli (Woude et al. 1996) and the fim gene cluster responsible for type 1 fimbriae production in Salmonella enterica (Baek et  al. 2011). Lrp is a positive regulator of the mrx operon in X. nematophila (He et al. 2004), further emphasising the importance of this regulator in its motility and pathogenicity. It has been demonstrated that X. nematophila, during growth in artificial medium, produces different factors, assessed by chromatographic separation that, when incubated, exhibited protease, cytolytic and/or haemolytic activities in vitro towards midgut epithelium and haemocytes from: G. mellonella (Pyralidae), Agrius convolvuli (Sphingidae), Mythimna unipuncta (=Pseudaletia unipuncta) (Noctuidae), Spodoptera litorallis (Noctuidae), Sl2b cell line from S. litorallis (INRA-France), and sheep and rabbit erythrocytes (Ribeiro et al. 1999; Ribeiro 1994, 2002). The same studies showed that FITC-labelled LPS from X. nematophila, E. coli and Serratia marcescens did not exhibit any direct cytolytic or haemolytic effects against any of the aforementioned types of cells and gut tissues. In fact, only plasmatocytes showed slight adherence to the FITC-labelled cell membrane of E. coli, with no induced mortality. Although LPS extracted from X. nematophila induce lysozyme expression in the fat body of S. exigua, infection with live bacteria in the same host suppresses expression of cecropin and other AMPs, probably through inhibition of the eicosanoid pathway (Bae and Kim 2003; Ji and Kim 2004; Hwang et al. 2013). The fractions that exhibited protease activity targeted and degraded the midgut epithelium and caused monolayers of haemocytes adherent to glass slides to become unstuck, impairing their ability to recognize and adhere to foreign substrates, whereas the fractions exhibiting cytolytic/haemolytic activities lysed all the aforementioned cells types, with the exception of rabbit erythrocytes (Ribeiro et  al. 1999). Massaoud et al. (2010) did not observe any correlation between the pathogenicity of Xenorhabdus strains and their protease production, but there are ­indications

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that they may indeed play a role in pathogenicity, being involved in the destruction of AMPs (Eleftherianos et al. 2018). Photorhabdus produces a serralysin-type metalloprotease, PrtA that targets and selectively cleaves 16 haemolymph immune proteins in vitro, collectively named PAT (PrtA target). Included in this group are the coagulation cascade effectors scolexins A and B, proteins involved in recognition processes such as β-1,3 glucan recognition protein 2, and proteins involved in immune signalling and regulation, such as the haemocyte aggregation inhibitor protein (HAIP), six serpin-1 variants, including serpin-1I, and serine proteinase homolog 3 (Felföldi et al. 2009). The inactivation of serine proteinase homolog 3 has been demonstrated to significantly reduce the ability of Manduca sexta larvae to overcome Photorhabdus infection, due to the strong downregulation of multiple antimicrobial effector genes and of the gene encoding prophenoloxidase (Felföldi et al. 2011). Brillard et  al. (2001) evaluated the haemolytic activity of several strains of Xenorhabdus and Photorhabdus, through assays in sheep blood agar plates, liquid haemolytic assays using culture supernatants and sheep and rabbit erythrocytes in suspension, and lepidopteran haemocytes in monolayers. The P. luminescens strains evaluated displayed different haemolysis patterns in the blood plates and, in the liquid assays showed either no extracellular cytolytic activity against any of the cell types, or only a weak activity against rabbit erythrocytes. Given the presence of several putative haemolysin sequences in the P. luminescens genome, the authors hypothesized that the lack of extracellular haemolytic activity observed could be due to factors such as the need for these compounds to be processed like toxin complexes or the fact that cytolysis may be contingent on contact between the bacteria and the target cells. The X. nematophila F1 strain tested produced a halo of total discoloration in the blood agar plates, and showed extracellular cytolytic activity against all three cell types used in the liquid assays. Two distinct and successive bursts of extracellular cytolytic activity were detected during the in vitro growth of X. nematophila F1, corresponding to two growth stage-­ specific extracts termed C1 and C2. These were predicted to represent two discrete haemolysins regulated according to the growth phase. The C1 cytolytic activity occurred when bacterial cells reached the stationary phase and the corresponding extract exhibited haemolytic activity against sheep, but not rabbit, erythrocytes. Both lepidopteran immunocompetent cells, known for their major role in cellular immunity (Givaudan and Lanois 2000), were susceptible to C1 activity, with granulocytes being much more sensitive than plasmatocytes. Lysis by necrosis of the haemocytes was preceded by a dramatic vacuolization of the cells, that appeared swollen and with dilated endoplasmic reticulum vesicles. In contrast, the second burst of cytolytic activity (C2) occurred late during the stationary phase and caused haemolysis of rabbit, but not sheep, erythrocytes. C2 activity was effective against granulocytes and plasmatocytes, with plasmatocytes being much more sensitive than granulocytes. The affected cells appeared shrunken and devoid of lamellipodia, with condensed chromatin evident in the cytoplasm. The C1 and C2 extracts were predicted to represent two discrete haemolysins, regulated according to the

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b­ acterium growth phase, with the former being flhD dependent and heat labile and the latter being flhD independent and heat resistant (Brillard et al. 2001). Ribeiro et al. (2003) reported the purification of a flhDC-dependent, heat-labile cytotoxin responsible for the C1 cytolytic activity, a cation-selective Ca-independent porin-like peptide called α-Xenorhabdolysin (αX), and demonstrated that the plasma membrane of insect haemocytes and sheep erythrocytes were the primary targets of this toxin. Electrophysiological observations indicated that the initial effect of αX on macrophage/granulocytes plasma membranes was an increase of monovalent cation permeability, sensitive to potassium channel blockers. As a result of this increase in permeability, several events occurred intracellularly, such as selective vacuolization of the endoplasmic reticulum, cell swelling, and cell death by colloid-osmotic lysis. These effects, inhibited by potassium channel blockers, were totally independent of Ca2+ cations. The size of the pores created by αX on the plasma membranes of insect granulocytes and sheep erythrocytes increased with the toxin concentration, leading to rapid cell lysis. Later, Vigneux et al. (2007) reported the molecular characterization of αX as the XaxAB binary cytotoxin, an α-pore-forming toxin (α-PFT) encoded by the xaxAB genes. This was the prototype of a new family of hemolysins with both necrotic and apoptotic activities in insect haemocytes and mammalian cells, and indicated that homologues of the xax operon were present in the insect pathogenic bacteria P. luminescens and Pseudomonas entomophila, as well as in the human pathogens Yersinia enterocolitica and Proteus mirabilis. Waterfield et al. (2008) also identified a XaxAB homolog as a potential virulence factor of P. asymbiotica in a rapid virulence annotation screening. More recently, homologous cytotoxins were found in Y. enterocolitica and named YaxAB (Wagner et al. 2013). Zhang et al. (2014) reported on the cytotoxic and insecticidal activity of the XaxAB-like cytotoxin Plu1961/ Plu1962 from P. luminescens, termed PaxAB (ffrench-Constant and Dowling 2014). Recent works describing the structures of YaxAB (Bräuning et al. 2018) and XaxAB (Schubert et al. 2018) provided further insight into the mechanisms involved in the formation of the pore complexes and the dynamics of the aggregation of the two subunits. Schubert et al. (2018) compared the available data and pointed out that, although the protomer structures of YaxAB and XaxAB are very similar, there may be a species-dependent size variability, since the YaxAB pore is comprised of 8 to 12 heterodimers, as opposed to the 12 to 15 heterodimers found in the XaxAB pore. The proposed sequence of events leading to pore formation also differs between the two cytotoxins. YaxA is hypothesized to enter the membrane first and then recruit YaxB (Bräuning et al. 2018), whereas XaxA and XaxB first heterodimerize/oligomerize in solution, and then associate with the membrane as heterodimers or oligomers (Schubert et al. 2018). XaxAB is one of the two Xenorhabdus haemolysins directly regulated by FliZ, a global regulatory protein that controls the expression of the master flagellar ­regulator

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operon flhDC (Lanois et al. 2008; Jubelin et al. 2013), the other being the cell surface-associated XhlA, encoded by the xhlBA operon and belonging to the two-­ partner secretion system family (Cowles and Goodrich-Blair 2005). The first member of the two-partner secretion system family to be characterized was the Serratia marcescens haemolysin ShlA and its transporter ShlB, encoded by the shlBA operon. ShlB belongs to the TpsB protein family and was shown to be essential to the secretion and activation of ShlA, which belongs to the TpsA protein family. Homologues of the shlBA operon were also found in other Enterobacteriaceae, such as P. mirabilis (hpmBA), Edwardsiella tarda (ethBA) and P. luminescens (phlBA) (Brillard et al. 2002) and, more recently, X. nematophila (xhlBA) (Cowles and Goodrich-Blair 2005). XhlA was reported to be cell surface-associated and exhibited cytotoxic activity against rabbit and horse erythrocytes, and against both immunocompetent lepidopteran haemocytes, with plasmatocytes being far more sensitive than granulocytes. It was hypothesized that XhlA might be responsible for the C2 activity reported by Brillard et  al. (2001), since the observed biological effects of XhlA were very similar to those reported for the C2 extract, in all the comparable activities (Cowles and Goodrich-Blair 2005). However, the fact that XhlA appears to be outer-membrane bound and FlhDC-dependent (Cowles and Goodrich-Blair 2005), whereas C2 was clearly secreted to the extracellular medium and was reported as being flhD independent (Brillard et al. 2001) makes it improbable that XhlA is the cytotoxin responsible for the C2 activity, hinting that there is a still uncharacterized haemolytic agent being produced by X. nematophila. The haemolytic activity of P. luminescens TT01 was evaluated and PhlA, a homologue of XhlA, was identified and characterized (Brillard et  al. 2002). Haemolysis was reported in sheep and horse blood agar plates, but couldn’t be attributed to PhlA since a phlA-null mutant still produced the same activity suggesting a second, distinct haemolysin being produced by this strain. Haemolytic activity was also reported in liquid haemolytic assays using exponentially growing bacteria, incubated with horse erythrocytes. This activity was attributed to PhlA, since no haemolysis was produced by the phlA-null mutant. The authors also reported that, when filter-sterilized bacterial supernatants of the strains tested were used instead of the cultures, no haemolytic activity was observed. These results are in accordance with those obtained by Brillard et al. (2001) that reported that either no extracellular cytolytic activity or only a weak activity against rabbit erythrocytes was observed when using culture supernatants of Photorhabdus strains. This lack of activity in the absence of bacterial cells could point to PhlA being cell membrane-associated, as XhlA was reported to be (Cowles and Goodrich-Blair 2005), but a more exhaustive study is needed before that conclusion can be drawn. Brillard et  al. (2002) also reported that PhlA was not a major virulence factor in the infection of S. littoralis by P. luminescens, whereas XhlA was reported to be essential for full virulence of X. nematophila against M. sexta (Cowles and Goodrich-Blair 2005).

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8.4.2  Killing the Insect Once the immune system has been neutralized, the pathogens start multiplying and producing a plethora of virulence agents and insecticidal toxins, in order to quickly kill the host. These toxins can be separated into four main groups (Rodou et  al. 2010): i) toxin complexes (TCs), ii) “makes caterpillars floppy” toxins (Mcf), iii) Photorhabdus insect-related proteins (PirAB) and iv) Photorhabdus virulence cassettes (PVC). TCs are large multimeric protein complexes, some of which are highly toxic to insects and destroy the larvae midgut both when injected and per os (Bowen et al. 1998; Waterfield et  al. 2001; Silva et  al. 2002). TCs were first described in Photorhabdus and Xenorhabdus, being encoded by tc (Waterfield et al. 2001) and xpt genes (Morgan et al. 2001), respectively. However, homologs are present in a wide range of other pathogens (ffrench-Constant and Waterfield 2005). TCs comprise three subunits, each belonging to a different class of proteins, termed A, B and C and grouped based on their size and sequence similarity (ffrench-Constant et al. 2007); biological activity depends on the presence of the three subunits (Lang et al. 2010; Sheets et al. 2011). The C subunit, consisting of a core domain, highly homologue between all members of the C class, and a hyper variable region (Lang et al. 2011), was identified as the actual functional component of the toxin. It has an ADP-ribosyltransferase activity and induces major changes on the actin cytoskeleton, which most likely cause the inhibition of haemocyte phagocytosis. Different variants of the C subunit target different proteins, with TccC3 and TccC5 being well document examples. The former modifies actin, largely disabling the monomeric actin sequestering mechanisms that prevent its polymerization under normal circumstances, leading to the extensive clustering of actin, while the latter may cause persistent activation of Rho GTPases, leading to the massive formation of stress fibers. The effect of these two molecules is synergistic, as application of both caused complete aggregation of the actin cytoskeleton, forming star-like clusters all over the cells (Lang et al. 2010, 2011; Aktories et al. 2011). TCs function differently from any other known toxins, forming discrete compartments for protein unfolding and processing, and translocating the cytotoxic payload in a novel way (Gatsogiannis et al. 2013; reviewed in Meusch et al. 2014). The A subunit forms a translocation channel to which a cocoon formed by the B and C subunits then attaches. The cytotoxic domain of C is then cleaved and translocated through the pore formed by the A subunit into the cytoplasm of the host cell. Mcf toxins, when injected, induce apoptosis in the insect’s midgut epithelium, disrupting osmoregulation and leading to the characteristic loss of body turgor that gives them their name (Daborn et al. 2002; Waterfield et al. 2003; Dowling et al. 2004, 2007). The Mcf1 toxin, a gut-active multidomain protein is taken up into target cells by endocytosis. It induces apoptosis by means of a BH3-like domain. BH3 is one of four specific regions of homology (BH1, BH2, BH3 and BH4) found in the Bcl-2 family of proteins. These domains are critical in the interactions with other apoptotic family members and regulatory pro- and anti-apoptotic proteins, and

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play a pivotal role in their function as cell death regulators. BH3 is a potent cell death mediator that triggers the mitochondrial pathway, releasing cytochrome c and altering the membrane potential (Lutz 2000). Mcf1 also affects insect haemocytes and appears to have an anti-phagocytic activity, causing unexpected alterations in the actin cytoskeleton of host phagocytes, quickly paralysing them upon internalization, effectively preventing cell migration and any further phagocytosis – “freezing” phenotype. The mechanism by which Mcf1 affects the actin cytoskeleton may be related to the inactivation of Rho GTPases, namely Rac (Vlisidou et al. 2009). Binary proteins, that include PirAB, are toxic to some insects and destroy the midgut epithelium of the larvae upon ingestion, causing the swelling and shedding of the apical membranes normally seen in presence of other gut active toxins (Duchaud et al. 2003; Waterfield et al. 2005; Blackburn et al. 2006). Also included in this group are the Plu1961/Plu1962 cytotoxins (Zhang et al. 2014), that are highly toxic to larvae when injected, causing necrosis of the midgut epithelium and are quite similar to the XaxAB haemolysin from X. nematophila (Vigneux et al. 2007), a pore-forming cytolysin (Ribeiro 2002; Ribeiro et al. 2003) causing necrosis and apoptosis of insect haemocytes. PVC are prophage-like loci predicted to encode 15-20 proteins, showing a different putative effector sequence each. Despite the fact that the structure of PVC products is highly similar to that of known bacteriocins, they show no antibacterial activity. Instead, the PVC-encoded effectors are toxic when injected into insect larvae, and are responsible for a dramatic condensation of the actin cytoskeleton of insect haemocytes, leading to cell death (Yang et al. 2006).

8.4.3  Settling in and Getting Rid of Competition After successfully evading the insect’s immune system and producing enough toxins to kill it, the next step is the production of hydrolytic enzymes to start converting the insect biomass into a nutrient-rich soup, with metabolites that facilitate the growth of both symbionts (Vizcaino et  al. 2014). Antimicrobial compounds are needed to keep other microorganisms (both present in the soil environment or living in the insect gut or on the nematode cuticle), from proliferating in the cadaver and competing with them for the readily available resources. To this end, Xenorhabdus produces a veritable arsenal (Bode 2009), that fall into one of two categories (Boemare et al. 1992): i) broad spectrum antibiotics (BSA) and ii) narrow-spectrum bacteriocins (NSB). BSA are active against a wide range of microorganisms. Foremost among them are  xenoamicins, large hydrophobic depsipeptides active against parasitic protozoa, without antibacterial activity (Zhou et al. 2013), xenocoumacins, benzopyranone derivatives that are highly active against Gram-positive bacteria and also exhibit antimycotic activity against several fungal species (McInerney et al. 1991b; Reimer et al. 2009), and PAX-peptides, that show antifungal and antibacterial activity (Gualtieri et al. 2009; Fuchs et al. 2011).

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NSB include the phage-derived xenorhabdicin (Thaler et al. 1995), and the colicin E3-type killer protein xenocin (proteins involved in signalling, regulation, pathogenicity, as well as in metabolism) produced by Xenorhabdus. NSB are active mainly against bacteria from the same genus or closely related genera, likely produced not only to eliminate direct competition for nutrients (Singh and Banerjee 2008), but also to ensure the bacteria vertical transmission to the nematode progeny, upon recolonization of the IJs, before the nematodes emerge from the insect cadaver to search for a new host (Thaler et al. 1995). Since Photorhabdus and Xenorhabdus have never been isolated from environmental samples, it is presumed that their entire life cycles unfold within a biological reservoir. Concomitantly, in their tripartite way of life (nematode gut - insect gut insect haemocoel) the EB display two different, perhaps opposed, life styles as mutualistic symbionts and as parasite/pathogenic symbionts. Insect guts present distinctive environments for microbial colonization, and the bacteria naturally present in the gut potentially provide many beneficial services to their hosts. They have also been shown to positively contribute to nutrition, protection from parasites and pathogens, modulation of immune responses, and communication (Engel and Moran 2013). After leaving the nematode, the insect gut represents the first challenging sector of their dual lifestyles. In fact, Xenorhabdus and Photorhabdus face a new task, because the indigenous microbiota poses an obstacle to the colonization by non-­ indigenous species, including pathogens, while also playing an important role in resisting infection. Gut bacterial consortia adapt by the transfer of plasmids and transconjugation between bacterial strains, and some insect species provide ideal conditions for bacterial conjugation, suggesting that the gut is a “hot spot” for gene transfer (Dillon and Dillon 2004). It is in this adverse environment that the EB will be introduced, to be challenged by various detrimental conditions, which they have to sense and adapt to, in order to prevail. To achieve this goal, several complex mechanisms of sensing, signalling, and regulation have evolved in both genera (Heermann and Fuchs 2008). A comparative analysis between P. luminescens and Y. enterocolitica uncovering candidate genes encoding proteins involved in signalling, regulation and pathogenicity, as well as in metabolism, employed in the gut invasion and subsequent insect exploitation of nutrients identified a set of factors shared by the two pathogens involved in the host infection process, in persistence within the insect, or in host exploitation. These results not only improved the understanding of the biology of both pathogens, but also revealed some implications on the evolution of invertebrate and vertebrate virulence factors. Carneiro et al. (2008) reported a dramatic decrease in the numbers and viability of natural enteric microbiota in the intestinal lumen of the lepidopteran Diatrea sacchralis, upon infection by Photorhabdus. Oral infection by Photorhabdus is known to trigger gut immune responses in the host, namely the expression of nitric oxide synthesis specifically in the gut, which prevents the pathogens from crossing into the haemolymph via the gut wall (Eleftherianos et al. 2009). However, the efficacy of these immune responses can vary dramatically and in a predictable way ­depending

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on the stage of development and/or age of the insect. Age-related variation in immunity in the adult stage has been frequently reported (DeVeale et al. 2004) and, in fact, older insect larvae have a reduced immune capacity, which is marked by an increased colonization of their gut (Eleftherianos et al. 2008). Joshi et  al. (2008) reported that X. nematophila secretes, through OMVs, an insecticidal protein, a GroEl homolog that confers oral toxicity to larvae of Helicoverpa armigera through gut membrane binding. This protein is made up of three different domains exhibiting various levels of binding and insecticidal activity. Similarly, Singh et al. (2013) reported the identification of the xenocin operon of X. nematophila that includes the ximB and xciA genes encoding, respectively, the 42-kDa immunity protein (IP) and a 64-kDa xenocin, possessing an RNase catalytic domain, with a specific intramolecular target in 16S rRNA (Zarivach et al. 2002), responsible for its strong lethal effect on the competing microbes present in the larval gut. These antibacterial xenocins are coexpressed with periplasmic IP, which protects Xenorhabdus from the lethal effects of their own xenocins. For this purpose, McQuade and Stock (2018) reported that the xenocins may be exported from the cytoplasm, through the cellular membrane, by the Sec translocase or Twin-arginine translocation (Tat) pathways into the periplasmic space. In certain physiological conditions, the external delivery of xenocins is processed by the type VI secretion system (Singh et al. 2013; McQuade and Stock 2018). Gram-negative bacteria use a well-known pathway to export products through their OMVs (Beveridge 1999). Khandelwal and Banerjee (2003) demonstrated the insecticidal potential of the outer membrane-associated proteins secreted by X. nematophilus. These proteins exhibited in vivo oral cytotoxicity, targeting the gut of neonate larvae and in vitro cytotoxicity to Sf-21 cells. The presence of chitinase activity together with bacteriocins, adhesins, and pore-forming proteins in the insecticidal multiprotein complex found in OMVs supports the role of this complex in pathogenicity, as these proteins are known to mediate host-pathogen interactions in other pathogenic bacteria. OMVs provide an efficient mechanism of protection, transport and delivery of effector molecules within the larval host, not only in the insect gut, but also in the haemocoel, where they target the haemocytes.

8.5  Conclusion EB and their associated nematodes, acting as biological reservoirs and delivery vectors to their insect pest targets, have great ecological and socioeconomic importance. Their genetic, molecular, cellular, metabolic, adaptive and co-evolutionary features make these nematode-bacterial symbionts excellent models for understanding the genes and molecular regulators that govern the fundamental biological processes in hosts and pathogens alike. Regardless of the end result, pathogenicity is a complex equation that balances the strategies of both pathogens and hosts, the first striving to overcome, the other

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Fig. 8.1  Schematic overview of the interaction between the symbiotic pairs Steinernema: Xenorhabdus and Heterorhabditis: Photorhabdus and their hosts, with emphasis on the different means by which the bacteria overcome and circumvent the insect immune system (Illustration created with BioRender)

to defend, and both trying to maximize their chances of survival, managing their mutual interactions and those with the surrounding environment. From the insect standpoint, we addressed the immune response to the pathogens challenge, and elucidated the structure and function of their immunocompetent cells, the haemocytes. We dissected the different ways by which the EPN bacterial symbiont circumvents and disables the insect immune system, rendering the various defence mechanisms at work during infection all but useless (Fig. 8.1). On the other side, looking at the life cycle of the EPN symbionts, we focused on their tripartite life, in which the bacterium shows mutualistic and pathogenic facets, modulating basic functions of the nematode host, facilitating its survival and, therefore, ensuring its own, ultimately influencing the insect response, only to overwhelm it.

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Chapter 9

Effects of Entomopathogenic Nematodes and Symbiotic Bacteria on Non-target Arthropods Ramandeep Kaur Sandhi and Gadi V. P. Reddy

Abstract  The concerns over environmental risks of chemical pesticides and their consequences for insect pest control have stimulated the search for alternative control measures such as biological control by entomopathogens. Nematodes of the families Steinernematidae and Heterorhabditidae are biological control agents that are non-toxic to humans, safer to environment than chemical pesticides, and can easily combined with other control tactics. However, there has always been concern regarding potential non-target effects of these parasitic nematodes. Only a few studies have examined the action of entomopathogenic nematodes on non-target organisms and these cases have shown effects ranging from negligible to harmful. In some cases, these nematodes can cause mortality to non-target organisms, usually on a temporary and restricted area basis. We review herein the direct and indirect effects of entomopathogenic nematodes on non-target organisms, from both laboratory and field studies. Keywords  Biological control · Entomopathogenic nematode · Steinernematidae · Heterorhabditidae · Non-target organism

9.1  Introduction The continuous use of chemical pesticides for insect pest control has led to various environmental problems including human illness, soil and water pollution, and insecticide resistance in insect pests. Frequent use of some insecticides may increase pesticide resistance in crop pests, increase the cost of crop production, and harm non-target organisms, including the natural enemies of pest insects. The negative effects of organophosphate and carbamate insecticides, for example, both on non-­ target organisms in soil (Cockfield and Potter 1983, Floate et al. 1989) and aquatic R. K. Sandhi · G. V. P. Reddy (*) Department of Research Centers, Western Triangle Agricultural Research Center, Montana State University-Bozeman, Conrad, MT, USA e-mail: [email protected] © Springer Nature Switzerland AG 2019 M. A. Khan, W. Ahmad (eds.), Microbes for Sustainable Insect Pest Management, Sustainability in Plant and Crop Protection, https://doi.org/10.1007/978-3-030-23045-6_9

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environments (Hurlbert et al. 1972) have been well documented. Development of integrated pest management (IPM) strategies with little or no reliance on chemical pesticides has become an important goal for modern agriculture. One of the important IPM strategies is to maximize the impact of biological control agents, such as predators, parasitoids, and pathogens (bacteria, fungi, viruses, and nematodes). Entomopathogenic nematodes (EPNs) (families Steinernematidae and Heterorhabditidae) are the insect parasites that kill their hosts with the help of mutualistic symbiotic bacteria (Xenorhabdus spp. and Photorhabdus spp. for steinernematids and heterorhabditids, respectively). These symbionts are motile Gram-negative bacteria in the family Enterobacteriaceae (Poinar 1990; Adams and Nguyen 2002; Lewis and Clarke 2012). Steinernematidae and Heterorhabditidae nematodes have the only one free-living stage, being the third juvenile stage, also called as infective juveniles (IJs). This stage enters the host through natural openings (mouth, anus, and spiracles), or in some cases, through the cuticle (Campbell and Gaugler 1991). These infective juveniles carry their symbiotic bacterium in a bi-lobed vesicle in the intestine (Martens et al. 2003). After entering the host’s haemocoel, nematodes release bacteria into the host hemolymph, causing septicemia, which kills the host in 24–48 h. Nematodes molt and complete up to three generations within the host body. Once the nutrient supply is depleted, IJs burst out of the cadaver and search to find new hosts (Poinar 1990; Lewis and Clarke 2012). For more details on EPN biochemistry see Chap. 8 of this Volume. EPN’s have been investigated worldwide for the management of a wide range of economically important pest insects because of their suitability for mass production, ease of application, host-seeking ability, ability to rapidly kill their hosts, and safety to mammals and other non-target organisms. They are also exempt from the U.S. federal pesticide regulation, making commercialization feasible. However, due to their sensitivity to UV radiation and unfavorable temperatures, EPNs have been successful primarily for the management of soil-inhabiting pests and pests in greenhouses or other forms of protected cultivation (Georgis and Gaugler 1991). However, as for any pest control tactic, concerns for potential non-target effects need to be understood. The ecological impacts of field application of EPNs on non-­ target arthropods is an important but relatively neglected topic. EPNs, which as a group have wide host ranges, are more likely to affect non-target arthropods (insect biocontrol agents such as parasitoids and predators). On the other hand some EPN species have limited host ranges. However, the risk of EPNs to non-target species under field conditions may be less than those implied by laboratory tests because of the limited dispersal and lack of persistence of these nematodes (van Lenteren et al. 2003; Lynch and Thomas 2000). Here we discuss the potential of EPNs to cause either direct or indirect harm to non-target organisms under laboratory or field conditions. We also discuss future prospects for further work needed to avoid important non-target impacts of EPNs.

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9.2  Symbiotic Relationships of EPNs with Bacteria Different species of EPNs (of Steinernema or Heterorhabditis) host different species of symbiotic bacteria (Table 9.1) (Boemare (2002); Ganguly (2006); Stock and Goodrich-Blair (2008); Plichta et  al. (2009); Askary (2010); Tailliez et  al. 2012; Kuwata et al. 2013; Ferreira et al. (2014). The EPN life cycle consists of two stages (Fig. 9.1): i) a free-living stage in the soil that carries the symbiotic bacteria in their gut and searches for new insect hosts and ii) a parasitic stage in which infective juveniles penetrate the host, release their bacterial symbionts, and reproduce (Emelianoff et al. 2007). The symbiotic bacteria require the nematode species for dissemination from one host to another (Akhurst and Boemare 1990; Boemare 2002), and in return nematodes rely on nutrients and metabolites produced by their symbionts for growth and reproduction (Goodrich-Blair 2007). Also, nematodes shelter the bacteria from unfavorable environmental conditions, and bacteria produces antibiotic and hydrolytic enzymes that degrade the insect body into a nutrient soup that provides nutrition to the nematodes. The bacteria also release toxins that prevent other pathogens or microorganisms from attacking tissues of the moribund host. These toxins can be insecticidal, antifungal, or antibacterial that inhibit the other pathogens to infect or colonize the hosts, thus making the host specific to the EPNs (Khandelwal and Banerjee-Bhatnagar 2003). This association characterizes the symbiotic relationship between nematodes and bacteria (Burnell and Stock 2000; Dowds and Peters 2002; Hazir et al. 2003). Other than steinernematids and heterorhabditids, nematode species in several other families (Phaenositylenchidae, Merminthidae, Sphaeririidae, Tetradonematidae and Allantonematidae) are parasitic on insects (Stock and Hunt 2005). Members of these families have been recovered from insects belonging to different orders (Coleoptera, Diptera, Thysanoptera, Lepidoptera, and Hymenoptera). However, nematodes in these families do not have the mutualistic bacteria found in steinernematids and heterorhabditids, and their inability to reproduce in artificial media make them of little use for pest control (Arthurs et al. 2004).

9.3  Bacterial Symbionts as Biocontrol Agents Xenorhabdus and Photorhabdus, the bacterial symbionts of steinernematid and heterorhabditid nematodes, are gram-negative bacteria belonging to family Enterobacteriaceae. They produce chemical compounds with antimicrobial, nematicidal, insecticidal, and even anti-cancer activity (Webster et al. 2002). Xenorhabdus species can infect species of both Lepidoptera (Ji and Kim 2004; Khandelwal et al. 2004; Banerjee et al. 2006; Kumari et al. 2013; Kalia et al. 2014) and Hymenoptera (Dudney 1997) when injected artificially into their bodies in the laboratory.

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Table 9.1  Bacterial species mutualistically associated with different EPN species Entomopathogenic nematodes Steinernematidae Steinernema affine Bovien; S. anatoliense Hazir, Stock and Keskin; S. feltiae Filipjev; S. intermedium Poinar; S. jollieti Spriridonov, Krasomil-Osterfeld and Moens; S. kraussei Steiner; S. litorale Yoshida; S. oregonense Liu and Berry; S. puntauvense Stock, Uribe-Lorio and Mora; S. silvaticum Sturhan, Spiridonov and Mrácek; S. weiseri Mrácek, Sturhan and Reid S. bicornutum Tallosi, Peters and Ehlers

Bacteria Xenorhabdus bovienii Akhurst

X. budapestensis Lengyel, Lang, Fodor, Szállás, Schumann, and Stackebrandt S. riobrave Cabanillas, Poinar and Raulston X. cabanillasii Tailliez, Pages, Ginibre, and Boemare S. diaprepesi Nguyen and Duncan X. doucetiae Tailliez, Pages, Ginibre, and Boemare S. serratum Li X. ehlersii Lengyel, Lang, Fodor, Szállás, Schumann, and Stackebrandt S. hermaphroditum Stock, Griffin and Chaenari X. griffiniae Tailliez, Pages, Ginibre, and Boemare S. karii Waturu, Hunt and Reid; S. monticolum Stock, Choo and X. hominickii Tailliez, Kaya Pages, Ginibre, and Boemare S. abbasi Elawad, Ahmad and Reid; S. thermophilum Ganguly and X. indica Somvanshi, Lang, Singh Ganguly, Swiderski, Saxena, and Stackebrandt S. scapterisci Nguyen and Smart X. innexi Lengyel, Lang, Fodor, Szállás, Schumann, and Stackebrandt S. aciari Qui, Yan, Zhou, Nguyen and Pang X. ishibashii Kuwata, Qiu, Wang, Harada, Yoshida, Kondo, and Yoshiga S. kushidai Mamiya X. japonica Nishimura, Hagiwara, Suzuki, and Yamanaka S. scarabaei Stock and Koppenhöfer X. koppenhoeferii Tailliez, Pages, Ginibre, and Boemare S. arenarium Artyukhovsky X. kozodoii Tailliez, Pages, Ginibre, and Boemare S. australe Edgington, Buddie, Tymo, Hunt, Nguyen, France, X. magdalenensis Tailliez, Merino and Moore Pages, Edginton, Tymo, and Buddie S. carpocapsae Weiser; S. websteri Cutler and Stock X. nematophila corrig. Poinar and Thomas (continued)

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Table 9.1 (continued) Entomopathogenic nematodes S. cubanum Mrácek, Hernandez and Boemare; S. glaseri Steiner S. puertoricense Román and Figueroa S. siamkayai Stock, Somsook and Kaya S. costaricense Stock, Uribe-Lorio and Mora; S. rarum de Doucet

Bacteria X. poinarii Akhurst X. romanii Tailliez, Pages, Ginibre, and Boemare X. stockiae Tailliez, Pages, Ginibre, and Boemare X. szentirmaii ehlersii Lengyel, Lang, Fodor, Szállás, Schumann, and Stackebrandt Not known

S. akhursti Qui, Hu, Zhou, Mei, Nguyen and Pang; S. apuliae Triggiani, Mrácek and Reid; S. ashiunense Phan, Takemoto and Futai; S. asiaticum Anis, Shahina, Reid, and Rowe; S. longicaudum Shen and Wang; S. leizhouense Nguyen, Qui, Zhou, and Pang; S. loci Phan, Nguyen, and Moens; S. neocurtillae Nguyen and Smart; S. pakistanense Shahina, Anis, Reid, Rowe and Maqbool; S. ritteri de Doucet and Doucet; S. robustispiculum Phan, Subbotin, Waeyenberge, and Moens; S. sangi Phan, Nguyen, and Moens; S. sasonense Phan, Spiridonov, Subbotin, and Moens; S. sichuanense Mrácek, Nguyen, Tailliez, Boemare, and Chen; S. tami Luc, Nguyen, Spiridonov, and Reid; S. thannhi Phan, Nguyen and Moens; S. yirgalemense Nguyen, Tesfamariam, Gozel, Gaugler, and Adams Heterorhabditidae Photorhabdus asymbiotica Heterorhabditis gerrardi Plichta, Joyce, Clarke, Waterfield, and Stock Fischer-Le Saux, Viallard, Brunel, Normand, and Boemare subsp. nov. H. zealandica Poinar (South Africa strains) P. heterorhabditis Ferreira, van Reenen, Endo, Tailliez, Pages, Sproer, Malan, and Dicks (strains SF41 and SF783) H. indica Poinar, Karunakar and David; H. marelata Liu and Berry P. luminescens subsp. Akhurstii Fischer-Le Saux, Viallard, Brunel, Normand and Boemare subsp. nov. H. bacteriophora Poinar (HP88) P. luminescens subsp. Laumondii Fischer-Le Saux, Viallard, Brunel, Normand and Boemare H. bacteriophora (Brecon) P. luminescens subsp. Luminescens Thomas and Poinar H. bacteriphora (NC1); H. downesi Stock, Burnell, and Griffin; H. P. temperata Fischer-Le megidis Poinar, Jackson, and Klein (Nearctic group); H. zealandica Saux, Viallard, Brunel, Normand, and Boemare Poinar (continued)

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Table 9.1 (continued) Entomopathogenic nematodes H. megidis (Palaearctic group)

H. amazonensis Andaló, Nguyen, and Moino; H. baujardi Phan, Subbotin, Nguyen, and Moens; H. brevicaudis Liu; H. floridensis Nguyen, Gozel, Koppenhofer, and Adams; H. Mexicana Nguyen, Shapiro-Ilan, Stuart, McCoy, James, and Adams; H. poinari Kakulia and Mikaia; H. taysearae Shamseldean, Abou El-Sooud, Abd-Elgawad, and Saleh

Bacteria P. temperata temperata Fischer-Le Saux, Viallard, Brunel, Normand, and Boemare Not known

Fig. 9.1  Life cycle of entomopathogenic nematodes

Similarly, species of Photorhabdus are virulent against a wide range of insects, including species of Lepidoptera (Mohan et al. 2003; Mahar et al. 2005; Jallouli et  al. 2013); Coleoptera (Blackburn et  al. 2005; Shrestha and Kim 2010), Thysanoptera (Gerritsen et al. 2005; Uma et al. 2010), Hemiptera (Blackburn et al. 2005; Fand et  al. 2012; Kumar et  al. 2014), Orthoptera (Mahar et  al. 2004),

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Hymenoptera (Bowen and Ensign 1998), and Diptera (Ahn et al. 2013) in the laboratory when hosts are infected orally or by injection. Bussaman et al. (2009) reported more than 80% mortality in female mushroom mites, Luciaphorus perniciosus Rack (Pygmephoridae) within 3 days of application of P. luminescens. Some field studies reported the insecticidal virulence of P. luminescens in a variety of hosts: the cabbage white butterfly, Pieris brassicae L. (Lepidoptera: Pieridae) (Mohan et al. 2003), Drosicha mangiferae Stebbins (Hemiptera: Margarodidae) (Mohan et  al. 2004), and pupae of diamondback moth, Plutella xylostella L. (Lepidoptera: Plutellidae) (Razek-Abdel 2003). Jallouli et al. (2013) reported near 100% mortality in a stored grain pest, Ephestia kuehniella Zeller (Pyralidae), when treated with P. temperata K122 used at the rate of 12 × 108 cells/ml. Few researchers have examined the effect of the symbiotic bacteria or their secondary metabolites on non-target arthropods, independent of their EPNs. Lalitha et al. (2012) studied the effect of P. luminescens (the symbiotic bacteria of H. bacteriophora strain PDBC Hbb1) on pupae and adults of the egg parasitoid Trichogramma chilonis Ishii (Hymenoptera: Trichogrammatidae) and eggs and larvae of the predator Chrysoperla zastrowi sillemi Esben-Peterson (Neuroptera: Chrysopidae). There was no physical changes in the eggs, larvae, and adults of T. chilonis and C. zastrowi sillemi, and no significant reduction was observed in egg hatch, adult emergence, or parasitism from the bacteria. Mohan and Sabir (2005) tested the virulence of P. luminescens (isolated from H. indica) against T. chilonis and Trichogramma japonicum Ashmead (Hymenoptera: Trichogrammatidae) inside eggs of their host Corcyra cephalonica Stainton (Lepidoptera: Pyralidae) in the laboratory at the rate of 1 × 105 cells per ml, and reported 84% reduction in emergence of Trichogramma adults from treated host eggs. However, despite biological activity of EPN symbiotic bacteria when applied artificially, to date, free-living forms of Xenorhabdus and Photorhabdus have not been found in the soil. Rajagopal and Bhatnagar (2002) and Mitani et al. (2004) reported that the nematodes alone can cause host mortality but cannot reproduce. Similarly, bacteria can cause mortality on their own but need nematodes for their reproduction and, apparently, cannot live alone in soil without nematodes (Burnell and Stock 2000). These observations suggest that there is little likelihood that either EPNs or their symbiotic bacteria can be effective for pest control, when used alone.

9.4  N  on-target Effects of EPNs and Bacterial Symbionts on Arthropods EPNs typically have broad host ranges, suppressing a wide range of insects, including species of Coleoptera, Lepidoptera, Diptera, Thysanoptera, Orthoptera and Isoptera (Grewal et  al. 2005; Půža and Mráček 2005; Arthurs and Heinz 2006; Barbara and Buss 2006; Malan and Manrakhan 2009; Khan et  al. 2016). The

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numbers of infective juveniles (IJs) applied in inundative programs are high, ranging from 2.5 to 7.5 × 109 IJs per hectare (Georgis et al. 2006). Despite being safer than conventional chemical pesticides to the environment, EPNs may have some potential to affect some non-target arthropods due to broad host ranges and the high doses used in the field. They have successfully controlled insect pests in several habitats, including soil, holes and galleries of borers, leaf mines, curled leaves, flowers, and buds, as well as, in some cases, on simple leaf surfaces. These habitats provide protection to EPNs from dryness and other unfavorable environmental conditions. In such habitats, non-target organisms may come in contact with EPNs and get affected. Also, EPNs may also have indirect effects on non-target organisms, as for example through the ingestion by predators of insects that are infected by EPNs, or the death of parasitoids inside infected hosts. It is known that predation of EPN-infected insects also impairs the development of EPNs themselves within the insect because, in addition to destroying the host, EPN development can be harmed by more rapid host dessication (Baur et al. 1998; Foltan and Puza 2009). Previous reviews on non-target effects due to EPNs include Poinar (1989), Bathon (1996), and Piedra-Buena et al. (2015), as well as various individual studies, most based on laboratory assays (Table 9.2), discussed here by taxonomic groupings of non-target species.

9.4.1  Coleoptera Most studies of EPN effects on non-target organisms assessed levels of mortality under laboratory conditions. Georgis et al. (1991) found that adult predators were not susceptible to EPN nematodes under laboratory conditions, but their immature stages were more prone to infection. In the field, Georgis et  al. (1991) did not observe any effects of EPNs on non-target predatory beetles in the Carabidae, Staphylinidae, or Histeridae in four crops (turfgrass, corn, cabbage, and cranberries) treated with S. carpocapsae, S. feltiae, or H. bacteriophora. Farag (2002) reported high mortality of larvae of Coccinella undecimpunctata L. (Coccinellidae) due to H. taysearae and S. carpocapsae strain S2 under laboratory conditions. Similarly, Rojht et al. (2009) found 93% mortality of twospotted lady beetle, Adalia bipunctata (L.) (Coccinellidae) in laboratory due to S. feltiae, S. carpocapsae, or H. bacteriophora. However, Shapiro and Cottrell (2005) found lower mortality (< 30%) to lady bird beetles, C. septempunctata, C. maculata, H. axyridis, and Olla v-nigrum as compared to 90% mortality to their host, Agrotis ipsilon (Hufnagel) (Lepidoptera: Noctuidae) due to H. bacteriophora. However, S. carpocapsae caused 80–85% mortality in C. septempunctata, C. maculata, and O. vnigrum which was still lower than the A. ipsilon mortality (100%). Shapiro and Cottrell (2005) predicted that EPNs applications have significantly less impact on lady beetle populations (relative to impacts on the target pest) in crop fields than suggested by studies in the laboratory or in greenhouses. Their reason for this prediction was that contact between nematodes and lady beetles would be limited

Insect Order Coleoptera

Coleoptera

Coleoptera

Coleoptera

EPNs species H. bacteriphora NC strain and S. carpocapsae All strain

S. carpocapsae

S. carpocapsae (Cxrd strain) and H. bacteriophora (VS strain),

S. feltiae, S. carpocapsae, and H. bacteriophora

Non-target insects Ground beetles, Harpalus sp. and Pterostaticus sp.; Cicindela sp., Tetracha sp., and Philonthus sp. Adults Carabid beetles, Bembidion properans Steph. and Pterostichus cupreus L. Beetles Coleomegilla maculate, Olla v-nigrum, Harmonia axyridis, and Coccinella septempunctata Larvae Two spotted lady beetle, Adalia bipunctata 0; 1.5 × 104; 3 × 104; 6 × 104 IJs

250 IJs/beetle

50, 250, and 500 IJs/larva

F

L

L

Laboratory (L)/Field (F)/ Greenhouse Insect life (G) Dose stage 20 nematodes Last larval or L per cm2 or 400 nymphal stages and IJs per larva/ adults adult

Table 9.2  Examples of virulence of EPNs towards non-target arthropods

Shapiro and Cottrell (2005)

Ropek and Jaworska (1994)

References Georgis et al. (1991)

(continued)

Rojht et al. Mortality rate at 25 °C was (2009) >93%; Infectivity of EPNs species increased with increasing temperature; H. bacteriophora was found to be least effective and other species and mixed suspension of species showed almost the same effects

Mortality was higher in lepidopterous host, Agrotis ipsilon than all 4 lady beetles;

No effect found

Comments Adults possessed low susceptibility to nematodes; Mortality rate was 25–44% in late instars

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S. carpocapsae “Mexican” strain

C. diaphaniae parasitized melonworm larvae

L

L

Eulophid parasitoid Larvae and adults wasp, Diglyphus begini

Hymenoptera Cardiochiles diaphaniae

L

Seven spotted lady beetle, Coccinella septempunctata

Coleoptera S. carpocapsae (BA2), S. carpocapsae (S2), H. sp. (D1), S. feltiae (SF), S. carpocapsae (All), S. riobravae (SR), S. scabtarsci (SS), S. glaseri (SG), H. bacteriophora (HP88), and H. marilatus (MAR) S. carpocapsae Hymenoptera

Insect life stage Larvae, pupae and adults

0, 5, 25, 50, and 250 IJs per larva and 0 to 320 nematodes/ adult 200 IJs/ larva

50, 25 and 12.5 IJs/larva

Laboratory (L)/Field (F)/ Greenhouse (G) Dose L 75, 150, 300 and 600 IJs/ insect stage

2nd instar larvae

Non-target insects Carabid beetle Calosoma granulatum

EPNs species Insect Order Coleoptera H. amazonensis isolate RSC 5 and H. amazonensis isolate JPM 4

Table 9.2 (continued)

Metwally et al. (2016)

References Mertz et al. (2015)

Only 0–10% immature parasitoids parasitized by nematodes within the hosts. Indirectly, affected the emergence rate of parasitoid in the host

Shannag and Capinera (2000)

100 per cent mortality of wasp Sher et al. (2000) larvae after 48 h; Adults showed no infection

Comments Only 1st instar predator larvae susceptible to EPNs at concentrations greater than 150 IJs; EPN isolates were found to be safer for predator S. feltiae (SF), S. carpocapsae (all), S. riobravae (SR), H. bacteriophora (HP88), S. carpocapsae (BA2), and S. scabtarsci (SS) caused 50–100% mortality; H. bacteriophora was least effective;

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Hymenoptera Bumblebee, Commercial products Bombus terrestris containing mixture of Steinernema and Heterorhabditis sp. and S. Kraussei S. feltiae, S. carpocapsae Neuroptera Green lacewing and H. bacteriophora (Chrysoperla carnea)

S. thermophilum, S. glaseri, and H. indica

L

Larva

50 IJs, 250, and 500 IJs/ larva

10, 25, and 50 IJs/cm2 of treatment box

100 IJs/ant

L

L

10 IJs/cm2 (152 IJs)

L

Adult bees

Larvae Hymenoptera Ichneumonid, Mastrus ridibundus and Liotryphon caudatus Hymenoptera Ants, Messor Adults himalayans

S. carpocapsae (Sal)

Insect life stage Worker and drone bees

Insect Order Non-target insects Hymenoptera Apis mellifera mellifera

EPNs species S. affinis, S. feltiae

Laboratory (L)/Field (F)/ Greenhouse (G) Dose 9–10 IJs In-situ in brood combs and L

Dutka et al. (2015)

Anes and Ganguly (2015)

Lacey et al. (2003)

References Zoltowska et al. (2003)

(continued)

Rojht and At 15 °C, S. feltiae was most virulent; at 20 °C and 25 °C, S. Trdan (2007) carpocapsae and two mixed suspensions (S. carpocapsae × S. feltiae and S. carpocapsae × H. bacteriophora) showed higher infectivity

Comments 20–71.4% mortality in both sexes; S. feltiae more invasive than S. affinis especially against worker bees. Reduction in protein content of worker bees larvae 70.7 and 85.2% mortality in Mastrus ridibundus and Liotryphon caudatus, respectively S. thermophileum (68–100% mortality) > H. indica (40–100% mortality) > S. glaseri (24–72% mortality) Both products caused very high mortality after only 72 h exposure, with the first deaths evident after 48 h

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EPNs species S. carpocapsae (BA2), S. carpocapsae (S2), H. sp. (D1), S. feltiae (SF), S. carpocapsae (All), S. riobravae (SR), S. scabtarsci (SS), S. glaseri (SG), H. bacteriophora (HP88), and H. marilatus (MAR) S. carpocapsae (BA2), S. carpocapsae (S2), H. sp. (D1), S. feltiae (SF), S. carpocapsae (All), S. riobravae (SR), S. scabtarsci (SS), S. glaseri (SG), H. bacteriophora (HP88), and H. marilatus (MAR) H. bacteriphora NC strain and S. carpocapsae all strain, and S. feltiae SN strain

Table 9.2 (continued)

Non-target insects Green lacewing, Chrysoperla carnea

Minute pirate bug, Orius albidipennis

Earwig, Labidura riparia

Insect Order Neuroptera

Hemiptera

Dermaptera

L

Last larval or L nymphal stages and adults

Immature and adults

Insect life stage 2nd instar larvae

S. carpocapsae caused 40% mortality; all other species were not virulent.

Comments S. carpocapsae (All) was the most effective with 55–88% mortality; Heterorhabditis sp. D1 was least effective (38–73% mortality)

20 nematodes No mortality observed per cm2 or 400 IJs per larva/ adult

50, 25 and 12.5 IJs/ml

Laboratory (L)/Field (F)/ Greenhouse (G) Dose L 50, 25 and 12.5 IJs/larva

Georgis et al. (1991)

Metwally et al. (2016)

References Metwally et al. (2016)

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Non-target insects Aphid predator, Aphidoletes aphidimyza

Spider, Neoscona theisi

Insect Order Diptera

Arachnids (Araneae)

EPNs species S. carpocapsae, S. feltiae, and H. bacteriophora

S. thermophilum, S. glaseri, and H. indica

Adults

Insect life stage 3rd instar larvae

L

100 and 500 IJs/adult

Laboratory (L)/Field (F)/ Greenhouse (G) Dose L and G 10, 100, and 1000 IJs/ larva in laboratory and 100 and 1000 IJs per/ larva in greenhouse Comments Mortality caused by all 3 species but comparatively less due to S. feltiae in laboratory; H. bacteriphora, not S. carpocapsae reduced pupal emergence; in greenhouse, adult emergence was reduced by both S. carpocapsae and H. bacteriophora; mortality (5–81%) in greenhouse was lower than laboratory mortality (8–93%) Maximum of 30, 20, and 10% mortality caused by H. indica, S. thermophilum, and S. glaseri, respectively Anes and Ganguly (2015)

References Powell and Webster (2004)

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because lady beetles spend their time looking for hosts on the crop foliage, while EPNs would be active only in the soil and were present briefly at most on foliage, at the time of treatment. Foltan and Puza (2009) also found that the carabid Pterostichus melanarius Illiger (Carabidae) avoided feeding on prey infected with Phasmarhabditis hermaphrodita Schneider (Rhabditidae) and Steinernema affine (Bovien) (Steinernematidae). The presence of either of the two nematodes deterred the beetles from consuming infected carabid cadavers. Mertz et al. (2015) reported that first instar larvae of Calosoma granulatum Perty (Carabidae) were susceptible (70% mortality) to two strains of H. amazonensis when this EPN was applied topically at concentrations greater than 150 IJs/ml. Mertz et al. (2015) also found a repellent effect of H. amazonensis infection (in the larvae of noctuid Spodoptera frugiperda Smith (Lepidoptera: Noctuidae) on feeding by third instar larvae or adults of C. granulatum. Larvae of C. granulatum that did feed on noctuids infected by H. amazonensis, however, died, although H. amazonensis had been reported to be safe to C. granulatum (Mertz et al. 2015). However, Metwally et al. (2016) observed that S. carpocapsae (All) was highly virulent (causing 72–100% mortality) to the larvae of the seven spotted lady beetle, C. septempunctata, under laboratory conditions. There are very few studies reporting the possible effects of EPNs on non-target beetles under field conditions. Ropek and Jaworska (1994) did not observe any mortality under field conditions to the carabid beetles Bembidion properans Stephens or Pterostichus cupreus L. (Carabidae) when fields of annual legumes were treated with S. carpocapsae. Similarly, Dillon et al. (2012) found no effects of S. carpocapsae on the richness or abundance of 65 species of non-target beetles, including 11 saproxylic species in a forest ecosystem. In contrast, reports suggest that field applications of H. megidis can reduce the population densities of the weevil Barypeithes sp. (Curculionidae) and that applications of S. feltiae reduced numbers of four non-­ target chrysomelids or carabids (Buck and Bathon 1993; Koch and Bathon 1993).

9.4.2  Hymenoptera EPNs have also been suggested as a potential threat to the pollinator bees. In principle, adult bees might be infected by EPNs while foraging on flowers for nectar or pollen (Morse and Flotum 1997). Some pollinator bees such as bumble bees (Bombus spp.) are reported to be hibernating under rotten tree stumps, piles, leaf litter, and soil (Alford 1969) which might be a route in EPN infection. Kaya et al. (1982) observed that larvae and adults of honey bees, Apis mellifera L. (Apidae), were susceptible to EPN nematodes, but EPNs did not adversely affect the whole colonies. Nguyen and Smart (1991) reported low mortality in honey bees due to Steinernema scapterisci. Zoltowska et al. (2003) observed sensitivity of honey bee

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larvae to infection by S. affinis and S. feltiae in both combs and petri dishes. Similarly, Dutka et al. (2015) found high mortality in Bombus terrestris L. (Apidae) due to two commercially available EPN products containing mixture of Steinernema and Heterorhabditis sp. and S. kraussei under laboratory conditions. Taha and Abdelmegeed (2016) observed no mortality in honey bees, A. mellifera, when hives were treated with H. bacteriophora to control wax worms, Galleria mellonella L. (Lepidoptera: Pyralidae). Shannag and Capinera (2000) reported the direct infection of hymenopteran parasitoid, Cardiochiles diaphaniae Marsh (Braconidae) by S. carpocapsae after they have emerged from infected hosts, melonworm, Diaphania hyalinata L., and pickleworm, D. nitidalis Stoll (both Lepidoptera: Crambidae), but before completion of the pupal cocoon. Similarly, Lacey et al. (2003) reported infection of developing larvae of two ectoparasitic ichneumonids, Mastrus ridibundus Gravenhorst and Liotryphon caudatus Ratzeburg (Ichneumonidae) by S. carpocapsae due to direct application in laboratory experiments. The potential compatibility of application of EPNs with parasitoid populations was observed because both parasitoids were able to avoid nematode-infected larvae and, in contrast, focused their oviposition on healthy cocooned host larvae of the codling moth, Cydia pomonella L. (Lepidoptera: Tortricidae) that escaped the nematode infection. Similarly, the eulophid parasitoid Diglyphus begini Ashmead (Eulophidae) avoided ovipositing in larvae of the agromyzid leafminer Liriomyza trifolii Burgess (Diptera: Agromyzidae) infected with S. carpocapsae (Sher et al. 2000). Anes and Ganguly (2015) found that the EPNs, S. thermophilum, S. glaseri, and H. indica were lethal to the beneficial ant Messor himalayanus Forel (Formicidae), causing 72–100% mortality under laboratory conditions, when applied topically.

9.4.3  Neuroptera Green lacewings (Chrysopidae) are predators of mites, aphids, whiteflies, scales, and other small insects. Some neuropterans spend some time in soil foraging for hosts, potentially bringing them into contact with EPNs. Rojht and Trdan (2007) studied the effects of three EPNs (S. feltiae, S. carpocapsae and H. bacteriophora) individually and in mixed suspensions on larvae of green lacewing, Chrysoperla carnea Stephens (Chrysopidae) at different temperatures in the laboratory with three doses: 50, 250, and 500 IJs/ larva. Steinernema feltiae was found to be virulent at 15 °C. At 20 and 25 °C, S. carpocapsae and two mixtures (S. carpocapsae × S. feltiae and S. carpocapsae × H. bacteriophora) caused the highest mortality in C. carnea. Metwally et al. (2016) reported high rates of mortality (30–100%) in 2nd instar larvae of C. carnea by different EPN strains at 50, 25 and 12.5 IJs/larva doses, under laboratory conditions.

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9.4.4  Hemiptera The minute pirate bug Orius albidipennis Rueter (Anthocoridae) is an insect predator in many agricultural crops. The nymphs and adults feed on a wide range of arthropods including aphids, chinch bugs, springtails, plant bugs, thrips, eggs and small larvae of corn earworms, whiteflies, spider mites. The adults of minute pirate bugs overwinter in plant debris or leaf litter that may bring them into contact with EPNs. Metwally et  al. (2016) reported S. carpocapsae as the most virulent EPN tested against O. albidipennis.

9.4.5  Diptera The cecidomyiid fly Aphidoletes aphidimyza Rondani (Cecidomyiidae) is an aphid predator that is a commercially available biological control agent, used in both greenhouses and orchards. The larvae of this predator drop from the plant foliage to the soil, where they pupate. The pupae will wiggle to the soil surface after emergence from the cocoon and emerge as adults. This time in soil provides opportunity for contact with EPNs. Powell and Webster (2004) observed mortality of A. aphidimyza following treatment with S. carpocapsae, S. feltiae, and H. bacteriophora. However, the levels of mortality found in a greenhouse study was lower than that in the laboratory test.

9.4.6  Other Arthropods The susceptibility of some isopods, diplopods, symphylans, and chelicerates to EPNs has been reviewed (Poinar 1989). Steinernema carpocapsae (DD 136) can infect the symphylan Scutigerella immaculata Newport (Symphyla: Scutigerellidae) under laboratory conditions (Swenson 1966). As symphylans live in the soil litter, there might be possibility of contact between them and EPNs. Similarly, the terrestrial isopods Porcellio scaber Latreille (Isopoda: Porcellionidae) and Armadillidium vulgare Latreille (Isopoda: Armadillidiidae) were susceptible to S. feltiae infection under laboratory conditions (Poinar and Paff 1985). The tick Boophilus annulatus Say (Acari: Ixodidae) was susceptible to S. carpocapsae under laboratory conditions. But no development of S. carpocapsae occurred in the cadavers of infected females (Samish and Glazer 1992; Glazer and Samish 1993). The nematode S. feltiae when applied at 80 nematodes/cm2 of soil reproduced successfully in the sowbug Porcellio scaber Latr. and in the millipede Blaniulus guttulatus Fabricius

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(Julida: Blaniulidae), under greenhouse conditions (Jaworska 1994). Zhioua et al. (1994) found penetration by S. carpocapsae and subsequent mortality of female ticks, Ixodes scapularis Say (Acari: Ixodidae). However, most of these studies were conducted under laboratory conditions in which high contact between nematodes and the target organism was assured, under highly favorable physical conditions. Fewer field studies on the above mentioned groups have been reported. Rahayu (1983) found reduction in numbers of the Collembolan Onychiurus armatus Tullberg (Collembola: Onychiuridae) in both field and laboratory studies from infection by S. carpocapsae. Similarly, Edwards and Oswald (1981) reported suppression of Collembola and mites by S. carpocapsae in sugar beet fields. In contrast, Georgis et  al. (1991) did not found any effects on non-target arthropods, including species of Gryllidae, Collembola, Gamasida, Actinedida, and Oribatida from exposures to S. carpocapsae, S. feltiae, or H. bacteriophora. The immature stages and adults of the earwig Labidura riparia Pallas (Dermaptera: Labiduridae) were found to be resistant to H. bacteriophora NC strain and S. carpocapsae infection, when treated in the laboratory with 400 IJs per larva or adult (Georgis et al. 1991). Wang et al. (2001) found the long term effect of H. bacteriophora, S. carpocapsae, and S. riobravis to be much less severe in Lycosidae, Staphylinidae, Araneae, Formicidae, Scelionidae, and Sminthuridae than the effect of the insecticide chlorpyriphos in turf grass, with the degree of impact varying considerably among non-­ target arthropod groups. Indeed, the population levels of oribatid mites were significantly higher in EPN treated plots than in untreated controls (Wang et  al. 2001). However, Peck (2009) found reduction in the abundance of insects and Collembola in plots of turf grass treated with H. bacteriophora. However, the apparent reduction of these aggregated groups was not detected when data were broken apart into more specific taxonomic groups, due to complications related to the tremendous amount of microarthropods collections produced by sampling. Anes and Ganguly (2015) observed 10–30% mortality in the spider Neoscona theisi Walckenaer (Araneae: Araneidae) following application of S thermophilum, S. glaseri, and H. indica under laboratory conditions. However, it is important to notice the possible lack of exposure of spiders to EPNs due to the spider habitat characteristics. Some other researchers have also found negative impacts of EPNs on terrestrial isopods, diplopods, and ticks under laboratory conditions (Jaworska 1991; Mauleon et al. 1993; Hill 1998). However, an increase in the population of isotomid collembolans, anysitid mites, and gnaphosid spiders under trees after nematode application was found. The increase in the anysitid mite population was apparently related to the increased foraging activity by the mite in the nematode-treated areas. The anysitid mites are important natural enemies in orchards, feeding on thrips, collembola, and nematodes (Goh and Lange 1989; Cuthbertson and Murchie 2004).

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9.5  Intraguild Predation Most of the above studies have focused on the direct effects of the nematode/bacteria complex on non-target speciess. EPNs can directly reduce the populations of some non-target organisms through significant non-target mortality. However, there are some indirect effects such as Intraguild Predation (IGP) that can affect the population densities of non-target organisms that ultimately minimize their input in biological control. Intraguild predation is defined as two species sharing a common host and engaging in predation or parasitism of each other (Holt and Polis 1997). IGP is relatively common among agents used for the biocontrol of arthropod pests, as compared to agents used against weeds or plant pathogens (Rosenheim et  al. 1995). The simultaneous application of EPNs with other types of biocontrol agents such as predators and parasitoids place large populations of both agents together, setting up conditions for IGP. Akhurst (1990) reported that when biocontrol agents such as parasitoids and entomopathogenic nematodes co-infect an insect, the nematodes either can infect immature parasitoids or inhibit their development. Factors such as the exact insect host, the parasitoid species, the host specificity of the nematode, and the relative timing of co-infestation of the agents can all affect these interactions. Predators can also be affected, as they can become infected with nematodes by eating a nematode-infected prey. IGP interactions have been described by Georgis and Hague (1982) and Battisti (1994) for Steinernema spp. interacting with sawflies and their parasitoids (ichneumonids). In both studies, the survival rate of parasitoids was lower in nematode-­ treated plots as compared to control. No infection of parasitoid larvae was found inside hosts (Battisti 1994). The parasitoid larvae inside cocoons were less susceptible to nematodes than the parasitoid parasitizing the sawfly in the laboratory tests (Georgis and Hague 1982). Sher et al. (2000) examined the interaction between S. carpocapsae and the eulophid parasitoid wasp, D. begini for the control of leafminer, L. trifolii on chrysanthemum. It was found that parasitoids were affected by nematodes either indirectly through host mortality or directly through nematode infection of the parasitoid larvae. The nematodes attacked paralyzed (by the parasitoid) leafminer larvae and healthy (non-parasitized) leafminer larvae equally, which suggested that behavioral avoidance by the nematode did not reduce infection of healthy leafminer larvae. Sher et al. (2000) also found that the total mortality of leafminers was higher when both agents were used together, than when either agent was used alone. Similarly, Powell and Webster (2004) studied the effects of S. carpocapsae, S. feltiae, and H. bacteriophora on the aphid predator A. aphidimyza. The adult emergence was reduced by H. bacteriophora and S. carpocapsae, but not by S. feltiae. Pupae of A. aphidimyza within their cocoons were non-susceptible to S. carpocapsae. The predator inhibited the development of EPNs and ultimately killed them in the host because of complete destruction of the host body, and its internal desiccation due to holes opened in the integument (Foltan and Puza 2009).

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9.6  Conclusion and Future Prospects EPNs have potential to control many pest arthropods. Some reports concerning effects of EPNs, their symbiotic bacteria or their metabolites on non-target species under natural conditions do exist. However, most of these studies were laboratory bioassays, with only a few studies evaluating field effects on non-target species, indicating a need for more field-based studies. EPN-bacterium complexes can affect non-target species either through direct infection or indirectly through effects such as depletion of the target host populations. The careful selection of host specific EPN strains for the target pests might help minimize the direct effects. While selecting any EPN species for use in a classical or inundative biological control program, the risk to non-target organisms like depletion of host food resources, competition with other pests in regard to scarcity of food in presence of EPNs, should always be considered. As discussed above, when EPNs compete with parasitoids inside hosts, death of immature parasitoids may occur. The degree of non-target impact will also depend on environmental factors that affect the EPN efficacy and persistence in the area of application. The use of pathogens that are highly virulent to some non-target organisms may not have these effects, if exposure of non-target species to the nematodes is unlikely to occur or limited in area. For example, chances of infection of adult parasitoids by EPNs is minimal because they mostly occur in above ground habitats, eliminating most contact chances with nematodes. Some factors affecting the degree of potential non-target risks posed by use of EPNs in biological control include the following: 1. Host Range: The host range plays an important role in the selection of EPN strains. In the past, most failures in use of EPNs in field are related to a poor match between the EPN species and target insect pest. It is a challenge to maintain balance between host specificity and broad host range for safety of non-­ target organisms. Steinernema scapterisci and S. neocurtillae Nguyen and Smart are host-specific species that have few, if any, non-target effects. However, broad host ranges are commercially desirable for nematodes marketed for application as pesticides. 2. Dose: The dose to be used in field should be determined in the laboratory first. Excessively high doses may be both costly and increase risk of non-target impacts. 3. Time of application: Strategies such as the avoidance of coincidence of EPN application with the peak activity of non-target organisms will reduce the chances of intraguild predation, that will indirectly limit the chances of migration of adult predators to other areas with more prey options, thus limiting the local extinctions of the natural enemies. 4. Dispersal: Harm to certain non-target organisms may be limited or avoided if exposure of the non-target species to the pathogen is outside the EPNs dispersal distance and beyond the treated area.

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5. Persistence: The long term persistence of particular EPNs must be determined as part of the risk assessment. The best example here would be the introduction of S. glaseri, which was applied on a large scale in New Jersey in 1939–42 to control Japanese beetle. This nematode could still be found after 50 years, but only at few locations and in the southern part of the area where nematodes were applied (Gaugler et al. 1992). However, along with requirement of persistence of these EPNs, non-target risks associated with this cannot be avoided. Little literature was found on possible negative effects on non-target species of EPN symbiotic bacteria when separated from their respective EPNs. These symbiotic bacteria require nematodes as carrier for their survival. Therefore, the possibility of establishment of these bacteria in absence of their particular host nematodes in natural conditions is highly limited. There is, however, still a need to study the virulence of these bacteria to non-target organisms in the field, without their EPN hosts. The formulation of symbiotic bacteria without nematodes, the persistence of such bacteria, their shelf life, production and application methods, and the effects of biotic and abiotic factors on these bacteria are all aspects that need to be explored and studied in detail for interpreting potential non-target effects. EPNs that show high virulence to non-target arthropods in laboratory studies may have much lower, or no, effects under natural conditions due to the lack of contact between the non-target species and the nematodes. Because the chances of direct contact between nematodes and organisms are maximized in laboratory tests, it is to be expected that impacts under field conditions will vary, and likely be lower. Bathon (1996) noted that mortality from EPNs will be temporary, spatially restricted, and will affect only part of a population. The physiological host range does not necessarily equal ecological host range of the entomopathogens (Hajek and Goettel 2000). The efficacy of EPNs does not only depend on the infectivity but is also affected largely because of the environmental factors surrounding the EPNs and the potential hosts. The host range of EPNs under favorable conditions i.e. adequate moisture, temperature and no effect of UV radiation will always be different from their ecological host range. The ecological host range of EPNs in natural environment can be determined after the physiological host range is known from laboratory studies. But in laboratory, it is difficult to maintain the ecological conditions as similar to field conditions and it is not reasonable to apply the laboratory results in field conditions. According to Bathon (1996), the susceptibility of hosts to EPNs in the field might be related to habitat requirements and searching behavior of nematode strains. This raises questions about the reliance only on data from laboratory studies to predict field non-target impacts. However, laboratory observations can be an important beginning for evaluating the potential effects of these nematodes in natural populations. The laboratory and semi-field studies should be conducted in a way to mimic the field conditions as much as possible. In-depth knowledge of field effects of these parasitic nematodes is required to assess the negative effects. Overall, the long-term impact of the EPNs on non-target arthropods is much less severe than that of chemical pesticides, with varying degree of impact among

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n­ on-­target arthropods. Current literature as a whole suggests that non-target effects of EPNs are negligible. Acknowledgements  This material is based upon work supported by the National Institute of Food and Agriculture, U.S.  Department of Agriculture, Multistate-W4185 project “Biological Control in Pest Management Systems of Plants” (Accn#1014043; Hatch Project#MONB00857).

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Rajagopal, R., & Bhatnagar, R. K. (2002). Insecticidal toxic proteins produced by Photorhabdus luminescens akhurstii, a symbiont of Heterorhabditis indica. Journal of Nematology, 34, 23–27. Razek-Abdel, A. S. (2003). Pathogenic effects of Xenorhabdus nematophilus and Photorhabdus luminescens (Enterobacteriaceae) against pupae of the Diamondback Moth, Plutella xylostella (L.). Journal of Pest Science, 76, 108–111. Rojht, H., & Trdan, S. (2007). Non-target effects of entomopathogenic nematodes on natural enemies: Example on (with) green lacewing (Chrysoperla carnea Stephens, Neuroptera, Chrysopidae) Zbornik predavanj in referatov, 8. Slovenskega postvetovanja o varstvu Rastlin, Radenci, Slovenija, 6–7 Marec, 2007. pp. 118–125. Rojht, H., Kac, M., & Trdan, S. (2009). Nontarget effect of entomopathogenic nematodes on larvae of twospotted lady beetle (Coleoptera: Coccinellidae) and green lacewing (Neuroptera: Chrysopidae) under laboratory conditions. Journal of Economic Entomology, 102, 1440–1443. Ropek, D., & Jaworska, M. (1994). Effect of an entomopathogenic nematode, Steinernema carpocapsae Weiser (Nematoda, Steinernematidae), on carabid beetles in field trials with annual legumes. Anz Sch~idlingskde, Pflanzenschutz, Umweltschutz, 67, 97–100. Rosenheim, J. A., Kaya, H. K., Ehler, L. E., Marois, J. J., & Jaffee, B. A. (1995). Intraguild predation among biological-control agents: Theory and evidence. Biological Control, 5, 303–335. Samish, M., & Glazer, I. (1992). Infectivity of Entomopathogenic Nematodes (Steinemematidae and Heterorhabditidae) to female ticks of Boophilus annulatus (Arachnida: Ixodidae). Journal of Medical Entomology, 29, 614–618. Shannag, H., & Capinera, J.  (2000). Interference of Steinernema carpocapsae (Nematoda: Steinernematidae) with Cardiochiles diaphaniae (Hymenoptera: Braconidae), a parasitoid of Melonworm and Pickleworm (Lepidoptera: Pyralidae). Environmental Entomology, 29, 612–617. Shapiro-Ilan, D. I., & Cottrell, T. E. (2005). Susceptibility of lady beetles (Coleoptera: Coccinellidae) to entomopathogenic nematodes. Journal of Invertebrate Pathology, 89, 50–156. Sher, R. B., Parrella, M. P., & Kaya, H. K. (2000). Biological control of the leafminer Liriomyza trifolii (Burgess): Implications for inraguild predation between Diglyphus begini Ashmead and Steinernema carpocapsae (Weiser). Biological Control, 17, 155–163. Shrestha, S., & Kim, Y. (2010). Differential pathogenicity of two entomopathogenic bacteria, Photorhabdus temperata subsp. temperata and Xenorhabdus nematophila against the red flour beetle, Tribolium castaneum. Journal of Asia-Pacific Entomology, 13, 209–213. Stock, S. P., & Goodrich-Blair, H. (2008). Entomopathogenic nematodes and their bacterial symbionts: The inside out of a mutualistic association. Symbiosis, 46, 65–75. Stock, S. P., & Hunt, D. J. (2005). Nematode morphology and systematics. In P. S. Grewal, R. U. Ehlers, & D.  I. Shapiro-Ilan (Eds.), Nematodes as Biological Control Agents (pp.  3–43). Wallingford: CAB International Publishing. Swenson, K.  G. (1966). Infection of the garden symphylan, Scutigerella immaculata, with the DD-136 nematode. Journal of Invertebrate Pathology, 8, 133–134. Taha, E. H., & Abdelmegeed, S. M. (2016). Effect of entomopathogenic nematodes, Heterorhabditis bacteriophora, on Galleria mellonella in bee hives of Apis mellifera. Journal of Biological Sciences, 16, 197–201. Tailliez, P., Pages, S., Edgington, S., Tymo, L.  M., & Buddie, A.  G. (2012). Description of Xenorhabdus magdalenensis sp. nov., the symbiotic bacterium associated with Steinernema australe. International Journal of Systematic and Evolutionary Microbiology, 62, 1761–1765. Uma, G.  P., Prabhuraj, A., & Vimala, S. (2010). Bioefficacy of Photorhabdus luminescens, a symbiotic bacterium against Thrips palmi Karny (Thripidae: Thysanoptera). Journal of Biopesticides, 3, 458–462. Van Lenteren, J.  C., Babendreier, D., Bigler, F., Burgio, G., Hokkanen, H.  M. T., Kuske, S., Loomans, A.  J. M., Menzler-Hokkanen, I., van Rijn, P.  C. J., Thomas, M.  B., Tommasini, M. G., & Zeng, O. (2003). Environmental risk assessment of exotic natural enemies using in inundative biological control. BioControl, 48, 3–38.

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Chapter 10

Granuloviruses in Insect Pest Management Pankaj Sood, Amit Choudhary, and Chandra Shekhar Prabhakar

Abstract Alternative strategies for insect pest management were looked and microbial pesticides in particular were noticed as attractive candidates. Among baculoviruses, the granulosis viruses are highly specific and safer to non target species. They are considered potential candidates for use as biological insecticides for control of economically important insects. These viruses are highly virulent, selective, stable and environmentally benign, once applied. However, usage and availability of granuloviruses is limited, worldwide. Slow action, restricted host range and low persistence are some of major drawbacks of such microbial pesticides, hindering their large scale usage. Methods must be developed for the unequivocal identification of these viruses. Their effects on non target species must be investigated at the cellular and molecular levels to enhance their role in pest management programmes. Efforts for identifying potent microbial agents, based on their rich biodiversity, must be applied extensively to expand the genetic make-up of baculoviruses. Recombinant DNA technology in baculoviruses could result in accomplishment of many achievements. In order to increase the uptake of baculoviruses, it is necessary to develop robust molecular biology techniques, to enhance safe food production for the nutritional and health security of growing population. Keywords  Biocontrol · GV · Baculovirus · Insect pest management · Organic agriculture

P. Sood (*) Krishi Vigyan Kendra, CSK Himachal Pradesh Krishi Vishvavidyalaya, Sundernagar, Mandi, Himachal Pradesh, India e-mail: [email protected] A. Choudhary Department of Entomology, Punjab Agricultural University, Ludhiana, Punjab, India C. S. Prabhakar Department of Entomology, Veer Kunwar Singh College of Agriculture, Dumraon (Bihar Agricultural University, Sabour), Buxar, Bihar, India © Springer Nature Switzerland AG 2019 M. A. Khan, W. Ahmad (eds.), Microbes for Sustainable Insect Pest Management, Sustainability in Plant and Crop Protection, https://doi.org/10.1007/978-3-030-23045-6_10

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10.1  Introduction In the recent years, biological control strategies have gained importance for the management of agricultural pests. This shift occurred due to the repeated failure of synthetic pesticides in managing agricultural pests, thereby causing several outbreaks. The growing knowledge and awareness on the negative impacts of chemical insecticide usage on the ecosystem and public health led to efforts directed towards a reduction in chemical control of insect pests, diseases and weeds (Haase et  al. 2015). Along with these issues, there are also the problems pertaining to target and non-target insect species. At the same time, human population is growing at a very fast rate. To fulfill the nutritional and other requirement of such a huge population we have to sustain an even more intensive agriculture. In such system, pest problems will continue to be a challenging factor. Hence, to operate such system in which synthetic pesticides have to be lowered without any negative effects on yields, we have to look for viable alternatives. These must confer the idea of sustainable crop yield by significantly managing the pest populations, helping in preserving the ecosystem health. Under the above situations, we look towards biological control of agricultural pests. These will, individually or in integration with other pest management strategies, help us in attaining the above mentioned objectives. Furthermore, under the concept of biocontrol we have to consider many agents such as macro and microorganisms. Among the microorganisms category, many entomopathogens can play a key role. Under natural conditions the populations of many pests are regulated by epizootics caused by various entomopathogens such as bacteria, fungi and viruses. Several examples are available showing how entomopathogens have been deployed successfully under classical biological control strategies, for management of insect-­ pests (Kalha et al. 2014). Among these, insect pathogenic viruses are promising and crucial components, devising suitable integrated pest management strategy. Viruses of only a few families are known to infect insects. Entomopathogenic viruses from a single family only, i.e. Baculoviridae, have been explored as biopesticides thus far. Insects belonging to orders Lepidoptera, Hymenoptera and Diptera are natural hosts for this virus family. These entomoptahogens are globally well known for application as biopesticides, to control a number of important insect pest populations of crops and forests (Sun and Peng 2007). Thus, their use represents an alternative and eco-friendly approach for pest management. They are safe to human health and the ecosystem, allowing innovative industrial productions and field applications (Sumathy et  al. 1996). Betabaculovirus, one of the four genera of family Baculoviridae, is a Lepidopteran-specific granulovirus (GV). The other three genera of Baculoviridae are Alphabaculovirus (Lepidopteran-specific nucleopolyhedrovirus), Gammabaculovirus (Hymenopteran-specific nucleopolyhedrovirus), and Deltabaculovirus (Dipteran-specific nucleopolyhedrovirus) (King et al. 2011).

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Baculoviruses have rod-shaped nucleocapsids, each comprising a single circular genome. They are enveloped, singly or in groups, by a membrane to form virions. Virions are occluded in a protein matrix, which forms the occlusion body (OB) (Williams et  al. 2017). The OB protects the virions, known as occlusion derived virions (ODVs), from environmental factors. These viruses can be readily distinguished into two groups: nucleopolyhedroviruses and granuloviruses that clearly differ in the structure of their OBs (Adams and McClintock 1991). Nucleopolyhedroviruses have polyhedral OBs (0.5–10 μm), mainly comprising of crystalline polyhedrin protein, which occludes large numbers of virions. In contrast, granuloviruses have smaller granule-like OBs (∼0.4  μm), that mainly comprise granulin protein. Each granulovirus OB contains a single virion (Williams et  al. 2017). Baculoviridae (Nucleopolyhedrosis and Granulosis viruses) are exploited for the development of commercial virus-based biopesticides for insect pest management. In the present chapter, we discuss the ecology and use of granulosis viruses in the management of insect pests.

10.2  Classification of Entomopathogenic Virus Earlier, viruses were classified using a Linnaean (Binomial) taxonomy, initially based on morphological characteristics and then stretched to genomic phylogeny with the development of modern molecular biology tools. Actual classification of viruses, known as the Baltimore classification, is based on nucleic acids and on the function of capsid (Sparks 2010). This classification separates viruses into seven classes based on the type of nucleic acids present in the capsid that represent the viral genome, and on the way the mRNA is coded, for protein production in the cell (Baltimore 1971). The Baltimore classes are as follows: I. double strand, dsDNA viruses (herpesviruses, baculoviruses, poxviruses) II. single strand, ssDNA viruses (+) sense DNA (parvoviruses) III. dsRNA viruses (reoviruses) IV. (+) ssRNA viruses (picornaviruses, togaviruses) V. (−) ssRNA viruses (orthomyxoviruses, filoviruses) VI. ssRNA-RT viruses (positive sense RNA with a DNA intermediate such as retroviruses) VII. dsDNA-RT (DNA viruses that utilize reverse transcription such as hepadnaviruses) In view of the general approval accorded by virologists to the uniform taxonomy of viruses, insect virus classification also follows the direction of the International Committee on Taxonomy of Viruses (ICTV) (Van Regenmortel et  al. 2000). The general characters considered in the classification of any virus include the type of genetic material (single- or double-stranded DNA, single- or double-stranded RNA, positive or negative strand), virion morphology and size (icosahedral, rod-shaped),

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the presence of an envelope surrounding the virion, and of an occlusion body engulfing the virions, host and host range. The same criteria are followed to classify the diversity of viral groups that attack insect species. Insect pathogenic viruses are named with acronyms, according to their host and the viral group to which they belong to. For example, the cabbage butterfly, Pieris brassicae granulosis virus, is named PbGV.  Therefore, all nucleopolyhedrosis viruses are named NPV while the granulosis viruses are named GV. Further, lepidopteran baculoviruses were divided in NPVs (group I and II) and GVs on the basis of genomic nucleic acids sequence data (van Oers and Vlak 2007). The entomopoxviruses are named EPV, the iridoviruses as IV, and the cytoplasmic polyhedrosis viruses (cypoviruses) as CPV.

10.3  Granuloviruses of Important Insect Pests GVs have been reported only from insects belonging to the order Lepidoptera, except one doubtful case reported in insect belonging to the order Hymenoptera. Insects may acquire GV inoculum through feeding. The GV infected insect tissues eventually disintegrate and the body fluid got filled with granulin inclusions. In India, GVs have been reported to be effectively used in the management of Chilo infuscatellus (Snell.) (Easwaramoorthy and David 1979; Mala and Solayappan 2001), Chilo sacchariphagus (Bojer) (Easwaramoorthy and Jayaraj 1993; Easwaramoorthy and Santhalakshmi 2000), Spodoptera litura (Fab.) (Narayanan 1985), Phthorimaea operculella (Zeller) (Pokharkar et al. 1999; Chandla and Verma 2000) and Plutella xylostella L. (Subramanian 2002; Kennedy et  al. 2003). GV infecting Pieris brassicae L. have been reported for the first time in India from Himachal Pradesh by Sood (2004) and Narayanan and Sood (2006). Several GVs have been reported from India and abroad (Table 10.1), but only a few have been studied in details for their potential as biopesticides, in the management of economically important insect pests.

10.4  Mode of Action Insect viruses, as they occur in nature, consist of either double-stranded or single stranded circular/straight genomes of viral DNA/RNA with/without a double membrane envelop. Mode of action of baculoviruses is well studied. Baculoviruses show double stranded and circular viral DNA, with a double membrane envelop. They typically have narrow host ranges, often limited to just one or a few related insect species, although the most intensely studied member of the family, Autographa californica multiple nucleopolyhedrovirus (AcMNPV), is able to infect as many as 30 species from several lepidopteran genera. These occlusion-derived viruses (ODV) contain one or multiple nucleocapsids with each capsid holding a single

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Table 10.1  Granuloviruses reported from main insect pests S. No. 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 36. 37. 38. 39. 40.

Insect Species Agrotis ipsilon Agrotis segetum Achaea janata Adoxophyes orana Artogeia rapae Clostera anachoreta Clostera anastomosis Choristoneura fumiferana Choristoneura murinana Chilo infuscatellus Chilo saccharigus indicus Cnaphalocrocis medinalis Cnephasis pumicana Cryptophlebia leucotreta Cydia pomonella Dendrolimus sibiricus Diatraea saccharalis Erinnyis ello Epinotia aporema Harrisina brillians Helicoverpa armigera Homona magnanima Hyphantria cunea Lacanobia oleracea Mythimna unipuncta Pericallia ricini Pieris brassicae Phthorimaea operculella Plodia interpunctella Plutella xylostella Pieris rapae Pseudalatia unipuncta Scripophaga excerptalis Scrobipalpa absoluta Spodoptera litura Spodoptera exigua Spodoptera frugiperda Trichoplusia ni Xestia c-nigrum Zeiraphera diniana

GV acronyms AiGV AsGV AjGV AdorGV ArGV CaGV ClanGV ChfuGV ChmuGV CiGV CsGV CnmeGV CnpuGV CrleGV CpGV DesiGV DsGV ErelGV EpapGV HabrGV HearGV HomaGV HycuGV LaolGV MuGV PeriGV PbGV PhopGV PiGV PlxyGV PrGV PsunGV ScexGV SaGV SlGV SeGV SfGV TnGV XcGV ZdGV

References Chaudhry et al. (1976) Zethner (1980) Pawar (1980) Aizawa and Nakazato (1963) – – – – Cunningham (1982) Easwaramoorthy and David (1979) Mehta and David (1980) Jacob et al. (1971) Glas (1991) Fritsch and Hubner (1986) Benz (1981) Baranovsky and Litvina (1978) Pavan et al. (1983) – – – Gitay and Palson (1971) Sato et al. (1980) Weiser et al. (1986) Crook and Brown (1982) – Jacob et al. (1972) Billotti et al. (1956); Sood (2004) Chandla and Verma (2000) Hunter et al. (1977) Kao and Rose (1976) Jaques (1973) – Singaravelu et al. (1999) Lipa (1998) Pawar and Ramakrishnan (1977) Narayanan (2003) – – Lipa and Ziemnicka (1971) Baltensweiler et al. (1977)

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viral genome. One or more ODVs, in turn, are ‘occluded’ within a protein matrix, referred to as occlusion bodies, that protect the virus from the environment, as the the occlusion bodies eventually slowly degrade under UV light (Cory and Myers 2003; Harrison and Hoover 2012). Baculoviruses are transmitted orally in nature, and the target of primary infection is the larval midgut epithelium. After being consumed, occlusion bodies get dissolved in the midgut lumen, releasing the embedded ODV, which infect midgut epithelial cells (Clem and Passarelli 2013). The ODV bind and fuse with midgut cells, releasing the nucleocapsids into the cytoplasm. The nucleocapsids are transported to the nucleus, uncoat and begin replication, producing budded virus particles (Miller 1997; Elderd 2013). Progeny BV bud from the basal surface of the epithelium, cross the basal lamina, and infect most of the remaining tissues of the larva. The process of ODV attachment and entry depends on several viral proteins found in the ODV envelope, referred to as PIFs (per os infectivity factors) (Clem and Passarelli 2013). Recent results indicate that some of the PIFs form a complex in the ODV envelope, likely interacting with an unknown midgut receptor, mediating fusion with the plasma membrane (Peng et al. 2012). Thus, the PIFs constitute a novel virus attachment/fusion protein complex. Interestingly, PIFs appear to be ancient genes that are conserved among three related insect virus families, as pif homologs are also found in the nudiviruses and the polydnaviruses, the latter of which form mutualistic relationships with parasitic wasps (Bezier et al. 2009; Burke et al. 2013). In most baculovirus infections, the budded virus then spreads throughout the larva via the tracheal system and haemocytes. Late in infection, the host’s tissues are filled with virions that are occluded in millions of occlusion bodies, which are released upon death when the host liquefies (Harrison and Hoover 2012). The gross GVs pathologies are similar to those of NPVs, but differences occur depending upon the type of tissues infected. The first indication of infection in the larva is the loss of appetite and a progressive color change from the normal color to a pale whitish or milky yellow appearance, especially on the ventral side (Huger 1966). The whiteness is due to the abundance of OBs in the hypertrophied fat bodies. When the infection is limited to the fat body, the larvae often increases in size, become white, opaque and mottled at an advanced stage of infection, and later show a brownish discolouration (Hamm and Paschuke 1963). Such larvae may live longer and become larger than an uninfected one. In the case of systemic granulosis, the larva usually dies in a brief period, much shorter than observed for an infection involving mainly the fat body. At death such a larva is smaller than a healthy one.

10.5  Genome Organization The super coiled ds DNA genome of GVs varies in size from 100 to 180 kilobase pair (kbp) with potential to code for over 100 genes. The lowest genome size (99,657  bp) has been detected in Adoxophyes orana GV (AdorGV), while the

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largest (178,733 bp) has been reported in Xestia c-nigrum GV (XcGV) (Hayakawa et al. 1999; Wormleaton et al. 2003). The estimated G + C content for betabaculovirus members ranged between 32.5% in Cryptophlebia leucotreta GV (CrleGV) and 45.2% in Cydia pomonella GV (CpGV) (Han et al. 2016). The granulin gene or a restriction site close to it have been used to define the zero point of GVs restriction maps. Hybridization studies from a large number of viruses show that most, if not all, granulin gene sequences are highly conserved. The promoter regions of granulin and polyhedron genes are of considerable interest because of the very high levels of expression of these proteins. The 5′ flanking region of these genes contains a highly conserved motif of 12 nucleotides (Rohrmann 1986) which includes the transcription initiation site, followed by an Adenine (A)Thyamine (T) rich sequence up to the start of coding region. There are additional two conserved nucleotides adjacent to this region in CpGV, PbGV and Trichoplusia ni GV, giving a 14- mer peptide with sequence TTTATAAGGAATTA. The region following this sequence is shorter and more variable in GVs than in NPVs, though with a similarly high A + T content of 82–93% and a complete absence of guanine (G) (Crook and Winstanley 1992). Studies by Bah et  al. (1997) showed that the granulin genes of TnGV, PbGV, CrleGV showed homologies ranging from 76.7 to 80.5% for nucleotide sequences and from 84.2 to 88.3% for amino acids, when compared to Choristoneura fumiferana GV (ChfuGV). The enhancin genes have been sequenced for TnGV and Pseudaletia unipuncta GV (PuGV). They have more than 90% amino acid homology. These genes also contain the consensus late promoter ATAAAG sequence just upstream the coding sequence (Table 10.2). The TnGV enhancing gene hybridizes under conditions of high stringency to DNA bands from some other GVs, indicating that these GVs also contain homologous genes.

10.6  Genetic Manipulation for Improved Efficacy In general, GVs are not stable under natural conditions, as they readily get inactivated under the influence of UV rays,. Enhancement of NPV infectivity and reduction in incubation time by the enhancing gene (viral enhancing factor) VEF have been demonstrated in certain GVs such as AcMNPV. This VEF is ten times more stable than virions to UV inactivation and also impart heat stability, apart from reducing the survival time (Granados and Corsaro 1990; Gallo et al. 1991). Studies with both native and recombinant enhancing showed inactivation by metal ions chelators and reactivation with divalent ions, thus identify the enhancing factor as a metalloprotease. In CpGV, an inhibitor of apoptosis (Iap) gene has been identified. It blocks apoptosis which is a natural defense mechanism in Spodoptera frugiperda, as shown in cells infected with a mutant AcMNPV lacking a functional p35 gene. The Iap gene encodes a polypeptide with a predicted relative molecular weight of 31.3 kd. The

Sequencea

Nucleotides before ATG 15 19 11

a

Gaps to align the sequences are indicated by dashes. The numbers of nucleotides between the conserved 14–mer and the ATG translation start site are shown

Virus PbGV CpGV TnGV

Table 10.2  Sequences of the promoter regions of granulin genes from PbGV, CpGV and TnGV

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region near the C- terminus of the polypeptide contains a distinctive zinc finger like motif which is similar to that found in several other genes, with potential to regulate apoptosis. Some other genes such as Bam H1’J’ have been subcloned and subjected to transposon mutagenesis in E. coli cells using a Tn3 derivative (Wonkyung et  al. 1997). There are a number of candidate genes available which, if expressed by the virus during infection, are likely to result in earlier host death or at least in reduced insect feeding. Though much work on these lines has been done in NPVs, very little information on these aspects pertains to GVs, mainly because of difficulties in developing a good cell culture system for selection and cloning of recombinant GVs. The insertion and expression of foreign genes in baculoviruses is mainly performed through the replacement of the granulin gene coding region with a foreign gene sequence. Granulin genes from TnGV and PbGV have been cloned and sequenced by Akiyoshi et  al. (1985) and Chakerian et  al. (1985), respectively. Crook and Winstley (1992) constructed a CpGV transfer vector containing the 5′ and 3′ flanking regions of the granulin gene including the entire promoter region, but the entire granulin coding region that was deleted and replaced by a Bam H1 cloning site. A cloned strain of CpGV 9-M1 using successive rounds of an in vitro limiting dilution method was obtained by Crook et  al. (1997). The region containing the granulin gene and an open reading frame (ORF) immediately upstream of it was sequenced. Lery et  al. (1997) established 12 in vitro clones from a Tunisian isolate of PhopGV.  Substitution of granulin for polyhedrin has also been demonstrated by Eason et al. (1998) for better infectivity. Smith and Goodale (1998) determined the nucleotide sequence and located the major in vivo transcript termini of the Lacanobia oleraceae granulosis virus (LoGV) egt gene.

10.7  Mass Production (i) Culturing of Host Insects So far, no medium has been developed to culture granuloviruses under laboratory conditions. The GV can be mass multiplied only in the host insect larvae. The host insect culture can be maintained on artificial/semi synthetic diet under laboratory conditions in most of the species. In some cases, i.e. in shoot borer C. infuscatellus, larvae from 30–90  days old sugarcane crop showing fresh dead hearts were collected in plastic containers with shoot bits, for rearing of larvae for GV mass production. (ii) Propagation Third or fourth instar larvae are suitable for virus multiplication. For inoculation, a virus suspension containing 107–108 OB/ml of water is prepared and the larvae are fed with a diet containing the viral inocula for 24 h. Thereafter, the virus fed larvae

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are reared on normal diet in individual plates or containers. The plastic containers are provided with filter paper for absorption of excess moisture, and the diet and filter paper are changed on alternate days. The infected larvae begin to show symptoms in 5–8 days and start dying from 8 up to 20 days, depending upon the species. Dead larvae which die due to infection are collected and stored at -20 °C. (iii) Purification The infected larvae are macerated in distilled water and filtered through muslin cloth to remove the debris. Thereafter the suspension is centrifuged at 500 rpm for 3 min and the sediment is discarded. The supernatant is centrifuged at 10000 rpm for 30 min to sediment the virus. The supernatant is discarded and the virus pallet is re-suspended in small volume of water to obtain fairly pure preparation of the virus. (iv) Storage The virus is stored, suspended in distilled water in amber coloured bottles in a cool dark place. If possible it can be stored in the refrigerator at 4–5 °C. The virus can be stored for three or more years without apparent loss in infectivity. (v) Safety Most of the GV isolates have been found to be safe to many adult parasitoids and predators present in the crop ecosystem, as well as to productive insects such as honey bee, mulberry etc. So far, no occupational hazard has been reported while working with GVs. (vi) Mass Production Baculoviruses have potential to be used as microbial control agents against insect pests due to their high pathogenicity, host specificity, environmental benefit and biosafety (Rahman and Gopinathan 2003; Simon et  al. 2004). Till date however, baculovirus-based products used on a large scale for insect pests control have been produced on infected larvae either field collected, or to the most reared in the laboratory on an artificial diet. It is very pertinent to determine the optimal parameters for baculovirus production under in vivo conditions, including diet composition, rearing conditions (containers used, temperature, humidity), and the age/developmental stage at which to infect larvae (Shapiro 1986; Hunter-Fujita et al. 1998). In addition, facilities such as equipments, processes for efficient infection of larvae, collection of cadavers, and isolation of viral occlusions are also a prerequisite (Wood and Hughes 1996; Black et al. 1997; van Beek and Davis 2007). However, in vivo mass production on host larvae suffers from certain problems such as the insurgence of diseases in the primary host production colony (McEwen and Hervey 1960) and contamination of the final product with other microorganisms and potentially allergenic insect parts (Podgwaite et al. 1983; McKinley et al. 1997). Hence, efforts have also been made for in  vitro mass production of baculoviruses using insect cell cultures. These do not only result in an improved quality product but also result less expensive, requiring less labour for production at a larger scale. They also allow for the production of baculoviruses that infect host species for which their

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in vivo production is not feasible or desirable (e.g., species with small larvae, such as P. xylostella, or species with urticacious hairs, such as L. dispar). The development of suitable, specific media for cell culture in large scale for in vitro production of baculoviruses has further been made more feasible (Maiorella et  al. 1988; Murhammer and Goochee 1988). Baculoviruses used as biocontrol agents, however, suffer from instability after exposure to solar irradiation, especially due to the quick inactivation of the virus by sunlight. UV-B (280–310 nm) in particular are responsible for the inactivation of insect pathogens in general and of insect viruses in particular, under natural conditions (Ignoffo et  al. 1977; Asano 2005). The resulting loss of biological activity affects the rate at which insect-pests are killed and in many instances the pathogens lose >90% of their original activity within days (Ignoffo et al. 1977; Shapiro et al. 2002). Sometimes, the efficacy of these microbials is greatly reduced within the first 24–48 h post spraying (Ignoffo and Batzer 1971; Young and Yearian 1986). These factors lower the satisfactory field utilization of GVs for pest management (Timans 1982; Shapiro et  al. 2009). Brightners, white carbon and antioxidants have been found to provide some level of UV protection for insect viruses (Shapiro and Robertson 1990; Shapiro 1992, Sood et al. 2013b). (vii) Formulation Formulation of biopesticides can improve their efficacy to acceptable levels of pest control, that can be achieved with lower doses, finally representing an important reduction in the cost of each application. The efficacy of bioinsecticides that act by ingestion can be improved by the use of phagostimulant formulations that increase the consumption by the pathogen. In the case of baculovirus-based insecticides, the use of feeding stimulants that encourage phytophagous larvae to consume foliage contaminated with OBs can result in an increased prevalence of infection and improved pest control. However, the added cost of such formulations may not always be justified by corresponding improvements in crop yields. The most common baculovirus formulations are water-dispersible liquids or powders that are applied to crops as aqueous sprays using conventional spray equipment. One limitation of baculoviruses as biopesticides is that they are rapidly deactivated by UV light exposure (Ignoffo et al. 1989). The incorporation of UV-protectant sunscreens into baculovirus formulations is therefore a subject of considerable interest (Shapiro et al. 1983; Williams and Cisneros 2001). Stilbene optical brighteners substantially enhanced the infectivity of baculoviruses (Hamm and Shapiro 1992; Dougherty et al. 1996). It appears that they change the pH of the insect midgut and also block the sloughing of primary infected midgut cells, leading to a higher probability of establishment of infection in larvae simultaneously fed with virus and optical brightener, compared with virus alone (Washburn et  al. 1998). Several optical brighteners are known to interfere with chitin fibrillogenesis, increasing the permeability of this structure which normally acts as a defensive barrier to microbial invasion (Wang and Granados 2000). The inclusion of optical brighteners in formulations may substantially increase the viability of baculoviruses used for biocontrol as higher pest mortality can be achieved with lower applications of virus (Hamm et al.

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1994; Thorpe et al. 1999). Although optical brighteners are widely used for ­domestic and industrial applications, they were not previously tested in the environment. If they were used in bio insecticide-based control systems, they could be applied to crops over large areas but little is known on the potential consequences of such actions, including possible effects on crops growth. The ingredients of a biopesticide formulation must: i) ensure stability during biopesticide production, processing and storage, ii) assist application, iii) protect the biopesticide from unfavorable environmental conditions iv) promote its activity at the target site. Formulations are composed of i) an active ingredient ii) carriers, often an inert material used to support and deliver the densely populated active ingredient to the target and iii) adjuvants, compounds that: promote and sustain the function of the active ingredient by protection from UV radiation, ensure rain fastness on the target, retain moisture or protect against desiccation, or promote the spread and dispersal of the biopesticide. Most of the viral biopesticides are formulated as dusts because of a number of advantages over liquid formulations. List of GV based products commercialized worldwide (Modified after Haase et al. 2015) is provided in Table 10.3. Table 10.3  List of GV based products commercialized worldwide (modified after Haase et al. 2015) Virus name Target insects Cydia Cydia pomonella GV pomonella, Grapholita molesta

Crops Apple, pear, walnut Apple, peach

Product names Carpovirus Plus, Madex, Carpovirusine, Madex Twin

Erinnyis ello GV

Erinnyis ello

Cassava, rubber trees

Baculovirus erinnyis

Phthorimaea operculella GV

Phthorimaea Potato operculella Tecia solanivora

Baculovirus, PTM baculovirus Corpoica Matapol Plus, Bacu-Turin

Phtorimaea Potato operculella Tecia solanivora Symmetrischema tangolias Codling moth Codling moth Vegetable Madmex, GV crops Granupom Phthorimaea operculella GV + Bacillus thuringiensis

Producer companies NPP-Arysta Life Science, Andermatt Biocontrol NPP-Arysta Life Science Empresa de Pesquisa Agropecuária e Extensão, Rural de Santa Catarina, BioCaribe, CORPOICA CORPOICA SENASA Peru, INTA Costa Rica PROINPA Foundation, INIAP

Andermatt Biocontrol,

Country Argentina, Uruguay Chile

Brazil, Colombia

Colombia, Peru, Costa Rica Bolivia, Ecuador

Switzerland, Germany

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10.8  Field Applications Since many years, GVs have been observed to be responsible for severe epizootics in various insect populations (Huger 1963). The GV of P. rapae excised an unusual high regulation degree of the pest during 1947–1948 and 1958 in New Zealand. The same GV was also observed in Hawaii to cause severe epizootics. Similar observations were made in Japan (Ito et al. 1960). Other GVs that cause high mortality in the field have been observed in populations of Choristoneura murinana on spruces, Hyphantria cunea on walnuts and Cydia pomonella on apples, pears and walnuts. In Chilo infuscatellus, the GV prevalence ranged from 16 to 36.8% in 1981 and from 17 to 42% in 1982 (Easwaramoorthy 1984). Similarly in Chilo sacchariphagus indicus, the GV infection ranged from 17.4 to 35% in 1982 and from 18.8 to 45.0% in 1982. A number of GVs underwent field testing for management of pests and the important ones are reviewed below. The most famous GV which has been developed as a commercial product is ‘Capex’ which is derived from AdorGV and got registered in Switzerland in 1989 (Andermatt 1991). A successful use of the GV (AsGV) has been reported against Agrotis segetum on root crops in Denmark (Zethner 1980), on maize in Spain (Caballero et al. 1990, 1991) and lettuces and dwarf asters in Germany (Fritzsche et al. 1991). Working on improvement of Virin-OS (AsGV), Kitik (1989) during 1987–1989 produced about 20 kg of formulated material which was used in Uzbekistan and Moldovia to control the pest on cotton, sugar beet, vegetables and winter cereals. A mixture of GV and NPV has also been tested with reasonable success in the Former Soviet Union. In a joint project between Denmark and Pakistan, a Danish strain of AsGV was tested in Pakistan on tobacco. Spraying tobacco seedlings with concentrations from 5 × 107 to 1 × 109 OBs/ml, at 60–250 ml/m reduced second instar larval damage by 72–100% (Chaudhry et al. 1976; Shah et al. 1979; Zethner et al. 1983). Potato is an important food crop which is infested with Phthorimaea operculella. A granulovirus of this insect (PhopGV) has been isolated from Sri Lanka and was later found also in Tunisia, Yemen, Egypt and elsewhere in Africa. Several isolates have been characterized (Vickers et al. 1991; CIP, 1993). Its production methodologies were developed in South America. These technologies appeared highly suitable and thus were successfully adopted in Tunisia and Egypt, where facilities capable of production enough virus to treat 10,000 tonnes of tubers were built (Abol-Ela et al. 1996). In Tunisia, trials were conducted to reduce infestation in tubers at harvest and in stores thereafter, by field applications of PhopGV to the soil surface before harvest. The spray formulation were reported to reduce field infestation of tubers by 73%. In storage, pest infestation failed to develop from tubers subjected to field treatment. Trails in Egypt demonstrated a much better rate of post-harvest protection (95%) using 2 LE (larval equivalent) in 10 ml water with 0.001% Tween 20 per kg of tubers. The virus was also found effective in Australia (Reed 1969; Reed and Springnett 1971), Peru (de Oliveira 1998) and Bolivia (Calderon and Andrews

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1994). In Peru, a PhopGV isolate was developed as a microbial pesticide through an initiative of the International Potato Center (CIP). This virus provided effective control of the pest when 20 virus infected larvae were mixed with one kg of talc, and used as a suspension in 1 L of water. A dry product applied at a dose of 5 kg per tonne of stored tubers also provided high levels of control (ca. 95% mortality) (Raman et al. 1987). Chilo infuscatellus is an important pest of sugarcane. The application of CiGV at 1 × 107 to 109 OBs/ml at 35, 50, 65 and 80 days after planting reduced pest incidence below the economic injury level (Easwaramoorthy 1984, 2002; Easwaramoorthy and Santhalakshmi 1988). Parameswaran et  al. (1991) reported that application of CiGV (107 OBs), twice at 35 and 50 days after planting, effectively reduced the shoot borer population. Application at a high volume was more effective than at a low volume (Easwaramoorthy and Jayaraj 1991). The CiGV was commercially produced by the Tamil Nadu Agriculture University (TNAU), Coimbatore, India for more than a decade. The GV of the codling moth, Cydia pomonella has been extensively tested in various countries. CpGV has been shown to be a potent agent for codling moth control. The application of CpGV has been especially more useful in regions where codling moth is univoltine, as in most of Europe. Where there is more than one annual generation e.g. in California, and especially where such generations have not exactly synchronized phenology, the pest appeared more difficult to manage. The use of pheromone trapping to reveal adult emergence patterns permits improved accuracy of time of spraying. Developmental programmes have been conducted in many countries such as USA and inside Europe in Austria, Germany, the Netherlands, United Kingdom and Switzerland. Under an IOBC/WPRS working group, the latter group of nations, together with Hungary tested the virus in more than a dozen orchards in 1976–1977, using material produced in Darmstadt, Germany (Huber 1981). A C. pomonella granulovirus (CpGV) identified by Tanada (1964) was capable of causing epizootics in populations infesting pome fruits and walnuts. Field tests data proved this virus to be a potential candidate in providing an effective alternative for control of codling moth larvae in integrated and organic pome fruit and walnut production in several countries (Huber and Dickler 1977). In Argentina, Carpovirus Plus® (Quintana and Alvarado 2004) and Madex® (Haase et al. 2015) showed highly satisfactory results when the virus was applied at doses of 1013 OBs/ ha at intervals of 8–10 days. An interesting separation of two strains of the AdorGV replicating in either the nucleus or the cytoplasm has been made, but it is not known which one was employed in field tests (Huber 1998). In field collected Cnephasia spp., Glas (1991) found a GV that infected both C. longana and C. pumicana. This author showed that it is possible to spread the virus disease by spraying GV suspensions on the bark of trees. A dose of 2 × 1010 OBs per trunk was sufficient to induce more than 90% larval mortality, leading to a complete breakdown of the pest population. In a field where the population was already

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p­ resent, spraying of GV on the eggs present on the trees was still useful, as the population collapsed 3 weeks earlier than in the untreated plots. Several commercial products of CpGV have been marketed in Europe. They include Carpovirisine (Burgerjon and Sureau 1985), Madex, Granupom, Granusal etc. In the Netherlands, Granusal was as effective as a chemical regime (Helsen et al. 1992). In Italy, field comparisons of Carpovirisine and Granupom suggested that both were as effective as a standard insecticide (Pasqualini et al. 1994). Studies on the non-chemical control of C. pomonella in France over a 10 year period were reviewed by Andemard (1986), who concluded that mating disruption and CpGV were the most promising management approaches. The fall armyworm Spodoptera frugiperda is another pest of economic importance. It is a polyphagous insect that causes economic losses in several important crops, such as maize, sorghum, rice, cotton, and pastures. Since this pest showed resistance to other control tactics, baculoviruses were evaluated for effectiveness in Argentina (Yasem de Romero et al. 2009), Brazil (Valicente and da Costa 1995), Mexico (Ríos-Velasco et al. 2012) and Peru (Vásquez et al. 2002). The efficacy in controlling the pest was similar to that of chemical insecticides, 22 days post emergence of the pest (Gómez et al. 2013). The epizootiology of a naturally occurring GV of Dendrolimus sibiricus was studied in the former Soviet Union by Baranovsky and Litvina (1978), and subsequently this GV was commercialized. Applications are best made in the autumn against early instars (Orlovskaya 1989). Erinnys ello, although known to attack over 35 plant species in South America, is known especially as a pest of cassava and rubber. In Brazil, the ErelGV has been used on over 2000 ha of cassava and rubber (de Oliveria 1998). A standardized formulation consists of 20 ml (18 g) of a crude and filtered macerate which is diluted for spraying in 200 L water/ha or for ULV in 3 L water/ha. In Colombia, 50–70 ml of larval macerated in 200 L water with 0.2 ml Triton-ACT is sprayed per ha. The speed of action in Colombia is phenomenal, as >80% mortality was recorded in 48 h. In Southern Brazil, a mortality of 90% was attained in 4 days after spraying of 20 ml of larval macerate. An increase in numbers of natural enemies following GV application is thought to explain the observed spread of infection from sprayed areas. Hypanthia cunea is a serious defoliator of popular and other deciduous trees. Experimental Hypanthia cunea granulosis virus (HycuGV) insecticides were developed and tested in Bulgaria with the commercial name of Hifantrin and in Russia and Ukraina with the name of Virin-ABB. Satisfactory results were reported from Ukraina (Sikura and Smetnik 1980; Kransnitskaya 1980). At present Virin-ABB-3 is in commercial production in Moldavia (Chkhirii 1993). Limited greenhouse trails indicated that Lacanobia oleracea GV has a strong potential as a pest control agent on tomato (Crook and Brown 1982). The effectiveness of spraying the homologous GV of Pieris rapae alone or in combination with Bacillus thuringiensis (Bt) var. kurstakii was reported in Taiwan (Su 1986, 1991). Control was effective for 14 days in the field with persistence of GV exceeding that of Bt in China, using 1 g of the powered GV product in 30 ml water, presumably sprayed to get a reasonable droplet coverage with 90% control of

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P. rapae on cabbage. Rituma (1990) reported that virin-KB gave 70–96% control and increased the yield value by 150 roubles/ha in Lativa. In North America, the GV is recommended at a dose of 7.5 × 1012 OBs/ha in about 100 L/ha. Two to six applications per year are made on cole crops, depending on date of planting and harvest. The control of P. rapae by GV measured by crop protection equals or exceeds control by either chemical insecticides or Bt (Cunningham 1998). A freeze-dried preparation was found to be effective for protection of almonds and raisins against attacks by the Indian meal moth, Plodia interpunctella Hubner. The virus was extensively tested on dried fruits and nuts and, to a lesser extent, on grains. It is used as a prophylactic material and persistence is adequate to provide protection from packaging of foods for consumption. The population of P. interpunctella was reduced by 95–100% and damage to products was reduced to a non-­ detectable level. A formulation of this virus has been patented by the USDA Agricultural Research Service. Cabbage butterfly, Pieris brassicae (Linnaeus) (Lepidoptera: Pieridae), is a serious pest of cabbage, cauliflowers and many crucifer crops of the world (Feltwell 1978; Bhandari et al. 2009). A P. brassicae granulovirus (PbGV) strain was isolated and characterized from Himachal Pradesh, India by Sood (2004), which was effective in managing P. brassicae larvae (Sood et al. 2011; Sood and Prabhakar 2012). The effective formulation technology is another challenge for the development of a stable insect virus product under field conditions. Usually, active ingredients i.e. virus OBs/POBs are mixed with various adjuvants which improve the efficacy, stability and handling of the pesticide. The most common process of formulating a virus is drying the infected larvae, either through dehydration, lyophilization or by an air flow, in order to generate a powder. Lactose was also added to improve the viruses stability and infectivity. A suitable carrier depending upon its compatibility with the virus particles is then added to achieve the uniform concentration. Silica and clays are commonly used for this purpose. Surfactants, adherents, thickeners, binders, phagostimulants, UV protectants or optical brighteners, boric acid as stress causative, etc. may also be added to make a good formulation (Sood et al. 2013a, b). Formulation in the form of microencapsulation has also been developed.

10.9  Conclusion Under natural conditions insect populations are regulated by epizootics caused by various entomopathogens such as bacteria, fungi and viruses etc. These may offer viable pest management solutions in most agroecosystems. The growing knowledge and awareness on the negative impacts of chemical insecticide usage on the ecosystem and public health led to efforts directed towards usage of biopesticides over synthetic pesticides. Among various entomopathogens, baculoviruses in particular are promising alternatives. Many virus-based formulations have been used in the past and are available worldwide, but their usage has been still restricted compared to other pesticides. Their increased utilization therefore requires enhanced virulence

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coupled with speed of kill and stability in the ecosystem. Accordingly, viruses initially found in nature need to be improved in terms of virulence and stability to make them more potent, before field usage. A number of granuloviruses infecting major insect pests have been isolated and tested for management, but only a few have been commercialized owing to their variable potency and stability. To counter this problem, some researchers manipulated and formulated stable products. However, still a lot has to be done to popularize their usage in large scale pest management programmes, which can be achieved by developing more virulent and stable formulations, apart from ease of the registration procedure.

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Chapter 11

Interactions of Entomopathogens with Other Pest Management Options Surendra K. Dara

Abstract  As the demand for sustainably produced food is increasing, the use of biopesticides has also been increasing in the recent years. While biopesticides based on entomopathogens have been successfully employed for pest management around the world for decades, there are still some concerns about their efficacy relative to chemical pesticides, limiting their widespread adaptation. Microbial pesticides could be an important part of all integrated pest management (IPM) strategies and understanding their interactions with other control options can help promote their use. Several studies demonstrated improved pest control efficacy when different kinds of entomopathogens were used. Entomopathogens were combined or rotated with chemical or botanical pesticides, or virulence of certain entomopathogens was synergized by chemical pesticides. Certain fungicides are also compatible with entomopathogenic fungi enabling pest and disease management treatments at the same time. Improved microbial control can help reduce chemical pesticide use and the associated environmental and human health risks. Keywords  Entomopathogens · Microbial control · Integrated Pest management · Synergy

11.1  Introduction Entomopathogens such as bacteria (e.g., Bacillus, Serratia, and Yersinia), fungi (e.g., Beauveria, Conidibolus, Entomophaga, Entomophthora, Erynia, Hirsutella, Isaria, Lecanicillium, Metarhizium, Neozygites, and Pandora), microsporidia (e.g., Brachiola, Endoreticulatus, Nosema, and Vairimorpha), nematodes (e.g., Heterorhabditis and Steinernema), and viruses (e.g., granuloviruses and nucleopolyhedroviruses) have been used for pest management in agriculture, forestry, animal husbandry, aquatic or urban environments, to manage a variety of arthropod S. K. Dara (*) University of California Cooperative Extension, San Luis Obispo, CA, USA e-mail: [email protected] © Springer Nature Switzerland AG 2019 M. A. Khan, W. Ahmad (eds.), Microbes for Sustainable Insect Pest Management, Sustainability in Plant and Crop Protection, https://doi.org/10.1007/978-3-030-23045-6_11

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pests. While some of these pathogens manage target pest populations through natural infections, others are mass-produced and applied through inundative releases. Several entomopathogens have been developed into a number of commercial biopesticide formulations and used for pest management, alone or in combination or rotation with other management options. Traditionally, entomopathogen-based microbial pesticides had limited use compared to chemical pesticides and were used in high value crops, greenhouse productions systems, or when control with chemicals or other conventional options was difficult. Some of the factors that may limit their widespread use are related to: a higher cost of microbial pesticides compared to chemical pesticides and the need of special storage and handling, shorter shelf life, poor quality control, slow rate of pest control, a lack of understanding of microbial control mechanisms, and the general perception of variability or unreliable control. While a few manufacturers of microbial pesticides maintained their share in some niche markets, complexities of mass-production of entomopathogens and the high cost of registering pesticide formulations restricted the development of new microbial pesticides, although research around the world identified several virulent isolates. However, indiscriminate use of chemical pesticides that frequently leads to the development of resistant pest populations as well as raises concern for environmental and human health, brought attention, in recent years, to sustainable food production using safer pest control options. Because of the increased demand for organically produced food, organic acreage has been steadily increasing around the world, ultimately creating a demand for non-chemical control options including microbial pesticides. For example, between 2007 and 2012, there was an 83% increase in the sales of organic products by the farms in the United States (USDA-NASS 2016). To maintain their share in the pesticide market, several large chemical pesticide companies are now adding microbial pesticides, among other non-chemical tools, to their product portfolios. An emphasis on sustainable food production is also promoting the use of microbial pesticides on conventional farms, as a part of integrated pest management (IPM), in addition to their common use in organic farms. In light of the demand for sustainably produced foods and increasing popularity and use of microbial pesticides, research studies are evaluating the efficacy of various microbial pesticides and their incorporation into IPM strategies. This chapter focuses on the interaction of entomopathogens among themselves and with other pest management options, to understand their efficacy and role in IPM.

11.2  Entomopathogen Combinations Similar to the combinations of chemical pesticides aiming at improving pest control efficacy, certain combinations of entomopathogens can also be employed for pest management with additive or synergistic effects. A number of studies evaluated the combination of entomopathogenic nematodes (EPN) with entomopathogenic fungi (EPF) or bacteria against various pests. Additive effect of Metarhizium anisopliae

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with Heterorhabditis indica at two application rates, and Beauveria bassiana with the bacterium Serratia marcescens at a higher rate, were observed against the pecan weevil, Curculio caryae resulting in increased pest mortality (Shapiro-Ilan et  al. 2004). Similarly, the combination of M. anisopliae and Steinernema kraussei as a soil drench resulted in 100% mortality of the overwintering larvae of the black vine weevil, Otiorhynchus sulcatus in strawberry compared to 50–88% mortality by M. anisopliae and 60–69% mortality by S. kraussei, when applied alone (Ansari et al. 2010). Acevedo et al. (2007) reported that the combination of a Brazilian isolate of H. bacteriophora (JPM4) and an isolate of M. anisopliae (LPP39) improved the mortality of the sugarcane borer, Diatraea saccharalis, with reduced LT50 and LT95 values. However, production of the infective juveniles (IJ) was reduced in the presence of M. anisopliae. In such cases, applying EPF prior to the application of EPN appeared to be ideal for IJ production (Shaurub et al. 2016). Application of B. bassiana before applying S. riobrave or H. bacteriophora improved IJ production in the Egyptian cotton leafworm, Spodoptera littoralis. Combination of different EPN or EPN with B. bassiana also resulted in increased S. littoralis mortality. Similar improved control from the co-application of EPF and EPN was seen in the large pine weevil, Hylobius abietis in Ireland (Williams et  al. 2013) and the chestnut weevil, Curculio elephas in Turkey (Asan et al. 2017). Fungal infections probably weaken the pest and increase infection by EPNs. The combination of EPF and bacteria was also found effective in different studies where bacterial infection reduced insect feeding, weakening the pest, delaying molting and increasing the chances of fungal infection. The combination of B. bassiana and B. thuringiensis subsp. tenebrionis increased the control efficacy by 6–35% in Colorado potato beetle, Leptinotarsa decemlineata in field studies in the United States (Wraight and Ramos 2005). Similarly, improved mortality in a shorter time was achieved by the combination of B. thuringiensis subsp. morrisoni (Btm) with lower conidial concentrations of M. anisopliae or B. bassiana vs L. decemlineata in Kazakhstan (Kryukov et al. 2009). Yaroslavtseva et al. (2017) demonstrated that sublethal doses of Btm var. tenebrionis significantly suppressed the cellular immunity and inhibited the detoxification enzymes in L. decemlineata resulting in a synergy with M. robertsii. Preliminary laboratory studies in China showed that B. bassiana infection in the second instar larvae of the spotted asparagus beetle, Crioceris quatuordecimpunctata, increased from 42% to 78% in the presence of a sublethal dose of Cry3Aa Bt toxin (10 μg/ml) (Gao et al. 2012). In an earlier study, synergism was seen between M. anisopliae and Serratia entomophila against the second instar larvae of the New Zealand grass grub, Costelytra zealandica (Glare 1994). In a Greek study, Mantzoukas et al. (2013) evaluated different combinations of Bt subsp. kurstaki (Btk), B. bassiana, and M. robertsii against the Mediterranean cornborer, Sesamia nanogrioides and found synergistic, additive, and negative interactions depending on the combination and time of observation. Sayed and Behle (2017) evaluated different proportions of Btk spores and crystals and B. bassiana blastospores against the cabbage looper, Trichoplusia ni, in the United States and found that 50:50 ratio was synergistic. A recent study in Pakistan ­demonstrated that EPF-Bt combination can also be used against sucking pests

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(Jugno et al. 2018). Commercial formulations of M. anisopliae and Bt were tested alone and in combination against the Indian cotton jassid, Amrasca biguttula and the cotton aphid, Aphis gossypii on eggplant. The combination killed 83% of the jassids and 72% of the aphids compared to 53% and 52% mortality by Bt and 67% and 59% by M. anisopliae in jassid and aphid populations, respectively. Combining entomopathogens with different modes of infection can enhance the pest control efficacy by taking advantage of multiple routes of infection. EPF infections are primarily by contact, bacterial infections are by ingestion, and EPN infections occurs through natural openings or cuticle by IJs actively seeking their host. Compared to younger pest larvae, older ones are more resistant to bacterial infections while EPF can be more effective against adults than immature stages, that might escape infection through molting. Combination of pathogens can also help address these situations by targeting multiple life stages.

11.3  Entomopathogens with Natural Enemies Natural enemies of arthropod pests coevolved with entomopathogens and developed their own ways of resisting or avoiding infections. However, the susceptibility of natural enemies to entomopathogens depends on the arthropod species or the strain of the pathogen. For example, the B. bassiana isolates from the native coccinellid predator, Olla v-nigrum in Southeastern United States were highly pathogenic to the native predator, but not to the invasive Harmonia axyridis (Cottrell and Shapiro-Ilan 2003). However, the commercial GHA isolate of B. bassiana was relatively less pathogenic to both lady beetle species. A laboratory study in Austria showed that a local isolate of B. bassiana was pathogenic to the bark beetle, Ips sexdentatus, but appeared safe to its predator, the European red-bellied clerid (Thanasimus formicarius) at the same concentration (1  ×  107 conidiophores/ml). Only when a very high dose of dry conidia of unspecified concentration was applied to the entire ventral side of the adult predators, 47% of them died from 24 to 48 days after inoculation (Steinwender et al. 2010). Similarly, Bourassa et al. (2001) reported that some African isolates of B. bassiana and M. anisopliae caused 77–85% mortality in adult larger grain borer, Prostephanus truncatus, compared to a 13–21% mortality in its predator, Teretriosoma nigrescens, 6 days after treatment. While the pest mortality went up to 91–96% in 14  days, that of the predator increased up to 42–56%. A Canadian study evaluated 10 isolates of B. bassiana from North America, Europe, and West Africa against L. decemlineata, the green peach aphid, Myzus persicae, and a predatory coccinellid, Coleomegilla maculata lengi (Todorova et al. 2000). While six of the isolates were highly pathogenic to both the pest and predatory species, two Canadian isolates, a Bulgarian and a Beninese isolate were highly pathogenic to the pest species, but not the coccinellid predator. Jacobson et al. (2010) reported that application of B. bassiana effectively controlled populations of the western flower thrips, Frankliniella occidentalis, in ­glasshouse cucumbers without having any negative impact on the predatory mite,

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Neoseiulus cucumeris, which was prophylactically released to prevent the buildup of whitefly populations. There was no difference in predatory mite numbers on treated plants, compared to untreated plants, even 3 weeks after B. bassiana application. Similarly, B. bassiana did not affect the predation of Phytoseiulus persimilis on the twospotted spider mite, Tetranychus urticae (Wu et al. 2018). Studies conducted in California strawberries also did not show any negative impact of B. bassiana on P. persimilis and Neoseiulus spp. when evaluated against T. urticae (Dara 2015a). No impact was also observed for B. bassiana, I. fumosorosea, and M. brunneum on a variety of arthropod natural enemies, when tested against the western tarnished plant bug, Lygus hesperus (Dara 2016). However, an isolate of M. brunneum was highly pathogenic to the predatory mites P. persimilis and Neoseiulus californicus (Dogan et al. 2017) while B. bassiana and I. fumosorosea caused 43 and 31% mortality in P. persimilis, respectively, and about 16% mortality in N. californicus in laboratory assays (Vergel et al. 2011). In studies conducted in Benin, the predatory mite Typhlodromalus aripo consumed equal numbers of healthy cassava green mites (Mononychellus tanajoa) and those infected with the entomophthoralean fungus Neozygites tanajoae (Ariori and Dara 2007; Agboton et  al. 2013). Combined use of T. aripo and N. tanajoae in screenhouse experiments showed reduced efficacy of T. aripo (Agboton et al. 2013) in the presence of the fungus. The preference of T. aripo to feed on fungus-infected pest mites and subsequent loss of survival and oviposition were thought to be responsible for the reduced efficacy of the predatory mite, in the presence of the fungus. In other studies, combined use of biological and microbial control agents improved the control efficacy. For example, the combination of the mosquito predator, Toxorhynchites brevipalpis and M. brunneum against Aedes aegypti larvae showed additive and synergistic effect at higher conidial concentrations (Alkhaibari et al. 2018). Since M. brunneum is pathogenic to T. brevipalpis, their combination was antagonistic at the concentrations of 105 and 106 conidia or blastospores/ml. However, the addition of the mosquito predator had an additive effect on the median LT50 of Ae. aegypti caused by the fungus at 107 conidia/ml and a synergistic effect at 108 conidia/ml. However, such an effect was not seen with blastospores because they were more virulent to the predator. In a 1981 study in the United States, parasitization of the gypsy moth (Lymantria dispar) by Apanteles melanoscelus increased 6–12 times in plots treated with Bt showing a synergistic effect (Weseloh et al. 1983). Bt reduces the size of surviving caterpillars and A. melanoscelus parasitism is higher in smaller caterpillars. Natural enemies can also help to spread an entomopathogen infection. Kryukov et al. (2017) reported that the ectoparasitoid Habrobracon hebetor, did not distinguish between healthy greater wax moth (Galleria mellonella) larvae and those infected with B. bassiana and significantly increased fungal infections through passive vectoring. It also appeared that envenomation by the parasitoid increased the conidial germination on the insect cuticle and reduced the fungal encapsulation (insect immune response) in the hemolymph. Roy and Pell (2000) discussed synergistic and antagonistic interactions of EPF and various natural enemies. For ­example, foraging natural enemies increased the infections of the entomophthora-

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lean fungi Pandora neoaphidis in pea aphid, Acyrthosiphon pisum, and Zoophthora radicans in the diamondback moth, Plutella xylostella, in laboratory studies, in some cases spreading the infection to healthy pest populations on different plants. Although some species of natural enemies are physiologically susceptible, especially under laboratory conditions, to hyphomycetous fungi that have a wide host range, they are ecologically less susceptible under field conditions. These studies show entomopathogen and natural enemy combinations that are either compatible or more effective. Understanding these interactions will help management decisions such as those considering the timing of natural enemies or entomopathogens release or certain combinations to avoid.

11.4  Entomopathogens with Semiochemicals, Traps, and Netting Unlike chemical and botanical pesticides, which kill the target arthropods either through direct contact or systemic activity in the plant, entomopathogens can persist, proliferate, and be disseminated in pest populations and habitats. Depending on the insect species, bait stations or pheromone traps equipped with entomopathogens, or insect netting treated with EPF, can be used to kill the visiting insects or inoculate them to disseminate inocula or infection in the rest of their populations. An earlier study in the United States by Nordin et al. (1990) explored autodissemination of two nucelopolyhedroviruses (NPV) from treated males to mated females and eventual transovarial transmission. This was a preliminary step to develop a technique using sex pheromones. Virus-treated males of tobacco budworm, Heliothis virescens, mated with females resulting in 69% virus-induced mortality in the progeny in the case of Autographa californica multiple nucleopolyhedrovirus (AcNPV), and 53% in the case of Helicoverpa zea single nucleopolyhedrovirus. However, in a two-year follow up field study using pheromone traps, male H. virescens transmitted AcNPV to females and caused subsequent larval mortality in the next generation, but to a limited extent (Jackson et al. 1992). Dispersal of H. virescens moths beyond the experimental plots was thought to be a reason for observing limited efficacy. Frankliniella occidentalis acquired twice as many M. anisopliae conidia in the presence of a semiochemical (5.0 × 104) in an autoinoculation device, compared to not having the semiochemical (2.2 × 104) in French bean fields, resulting in a 42% increase in thrips mortality in a Kenyan study (Niassy et al. 2011). However, semiochemical volatiles drastically reduced the conidial viability within 7 days, with a gradual decline in daily mortality. In a field study in Canada, control of the artificial infestations of the common click beetle, Agriotes obscurus, by band applications of M. brunneum conidia on rice granules significantly improved when pheromone-­ impregnated cellulose-based granules were also applied (Kabaluk et  al. 2015). Some studies evaluated the mating competitiveness of males treated with EPF and

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found that fungus-treated males were equally competitive as untreated males for the first few days after inoculation and that their competitiveness declined as the infection progressed (Dimbi et  al. 2009; Thaochan and Ngampongsai 2018). In their laboratory study conducted in Spain with an attractant and M. anisopliae against the Mediterranean fruit fly, Ceratitis capitata, San Andrés et al. (2014) also showed that the fitness and attractiveness of sterile or wild type males were not affected by fungal infection. Entomopathogens such as EPF can be used for managing non-agricultural pests such as ticks, flies, and mosquitoes. In Kenya, a 64% reduction in field populations of the tropical bont tick, Amblyomma variegatum, was achieved by using M. anisopliae in a semiochemical-baited trap (Nchu et al. 2010). A simple contamination device made with a clear plastic water bottle helped infect >90% of the visiting tsetse flies (Glossina spp.) with a Kenyan isolate of M. anisopliae (Maniania 2002). Conidial viability lasted for 31 days under field conditions in this study. Mosquito nets treated with M. anisopliae and B. bassiana were successfully used for controlling different species of mosquitoes (Mnyone et al. 2010; Howard et al. 2010).

11.5  Entomovectoring Entomovectoring refers to the use of pollinators such as bees to deliver entomopathogen inocula to pest habitats. Although pollinators are not directly involved in pest management, this section is included to provide a brief overview of how the foraging behavior of bees can be used for microbial control. Beehives or nesting units are attached with compartments containing entomopathogen inocula at the exit points. Bees walk through and pick up inoculum as they leave the hives and passively disperse it as they visit different plants. When bees return to the hives, they carry very low levels of inoculum and the high temperatures in the hives are generally detrimental to the entomopathogens, preventing infections in bee colonies. While this technology is now commercially available (e.g. Flying Doctors® by Biobest) to deliver beneficial microbes for pest and disease control, several earlier studies explored different entomovectoring concepts initially for controlling plant pathogens, and later for arthropod pests. In 1989, Gross et al. (1994) developed a ‘honey bee hive compatible pathogen applicator device’ to deliver Heliothis NPV (HNPV) in crimson clover (Trifolium incarnatum), in Southeastern United States. HNPV-induced mortality in corn earworms (Helicoverpa zea) was significantly higher (74–87%) in fields foraged by Apis mellifera with HNPV compared to control fields (11–14%). HNPV-induced mortality was also higher in Heliothis spp. collected from the treated fields and held in the lab (27–42%) compared to those from the control fields (2%). Similarly, mortality of H. zea that fed on cranesbill (Geranium carolinianum) fruits and flowers from the treated fields or on the honey from HNPV exposed beehives was also very high compared to controls. In 1996 and 1997 Jyoti and Brewer (1999) used A. ­mellifera to deliver Bt in North Dakota sunflower fields to control the banded sun-

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flower moth, Cochylis hospes. They did not evaluate the pest mortality in the field, but fed the larvae with diet containing wash water from sunflower capitula collected in fields foraged by A. mellifera carrying Bt, fields sprayed with Bt, and untreated fields resulting in 87–88%, 58–68% and 8–10% mortality, respectively. Seed yield, seed weight, oil content, and damage control from bee-vectored Bt fields were equal or superior to those measured in Bt sprays. Butt et al. (1998) reported using A. mellifera to deliver M. anisopliae to control the pollen beetle, Meligethes aeneus, in oilseed rape during the winter and spring of 1997, in field cages (2.7 × 2.7 × 1.8 m) in the United Kingdom. When beetles collected from the cages were incubated in the lab, the mortality on different sampling dates during the two seasons varied from 23–99% in the presence of bees with M. anisopliae, compared to 0–23% in the presence of bees without the fungus. In a Canadian study, when B. bassiana was delivered by the bumble bee Bombus impatiens in greenhouse sweet peppers, it resulted in 34–45% mortality in the tarnished plant bug, Lygus lineolaris, and 34–40% in F. occidentalis, compared to 9–15% and 3% in controls, respectively (Al-mazra’awi et al. 2006a). In another Canadian study in greenhouse tomato and sweet pepper, B. bassiana delivered by B. impatiens provided a significant control of the greenhouse whitefly, Trialeurodes vaporariorum (49%) in tomato, and of L. lineolaris (73%) in sweet pepper (Kapongo et  al. 2008). Bees also delivered another fungus, Clonostachys rosea (Gliocladium roseum), resulting in a significant suppression of the grey mold caused by Botrytis cinerea in tomato and sweet pepper. In a follow up study in greenhouse tomato and sweet pepper, Shipp et al. (2012) found no negative impact of B. bassiana delivered by B. impatiens on parasitoids Encarsia formosa, Eretmocerus eremicus, and Aphidius colemani and the predatory mite Amblyseius swirskii. Although mortality of the predator Orius insidiosus was higher in B. bassiana treatment, there was no difference in overall parasitism or predation by different species. Bumble bee B. terrestris picked up 9.3 × 106 M. anisopliae conidia on their body as they walked through the inoculum containing 107 conidia/g and retained 28% of those spores after flying for a minute, with no negative impact on survival (Smagghe et al. 2013). Similarly, Lin et al. (2017) explored the idea of delivering B. bassiana with predatory mites Stratiolaelaps scimitus, N. cucumeris and Amblyseius swirskii. Predatory mites retained conidia mixed in the substrate (inert material used for transporting and delivering predatory mites) with minimal or no negative effect on their survival by the fungus. However, evaluation of the efficacy of B. bassiana on the pest mite was not within the scope of this study. In a Canadian study, B. bassiana transported by honey bee, A. mellifera, in canola (Brassica napus) resulted in a higher mortality (48–56% in 2002 and 22–45% in 2003) in L. lineolaris, compared to those in control cages visited by fungus-free bees (9–10% in 2002 and 15–22% in 2003) (Al-Mazra’awi et al. 2006b). Entomovectoring is now more common in greenhouses than in open fields. Since bee pollination is utilized in a number of field crops and orchard systems, growers can take advantage of the bees to deliver inocula of entomopathogens for arthropod pests and other beneficial microbes for disease control. This practice can reduce the cost of applying pesticides and increase the control efficacy.

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11.6  E  ntomopathogens with Botanical and Chemical Pesticides Using entomopathogens with botanical and chemical pesticides is a good strategy to: (i) maintain pest control efficacy, (ii) reduce the risk of resistance development, (iii) extend the life of effective chemical pesticides, (iv) minimize the impact on nontarget organisms, (v) optimize the cost of pest management, and (vi) maintain environmental and human health. Employing a variety of control options is an important component of IPM, and entomopathogens can be important for many IPM programs. Several studies demonstrated improved pest management with combining or rotating entomopathogens with botanical and chemical pesticides. Shi and Feng (2006) reported that combining ~5% of the labeled rate of pyridaben 15% with an emulsifiable formulation of B. bassiana provided a very effective and prolonged control of the citrus red mite, Panonychus citri, in citrus orchards in southeastern China, compared to pyridaben treatment alone at the labeled rate. Exposing the house fly, Musca domestica, to sequential applications of imidacloprid and B. bassiana at LC30 rates resulted in a significant increase in the mortality than the individual treatments in both insecticide-resistant and susceptible adults, especially when B. bassiana followed imidacloprid application (Farooq et  al. 2018). Applying imidacloprid after B. bassiana increased the mortality by 7–9% in the susceptible strain and by 8–10% in the resistant strain, whereas fungal treatment after imidacloprid increased 16% mortality in the susceptible strain and 18–22% in the resistant strain. Although the resistant strain had higher expression of detoxification genes and elevated levels of detoxification enzymes, adding the fungus significantly improved fly mortality. In a West African study, imidacloprid also improved the conidial germination of another hypocrealean fungus, Hirsutella thompsonii, but did not significantly improve the mortality of M. tanajoa, in combination (Dara and Hountondji 2001). Assays conducted in Georgia showed that simultaneous or sequential treatment of B. bassiana and the insect growth regulators (IGRs) diflubenzuron and novaluron had additive effects on the mortality of the second instar nymphs of the migratory locust, Locusta migratoria migratorioides (Bitsadze et al. 2013). Nymphs were exposed to B. bassiana by dipping in conidial suspensions and to IGR by feeding pieces of IGR-treated corn leaves. Except when diflubenzuron-­ treated leaves were fed first followed by the fungal treatment after 48 hours, simultaneous exposure or exposing the nymphs to B. bassiana first resulted in additive effect. Hiromori and Nishigaki (1998) found improved control of the first instar larvae of the oriental beetle, Anomala cuprea, when a Japanese isolate of M. anisopliae was used with some chemical insecticides and IGRs in laboratory assays, although such an improvement could not be noticed in the field. Studies conducted in California on strawberries and vegetables, and the feedback received from several vegetable growers showed that combining azadirachtin – a botanical insect growth regulator, insecticide, and antifeedant  – with B. bassiana is more effective than individual treatments of the fungus or azadirachtin (Dara 2015b, 2016). Infestations of the honeysuckle aphid, Hyadaphis foeniculi, and the rice root aphid,

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Rhopalosiphum rufiabdominale, in organic celery showed 18 and 129% increase after two applications of azadirachtin or B. bassiana alone, respectively, while their combination resulted in a 62% reduction (Dara 2015b). Similarly, a better control of aphids (unknown species) and L. hesperus was achieved in strawberry by the combination of B. bassiana and azadirachtin or a higher label rate of azadirachtin alone than individual treatments of B. bassiana or a lower rate of azadirachtin (Dara 2016). When the invasive Bagrada bug, Bagrada hilaris, infested several vegetable crops in California, several organic growers who were advised to use the combination of B. bassiana and azadirachtin reported satisfactory control (Dara, pers. communication with farmers). Mohan et al. (2007) tested the compatibility of neem oil (with 1.5% azadirachtin) with 30 B. bassiana isolates from around the world. They found that some isolates of B. bassiana were sensitive to neem oil (with 0.15% azadirachtin) and among compatible isolates, one had a synergistic impact on the mortality of second instar larvae of the tobacco cutworm, Spodoptera litura. Combination of B. bassiana and azadirachtin also improved the control of the sweetpotato whitefly, Bemisia tabaci, on eggplant with improved egg and nymphal mortality and lower LT50 values, than their individual treatments, in China (Islam et al. 2010). Since immature stages of insects can escape fungal infection due to molting before cuticle penetration of the germ tubes, a combination of chemical or botanical IGRs with EPF can work in two ways: i) IGRs target immature stages while EPF target adults and susceptible immatures, and ii) IGRs interfere with molting and make immatures more vulnerable to fungal infections. Some pesticide manufacturers are now marketing B. bassiana formulations with pyrethrums and/or azadirachtin (e.g. BotaniGard MAXX, XPECTRO by LAM International) or pyrethrins with azadirachtin (e.g. Azera by Valent USA) among other active ingredient combinations. Laboratory studies in Russia found synergy between lower rates of M. robertsii (at 5 × 105 conidia/ml) and a commercial formulation of avermectins (metabolites of Streptomyces avermitilis at 0.005%) against L. decemlineata larvae (Tomilova et al. 2016). Avermectins significantly suppressed the insect immune responses and helped with increased fungal infections resulting in 1.2 to 1.6-fold increases in control efficacy. Niu et al. (2016) reported additive and synergistic interaction of S. marcescens with spirotetramat and thiamethoxam, depending on the concentration, in laboratory and greenhouse studies in China, against the third instar nymph of the brown plant hopper, Nilaparvata lugens. In a study conducted in the United States, Wang et  al. (2013) found out that feeding a low dose of the chitin synthesis inhibitor, lufenuron to the Formosan subterranean termite, Coptotermes formosanus, followed by the exposure to Pseudomonas aeruginosa had a synergestic effect. However, lufenuron did not have such an interaction with S. marcescens and Bt subsp. israelensis. Similarly, a chitinase producing bacterium, Paenibacillus sp. D1 and chitinase extracted from the bacterium, had a synergistic interaction with acephate against the cotton bollworm, Helicoverpa armigera, in laboratory studies in India (Singh et al. 2016).

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Laboratory, greenhouse, and field studies in California evaluated entomopathogen-­ based, botanical, and chemical pesticides as standalone applications or combinations and rotations to develop IPM strategies against various pests, with emphasis on the western tarnished plant bug, Lygus hesperus (Dara et al. 2013, 2018; Dara 2016). When a low rate of B. bassiana (0.2 lb./acre) was used with 1/5 the label rates of azadirachtin, fenpropathrin, naled, and thiamethoxam in the 2010 laboratory assays, the combination with fenpropathrin and thiamethoxam was higher than their individual treatments, during the first 3 days after application (Dara et al. 2013). In the follow up field studies in 2012, 2013 and 2014, various combinations and rotations of chemical pesticides, azadirachtin, pyrethrins, B. bassiana, M. brunneum and the bacterium Chromobacterium subtsugae were applied thrice at weekly intervals (Dara 2016). In the 2012 study, while L. hesperus numbers increased by 12% in untreated control by the end of the study, B. bassiana with fenpropathrin, azadirachtin, and 0.5 rate of acetamiprid caused 79, 69 and 67% reductions, respectively, compared to a 7.4% reduction by B. bassiana alone, 63% by novaluron + bifenthrin, and 86% by the full rate of acetamiprid. In the 2013 study, L. hesperus increased by 47% in untreated control, 17% in the chemical standard, acetamiprid, and 56% in B. bassiana  +  azadirachtin, followed by C. subtsugae and flonicamid. However, B. bassiana applied with low rates of acetamiprid, flonicamid, and avermectin in consecutive weeks, resulted in a 61% reduction in L. hesperus followed by 50% reduction from one application of novaluron + bifenthrin and two applications of B. bassiana + azadirachtin, and 47% from two applications of sulfoxaflor high rates followed by B. bassiana  +  C. subtsugae. Other chemical pesticide rotations provided 46–53% control in this study. In a 2014 study, none of the treatments was able to reduce L. hesperus populations and only their ability to limit the population buildup compared to pre-treatment counts could be observed. Three applications of the high rate of sulfoxaflor restricted the increase in L. hesperus to 14% and the high rate of diatomaceous earth (DE) followed by low rate of B. bassiana + acetamiprid and M. brunneum + azadirachtin to 17%, compared to 383% increase in untreated control and 1083% in the chemical standard of acetamiprid. Two applications of B. bassiana + azadirachtin after novaluron + bifenthrin limited the L. hesperus buildup to 54%. In a latter study, Dara et al. (2018) expanded the control options to multiple EPF (B. bassiana, I. fumosorosea, and M. brunneum), botanical pesticides (azadirachtin and pyrethrum), chemical insecticides from different mode of action groups, and mechanical removal by vacuuming. While a rotation of three chemicals (sulfoxaflor, flupyradifurone, and flonicamid) was the only treatment that reduced the L. hesperus numbers by 22%, the remaining treatments varied in their ability to limit or not limit the population buildup after three weekly applications of treatments. Compared to untreated control where L. hesperus increased by 190.8% post-­ treatment, the grower standard of acetamiprid limited the increase to 54.9%. While two sulfoxaflor applications followed by vacuuming resulted in the highest increase (403.6%), two flupyradifurone applications followed by vacuuming limited the increase to 25.5%. There was only 9.5% increase in L. hesperus numbers when a

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formulation of B. bassiana with pyrethrum was followed by vacuuming and ­application of novaluron + bifenthrin. Two other treatments that included two applications of M. brunneum or I. fumosorosea with azadirachtin products limited the pest build up to 59–79%. These data show that chemical pesticides are not always effective and EPF can be effectively used with other control options. Dara et  al. (2018) also observed, in a different study, that the combination of B. bassiana and bifenazate at the lowest label rates resulted in a 46.7% reduction in T. urticae after two applications compared to 39.9% by the full rate of bifenazate, 33.3% by abamectin, 47.2 and 50.9% by two formulations of fenpyroximate, and 62.4% by cyflumetofen. These extensive studies show that there are unlimited possibilities for chemical and non-chemical pesticide combinations and that all control options vary in their efficacy from time to time. They also demonstrated that EPF, when used with certain chemical or botanical pesticides at regular or lower rates, provided control similar or even superior to chemical pesticides. Synergistic combinations of EPF with DE and chemical insecticides have been effectively used for controlling a variety of stored-product pests as discussed in a recent review of Rumbos and Athanassiou (2017). While diatomaceous earth is abrasive to the insect cuticle and increase the chances of conidial adhesion and penetration, chemical insecticides weaken the insect and increase the chances of fungal infections. Some of the examples that Rumbos and Athanassiou discussed include DE with B. bassiana against the lesser grain borer (Rhyzopertha dominica), the red flour beetle (Tribolium castaneum), and weevils (Sitophilus spp.), DE with M. anisopliae against R. dominica and the rice weevil, S. oryzae, and fenitrothion with B. bassiana and M. anisopliae against S. oryzae. It is also important to understand and avoid antagonistic combinations such as B. bassiana with the acaricide, triflumuron (benzoylphenyl urea) which reduced the fungal efficacy against T. urticae (Irigaray et al. 2003). Similarly, imidacloprid was antagonistic to N. tanajoae (= N. floridana) and reduced the germination of primary conidia and formation of infective capilliconidia (Dara and Hountondji 2001). Studies conducted by Neves et al. (2001) evaluated the impact of neonecotinoids acetamiprid, imidacloprid, and thiamethoxam against B. bassiana, M. anisopliae, and Paecilomyces sp. at the field application rate, a rate above and below. Depending on concentration, these chemicals either improved or negatively affected the conidial production, germination and vegetative growth. One of the concerns for using EPF for pest control is their compatibility with fungicides in cropping systems where the latter are frequently applied for disease control. To save the cost of application, growers frequently use tank-mix pesticides and fungicides. While some combinations are compatible, others affected EPF. Multiple studies explored the compatibility of B. bassiana and M. anisopliae with fungicides (Samson et  al. 2005; Bruck 2009; Akbar et  al. 2012; Dara et  al. 2014; Roberti et al. 2017). Samson et al. (2005) reported adverse effect of methoxy ethyl mercuric chloride, flusilazole, prochlorax, and propiconazole on the readial growth of two isolates of M. anisopliae in vitro, but the conidial viability and ­recovery in the soil were not affected. They also found that when the greyback canegrub, Dermolepida albohirtum, and the negatoria canegrub, Lepidiota negatoria,

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were exposed to the soil treated with M. anisopliae and fungicides, the mortality of D. albohirtum was affected, but not that of L. negatoria. Bruck (2009) reported that several fungicides, except etridiazole, propamocard, and mafanoxam, significantly inhibited the conidial germination and mycelial growth of M. brunneum in vitro. When Norway spruce (Picea abies) cuttings were planted in soil containing M. brunneum and two applications of the fungicides were applied, there was no negative impact on M. anisopliae in bulk soil, but only captan and triflumizolet (which had shorter intervals between the applications) had a negative impact on M. anisopliae in the rhizosphere. Akbar et al. (2012) reported that a metalaxyl + mancozeb combination was highly toxic to the conidial germination and mycelial growth of M. anisopliae. In studies with B. bassiana and some common fungicides used in strawberry, Dara et  al. (2014) found that captan and thiram affected the fungus. When mealworm (Tenebrio molitor) larvae were exposed to B. bassiana with fungicides applied at different time intervals, captan caused a 43–70% reduction and thiram caused a 26–67% reduction in larval mortality regardless of the interval. Other fungicides (fluxapyroxad + pyraclostrobin, sulfur, pyraclostrobin + boscalid, myclobutanil, iprodione, and cyprodinil + fludioxonil) from different mode of action groups were compatible with B. bassiana (BotaniGard ES, strain GHA) resulting in 93–100% larval mortality. It also appeared that larval mortality was higher in treatments with fluxapyroxad + pyraclostrobin and cyprodinil + fludioxonil, during the first 2  days after treatments, compared to B. bassiana alone. In a different study, Roberti et al. (2017) observed that mepanipyrim and spiroxamine reduced B. bassiana (Naturalis, strain ATCC 74040) colony growth by 9.7 and 6.9%, respectively. When the fungicides and B. bassiana were sprayed on zucchini plants for controlling T. vaporariorum, boscalid + pyraclostrobin and cyprodinil + fludioxonil affected the pathogenicity by reducing the efficacy against nymphs by 91 and 87%, respectively. These studies show that several fungicides are compatible with EPF depending on species and strains.

11.7  Conclusion As the world is gradually moving towards sustainable pest management approaches, entomopathogens are becoming increasingly popular as control options. To develop an effective and long-lasting IPM strategy, one should consider a variety of tools including chemical and non-chemical alternatives. Understanding their interactions to identify additive and synergistic combinations and avoiding antagonistic ones helps with effective pest management and reinforces trust in the potential of non-­ chemical alternatives.

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Tomilova, O. G., Kryukov, V. Y., Duisembekov, B. A., Yaroslavtseva, O. N., Tyurin, M. V., Kryukova, N. A., Skorokhod, V., Dubovskiy, I. M., & Glupov, V. V. (2016). Immune-physiological aspects of synergy between avermectins and the entomopathogenic fungus Metarhizium robertsii in Colorado potato beetle larvae. Journal of Invertebrate Pathology, 140, 8–15. United States Department of Agriculture-National Agricultural Statistics Service (USDA-NASS) (2016). Organic survey (2014). In T. Vilsack & J. T. Reilly (Eds.), 2012 census of agriculture (pp. 592). Vergel, S. J. N., Bustos, R. A., Rodríguez, C. D., & Cantor, R. F. (2011). Laboratory and greenhouse evaluation of the entomopathogenic fungi and garlic-pepper extract on the predatory mites, Phytoseiulus persimilis and Neoseiulus californicus and their effect on the spider mite Tetranychus urticae. Biological Control, 57, 143–149. Wang, C., Henderson, G., & Gautam, B.  K. (2013). Lufenuron suppresses the resistance of Formosan subterranean termites (Isoptera: Rhinotermitidae) to entomopathogenic bacteria. Journal of Economic Entomology, 106, 1812–1818. Weseloh, R. M., Andreadis, T. G., Moore, R. E. B., Anderson, J. F., Dubois, N. R., & Lewis, F. B. (1983). Field confirmation of a mechanism causing synergism between Bacillus thuringiensis and the gypsy moth parasitoid, Apanteles melanoscelus. Journal of Invertebrate Pathology, 41, 99–103. Williams, C.  D., Dilon, A.  B., Harvey, C.  D., Hennessy, R., Mc Namaara, L., & Griffin, C.  T. (2013). Control of a major pest of forestry, Hylobius abietis, with entomopathogenic nematodes and fungi using eradicant and prophylactic strategies. Forest Ecology and Management, 305, 212–222. Wraight, S. P., & Ramos, M. E. (2005). Synergistic interaction between Beauveria bassiana- and Bacillus thuringiensis tenebrionis-based biopesticides applied against field populations of Colorado potato beetle larvae. Journal of Invertebrate Pathology, 90, 139–150. Wu, S., Xing, Z., Sun, W., Xu, X., Meng, R., & Lei, Z. (2018). Effects of Beauveria bassiana on predation and behavior of the predatory mite Phytoseiulus persimilis. Journal of Invertebrate Pathology, 153, 51–56. Yaroslavtseva, O. N., Dubovskiy, I. M., Khodyrev, V. P., Duisembekov, B. A., Kryukov, V. Y., & Glupov, V.  V. (2017). Immunological mechanisms of synergy between fungus Metarhizium robertsii and bacteria Bacillus thuringiensis ssp. morrisoni on Colorado potato beetle larvae. Journal of Insect Physiology, 96, 14–20.

Chapter 12

Toxicological Prospects on Joint Action of Microbial Insecticides and Chemical Pesticides A. R. N. S. Subbanna, J. Stanley, V. Venkateswarlu, V. Chinna Babu Naik, and M. S. Khan

Abstract  Microbial insecticides or entomopathogens are effective and eco-friendly insect pest management options. But slow mode of action and lack of a visual pest control, as expected by a farmer, mostly limits their wide commercial usage. The present day regular and high incidences of insect pests, due to intensive monocultures, warrant inevitable use of high doses of chemical pesticides. However, their judicious application depends on the diverse environmental threats associated. So, deployment of both entomopathogenic microbes and chemical pesticides together is considered to reduce the risk to the environment. Various studies also reported more efficient synergistic interactions in combined use than for independent applications. Synergism has the ability to reduce the pesticide doses. Most importantly, the combined application due to synergism can effectively tackles the pest problem and also helps in establishment of an entomopathogen in a given ecosystem. Once estabA. R. N. S. Subbanna (*) Crop Protection Section, ICAR-Vivekananda Institute of Hill Agriculture (VPKAS), Almora, Uttarakhand, India Department of Entomology, College of Agriculture, Govind Ballabh Pant University of Agriculture and Technology (GBPUA&T), Pantnagar, Uttarakhand, India e-mail: [email protected] J. Stanley Crop Protection Section, ICAR-Vivekananda Institute of Hill Agriculture (VPKAS), Almora, Uttarakhand, India V. Venkateswarlu Division of Crop Protection, ICAR-Central Tobacco Research Institute, Rajahmundry, Andhra Pradesh, India V. Chinna Babu Naik Division of Crop Protection, ICAR-Central Institute of Cotton Research, Nagpur, Maharashtra, India M. S. Khan Department of Entomology, College of Agriculture, Govind Ballabh Pant University of Agriculture and Technology (GBPUA&T), Pantnagar, Uttarakhand, India © Springer Nature Switzerland AG 2019 M. A. Khan, W. Ahmad (eds.), Microbes for Sustainable Insect Pest Management, Sustainability in Plant and Crop Protection, https://doi.org/10.1007/978-3-030-23045-6_12

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lished, the entomopathogens can effectively manage the pest population build up in an eco-friendly manner, and over the years they can evade the use of pesticides or, if not so, reduce their dosage. The present chapter critically discusses possible synergism between entomopathogens and chemical pesticides and the present status of pest management achieved through this approach, in the context of latest research findings. Keywords  Entomopathogens · Chemical pesticides · Joint action · Compatibility · Synergism

12.1  Introduction Insect pests continue to be limiting factors of agricultural production destroying an estimate around one-fifth of total global crop production annually, making crop protection an inevitable issue for agricultural production systems. An annual investment of US$ 40 billion on 3 million metric tonnes of pesticides (Atwal and Dhaliwal 2015) was unable to manage this level of damage. Application of this huge amount of pesticides raised complicate environmental and human health issues, besides becoming economically demanding against resistant populations of pest species. Despite of all these concerns, from the farmers’ point of view chemical pesticides are the competent pest management options available, due to their easy accessibility, storability, reliability of results, ready to use formulations and, especially, the ‘fast solution’ they provide against any pest problem. Keeping in view the different environmental threats associated with the use of chemical pesticides, the concept of pest control was tuned towards pest management by combination of different tactics for pest suppression. This underpins the idea of Integrated Pest Management (IPM) based on economic thresholds. The essence of IPM is the use of all the tactics of pest suppression including cultural, mechanical, physical, biological and chemical methods. In the present context of increased pest problems, in most cases IPM relies on mixtures or sequential applications of pesticides, with different modes of action. Although non-intensive, the application of alternative non-chemical tactics require through knowledge about biology and ecology of target pests, which might also be location-specific. So, use of pesticides is continued to be a key option for pest management in current chemical intensive cropping systems. Besides, pesticides are a global million dollar industry and in situations like emergencies and epidemics they are the only quick option available. Still, their use should be minimized to safeguard both biotic and abiotic environments. Although some chemical synergists are reported to improve bioefficacy and lower field doses, their impact on environment is also deleterious. Insect pathogens such entomopathogenic bacteria (EPB), fungi (EPF) and nematodes (EPN) are thought to be coevolved with insect pests and have the capability to cause epizootics. They are considered as one of the major natural regulation factors, maintaining the pest population density in a given ecosystem. Biological control using entomopathogens is considered as a biologically safe and environment-­

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friendly alternative to pesticides. In organic and sustainable agriculture systems, these approaches represent the major strategy of pest management and are gradually gaining significance. Although potentially bioactive, major drawbacks is their slow and imperceptible mode of action. Besides, indiscriminate use of pesticides definitely over mask their existence and sometimes may be also lethal. It is also important to note that some strains of entomopathogens are capable to resist lethal action of pesticides, sometimes found to be compatible or even synergizing each other in killing target pest (Feng and Pu 2005; Jia et al. 2016; Ali et al. 2017; Meyling et al. 2018). Entomopathogens, besides being direct lethal agents, can also be used as biosynergists to chemical pesticides. A clear understanding of biology, joint ecology and resulting physiological pathogenicity effects may give an idea about their combined utilization in successful pest management programs. The insecticide resistance developed by the majority of key pests also demands an alternative strategy with improved efficacy. It is also known that resistance against entomopathogens is ostensible in insect pests and can provide stable pest control under favorable environments. Keeping in view the reliability of both entomopathogens and chemical pesticides, their joint application offers an excellent strategy against many pest problems with reduction in doses (Wang et al. 2013; Singh et al. 2016). In this chapter, we discuss the possible combinations, synergism and modes of action of synergism, apart from further implications and knowledge gaps.

12.2  Methodologies Used for Compatibility Analysis In vitro assays are the first dependable methodology to envisage the levels of compatibility between two pesticidal components. However, dealing with a broad range of unicellular (i.e. EPBs), multi-cellular (EPF and EPN), non-cellular [entomopathogenic viruses (EPVs)] and symbionts (EPN), different entomopathogenic groups warrant specific methodologies for compatibility assays. The laboratory compatibility of common microbes with that of pesticides is measured by median inhibitory concentration (IC50) (Stanley and Preetha 2015). This section portrays different methodologies adopted by various authors in compatibility analysis between diverse entomopathogens with chemical insecticides. (i) For EPF In general, compatibility of EPF with insecticides was estimated by evaluating conidia germination after their incubation (about 24 h) in required concentrations of the pesticide to test. Alizadeh et al. (2007) reported use of field recommended doses (FRD) of insecticide at 0.5× FRD, 1× FRD and 2× FRD, in combination with conidia (at 106 conidia/ml) in potato dextrose broth at 25 ± 2 °C. After 24 h, number of conidia germinated out of total counted was estimated to get percent germination in comparison with the control treatment, without insecticide used. Yii et al. (2015) proposed a plate assay where a 107 conidia/ml solution was evenly spread onto dif-

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ferent concentrations in intoxicated potato dextrose agar (PDA), and allowed to germinate. Subsequently, 5–6 replications of 1 cm2 discs were cut and counted for germination percent. Poisoned food technique (Moorhouse et al. 1992) is one of the most widely used methodologies to estimate effects of pesticide poising on vegetative growth of fungi. In this technique, a required concentration of pesticide is aseptically mixed with warm PDA (± 45 °C) and plated in a petri dish. After solidification, a disc (~8 mm) of actively growing fungal mycelium is placed at the center (Stanley et al. 2010) and allowed to grow at 25 ± 2 °C. The data on radial growth of fungal mycelium is measured either at regular intervals or when the mycelium in control plates reaches borders, to estimate percent inhibition. Based on conidia germination, radial vegetative growth inhibition and inhibition in conidia production on intoxicated media, Alves et al. (2007) proposed the following biological index (BI) formula, to categorize toxicological effects of pesticides on EPF.



BI =

10 ( GR ) + 47 ( VG ) + 43 ( SPR ) 100



in which GR  =  reduction in conidia germination, VG  =  reduction in vegetative growth, and SPR = reduction in conidia production or sporulation. All these parameters were initially corrected for the controls and then fed to the formula for calculating BI. A BI value less than 42 represents toxicity, between 42 and 60 is considered moderately toxic, and above 60 represents compatible EPF and pesticide. The BI formula represents the actual toxicity status of a given pesticide by considering all the three growth stages of EPF, i.e. germination, vegetative growth, sporulation to an extent of 10, 47 and 43%, respectively, instead of considering their maximum levels. Due to this the formula is widely accepted and used in estimating compatibility between EPF and pesticides (Silva et al. 2013; Yii et al. 2015). (ii) For EPN The toxicological effects of pesticides on EPNs are manifested in the form of reduced viability and infectivity (Negrisoli et al. 2010a). Viability of EPN was estimated by incubating them in an equal volume of test concentration of the pesticide. After 48 h, the viability is estimated under a stereomicroscope by observing movements of infective juveniles (IJs). The infectivity of IJs at different incubation periods is estimated after removing the pesticide component by repeated washings and decanting. Thus obtained IJs are used for bioassay against Galleria mellonella (about 250 IJs per 10 final instar larvae) and compared with control mortality, once confirming the infection by EPNs after cadaver dissection. The toxicological effects (E %) of pesticides on EPN are calculated as follows.

E% = 100 - (100 - %corrected mortality ) × (100 - RI )

where RI = reduction in infectivity of IJs, which was calculated as



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RI = (1 - M t / M c ) × 100

321



where Mt = mortality in treatments and Mc = mortality in control. The E% value represents the toxicological classification of test pesticide viz., < 30% is non toxic, between 30–79% is slightly toxic, between 80–99% is moderately toxic and > 99% is highly toxic (Negrisoli et al. 2010a). (iii) For EPB No standard and specific protocols were reported to estimate compatibility of EPBs with chemical pesticides. However, some qualitative techniques as that of EPF by intoxicated media are reported (Mishra and Tandon 2003). A required insecticide concentration was mixed in a specific growth medium, after solidification, 100 μL of approximately 104 cfu/ml solution was evenly spread on to plates. After incubation, the colonies formed were counted and compared with control plates. For Pseudomonas, its fluoresce nature was measured every 24 h of incubation for 3 days and compared with that of the control (Stanley et al. 2010). In a similar qualitative assay, specific media is spread with 100 μL of bacterial culture or mixed with 2% of actively growing bacterial cells and, after solidification, a paper disc impregnated with required concentration of pesticide is placed above. After incubation, any inhibition zone of bacterial growth in and around the disc represents incompatibility.

12.3  Methodologies Used for Estimation of Joint Action Majority of studies reported joint action of entomopathogens with commercial formulations of insecticides. The entomopathogens are laboratory prepared for their infective stages viz., colony forming units (CFUs) for EPB, conidia for EPF, IJs for EPN and POBs for EPV. Different combinations are tested against lepidopteran, homopteran and coleopteran pests using specific Insecticide Resistance Action Committee (IRAC) approved methods. In all the bioassays, especially with EPB and EPF, a detergent (i.e. Tween 80 or Triton X100 at a concentration of 0.01%) is included for measuring equal dispersal of conidia or CFUs and wetting of feed surface area. Against termites, Yii et  al. (2015) proposed a bait formulation for estimating joint action of EPF and insecticides. In this bioassay, sawdust was initially immersed in required concentrations of an insecticide, following air drying the saw dust was mixed with dry harvest conidia at required concentration. Thus prepared bait formulation was fed with worker termites and data on mortality was recorded up to 8 days. The dead insects were also kept for development of mycosis. Wang et al. (2013) also used similar methodology by using EPB (Pseudomonas aeruginosa, Serratia marcescens and Bacillus thuringiensis) with a stack of 6 filter paper discs (9 cm diameter) as feed instead of sawdust baits.

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M E = M I + M F (1 - M I / 100 )



Where MI = mortality by insecticide alone and MF = mortality caused by EPF alone. Mansour et al. (1966) proposed a cooperative virulence index (cf) to classify the joint action of insecticides as cf =



Mm - Mi - Me Mi - Me

Where Mm  =  mortality caused by mixture, Mi  =  mortality caused by insecticide alone, Me = mortality caused by the entomopathogen alone. This can also be written as co-toxicity factor, as given by Mansour et al. (1966) and Abbassy et al. (1979) as Co　 toxicity factor =

Observed mortality - Expected mortality Expected mortality



A co-toxicity factor value < −20 indicates antagonism, > 20 indicates synergism and between −20 to +20 indicates additive interaction. Although different co-toxicity and synergistic coefficients are proposed, Mansour’s cooperative virulence index was widely used for estimating joint action between insecticides and entomopathogens (Zou et al. 2014). This is due to the use of lethal concentrations by former methods where an estimate of active ingredient (ai) is based on weight upon volume (mg/lt or mg/kg), which is suitable for chemical pesticides. But in case of entomopthogens active ingredients are CFU/ml (in case of EPB) or conidia/ml (in case of EPF) or POB/ml (in case of EPV). The Mansour’s method relies on mortalities of individual components and mixtures, to estimate joint action which fits with both insecticides and entomopathogens. Majority of the studies reported linear relationship between concentration-­ mortality (CM). However, time factor is also important to realize the effects of toxin components due to the use of lowered concentrations than those recommended (Nowierski et al. 1996). Feng and Pu (2005) proposed a time–concentration–mortality (TCM) modeling, which is both statistically and biologically robust, to estimate synergistic interactions between insecticide and entomopathogens. In some instances, the TCM model showed an acceptable goodness of fit with reliable estimates of LC50 and LC90, where the commonly used CM model based on probit analysis failed, due to uncontrolled bias. Besides, it is also important that scaled-up range of toxins should be investigated for realizing the exact interaction effects between pesticides and entomopathogens. Using overdoses of insecticide easily overwhelm the activity of entomopathogen counterpart and at the same time the maximum mortality realized cannot exceed 100% in any combinations, which may lead to misinterpretation of antagonism. Feng and Pu (2005) also indicated that sublethal doses of pesticides below LC50 should be considered first, and reported a reduction in imidacloprid field dose from 300 to 30 g ai/ha in combination of EPF, Beauveria bassiana, to realize a similar effect.

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Different authors used various combinations and mixtures of insecticides and entomopathogens for field evaluations. For example, Negrisoli et al. (2010b) used half the recommended concentrations of insecticides and entomopathogens in combination. Morales-Rodriguez and Peck (2009) used ½ and ¼ doses of pesticides against white grub species, Amphimallon majale and Popillia japonica. Feng and Pu (2005) used sublethal doses of the insecticide imidacloprid against Nilaparvata lugens. Niu et al. (2018) evaluated approximate LC20 concentrations of S. marcescens (108 to 109 CFU/ml) with two insecticides (spirotetramat and thiamethoxam), at LC25 and LC50 concentrations, against N. lugens, with mortality data up to 9 days. The primary objective of using mixtures of insecticides and entomopathogens is the minimization of chemical pesticides and promotion of entomopathogens activity, in a given ecosystem. In view of this we recommend field applications/evaluations of mixtures containing insecticides at their sublethal doses at which they can simply act as stressor against target pest, to enhance the infectivity of the entomopathogen component rather than acting as one of the lethal factor.

12.4  Compatibility Between Entomopathogens with Chemical Pesticides Studies on the compatibility between targeted combinations of entomopathogens and pesticides under in vitro conditions are one of the preliminary requirements to assess the fate of both pest management options. However, mixing entomopathogens (biological organisms) with toxic chemical compounds, in a huge variety of modes of action, appears as a complicated issue and requires through evaluation (Zou et al. 2014). Field application of bacteria, fungi and nematodes usually relied upon environment-resistant stages (i.e. spores, conidia and infective juveniles), which usually show a minimal tolerance level to different stresses including toxicological compounds like pesticides. However, their combined use requires through investigations on retaining their native activity, survival and bioactivity. Many researchers proposed different methodologies/techniques (discussed in previous headings) which targeted these basic issues. Actually, most compatibility studies were reported with fungal bioagents. Conidia viability, growth of mycelium in intoxicated media, further sporulation and infectivity are the critical factors considered for fungal bioagent compatibility (Alves et al. 2007). Asi et al. (2010) reported that vegetatively growing fungi are more resistant than their conidial stages. However, variations in methodologies may contribute to differential results. For example, Yii et al. (2015) reported that solvent used for insecticides, i.e. acetone, significantly reduced spore germination and increased vegetative growth and spore yield of Metarhizium anisopliae. Similar alterations in results can be expected when using commercial formulations which contain a variety of additives (Anderson and Roberts 1983), apart from the technical a.i. tested. Studies also reported that entomopathogenic nematodes are sensitive to

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surfactants which are commonly used in most commercial formulations of insecticides (Kaya et al. 1995; Krishnayya and Grewal 2002). The in vitro compatibility tests generally use direct exposure of bioagents to high concentrations of toxin, as well as allowing long term contact. The latter is mostly not a situation often encountered under field conditions (Neves et  al. 2001). Additionally, fate of an insecticide under field conditions (half life, degradation factors, plant absorption, relative contact with bioagent etc.) ultimately decides its deleterious effects on bioagent. As a consequence, the realized laboratory toxicity does not always mean that the pesticide is detrimental. Sometimes, the mode of action of a given pesticide can also signify possible impairment to the target bioagent. For example, abamectin damages sensory organs of nematodes leading to poor infectivity of IJs (Head et al. 2000). The primary constituents of nematode cuticle include collagens, cuticulins and other proteins (Negrisoli et  al. 2010a, b) which are not affected by pesticides with chitin synthesis inhibition activity (i.e. triflumurom, diflubenzuron etc.) (Hara and Kaya 1982; Rovesti and Deseo 1990; De Nardo and Grewal 2003). Similarly, the presence of butyrylcholinesterase, instead of acetylcholinesterase, in synapses of EPNs makes them compatible with organophosphous insecticides such as chlorpyrifos (Zimmerman and Crashaw 1990; Selkirk et  al. 2001; Alumai and Grewal 2004; Gutierrez et al. 2008). Such interpretation in respect of basic knowledge about the pesticide and bioagents may avoid tedious experimental procedures on compatibility and joint action. Various studies on compatibility reported contentious results with respect to same species of bioagents and insecticide. The reason is mainly related to the use of native strains or isolates, which may have an innate capacity to counteract the toxic effects resulting from environment and local isolation conditions. Such physiological mechanisms can be expressed in the form of increased survival (Shumacher and Poehling 2012), reproduction and even utilization of pesticides or metabolites as nutrients (Moino and Alves 1998). Adaptability of a bioagent to toxic chemicals is also an important factor which warrants time lapse studies and further understanding of biological adaptations. The toxicological effects of pesticides on bioagents is manifested in form of reduced spore viability, inhibition in germination, lower vegetative growth, limited sexual reproduction or loss of infectivity etc. All these noxious effects appear dose-­ dependent and require intensive investigations to understand dose-damage relationships. These studies may also provide required informations about temporal and spatial separation of both pest management options, for synergism to be observed.

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12.5  J oint Action of Entomopathogens with Chemical Pesticides In general, most studies reported the use of compatible chemical pesticides for further evaluation of their joint action against insect pests. However, the negative results in laboratory compatibility tests may not always lead to poor biotoxicities, especially under field conditions, which is governed by a variety of ecological and biological parameters (some examples of positive and negative interactions between entomopathogens and pesticides are detailed in Table 12.1 and 12.2, respectively). Moreover, use of minimal concentrations of chemical pesticides ― approximately between LC10 - LC30 ― reported high synergistic interactions under both laboratory and field conditions. Field evaluation of these negative interactions or further testing at lesser dosage of pesticide may hence yield a real picture of bioactivities. Joint action studies should hence be planned with a range of pesticide and bioagent doses starting from lower sublethal doses to a maximum of individual mortality up to 50%. It is also important to note that no generalized behavior can be expected from single chemical group of insecticides to a single bioagent. Differential synergistic interactions between imidacloprid and clothianidin (neonicotinoid group) against P. japonica and A. majale (Morales-Rodriguez and Peck 2009) or of imidacloprid and thiamethoxam with Heterorhabditis bacteriophora against Cyclocephala hirta, C. pasadenae and Exomala orientalis (Koppenhöfer et al. 2002) evidently showed this feature. A synergistic joint action can also be achieved by a temporal separation of the insecticide and bioagent (Negrisoli et al. 2010b). Meyling et al. (2018) reported synergism of entomopathogens and insecticide as a function of sequence and time of application. Conventional mortality analysis carried out using factorial experiments may provide data on target pest mortality over a fixed and minimal time period (generally 72 h). However, the lethal action of both mortality factors is a function of the interactions between time and concentration (Nowierski et  al. 1996). Most studies ignored time factors, which may yield a bias when compared with individual mortalities (Nowierski et al. 1996; Feng and Pu 2005). This bias is greatly overcome by a robust TCM relationship, that takes into account the time and concentration into a single model, enabling us to separate the interaction effects from the individual effect of time and concentration on pest mortality. This estimate of TCM is not only robust from the mathematical point of view but also have a high biological relevance on the relative potency of both the entomopathogen and the chemical. The median lethal time (LT50) is estimated by linear interpolation in such TCM models. Relative potential are also tested using LC50 values of entomopathogens and chemicals (Feng and Pu 2005). In case of virus-insecticide mixtures, the standard probit or logit analysis do not holds good as the concentration mortality data do not follow a logistic or Gaussian distribution, due to the differences in the mode of action and/or interactions between the virus and the insecticide.

S. carpocapsae

H. indica, S. carpocapsae, S. glaseri

H. bacteriophora, H.megidis, H. marelatus, S. glaseri, S. feltiae. Heterorhabditis sps., S. carpocapsae

Synergism was discernible from laboratory, greenhouse and field trials

Morales-­ Rodriguez and Peck (2009) Polavarapu et al. (2007) Koppenhöfer et al. (2002)

Amphimallon majale, Popillia japonica A. orientalis

Additive (H. indica with cypermethrin, spinosad, methoxyfenozide and deltamethrin+triazofos; S. carpocapsae with lufenuron, chlorpyrifos and cypermethrin) and synergistic (H. indica with chlorpyrifos; S. glaseri with chlorpyrifos and lufenuron) Synergism

Negrisoli et al. (2010a)

S. frugiperda

Azadiractin

Western flower thrips

Compatible with both nematode species

Sabino et al. (2014)

Tuta absoluta

Otieno et al. (2016)

Koppenhöfer et al. Synergism (2002)

Triflumuron, Deltamethrin, dimethylamino-propyl, lambda-­ Cyhalothrin + chlorantranilprole, Clorantranilprole, Thiamethoxam + lambda-cyhalothrin. Chlorpyrifos, Cypermethrin, Spinosad, Methoxyfenozide, Deltamethrin+Triazofos, Lufenuron

Imidacloprid

P. japonica, Cyclocephala hirta, C. pasadenae, E. orientalis E. orientalis

Synergism was with low dose but not with high doses Imidacloprid provided stronger and more consistent synergism with nematodes than thiamethoxam.

Remarks

References

Target Pest

Imidacloprid Thiamethoxam

Imidacloprid

Entomopathogens Insecticide Entomopathogenic Nematodes H. bacteriophora Imidacloprid and Clothianidin

Table 12.1  Example of positive interactions between entomopathogens and insecticides

326 A. R. N. S. Subbanna et al.

M. brunneum

Rice stalk stink bug

Blatella germanica

Diaprepes abbreviates Diaprepes abbreviates C. bergi

Coptotermes curvignathus Popillia japonica

German cockroach

Endosulfan, Imidacloprid, Lufenuron, Spilarctia obliqua diflubenzuron, Dimethoate, Oxydemeton methyl Imidacloprid Anoplophora glabripennis

Thiamethoxam

Imidacloprid

Imidacloprid

Imidacloprid

Imidacloprid, Clothianidin

Chlorpyrifos, Propetamphos, Cyfluthrin Fipronil

Entomopathogens Insecticide Target Pest Entomopathogenic fungi M. anisopliae Permethrin, Imidacloprid, NeemAzal, NA Amitraz Thiamethoxam, lambda-cyhalothrin Rice pests

Russel et al. (2010)

Morales-­ Rodriguez and Peck (2009) Quintela and McCoy (1997) Quintela and McCoy (1998b) Jaramillo et al. (2005) Pachamuthu and Kamble (2000) Quintela et al. (2013) Purwar and Sachan (2006) Compatible

Synergism

Synergism

(continued)

Low conidial attachment to larval cuticle at higher dose of chemical

Synergism

Discernible in the laboratory and greenhouse, but not in field.

Greatest synergism

Compatible with conidial germination, vegetative growth and sporulation No effects on conidial germination, vegetative growth and sporulation Additive and syngergistic

Schumacher and Poehling (2012) Silva et al. (2013) Pachamuthu and Kamble (2000) Yii et al. (2015)

Remarks

References

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M. Anisopliae, B. bassiana B. Bassiana

Entomopathogens M. anisopliae, B. bassiana M. anisopliae, B. bassiana, Paecilomyces M. Anisopliae, B. bassiana

Table 12.1 (continued)

NA

Tenebrio molitor

Spodoptera litura Spilarctia oblique Popillia japonica

Diaprepes abbreviatus

Imidacloprid

Alpha-cypermethrin

Neem

Endosulfan

Imidacloprid and clothianidin

Imidacloprid

Imidacloprid, Fipronil

Amphimallon majale and Popillia japonica NA

Target Pest Western flower thrips NA

Imidacloprid, Clothianidin

Acetamiprid, Imidacloprid, Thiamethoxam

Insecticide Azadiractin

Mohan et al. (2007) Purwar and Sachan (2006) Morales-­ Rodriguez and Peck (2009) Quintela and McCoy (1998b)

Morales-­ Rodriguez and Peck (2009) Moino and Alves (1998) Neves et al. (2001), Alizadeh et al. (2007) and Abidin et al. (2017) Meyling et al. (2018)

Low conidial attachment to larval cuticle at higher dose of chemical

Discernible in the laboratory and greenhouse, but not in the field.

4.9 times increased toxicity

Synergistic effect when only the interval between applications was >48 h. with 72 h between exposures, mortality had increased to 100% after 8 days Compatible and synergistic

No effects on conidial germination, vegetative growth and sporulation

No negative effect

Synergism

References Remarks Otieno et al. Synergism (2016) Neves et al. (2001) No negative effect on conidia germination, growth and vegetative growth

328 A. R. N. S. Subbanna et al.

Khalique and Amhed (2005) Niu et al. (2018) Nathan et al. (2004)

Bemisia tabaci B. tabaci

H. armigera

Matrine

Spirotetramat, Acetamiprid, Imidacloprid, Thiamethoxam

Spirotetramat, Thiamethoxam Botanicals

Entomopathogenic virus Nuclear polyhedrosis Spinosad virus Nuclear polyhedrosis Spinosad virus Nuclear polyhedrosis Carbamates, Methomyl and virus Pyrethroids

S. marcescens B. Thuringiensis

Synergism

Synergism

Mendez et al. (2002) Mccutchen et al. (1997)

S. frugiperda Heliothis virescens

Reduction in lethal dose

Khattab (2007)

Synergism observed at high thiodicarb and low Bt-toxin concentration Both synergistic and additive effects Synergism

Compatible

Lethal time and pathogenicity increased

Synergism

Remarks Synergism

S. littoralis

Nilaparvata lugens Cnaphalocrocis medinalis

NA

Zou et al. (2014) and Tian et al. (2015) Vyas et al. (1992)

Ephestia kuehniella

Botanical compounds

References Purwar and Sachan (2004) Shakarami et al. (2015) Ali et al. (2017)

Target Pest Lipaphis erysimi

Insecticide

B. brongniartii, M. Azadiractin anisopliae Entomopathogenic bacteria B. thuringiensis Thiodicarb

Lecanicillium muscarium Isaria fumosorosea

Entomopathogens

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Table 12.2  Examples of negative interactions between entomopathogens and insecticides Entomopathogens H. bacteriophora, S. carpocapsae

M. anisopliae

Insecticide Thiophanatemetil, Thiametoxam, Imidacloprid, Aldicarb, Carbofuram Fipronil Imidacloprid

B. Bassiana

Imidacloprid

Deltamethrin, Chlorpyrifos, Thiodicarb, Imidacloprid, Cypermethrin Fenitrothione

References Negrisoli et al. (2008)

Remarks Reduced the viability and infectivity of EPNs

Schumacher and Poehling (2012) Abidin, et al. (2017) and Shumacher and Poehling (2012) James and Elzen (2001) and Abidin et al. (2017) Abidin et al. (2017)

Moderate toxicity at higher doses Moderately inhibited the conidia germination and vegetative growth Antagonism

Conidia germination inhibited

Zibaee et al. (2009) Fungal infection decreased the susceptibility.

In view of available data, joint action studies of different bioagents with pesticides resulted in maximum additive and few antagonistic or synergistic interactions (Morales-Rodriguez and Peck 2009). Bacterial bioagents yield minimal number of synergistic interactions and are very particular with pesticide. Overall maximum compatibility and synergism responses of bioagents was in the order: EPN > EPF > EPB. At the same time, maximum number of studies was in the order of EPF  >  EPN  >  EPB.  The strength of synergism diminishes from laboratory to greenhouse, to field (Morales-Rodriguez and Peck 2009). Amongst EPB, B. thuringinsis is well studied, having more than a century of history in pest management. However, only minimal studies reported its synergism with chemical pesticides (Chen et al. 1974; Salama et al. 1984; Morales-Rodriguez and Peck 2009; Subbanna et al. 2019). With respect to chemical groups, B. thuringiensis is compatible and showed more synergism with pyrethroids than with carbamates and organophsphous insecticides (Salama et al. 1984). The carbamate group had no significant effect on bacterial growth at lower concentrations, and no significant synergism was observed (Sutter et al. 1971). Likewise, the sister species, B. popilliae and B. sphaericus were found to be susceptible to most commonly used pesticides at high concentrations (Dingman 1994; Mishra and Tandon 2003). Other insect pathogens such as Serriatia marcescens should be studied for synergistic properties in view of their reported field performance, along with spirotetramat or thiamethoxam (Niu et al. 2018). EPFs are highly variable in their synergistic potential with pesticide groups, which is a function of the strain or isolate under testing. Abidin et al. (2017) opined

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that the EPF adaptability to toxins should also be taken into consideration in designing joint applications. Neonicotinoids (especially imidacloprid) are reported to be synergistic with EPFs against the white grubs such as P. japonica (Morales-­ Rodriguez and Peck 2009), the weevil Diaprepes abbreviatus (Quintela and McCoy 1997, 1998a), the caterpillar Spilarctia obliqua (Purwar and Sachan 2006), the bug Cyrtomenus bergi (Jaramillo et  al. 2005), the termites Heterotermes tenuis (and Reticulitermes flavipes (Ramakrishnan et  al. 1999; Moino and Alves 1998), the German cockroach (Kaakeh et  al. 1997), the burrower bug Cyrtomenus bergi (Jaramillo et al. 2005) and the rice stalk stink bug Tibraca limbativentris (Quintela et  al. 2013, with thiomethoxam). These synergistic interactions can reduce the insecticide dose from 25% to 14% or even less of the recommended rate (Feng and Pu 2005; Quintela et al. 2013). Sometimes, compatible insecticides may show positive correlation between insecticide dose and induced mortality. Other insecticide groups are also safe and can be used with EPF at appropriate concentrations (Li et al. 1996; Pachamuthu and Kamble 2000; Chen and Feng 2003). Synergistic interactions are also prominent between sucking pest specific EPFs and pesticides such as Isaria fumosorosea with spirotetramat (Zou et al. 2014), or Lecanicillium muscarium with matrine against Bemesia tabaci (Ali et al. 2017). The commercial use of EPNs is mainly concentrated towards white grubs management. Most joint action studies using EPNs and insecticides was restricted against white grubs (Negrisoli et  al. 2010b). In case of EPNs also, imidacloprid showed high synergistic interactions at appropriately lower doses. Higher doses of imidacloprid and nematodes sometimes showed some antagonistic interactions (Koppenhöfer et al. 2000; Polavarapu et al. 2007; Sabino et al. 2014). Nematode strain and species-dependent responses were highly prevalent. Major studied with white grub species involved P. japonica, Cyclocephala hirta, C. pasadenae, Anomala orientalis, Exomala orientalis, A. majale, Maladera castanea etc. All these species showed differential responses to varying combinations of neonicotioids with EPNs (Koppenhöfer et al. 2002; Koppenhöfer and Fuzy 2003). Minimal studies reported joint action of EPVs with insecticides. So the potential interaction of EPVs with pesticides is little understood. McCutchen et al. (1997) opined that insecticide acting on nervous system (such as pyrethroids, carbamates etc.) perform well along with EPVs even though their mode of action is not clear. Spinosad is also reported to act synergistically under field conditions against Spodopetra frugiperda (Mendez et  al. 2002) and S. littoralis (Khattab 2007) although minimal positive interactions were observed in laboratory. Majority of the antagonistic interactions in entomopathogen-insecticide complex are the result of high doses of chemical pesticides which may engulf the activity of the bioagent counterpart. At the same time it may not be safe to test only low doses, as often reported. It is also important to note the subsequent effects of an insecticide on a bioagent. For example, Russel et al. (2010) showed reduction in fungal mycosis and conidia yield by M. brunneum when combined with low doses of imidacloprid, although the mixture was synergistic against Anoplophora glabripennis. In EPNs this antagonism is manifested in the form of reduced parasitism (Baweja and Sehgal 1997). Besides, the laboratory antagonism may not always yield a negative

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response under field conditions. Neonicotinoid insecticides (imidacloprid and ­thiomethoxam) were also found to be antagonistic in many cases (James and Elzen 2001; Koppenhöfer et al. 2002; Morales-Rodriguez and Peck 2009). The joint action of entomopathogens was mostly studied with neem products such as commercially available formulations, extracts, seed kernel extracts, oils etc. The target pests used for estimating a joint action belonged to diverse groups, including sucking pests such as B. argentifolii (James 2003), Thrips tabaci (Mohan et al. 2007), B. tabaci (Shakarami et al. 2015) and Frankliniella occidentalis (Otieno et al. 2016), lepidopteran larvae such as Spodoptera litura (Shakarami et al. 2015) and storage pests such as Tribolium castaneum (Akbar et al. 2005). Most studies targeted the joint action of EPFs with neem products and supported improvements in lethal concentrations and time of kill (Mohan et al. 2007). Despite of these synergistic interactions, the active botanical insecticidal compounds, i.e. phytoalexines, triterpenoids and sulfurade compounds etc., have a mycotoxic capacity and other associated compounds may also influence germination and growth of bioagents. Entomopathgenic bacteria such as B. thuringiensis (Nathan et al. 2004), with nematodes (Otieno et al. 2016) and viruses (Muralibaskaran et al. 1999; Bhandari et al. 2009) also showed substantial synergistic activity with neem products, although dose- dependent. Extracts from other pesticidal botanicals from Annona squamosa, Prosopis juliflora, Eupatorium and Artemisia also showed relative synergism with different bioagents.

12.6  Mechanism/Mode of Action of Synergisms The combined mode of action of two different pest management options based on a pure biological and a chemical toxin is difficult to ascertain. However, some models and predictions can be done after knowing the individual biological targets of the associated components. In general, majority of the studies reported apparent decrease in lethal concentrations and time required to kill target pests. Both independent and combined actions of the components can explain the reduction in time required to kill. In independent action, the insecticides generally kills insect pest before 72 h of exposure, a period during which the bioagents colonize the site of action but are not lethal. Subsequent lethal actions by bioagents further improve mortality, which is not the situation in independent actions. On the other hand, biological explanations on synergistic joint action may involve similar target sites (nervous system, ion exchange channels etc), concurrent actions on different tissues or binding sites, diverse actions on the same tissue or different sites, and other physiological irregularities etc. Physiological interactions play an important role where one component wanes a biological system or pest mechanism, thereby enhancing or providing a way for an elevated lethal action of the associated component. For instance, any pathogenic infection to nerve cells improves their sensitivity to insecticides. Similarly, pathogenic S. marcescens strains (Ishii et al. 2012) kill host immune cells. A low dose of

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imidacloprid was found to affect the grooming behavior of soil inhabiting pests (termite, Heterotermes tenuis and white grub, Diaprepes abbreviatus) thus facilitating the attachment of EPF conidia and further infection (Quintela and McCoy, 1998a, 1998b). Type, concentration and formulation of an insecticide are the architectural factors governing the joint action and its extent. Other effects include extended developmental periods, reduced larval feeding, instar intermediates, abnormal pupae, malformed adults, reduced adult longevity and fecundity, poor survival of new born etc. which weaken insect biological systems promoting the toxic actions. However, reduced food intake is a physiological mechanism to resist or avoid the action of systemic insecticides, and some enterobacteria (Morales-­ Rodriguez and Peck 2009). Some physiological interactions induced by insecticides may also favor the entomopathogenic counterpart, thereby exerting synergism. Sublethal doses of some insecticides weaken the immune system, facilitating fungal invasion (Hiromori and Nishigaki 2001). Quintela et  al. (2013) reported that sublethal concentrations of thiamethoxam reduced fungistatic aldehyde production by rice stalk stink bug, thus improving infectivity by M. anisopliae. Some indirect interactions may also facilitate survival, development and infectivity of entomopathogens. For example, Anderson et al. (1989) reported growth enhancement of B. bassiana by some pesticides due to the adjuvants in the formulation, which helped in scattering of conidial piles thereby promoting propagule numbers. Similarly surfactants and solvents may also be involved in synergistic interactions. The basic biological functions of any given living organisms rely on enzymes, which is also true for insects. Any external toxicological intervention definitely alters different enzymes associated to target systems, viz. digestive tract, nervous system etc. In insects, tolerance towards toxic compounds is a function of the efficiency in their detoxification by carboxylesterases and glutathione-S-transferase (Claudianos et al. 2006). Other enzymes involved in repair of damage to biomembranes are superoxide dismutase, catalase and peroxidase. Enzymes such as chitinases are directly involved in defense against invading pathogens, especially against EPF. Studies reported high levels of these enzymes in response to pesticides and entomopathogen applications in different insects i.e. B. tabaci (Tian et al. 2015; Jia et al. 2016), Plutella xylostella (Luo and Zhang 2003) etc. The increase in enzyme activity is prominent during initial periods of application (mostly up to 72 h), later restrained due to overhaul by toxicity of the external agent (pesticides or entomopathogen). In joint applications the sublethal or even non-lethal doses of insecticides act as stressor facilitating the entomopathogen invasion (Vallet-Gely et  al. 2008). Metabolism of acetylcholine is also a primary target for most of the insecticides, particularly neurotoxic. Synergistic interaction between nerve toxins and entomopathogens can be explained by chemical disturbance in acetylcholine balance and its degrading enzyme, acetylcholine esterase (Liu et al. 2008; Zibaee et al. 2009; Ali et al. 2017). Insect immune system is governed by hemocytes, which attack any foreign bodies’ directly, and phenoloxidase cascade, a major defense of humoral reaction (for

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more details see Chaps. 8 and 13). Amongst hemocytes, the plasmatocytes and granular cells are responsible for phagocytosis and encapsulation of intruders. Hiromori and Nishigaki (2001) reported decrease in granulocytes and inhibition in humoral activity by reduced phenoloxidase activity in the whitegrub A. cuprea, in response to mixed application of M. anisopliae and an insecticide. Insect growth regulators used as insecticides also have synergistic interactions, causing disruptions in chitin lamellae thereby facilitating infection by EPF (Hassan and Charnley 1987, 1989).

12.7  Conclusion and Future Prospects Both compatibility and joint interactions of any given pesticide and entomopathogens, under both laboratory and field conditions, represent the outcome of complex systems and cannot be generalized. Their interaction is function of the dynamic uniqueness of both components. The determining factors of entomopathogen are species, strain/isolate, insecticide resistance level, existing biotic and abiotic conditions in the area of isolation, innate biochemical and physiological potency etc. The pesticide factors to be considered are chemical group, mode of action, concentration, type of formulation, ingredients of formulation etc. Target species, evaluation methods, timing of assay can also be influential parameters which are under the researcher’s hand. Scientific understanding of all these factors could predict the interaction but not the actual performance. Amongst entomopathogens, to the best of our knowledge no study reported compatibility and synergism of EPVs with insecticides which might be due to their obligate parasitism. However, they represent an environmentally competent group of biopesticides due to their ability to cause epizootics and ease of formulation preparations. Intensification of research efforts in this area is hence required. The synergistic interactions always showed waning trends towards field, which might be due to increased size of experimental units as well as other biotic and abiotic factors which are excluded under laboratory conditions. Correspondingly, some antagonistic and additive interactions also proved to be effective under field conditions. Although biochemical and physiological interactions of synergism are known, performance of minimum possible interactions, including antagonistic and additive effects under field conditions, could give a clear picture of a joint action between pesticides and entomopathogens. Similar studies on temporal and spatial separation of incompatible mixtures may also explain possible combinations for enhanced and ecofriendly pest management. The possible effects on non-target species and natural enemies should not be ignored, as the final applications are targeted to agroecosystems where a complex of species coexists. Understanding the entomopathogens and insecticides interplay may lead to possible new strategies for management. This strategy uses manifold and complex attack approaches in killing target insect pests. Besides, due to direct toxicity effects on pest populations after receded insecticidal activity of pesticides, the entomo-

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pathogen may establish on the residual host population offering subsequent pest management in targeted agro-ecosystem. This process may also eliminate subsequent pesticidal applications and safeguard the biodiversity. There is a possible development of resistant or tolerant strains of entomopathogens that can survive at high pesticide concentrations. These strains offer possible development of a composite commercial formulation containing combined preparations, that can also be used in insecticide resistance management. Despite of relative, partial successes, no study reported large scale field successes of joint applications, which needs to be done to exploit their full potential. Acknowledgments  This study was supported by the Indian Council of Agricultural Research (ICAR), New Delhi. Authors are thankful to Director, ICAR-VPKAS, Almora.

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Chapter 13

Entomopathogen and Synthetic Chemical Insecticide: Synergist and Antagonist Arash Zibaee

Abstract  The use of synthetic insecticides and biological agents are the main tools to control agricultural pests, each with its own advantages and disadvantages. Although the use of chemical compounds is inevitable in some cases, attitudes of consumers and agricultural experts towards healthy products and lower environmental contamination have increased the prevalence and preference for biological agents. Among them, insect pathogens have been considered as the unique and widely distributed components in many ecosystems, due to their diverse virulent mechanisms. The entomopathogenic fungi (EF) and nematodes have been commercialized as biologically active insecticides against a wide range of pests. Although many environmental benefits for these compounds have been identified, the disadvantages such as low virulence due to behavior or habitat of target pest, delayed killing performance and sensitivity to environmental factors, lead to simultaneous use of entomopathogens with one or more chemical insecticides in reduced doses. In this review, the possibility of simultaneous use of chemical insecticides from different classes with EF and nematodes were discussed by indicating severally up-to-­ date studies. The effects of insecticides on cessation or induction of germination and conidiation of fungi have been reported, depending on the concentrations of used chemicals. Field or laboratory experiments have shown synergism or antagonism of EF with some insecticides. In case of entomopathogenic nematodes, the effects of insecticides from different classes have been investigated on mobility and survival of nematodes, as well as on synergistic or additive effects. Generally, the possibility of simultaneous use of chemical insecticides with these two groups of entomopathogens depends on target pest, spraying method, insecticide class or formulation and origin of entomopathogens. Keywords  Entomopathogen · Insecticide · Interaction · IPM

A. Zibaee (*) Department of Plant protection, Faculty of Agricultural Sciences, University of Guilan, Rasht, Iran e-mail: [email protected] © Springer Nature Switzerland AG 2019 M. A. Khan, W. Ahmad (eds.), Microbes for Sustainable Insect Pest Management, Sustainability in Plant and Crop Protection, https://doi.org/10.1007/978-3-030-23045-6_13

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13.1  Introduction To produce healthy food for growing population of world has been one of the most important requirements in recent decades. In addition to limiting factors such as proper planting, climate and processing costs, insect pests, plant diseases and weeds have caused the significant negative effects on agricultural production (Oerke et al. 1994). Insects as main, occasional and secondary pests have had enormous impacts on the yields of agricultural products and induce much of production costs (Chandler et al. 2011). Although different methods have been applied to reduce insects economic impact over years, biological and chemical controls have been assigned the largest share (Pretty 2008; Lacey et al. 2015). Although the use of biological agents has led to production of healthy or so-called organic products, chemicals from different groups of organochlorine, organophosphorus, carbamates, neonicotinoids, insect growth regulators, botanical insecticides and other nature-based insecticides have been inevitable in many cases (Talebi-Jahroumi 2012). Data gained based on the essential chemical industry-online indicate that chemical insecticides comprise 24% of used agrochemicals, in which different classes have their own proportions from 3% to 22% (Fig. 13.1 a, b, c). Despite fast and efficient impacts of chemicals on suppressing pest population outbreaks, some issues led to limitations on their use as summarized by Chandler et al. (2011): (i) damages on environment and human during manufacturing, handling and application because of not judicious use of broad-spectrum insecticides, (ii) issues such as pest resurgence, emergence of secondary pests and development of heritable resistance following prophylactic use of insecticides, (iii) withdrawal of several groups of insecticides because of new health and safety legislations, (iv) concerns on chronic toxicity of insecticides due to residuals in food, on all consumers (human and livestock). Such concerns and disadvantages of synthetic chemical insecticides forced trends to use other control procedures against insect pests. One of the promising and long-lasting ways could be to use natural enemies in different agroecosystems. Although natural enemies may result in 50–90% efficiency of pest control in crop fields, potential application and reliability require the extended scientific knowledge and concerned economic and social attitudes (Pimentel 2005). Nevertheless, there are several advantages that make biological control as one of the major control procedures in sustainable agriculture including: (i) the high efficiency at low cost, (ii) self-perpetuation at little or no cost, (iii) free of harmful effects on non-target organisms, (iv) ability of biocontrol agents to rapid reproduction, (v) self-searching the target hosts, (vi) survival at relatively low host densities in addition to (vii) adaptability to several climatic conditions (Mason and Huber 2002; Neuenschwander et al. 2003; Omkar 2016; Khan and Ahmad 2015; Khan et al. 2016). Smith (1919) was the first to define the term of biological control (biocontrol) as the “top-down” action of natural enemies to equilibrate the density of pest population at a level lower than when they are absent. Later, DeBach (1974) represented it as “biocontrol is the use of living organisms/natural enemies to suppress the population density or impact of a specific pest organism, making it less abundant or less damaging than

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Fig. 13.1  The percentage of chemical insecticides used vs other chemicals and their different classes compared to groups of entomopathogenns. (a) Proportion of pesticide used in agroecosystems, (b) Proportion of different classes of insecticides used in agroecosystems, (c) Commercialization proportion of entomopathogens against insect pessts. (Based on: www.essentialchemicalindustry.org/materials-and-applications/crop-protection-chemicals. Html; The 2010 Worldwide Biopesticides Market Summary, vol. 1. CAB International Centre, Wallingford.)

it would otherwise be”. Based on these definitions, biological control relies on proper action of organisms which are known as biocontrol agent acting vs pests. Predators, parasites, parasitoids and entomopathogens are the biocontrol agents engaged in biological control tactics over decades. Each agent adopts its unique way to concur and subsequently decrease population density of hosts, but several factors such as their high adaptability, host specificity (monophagous), voracious feeding capacity, synchronized development with pest species, more spatial and temporal dispersal, multiplication capacity, short life cycle with multiple generations, unending host search capacity, being non-palatable for predators, and resistance to other pathogens and parasites do influence their appropriate effectiveness in (IPM) (Omkar 2016). Among the biocontrol agents, entomopathogens are the infectious microorganisms that invade insects through different portal of entries, e.g. integument and mouth, reproduce within hemocoel and spread to infect other individuals or insects. Spanning over the prokaryotic and eukaryotic Kingdoms they include viruses, bacteria, microsporidia, fungi and nematodes (Kaya and Vega 2012). These agents are used to control several insect pests and have shown to be successful

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depending on virulence, ease of application and production, low cost, good storage properties, safety to farmers, and colonization in environment (Lacey et al. 2015; Omkar 2016). Among the entomopathogens, the bio-insecticides based on the bacterium Bacillus thuringiensis Berliner and its derivate comprise the highest ratio of the market, with fungi in the second rank and nematodes in the least ratio (Fig. 13.1c). In this chapter, initially a background on use of entomopathogenic fungi (EF) and nematodes as well as their infectious mechanisms in insects will be presented, then, their limitations in biological control programs and reasons for integration with insecticides will be discussed by reporting the results of laboratory and field studies. Eventually, the synergistic and antagonistic mechanisms between EF/nematodes with chemical insecticides will be compared in several case studies.

13.2  Potential of Entomopathogenic Fungi Against Insects EF are the dominant pathogens among populations of insect pests which are almost ubiquitous in both terrestrial and aquatic ecosystems. These biocontrol agents have shown a high potential to regulate population densities of insects not only agricultural pests but also insect-vector like mosquitoes. EF cause infections throughout insect populations as obligatory to facultative pathogens but their applications in ecosystems might be possible through mass production, formulation and proper dosage in the framework of microbial insecticides (Lacey et al. 2015). EF are categorized into several groups that are located in the Hypocreales order of Ascomycota, including imperfect taxa such as Beauveria, Metarhizium, Nomuraea, Isaria. and Hirsutella spp. Other important species such as Entomophthora, Zoophthora, Pandora and Entomophaga spp. belong to the order of Entomophthorales from Zygomycota (Araujo and Hughes 2016; see Chaps. 2 and 3 for more details). Species belonging to Hypoceales such as Beauveria spp. and Metarhizium spp. are suitable for mass production and to be used against insect pests of crops, orchards, greenhouses and forests because of their possibility to grow on low-cost cultivated environments, utilizing simple technologies to be formulated and marketed (Jenkins et al. 1998; Maniana et al. 2002). Once a conidium attaches to the insect cuticle, it germinates through it and multiplies after reaching the body general cavity. Ultimately, blastospores are proliferated and lead to host death by production of secondary metabolites and consumption of food sources within tissues, mainly fat bodies (Brownbridge et al. 2001; Ortiz-­ Urquiza and Keyhani 2013). The pathogenic fungi which are able to penetrate faster the hemocoel can quickly spread within an insect population and cause earlier death. They utilize an enzymatic complex with main roles of chitinases and proteases to penetrate through integument (St. Leger et  al. 1986; Zibaee and Bandani 2010; Ortiz-Urquiza and Keyhani 2013; Firouzbakht et al. 2015). Typically, at the

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beginning of infection, trypsin- and subtilisin-like proteases synthesize and infiltrate the fungus hypha to the body while after several hours, the synthesis of chitinases increases the penetration and contamination efficiency (St. Leger et al. 1986). Therefore, in addition to bioassay of fungal pathogens on insects, it is possible to determine their efficacy by comparing expression of the involved genes, especially proteases and chitinases (Pedrini et al. 2007; Ortiz-Urquiza and Keyhani 2013). There are several advantages which make EF suitable candidates in biological control programs: (i) they are easily cultured on standard artificial media such as potato-dextrose agar or malat extract agar (however Entomophthora species require a medium containing animal materials), (ii) they are safe for humans and livestock because the best growth temperatures are between 20 and 25 °C, and their growth is inhibited at high temperatures, till 37 °C, (iii) ease of release in a culture medium or on hosts in wet conditions that facilitates the production of non-sexual spores, that are easily dispersed by wind, rain, and water droplets, (iv) resistance to peripheral conditions by producing highly resistant or inactive spores. Nevertheless, there are some disadvantages which make it difficult to recommend EF as the most powerful biocontrol agents in agroecosystems. Several environmental factors are involved in formation of a fungal epizootic in a pest infested area, thus providing favorable environmental conditions is essential for inoculum success in agroecosystems (Lacey et al. 2015). The spores of EF are sensitive to ultraviolet radiation and drought. Another problem is a provision of appropriate types of inoculum to be used in fields, although those can persist in soil and organic matter residues (Goettel et al. 2010). Moreover, several EF may be susceptible to high doses of conventional fungicides. Finally, specificity to target insect, dose of conidia and slow killing are the other factors which limit use of EF as myco-­ insecticides, in some cropping systems. Hence, the use of these agents should be reasonably made along with a complete understanding of where they are aimed to be used (Goettel et al. 2010; Lacey et al. 2015). It is imperative to highlight that advances in molecular biology of EF provide a fundamental platform on improvements of fungal survival and virulence by transforming techniques (Ortiz-Urquiza et al. 2015; Zhao et al. 2016). In fact, these techniques produce recombinant fungi with new characteristics which requires appropriate transformation systems to isolate specific pathogenic genes, investigate virulence determinants leading strains with enhanced virulence or tolerance to environmental stress (Ortiz-Urquiza et al. 2015; Zhao et  al. 2016). But these techniques are costly, require a lot of time to study biometrics and hosting allocation, finally including the legal rules governing genetically modified organisms. Therefore, the use of other methods is necessary to increase efficacy or durability of EF in agroecosystems. In this step, combining the application of EF with reduced concentrations of authorized insecticides can effectively control insect pests and reduce the environmental and biological risks due to chemical compounds.

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13.3  I n vitro Interactions of Entomopathogenic Fungi and Chemical Insecticides Like natural environmental factors, chemical insecticides do directly affect life characteristics of EF. This would be crucial if a compound turned into an obstacle on fungal inoculum establishment in agroecosystems. Such incompatible insecticides suppress vegetative and reproductive performance of fungi and decrease fungal efficiency in biological control or IPM (Zibaee et al. 2009). It is hence mandatory to determine potential compatibility of EF and chemical insecticides prior to field application by assessing germination, vegetative growth and conidia production. These parameters may vary depending on species and isolates, as well as on the nature of the insecticide (Goettel and Inglis 1997; Pachamuthu et al. 1999; Antonio et al. 2001). Germination is an important factor for fungal penetration and subsequent host colonization as any decline in its percentage may negatively influence fungal induced mortality in the host population (Alves 1998). Increasing the yield of conidia and improving their quality are the main objectives to consider EFs as suitable biocontrol agents. This is of highly importance as conidia are the main infective units against insect pests and contribute significantly in fungal survival and dispersal in the environment (Muñiz-Paredes et al. 2017). In vitro studies on interactions between chemical insecticides and EF have been concentrated on possible inhibition of mycelial growth, sporulation and conidial germination although these traits are not necessarily correlated together (da Silva et  al. 2013). For example, Tamai et al. (2002) reported inhibition of mycelial growth in B. bassiana whereas conidia production increased due to stress imposed by the chemical pesticides or initial reduction of mycelial growth. Even, a chemical may have no negative effects on mycelial growth but may inhibit conidiation. Among the mentioned fungal traits in response of chemical treatments, conidial germination may be an indicator of successful epizootics among insect populations (da Silva et al. 2013). Hence, any suppression in conidial germination may lead to lower efficiency and persistence of EF in the target insect habitat. In the following, some examples are given to elucidate in vitro interactions between chemical insecticides and EF in laboratory assessments. Li and Holdom (1994) showed that organochlorine or organophosphorus insecticides were more toxic to entomopathogens than other insecticides. They reported that carbamate insecticides such as carbofuran, methoxyl and oxymyl were moderately toxic while chloropyriphos, malathion and tempephos were extremely toxic. Asi et  al. (2010) evaluated the effects of 13 insecticides including chlorpyrifos, methomyl, thiodicarb, chlorfenapyr, indoxacarb, emamectin benzoate, lufenuron, profenofos, abamectin, triflumuron, flufenoxuron, methoxyfenozide and spinosad on mycelial growth and conidial germination of Metarhizium anisopliae (Metchnikoff) and Paecilomyces fumosoroseus (Wize). Although all used insecticides significantly decreased mycelial growth and conidial germination, with highest toxicity showed by chlorpyrifos. In contrast, spinosad had the least effects on mycelial growth and conidial germination of both fungi (Asi et al. 2010). Marzieh

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et  al. (2010) reported drastic reduction in conidial germination of M. anisopliae when fipronil, pyriproxyfen and hexaflumuron were amended to Sabourad Dextrose Agar (SDA) at the highest concentration (0–15%). They also found higher negative effects on hexaflumuron than other two insecticides (fipronil and pyriproxyfen) indicating its incompatibility with M. anisopliae. Archana and Ramaswamy (2012) added different concentrations of three organophosphorus insecticides (phorate, malathion and chlorpyrifos) and the two synthetic pyrethroids (deltamethrin and permethrin) into the three culture media (Potato Dextrose Agar, Sabourad Dextrose Agar and Czapek Dox Agar) of P. fumosoroseus to check their effects on vegetative growth and colony development. At first, the authors reported that PDA and SDA were the more appropriate media for vegetative growth and spore production of P. fumosoroseus than CDA. Then, they found phorate as the most incompatible insecticide compared to other ones as it caused the least colony growth and spore production. A moderate growth of the fungus was observed in PDA plates treated by melathion, permethrin, deltamethrin and chlorpyriphos, while the growth of P. fumosoroseus in SDA plates treated by the mentioned insecticides showed no significant difference, compared to control. These four insecticides had no effects on spore production in both PDA and SDA media (Archana and Ramaswamy 2012). In vitro toxicity of conventional agrochemicals used in rice crops including eight insecticides (fipronil in two formulations, methyl parathion, cypermethrin, thiamethoxam, thiamethoxam+Lambsa-cyhalothrin, lambda-cyhalothrin, methamidophos), four fungicides (difenconazole, propiconazole, trifloxystrobin, azoxystrobin) and five herbicides (glyphosate, 2,4-dichlorophenoxyacetic acid, bentazon, imazapic + imazapyr, pendimethalin) were investigated on germination, vegetative growth and conidia production in a Brazilian strain of M. anisopliae (CG 168) (da Silva et al. 2013). The field recommended doses of each chemical was added into PDA culture media. Results demonstrated that the fungicides difenoconazole, propiconazole, trifloxystrobin and azoxystrobin had the most toxic effects on the given parameters of M. anisopliae. In contrast, all used insecticides showed the least toxicity and the herbicides only decreased mycelial growth. The authors concluded that methyl parathion, thiamethoxam, lambda-cyhalothrin, glyphosate, bentazon and imazapic + imazapyr were the compatible chemicals with the Brazilian strain of M. anisopliae and could be used in IPM of rice pests. Shaabani et al. (2015) investigated the potential compatibility of an Iranian isolate of M. anisopliae (DEMI 001 strain) with imidacloprid through determining conidial germination, mycelial growth and sporulation. Different concentrations of imidacloprid were added into SDA and yeast extract prior to add 0.5 mL of conidial suspension (105 conidia/mL). The authors reported that imidacloprid had no negative effects on conidial germination, mycelial growth and sporulation, compared with control. In a recently published study, the in vitro interaction between five insecticides of Archer PlusTM (gamma-cyhalothrin 15% [w/v]), LambdaTM (lambda-cyhalothrin 25% [w/v]), CoragenTM (rynaxypyr 20% [w/v]), MatchTM (luphenuron 5% [w/v]) and IntrepidTM (methoxyfenozide 24% [w/v]), and different strains of Beauveria bassiana (Balsamo-Crivelli), M. anisopliae and M. robertsii Bisch et  al. were

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s­ tudied under laboratory conditions (Pelizza et al. 2018). The significant differences were found via conidial-viability tests among the assayed insecticides and the fungal strains. Germination of conidia in all fungal isolates significantly decreased along with increasing insecticide concentrations although γ- and λ-cyhalothrin showed the least effects on germination. Also, the vegetative growth of different fungal strains was influenced by chemical insecticides with the most inhibitory effects on the colony area of B. bassiana strains LPSc 1082, LPSc 1098 and M. anisopliae strain LPSc 907, when exposed to luphenuron. In addition, the highest concentrations of γ-cyhalothrin, λ-cyhalothrin, and rynaxapyr had no effects on the vegetative growth of strain LPSc 1067. Finally, the authors found the highest inhibitory effects on conidia production for methoxyfenozide, whereas γ-cyhalothrin and λ-cyhalothrin showed the lowest effects.

13.4  I n vivo Interactions of Entomopathogenic Fungi and Chemical Insecticides 13.4.1  Selected Case Studies Because of some potential deficiencies of EF to control insect pests, combination with insecticides (mainly low risk ones) might be recommended in agroecosystems. Although such a combination has been shown through synergistic or additive mechanisms, there are some studies indicating antagonistic interactions between EF and synthetic insecticides. In this section, these findings will be exemplified and their mechanisms will be discussed. Earlier studies demonstrated that combinations of dichlorodiphenyltrichloroethane (DDT), azinphos-ethyl, carbaryl, fenvaralate, abamectin, triflumuron, and thuringiensin with B. bassaina caused no synergistic or additive effects on mortality of Colorado potato beetle, Leptinotarsa decemlineata (Say) (Coleoptera: Chrysomelidae) (Fargues 1973, 1975; Anderson et al. 1989). In contrast, Quintela and McCoy (1997) prepared different concentrations of B. bassiana and M. anisopliae, and treated them via dipping assay against first instar larvae of weevil, Diaprepes abbreviates L. (Coleoptera: Curculionidae). Meanwhile, the larvae treated with these EF were orally and topically exposed to 0 and 100 ppm concentrations of active ingredient as well as 500 ppm of formulated imidacloprid. The authors reported no mortality caused by the highest concentrations of the fungi while the larval mortality and mycosis gradually increased once the concentrations were applied with imidacloprid, in both contact and oral exposures. Such a synergism was more significant when the highest concentrations of fungus and insecticides were administrated against the larvae. Also, the authors suggested additive combination of the fungus and imidacloprid when ecdysis of the contact-treated larvae decreased by increasing both insecticide and EF doses. No larval ecdysis was found in oral exposure of imidacloprid. The LT50 value of the fungus significantly

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decreased in both contact and oral administrations with imidacloprid although oral administration led to the lower LT50 value compared to contact exposure. The findings on combination of formulated imidacloprid with M. anisopliae revealed that carriers in formulation had negligible effect on larval mortality, suggesting that the active ingredient was the key component causing synergism between imidacloprid and the EF. The sole and combined effects of M. anisopliae and sublethal concentrations of chlorpyrifos, propetamphos and cyfluthrin, all in commercial formulations, were determined against Blattella germanica L. (Blattodea: Ectobiidae) by Pachamuthu and Kamble (2000). The authors topically treated the adult cockroaches with 108 conidia/ml of M. anisopliae along with low and high concentrations of each insecticide. The combination of M. anisopliae with the three mentioned insecticides led to the significant higher mortalities in all eight concentrations except for 100 ppm. The mortality increased in a dose-dependent manner so that the highest values were found in the combination of M. anisopliae with 300 ppm concentration of chlorpyrifos, propetamphos and cyfluthrin. In details, the combination of M. anisopliae with 200 and 300  ppm of chlorpyrifos, 300  ppm of propetamphos, and 20, 30, and 40 ppm of cyfluthrin led to the faster mortalities than M. anisopliae alone, while other concentrations of the insecticides caused no statistical difference or even antagonistic effects on LT50 value. Finally, the authors concluded that the increased mortality was influenced by the concentration and not the type of each insecticide, showing an additive interaction with the EF. James and Elzen (2001) reported antagonism in the combined application of B. bassiana and imidacloprid against Bemisia argentifolii Bellows (Hemiptera: Aleyrodidae) as the fungus inhibited the insecticide effectiveness. Collectively, they found the mixture of imidacloprid and B. bassiana increased whitefly mortality but less than in a direct additive way, whereas the insecticide showed no negative impacts on spore germination and colony formation. The authors concluded that B. agnetifolii treated by B. bassiana recruited a behavioral response to decline feeding and uptake of imidacloprid so that it might be useful to apply imidacloprid in soil rather than foliage. Their data were in contrast to findings of Steinkraus (1996) and Brown et al. (1997) on the combined effects of these two components against tarnished plant bug. Shapiro-Ilan et al. (2011) set a series of laboratory experiments to find potential combination of B. bassiana with the two common commercial insecticides, cypermethrin and carbaryl, to increase control efficiency of the pecan weevil, Curculio caryae (Horn) (Coleoptera: Curculionidae). After initial experiments to find effective concentrations of both microbial and chemical agents against larvae and adults of weevil, different combinations were considered using low and high concentrations. Mortality was monitored for 9 and 5 days for the larvae and adults, respectively. Results of combined effects on both stages showed synergistic and antagonistic interactions among B. bassiana, cypermethrin and carbaryl, respectively. B. bassiana and carbaryl led to synergistic mortality on the pecan weevil while an antagonistic interaction was found once a combination of B. bassiana and cypermethrin was used against the larvae. The authors found their results differed

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from some other reports and concluded that the type of interaction between microbial agents and chemical insecticides might be dependent on microbial species, chemical structure of insecticides, host species and habitat. In a factorial study, different combinations were prepared by Sharififard et al. (2011) using two concentrations of M. anisopliae and the three concentrations of spinosad to find their effects on the larvae and the adults of Musca domestica L. (Diptera: Muscidae). All treatments including fungal inoculation, insecticide and their combinations led to significant mortalities of adults with the highest values caused by the fungus-insecticide mixture. In details, a fungal concentration of 105 conidia/g with 0.5, 1 and 1.5 mg/g of spinosad led to the highest mortality compared to other fungus concentrations. These findings indicate synergism between spinosad and M. anisopliae at 107 conidia/g with additive effects on adults mortality. Apart from mortality, LT50 values of the fungus/spinosad combinations were significantly lower than each component alone. Likewise, the highest larval mortality was evaluated the combination of M. anisopliae at 108  conidia/g and 0.002, 0.004 and 0.006 μg/g of spinosad, showing synergism. Apart from chemical insecticides, there are a few studies indicating synergism of using EF and horticultural oils against some hemipterans. Initially, Cuthbertson and Collins (2015) demonstrated that mixed application of a petroleum horticultural oil, Tri-Tek, and B. bassiana increased mortality of Bemisia tabaci (Gennadius) compared to sole use of these components under glasshouse conditions. Kumar et al. (2017) tested possible compatibility and efficiency of Isaria fumosorosea Wize with six horticultural oils, vs the Asian Citrus Psyllid, Diaphorina citri Kuwayama (Hemiptera: Liviidae). A blastospore formulate of the fungus was combined with Orchex 796, Sun Pure Spray Oil 435, Conoco Blend Spraybase 435 Oil (90 and 100%), JMS Stylet Oil and PFR-97 and applied against D. citri via leaf disk assay, under laboratory condition. The results clearly demonstrated that horticultural oils significantly increased the fungus efficiency by 100% mortality caused by Orchex 796 and JMS, at 8 and 12 days. Meanwhile, other oils significantly decreased survival time of the psyllids mainly Conoco Blend 435 (100%) although no cadaver colonization was observed in D. citri treated by the oil. The authors concluded that these oils were compatible with I. fumosorosea, increased its efficiency against adult psyllids and fungal development index (FDI), the latter being crucial for fungal persistence in the site of spraying, for horizontal transmission of spores to other individuals (Kumar et al. 2017). Pelizza et al. (2018) evaluated the combined effects of five insecticides and three EF (five strains) against soybean defoliating pest, Rachiplusia nu Guenee (Lepidoptera: Noctuidae). Three concentrations of γ-cyhalothrin, λ-cyhalothrin, rynaxypyr, luphenuron, and methoxyfenozide were prepared based on 100, 50 and 25% of the average field recommended doses and combined 1:1 with the three concentrations (104, 106 and 108 conidia/ml) of B. bassiana (isolates LPSc 1067, LPSc 1082, LPSc 1098), M. anisopliae (LPSc 907), and M. robertsii (LPSc 963) to test their potential synergistic, antagonistic or additive effects on larvae. Results revealed significantly different mortalities caused by insecticides/fungi combinations on the third larval instars after 10-day exposure using half concentration of the field

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r­ecommended dose. Larval exposure to LPSc 1067, LPSc1082, LPSc 1098, LPSc 907 and LPSc 963, combined with all insecticides at the highest field concentration, led to 100% mortality showing additive interactions. Similar results were observed except for the combination of λ-cyhalothrin/LPSc-1098 and M. robertsii strain LPSc 963. At the same concentration, the synergistic interaction was found in case of γ-cyhalothrin/LPSc-1067 or LPSc-963, λ-cyhalothrin/LPSc-1067, rynaxypyr/ LPSc-1067 or LPSc-1098, and luphenuron/LPSc-1067. Moreover, antagonistic interaction was obtained when the combinations of λ-cyhalothrin/LPSc-1098 or LPSc-963, rynaxypyr/LPSc-963, luphenuron/LPSc-963, and methoxyfenozide/ LPSc-907 or PSc-963 were used against the larvae of soybean defoliator. The combination prepared based on 25% of the highest field concentration showed an additive interaction except for γ-cyhalothrin/LPSc-1067, λ-cyhalothrin/LPSc-1067, rynaxypyr/LPSc-1067, with methoxyfenozide/LPSc-1082 indicating synergism and rynaxypyr/LPSc-1098 showingantagonism.

13.4.2  M  echanisms Underlying EF and Insecticides Interaction Although fungal species, sensitivity to environmental factor, conidia formulation, target insect species, chemical structure of insecticide and mode of action are among the factors influencing outcomes of fungi/insecticides combinations, there are some physiological traits within insects which determine whether these combinations may be synergistic or antagonistic. Serebrov et al. (2001) infected the fifth larval instars of greater wax moth, Galleria mellonella L. (Lepidoptera: Pyralidae) with B. bassiana, M. anisopliae and P. fumosoroseus.Hemolymph of the larvae was then collected, determining esterase activity with specific substrates.. In next step, the authors treated the larvae with a concentration of deltamethrin via topical application to determine their sensitivity. The authors found that all fungi induced proteinaceous patterns related to esterase activity in gel electrophoresis, related to the virulence of each entomopathogenic fungus and treatements with deltamethrin. Such changes were observed during mycosis, mechanical damage of cuticle, chilling at –4 °C for 30 min and larval treatment by deltamethrin. Moreover, the LC50 concentration for deltamethrin on the intact larvae was 1.8 mg/g and 2.1 mg/g in the larvae infected by highly virulent strain. Serebrov et  al. (2006) tried to determine whether fungal infection could elicit insect resistance against malathion, an organophosphorus insecticide. Initially, the fifth larval instars of G. mellonella were infected by M. anisopliae, then hemolymphs of control and infected larvae were collected to assay activities of esterase, glutathione S-transferase and phosphatases. In a further assay, the control and already infected larvae were exposed to malathion. Finally, enzyme inhibitors were used to confirm the role of detoxification enzymes in the insect resistance to the entomopathogenic fungus. The activity of esterases significantly increased almost

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two folds in the hemolymph of the infected larvae by M. anisopliae, while no significant changes were observed in fat body and intestine preparations. Similar results were found in the activity of glutathione S-transferase while acid- and alkaline phosphatases showed no significant changes (Serebrov et al. 2006). Results on malathion treatment against control and M. anisopliae infected larvae revealed that LC50 required to kill 50% of the infected larvae was 1.5 times higher than in control, showing a certain degree of resistance to malathion in these larvae. Results on using inhibitors of detoxifying enzymes including S,S,S-tributyltrithiophosphate (tribufos, a non-specific esterases inhibitor), piperonyl butoxide (cytochrome P450-­ dependent monooxygenases inhibitor), and diethyl maleate (glutathione S-transferases inhibitor) showed that the M. anisopliae-infected larvae exposed to tribufos and piperonyl butoxide had higher mortality compared to control. No significant difference was observed when diethyl maleate was used against the larvae. Consequently, the authors attributed the higher activities of detoxifying enzymes during mycosis to intoxication with metabolites of M. anisopliae or with products present in the degraded host tissues (Serebrov et al. 2006). In a similar study, Zibaee et al. (2009) investigated the effects of B. bassiana and its secondary metabolite on toxicity of fenitrothione against sunn pest, Eurygaster integriceps Puton (Hemiptera: Scutelleridae). Both fungal spores and secondary metabolites significantly increased glutathione S-transferase activity using CDNB and DCNB as substrates, 3–5 days after inoculation. Similar results were found in the activity of esterase in the infected hemipteran compared to control, using α- and β-naphtyl acetate as substrates. In the last experiment, results on fenitrothione treatment on control and treated hemipteran with B. bassiana spores and secondary metabolites showed LC50 values of 9.4 ml/mg and 22.5 ml/mg compared to 5.9 mL/mg of control, almost two to fourfold higher. These results clearly showed resistance of E. intergriceps to fenitrothione after fungal treatment. Apart from above-mentioned findings indicated the possible mechanisms underlying antagonistic interactions between EF and insecticides, there are studies on explaining why synergism occurs between these agents. In the sets of experiments, Jia et al. (2016) tried to find whether chlorantraniliprole and a M. anisopliae strain had synergism in control of the migratory locust, Locusta migratoria L. (Orthoptera: Acrididae). Then they determined what might be the mechanism underlying such an interaction. In the first two experiments, the efficiency of M. anisopliae and chlorantraniliprole was evaluated separately. In the third assay, different proportions of M. anisopliae and chlorantraniliprole were prepared to test their combination efficiency against L. migratoria. In the last experiment, the insects were initially treated with M. anisopliae conidia then after 4 days, and fed on wheat sprouts treated with different concentrations of chlorantraniliprole, under laboratory conditions. The insects exposed to combined proportion of the entomopathogenic fungus and insecticides were considered for biochemical analysis of esterase, glutathione S-transferase, monoxygenase, acetylcholine esterase, phenoloxidase, superoxide

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dismutase, peroxidase, catalase, chitinase and aryl acylamidase. A higher mortality was found when the mixed application of M. anisopliae and chlorantraniliprole was used against L. migratoria with LC50 values of 0.01 and 0.02 mg/L in comparison with 0.15 mg/L of M. anisopliae alone. Also, the locusts initially treated with the fungus showed co-toxicity after treatment with chlorantraniliprole. Separately treated L. migratoria with M. anisopliae and chlorantraniliprole showed the higher activity of esterase compared to control, but the enzyme activity significantly decreased following treatment with the mixed solution. A similar trend was reported for glutathione S-transferase only in the earlier periods after treatments. L. migratoria treated with the mixture of M. anisopliae and chlorantraniliprole showed the lower activity of phenoloxidase at earlier periods that increased later. Separate treatment with these two agents increased the enzyme activity compared to control. The sole and mixed exposure on L. migratoria led to the higher activity of monoxygenase somehow similar to the activities of acetylcholinesterase, chitinase and aryl acylamidase. The activities of superoxide dismutase, catalase and peroxidase increased in the locusts treated with M. anisopliae and chlorantraniliprole alone, but the mixed combination increase the activities of superoxide dismutase and peroxidase, with a lower activity of catalase. The authors concluded that L. migratoria recruits enzymatic defenses against M. anisopliae or chlorantraniliprole, while their mixed application may disable these enzymes regarding time after exposure. The authors believed that these agents were able to disrupts calcium ion within insect cells which was critical to normal activity of enzymes, mainly phenoloxidase. Moreover, mixed action of M. anisopliae and chlorantraniliprole disabled a complex of biochemical processes leading to synergistic interactions in L. migratoria (Jia et al. 2016). In addition, Ali et al. (2017) reported data on toxicological and biochemical characteristics of synergism between Lecanicillium muscarium Zare & Gams and matrine, following treatment on B. tabaci. Experimental design and assayed biochemical enzymes were similar to Jia et al. (2016). Results showed higher mortality with the lower LC50 values when different proportions of L. muscarium and matrine were prepared and applied on B. tabaci, showing synergistic interactions between the two agents. The esterase and glutathione S-transferase activities initially significantly increased after L. muscarium and matrine treatment. However the enzyme activity sharply decreased in B. tabaci treated with mixture of both agents. In contrast, the activity of acetylcholinestrase decreased in B. tabaci treated with the sole or mixture of both agents, regardless of post-treatment intervals. Finally, the activities of chitinase and the three enzymes involved in antioxidant responses (superoxide dismutase, catalase and peroxidase) increased in the insects treated with sole or mixture of L. muscarium and matrine, 3 and 4 days after treatments, respectively. The authors attributed the synergistic interactions of L. muscarium and matrine to alterations of acetylcholineesterase activity, leading to disturbance of the acetylcholine balance in the treated B. tabaci.

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13.5  P  otential of Entomopathogenic Nematodes Against Insects Entomopathogenic nematodes (EPN) have a unique symbiotic relationship with bacteria (Shapiro-Ilan et al. 2017). They enter host body via natural openings like mouth and anus then continue pathogenesis inside the body as endoparasites (Lacey et al. 2015, see also Chap. 9). These agents are found in different ecological habitats infecting several agricultural insect pests although their relationship with insects varies from phoresis to parasitic/virulent interactions (Shapiro-Ilan et al. 2017). The entomopathogenic nematodes have been described in 23 families of which seven families contain species with potential for biological control of insects (see also Chap. 9). It is notable that species from the two most important families Steinernematidae and Heterohabditidae have been commercialized for use in agroecosystems (Shapiro-Ilan et al. 2017). It has been reported that EPN distribution is limited by the availability of sensitive hosts although virulent have the ability to quickly kill their hosts (1–4 days depending on nematode and host species), coexisting with bacteria (Xenorhabdus bacteria for steinernematidae and Photorhabdus bacteria for heterohabditidae) (Dowds and Peters 2002; Shapiro-Ilan et al. 2017). The common feature of all EPNs is to have a third stage infective juvenile (Dauer Juvenile) which exists outside of host body, carries the cells of symbiont bacteria, finds the host and enters into hemocoel. In host body, the abandoned bacteria proliferate and kill the host through several mechanisms. Nematodes within infected body will produce 1–3 generations, and then in the next generation, the contaminated juveniles will exit to infect new hosts (Kaya and Gaugler 1993). Infectious nematodes can be identified with different symptoms. In case of steinernematids, the killed host larvae are brown, red, and black, while the larvae killed by heterohabditids are red, brick, purple, orange, yellow, and green with differences depending on the bacterial species (Shapiro-Ilan et al. 2017). The range of insect hosts for the entomopathogenic nematodes is extensive in laboratory, but it is much more limited in natural environments, depending on host and environmental conditions. Nevertheless their impact on non-target organisms is negligible (Lacey et al. 2015). One of the factors limiting their host range is the type of behavior required to get into the host body. Some species have a sit-and-wait strategy (ambush) close to the soil surface, where they infect hosts with a special nictation and jumping behavior. Other species are explorers, active in a wide range compatible with relatively low-mobility hosts. Some species may also choose between these two strategies (Shapiro-Ilan et al. 2017).

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13.6  I nteractions of Entomopathogenic Nematodes and Chemical Insecticides The EPN that are used against agricultural pests will definitely be exposed to other materials applied in agro-ecosystems affecting their survival, distribution and efficiency (Shapiro-Ilan et al. 2017). Irrigation methods, type and amount of fertilization, type of pesticides used and their formulation are among the factors that affect interactions of insects and their pathogenic nematodes (Lacey et  al. 2015). Depending on time and type of chemical used, and physical and chemical characteristics of soil, EPN can show antagonistic to synergistic interactions with insecticides which influence effectiveness of the control method and its cost. Nishimatsu and Jackson (1998) evaluated the combined effects of terbufos, fonofos and tefluthrin with the entomopathogenic nematode Steinernema carpocapsae Weiser against the larvae of western corn rootworm, Diabrotica virgifera virgifera LeConte (Coleoptera: Chrysomelidae). The combination of nematode and terbufos or fonofos resulted in additive effects on mortality of D. virgifera while tefluthrin and S. carpocapsae showed synergism by increasing 24% of total mortality. Similar findings were reported in combination of tefluthrin and Heterorhabditis bacteriophora Poinar. The authors proposed that the paralytic and convulsive responses of D. virgifera due to the insecticide treatment might enhance the insect susceptibility to nematode infection or establishment within the host. In fact, the increased metabolic activity and reduced directional movement led to higher production of CO2 which facilitates host finding by the nematode (Nishimatsu and Jackson 1998). Koppenhöfer et al. (2000) found the synergistic interaction between imidacloprid and the two entomopathogenic nematodes, Steinernema glaseri (Steiner) and H. bacteriophora as well as additive interaction with S. kushidai Mamiya against the larvae of white grubs.Imidacloprid negatively affected mortality, speed of kill and establishment of S. kushidai while these parameters were positively affected in case of S. glaseri and H. bacteriophora. The authors reported the general disruption of normal nerve function leading to lower biological activity of larvae, including their grooming and evasive behavior, as the main phenomenon responsible for the synergistic interactions observed. Radova (2010) determined the interaction of Steinernema feltiae Filipjev with eight insecticides (kinoprene, lufenuron, methomyl, metoxyfenozide, oxamyl, piperonyl-butoxide, pyriproxyfen, tebufenozide), seven acaricides (azocyclotin, clofentezin, diafenthiuron, etoxazole, fenbutatinoxide, fenpyroximate, tebufenpyrad) and four fungicides (captan, fenhexamid, kresoxim-methyl, nuarimol) evaluating survival and infectivity of the infective juveniles. Treated insecticides caused a low mortality (2–18%) on infective juveniles of S. feltiae after 3 days of application, in which tebufenozide and piperonylbutoxide showed the lowest and highest mortalities, respectively. Similar findings were reported in case of acaricides except

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for fenpyroximate that caused a >20% mortality after 3 days. Mortalities around 7–8% were observed in S. feltiae after treatment with fungicides, with a lower value than control for nuarimol. No negative effects were observed in virulence of S. ­feltiae after insecticidal treatments as the mortality of 80–100% were imposed on red flour beetle, Tribolium castaneum Herbst (Coleoptera: Tenebrionidae) after 5 days, similar to fungicides, showing no significant effects on EPN virulence. In contrast, the two acaricides, fenpyroximate and tebufenpyrad, significantly decreased S. feltiae virulence to 95% and 85%, respectively (Radova 2010). Negrisoli et al. (2010a) determined the compatibility of the three entomopathogenic species including Heterorhabditis indica Poinar, Steinernema carpocapsae Nguyen & Smart and S. glaseri Nguyen & Smart with 18 conventional insecticides used against fall armyworm, Spodoptera frugiperda (Smith) (Lepidoptera: Noctuidae) [namely betacyfluthrin, cypermethrin, chlorpyrifos (two commercial products), deltamethrin, deltamethrin + Triazofos, diflubenzuron, gamacyhalothrin, lambdacyhalothrin, lufenuron, methoxyfenozide, methylparathrion, permethrin (two commercial products), spinosad, triazofos, triflumuron and alphacypermethrin]. Initially, the infective juveniles of each nematode species were exposed to insecticides, then their viability and infectivity against S. frugiperda were evaluated in laboratory conditions. One of the commercial formulations of chlorpyrifos (LorsbanTM) and lufenuron showed the lowest mortality on the infective juveniles of all EPN. A 88% after exposure to chlorpyrifos (LorsbanTM) and lufenuron, similar to control, while the lowest one was scored for H. indica after exposure to methylparathion. The infectivity of S. carpocapsae was high after exposure to gamacyhalothrin, chlorpyrifos (LorsbanTM) and lufenuron while it was significantly lower than control in response to triazofos. Finally, the insecticides cypermethrin, lambdacyhalothrin, triflumuron, methoxyfenozide, triazofos and alphacypermethrin caused the highest S. glaseri infectivity (Negrisoli et al. 2010a). These authors attributed the insensitivity of nematodes to some insecticides to the presence of butyrylcholinesterase in their synapses, that protects acetylcholinesterase toward inhibitors such as organophosphorus insecticides. Also, they found that nematodes had no chitin in their cuticle so they are insensitive to diflubenzuron. These findings and conclusions indicated compatability of H. indica, S. carpocap-

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sae and S. glaseri with some registered chemicals to be used against S. frugiperda at least in laboratory conditions (Negrisoli et al. 2010a). In a subsequent study, Negrisoli et al. (2010b) tested the efficacy of the above-­ mentioned insecticides and the EPN to control S. frugiperda in laboratory and field conditions. All used insecticides showed equal mortalities against S. frugiperda larvae with values significantly higher than control. After 2 days insecticides including cypermethrin, spinosad, methoxyfenozide and deltamathrin + triazofos led to additive interaction with H. indica while chlorpyrifos (LorsbanTM) showed a synergic interaction. Similar results were observed in combination of S. carpocapsae with lufenuron, chlorpyrifos (LorsbanTM), cypermrthrin, methoxyfenozide, lambdacyhalothrin and chlorpyrifos (VexterTM). S. glaseri demonstrated synergism with chlorpyrifos (VexterTM), chlorpyrifos (LorsbanTM) and methoxyfenozide while the nematode had antagonism with deltamethrin + triazofos, diflubenzuron, matylparathion and alphacypermethrin. Other insecticides caused additive interactions with S. glaseri (Negrisoli et al. 2010b). After 4 days, chlorpyrifos (VexterTM) showed the additive interaction with S. glaseri whereas the other insecticides showed nematode antagonism. Chlorpyrifos (LorsbanTM) and gamacyhalothrin showed synergism with H. indica while the nematode had antagonism with deltamethrin and methylparathion. A synergism was found in the interaction of S. carpocapsae with alphacypermethrin and gamacyhalothrin. The authors suggested that time of exposure would definitely affect the EPN interaction type with insecticides. Such synergistic or additive effects have been already reported in combination of nematodes with malathion (Baweja and Sehgal 1997), organophosphorus, carbamates, pyrethroids, cartap and imidacloprid (Zhang et al. 1994; Koppenhöfer and Kaya 1998; Koppenhöfer et al. 2002). In field experiments, the combined application of the insecticides and the EPN caused higher mortalities against S. frugiperda compared to the nematodes alone, with additive interactions. It is essential to indicate that chlorpyrifos (LorsbanTM) and methoxyfenozide showed no statistical difference from EPN alone (Negrisoli et al. 2010b). Bortoluzzi et al. (2013) evaluated the efficiency of sixteen steinernematids and heterohabditids EPN against banana weevil borer, Cosmopolites sordidus Germar (Coleoptera: Curculionidae) and their interactions with a conventional insecticide, carbofuran. In combination experiments, 400 ml/L of the insecticide were mixed with a solution containing 2000 infective juvenile of each species to check for potential interactions. All nematode species caused significant mortality on C. sordidus. No negative effects of carbofuran was observed on viability of infective juveniles but a significant decrease by 72% was recorded in the nematodes exposed to the insecticide. The authors concluded that carbuforan as a carbamate insecticide inhibited acetylcholinesterase and disrupting the foraging behavior and entering capability of the treated nematodes toward host body. In a regional study, the interactions among three conventional insecticides, dinotefuran, indoxacarb and imidacloprid, were investigated to the two Arizona-­ native entmopathogenic nematodes, Heterorhabditis sonorensis Stock et  al. and Steinernema riobrave Cabanilas et al., by measuring survival, virulence and reproduction rates (Navarro et  al. 2014). The authors set up three bioassay including

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initial EPN application prior to insecticidal exposure after 24 h, initial exposure to insecticides prior to nematode application after 24 h and simultaneous application of all agents. The survival of both nematodes was not affected by the insecticides. Even one of them, indoxacarb, showed a synergistic effect on the S. riobrave ­virulence, although the reproduction of both nematodes decreased by two-fold, compared to control. Antagonistic interactions on virulence were observed in combinations of imidacloprid with S. riobrave compared to an additive effect of the insecticide on H. sonorensis virulence. Finally, dinotefuran promoted additive effects on virulence of both nematodes against corn earworm, Helicoverpa zea Boddie (Lepidoptera: Noctuidae). The authors concluded that synergism between indoxacarb and the EPN might be due to non-systemic effects of the insecticide on the insects nervous system, by blocking Na+ channels, leading to larval paralysis that makes them more susceptible to nematodes. Additionally, antagonism between imidacloprid and one of the used nematodes was due to its unsuitability against lepidopteran insects (Navarro et al. 2014). Laznik and Trdan (2014) determined the possible compatibility of eight insecticides (abamectin, lufenuron, toxin of Bacillus thuringiensis var. kurstaki, pymetrozine, azadirachtin, imidacloprid, λ-cyhalotrin and pirimicarb) with commercial products of S. feltiae, S. carpocapsae and S. kraussei Steiner. The two species of S. carpocapsae and S. kraussei appeared incompatible with all assayed insecticides while only abamectin and lufenuron negatively affected viability of H. bacteriophora. In contrast, S. feltiae was compatible with azadirachtin, toxin of Bacillus thuringiensis var. kurstaki and imidacloprid. The authors believed that compatibility might be even strain-specific rather than species-specific. The combination of S. feltiae and H. bacteriophora were compatible with azadirachtin and pirimicarb leading to a cost-effective alternative to control vegetable pests. Sabino et al. (2014) studied the interaction of S. carpocapsae and H. amazonensis with insecticides (abamectin, triflumuron, deltamethrin, dimethylamino-propyl, chlorpyriphos, lambda-cyhalotrin +chlorantranilprole, chlorantranilprole and thiamethoxan + λ-cyhalothrin) in field condition. No significant effect was found on EPN viability except for dimethylamino-propyl which caused a mortality higher than control (24.6%). Abamectin and chlorpyrifos caused 26–30% and 20–50% mortalities against the larvae of G. mellonella in combination with H. amazonensis and S. carpocapsae, respectively. No significant differences for EPN viability and infectivity were observed after combination with thiamethoxan + λ-cyhalothrin and anthranilamide, while all pyrethroids imposed significant effects compared to control. This was likely due to the presence of surfactants in the formulations of these insecticides. Similarly, Guo et al. (2017) designed the laboratory and field experiments to find interactions of chlorantraniliprole, diflubenzuron and imidacloprid with S. longicaudum Shen & Wang and H. bacteriophora, against the white grub, Holotrichia oblita Faldermann (Coleoptera: Scarabaeidae). Different insecticide concentrations showed no effect on survival or virulence of S. longicaudum and H. bacteriophora. In laboratory experiment evaluating the effect of nematode–insecticide combinations against larvae of H. oblita, mortality rates of the larvae in all nematodes treatments were significantly higher than that found in the insecticide

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treatments, at 4, 7, and 14  days. Combinations of nematodes and insecticides showed higher mortalities of target insect compared to the nematodes alone, showing synergism after 4 days. The combinations of S. longicaudum–imidacloprid, H. bacteriophora  – imidacloprid, or H. bacteriophora  – chlorantraniliprole ­demonstrated synergistic effects in peanut fields, leading to reductions of grub larvae and percent of injured legumes. Finally, cost–benefit analysis revealed that a combination of H. bacteriophora – imidacloperid might be practical to field management of H. oblita in the peanut production.

13.7  Conclusion IPM nowadays is one of the most important systems to control pest damages and to produce high quality and healthy crops. Different methods are used to control insects, in which biological and chemical control are most prominent techniques. Although the sole use of these methods has given satisfactory results, IPM should include all reasonable control methods to achieve best results at a minimum cost. Since the use of biological control methods, with a focus on entomopathogens and chemical insecticides, has its own disadvantages and benefits, it is advisable to create a framework for their combined use. Such a framework requires laboratory and field studies to examine positive or negative interactions when these two methods are applied together. As shown in this chapter, some chemical compounds have synergistic or additive effects on entomopathogens, while others cause some antagonistic responses. Therefore, before combining these two methods, screening between conventional chemical insecticides to control a specific pest and EPN should be done in both laboratory and field. Such an experiment involves the selection of insecticides from different classes, taking into account their diverse formulations, spraying time and spray location. If these experiments are carried out carefully, it seems that the proper combination of a low-risk insecticide with an entomopathogen can successfully control the target pests, with lowest costs and optimal outcomes.

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Chapter 14

Current State of Fungal Antagonists with Special Emphasis on Indian Scenario Purnima Das, Lakshmi Kanta Hazarika, Surajit Kalita, and Somnath Roy

Abstract  The major objective of shifting from conventional to eco-friendly and sustainable agriculture is to minimize the load of chemical inputs. The replacement of chemical pesticides by biologicals is a prime step for achieving this objective. Entomopathogenic fungi (EPF) can act as effective management tools for insect and mite pest regulations without harming the biodiversity and environment. About 49 EPF species, predominantly from Deuteromycotina and Zygomycotina, have so far been commercially exploited world-wide as biopesticides, among which the most widely used species belong to genus Beauveria and Metarhizium. In India, a country of rich biodiversity, research on EPF has given rise to many products, out of which about 638 EPF formulations have so far been registered. Despite of huge resources and many advantages over conventional synthetic pesticides, EPF remain relatively underutilized in Indian agriculture. Biopesticides represent only around 2–3% of the overall pesticide market in India, a share that is expected to increase manifold in coming years. In this review, we provide an overview on promising EPF as bio-control agents of the key agricultural pests and also the scope for future studies for their better utilization. Keywords  Entomopathogenic fungi · Pest control · Biodiversity · Fungal antagonists

P. Das · S. Kalita Department of Entomology, Assam Agricultural University, Jorhat, Assam, India L. K. Hazarika Assam Women’s University, Jorhat, Assam, India S. Roy (*) Division of Entomology, Tea Research Association, Jorhat, Assam, India e-mail: [email protected] © Springer Nature Switzerland AG 2019 M. A. Khan, W. Ahmad (eds.), Microbes for Sustainable Insect Pest Management, Sustainability in Plant and Crop Protection, https://doi.org/10.1007/978-3-030-23045-6_14

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14.1  Introduction In nature, a pool of microorganisms including 1000 species of entomopathogenic fungi (EPF) under phyla Entomopthoromycota, Blastocladiomycota, Mycosporidia, Basidiomycota and Ascomycota (Vega et al. 2012; Wang and Wang 2017), 90 species of bacteria, 1600 viruses (Moazami 2009) and 111 species of nematode (Hunt and Nguyen 2016) regulate insect populations. However, about 49 EPF species predominantly from Deuteromycotina and Zygomycotina have so far been commercially exploited as biopesticides representing numerically 44% of the antagonistic microorganisms pool in number. The most widely used species is Bacillus thuringiensis Berliner. Microorganisms exploited for biological control of insect pests through introduction and augmentation contribute 1.4–2.5% of the total $28 billion global pesticide market, amounting for $392–700 millions. Bacteria alone occupy 74% (Thakore 2006). Figure 14.1 shows the status of the remaining groups. Moreover, EPF as alternatives to insecticides, or their combined applications with insecticides could be very useful for managing resistant pests (Hoy 1999) as well as for formulating sustainable integrated pest management strategies of various crops and public health pests. Normally biopesticides are eco-safe alternatives and inherently less harmful in compared to chemical pesticides. They are target specific, effective in small quantities, easily degradable, do not leave problematic residues, and their production cost is lower compared to synthetic pesticides. It is less likely that target pests develop resistance. Another advantage is that unlike bacteria and viruses, EPFs behave like contact insecticides. However, main disadvantages are unavailability of quality products in sufficient quantity in the local market, poor shelf life of majority of formulations, slow in action, inconsistent or variable efficacy under field conditions due to the influences of biotic and abiotic factors. In recent times, importance has been given to some technical aspects of biopesticide development, such as mass production, formulation, and selection of more virulent strains with consistent efficacy under field 80

74%

70

Share (%)

60 50 40 30 20

10%

10 0

Bacteria

Fungus

5%

Virus

8%

Predators

3%

Others

Biocontrol agents Fig. 14.1  World’s biopesticide market as shared in percentage by different groups

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c­ onditions. These are of course essential, but identification and modification of fungal virulence determinants or genes must be undertaken to produce efficient mycoinsecticides.

14.2  Historical Background Earliest studies of EPF dated back to eighteenth century. While developing ways to control diseases that devastated the silkworm industry in France, Agostino Bassi (1773–1856) demonstrated that Beauveria bassiana (Bals.-Criv.) Vuill.  was the infectious agent causing the muscardine disease of silkworms. But the use of B. bassiana for insect pest control was first studied by Audoin (1837). Subsequently, many discoveries were made to isolate several EPFs, out of which the most important fungi are Metarhizium anisopliae Sorokin, Nomuraea rileyi  (Farl.) Samson, Lecanicillium lecanii R. Zare & W. Gams, Hirsutella thompsonii Fisher and Isaria fumosorosea (Wize) A.H.S. Br. & G. Sm. (Table 14.1). B. bassiana isolated from a variety of insects worldwide is a filamentous imperfect fungus with high host specificity. Host range includes Leptinotarsa decemlineata Say, Cydia pomonella Linnaeus, several genera of termites, and Helicoverpa armigera Hubner (Thakur and Sandhu 2010), Dicladispa  armigera Olivier (Hazarika and Puzari 1997; Hazarika et  al. 2005), Corcyra cephalonica Stainton (Das 2015), Periplaneta americana L. (Mudoi et al. 2017) and several other vectors of tropical infectious diseases such as Phlebotomus, Glossina morsitans and bugs of genera Triatoma and Rhodnius. Likewise, M. anisopliae is one of the commercially exploited EPFs against (Sandhu et  al. 2012) locusts, grasshoppers, cockroaches in both developed and developing countries of Africa, America and Australia. It is formulated in oils, under high humid conditions and applied, which resulted 90% kill of locusts within 7–21 days. It can be mass produced on rice grains. L. lecanii is another widely distributed EPF, which causes widespread epizootics in tropical and subtropical regions (Nunez et al. 2008). Being a dimorphic hypomycete, N. rileyi can cause epizootics in lepidopterans (noctuids) and coleopterans (Ignoffo 1981; Ignoffo et  al. 1989; Vargas et al. 2003). Its host specificity and eco-friendly nature encourage its use in insect pest management. Limitations of EPFs are that they are unpredictable and geographical locations and hosts play important roles in determining specificity and virulence (Boucias et al. 1982; Vimaladevi et al. 2003). In a review, Feng et al. (1994) summarized the progress and achievements made in mass production of B. bassiana using different media and techniques by highlighting advantages and disadvantages of solid and liquid medium or submerged culture. Methods were indicated to overcome problems associated with mechanical production system. Solid- and liquid-state fermentation have been developed for mass production of B. bassiana, based on diphasic and submerged techniques (Rombach et al. 1988; Feng et al. 1994). Whatever the technique, it must result in high productivity consuming less energy and produce low wastewater (Lonsane

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Table 14.1  Entomopathogenic fungi isolated from insect pests Fungal spp. Metarhizium anisopliae

Beauveria bassiana

Nomurea rileyii

Verticillium lecanii

Stage of infection Larvae Grub

Insect spp. Cabbage butterfly (Pieris brasicae) White grub (Anomela spp, Apogonia spp) Cutworm (Agrotis ipsilon) Brinjal aphid (Myzus persicae)

Order Lepidoptera Coleoptera

Cowpea aphid (Aphis craccivora)

Hemiptera

Termite (Odontotermes obesus) Rhinoceros beetle (Oryctes rhinoceros) Sugarcane shot borer (Chilo infuscatellus) Rice Hispa (Dicladispa armigera) White fly (Bemisia tabaci) Tea mosquito bug (Helopeltis theivora) Dung beetle (Catharsius molossum) Black ant (Diacamma rugosum) Rice case worm (Nymphula depunctalis) Rice weevil (Sitophilus oryzae) Tiger moth (Creatonotos gamgis) Yellow tussock moth (Somenia scintillans) Black fly (Aleurocanthus wouglumi) Cabbage looper (Spodoptera litura) Velvetbean moth (Anticarsia gemmatalis) Cotton bollworm (Helicoverpa armigera) Cabbage Looper (Trichoplusia ni) Clover leaf weevil (Hypera punctata) Black cutworm (Agrotis ipsilon) Armyworm moth (Mythimna unipuncta) Red spider mite (Oligonychus coffeae)

Isoptera Coleoptera

Grub Nymph and adult Nymph and adult All stages Grub

Lepidoptera

Larvae

Coleoptera Hemiptera Hemiptera

All stages Adult Adult

Lepidoptera Hemiptera

Coleoptera Adult Hymenoptera Adult Lepidoptera Larvae Coleoptera Lepidoptera Lepidoptera

Adult Larvae Larvae

Hemiptera Lepidoptera Lepidoptera

Nymph Larvae Larvae

Lepidoptera

Larvae

Lepidoptera Coleoptera Lepidoptera Lepidoptera

Larvae Adult Larvae Larvae

Acarina

Adult

Source: Hazarika et al. (2005), Roy and Muraleedharan (2014), Dutta et al. (2012), and Das (2015)

et al. 1992; Pandey et al. 2001). Many agricultural wastes were utilized for production of EPF which include sugarcane bagasse, rice bran and rice husk (Mazumder et al. 1995; Puzari et al. 1997). Besides wheat bran, soybean chunks, rice, maize and oats were also utilized by many workers to mass produce B. bassiana and

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M. anisopliae. Amongst the liquid medium, rice grewel, coconut water and potato broths were the most common. Das et al. (2017) improved some of the media by adding different carbon and nitrogen sources. It was also observed that addition of crustacean chitin further improves the productivity of the liquid medium.

14.3  Mode of Action of Fungal Antagonists The insect cuticle is the foremost barrier for any biological insecticides. It is mainly composed of epicuticle, procuticle and epidermis. The outermost surface layer of the epicuticle is the cement and lipid layer which are mostly resistant to enzymatic degradation, and act as barrier to biotic and abiotic agents (Hadley 1981). Unless physically disrupted, it can prevent entry of EPF. Recently, Wang and Wang (2017) reviewed comprehensively the molecular mechanism of entomopathogenesis based on researches conducted on species of Beauveria and Metarhizium. Different genes mediating infection by an EPF have been functionally characterized and grouped under adhesion, cuticle degradation, nutrient assimilation and stress management (Butt et al. 2016). In general, spore adhesion, differentiation of infection structures, detoxification, hemocoel adaptation, starvation and evasion of innate immunity are different phases of an infection of a susceptible host by an EPF (Figs.  14.2 and 14.3a-d). Hydrophbin genes, hyd1 and hyd2, mediate adhesion of a B. bassiana spore to lipid layer of the insect epicuticle (Boucias et al. 1988; Lord 2001; Goettel et al. 2005; Zhang et al. 2011). Following this stage, the spore germinates in a high humid and moderately hot environment as well as in presence of specific

Fig. 14.2  Schematic diagram of mode of action of entomopathogenic fungi

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Fig. 14.3  Mode of action of B. bassiana on D. armigera. Conidia in contact with integument (a). Germination of conidia (b). Appresoria formation and penetration of germ tube through the cuticular depression on prothorax (c). Penetration of germ tube through membranous cuticular depression. Scale bars: 10 μm (a, c, d); 1 μm (d). (d)

epicuticular hydrocarbons (Feng et al. 1994; Napolitano and Juarez 1997) to form a germ tube (Wang and Wang 2017). Subsequently, an infection structure, the appressorium, is formed (Hajek and St. Leger 1994; Wang and Wang 2017), which coincides with secretion of aminopeptidases. Likewise, MAD 1 acts as hydrophobins of Metarhizium. Long chain C18 cuticular fatty acids enhance the process (Bidochka and Khachatourians 1991), while short chain fatty acids and aldehydes inhibit growth of EPF (Smith and Grula 1982; Sosa-Gomez et  al. 1997). However, it is hypothesized that besides adhesion, these genes perform other functions, such as cell surface hydrophobicity, virulence, and formation of rodlet layer (spore coat) (Zhang et al. 2011). In addition to release of proteases and chitinases, the mother spore translocates lipid droplets (LDs) to the appressorium. As a result of hydrolysis of LDs, accumulation of high concentration of glycerol takes place, which generates a high turgor pressure to breach the epicuticle (Wang and Wang 2017) in presence of cyclic anti microbial peptides (cAMPs) (St. Leger et al. 1991). Among the proteases found in EPF, the spore bound PR1 and PR2 endoproteases have been well characterized from M. anisopliae infecting Calliphoran vomitoria L. and Manduca sexta L. The secretion of PR1 and PR2 and their role in cuticle invasion has also been established (St. Leger et  al. 1994; Hussain et  al. 2010).

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Subsequently, an appressorium undergoes a series of morphological changes at ­different layers such as infection pegs (epicuticle), penetrant hyphae and penetrant plates (procuticle), and hyphal bodies or blastospores (hemocoel), in order to disperse through the hemocoel (Hajek and St. Leger 1994; Bhattacharyya et al. 2004). Though molecular mechanisms of spore adhesion and host recognition have been understood to a great extent, host signals/ligands and transcriptional factors as activated by protein kinase signal pathways are yet to be researched. Different fungal species obtain energy from their insect hosts through biotrophy (nutrition derived only from living cells, which ceases once the cell dies), necrotrophy (killing and utilization of dead tissues), and hemibiotrophy (initially biotrophic and then becoming necrotrophic). Generally deuteromycetes kill cells through the release of toxins. Beauvericin, bassianolide, isarolides, and beauverolides are some of the toxins isolated from B. bassiana (Hamill et  al. 1969; Elsworth and Grove 1977), while destruxins (DTXs) A, B, C, D, E, F and cytochalasins are isolated from M. anisopliae-infected hosts. DTXs depolarize the muscle membrane by activating Ca+ channels and inhibit hemocytes (Bradfisch and Harmer 1990). Neuroactive mycotoxins such as penitrem A, verruculogen and alfatrem appear to interact with glycine- and GABA-receptors, thus reducing glycine production resulting in tremorgenic effects. Evasion of host immunity in insects is an exciting field, less researched in India. B. bassiana infected and killed adults, larvae and eggs of Dicladispa armigera Olivier (Coleoptera: Chrysomelidae) as the process of pathogenesis have been studied and described (Figs. 14.2 and 14.3a–d) (Hazarika and Puzari 1990, 1995; Puzari et  al. 1994). In an interaction study between pathogen and hemocytes, it was observed that B. bassiana colonized D. armigera hemocoel by evading host acute immune system including hemocyte recognition, encapsulation, melanization and phagocytosis as well as AMP production (Phukan et al. 2008). In this insect it was revealed that granulocytes performed phagocytosis, encapsulation and nodulation of blastospores (Phukan et al. 2008). However, cells react after a specific period of infection time. Das (2015) reported that B. bassiana (Strain KR855715) at 1 × 109 conidia/ml caused 60% mortality of Corcyra cephalonica Stainton, whereas, the same strain at 1 × 107 conidia/ml caused 64–72%, 52–68% and 16–36% mortality in 3rd, 4th and 5th instars of Periplanta americana, respectively, within 10 days of treatment (Mudoi et al. 2017). Several studies showed that high hemocyte loads led to successful encapsulation not only in Drosophilla melanogaster (Basset et al. 2000), but also in Helopeltis theivora (Baruah and Hazarika 2006). When H. theivora hemocytes are challenged by B. bassiana, cell disintegration followed by clumping, capsule and nodule formation around B. bassiana took place (Baruah and Hazarika 2006). Hyphae of Aspergilus flavus link are engulfed by immune system cells of Blatella (Kulshrestha and Pathak 1997). However, cells react after a specific period of infection time (Gray and Anderson 1983; Baruah and Hazarika 2006). When the object is big enough to phagocytose, encapsulation occurs by forming multi-layered aggregates (Lackie 1988). Similar to our observation, in response to B. bassiana inoculation, hemocytes of Leptinotarsa decemlineata  (Say) also disintegrate (Sirotina 1961).

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Sewify and Hashem (2001) reported an increase in the hemocyte population of G. mellonella following infection by Metarhizium up to 96 h. Drosophila that is naturally infected by EPF exhibits an adapted response by producing antifungal peptides like dorsomycin. Based on these findings and others mainly by Schmid-Hempel (2005), a diagram is constructed to show how the fungus and insect interacts (Fig.  14.2). Bangham et  al. (2006) also developed a similar scheme describing four immune responses of D. melanogaster against major challenges faced by insects, in which (1) differentiation of LA forms a melanized capsule around the egg of a parasitoid, (2) triggering of Toll pathway occurs in response to Gram+ bacteria and fungi, (3) Gram-bacteria trigger Imd pathway, and (4) viruses trigger Jak-STAT pathway, resulting in transcription of antiviral genes.

14.4  Molecular Techniques for Species Characterization Being ubiquitous, EPF may have a massive number of strains. In general, species are separated based on morphological characteristics of spores, colonies growth or nutrient requirements from earlier days, however, molecular identification techniques are actually more appropriate. PCR-based tools have made it possible to understand the phylogenetic characterization. A number of unspecific DNA based methods have been used specially in Beauveria (Glare et al. 2008). DNA polymorphism using RAPD-PCR (Random Amplified Polymorphic DNA  – Polymerase Chain Reaction) is a widely used molecular technique in identification of this group (Joshi and St Leger 1999). It was introduced in 1990 (Samsinakova et al. 1983) to reveal polymorphism within completely unknown samples without probe hybridization or DNA sequencing, using short oligonucleotide primer (6–12 bases). RAPD has already been used to estimate the diversity of a population, genotype characterization or constructing the molecular phylogeny of closely related taxa (Tigano-Milani et al. 1995). This is based on a product which may be a spectrum of DNA fragments differing from each other in terms of length and nucleotide sequence. The application of RAPD markers is similar to those of other DNA polymorphism detection methods and can be used for characterization of a fungal isolate by constructing a specific fingerprint, or for genetic stability testing of an individual isolate. In India, using a 28S rDNA technique Uday et al. (2017) identified a potent xylanase producing strain of A. niger Tiegh. (KP874102.1) growing in various substrates such as beech wood xylan, oat spelt xylan and CM cellulose, which is a potential candidate for enzymatic hydrolysis of orange peel. Till now, 26 species of insect pathogens including 23 species of Ascomycete and 3 species of Entomopothoromycota have been sequenced (Wang and Wang 2017), of which Metarhizium spp. predominate. This kind of molecular studies will help in studying fungus-environment interaction that benefit agriculture, environment and human health.

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Universally primed (UP)-PCR has been used to separate genetically sympatric isolates of Beauveria (Meyling and Eilenberg 2006). Thakur et al. (2005) studied 48 isolates of indigenous strains of B. bassiana employing protease zymography and RAPD analysis, revealing high genetic and biochemical diversities amongst the strains collected from lepidopterans and coleopterans. RFLPs (Restriction Fragment Length Polymorphism), AFLPs (Amplified Fragment Length Polymorphism), ISSR (Inter-Simple-Sequence Repeats), SSRs (Simple Sequence Repeats), or microsatellites are other molecular techniques utilized for the characterization of both Beauveria and Metarhizium (Sandhu et al. 2012). Recent development of microsatellite markers (Rehner and Buckley 2003; Enkerli et al. 2005) linked B. bassiana to plants as an endophyte (Arnold and Lewis 2005), as M. anisopliae, that was associated with the rhizosphere of plants (Hu and Leger 2002).

14.5  Commercialization of Entomopathogenic Fungi Due to their variable performances under different environmental conditions, commercialization and large scale application of biopesticides have been slowed down during the last decade (Wang et  al. 2003; Fravel 2005). The global biopesticide market is continuously progressing, and it is estimated to be valued at $1.16 billion in 2015. It is expected to reach $3.18 billion by 2021 at a Compound Annual Growth Rate (CAGR) of 18.3% from 2016 to 2021. North America shares 44% of biopesticides market, followed by the European Union (20%) and Oceania (20%), South and Latin American countries (10%), India and other Asian countries (6%) (Khachatourians 2009; Bailey et al. 2010). Out of about 1400 biopesticides available worldwide, 202 products including 102 microorganisms were registered in USA (Chandler et al. 2011). H. thompsonii was the first mycoinsecticide registered in the U.S. under the trade name of Mycar, used against spider mites (Kenneth et al. 1979). Amongst all the 171 products of EPF so far developed, B. bassiana-based products dominate with 58 formulations representing 33.9% of the total, followed by 49 products of L. lecanii and 11 products of M. anisopliae representing 33.9%, with I. fumosorosea and B. brongniartii representing 5.8 and 4.1 per cent, respectively (de Faria and Wraight 2007): They are used against insect pests of coffee, bean, cabbage, corn, potato, and tomato and mosquitoes and flies (Florez 2002). Some of the commercial products registered across the globe are listed in Table 14.2. Compared to bacteria (285 species and strains), EPF registered in India are more (638 species and strains) along with nuclear polyhedrosis viral (37), pheromone (2), botanical products (1) and 7 others (Central Insecticide Board and Registration committee, New Delhi, http://cibrc.nic.in/) (Fig. 14.4). Amongst the EPF antagonists registered for agricultural pest management, 108 products are based on B. bassiana, followed by 95 with L. lecanii, 38 with M. anisopliae, 31 with P. lilacinus, 5 with P. chlamydosporia and 1 with H. thompsonii (Table 14.3).

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Table 14.2  Commercial microbials from various countries Country Beauveria bassiana France India India India India India India India

Trade name Ostrinil BioGuard Rich Bio-Power Racer Daman Beavera Brigade Bio-Be-Ba

India India India India India India Spain

Baba Mycojaal Metabeave Jas Beesi BBC Toxin Trichobass-L & Trichobass-P Africa Bb Plus & Bb Weevil USA Balance USA BotaniGuard & Mycotrol USA CornGard USA Naturalis L USA Naturalis H & G USA Naturalis T & O USA Organigard Colombia Agronova

Lecanicillium lecanii

UK

Mycotal

Russia Spain India India

Verticillin Trichovert Bio-Catch Biovert Rich

India India India India India India India

Mealikil Vertimust Biogade-V Vertifire-L Cropfit Varsha Biosar

Manufacturer Natural Plant Protection (NPP) Plantrich Chemicals & Biofertilizers Ltd. T. Stanes & Company Limited Agri Life International panacea Ltd. Jai Biotech Industries Kan Biosys Pvt Ltd. Microplex, Nagarjuna Agro Chemicals Multiplex Bio Tech Pvt. Ltd. Pest Control (India) Pvt. Ltd. R. B. Herbal Agro Shri Ram Solvent Extractions Sri Biotech Laboratories India Ltd. Varsha Bio Science & Technology Trichodex S. A. Biological Control Products SA Ltd. Jabb of the Carolinesn Inc. Laverlam International Corporation, USA Mycotech Corp., USA Troy BiosciencesInc., USA Troy BiosciencesInc., USA Troy BiosciencesInc., USA Emarald BioAgriculture Corp., Live Systems Technology S. A., Colombia Koppert Biological Systems, Netherlands Biodron, Russia Trichodex S. A., Spain T. Stanes & Company Limited, India Plantrich Chemicals & Biofertilizers Ltd., India Agri Life, India Jai Biotech Industries Kan Biosys Pvt. Ltd. International panacea Ltd. Microplex Multiplex Bio Tech Pvt. Ltd. R. B. Herbal Agro (continued)

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Table 14.2 (continued)

Metarhizium anisopliae

Country India

Trade name Jasverti

India India Brazil Colombia Holland Spain

Spider Shock Vertinat Ago Biocontrol Vertalec Trichomet

India India

Bio-Magic Biomet Rich

India India India India India India India

Pacer Kalichakra Cropmet Metrocid Metaz Metarhizium Jasmeta

Manufacturer Shri Ram Solvent Extraction Pvt. Ltd. Sri Biotech Laboratories India Ltd. Varsha Bioscience & Technology Natural Rural, Brazil Ago Biocontrol, Colombia Koppert, Holland Trichodex S. A., Spain

India Biostorm India Multiplex Australia BioCane & Chafer Guard USA Tacnure USA Bio-BlastTM Germany BIO 1020

T. Stanes & Company Ltd., India Plantrichs Chemicals & Biofertilizers Ltd. Agri Life, India International panacea Ltd. Microplex Sri Biotech Laboratories India Ltd. Jai Biotech Industries Multiplex Bio Tech Pvt. Ltd. Shri Ram Solvent Extraction Pvt. Ltd. Varsha Bioscience & Technology Multiplex Bio Tech Pvt. Ltd Becker Underwood Inc., USA- Australia Novozymes Biologicals Inc., USA EcoScience, USA Bayer, Germany

Source: Modified after Reddy et al. (2013)

Fig. 14.4 Microbial product registered for use in India

Bacterial 30%

NPV 4% Fungus 66%

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Table 14.3  Entomopathogenic fungal antagonists registered in India Sl. No. 1 2 3 4 5 6

Name of the entomopathogen Beauveria bassiana Lecanicillium (Verticillium) lecanii Metarhizium anisopliae Paecilomyces lilacinus Pochonia chlamydosporia Hirsutella thompsonii

Numbers of registered products 108 95 38 31 5 1

14.6  Use of Fungal Antagonists in Pest Management Ecology based pest-management strategies, biological control agents and environment-­friendly natural pesticides, of either indigenous origin or introduced, may create a sustainable crop production system in which pest species densities are maintained below the economic injury level. Utilization of microorganisms or their by-products for the control of insect pest species are of prime importance in this context. Beauveria and Metarhizium are the two important genera that are commercially exploited targeting 700 species under Lepidoptera, Hemiptera, Homoptera, Orthoptera, Diptera and Coleoptera (Moore and Prior 1996; Moore et al. 1996; Hazarika and Puzari 1995, 1997; Hazarika et al. 2005; Khan and Ahmad 2015), of which, however, M. anisopliae is dominating (Li et al. 2010; Dutta et al. 2012). They are found to be most effective against immature stages of Hemiptera, Diptera, Coleoptera, Lepidoptera, Orthoptera and Hymenoptera than vs the adult stages, while some others such as Aschersonia aleyrodis Webber and M. rileyi Farlow have restricted host ranges of only whiteflies and lepidopteran larvae respectively. Metarhizium (formerly Nomuraea) rileyi is mostly used against Spodoptera litura F. (Ignoffo 1981), Trichoplusia ni Hubner, Heliothis zea Boddie, Hypena scabra F. and Anticarsia gemmatalis Hubner (Mathew et  al. 1998). Lecanicillium. lecanii was found to be effective against whitefly and several aphids species, including the green peach aphids (Myzus persicae Sulzer) in th greenhouse chrysanthemums during the ‘70s (Hamlen 1979). The application of B. bassiana to control the pine moth, Dendrolimus spp, in China probably represents one of the largest uses of a biocontrol agent over 1 million hectares of pine forest. Strain Bb-147 of B. bassiana is registered on maize for controlling Ostrinia nubilalis Hübner and O. furnacalis Guenee. In addition to this, the strain GHA is registered in US for controlling aphids, thrips, whitefly and mealybugs. Strain ATCC 74040 is registered against many soft-bodied Homoptera, Heteroptera and Coleoptera. Beauveria brongniartii Saccardo is registered on sugarcane and barley for controlling white grubs and cockchafers. Various studies revealed that M. anisopliae (var. acridum) is effective against the brown locust, Locustana pardalina Walker in Africa, Locusta migratoria L. in Madagascar and the Australian plague locust, Chortoicetes terminifera Walker and L. migratoria in Australia. M. flavoviride Gams and Roszypal has also been tested

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against the tree locust, Anacridium melanorhodon Walker in Sudan, the rice ­grasshopper, Hieroglyphus daganensis Krauss in Benin, Mali and Senegal and the desert locust, Schistocerca gregaria Forsskal in Mauritania (Rosell et al. 2008). The National Bureau of Agriculturally Important Insects (NBAII) in Bangalore is maintaining 77 isolates of B.  bassiana, 3 isolates of B. brongniartii, 39 of M. anisopliae, 35 of Lecanicillium spp., 37 of N. rileyi, 3 of I. fumosorosea and 3 of I. farinose, collected from various insect hosts from different geographical regions in the country. Ramanujam et al. (2014) have identified several potential strains of B. bassiana, M. anisopliae, Lecanicillium spp. and N. rileyi active against H. armigera, S. litura, Aphis craccivora Koch, Aphis gossypii Glover, Rhopalosiphum maidis Fitch, Brevocoryne brassicae Linnaeus, Myzus persicae Sulzer and Bemisia tabaci Gennadius across the country. Moreover, some promising strains of N. rileyi, B. bassiana and M. anisopliae have been identified against pests of soybean, groundnut and sugarcane (Table 14.4). In order to have improved fungal antagonist formulations, technologies pertaining to oil formulation of B. bassiana for management of pests of oilseed crops have also been developed. Several technologies including B. bassiana based mycoinsecticide for D. armigera management at Assam Agricultural University, Jorhat (Das et  al. 2017) and M. anisopliae-based formulation for Oryctes rhinoceros L. at CPCRI, Kayangulam were developed (Ramanujam et al. 2003). B. bassiana (Strain No. KR855715) at the rate of 1 × 109 conidia/ml showed effective results against Table 14.4  Commercial products of entomopathogenic fungi and their target pests Fungus B. bassiana

B. brongniartii

M. anisopliae

M. flavoviride V. lecanii

P. fumosoroseus

Brand name Mycotrol Naturalis Conidia Ostrinol Myco-Jaal Biosoft Biowonder Betel Engerlingspilz Melocont BIO 1020 Bio-BlastTM Bio magic Multiplex BioGreen Vertalec Biocatch Verticare PFR-97TM Prioroty

Target pests Whiteflies/aphids/thrips Sucking insects Coffee berry borer Corn borer Diamond backback moth Helocoverpa & sucking pests Rice pests Scarab beetle larvae Scarab beetle larvae Scarab beetle larvae Black Vine weevil Termites Brown plant hopper Root grubs Red-headed cockchafer Aphids, whiteflies Whiteflies Mealybugs & scales Whiteflies/thrips Mites

Crop Field crops Cotton, glasshouse crops Coffee Maize Cabbage Several crops Rice Sugarcane Pasture Pasture Glasshouse ornamental House Rice Several crops Pasture/turf Glasshouse crops Cotton Citrus Glasshouse crops Wide range of crops

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P. Das et al.

sixth instars of C. cephalonica, as reported by Das (2015). Wettable powder ­formulation of B. bassiana at 10 g/kg of rice seeds showed the highest mortality of C. cephalonica (75%) at 7  days of treatment. On the other hand, Mudoi (2016) observed that the same B. bassiana (Strain No. KR855715) was also effective against the 1st, 2nd and 3rd instars of P. americana, at a concentration of 1  × 107 conidia/ml. In the light of increasing awareness about adverse effects of pesticide residues in food on human health, microbial product based pesticides play an important role in India. Several works on using mycoinsecticides have been carried out in the country for controlling different pests of rice, pulse, oilseeds, tea, sugarcane, coffee, coconut, house hold pests and storage pests as well (Table 14.5). Table 14.5  Biological Control of pests using entomopathogenic fungi in India Fungus B. bassiana

Crop Rice Coffee Tea Sunflower Green gram Storage

Insect controlled Rice Hisp, Dicladispa armigera Coffee berry borer, Hypothenemus hampei Tea looper, Buzura suppressaria Head borer, Helicoverpa armigera White grubs species

Rice moth, Corcyra cephalonica Household Cockroach, Periplaneta americana B. Brongniarti Sugarcane White grub, Holotrichia serrata M. anisopliae Coconut Rhinoceros beetle, Oryctes rhinoceros Sugarcane White grub species Pigeon pea Pod borer, H. armigera Potato White grub, Brahmina sp. Soyabean White grub, Holotrichia longipennis L. Lecanii Coffee Green scale, Coccus viridis Citrus Green scale, C. viridis Mustard N. rileyi

Castor Soybean

Mustard aphid, Lipaphis erysimi Tobacco caterpillar, Spodoptera litura S. litura, H. armigera, Thysonoplusia orichalcea

Dose 1 × 107 spores/ml 1 × 107 spores/ml + 0.1% sunflower oil + 0.1% wetting agent 2.5 g/l 200 mg/l 5 × 1013 conidia/ha 1 × 109 conidia/ml 1 × 107 conidia/ml + Tween- 80 (0.023%) 1 kg/acre 5 × 1011 spores/m3 1 × 1013 spores/ha – 5 × 1013 conidia/ha 5 × 1013 conidia/ha 16 × 106 spores/ml + Tween-80 2 × 106 spores/ml + quinalphos (0.005%) + Teepol (0.05%) 1 × 106 spores/ml 10 × 1010 spores/ml 2 × 108/ml

Source: Modified after Ramanujam et al. (2014), Das (2015), and Mudoi (2016)

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14.7  Economics For long-term positive effects of bio-pesticides use in sustainable pest management program, there is a growing demand and awareness among farmers and policy makers. One of the strongest benefits of bio-pesticides is that they could be combined with other pest management tactics. They are also relatively cheap to develop and need to be re-developed less frequently, saving expenditures on research and development. The efficacy of many of the bio-pesticides is equal to that of conventional chemical pesticides. However, the mode of action is different. With many of the bio-pesticides, the time from exposure to morbidity and death of the target insect may be 2–10 days in general. Due to application of bio-pesticides in tea crops (70% crop coverage) almost 9 kg of made tea increase could be achieved with a chemical cost saving of Rs. 3750 (Hazarika et al. 2001). One of the multi-location field trial studies against major pest of rice with B. bassiana and conventional insecticides revealed that the mycoinsecticides were superior to monocrotophos in controlling D. armigera, leading to an increased cost benefit ratio of 1:7.66 for B. bassiana against 1:2.92 for monocrotophos (Hazarika and Puzari 1977).

14.8  Conclusion Adverse effects of chemical pesticides on non-target organisms, food safety, environmental issues and development of pest resistance have forced the scientific communities to focus on the development of alternative eco-friendly measures. EPF have a key role in sustainable pest management programs in many agricultural systems. However, the large-scale use of EPF as bio-pesticides can be hindered by a number of issues. The major barriers include the inoculums short or poor shelf life span, requiring a couple of weeks to kill the target pests, the need for a fungicide-­ free environment, high relative humidity (> 80% RH), poor quality of some commercial products, high-cost of commercial formulations and the possibilities of contamination with mycotoxins (aflatoxins, trichothecenes, zearalenone, fumonisins, citrinin, etc.). Hence, there is a need to promote R & D activities on EPF to increase shelf-life of their formulation up to 12–18 months, by adding additives and adjuvants including oil, sunscreen UV- and IR-protectants to make products more active and durable in the field (wet land and dry land), better storage conditions under different environments and ecosystems. Moreover, there is a tremendous scope to utilize the biodiversity of Entomophthora, Zoophthora, Neozygites fungi, belonging to the Entomophthorale group, which have a high potential for management of sucking and lepidopteran pests. Advanced techniques in molecular biology have the potential to manipulate desirable traits of EPF to improve overall field activity. There is also the further need for understanding the host-parasite interactions, in relation to crop ecosystem. This is vital as every crop ecosystem is unique and provides different microclimatic

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c­ onditions in which the target pests evolved a number of mechanisms to keep the pathogen at bay. Though EPF have great potential, R & D activities for their commercial exploitation as inundative, inoculative and classical biopesticides are not adequate to meet the actual huge demand. In order to achieve a successful EPFbased pest management strategy, a paradigm shift in the mindset of farmers is needed. The Government and the industry jointly may develop a mission for the popularization of EPF. Acknowledgements  The authors are grateful to Assam Agriculture University and Tocklai Tea Research Institute, Jorahat, Assam, India, for the facilities provided for conducting the study. Declaration Statements  Ethics approval and consent to participate This article does not contain any studies with human participants or animals performed by any of the authors, so not applicable. –– Consent for publication: Not applicable Disclosure-All the experiments undertaken in this study comply with the current laws of the country where they were performed. –– –– –– ––

Availability of data and material: As it is review article it is not applicable Competing interests: The authors declare that they have no competing interests. Funding: As it is review it is not applicable Authors’ contributions (Do not include any authors’ information)

LKH with PD and SR conceived the idea of the manuscript. PD, SK and SR participated in preparing first draft of the manuscript. PD, SK and SR conducted literature surveys. All authors read and approved the final manuscript.

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Index

A Abacarus hystrix, 166 Abamectin, 310, 324, 346, 348, 358 Acetamiprid, 309, 310, 329 Acetylcholine esterase, 333, 352 Acrididae, 352 Actinedida, 263 Acylamidase, 353 Adalia bipunctata, 254, 255 Adoxophyes honmai, 65, 76 Adoxophyes orana, 65, 69, 76, 279 Adoxophyes orana GV (AdorGV), 69, 279, 280, 287, 288 Agrius convolvuli, 224 Agroecosystem, 14, 24, 132, 134, 139, 290, 334, 342, 343, 345, 346, 348, 354 Agromyzidae, 261 Agrotis ipsilon, 57, 65, 254, 255, 279, 368 Agrotis segetum, 57, 59, 65, 128, 279, 287 Aleurocanthus spiniferus, 165 Aleyrodes citri, 165 Alfa alfa looper, 44, 56 Allantonematidae, 249 Alphabaculovirus, 276 Alternaria alternata, 172 Amblyseius swirskii, 306 American bollworm, 55, 161 Aminopeptidases, 156, 370 Amsacta moorei, 71 Anagasta kuehniella, 26 Anagrapha falcifera, 56 Anamorph, 33, 34, 92, 94, 96, 127, 133, 134, 138, 155, 161, 165, 186 Annona squamosa, 332 Anomala cuprea, 307, 334

Anopheles aegypti, 170 Anopheles gambiae, 87, 171 Antagonists, 94, 173, 208, 342–359, 366–380 Antheraea proylei, 72 Anthocoridae, 262 Anticarsia gemmatalis, 55, 64, 120, 164, 368, 376 Antifeedant, 156, 307 Antioxidant, 190, 198–202, 285, 353 Aphidius colemani, 306 Aphidoletes aphidimyza, 259, 262, 264 Aphis craccivora, 164, 166, 368, 377 Apidae, 54, 260, 261 Apis mellifera, 54, 91, 130, 166, 257, 260, 305, 306 Arabidopsis thaliana, 213 Arabinose, 200 Arabitol, 190 Araneidae, 263 Archaea, 77, 79, 88, 170, 212 Archiascomycetes, 154 Arctiidae, 76 Armadillidiidae, 262 Armadillidium vulgare, 262 Aschersonia aleyrodis, 160, 166, 168, 189, 376 Aschersonia placenta, 189 Ascomycota, 24, 28, 31, 48, 90–92, 94, 133–134, 152–173, 344, 366 Ascosphaera, 91, 133 Ascosphaera aphis, 91 Ascoviruses, 72 Asexual spore, 25, 89, 157, 186 AsGV, 279, 287 Aspergillus, 133, 155, 162 Aspergillus niger, 190, 372

© Springer Nature Switzerland AG 2019 M. A. Khan, W. Ahmad (eds.), Microbes for Sustainable Insect Pest Management, Sustainability in Plant and Crop Protection, https://doi.org/10.1007/978-3-030-23045-6

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388 Autographa californica, 56, 62, 64, 76, 304 Autographa californica multiple nucleopolyhedrovirus (AcMNPV), 58, 62, 64, 73, 76, 278, 281 Avermectin, 121, 308, 309 Azadirachtin, 307–310, 358 Azinphos-ethyl, 348 Azocyclotin, 355 Azoxystrobin, 347 Azygospore, 31, 131, 137 B Bacillaceae, 77, 79, 88, 117, 120 Bacilli, 77, 79, 81 Bacillus spp., 26, 35, 77, 81–84, 88, 116, 117, 120, 125, 299 B. anthracis, 83–85 B. larvae, 81 B. licheniformis, 83 B. popilliae, 78, 79, 81, 82, 330 B. sphaericus, 47, 78, 80, 81, 83, 119, 330 B. thuringiensis, 12, 26, 27, 35, 46, 48, 51, 77–79, 81–87, 116, 117, 119, 125, 126, 167, 168, 289, 301, 321, 329, 330, 332, 344, 358, 366 B. thuringiensis thompsoni, 78 B. weihenstephanensis, 84 Bacteriophages, 78, 82 Baculoviridae, 55, 60–62, 276, 277 Baculovirus, 45, 54–58, 61–70, 72–75, 277, 278, 280, 283–286, 289, 290 Bagrada hilaris, 308 Banana weevil, 167 Basidiobolus ranarum, 96 Basidiomycota, 24, 28, 91, 92, 132, 154, 366 Bassianolide, 156, 159, 371 Beauveria, 31, 89, 90, 94, 128, 134, 138, 155, 161, 167, 171, 172, 186, 187, 344, 369, 372, 373, 376, 377 Beauveria brongniartii, 92, 138, 153, 167, 168, 329, 373, 376, 377 Beauveria pseudobassiana, 201 Beauvericin, 156, 158, 371 Beauverolides, 156, 371 Bembidion properans, 255, 260 Bemisia tabaci, 12, 164, 308, 329, 331–333, 350, 353, 368, 377 Benzoate, 346 Benzylideneacetone (BZA), 222 Betabaculovirus, 276, 281 Bifenthrin, 309, 310 Biodiversity, 24, 173, 335, 379

Index Biopesticides, viii, 34–36, 44, 51, 78, 81, 82, 92, 116, 117, 128, 138, 139, 158, 163, 167, 276–278, 285, 286, 290, 334, 343, 366, 373, 380 Biotrophic, 25, 31, 34, 90, 371 Black vine weevil, 301, 377 Blaniulidae, 263 Blaniulus guttulatus, 262 Blastocladiomycota, 24, 91, 92, 130–131, 366 Blastopore, 135, 138, 158, 167, 301, 303, 344, 350, 371 Blattella germanica, 327, 349 Blattodea, 33, 349 Bombus spp., 260 B. impatiens, 306 B. terrestris, 257, 261, 306 Bombycidae, 54, 76 Bombyx mori, 26, 46, 54, 58, 63, 64, 76, 86, 116, 164 Bombyx mori nucleopolyhedrovirus (BmNPV), 54, 64 Boophilus annulatus, 262 Botrytis cinerea, 306 Brevibacillus, 88 Brevibacillus laterosporus, 26, 27, 48, 120–121, 125 Brevipalpus phoenicis, 166 Brown plant hopper, 308, 377 Bumble bees, 44, 260, 306 C Cabbage moth, 55, 56 Caenorhabditis elegans, 87, 213 Calacarus heveae, 166 Calosoma granulatum, 256, 260 Carabidae, 254, 260 Carbamate, 2, 3, 12, 13, 247, 329–331, 342, 346, 357 Carboxymethylcellulose, 191 Carboxypeptidase, 157, 158 Catalase, 120, 190, 199, 200, 202, 333, 353 Cecidomyiidae, 262 Cellular immunity, 225, 301 Cereal rust mite, 166 Chestnut weevil, 301 Chilo spp. C. infuscatellus, 278, 279, 283, 287, 288, 368 C. iridescent, 71 C. sacchariphagus, 278, 287 Chitinase, 123, 138, 156–159, 170, 231, 308, 333, 344, 345, 353, 370

Index Chlorantraniliprole, 352, 353, 358, 359 Chlorpyriphos, 2, 263, 347, 358 Choristoneura fumerferana, 281 Chromobacterium substugae, 26 Chrysanthemum, 162, 264, 376 Chrysoperla zastrowi sillemi, 253 Chrysopidae, 253, 261 Chymoelastase, 158 Chymotrypsins, 157 Chytridiomycota, 24, 28, 90–93, 130, 154 Cicadellidae, 165 Citrus white fly, 165 Clavicipitaceae, 33, 93, 97, 133, 134, 186 Clofentezin, 355 Clostridium, 81 Clostridium bifermentans, 26, 48, 121 Clothianidin, 325–328 Cnephasia spp., 288 Cocci, 79 Coccinella undecimpunctata, 254 Coccinellid, 44, 302 Codling moth, 44, 56, 128, 152, 161, 261, 286, 288 Coleomegilla maculata lengi, 302 Coleoptera, 29, 32, 33, 51, 60, 77, 81, 83, 86, 87, 89, 120, 123, 124, 127, 155, 159, 160, 162, 164, 166, 208, 249, 252–260, 348, 349, 355–358, 368, 371, 376 Colias eurytheme, 54 Collembola, 263 Colorado potato beetle, 26, 123, 152, 161, 301, 348 Conidia, 30, 31, 33, 89, 132, 152, 186, 198, 302, 319, 345, 371 Conidiobolus coronatus, 96 Conidiophores, 30, 132, 159, 302 Coptotermes formosanus, 308 Cordyceps spp., 31, 33, 94–97, 127, 133, 154, 165 C. bassiana, 33, 161 C. militaris, 95, 96 Cordycipitaceae, 33, 93, 133, 134, 161, 186 Cosmopolites sordidus, 167, 357 Crioceris quatuordecimpunctata, 301 Cry genes, 86, 119 Cry proteins, 27, 78, 81, 121, 126, 127 Crypticola entomophaga, 90 Crystalliferous, 26 Culex nigripalpus, 70, 73 Culex pipiens, 87, 164 Culicinomyces, 155 Curculio elephas, 301 Curculionidae, 260, 348, 349, 357

389 Cuticle, 31, 33, 49, 90, 92, 116, 135, 136, 138, 155–160, 169, 186, 198–200, 209–212, 218, 229, 248, 302, 303, 308, 310, 324, 327, 328, 344, 351, 356, 369, 370 Cydia pomonella, 51, 55, 56, 69, 86, 128, 261, 279, 281, 286–289, 367 Cydia pomonella granulovirus (CpGV), 69, 279, 281–283, 288, 289 Cyfluthrin, 327, 349 Cypermethrin, 2, 326, 328, 330, 347, 349, 356, 357 Cypovirus, 45, 54, 62, 63, 278 Cytolysin, 220, 223, 229 Cytoplasmic polyhedrosis virus (CPV), 45, 59, 62, 63, 70, 278 Cytotoxic factors, viii, 208–232 Cytotoxins, 220, 223, 226, 227, 229 D Daktulosphaira vitifoliae, 152 Danio rerio, 213 Dasyhelea obscura, 154 Decomposers, 24, 133, 153 Deltabaculovirus, 276 Deltamethrin, 2, 14, 326, 330, 347, 351, 356–358 Dendrolimus punctatus, 63 Dendrolimus spp., 161, 167, 376 Dermaptera, 33, 258, 263 Deuteromycota, 154 Dextruxin, 158 Diafenthiuron, 355 Dialeurodis citri, 165, 350 Diamondback moth, 55, 120, 122, 166, 168, 253, 304 Diaphania nitidalis, 261 Diaprepes abbreviatus, 328, 331, 333 Diatraea saccharalis, 69, 279, 301 Dichloro-diphenyl-trichloroethane (DDT), 2–4, 6, 7, 10, 348 Difenoconazole, 347 Diglyphus begini, 256, 261, 264 Dikarya, 91, 93, 154 Diptera, 29, 32, 33, 51, 56, 60, 62, 63, 70, 73, 77, 81, 86, 87, 89, 120, 123, 127, 130, 160, 166, 208, 249, 253, 259, 261, 262, 276, 350, 376 DNA-DNA hybridization, 52, 81, 84 DNA helicase, 73 DNA hybridization, 73, 81 DNA polymerase, 73 DNA polymorphism, 372

390 DNA sequencing, 74, 80, 372 DNA viruses, 277 Double-stranded DNA (dsDNA), 60, 71, 86, 277 Double-stranded RNA (dsRNA), 59, 277 Drosicha mangiferae, 253 Drosophila, 46, 62, 213–220, 372 Drosophila melanogaster, 123, 213, 214, 220, 372 dsRNA viruses, 277 Duponchelia fovealis, 165 E Ecosystems, 2, 4, 7, 15, 33, 55, 89, 94, 153, 155, 171, 208, 260, 276, 284, 290, 318, 323, 344, 379 Ectobiidae, 349 Edwardsiella tarda, 227 Egg parasitoid, 253 Eicosanoids, 222, 224 Electron microscopy, 54, 202 Endochitinas, 157 Endophytes, 31, 82, 94, 97, 116, 125, 134, 153, 373 Endoproteases, 156, 158, 370 Enterobacteriaceae, 78, 79, 117, 122–123, 208, 224, 227, 248, 249 Entomophaga aulicae, 96 Entomophthorales, 24, 25, 28–34, 94, 96, 116, 131, 134–138, 344 Entomophthora spp., 32, 48, 90, 94, 132, 299, 344, 345, 379 E. anisopliae, 157 E. virulenta, 167–169 Entomophthoromycetes, 24, 29–31 Entomophthoromycota, 24, 28, 29, 90, 91, 93, 95, 96, 131–132 Entomopoxviruses (EPV), 45, 59, 62, 71, 278, 319, 321, 322, 331, 334 Entomopthoromycotina, 24, 29–31 Ephestia kuehniella, 253, 329 Epicuticle, 31, 158, 159, 198, 200, 369, 371 Epidermis, 218, 369 Epithelial cells, 119, 224, 280 Epithelium, 27, 28, 48, 119, 120, 171, 208, 210, 224, 228, 229, 280 Epizootics, 29, 31, 55, 56, 82, 92, 94, 120, 162, 164, 186, 188, 276, 287, 288, 290, 318, 334, 345, 346, 367 ErelGV, 69, 279, 289 Erinnys ello, 279, 286, 289 Erynia, 32, 132, 137, 299

Index Erythritol, 190 Erythrocytes, 223–227 Escherichia coli, 85, 224, 283 Esterases, 156, 158, 170, 333, 351–353 Etoxazole, 355 Etridiazole, 311 Eubacteria, 79, 81, 88 Eukaryotes, 44, 79, 95, 208 Eulophidae, 261 Eurotiomycetes, 91, 133 Eutectona machaeralis, 161, 163 Exomala orientalis, 325, 326, 331 F False spider mite, 166 Fenpropathrin, 309 Filoviruses, 277 Fire ants, 171 Firmicutes, 77, 79, 117 Flonicamid, 309 Flupyradifurone, 309 Formicidae, 261, 263 Frankliniella occidentalis, 302, 304, 306, 332 Fructose, 200 Fungi imperfecti, 154 Fusarium, 89 G Galactose, 200 Galleria mellonella, 86, 170, 171, 210, 217, 223, 224, 261, 303, 351, 356, 358, 372 Gamacyhalothrin, 356, 357 Gammabaculovirus, 276 Gamma-cyhalothrin, 347 Gelechiidae, 76 Genome sequencing, 63, 83, 84, 96 Genomics, 52, 61–76, 80, 82–89, 94–97, 277, 278 Genotypes, 57, 58, 62, 73, 372 Geometridae, 76 Geranium carolinianum, 305 Gilpinia hercyniae, 55 Glassy winged sharpshooter, 165 Glomeromycota, 91, 92, 131 Glutaredoxins (grx), 199 Glycerol, 190, 370 Glycosphingolipids, 217 Glyphosate, 347 Golgi apparatus, 215–217, 219, 220 Golgi complexes, 216 Gracilicutes, 79

Index Gram-negative, 78, 79, 87, 122, 123, 208, 221, 223, 231, 248, 249 Gram-positive, 78, 79, 88, 117, 223, 229 Gram stain, 27, 79, 81 Granulocytes, 213, 214, 216–217, 225–227, 334, 371 Granulosis, 277, 278, 280, 283, 289 Granuloviruses (GVs), viii, 51, 54–56, 59, 61, 62, 72–74, 76, 276–291, 299 Grape berry moth, 55 Grape phylloxera, 152 Greater wax moth, 210, 303, 351 Green lacewing, 257, 261 Gryllidae, 263 Gylpinia herciniae, 55 Gypsy moth, 44, 303 H Habrobracon hebetor, 303 Haematopoietic tissues, 219 Haemocoel, 48, 156, 157, 159, 198, 208, 210–212, 220, 230, 248 Haemocytes, viii, 47, 208–232 Haemolymph, 47, 155, 157–160, 170, 210, 213, 216, 217, 220–222, 225, 230 Haemolysin, 211, 212, 220, 223, 225–227, 229 Harmonia axyridis, 255, 302 Harposporium, 165 Harrsinia billions, 76 Heat-shock proteins (HSPs), 172, 190 Helicoverpa spp., 12, 55, 56 H. armigera, 11, 12, 66, 70, 121, 161, 164, 231, 279, 308, 367, 368, 377, 378 H. zea, 51, 66, 304, 305, 358 Heliothis, 55, 62, 305, 329 Heliothis virescens, 72, 86, 304, 329 Helopeltis spp., 162 Hemiascomycetes, 154 Hemibiotroph, 25, 33, 90, 371 Hemiptera, 13, 14, 29, 32, 33, 51, 60, 90, 124, 138, 160, 162, 165, 166, 252, 253, 258, 262, 349, 350, 352, 368, 376 Hemocoel, 27, 30, 31, 90, 92, 119, 120, 122, 123, 138, 170, 198, 343, 344, 354, 369, 371 Hepadnaviruses, 277 Hepialidae, 76 Herpesviruses, 74, 277 Heterorhabditidae, 48, 78, 208, 248, 251 Heterorhabditis spp., 49, 79, 88, 122, 208–210, 232, 249, 257, 261, 299

391 H. bacteriophora, 51, 78, 208, 251, 253–257, 259, 261–264, 301, 325, 326, 330, 355, 358 H. indica, 88, 251, 253, 257, 259, 261, 263, 301, 326, 356, 357 H. taysearae, 252, 254 Heterotermes tenuis, 331, 333 Heterotrophic, 28, 89, 154 Hirsutella spp., 51, 94, 155, 161, 165–166, 299, 344 H. gregis, 165 H. kirchneri, 165 H. necatrix, 165 H. nodulosa, 165 H. thompsonii, 51, 96, 128, 159, 165, 166, 168, 169, 307, 367, 373, 376 Histeridae, 254 Homalodisca coagulata, 165 Homoptera, 29, 32, 78, 89, 127, 162, 376 Horizontal gene transfer, 58, 78, 82, 84, 97, 170 Humoral immune responses, 212, 222 Humoral response, 209, 217, 221–223 Hyadaphis foeniculi, 307 Hyblaeapara, 161 Hydrocarbons, 198, 200, 202, 370 Hydrolytic enzymes, 158, 229, 249 Hylobius abietis, 301 Hypanthia cunea, 279, 289 Hyperoxidant, 198, 201 Hypocreales, 24, 25, 28, 29, 31–34, 89–91, 93, 116, 134–138, 165, 186, 344 I Ichneumonidae, 14, 261 Ichneumonids, 257, 261, 264 Imidacloprid, 173, 307, 310, 322, 325–327, 330, 331, 333, 347–349, 355, 357, 358 Immune reaction, 14, 213, 221–222 Immune system, 10, 45, 53, 95, 158, 212, 217, 221–229, 232, 333, 371 Immunity, 95, 171, 212–220, 301, 369, 371 Infective juveniles (IJs), 49, 209–211, 230, 248, 249, 254–257, 259–261, 263, 301, 302, 320, 321, 323, 324, 354–357 Inositol, 200 Insect growth regulators (IGRs), 14, 307, 308, 334, 342 Integrated pest management (IPM), viii, 25, 36, 152–173, 248, 276, 300, 307, 309, 311, 318, 343, 346, 347, 359, 366 Ips sexdentatus, 302

392 Isaria, 31, 94, 134, 156, 161, 164, 172, 186, 188, 299, 344 Isaria fumosorosea, 51, 92, 153, 156, 166, 168, 187–190, 200, 303, 310, 329, 331, 350, 367, 373, 377 Isarolides, 156, 371 Isoptera, 89, 90, 208, 253, 368 Ixodes scapularis, 263 Ixodidae, 262, 263 J Jassid, 302 Junonia orithya, 164 K Kickxellomycotina, 24, 91, 131 Kinesin, 157 Kinoprene, 355 L Labidura riparia, 258, 263 Laboulbeniomycetes, 89, 91, 133, 154, 155 Lacanobia oleracea, 279, 283, 289 Lactobacillaceae, 79 Lactose, 200, 290 Lambda-cyhalothrin, 326, 327, 347 Lasiocampidae, 76 Leafminer, 261, 264 Lecanicillium, 31, 134, 155, 158, 166, 172, 299, 376, 377 Lecanicillium aphanocladii, 189 Lepidiota negatoria, 310 Lepidoptera, 11, 12, 15, 29, 32, 33, 45, 51, 54, 56, 58, 60–65, 69, 72–74, 76, 77, 81, 83, 86, 123, 124, 126, 127, 159, 160, 162, 164–166, 168, 169, 208, 213–220, 249, 253, 254, 260, 261, 276, 278, 290, 350, 351, 356, 358, 368, 376 Leptinotarsa decemlineata, 26–27, 87, 123, 301, 302, 308, 348, 367, 371 Leucoma salicis, 57, 58 Light microscopy, 59, 216 Liotryphon caudatus, 257, 261 Lipases, 138, 156, 157, 159, 211, 220 Lipopolysaccharides (LPS), 221, 223, 224 Liriomyza trifolii, 261, 264 Liviidae, 350 Loculoascomycetes, 154 Locusta migratoria, 307, 352, 353, 376 Locustana pardalina, 376 Luciaphorus perniciosus, 253

Index Lufenuron, 308, 326, 327, 346, 355–358 Lycosidae, 263 Lygus hesperus, 303, 308, 309 Lymantria dispar, 51, 55, 63, 67, 285, 303 Lymantriidae, 58, 76 Lysinibacillus, 88, 119 Lysinibacillus sphaericus, 26, 48, 78, 81, 82, 119 M Mafanoxam, 311 Mahanarva posticata, 163 Mahanarva spp., 167 Mallophaga, 78, 127 Mamestra brassicae, 55, 56, 67 Manduca sexta, 86, 170, 171, 225, 227, 370 Mannitol, 189, 190 Mantodea, 33 Mastrus ridibundus, 257, 261 Mediterranean flour moth, 26, 77, 116 Melanin, 199, 217, 218, 220, 222 Melanoplus sanguinipes, 71 Mermithidae, 49 Messor himalayanus, 261 Metarhizium spp., 31, 89, 90, 94, 95, 97, 127, 134, 138, 157–158, 161, 163–164, 167, 169–172, 199, 299, 344, 369, 370, 372, 373, 376 M. acridum, 94, 170, 188–190, 201 M. album, 94, 97, 138 M. anisopliae, 48, 91, 92, 95, 128, 135, 138, 153, 157, 160–163, 166–168, 170–172, 187–191, 199–201, 300–302, 304–307, 310, 311, 323, 327–330, 333, 334, 346–353, 367–371, 373, 375–378 M. brunneum, 51, 95, 189, 201, 303, 304, 309, 311, 327, 331 M. guizhouense, 94 M. majus, 94 M. robertsii, 95, 138, 170–172, 188–191, 200, 201, 301, 308, 347, 350 Methomyl, 2, 329, 355 Metoxyfenozide, 355 Micrococcaceae, 79 Miridae, 162 Mithymna unipuncta, 218 Mitochondrial DNA (mtDNA), 52, 154 Monophagous, 343 Monosporella unicuspidata, 154 Monoxygenase, 352, 353 Musca domestica, 87, 120, 137, 307, 350 Muscidae, 350 Mutations, 11, 50, 52, 58, 82

Index Mycelium, 121, 155, 160, 162, 320, 323 Mycoinsecticides, 31, 152, 153, 163, 166–169, 171, 172, 198–202, 367, 373, 377–379 Mycoparasites, 31, 97, 131, 134, 158 Mycorrhizae, 153 Mycosporidia, 28, 366 Mycotoxicity, 158, 173, 332, 379 Myriangiales, 133, 154 Myzus persicae, 13, 162, 166, 170, 302, 368, 376, 377 N N-acetyl-D-glucosamine, 218 N-acetyl glucosamine, 158 Necrotrophy, 90, 371 Neisseriaceae, 79, 117, 122, 124 Neocallimastigomycota, 91, 92 Neodiprion lecontei, 70, 72 Neodiprion sertifer, 62, 70 Neolecta, 154 Neonicotinoid, 325, 331, 332, 342 Neoscona theisi, 259, 263 Neoseiulus cucumeris, 303, 306 Neozygites, 299 Neozygites tanajoae, 303, 310 Nephotettix spp., 161 Neuroptera, 253, 257, 261 Nilaparvata lugens, 308, 323, 329 Noctuidae, 11, 12, 55, 76, 224, 254, 260, 350, 356, 358 Nomuraea, 94, 155, 159, 161, 162, 164, 344, 376 Nomuraea rileyi, 159, 160, 164, 168, 169, 186, 367, 368, 377, 378 Non-crystalliferous, 26 Non-spore-forming, 48, 79 Nosema, 129, 299 Notodontidae, 76 Nucelocapsids, 59 Nucleopolyhedrosis, 277, 278 Nucleopolyhedrovirus (NPVs), 51, 54–57, 59, 61, 62, 64, 65, 68–70, 72–74, 76, 276–278, 280, 281, 283, 287, 299, 304 Nudiviruses, 280 Nutrient agar, 27 Nutrient broth, 27 Nymphalidae, 76 O Occlusion bodies (OBs), 54, 56, 59, 61, 70, 72, 73, 277, 280, 285, 288, 290

393 Occlusion-derived virus (ODV), 58, 277, 278, 280 Odonata, 33, 60 Oenocytoids, 213–215, 217, 218, 220 Onychiuridae, 263 Onychiurus armatus, 263 Oomycetes, 90, 128 Oomycota, 90, 128–129 Ophiocordyceps, 90, 127 Ophiocordyceps sinensis, 94–96 Ophiocordycipitaceae, 33, 93, 133, 134 Ophiostoma, 89 Organochlorines, 2, 3, 6, 7, 10, 15, 342, 346 Organophosphates, 2, 3, 10, 12, 13, 247 Organophosphous, 324 Orgyia pseudotsugata, 55, 57, 58, 65 Oribatida, 263 Orius albidipennis, 258, 262 Orthomyxoviruses, 277 Orthoptera, 29, 33, 51, 60, 78, 123, 127, 160, 162, 166, 208, 253, 352, 376 Oryctes rhinoceros, 62, 128, 164, 368, 377, 378 Ostrinia nubilalis, 86, 128, 161, 376 Otiorhynchus sulcatus, 301 Oxamyl, 355 Oxidative stress, vii, 160, 171, 190, 198–202 Oxindole, 222 P Paecilomyces spp., 94, 155, 164, 310 P. farinosus, 161 P. fumosoroseus, 164, 166, 167, 187, 346, 347, 351, 377 P. lilacinus, 35, 51, 167, 169, 373, 376 Paenibacillaceae, 79, 117, 120–121 Paenibacillus spp., 82, 88, 120, 308 P. lentimorbus, 26, 47, 120 P. popilliae, 26, 47, 51, 78–79, 82, 120, 125 Pandora, 32, 94, 132, 137, 299, 344 Paralobesia viteana, 55 Parasporal crystals, 27, 47, 117, 126 Parvoviruses, 277 Periplaneta americana, 367, 371, 378 Peroxidase, 199, 202, 333, 353 Peroxisome, 198, 200 Pezizomycotina, 24, 31–34, 133, 154 Phaenositylenchidae, 249 Phagocytosis, 47, 199, 209, 213–216, 221, 228, 229, 334, 371 Phasmatodea, 33

394 Phenoloxidase, 222, 223, 333, 352, 353 Phorate, 2, 5, 347 Photorhabdus spp., 48, 49, 79, 81, 88, 117, 208, 209, 220–223, 225, 227, 228, 230, 232, 248, 249, 252, 253, 354 P. asymbiotica, 88, 226, 251 P. luminescens, 78, 86, 88, 89, 122, 225–227, 230, 251, 253 P. temperata, 88, 89 Phthorimaea operculella, 55, 70, 278, 279, 286, 287 Phyllocoptruta oleivora, 128, 159 Phylogeny, vii, 44–98, 133, 277, 372 Phytoalexines, 332 Picornaviruses, 277 Pieridae, 54, 76, 253, 290 Pieris brassicae, 54, 253, 278, 279, 290 Pine weevil, 301 Piperonyl-butoxide, 352, 355 Plasma membranes, 126, 217, 226, 280 Plasmatocytes, 14, 213–216, 218–220, 224, 225, 227, 334 Plasmids, 46, 78, 82–84, 86, 230 Plathypena scabra, 164 Pleosporales, 133, 154 Plutella xylostella, 32, 55, 56, 65, 86, 120, 166, 200, 253, 278, 279, 285, 304, 333 Plutellidae, 76, 253 Pollinators, 14, 44, 260, 305 Polydnavirus (PDV), 45, 280 Polymerase Chain Reaction, 372 Popillia japonica, 51, 79, 83, 87, 120, 128, 323, 325–328, 331 Porcellionidae, 262 Porcellio scaber, 262 Potato dextrose broth, 319 Potato tuber moth, 55 Poxvirus, 71, 277 Profenofos, 346 Prohaemocytes, 214, 215, 219 Prokaryotes, 44, 77, 87, 88 Propamocard, 311 Propiconazole, 310, 347 Prostephanus truncatus, 302 Proteases, 27, 28, 123, 126, 138, 156–158, 170, 212, 220, 223, 224, 344, 370, 373 Proteinases, 159, 225 Proteobacteria, 78, 117, 122, 223 Proteus mirabilis, 224, 226, 227 Protoplasts, 25, 31, 34, 90, 137, 173 Protozoa, 13, 24, 44, 49, 78, 127, 129, 213, 229 Pseudaletia unipuncta, 70, 224, 279, 281

Index Pseudomonadaceae, 79, 88, 122, 123 Pseudomonas spp., 81, 123, 321 P. entomophila, 48, 88, 123, 226 P. putida, 88, 123, 125 Pseudoplusia, 164 Pseudopodia, 215, 216, 219, 220 Pterostichus cupreus, 255, 260 Pucciniomycotina, 91 Pyralidae, 54, 76, 224, 253, 261, 351 Pyrenomycetes, 154 Pyrethrins, 3, 308, 309 Pyrethroids, 2, 3, 11, 12, 14, 329–331, 347, 357, 358 Pyrilla purpusilla, 163 Pyriproxyfen, 347, 355 R Rachiplusia nu, 350 Rachiplusia ou, 65, 76 Randomly amplified polymorphic DNA (RAPD), 52, 372, 373 Reactive oxygen species (ROS), 171, 172, 190, 198, 200–202 Restriction fragment length polymorphism (RFLP), 52, 83 Retroviruses, 277 Rhabditida, 78 Rhizoctonia solani, 201 Rhizosphere, 81, 94, 135, 373 RNA viruses, 45, 59, 62 Rphopalosiphum rufiabdominale, 308 Rubber tree mite, 165–166 S Sabouraud Dextrose Agar, 188 Sabouraud Dextrose Broth (SDB), 188 Sabouraud Maltose Agar (SMA), 188 Saccharomyces cerevisiae, 153 Saccharomycotina, 133, 154 Salmonella enterica, 224 Saturniidae, 76 Scelionidae, 263 Schistocerca gregaria, 163, 377 Schizosaccharomyces, 154 Sclerotia, 201 Sclerotinia minor, 201 Sclerotinia sclerotiorum, 201 Sclerotium rolfsii, 201 Scutigerella immaculata, 262 Semiochemicals, 304–305 Septobasidiobasidiaceae, 91

Index Serotype, 78, 81 Serratia spp., 81, 82, 88, 122, 125, 299 S. entomophila, 26, 48, 87, 88, 122, 125, 301 S. marcescens, 48, 81, 87, 122, 224, 227, 301, 308, 321, 323, 329, 330, 332 S. proteamaculans, 87, 122, 125 Sexual spores, 25, 89, 128, 133, 153 Simplicillium lanosoniveum, 189 Sminthuridae, 263 Soil-borne bacteria, 24–36 Soil-borne entomopathogen, 24–36 Solenopsis invicta, 171 Sordariomycetes, 24, 31–34, 91, 133 Sorosporella, 155 Spherulocytes, 213–215, 217–218 Sphingidae, 76, 224 Spinosad, 326, 329, 331, 346, 350, 356, 357 Spirilla, 79 Spodoptera exigua MNPV (SeMNPV), 58, 59, 68 Spodoptera spp. S. exigua, 51, 55, 58, 68, 86, 125, 200, 224, 279 S. frugiperda, 12, 68, 70, 72, 86, 260, 279, 281, 289, 329, 331, 356, 357 S. littoralis, 69, 86, 210, 227, 301, 329, 331 Sporangium, 27 Spore-forming bacteria, 46 Sporulation, 25, 47, 48, 54, 90, 117, 119, 120, 137, 158, 171, 187, 198, 320, 323, 327, 328, 346, 347 ssDNA viruses, 277 Staphylinidae, 254, 263 Steinernema spp., 49, 79, 208–211, 250, 257, 260, 264, 299 S. carpocapsae, 51, 208, 250, 254–257, 259–264, 326, 330, 355–358 S. kraussei, 250, 257, 261, 301, 358 Sugarcane spittlebugs, 167 Superoxide dismutase (SOD), 160, 171, 199, 202, 333, 353 Symbionts, 77–79, 116, 117, 208–212, 220–232, 248–254, 319, 354 Symbiotic bacteria, 48, 86, 209–212, 247–267 Synergy, 173, 301, 308 T Taphrina, 154 Taphrinomycotina, 133, 154 Tebufenozide, 355

395 Tebufenpyrad, 355, 356 Teleomorph, 33, 34, 94, 133, 138, 155, 161 Tenebrio molitor, 202, 311, 328 Tenericutes, 79, 117 Teretriosoma nigrescens, 302 Tetradonematidae, 249 Thanasimus formicarius, 302 Thermotolerance, vii, 172, 186–191, 199 Thiamethoxam, 308–310, 323, 325, 326, 328–330, 333, 347 Thiolutin, 223 Thiomethoxam, 331, 332 Thioredoxins (trx), 199 Thrips, 136, 158, 160, 163, 167, 168, 172, 262, 263, 302, 304, 326, 328, 332, 376, 377 Thysanoplusia orichalcea, 65, 76, 378 Thysanoptera, 33, 51, 249, 252, 253 Tipula paludosa, 62 Togaviruses, 277 Tolypocladium cylindrosporum, 189 Tolypocladium inflatum, 96, 189 Torrubiella, 165 Tortricidae, 76, 261 Trehalase, 157, 158, 170 Trialeurodes spp., 162, 164 Trialeurodes vaporariorum, 162, 311 Tribolium castaneum, 12, 202, 310, 332, 356 Trichogramma chilonis, 253 Trichogrammatidae, 253 Trichoplusia ni, 63, 69, 70, 72, 73, 76, 86, 164, 279, 281, 301, 368, 376 Trifloxystrobin, 347 Triflumizolet, 311 Triterpenoids, 332 Trypsins, 157, 345 Tuber magnatum, 153 Typhlodromalus aripo, 303 Tyrosine, 223 U Ultraviolet (UV) radiation, 169, 171, 172, 189, 198, 248, 266, 286, 345 V Vairimorpha, 49, 299 Varroa destructor, 166 Verticillium spp., 155, 158, 161, 162, 166, 368, 376 V. lacanii, 162 Vibro, 79

396 W White grubs, 128, 138, 167, 168, 323, 331, 333, 355, 358, 368, 376, 378 White truffle, 152–153 Wiseana cervinata, 76 X Xenorhabdus spp., 48, 49, 79, 86, 88, 117, 208–211, 220–226, 228–232, 248–250, 253, 354 X. bovienii, 86, 250 X. nematophila, 86, 88, 211, 222, 224, 225, 227, 229, 231, 250 Xestia c-nigrum, 70, 72, 76, 279, 281 Xylella fastidiosa, 165

Index Y Yellow muscardine, 164 Yersinia, 88, 123, 226, 299 Yersinia enterocolitica, 123, 226, 230 Z Zinc-chelator, 223 Zoophthora, 32, 94, 132, 304, 344, 379 Zoophthora radicans, 304 Zoospore, 91, 128, 130 Zygomycetes, 28, 93, 131 Zygomycota, 24, 94, 131, 154, 344 Zygomycotina, 366