Entomopathogenic Nematodes and Their Symbiotic Bacteria: A Laboratory Manual (Springer Protocols Handbooks) 1071614444, 9781071614440

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S. Sivaramakrishnan M. Razia

Entomopathogenic Nematodes and Their Symbiotic Bacteria A Laboratory Manual

SPRINGER PROTOCOLS HANDBOOKS

For further volumes: http://www.springer.com/series/8623

Springer Protocols Handbooks collects a diverse range of step-by-step laboratory methods and protocols from across the life and biomedical sciences. Each protocol is provided in the Springer Protocol format: readily-reproducible in a step-by-step fashion. Each protocol opens with an introductory overview, a list of the materials and reagents needed to complete the experiment, and is followed by a detailed procedure supported by a helpful notes section offering tips and tricks of the trade as well as troubleshooting advice. With a focus on large comprehensive protocol collections and an international authorship, Springer Protocols Handbooks are a valuable addition to the laboratory.

Entomopathogenic Nematodes and Their Symbiotic Bacteria A Laboratory Manual

S. Sivaramakrishnan Department of Biotechnology, Bharathidasan University, Tiruchirappalli, Tamil Nadu, India

M. Razia Department of Biotechnology, Mother Teresa Women's University, Kodaikanal, Tamil Nadu, India

S. Sivaramakrishnan Department of Biotechnology Bharathidasan University Tiruchirappalli, Tamil Nadu, India

M. Razia Department of Biotechnology Mother Teresa Women’s University Kodaikanal, Tamil Nadu, India

ISSN 1949-2448 ISSN 1949-2456 (electronic) Springer Protocols Handbooks ISBN 978-1-0716-1444-0 ISBN 978-1-0716-1445-7 (eBook) https://doi.org/10.1007/978-1-0716-1445-7 © The Editor(s) (if applicable) and The Author(s), under exclusive licence to Springer Science+Business Media, LLC, part of Springer Nature 2021 This work is subject to copyright. All rights are solely and exclusively licensed by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Springer imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

Foreword The use of chemical pesticides in agriculture raises the concern about their hazardous effects on the environment. There is a need for a change in crop protection methods that are safe to humans and environment. Entomopathogenic nematodes with their symbiotic bacteria play an important role in the field of insect pest management. As these nematodes are eco-friendly, they are used worldwide as biocontrol agents as an effective alternative solution to chemical pesticides. Entomopathogenic nematodes are currently receiving most attention for their bio-efficacy. This book is a complete systematic approach to various entomopathogenic nematode procedures which help in identifying and characterizing them for agricultural pest management. The commitment and the efforts put forth by the authors Dr. S. Sivaramakrishnan, Professor and Head, Department of Biotechnology, Bharathidasan University, Tiruchirappalli, and Dr. M. Razia, Assistant Professor, Department of Biotechnology, Mother Teresa Women’s University, Kodaikanal, in designing this manual is highly appreciated and welcomed. Thoughtfully planned and expertise care is taken to pool all the relevant features of entomopathogenic nematodes and their symbiotic bacteria in a single manual in a wellorganized manner, signifying the uniqueness of this manual. The contents are lucid and noteworthy. The protocols included are clearly defined and stepwise. This book will surely meet the laboratory challenges and will be a valuable resource to the readers. This laboratory manual is an excellent contribution of the authors to the discipline of nematology and will definitely do good to the hands of bio-application.

Bharathidasan University, Tiruchirappalli, Tamil Nadu, India

v

P. Manisankar

Preface Realizing the importance of the entomopathogenic nematodes and their symbiotic bacteria as biocontrol agents, it was our desire and dream to pen our knowledge and research experience and give it a shape in the format of this laboratory manual. This manual focuses on the potential application of an array of laboratory methods and techniques for studying and evaluating entomopathogenic nematodes and their symbiotic bacteria. The available study materials on practical approach for work with the entomopathogenic nematodes are very much scattered. The descriptions provided in other chapters and books are in a broader sense. At times, there is a difficulty to find a set of techniques in a required time in a single book. In some instances, the protocol varies with different scientists. So a necessity was felt to produce a new resource in this area, and this manual was intended to fill the need. In this manual, we provide simple procedures to execute the experiments in an easiest way. This book is a compilation of the basic works necessary for insect pest management using entomopathogenic nematodes research. The important aspects and techniques for EPN and their symbiotic bacteria are compiled into a single book with modifications to suite our laboratory conditions in a nutshell, described in a simplified, clear, and understanding manner. The protocols are adapted from various published protocols. This book concisely deals with host insect rearing, nematode sampling, isolation techniques, characterization including morphological, molecular, and ecological studies, mass production, virulence bioassay, field application, and efficacy. It also includes similar methods and techniques for their associated symbiotic bacteria. This will not only give a comprehensive knowledge to the reader but also provides a practical guideline to those who are seeking to learn and apply the techniques. It is highly expected that this book will definitely meet the requirements of researchers, nematologists, entomologists, microbiologists, and any scientist who are working at insect pest management research laboratories as well as different industries and academic institutions. This book will surely find its wider application for current as well as for future researchers as an easy reference handy material. Tiruchirappalli, Tamil Nadu, India Kodaikanal, Tamil Nadu, India

S. Sivaramakrishnan M. Razia

vii

Acknowledgments It is our immense pleasure to express our respectful deep sense of heartfelt gratitude to Distinguished Professor, Emeritus Dr. Harry K. Kaya for his kind consideration, timely valuable suggestions, review, and feedback, and for his wishes in our endeavor that has helped us to give a form and life to this book; and to him we lovingly dedicate our book. We thank all the nematologists and the scientists for their contributions which served as a basement for our book. Dr. S.S. extends his sincere appreciation to his student, Dr. R. Karthik Raja for his diligence, loving assistance, and cooperation in this project and at all times.

ix

Contents Foreword. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgments. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1 2 3 4 5 6 7 8 9 10 11 12

v vii ix

Safety Guidelines. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Historical Aspects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nematode-Bacterium Symbiosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Biology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nematode Behavior . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Chemical Ecology. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Laboratory Techniques for Entomopathogenic Nematodes . . . . . . . . . . . . . . . . . . Laboratory Techniques for Symbiotic Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Field Application Techniques for Control of Insect Pests . . . . . . . . . . . . . . . . . . . . Application Efficacy Against Insects in Other Than Soil Habitats . . . . . . . . . . . . . Recent Advances and Future Prospect . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1 5 7 15 19 27 31 47 113 145 153 159

References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

163 175

xi

About the Authors S. SIVARAMAKRISHNAN is the Professor and Head, Department of Biotechnology, Bharathidasan University, Tiruchirappalli, India. He is engaged in research and teaching for over 20 years. His specialization is nematology, agricultural biotechnology, biopesticides, and nanotechnological applications. He has won Young Scientists Award four times. He is a reviewer for nearly ten journals. He is a member of Nematological Society of India. He has scientific publications in national and international journals. He has been the principal investigator of major DBT, DST, and UGC projects. He has international collaborations, signed MoU between USA, Korea, Iran and Turkey. He has organized international conferences and national awareness programmes. M. RAZIA is an Assistant Professor in the Department of Biotechnology, Mother Teresa Women’s University, Kodaikanal, India. She has expertise in the areas of nematology, microbial biotechnology, and phytochemistry. She is engaged in teaching and research activities for 15 years. She has published several scientific papers in reputed national and international journals. She has carried out major projects of UGC and DST-SERB. She has organized conferences, seminars, and workshops. She is a life member of Lichen Society of India, The Indian Science Congress Association, The Biotechnology Society of India, and Association of Microbiology India. She has won the Young Scientists Award.

xiii

Chapter 1 Safety Guidelines Abstract Safety guidelines enable a safe and healthy working environment in scientific laboratories during experimentation, observation, or research in a field of study. It is essential to ensure laboratory safety with care towards possible hazards and to protect from physical, chemical and biological injuries. Students, research scholars and technicians must be aware of probable contamination and follow proper safety measures in the laboratory. Keywords Practices, Chemicals, Glassware, Protection, Facilities

Scientific laboratories providing opportunities for experimentation, observation, or research in a field of study can be a hazardous environment to work. Medical or plant parasitic or insect parasitic nematodes and microbes working laboratory is a place where a number of serious scientific safety hazards like physical, chemical, and biological injuries may occur. The main safety concerns in the laboratory are electrical circuits, hazardous chemicals and reagents, heat sources, and infectious biological agents like pathogenic microbes. It is essential to ensure laboratory safety with care toward possible hazards and observation of proper safety procedures. Any lack of knowledge or a moment’s inattention could lead to an injury. Students, research scholars, and technicians must be aware of probable contamination and follow proper safety measures in the individual research laboratory. Safety rules help in safe culture practices of both nematodes and bacteria in the lab. Follow proper safety instructions on handling the equipment and knowing the materials and procedures required to carry out each laboratory analysis.

S. Sivaramakrishnan and M. Razia, Entomopathogenic Nematodes and Their Symbiotic Bacteria, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-1445-7_1, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2021

1

2

1

2

1 Safety Guidelines

General Instruction of Laboratory Practices l

Familiarize yourself with the working place, procedures, careful handling of fire extinguishers, eyewash fountains, safety showers, first aid kit, and other safety apparatus in the laboratory.

l

Know the place of emergency exits and the evacuation routes in the building to use in an emergency.

l

Close doors and windows to keep away from contaminants in the air.

l

Keep your work area tidy and clean.

l

Disinfect the work bench surface with 2% phenol or polysan before and after use.

l

Keep a record of all the experiments and observations.

l

Report all injuries, spills, accidents, and broken equipment or glass to your instructor or laboratory in-charge immediately no matter how small they seem.

l

Never take out any experimental samples or any other objects from the laboratory.

l

Follow the right methods for disposing lab waste.

l

Never raise any glassware, solutions, or other types of equipment over the eye level.

l

Avoid using extension cords as a substitute for permanent wiring.

l

Do not use contact lenses, put on cosmetics, smoke, drink, eat, or store foodstuffs and beverages in the laboratory.

l

Keep all laboratory equipment and other items at their respective places after use.

Safety Measures While . . .

2.1 Handling Instruments

l

Know the working of an instrument prior to handling it independently.

l

Cleanse all the laboratory apparatus regularly whether they are clean or contaminated.

l

Handle hot objects using protective mittens or tongs.

l

Take care while using gas burners and electric hot plates to keep away from severe injuries.

l

Check for electrical equipment with damaged or bare wires or with broken switches or sockets.

Lab Infrastructural Facilities

2.2 Handling Chemicals

2.3 Handling Glassware

3

4

3

l

Treat every chemical as though it is dangerous.

l

Label all chemicals.

l

Replace caps on reagents and solution bottles.

l

Never smell or taste chemicals.

l

Check the glassware each time you use for any chips and cracks.

l

Dispose all broken glasses in a cardboard box or a container designated as a glass receptacle.

Personal Protection Safety Rules l

Always wear lab overcoat/apron while performing laboratory experiments.

l

Always wear appropriate gloves if there are cuts on hands and also while experimenting with hazardous or toxic chemicals.

l

Always put on safety glasses or whichever suitable protecting eye wear while conducting or recording experiments or presentations.

l

Use toe-covered footwear at all times in the laboratory.

l

Wash your hands before and after every experiment thoroughly with soap and water.

l

Do not touch your face, eyes, mouth, neck, and any other body part with your hands while working with laboratory equipment and chemicals.

l

Be careful while transferring liquids since aerosols can develop and they may be harmful to eyes and lungs.

l

Report promptly to your instructor if any contact with human pathogens to limit your exposure to possible infection.

Lab Infrastructural Facilities The basic equipment facilities for nematology work include: l

Laminar flow chamber.

l

Biological Oxygen Demand incubator.

l

Compound microscope.

l

Phase contrast microscope.

l

Dissection microscope.

l

Stereoscopic microscope.

l

Electrophoresis unit.

4

1 Safety Guidelines l

Gel documentation system.

l

Polymer chain reaction machine.

l

Incubator with shaker.

l

Spectrophotometer.

l

Fermentor.

l

Chromatography.

l

SDS-PAGE.

l

Water bath.

l

Centrifuge.

l

Refrigerator.

l

UV light lamp.

l

PCR machine.

l

Vortex machine.

l

pH meter.

Chapter 2 Introduction Abstract Entompathogenic nematodes (EPNs) are created in the nature with significant unique characteristics. The efficacy of EPN application in the field as bio-insecticides depends on its virulence, host finding, environmental tolerance and persistence, and also in the selection of the appropriate nematode that matches with the particular target pest. Understanding the biology, adaptability and identification of these entomopathogenic nematodes with their bacterial symbionts play a key role in their successful application. Keywords entomon, Pathogenic, Model system, Infective juveniles, Application

Entomopathogenic nematodes (EPNs) are created in the nature with significant unique characteristics. The name entomopathogenic is derived from the Greek word entomon, indicating insect, and pathogenic indicates creating disease. As the name implies, EPNs infect only insects. They are an excellent economically important bioagents. It is biologically fascinating to study the highly diverse, complex, and specialized entomopathogenic nematodes. They are obligate or sometimes facultative insect parasites and are microscopic roundworms with non-segmented soft body. They are present in nature in terrestrial habitats. The widely recognized entomopathogenic nematodes are organisms that can be applied in the biological management of insects, the members of Steinernematidae and Heterorhabditidae [1]. These are the only insect-parasitic nematodes with an optimal balance of biological regulation traits [2]. Entomopathogenic nematodes of the genera Steinernema and Heterorhabditis together with their respective symbiotic bacteria Xenorhabdus and Photorhabdus facilitate the infective juveniles (IJs) of nematodes to infect and kill their insect hosts by the host–vector–symbiont interactions. Due to this insect-killing capabilities, EPNs are utilized as a method of biological control agents of economically significant insect pests and a model system to study host–parasite interactions. The metabolites, insecticidal toxins, and

S. Sivaramakrishnan and M. Razia, Entomopathogenic Nematodes and Their Symbiotic Bacteria, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-1445-7_2, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2021

5

6

Introduction

peptides produced by the association of EPNs, their bacterial symbionts, and the infected host play varied roles in EPN ecology, development, and reproduction. The efficacy of EPN application in the field as bio-insecticides depends on its virulence, host finding, environmental tolerance and persistence, and also in the selection of the appropriate nematode that matches with the particular target pest. Understanding the biology, adaptability, and identification of these entomopathogenic nematodes with their bacterial symbionts plays a key role in their successful application. Before using the EPNs for bio-assay, they need to be quantified. This book provides the suitable protocols for its use in the study of EPNs. The protocols described here are compiled from the previous works of published sources and are modified to suit our laboratory conditions.

Chapter 3 Historical Aspects Abstract Entomopathogenic nematodes of the genus Steinernema and Heterorhabditis do not have a select shared ancestry. They independently established relationships with bacteria and insects from disparate, unrelated ancestors. The diversity and distribution of the families Steinernematidae and Heterorhabditidae entomopathogenic nematodes are widespread in every continent except in Antarctica. In 1923, Steiner described the first entomopathogenic nematode. All species of Steinernema have mutualistic associations with Xenorhabdus species and all species of Heterorhabditis have symbiotic associations with Photorhabdus species. S. glaseri was the first mass produced nematode and used under field conditions for biological control of an insect pest. Keywords proto-Rhabditonema, Biogeography, Habitat, Steinernema, Heterorhabditis, Xenorhabdus, Photorhabdus, Systematic

1

Evolution l

The bio-safe eco-user entomopathogenic nematodes Steinernema and Heterorhabditis are one among the ancient species which are considered to be evolved 375 million years ago from a mid-Paleozoic origin.

l

The independent, converging evolution of Steinernema and Heterorhabditis produces alike morphology and life history attributes.

l

Based on the similarity between the male tail morphology and the buccal capsule, it is believed that Steinernema arose from the “proto-Rhabditonema” ancestor in the terrestrial environment and Heterorhabditis developed from the “Pellioditis-like” ancestor in the sandy, aquatic environment [3, 4].

l

Steinernema and Heterorhabditis do not have an exclusive shared ancestry. They independently established relationships with bacteria and insects from disparate, unrelated ancestors [5–8].

S. Sivaramakrishnan and M. Razia, Entomopathogenic Nematodes and Their Symbiotic Bacteria, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-1445-7_3, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2021

7

8

Historical Aspects l

The widespread presence of these wonderful organisms shows that their lineages occurred when all land areas were connected as the Pangaea supercontinent [9].

l

The symbiotic bacteria Xenorhabdus and Photorhabdus are phylogenetically close [7, 10–12] and form tight sister groups.

l

A common ancestor of these bacteria dates back to around 200–500 million years ago and is associated with both Steinernema and Heterorhabditis nematode hosts [13].

l

In 1917: Dr. Krausse in Germany collected the first EPN species.

l

1923: Steiner described the first entomopathogenic nematode Aplectana kraussei (now Steinernema kraussei).

l

1929: Steiner identified the second entomopathogenic nematode Neoaplectana glaseri from material isolated by Glaser and Fox [14].

l

1955: Jaroslav Weiser [15] described Neoaplectana carpocapsae.

l

1955: Dutky and Hough [16] isolated the DD-136 strain of an undescribed steinernematid.

l

1955: Steinernema carpocapsae was obtained from Weiser.

l

1965: Poinar and Thomas described the symbiotic bacterium, Achromobacter nematophilus associated with S. carpocapsae.

l

1966, 1967: Poinar Jr and Thomas [17] explained the role of the bacterium in the growth of the nematode and the host’s death.

l

1967: Poinar showed that the North American DD-136 nematode and the Czechoslovakian strain of S. carpocapsae were of same species.

l

1979: Thomas and Poinar later transferred the bacterium to a new genus, Xenorhabdus.

l

1937: Pereira [18] described the first species of heterorhabditid to be Rhabditis hambletoni, which is currently regarded as a species inquirenda.

l

1976: However, Heterorhabditis was first definitively described by Poinar.

l

1979: Thomas and Poinar characterized the symbiotic bacterium of H. bacteriophora as Xenorhabdus luminescence.

l

1993: Boemare later transferred this bacterial species to the genus Photorhabdus.

l

2011: Proheterorhabditis burmanicus, an old fossil that is 100 million years old, was found in early Cretaceous Burmese amber, and the ancient age of the clade of Heterorhabditis was calculated [19].

Biogeography

9

It is clear that all species of Steinernema have mutualistic associations with Xenorhabdus species and that all species of Heterorhabditis have symbiotic associations with Photorhabdus species [20, 21].

2

l

Early 1930: The biocontrol potential of these nematodes was first investigated by Glaser and his colleagues who investigated S. glaseri [9]. Neoaplectana glaseri, now known as Steinernema glaseri, was the first mass produced nematode and used under field conditions for biological control of an insect pest [22].

l

1959: Dutky first noticed the antibiotic properties of the bacterium associated with S. carpocapsae, which described how the foreign bacteria that penetrated the insect cadaver bearing the growing nematodes could be killed [23].

l

1970: The first entomopathogenic nematodes for field testing was produced by using live insects. Biotrol NCS-DD-136 was produced for experimental purpose using larvae of the wax moth, Galleria mellonella by the Nutrilite Corporation in Lakeview, California.

l

2020: One hundred valid Steinernema species and 21 Heterorhabditis species are identified from different countries of the world [24].

l

2020: Twenty-six species of the genus Xenorhabdus and 19 species of the genus Photorhabdus are currently recognized [13].

l

Entomopathogenic nematodes are widespread in every continent except in Antarctica.

l

Steinernema and Heterorhabditis are found in different countries including India, Pakistan, China, Iran, Thailand, Vietnam, Japan, Korea, Nepal, North America, Central America, South America, Germany, Denmark, France, Spain, Turkey, Russia, Egypt, Kenya, Ethiopia, Tanzania, Benin, Morocco, South Africa, Rwanda, Algeria, Cameroon, Nigeria, Jordan, Oman, United Arab Emirates, Saudi Arabia, Australia, and New Zealand [24].

l

Several new species are identified from Asia.

l

Steinernematids are heterorhabditids.

l

Steinernema carpocapsae, Steinernema feltiae, and Heterorhabditis bacteriophora have the widest geographical distribution (Fig. 1 and Table 1).

Biogeography

2.1 Geographical Distribution

more

diversified

compared

to

10

Historical Aspects

Fig. 1 Geographical distribution of entomopathogenic nematodes 2.2 Habitat Distribution

l

S. affine, S. feltiae, and S. intermedium inhabit mostly the grassland ecosystems [29, 31, 45].

l

S. kraussei occurs in deciduous forests, coniferous forests, and grasslands [28, 46–48].

l

S. carpocapsae was found mainly near the soil surface [49].

l

Certain species of Heterorhabditidae family are widespread in coastal sandy soils [31, 50, 51].

l

H. bacteriophora is widespread in turf and weedy habitats [52, 53].

l

H. indica occurs mainly in calcareous sands [54].

l

Irish type of Heterorhabditis confined to the coastal areas of Ireland and Britain are found in Central and Northern Europe grasslands [55].

Biogeography

11

Table 1 Geographical distribution of some EPN species EPN species

Country

References

Steinernema kraussei

Germany

[25, 26]

Czech Republic

[27]

Switzerland

[28]

The Netherlands

[29]

United Kingdom Spain

[30]

North America

[31]

Steinernema feltiae

Denmark

[32, 33]

Steinernema affine

Denmark

[33, 34]

Steinernema intermedium

USA

[35, 36]

Steinernema longicaudum

China

[37]

Korea Western USA Steinernema bertusi

South Africa

[38]

Heterorhabditis bacteriophora

USA

[39]

Australia East Asia (China, Japan, Korea) Central and Southern Europe Heterorhabditis zealandica

New Zealand

[40]

Heterorhabditis indica

Southern India

[41]

Sri Lanka Indonesia North Australia West Malaysia Caribbean region Egypt Kenya Japan Heterorhabditis marelatus

California

[42, 43]

Oregon USA Heterorhabditis noenieputensis

South Africa

[44]

12

3 3.1

3.2

Historical Aspects

Systematic Position Steinernema

Phylum:

Nematoda Rudolphi, 1808

Class:

Secernentea von Linstow, 1905

Order:

¨ rley, 1880) Chitwood, 1933 [56, 57] Rhabditida (O

Suborder:

¨ rley, 1880) Chitwood, 1933 [56, 57] Rhabditina (O

Super family:

Strongyloidoidea Travassos, 1920 [58]

Family:

Steinernematidae Chitwood and Chitwood, 1937 [59]

Type genus:

Steinernema Travassos, 1927 [60]

Syn.:

Neoplectana Steiner, 1929 [61]

Type species:

Steinernema kraussei (Steiner, 1923) Travassos, 1927 [25, 60]

Syn.:

Aplectana kraussei (Steiner, 1923) Travassos, 1927 [25, 60]

Other species:

99 valid species

Other genus:

Neosteinernema Nguyen and Smart, 1994 [62]

Type and only species:

Neosteinernema longicurvicauda Nguyen and Smart, 1994 [62]

Heterorhabditis

Phylum:

Nematoda Rudolphi, 1808

Class:

Secernentea von Linstow, 1905

Order:

¨ rley, 1880) Chitwood, 1933 [56, 57] Rhabditida (O

Suborder:

¨ rley, 1880) Chitwood, 1933 [56, 57] Rhabditina (O

Super family:

¨ rley, 1880) Travassos, 1920 [57, 58] Rhabditoidea (O

Family:

Heterorhabditidae Poinar Jr, 1976 [39]

Type and only genus:

Heterorhabditis Poinar Jr, 1976 [39]

Syn.:

Chromonema Khan, Brooks and Hirschmann, 1976 [63]

Type species:

Heterorhabditis bacteriophora Poinar Jr, 1976 [39]

Syn.:

Chromonema heliothidis Khan et al., 1976 [64] (continued)

Systematic Position

13

Heterorhabditis heliothidis (Khan et al., 1976; Poinar et al., 1977) [64] Other species:

3.3

3.4

20 valid species

Xenorhabdus

Domain:

Bacteria

Phylum:

Proteobacteria

Class:

Gammaproteobacteria

Order:

Enterobacterales

Family:

Morganellaceae

Genus:

Xenorhabdus (Thomas and Poinar Jr, 1979) [65]

Type species:

Xenorhabdus nematophila

Other species:

25 species

Photorhabdus

Domain:

Bacteria

Phylum:

Proteobacteria

Class:

Gammaproteobacteria

Order:

Enterobacterales

Family:

Morganellaceae

Genus:

Photorhabdus (Boemare et al., 1993) [66]

Type species:

Photorhabdus luminescens

Other species:

18 species

Chapter 4 Nematode-Bacterium Symbiosis Abstract Entomopathogenic nematodes of the families Steinernematidae and Heterorhabditidae are lethal insect endoparasites signified by their mutualism with symbiotic bacteria in the genera Xenorhabdus and Photorhabdus, respectively. This association is an obligate mutualism in nature with each partner wanting the other to complete its life cycle. Steinernema species form symbiosis with only one Xenorhabdus species, whereas Photorhabdus species form symbiosis with many Heterorhabditis species. Steinernema carry their symbiotic bacteria in a specialized vesicle called the receptacle. Heterorhabditis use their intestinal lumen to harbor its symbiotic bacteria. These bacteria produce anti-immune proteins to help the nematode in evading host defenses, and antimicrobials to subdue competitors. Keywords Obligate mutualism, Gut, Intestinal lumen, Symbiosis, Nematode-bacterium complex

l

Entomopathogenic nematodes of the families Steinernematidae and Heterorhabditidae are lethal insect endoparasites signified by their mutualism with symbiotic bacteria in the genera Xenorhabdus and Photorhabdus, respectively.

l

This association is an obligate mutualism in nature with each partner wanting the other to complete its life cycle.

l

There are no evidence for the isolation of Xenorhabdus and Photorhabdus from soil assuming that these bacteria cannot exist in the soil environment in the absence of their nematode associates.

l

The genus Steinernema carry symbiotic bacteria in a specialized vesicle called the receptacle, which is located in the anterior part of the gut [67].

l

Heterorhabditis do not have such a specialized structure and use their intestinal lumen to harbor its symbiotic bacteria in the foregut and midgut [68].

l

The nematode infective juvenile carries between 0 and 2000 cells of its symbiont bacterium in the anterior part of the intestine [69, 70].

S. Sivaramakrishnan and M. Razia, Entomopathogenic Nematodes and Their Symbiotic Bacteria, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-1445-7_4, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2021

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1

Nematode-Bacterium Symbiosis l

Nematodes assist the bacteria to find suitable hosts and invade them.

l

The development and multiplication of the nematode depends on the conditions created by the bacterium in the insect host.

l

The bacteria further produce anti-immune proteins to help the nematode in evading host defenses, and antimicrobials to subdue competitors [71].

l

The nematode–bacterium complex is a significant natural enemy of soil insects and holds an important part as biological insecticides [72].

Steinernema and Their Xenorhabdus Symbiont Each Steinernema species forms symbiosis with only one Xenorhabdus species, whereas Xenorhabdus species associates with many nematode species (Table 1).

Table 1 Steinernema species and their symbiont Steinernema nematode

Xenorhabdus symbiont

References

S. carpocapsae

X. nematophila

[65, 73]

S. feltiae

X. bovienii

[66, 74]

S. intermedium S. kraussei S. weiseri S. silvaticum S. sichuanense S. nguyeni S. poinari S. tbilisiensis S. jollieti S. puntauvense S. oregeonense S. litorale S. glaseri

X. poinarii

S. cubanum S. longicaudum

X. beddingii

[75, 76]

S. kushidai

X. japonica

[77] (continued)

Heterorhabditis and Their Photorhabdus Symbiont

17

Table 1 (continued) Steinernema nematode

Xenorhabdus symbiont

References

S. bicornutum

X. budapestensis

[78]

S. ceratophorum S. scapterisci

X. innexi

S. rarum

X. szentirmaii

S. serratum

X. ehlersii

S. riobrave

X. cabanillasii

S. diaprepesi

X. doucetiae

S. hermaphroditum

X. griffiniae

S. karii

X. hominickii

[79]

S. monticolum S. scarabaei

X. koppenhoeferi

S. arenarium

X. kozodoii

S. apuliae S. puertoricense

X. romanii

S. siamkayai

X. stockiae

S. abbasi

X. indica

[79, 80]

S. australe

X. magdalenensis

[81]

S. aciari

X. ishibashii

[82]

S. khoisanae

X. khoisanae

[83]

S. eapokense

X. eapokensis

[84]

S. sangi

X. thuongxuanensis

S. yirgalemense

S. jeffreyense S. saccharii

2

Heterorhabditis and Their Photorhabdus Symbiont In Heterorhabditis–Photorhabdus symbiosis, several species of both nematodes and bacteria can involve in symbiotic associations with multiple species of symbiotic partners (Table 2).

18

Nematode-Bacterium Symbiosis

Table 2 Heterorhabditis species and their symbiont Heterorhabditis nematode

Photorhabdus symbiont

References

H. bacteriophora Brecon

P. luminescens

[65, 85, 86]

H. indica

P. akhurstii

[85, 87]

H. bacteriophora

P. laumondii

[85]

P. kayaii

[88]

H. indica

P. thracensis H. zealandica

P. temperata

[85]

P. australis

[87, 89]

P. cinerea

[87, 90]

H. bacteriophora

P. caribbeanensis

[12, 87]

H. zealandica

P. tasmanensis

H. bacteriophora NC1 H. megidis H. gerrardi H. indica H. downesi H. megidis H. bacteriophora

H. marelatus H. georgiana

P. kleinii

H. bacteriophora

P. stackebrandtii

H. sonorensis

P. sonorensis

[92]

H. beicherriana

P. bodei

[87]

H. mexicana

P. mexicana

[86]

H. atacamensis

P. guanajuatensis

[87, 91]

Chapter 5 Biology Abstract The life cycle of the entomopathogenic nematodes Steinernema and Heterorhabditis consists of two phases namely a free-living phase in the soil and a parasitic phase inside the insect host. Steinernema IJs use ambusher strategy and Heterorhabditis IJs use cruiser strategy to locate their insect host. IJs enter the insect host through natural openings like mouth, anus and spiracles or cuticle. The metabolites produced by bacteria are lethal to the larvae and the host insect is killed by septicemia or toxemia. The bacterial metabolites have a wide-spectrum of antimicrobial activity. Biocontrol potential of EPNs depends on the mutualism amongst a host-seeking nematode and a lethal insect-pathogenic bacterium. Keywords Life cycle, Infective juveniles, Host, Septicemia, Metabolites, Biocontrol

l

Three special features of Steinernema and Heterorhabditis nematodes present them as an amazing study platform in biocontrol purposes.

l

First, they form a complex nematode–bacterium mutualistic symbiosis. The nematodes carry the bacteria in their body and release them into the insect hosts [40].

l

Second, they are insect pathogens with a very wide host range that covers most of the insect orders.

l

Third, they can be mass produced either in vivo or in vitro.

l

Although the two nematode groups show similarity in infecting, killing, and emerging as a new generation from insect hosts, they exhibit difference in their life cycles [93].

l

The life process of the nematode–bacteria complex explains the mode of action of these potent biological insecticides.

S. Sivaramakrishnan and M. Razia, Entomopathogenic Nematodes and Their Symbiotic Bacteria, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-1445-7_5, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2021

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Biology

Life Cycle l

The life cycle of the entomopathogenic nematodes Steinernema and Heterorhabditis consists of two important phases: a freeliving phase in the soil and a parasitic phase inside the insect host.

l

The life cycle takes place 10–15 cm below the soil.

l

The life cycle consists of the stages of the egg, juvenile 1, 2, 3, and 4 (J1, J2, J3, and J4), and adult.

l

The third stage (J3) infective juvenile (IJ) or dauer juvenile (DJ) is the only free living non-feeding stage of the nematode outside of their insect host.

l

IJ with its closed mouth and anus is sheathed in a two-layered cuticle and can persist for months in the soils in the absence of a suitable host.

l

During favorable environmental conditions, IJs can search for susceptible hosts in the environment. Steinernematidae IJs use ambusher strategy and Heterorhabditidae IJs use cruiser strategy to locate their insect host.

l

Once the insect is detected, the IJ expels its external cuticle to expose the mouth and anus, and the parasitic phase starts when IJ enters the insect host through natural openings like mouth, anus, and spiracles or cuticle.

l

Once inside the host, the nematodes invade along the intestinal wall into the insect hemocoel or blood cavity and release their symbiotic bacteria.

l

The bacteria produce secondary metabolites that suppress the immune function of the insect larvae.

l

The metabolites are also lethal to the larvae and the host insect is killed by septicemia or toxemia within 24–48 h [94] after infection.

l

The bacteria rapidly proliferate, breaking down the host tissues, and convert the insect cadaver into an effective growth base for the nematode. The bacteria and the degrading tissues of the host cadaver serve as food sources for nematode growth and reproduction [21].

l

Owing to the association of various symbiotic bacteria with EPN, steinernematid nematodes turn the insect cadaver tan, ochre, gray, or dark gray, whereas heterorhabditid nematodes turn the host cadaver red, purple, orange, yellow, brown, or sometimes green.

l

In the process, the bacteria create a secured niche for the nematode and themselves by developing antibiotics with bactericidal, fungicidal, and nematicidal properties that inhibit other microorganism competitors [95].

Life Cycle

21

l

The IJ leaves the pathogenic phase in a growth stage that is known as recovery and changes into the fourth stage (J4) and then the first-generation adults.

l

J4 stage nematodes mature into egg laying female or male adults in the insect cadaver and with this undergo four juvenile stages (J1–J4) and the adult stage has up to three generations [1].

l

Steinernematid IJs develop into males or females in all successive generations, while heterorhabditids become self-fertilizing hermaphrodite females in the first generation, and in subsequent generations develop into amphimictic males and females or into automictic hermaphrodite, or both can happen concurrently [96].

l

Therefore, in general, the two nematodes steinernematids and heterorhabditids differ slightly in their reproduction in the host insect. If it is Steinernema, it needs two individuals, one male and one female, to be present in the insect host before they can reproduce, whereas for a Heterorhabditis, a single IJ is enough to invade a host insect to reproduce and develop [97].

l

The progeny of the first-generation adults evolves either through all the four juvenile stages to the second-generation adults or into third-stage IJ, depending primarily on the availability of nutrition and density of population.

l

Several hundred thousands of new generation IJs are developed depending on the size of the insect prey in around 2 weeks from a single insect cadaver.

l

After two to three reproduction cycles and depletion of all the nutrition in the cadaver, an increased population density of nematodes triggers their development into IJs again.

l

The infective juvenile nematodes re-associate with bacterial symbionts and reinitiate the symbiosis.

l

Finally, the IJs emerge from the empty insect carcass into the soil environment from where they can locate and invade the next insect hosts.

l

The life cycle of EPNs from infective juvenile penetration to infective juvenile emergence is completed within 6–11 days for Steinernematids and 12–14 days for Heterorhabditids [98].

l

Growth of nematode under optimum conditions (soil temperatures 77–82 ºF) takes about 3–7 days for one life cycle from egg to egg inside an insect host (Fig. 1) [99].

22

2

Biology

Bacterial Metabolites l

Over 30 bioactive secondary metabolites from different chemical groups are produced by the symbiotic bacteria Xenorhabdus and Photorhabdus.

l

The metabolites from Xenorhabdus species are more complex than that of Photorhabdus.

l

Several Xenorhabdus and Photorhabdus species produce more than one group of bioactive secondary metabolites.

l

The bacterial metabolites have a wide spectrum of antimicrobial activity against bacterial and fungal species including human pathogenic bacteria, fungi and yeasts, and multidrug resistant [100–102] and are anti-cancerous [103].

Fig. 1 (a) Life cycle of entomopathogenic nematodes and their symbiotic bacteria. (b) Penetration, infection and reproduction of entomopathogenic nematodes inside termite

Bacterial Metabolites

23

Fig. 1 (continued) l

The mechanism involved in biosynthesis and regulation of possible metabolites having insecticidal properties is an essential role in nematode–bacteria [104].

l

They possess a broad spectrum of bioactivities of agricultural and medicinal interest, like antibiotic, antimycotic, antiviral, insecticidal, nematicidal, antiulcer, and antineoplastic.

l

The antibiotic action is effective toward microorganisms of the Gram-positive Micrococcus, Staphylococcus, and Bacillus; the Gram-negative Escherichia, Shigella, Enterobacter, Serratia, Proteus, Erwinia, Flavobacterium, and Pseudomonas; and the yeasts Candida and Saccharomyces [105].

l

Antimycotic activity is effective in a number of fungi including Botrytis cinerea, Fusarium oxysporum, Fusarium solani, Mucor piriformis, Pythium coloratum, Pythium ultimum, Penicillium spp., Rhizoctonia solani, Trichoderma pseudokoningii, and Verticillium dahliae.

l

Xenocoumacin doses given orally to rats show potent activity toward stress-induced ulcers [102].

l

The compound cytotoxicity [106].

protein

CyA

is

responsible

for

24

Biology l

3

The genomic DNA containing several genes is used in the synthesis of insecticidal and antimicrobial bacterial metabolites (Table 1) [107, 108].

Biocontrol Potential l

EPNs are possible biocontrol agents as they function as bacterial vectors, rapidly destroy target insect pests, have a wide variety of hosts, are extremely virulent, contain chemoreceptors, can be easily grown in vitro, are healthy for vertebrates, plants, and non-targets, have been excluded from registration, are easily applied using regular application tools, are compatible with many chemical insecticides, and are suitable for genetic selection.

l

Currently, hundreds of researchers from more than 40 countries are focusing on EPNs as biocontrol agents, and their numbers are growing.

l

This strong curiosity relies on the amazing characteristics of these beneficial nematodes, which include ease of mass processing, ease of application, host specificity, high lethality, and protection for non-target species [1, 123].

l

Biocontrol potential of EPNs depends on the mutualism among a host-seeking nematode and a lethal insect-pathogenic bacterium.

l

The key element for their successful role in pest management is the potential of EPN IJs to scatter effectively in the soil and find a host [124].

l

Since the first use of EPNs (S. glaseri) for controlling beetles of the species Popillia japonica in USA [125], so far, no reports of environmental harms by these bioagents have been documented.

l

These nematode biopesticides are widely used globally to manage main agricultural insect pests including chafers, weevils, fungus gnats, various caterpillars, and mole crickets.

Biocontrol Potential

25

Table 1 Bioactive metabolites of EPN-symbiotic bacteria Species

Bacterial metabolites

Bioactivities

References

Xenorhabdus beddingii R-type bacteriocins

Antibiotic

[75, 76, 109]

X. budapestensis

Antibiotic

[110–113]

Antibiotic Antimycotic Insecticidal Antiulcer Anticancer

[114, 115]

Fabclavine Bicornitun

X. doucetiae

Xenoamicin Xenocoumacin Xenorhabdin

X. kozodoii

Xenocoumacin

Antibiotic Antimycotic Antiulcer

[113, 116]

X. bovienii

Xenorhabdin

Antibiotic Antimycotic Insecticidal Anticancer

[102, 107, 117, 118]

Antibiotic Antimycotic Antiulcer

[101, 102, 119]

Indoles

Antibiotic Antimycotic Antiviral

[100]

Isoflavonoids

Antibiotic

Dithiolopyrrolone

Cytotoxic

[103]

Photorhabdus luminescens

3,5-Dihydroxy-4isopropylstilbene

Antifungal Nematicidal

[120, 121]

Photorhabdus spp.

Siderophore photobactin

Antibacterial

[122]

Carbapenem

Antibiotic

Hydroxystilbene

Antibiotic Antimycotic Nematicidal

Anthraquinones

AntibioticAntimycotic Anticancer Antiviral

Nucleosides

Antibiotic Antimycotic

Amicoumacin Xenomin Xenorxides Xenorhabdin Xenematide X. nematophilus

Xenocoumacins Nematophin

Xenorhabdus spp.

(continued)

26

Biology

Table 1 (continued) Species

Bacterial metabolites

Bioactivities Anticancer Antiviral

Macrolides

Antibiotic

References

Chapter 6 Nematode Behavior Abstract Entomopathogenic nematodes (EPNs) use foraging strategies to find their suitable insect hosts. They respond to insect or plant host-associated volatile and chemical cues including CO2 through chemosensation. They depend on chemical signals in their surroundings to locate food sources, suitable hosts, and reproductive partners. The intensity of the nematode’s behavior towards chemical stimuli in its natural environment is greatly dependent on the diffusion rate of the chemical compound and on the structural heterogeneity of soil. Foraging strategies such as ambush, standing, jumping, cruise, crawling behavior and intermediate strategies may differ from species to species. Foraging assays are carried out to study the nematode behavior. Keywords Foraging strategies, Chemosensation, Ambushers, Cruisers, Foraging assays

1

Host-Seeking Behavior l

Host-finding behavior of entomopathogenic nematodes helps to identify which species can be used in the biological pest management programs.

l

EPNs use foraging strategies to find their insect hosts.

l

They respond to host-associated volatile cues and chemical cues including CO2 through chemosensation.

l

Chemosensation is a means of sensory function which the nematodes use to direct themselves toward their insect hosts.

l

They depend on chemical signals in their environment to locate food sources, suitable hosts, harmful compounds, reproductive partners, and sometimes, allow them to choose between alternative developmental stages [126].

l

The intensity of the nematode’s behavior toward chemical stimuli in its natural environment is greatly dependent on the diffusion rate of the chemical compound and on the structural heterogeneity of soil [127].

l

When a foraging nematode is presented with a range of signals arising from similar general region, the behavior can rely on the

S. Sivaramakrishnan and M. Razia, Entomopathogenic Nematodes and Their Symbiotic Bacteria, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-1445-7_6, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2021

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Nematode Behavior

intensity and time of exposure and the nature of the stimulus [128]. l

2

Foraging strategies differ from species to species.

Foraging Strategies

2.1 Ambush Foraging Strategy

Ambushing EPNs remain at one place or close to the soil surface and detect host insects by direct contact. They are attracted toward CO2 and host-specific odorants, whereas volatiles find no major role in host-finding. Ambushers successfully manage insect pests which are extremely mobile at the surface of the soil like cutworms, armyworms, and mole crickets. Ambusher EPN species include S. carpocapsae and S. scapterisici.

2.1.1 Standing Behavior

An infective juvenile of ambusher attaches to the mobile hosts at the soil surface by raising the anterior portion of its body and waves it back and forth and balances on its tail, referred to as “standing” [129], “nictation” [130], and “winken” (Fig. 1) [131].

2.1.2 Jumping Behavior

IJs of ambushers also exhibit jumping behavior, where the IJ stands on its tail, curls, and propels itself into the air to attach to the hosts [132].

2.2 Cruise Foraging Strategy

Cruising EPNs actively move about through the soil to find their host by detecting carbon dioxide or volatiles emitted by the host. Cruisers efficiently monitor sedentary and slow-moving insect pests at different soil depths, such as white grubs and root weevils. Cruiser EPN species include H. bacteriophora and S. glaseri.

2.2.1 Crawling Behavior

IJs of cruisers crawl by sinusoidal movement on the substrate using the surface tension forces to propel them forward or backward to locate their hosts [133].

2.3 Intermediate Foraging Strategy

Intermediating EPNs use both ambushing and cruising strategies to locate their host. These foragers are less successful when compared to ambushers and cruisers at parasitizing hosts either on the soil surface or deep in the soil profile. Intermediate EPN species include S. feltiae and S. riobrave.

3

Foraging Assays

3.1 Standing and Jumping Behavior

1. Sprinkle 0.5 g of sand particles on a 9-cm petri dish surface, lined with double-moisten filter paper. 2. Apply 100 IJs on the filter paper surface.

Foraging Assays

29

Fig. 1 Nictating infective juvenile of S. siamkayai

3. Observe the behavior of the IJs under a dissecting microscope with 45 magnification. 3.2 Attachment Rate [134]

Nematode foraging strategies are assessed by the use of a mobile host (G. mellonella) compared to a sit-and-wait forager (ambusher) and an active forager (cruisers). 1. Sprinkle 0.5 g of sand on a petri dish lined with moist filter paper. 2. Add 1000 IJs/200 μl and allow for 15 min for the dispersal of nematode and add one Galleria larva. 3. Using a probe, the larva should crawl continuously. 4. Remove the larva after 10 min and wash with distilled water. 5. Using a dissecting microscope, count the number of IJs.

3.3 Effect of Substrate [134]

1. Fill 2 g of moist sand in a petri dish. 2. Add 50 IJs in 60 μl of water followed by a single G. mellonella larva. 3. Dissect the dead larvae after 3 days and then digest in pepsin solution. 4. Count the nematodes.

3.4 Host Finding [134]

The potential of the IJs to detect and infect a sedentary host at various soil depths is analyzed. 1. Fill a 15-cm plastic container with sandy loam soil up to 10 cm.

30

Nematode Behavior

2. Place G. mellonella in aluminium mesh cages to inhibit its crawling. 3. Keep the cages on the surface of the soil at 2, 6, or 10 cm depth. 4. Add IJ suspension of 100 IJs/1 ml on each column surface. 5. Remove the columns after 3 days. 6. Dissect the dead larvae under a microscope and record the number of nematodes.

Chapter 7 Chemical Ecology Abstract The third stage of Infective juveniles (IJs) habitually traces suitable hosts when the insects feed, through chemical cues produced from plant roots damaged by herbivores. EPNs are attracted to CO2 and volatile substances emitted by soil insects and plant roots. Besides, other chemical mixtures, belonging to diverse chemical categories persuade chemotaxis in nematodes. Multitrophic interactions that use and exchange this chemical information play key role in protection of plant and predator-prey mechanics in the chemical ecology of EPNs. Various protocols are examined to study the complexity of adaptations in a chemical ecology of a chemically mediated tritrophic interactions among plants, insects and nematodes. Keywords Chemotaxis, CO2, Sensory neurons, Chemical attractant, Volatile cues, Semiochemical, Olfactometer

1

l

Infective juveniles of entomopathogenic nematodes habitually trace suitable hosts when the insects feed, through chemical cues produced from plant roots damaged by herbivores [135, 136].

l

The unique tripartite complex of EPN-symbiont-host forms pheromones, compounds which are insecticidal in nature, antipathogenic compounds, and scavenging deterrents that are important for the EPNs [121, 137, 138].

l

Such metabolites offer abundance of chemical material to other habitants in the ecological community such as insect prey of nematodes and other plants.

l

Multitrophic interactions that use and exchange these chemical information play key role in protection of plant and predator–prey mechanics in the chemical ecology of EPNs.

Chemical Cues Various EPNs are attracted to CO2 emitted by soil insects and plant roots. CO2 acts in a combined manner with root volatiles to appeal EPNs [139]. Besides CO2, other chemical mixtures, belonging to

S. Sivaramakrishnan and M. Razia, Entomopathogenic Nematodes and Their Symbiotic Bacteria, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-1445-7_7, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2021

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Chemical Ecology

diverse chemical categories, persuade chemotaxis in nematodes. Insects are generally infected by EPNs as they are capable of locating CO2, insect odorants, and plant odorants [140]. Numerous sulfur-containing compounds, like dimethyl sulfide, dimethyl trisulfide, methanethiol, and glucosinolate catabolic substances like thiocyanates and isothiocyanates, are discharged from the roots responding to insect incursion. Organic acids, proteins, saccharides, amino acids, lipids, coumarins, flavonoids, enzymes, and aliphatic and aromatic substances are samples of the chief compounds present in the root rhizosphere. The sensory neurons, otherwise called as the BAG neurons, that are located in the head can control free-living and parasitic nematodes’ reactions for CO2 (Table 1) [141].

2

Tritrophic Interactions [148]

2.1 Materials Required

2.2 Plants, Insects, and Nematodes 2.2.1 Procedure

Diverse ecological interactions thrive on chemical cues. This assay helps to study the complexity of adaptations in a chemical ecology of a chemically mediated tritrophic interactions among plants, insects, and nematodes. l Plant potato. l

EPN‐infected cadavers.

l

EPNs.

l

EPN cues. 1. Expose roots of plants (potato or any plant) to living EPNs, EPN chemical cues, or apt controls. 2. To identify the direct responses of plant on EPNs exposure. (a) Add 90 ml of water containing 35,000 IJs into soil to each potted plant and 90 ml of water to control pot plants. (b) Note: Add 90 ml per day, for 2 days, this allows the soil to retain the required water. 3. To identify the direct plant responses on EPN chemical cues exposure. (a) Expose the roots of the plants to EPN chemical cues limiting physical contact to EPNs. 4. Expose the headspace of the infected carcass by EPN (G. mellonella with nematodes) and roots of plants to filtered clean air. 5. Transplant the plants prior to 2 weeks of the experiment into chambers made of clean glass that are filled with peat‐based potting mix.

Tritrophic Interactions

33

Table 1 Attractant compounds of entomopathogenic nematodes Nematode species

Chemical attractant

References

Steinernema carpocapsae

Carbon dioxide

[141, 142]

2-Nonanone 4,5-Dimethylthiazole Octyl acetate Heptanol Hexanol Nonanol Octanol Pentanol Methyl salicylate Undecyl acetate 6-Methyl-5-hepten-2-one

[143]

ß-Caryophyllene Linalool α-Pinene Bornyl acetate Borneol 2,4-Di-tetra-butylphenol 2-Ethyl-hexanol Terpinolene S. glaseri

Carbon dioxide

[144]

S. diaprepsi

α-Santalene

[145]

S. riobrave

α-Santalene

[145]

S. feltiae

α-Santalene

[145]

ß-Caryophyllene

[143]

α-Pinene Bornyl acetate Borneol 2,4-Di-tetra-butylphenol 2-Ethyl-hexanol Terpinolene (continued)

34

Chemical Ecology

Table 1 (continued) Nematode species

Chemical attractant

S. kraussai

Decanal

References

Undecane ß-Caryophyllene α-Pinene Bornyl acetate Borneol 2,4-Di-tetra-butylphenol 2-Ethyl-hexanol Terpinolene Heterorhabditis bacteriophora

Carbon dioxide

[146]

1-Heptanol 1-Hexanol 1-Nonanol 1-Octanol 1-Pentanol 2-Acetylthiazole 2-Heptanol 2-Isobutylthiazole 2-Methylpyrazine 2-Nonanol 3-Nonanol Benzothiazole Caproic acid Caprylic acid Methylvaleric acid 4,5-Dimethylthiazole

[141, 146]

Methyl salicylate

[141]

p-Cymene Propanol Undecyl acetate (E)-β-Caryophyllene

[136] (continued)

Tritrophic Interactions

35

Table 1 (continued) Nematode species

Chemical attractant

References

Nonanal

[143]

Octanal Decanal ß-Caryophyllene α-Pinene Bornyl acetate Borneol 2,4-Di-tetra-butylphenol 2-Ethyl-hexanol Terpinolene H. megidis

(E)-β-Farnesene

[147]

(E)-Nerolidol

H. indica

ß-Caryophyllene

[143]

Geijerene

[145]

Pregeijerene

6. Add to the EPN exposure, treatment chambers that contain autoclaved dampened sand, the EPN‐infected cadavers. Interconnect these chambers with plant chambers using a sand-filled glass arm. 7. Expose plants in the control to headspace chemical compounds either from G. mellonella in sand that is thawed and freeze‐ killed (control carcass), or only sand. 8. Separate roots of plants in every treated chambers from the exposure‐source chambers using 400 mesh film having a distance of 28 cm. 9. Expose the plants to the several treatments for 48 h earlier to obtain any measurements and uninterruptedly throughout the period of the sampling process. 2.3 Collection and Assaying of EPN Volatiles

1. Collect volatile cues discharged by cadavers infected with EPN by dynamic headspace sampling. 2. Analyze through gas chromatography coupled to mass spectrometry (GC/MS).

36

Chemical Ecology

2.4 Larval Performance Analysis

A no‐choice feeding bioassay can be directed for ascertaining the impact of plant disclosure toward EPN cues on performance of the herbivore. 1. Relate the larval mass increase and foliage feeding in exposed plants to cues from EPN carcass, control carcass, or empty sand controls. 2. Cage five neonates on undamaged foliage succeeding the initial exposure. 3. Examine the larvae each day and if the larvae had fed on more than half of the leaf tissue in the cage, then transfer to fresh leaves. 4. Take out larvae after 5 days and weigh. 5. Scan the leaves of individual plant and calculate the total leaf area fed by larvae with Adobe Photoshop software.

2.5 Insect Herbivore Oviposition Preference Analysis

To assess the effect of EPN chemical cues on herbivore oviposition, a three‐way choice test is piloted with female insect. 1. Place a single insect in a cage consisting of three separately planted potato plants. 2. Use soil as control, and place a pot with three EPN‐infected cadavers (G. mellonella with IJs) and another pot with three control cadavers (G. mellonella). 3. Place female insect in the middle of the pot and allow to oviposit for 3 days. 4. Count the total number of eggs in each treatment.

3

Volatile Cues

3.1 Detection of Volatiles Emitted by Insect Larval Hosts

Entomopathogenic nematodes are attracted to volatile substances released by insect-damaged plants. These volatile compounds work as a kind of chemical signals (semiochemical). By appealing EPNs to an affected plant, the volatile compound provides indirect fortification to the plant. The exposure period of an EPN to volatile compounds is of crucial significance for recognizing chemical stimuli. EPN reaction to diverse volatile cues is a species-specific feature [143]. To examine the influence of odors on host-locating behaviors, use thermal desorption-gas chromatography-mass spectroscopy (TD-GC-MS) to recognize insect emanated odorants.

3.1.1 Materials Required

l

Flask.

l

Last-instar larvae of Galleria mellonella / Tenebrio molitor.

l

Thermal desorption tube.

Volatile Cues

3.1.2 Procedure

l

Gas chromatography (GC).

l

HP 6890 GC-5973 MS system.

l

Eclipse 4660 purge.

l

Trap sample concentrator.

l

HP-624 capillary column.

l

Quadrupole mass spectrometer.

37

1. Take 125-ml flask. 2. Place last-instar larvae (Galleria mellonella and Tenebrio molitor) within the flask. 3. Flow into the flask a current of air (10% oxygen, 90% nitrogen) and out through a thermal desorption tube at nearly 104 ml/ min flow rate. Note: Experiments are carried out in sets, with a vacant control flask being run every time. 4. Transfer the substances from the thermal desorption tube to the HP 6890 GC-5973 MS system with an Eclipse 4660 purge and trap sample fitted with an air tube desorber accessory. 5. Desorb the tubes at 200
C for 15 min and transfer through a helium flow to an inner trap detained at room temperature. 6. Heat the inner trap to 200
C subsequent to desorption. 7. Bring this trap in line through the GC carrier gas flow as the trap reaches 180
C. 8. Take the trap offline and run it through a bake out process. 9. Flow the sample to a GC through a transfer line maintained at 120
C, and it moves in a split-splitless injector apprehended at 200
C. 10. Operate the injector in split mode of split ratio at 30:1, later install a 1 mm liner to augment chromatographic resolution. 11. Attain separation with HP-624 capillary column (30 m  0.320 mm) in which a volumetric flow of 1 ml/min is upheld by electronic pressure control. 12. Clutch the transfer line to the mass spectrometer at 200
C, the ion source at 250
C, and the quadrupole at 100
C. The mass spectrometer is fitted by an electron impact source. 13. Establish electron energy to 70 eV in order to acquire the best conceivable library spectrum matches. 14. Operate the quadrupole mass spectrometer with full width at 0.65 m/z half maximum. 15. Authenticate mass calibration each week. 16. Ramp the GC oven from 30 to 260
C and run for 42 min.

38

Chemical Ecology

17. Analyze the data with Chemstation as well as MassHunter software. 18. Search mass spectra against the Wiley library (275,000 spectra) of electron impact mass spectra. 19. Substances recognized are certainly established by running the pure substance and relating the retention time and mass spectra of the assay-identified substance to the known substance. 20. In situations at which the retaining time is off by 0.5 min or the mass spectra do not match, the assay-identified substance is regarded unknown and never used in behavioral assessments.

4

Olfactometer Assay [139]

4.1 Materials Required

The captivation of nematodes to plant-produced substances is tested using an olfactometer. l Olfactometer. Apparatus description: – The olfactometer comprises a central glass chamber including six similarly arranged side arms fitted to a female connector. These arms are coupled to the center chamber by six glass pots where plants or other sources of attractants are kept. – Every single pot has a female connector. – The connecting arms comprise two removable sections: one is a glass tube with ground-glass connectors on each sides, while the other part is a Teflon connector used to connect the glass tube with the odor source pot. – The custom-made Teflon connectors comprise an ultra-fine metal screen (2300 mesh), averting the nematodes from attaining the odor source pots.

4.2

Procedure

l

Nematodes.

l

Sterile sand.

l

Attractant.

l

Filter disk.

l

Bearmann extractor.

l

Dissection microscope. 1. Fill the six outer pots and the central pot of the olfactometer with sterile sand approximately up to 5 cm from the rim of the pots. 2. Place the attractant into three outer pots alternated with three control pots that contained no attractant.

Chemotaxis Assay

39

Fig. 1 Six-arm olfactometer to test nematode attraction

3. Release the nematodes in a water drop exactly at mid-point in the center pot. 4. Allow for good passive diffusion of the attractants from the surrounding pots to the central arena. 5. Disassemble the olfactometer after 1 day of the nematode release. 6. Place the sand in individual removable glass tube on a distinct 19-cm diameter cotton filter disk. 7. Place the disk containing sand inside a Bearmann extractor. 8. Count the number of nematodes in the collection tube using dissection microscope. 9. Use these numbers as a measurement of the attractiveness of each tested odorant (Fig. 1).

5

Chemotaxis Assay EPNs respond differently to different host odors. Some odorants are appealing and revolting, and some have no specific response. The entomopathogenic Steinernema can jump, that is supposed to have a vital function in host-seeking and dispersal [149]. Jumping and chemotaxis assays are efficaciously applied to identify odorants which kindle these behaviors in a number of nematodes [150].

40

Chemical Ecology

5.1 Chemotaxis for EPN Behavior [146, 151, 152]

This assay is carried out to ascertain if chemotaxis is species and strain specific and to evaluate behavioral influence of EPNs toward volatile compounds.

5.1.1 Materials Required

l

Infective juveniles.

l

9-cm diameter petri dishes.

l

1.6% Technical agar.

l

Potassium phosphate (pH 6.0).

l

CaCl2.

l

MgSO4.

l

95% Linalool.

l

98% α-caryophyllene or 98% β-caryophyllene.

l

M9 buffer.

l

Parafilm.

l

Dissecting microscope.

5.1.2 Procedure

1. Add 25 ml agar, 5 mM potassium phosphate, 1 mM CaCl2, and 1 mM MgSO4 to the petri dish. 2. Make three spherical marks of 1 cm diameter on the underside of the petri dish—one in the center, one on the right side, and the other on the left side (1.5 cm from its edge). 3. Place 50 μl drop of 100 IJs in the midpoint of the agar surface. 4. Place 10 μl drop of linalool, α-caryophyllene, β-caryophyllene on the right side of the agar surface.

or

5. Place 10 μl of M9 buffer as control on the left side of the agar surface. 6. Seal the petri dishes with parafilm. 7. Once sealed, keep the petri dishes within a rearing chamber at 25
C and 75% RH without light. 8. Allow the nematodes to freely wander for 3 or 22 h. 9. Place these petri dishes inside a freezer for 5 min at 20
C and restrain the nematodes. 10. Count the total nematodes in the control and treatment part under a dissection microscope. 11. Repeat the experiments three times. 12. Include five replicates for each treatment. 13. Calculate the specific chemotaxis index (CI) [153] as given: (Number of nematodes in the treatment area  Number of nematodes in the control area)/Total number of nematodes in the assay.

Chemotaxis Assay

41

14. Chemotaxis index can alter from 1.0 (perfect attraction) to 1.0 (perfect repulsion). 5.2 Chemotaxis to Host Volatiles [141, 154, 155] 5.2.1 Materials Required

5.2.2 Procedure

l

Chemotaxis media plates.

l

KD scientific pump.

l

PVC (polyvinyl chloride) tube.

l

Nematodes (IJs).

l

Hamilton gastight syringes (50 ml).

l

Petri dish (9 cm diameter).

l

Media: 2% agar, 5 mM potassium phosphate (pH 6.0), 1 mM CaCl2, and 1 mM MgSO4. 1. Prepare chemotaxis media plates at room temperature at least 12 h before experiments start. 2. Fill test syringes with five alive, non-infected G. mellonella larvae, and leave the control syringes blank. 3. Load the syringes inside a KD scientific pump. 4. Drill two 10-mm holes on each side of the lid roughly 10 mm apart from the edges. 5. Attach PVC tube into holes and onto the syringes in order to join the two. 6. Air in the syringes travels onto the scoring circles through these tubes that are joined to the bottom of the chemotaxis plate. 7. Place 250 μl IJs (pellet) onto the midpoint of the chemotaxis plate. Note: Place an altered lid on top of the plates and position the tubing placed over its equivalent scoring circle. 8. Set the plates on a vibration-reducing stand for 1 h. 9. Score the assay with the scoring template fastened to the bottom of the chemotaxis plates.

10. Run at least three plates for every experiment at every time point, and individual experiment consists of nine technical replicates. 11. Run one complete experiment with unexposed IJs to the host cuticle, while another complete experiment is conducted with exposed IJs. 12. Determine the chemotaxis index (CI) values by counting the number of IJs within each scoring circle, which are being exposed to the host odors and control. CI ¼ No. in host circle  No. in control circle/Sum of all individuals in both circles (Fig. 2).

42

Chemical Ecology

Fig. 2 Template used to score chemotaxis assays for chemotaxis index 5.3 Jumping Assay [156]

l

Nematodes.

l

Tap water.

5.3.1 Materials Required

l

Odorants.

l

CO2.

l

Petri dish.

l

Whatman 1 filter paper.

l

Hamilton gastight syringe.

l

Luer-Lok Becton Dickinson syringe.

5.3.2 Procedure

1. Drill a 1.25-mm hole from side to side of a 5-cm petri dish and lid to permit odors to be added with the lid on, which prevents drying or the confounding effects of drafts. 2. Keep a 55-mm Whatman 1 filter paper on the underside of the dish to attract and hold moisture, thus supplying a fibrous material to make jumping easy. The filter paper performs as a soil-like layer. 3. Apply to this ~200 μl of tap water suspended with 800 μm Body length of IJ 1000 μm (1034–1171); male tail 4 without mucron Average length of IJ less than 1000 μm (849–951); male 8 tail with or without mucron 4. Tail of IJ 106 (119–160); female without epiptygma

5

Tail of IJ averaging >90 μm (95–95); E% 72 μm (77–84); E% 6 50

9

9. IJ body length averaging 951 μm (797–1102); spicule S. kraussei length about 49 μm (42–53) IJ body length averaging 849 μm (736–950); spicule length about 70 μm (65–77)

S. feltiae

10. Average length of IJ >600 μm (622–693)

11

Average length of IJ 540 μm

S. intermedia S. riobravis 14 15

14. First-generation male without mucron; spicule length S. ritte 69 μm (58–75), SW ¼ 1.56 (1.44–1.57); in IJ, E% averaging 88 First-generation male with mucron; spicule length 47 μm (42–52), SW ¼ 0.94 (0.91–1.05); in IJ, E% averages 72 15. First-generation male without mucron; in IJ, E% averaging 92 First-generation male with mucron; in IJ, E% 700 μm (736–800) Average body length of IJ 80 μm (84–119), E% ¼ 127 3 or less 3. IJ body length averaging >600 μm

4

IJ body length averaging 120, c >7; lamina of spicule with ventral expansion In IJ, E% cryptic foliage > exposed foliage) and trial location (greenhouse > field studies).

l

Comparative humidity and temperature during and up to 8 h after application have an impact on the nematode infection rates.

l

The greenhouse whitefly (GHWF), Trialeurodes vaporariorum (Westwood) (Hemiptera: Aleyrodidae), is a cosmopolitan, polyphagous, and grave insect pest all over the world. This whitefly is a critical and vivacious pest on several greenhouse crops like cucumbers, tomatoes, peppers, and an assortment of other plants [249, 250]. Around 250 plant species from diverse regions of the world are recorded as its host [251].

2.1 Greenhouse Experiment [252]

S. feltiae and H. bacteriophora have the ability to diminish the survival of T. vaporariorum. To enhance pest management and crop health within greenhouse productions.

2.1.1 Materials Required

l

Greenhouse whitefly.

l

Greenhouse crops cucumber and pepper.

l

Nematode species (IJs).

l

Cylindrical clip cage.

l

Petri dish (90 mm diameter).

l

1.5% Agar.

l

Triton X-100 (0.1% v/v).

l

Parafilm.

l

Stereomicroscope.

l

2.1.2 Procedure

1. Introduce nearly 30 adults with a sex ratio of 2:1 females to males of greenhouse whitefly (T. vaporariorum) in every cylindrical clip cage (10 cm diameter  11 cm height).

156

Application Efficacy Against Insects in Other Than Soil Habitats

2. Permit to mate and lay eggs. 3. Take out the adults after a 48 h oviposition period. 4. Preserve those plants within the greenhouse at 25  1 C, 65% RH for 16:8h (L:D). 5. Retain the plants in the greenhouse for a further 12 days until the presence of the second nymphal instar [253]. 6. Choose the second nymphal instar since the second nymphal stages are the most vulnerable to nematode infection. 7. Pick randomly 5–6 leaves from each plant (cucumber or pepper) infested with T. vaporariorum second nymphal instars. 8. Relocate each leaf into the petri dish filled with a thin layer of 1.5% agar. 9. Amend the nematode suspensions to yield the concentrations 0-control, 25, 50, 100, 150, 200, and 250 IJ/cm2 by volumetric dilutions in distilled water to an absolute volume of 1 ml. 10. Mix Triton X-100 to the EPN concentrations and the control comprising only distilled water with a handheld sprayer. 11. Spray each suspension onto the surface of each petri dish till runoff by means of a handheld sprayer. Have four replicates for each treatment, and repeat the whole experiment twice, with diverse batches of fresh nematodes, insects, and plants. 12. Execute the spraying at greenhouse in the afternoon. 13. Conceal the petri dish after inoculation of nematodes. 14. Maintain the plants at 20  1  C, 85% RH, and a 12:12 h L:D period for 72 h [254]. 15. Check the infection of S. feltiae and H. bacteriophora after 72 h treatment. 16. Have three replicates for each treatment, and repeat the whole experiment twice. 17. Dissect the cadavers using a stereomicroscope to check that mortality ensued from EPN infection. 2.2 Diaspora of Infective Juveniles from Greenhouse Cadavers [255]

To demonstrate the efficacy of formulated IJs against Hoplia philanthus in greenhouse.

2.2.1 Materials Required

l

Hoplia philanthus grub.

l

Greenhouse crop ryegrass.

l

Cadaver (HbK15-5).

l

PVC pipe (3.6 cm inner diameter).

l

Plastic pot.

Greenhouses

2.2.2 Procedure

l

Silicone adhesive.

l

Sterilized sandy soil.

l

24-Well plates.

l

Tap water.

l

Stereomicroscope.

157

1. Attach the PVC pipe to a hole (4 cm diameter) on the lateral surface of the plastic pot (8.2  6.7  9 cm; surface area 52.8 cm2) and fix by means of a silicone adhesive. 2. Close the left-out end of the pipe by a detachable cap. 3. Fill the pot and pipe with humid sanitized sandy soil. 4. Seed the pots consequently with perennial ryegrass. 5. Permit the plant to rise for 4 weeks earlier to infestation with H. philanthus grubs. 6. Keep the grubs separately in moist sand in 24-well plates at 15 ºC for 1 week. 7. Remove the dead grubs during this period. 8. Add six vigorous grubs on the surface of every pot 4 h before placement of one recently prepared cadaver (HbK15-5) at the opposite end of the pipe. 9. In the control treatment, place in every pot a single prepared cadaver in a hole made to a depth of 5 cm in the center of the pot and conceal with soil. 10. Remove the grubs that did not pass into the soil within 2 h and replace with healthy ones. 11. Follow five experimental treatments with five replications of each treatment. 12. Add 50 ml tap water on alternate days to every pot to sustain soil moisture. 13. Maintain the average air temperature within the greenhouse at 26 ºC (range 20–30 ºC). 14. After a fortnight, sample the pots detrimentally. 15. Count the live and deceased grubs. 16. Dissect the dead larvae under the stereomicroscope to check that mortality resulted from EPN infection. 17. Duplicate the experiment.

2.3 Greenhouse Pot Trial [256]

To study the efficacy and dispersal of EPN for control of black vine weevil (BVW).

158

Application Efficacy Against Insects in Other Than Soil Habitats

2.3.1 Materials Required

2.3.2 Procedure

l

Black vine weevil (BVW).

l

Composted green waste (CGW).

l

H. bacteriophora UWS 1.

l

Plastic pots (2 L).

l

Peat, bark, coir, and peat blends.

l

Tap water. 1. Fill plastic pots (13 cm height, 16 cm diameter, 201 cm2 surface area) with 10% peat, bark, coir, and peat blends. 2. Transplant 20% CGW. 3. Place eight third-instar BVW larvae all over the base of each plant 4 h before the start of an experiment. 4. Substitute the larvae that had not entered into the substrates within 2 h. 5. Smear H. bacteriophora as an inundate at three concentrations (1) low (1.2  109 IJs/ha), (2) medium (2.5  109 IJs/ha), and (3) high (5.0  109 IJs/ha). 6. Apply the EPN suspensions in 50 ml of tap water per pot trailed by another 50 ml of water to wash the EPN into the substrates. 7. Treat the control pots with 100 ml water only. 8. Arrange the pots in a random block design with 10 pots per treatment. 9. Sample the pots destructively after 2 weeks.

10. Record the EPN surviving BVW.

application

and

the

number

of

11. During the experiment, maintain the air temperature in the greenhouse 14.5–28.4 ºC (average 19.5 ºC). 12. Conduct the entire experiment twice.

Chapter 12 Recent Advances and Future Prospect Abstract After the discovery of entomopathogenic nematodes, they have proven to be excellent biocontrol agents for insect pests. Study is being performed in several parts of the world on entomopathogenic nematodes Steinernema and Heterorhabditis and their symbiotic bacteria Xenorhabdus and Photorhabdus. The future of entomopathogenic nematodes as potential biopesticides is promising. Their success is bound with the innovative ideas of incorporating EPN methodologies in different fields of study. Progressive advances in analysis and study on EPN promote the extensive use of EPNs. Keywords Genome sequence, Benzoxazinoid resistance, Cockchafer, Drosophila, Microgravity

1

Recent Advances Research on entomopathogenic nematodes Steinernema and Heterorhabditis and their symbiotic bacteria Xenorhabdus and Photorhabdus is being conducted in many parts of the world. Recent advances in molecular phylogeny associated with the increasing number of available genes and genome sequences have reassessed the taxonomy of the symbiotic bacteria of entomopathogenic nematodes with elevation of many subspecies to the species level and creation of new taxa [13]. Around 100 valid species of Steinernema and 21 species of Heterorhabditis have been identified till date from different countries of the world. Utilization of native EPN species would serve as an alternative to chemical pesticides and fit well in integrated pest management program [24]. EPN from Mexican maize fields are identified to overcome western corn rootworm defence metabolites and gains insights into the prevalence and relative importance of benzoxazinoid resistance in these EPN isolates [257]. H. indica, S. carpocapsae, and S. longicaudum are associated with higher mortality, penetration rate, and reproduction rate and effectively control Spodoptera litura, a polyphagous pest that is seriously affecting various crops [258].

S. Sivaramakrishnan and M. Razia, Entomopathogenic Nematodes and Their Symbiotic Bacteria, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-1445-7_12, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2021

159

160

Recent Advances and Future Prospect

An indigenous strain of Heterorhabditis bacteriophora Poinar has a high potential for biocontrol of white grub in the strawberry fields. The beneficial traits have a marked influence on EPN foraging behavior, persistence, and movement direction with implications for harnessing them as biological pest control agents [259]. The bacteria colonizing the midgut of the infected insect cockchafer (Melolontha melolontha) exhibit antagonistic activity against the selected species of the genera Xenorhabdus and Photorhabdus, thus opening avenues for increasing the efficacy of pest management programs [260]. EPNs present themselves as a model system in Drosophila to understand the host’s anti-nematode response furthering insight into clotting factors, immune factors, or hemocyte behavior, increased temporal and spatial infection kinetics to have a knowledge regarding a threshold for overcoming the host immune system, which has implications for both the agricultural industry and human health [261]. The first agricultural biocontrol experiment in space gives insight to dynamics of EPN foraging and infectivity in microgravity, long-term space flight for symbiotic organisms, parasite biology, and the potential for sustainable crop protection in space [262].

2

Future Prospect Entomopathogenic nematodes have been ecologically successful as exemplified by their wide distribution throughout the world [263]. Frequent surveys are done on isolation and identification of EPNs in different continents of the world to explore untouched geographic areas and climatic conditions, in both plantations and indigenous forests with an aim to identify and exploit additional EPN species. The study of symbiont properties, such as cellular exportation, exoenzymatic activities, special metabolites, pathogenic processes, capabilities to differentiate into multicellular populations for the colonization of different habitats, are bringing new insights for microbiology. Progress in the areas of nematode-bacteria mutualism makes EPNs as an outstanding model for research in symbiosis, ecology, evolution, molecular genetics, and biochemistry. The emerging new, natural bioactive compounds from the EPN bacterial symbionts, Xenorhabdus and Photorhabdus are anticipated to be used directly as pharmaceutical drugs. From an applied perspective, numerous technological innovations are accomplishing in relation to their implementation in biocontrol. The future of entomopathogenic nematodes as potential biopesticides is promising. Their success is bound with the innovative ideas of incorporating EPN methodologies in insect pest

Future Prospect

161

management programs. Present technologies like transposon mutagenesis, EST (expressed sequence tags) screening, RNAi technology, genetic engineering, and genetic improvement of strains used in EPN study are developing over the time. EPNs definitely have a very bright scope as an alternative to chemical pesticides. The EPN commercialization is likely to increase over the years in the future as they are eco-friendly and user-friendly. The application efficacy of EPNs is progressing by the current development in nematode formulation, application equipment or approaches, and strain improvement. Further studies in lowering product costs, increasing product availability, enhancing ease-ofuse, and improving efficacy and carryover effect will encourage the wide use of EPNs in biomanagement. These advances help EPNs to minimize chemical insecticide hazards and add to the maintenance of crop yields and the environment. Advanced research studies can be undertaken to explore the truth that may be concealed in EPNs. Further studies may include an in-depth study of host defences and immune responses to EPN infection, focusing on various abiotic and biotic factors that may affect the survival of soil IJs or EPN populations, to enhance the optimization of EPNs for greater use in the biological control of insect pests in the soil environment, mass formulation of nematodebased products, stability and application strategies to be economical with chemical insecticides, to understand the conservation and manipulation of the natural EPN populations strategy required in diverse habitats, to study and identify the novel valuable bioactive compounds from symbiotic bacteria and their various applications in different agricultural sectors, as well as medicinal value for dreadful diseases, to study the utility and importance of gene knock and gene knock in EPNs and their symbiotic bacteria, to employ EPNs as novel species to generate abiotic and biotic resistance to transgenic nematodes, to use EPNs as a model for human genetic diseases, and to use EPNs as an indicator of environmental pollution and climate change. Entomopathogenic nematodes can compete as stable potential biocontrol agents with any other organism.

References

1. Kaya HK, Gaugler R (1993) Entomopathogenic nematodes. Annu Rev Entomol 38:181–206 2. Cranshaw WS, Zimmerman R (2013) Insect parasitic nematodes. Fact sheet no. 5.573. Colorado State University extension 3. Poinar GO (1983) The natural history of nematodes. Prentice-Hall, Englewood Cliffs, NJ, 323 pp 4. Poinar GO Jr (1993) Origins and phylogenetic relationships of the entomophilic rhabditids, Heterorhabditis and Steinernema. Fundam Appl Nematol 16:333–338 5. Adams BJ, Burnell AM, Powers TO (1998) A phylogenetic analysis of Heterorhabditis (Nemata: Rhabditidae) based on internal transcribed spacer 1 DNA sequence data. J Nematol 30(1):22–39 6. Blaxter ML, De Ley P, Garey JR, Liu LX, Scheldeman P, Vierstraete A, Vanfleteren JR, Mackey LY, Dorris M, Frisse LM (1998) A molecular evolutionary framework for the phylum Nematoda. Nature 392:71–75 7. Liu J, Berry R, Poinar G, Moldenke A (1997) Phylogeny of Photorhabdus and Xenorhabdus species and strains as determined by comparison of partial 16S rRNA gene sequences. Int J Syst Bacteriol 47(4):948–951 8. Sudhaus W (1993) Redescription of Rhabditis (Oscheius) tipulae (Nematoda: Rhabditidae) associated with leatherjackets, larvae of Tipula paludosa (Diptera: Tipulidae). Nematologica 39:234–239 9. Poinar GO, Grewal PS (2012) History of entomopathogenic nematology. J Nematol 44(2):153–161

10. Koppenho¨fer HS (2007) Bacterial symbionts of Steinernema and Heterorhabditis. In: Nguyen KB, Hunt DJ (eds) Entomopathogenic nematodes: systematics, phylogeny and bacterial symbionts. Brill NV, Leiden 11. Stock SP (2015) Diversity, biology and evolutionary relationships. In: Campos-Herrera R (ed) Nematode pathogenesis of insects and other pests: ecology and applied technologies for sustainable plant and crop protection. Springer International Publishing, Neuchaˆtel, Switzerland, pp 3–27 12. Tailliez P, Laroui C, Ginibre N, Paule A, Pages S, Boemare N (2010) Phylogeny of Photorhabdus and Xenorhabdus based on universally conserved protein-coding sequences and implications for the taxonomy of these two genera. Proposal of new taxa. X. vietnamensis sp. nov., P. luminescens subsp. caribbeanensis subsp. nov., P. luminescens subsp. hainanensis subsp. nov., P. temperata subsp. khanii subsp. nov., P. temperata subsp. tasmaniensis subsp. nov., and the reclassification of P. luminescens subsp. thracensis as P. temperata subsp. thracensis comb. nov. Int J Syst Evol Microbiol 60 (8):1921–1937 13. Sajnaga E, Kazimierczak W (2020) Evolution and taxonomy of nematode-associated entomopathogenic bacteria of the genera Xenorhabdus and Photorhabdus: an overview. Symbiosis 80:1–13 14. Glaser RW, Fox H (1930) A nematode parasite of the Japanese beetle (Popillia japonica Newm.). Science 71:16–17 15. Weiser J (1955) Neoaplectana carpocapsae n. sp. (Anguillata, Steinernematidae) novy

S. Sivaramakrishnan and M. Razia, Entomopathogenic Nematodes and Their Symbiotic Bacteria, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-1445-7, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2021

163

164

References

cizopasnik housenek obalece jablecneho, Carpocapsae pomonella L. Vestnik Ceskoslovenske Spolecnosti Zoologicke 19:44–52 16. Dutky SR, Hough WS (1955) Note on a parasitic nematode from codling moth larvae, Carpocapsae pomonella. Proc. Entomol. Soc. Wash. 57:244 17. Poinar GO Jr, Thomas GM (1967) The nature of Achromobacter nematophilus as an insect pathogen. J Invertebr Pathol 9 (4):510–514 18. Pereira C (1937) Rhabditis hambletoni n.sp. nema apparentemente semiparasito da “broca do algodoeiro” (Gasterocercodes brasiliensis). Arch Inst Biol 8:215–230 19. Poinar GO Jr (2011) The evolutionary history of nematodes. Brill, Leiden. 439p 20. Akhurst RJ, Boemare NE (1990) Biology and taxonomy of Xenorhabdus. In: Gaugler R, Kaya HK (eds) Entomopathogenic nematodes in biological control. CRC Press, Boca Raton, FL, pp 75–90 21. Boemare NE (2002) Biology, taxonomy and systematics of Photorhabdus and Xenorhabdus. In: Gaugler R (ed) Entomopathogenic nematology. CABI Publishing, Wallingford, UK, pp 35–56 22. Hunt DJ, Subbotin SA (2016) Taxonomy and systematics. In: Hunt DJ, Nguyen KB (eds) Advances in entomopathogenic nematode taxonomy and phylogeny, Nematology monographs and perspectives, vol 12. Brill, The Netherlands, pp 13–58 23. Dutky SR (1959) Insect microbiology. Adv Appl Microbiol 1:175–200 24. Bhat AH, Chaubey AK, Askary TH (2020) Global distribution of entomopathogenic nematodes, Steinernema and Heterorhabditis. Egypt J Biol Pest Control 30:31 25. Steiner G (1923) Aplectana kraussei n. sp., einer in der Blattwespe Lyda sp. parasitierende Nematodenform, nebst Bemerkungen u¨ber das Seitenorgan der parasitischen nematoden. Zbl Bakt II Natur 2(59):14–18 26. Mra´cˇek Z, Hernandez EA, Boemare NE (1994) Steinernema cubana sp. n. (Nematoda: Rhabditida: Steinernematidae) and the preliminary characterisation of its associated bacterium. J Invertebr Pathol 64:123–129 27. Mra´cˇek Z (1977) Steinernema kraussei, a parasite of the body cavity of the sawfly, Cephaleia abietis, in Czechoslovakia. J Invertebr Pathol 30(1):87–94 28. Steiner WA (1994) The influence of air pollution on moss-dwelling animals: methodology and composition of flora and fauna. Rev Swis Zool 101:533–556

29. Hominick WM, Reid AP, Briscoe BR (1995) Prevalence and habitat specificity of steinernematid and heterorhabditid nematodes isolated during soil surveys of the UK and the Netherlands. J Helminthol 69(1):27–32 30. Del Pino FG, Palomo A (1996) Natural occurrence of entomopathogenic nematodes (Rhabditida: Steinernematidae and Heterorhabditidae) in Spanish soils. J Invertebr Pathol 68(1):84–90 31. Stock SP, Pryor BM, Kaya HK (1999) Distribution of entomopathogenic nematodes (Steinernematidae and Heterorhabditidae) in natural habitats in California, USA. Biodivers Conserv 8:535–549 32. Filipjev IN (1934) Miscellanea Nematologica 1. Eine neue Art der Gattung Neoaplectana Steiner nebst Bermerkungen u¨ber die systematische Stellung der letzteren. Parazitologich 4:229–240 33. Wouts W, Mracek Z, Gerdin S, Bedding RA (1982) Neoaplectana Steiner, 1929 a junior synonym of Steinernema Travassos, 1927 (Nematoda: Rhabditida). Syst Parasitol 4:147–154 34. Bovien P (1937) Some types of association between nematodes and insects, vol 101. Videnskabelige Meddelelser fra Dansk Naturhistorisk Forening, København, pp 1–114 35. Poinar GO Jr (1985) Neoaplectana intermedia n. sp. (Steinernematidae: Nematoda) from South Carolina. Rev Nematol 8:321–327 36. Mamiya Y (1988) Steinernema kushidai n. sp. (Nematoda: Steinernematidae) associated with scarabaeid beetle larvae from Shizuoka, Japan. Appl Entomol Zool 23:313–320 37. Shen CP, Wang GH. (1992). Description of an entomopathogenic nematode, Steinernema longicaudum sp. nov. and its application. In Proceedings of the XIX international congress of entomology, Beijing, China, pp 220–231 38. Katumanyane A, Malan AP, Tiedt LR, Hurley BP (2020) Steinernema bertusi n. sp. (Rhabditida: Steinernematidae), a new entomopathogenic nematode from South Africa. Nematology 22(3):343–360. https://doi. org/10.1163/15685411-00003309 39. Poinar GO Jr (1976) Description and biology of a new insect parasitic rhabitoid, Heterorhabditis bacteriophora n. gen. n. sp. (Rhabditida; Heterorhabditidae n. family). Nematologica 21:463–470 40. Poinar GO (1990) Taxonomy and biology of Steinernematidae and Heterorhabditidae. In: Gaugler R, Kaya HK (eds) Entomopathogenic nematodes in biological control. CRC Press, Boca Raton, FL, pp 23–61

References 41. Poinar GO Jr, Karunakar GK, David H (1992) Heterorhabditis indicus n. sp. (Rhabditida, Nematoda) from India: separation of Heterorhabditis spp. by infective juveniles. Fund Appl Nematol 15:467–472 42. Liu J, Berry RE (1996) Steinernema oregonensis n. sp. (Rhabditida: Steinernematidae) from Oregon, USA. Fundam Appl Nematol 19 (4):375–380 43. Stock SP, Choo HY, Kaya HK (1997) An entomopathogenic nematode Steinernema monitocolum sp. n. (Rhabditida: Steinernematidae) from Korea with a key to other species. Nematologica 43:15–29 44. Malan AP, Knoetze R, Louwrens T (2014) Heterorhabditis noenieputensis n. sp. (Rhabditida: Heterorhabditidae), a new entomopathogenic nematode from South Africa. J Helminthol 88:139–151 45. Boag B, Neilson R, Gordon SC (1992) Distribution and prevalence of the entomopathogenic nematode Steinernema feltiae in Scotland. Ann Appl Biol 121:355–360 46. Mracek Z, Becvar S, Kindlann P (1999) Survey of entomopathogenic nematodes from the families Steinernematidae and Heterorhabditidae (Nematoda: Rhabditida) in the Czech Republic. Folia Parasitol 46:145–148 47. Stock SP, Mra´cek Z, Webster J (2000) Morphological variation between allopatric populations of Steinernema kraussei (Steiner, 1923) (Rhabditida: Steinernematidae). Nematology 2(2):143–152 48. Sturhan D (1999) Prevalence and habitat specificity of entomopathogenic nematodes in Germany. In: Proceedings of a workshop held at Todi, Perugia, Italy, 16 to 29 May 1995. European Commission, COST, Luxembourg, vol 819, pp 123–132 49. Campbell PM, Newcomb RD, Russell RJ, Oakeshott JG (1998) Two different amino acid substitutions in the ali-esterase, E3, confer alternative types of organophosphorus insecticide resistance in the sheep blowfly, Lucilia cuprina. Insect Biochem Mol Biol 28 (3):139–150 50. Griffin CT, Joyce SA, Dix I, Burnell AM, Downes MJ (1994) Characterisation of the entomopathogenic nematode Heterorhabditis (Nematoda: Heterorhabditidae) from Ireland and Britain by molecular and crossbreeding techniques, and the occurrence of the genus in these islands. Fundam Appl Nematol 17:245–253 51. Yoshida M, Reid AP, Briscoe BR, Hominick WM (1998) Survey of entomopathogenic nematodes (Rhabditida: Steinernematidae

165

and Heterorhabditidae) in Japan. Fundam Appl Nematol 21(2):185–198 52. Stock SP, Kaya HK (1996) A multivariate analysis of morphometric characters of Heterorhabditis species (Nemata: Heterorhabditidae) and the role of morphometrics in the taxonomy of species of the genus. J Parasitol 82:806–813 53. Stuart RJ, Gaugler R (1994) Patchiness in populations of entomopathogenic nematodes. J Invertebr Pathol 64(1):39–45 54. Constant P, Marchay L, Fischer-Le-Saux M, Briand-Panoma S, Maule´on H (1998) Natural occurrence of entomopathogenic nematodes (Rhabditida: Steinernematidae and Heterorhabditidae) in Guadeloupe islands. Fundam Appl Nematol 21(6):667–672 55. Griffin CT, Dix I, Joyce SA, Burnell AM, Downes MJ (1999) Isolation and characterisation of Heterorhabditis spp. (Nematoda: Heterorhabditidae) from Hungary, Estonia and Dennmark. Nematology 1:321–332 56. Chitwood BG (1933) Notes on nematode systematics and nomenclature. J Parasitol 19:242–243. (W.L. 11428) ¨ rley L (1880) Az anguillulida´k maga´nrajza. A 57. O kir. m. terme´szettudom. ta´rsulat a´ltal a bugat dijjal jutalmazott pa´lyamu ¨ . (Monographic der Anguillulidcn. Eine von der k. ung. naturhistorischen Gescllschaft gekronte Prcisschrift). Terme´szetr Fuz 4:16–150. (German text): 154–177; pls. 1–7. (W.L. 21105) 58. Travassos LP (1920) Esboc¸o de uma chave geral dos nematodes parasitos. Rev Vet Rio de J 10:59–70. vis. (W.L. 18971) 59. Chitwood BG, Chitwood MB (1937) “Outline classification of Nematoda”. [Appendix to] An introduction to nematology, sect. I, pt. I, pp 49–52 60. Phylogenetic relationships of Steinernema Travassos, 1927 (Nematoda: Cephalobina: Steinernematidae) based on nuclear, mitochondrial and morphological data 61. Steiner G (1929) Neoaplectana glaseri n. g., n. sp. (Oxyuridae) a new nemic parasite of the Japanese beetle (Popillia japonica Newm.). J Wash Acad Sci 19:436–440 62. Nguyen KB, Smart GC Jr (1994) Neosteinernema longicurvicauda n. gen., n. sp. (Rhabdifida: Steinernematidae), a parasite of the termite Reticulitermes flavipes (Koller). J Nematol 26:162–174 63. Brooks, Hirschmann (1976) Chromonema heliothidis n. gen., n. sp. (Steinernematidae, Nematoda), a parasite of Hellothis zea (Noctuidae, Lepidoptera), and other insects. J Nematol 8(2):159–168

166

References

64. Khan A, Brooks WM, Hirschmann H (1976) Chromonema heliothidis n. gen., n. sp.(Steinernematidae, Nematoda), a parasite of Heliothis zea (Noctuidae, Lepidoptera), and other insects. J Nematol 8(2):159 65. Thomas GM, Poinar GO Jr (1979) Xenorhabdus gen. nov., a genus of entomopathogenic nematophilic bacteria of the family Enterobacteriaceae. Int J Syst Bacteriol 29:352–360 66. Boemare NE, Akhurst RJ, Mourant RG (1993) DNA relatedness between Xenorhabdus spp. (Enterobacteriaceae), symbiotic bacteria of entomopathogenic nematodes, and a proposal to transfer Xenorhabdus luminescens to a new genus, Photorhabdus gen. nov. Int J Syst Evol Microbiol 43(2):249–255 67. Bird AF, Akhurst RJ (1983) The nature of the intestinal vesicle in nematodes of the family Steinernematidae. Int J Parasitol 13 (6):599–606 68. Boemare N, Laumond C, Mauleon H (1996) The entomopathogenic nematode-bacterium complex: biology, life cycle and vertebrate safety. Biocontrol Sci Tech 6(3):333–346 69. Endo BY, Nickle WR (1994) Ultrastructure of the buccal cavity region and oesophagus of the insect parasitic nematode, Heterorhabditis Bacteriophora1. Nematologica 40 (1–4):379–398 70. Spiridonov SE, Akhmedov EN, Belostotskaya FN (1991) Proliferation of symbiotic bacteria in the intestinal vesicles of invasive larvae of Neoaplectana spp. (Nematoda, Steinernematidae). Helminthologia 28:141–142 71. Forst S, Clarke D (2002) Bacteria-nematode symbiosis. In: Gaugler R (ed) Entomopathogenic nematology. CABI Publishing, Wallingford, UK, pp 57–77 72. Koppenho¨fer HS, Gaugler R (2009) Entomopathogenic nematode and bacteria mutualism. In: White J, Torres M (eds) Defensive mutualism in microbial symbiosis. CRC Press, Boca Raton, FL, pp 99–116 73. Poinar GO Jr, Thomas GM (1965) A new bacterium, Achromobacter Nematophilus sp. nov. (Achromobactericeae Eubacteriales) associated with a nematode. Int Bull Bact Nomen Tax 15:249–252 74. Akhurst RJ (1983) Taxonomic study of Xenorhabdus, a genus of bacteria symbiotically associated with insect pathogenic nematodes. Int J Syst Bacteriol 33(1):38–45 75. Akhurst RJ (1986a) Xenorhabdus nematophilus subsp. poinarii: its interaction with insect pathogenic nematodes. Syst Appl Microbiol 8 (1–2):142–147

76. Akhurst RJ (1986b) Xenorhabdus nematophilus subsp. beddingii (Enterobacteriaceae): a new subspecies of bacteria mutualistically associated with entomopathogenic nematodes. Int J Syst Bacteriol 36:454–457 77. Nishimura Y, Hagiwara A, Suzuki T, Yamanaka S (1994) Xenorhabdus japonicus sp. nov. associated with the nematode Steinernema kushidai. World J Microbiol Biotechnol 10 (2):207–210 78. Lengyel K, Lang E, Fodor A, Szallas E, Schumann P, Stackebrandt E (2005) Description of four novel species of Xenorhabdus, family Enterobacteriaceae: Xenorhabdus budapestensis sp. nov., Xenorhabdus ehlersii sp. nov., Xenorhabdus innexi sp. nov., and Xenorhabdus szentirmaii sp. nov. Syst Appl Microbiol 28(2):115–122 79. Tailliez P, Pages S, Ginibre N, Boemare N (2006) New insight into diversity in the genus Xenorhabdus, including the description of ten novel species. Int J Syst Evol Microbiol 56(12):2805–2818 80. Somvanshi VS, Lang E, Ganguly S, Swiderski J, Saxena AK, Stackebrandt E (2006) A novel species of Xenorhabdus, family Enterobacteriaceae: Xenorhabdus indica sp. nov., symbiotically associated with entomopathogenic nematode Steinernema thermophilum Ganguly and Singh, 2000. Syst Appl Microbiol 29(7):519–525 81. Tailliez P, Pages S, Edgington S, Tymo LM, Buddie AG (2012) Description of Xenorhabdus magdalenensis sp. nov., the symbiotic bacterium associated with Steinernema australe. Int J Syst Evol Microbiol 62(8):1761–1765 82. Kuwata R, Qiu LH, Wang W, Harada Y, Yoshida M, Kondo E, Yoshiga T (2013) Xenorhabdus ishibashii sp. nov., isolated from the entomopathogenic nematode Steinernema aciari. Int J Syst Evol Microbiol 63:1690–1695 83. Ferreira T, van Reenen CA, Endo A, Sproer C, Malan AP, Dicks LMT (2013) Description of Xenorhabdus khoisanae sp. nov., the symbiont of the entomopathogenic nematode Steinernema khoisanae. Int J Syst Evol Microbiol 63 (9):3220–3224 84. K€ampfer P, Tobias NJ, Ke LP, Bode HB, Glaeser SP (2017) Xenorhabdus thuongxuanensis sp. nov. and Xenorhabdus eapokensis sp. nov., isolated from Steinernema species. Int J Syst Evol Microbiol 67(5):1107–1114 85. Fischer-Le Saux M, Viallard V, Brunel B, Normand P, Boemare NE (1999) Polyphasic classification of the genus Photorhabdus and proposal of new taxa: P. luminescens subsp.

References luminescens subsp. nov., P. luminescens subsp. akhurstii subsp. nov., P. luminescens subsp. laumondii subsp. nov., P. temperata sp. nov., P. temperate subsp. temperata subsp. nov. and P. asymbiotica sp. nov. Int J Syst Bacteriol 49 (4):1645–1656 86. Machado RAR, Bruno P, Arce CCM, Liechti N, Ko¨hler A, Bernal J, Bruggmann R, Turlings TCJ (2019) Photorhabdus khanii subsp. guanajuatensis subsp. nov., isolated from Heterorhabditis atacamensis, and Photorhabdus luminescens subsp. Mexicana subsp. nov., isolated from Heterorhabditis mexicana entomopathogenic nematodes. Int J Syst Evol Microbiol 69 (3):652–661 87. Machado RAR, Wu¨thrich D, Kuhnert P, Arce CCM, Tho¨nen L, Ruiz C, Zhang X, Robert CAM, Karimi J, Kamali S, Ma J, Bruggmann R, Erb M (2018) Wholegenome-based revisit of Photorhabdus phylogeny: proposal for the elevation of most Photorhabdus subspecies to the species level and description of one novel species Photorhabdus bodei sp. nov., and one novel subspecies Photorhabdus laumondii subsp. clarkei subsp. nov. Int J Syst Evol Microbiol 68 (8):2664–2681 88. Hazir S, Stackebrandt E, Lang E, Schumann P, Ehlers R-U, Keskin N (2004) Two new subspecies of Photorhabdus luminescens, isolated from Heterorhabditis bacteriophora (Nematoda: Heterorhabditidae): Photorhabdus luminescens subsp. Kayaii subsp. nov. and Photorhabdus luminescens subsp. thracensis subsp. nov. Syst Appl Microbiol 27(1):36–42 89. Akhurst RJ, Boemare NE, Janssen PH, Peel MM, Alfredson DA, Beard CE (2004) Taxonomy of Australian clinical isolates of the genus Photorhabdus and proposal of Photorhabdus asymbiotica subsp. asymbiotica subsp. nov. and P. asymbiotica subsp. australis subsp. nov. Int J Syst Evol Microbiol 54 (4):1301–1310 90. To´th T, Lakatos T (2008) Photorhabdus temperata subsp. cinerea subsp. nov., isolated from Heterorhabditis nematodes. Int J Syst Evol Microbiol 58(11):2579–2581 91. An R, Grewal PS (2011) Photorhabdus luminescens subsp. kleinii subsp. nov. (Enterobacteriales: Enterobacteriaceae). Curr Microbiol 62(2):539–543 92. Orozco RA, Hill T, Stock SP (2013) Characterization and phylogenetic relationships of Photorhabdus luminescens subsp. sonorensis (γ- Proteobacteria: Enterobacteriaceae), the bacterial symbiont of the entomopathogenic nematode Heterorhabditis sonorensis

167

(Nematoda: Heterorhabditidae). Curr Microbiol 66(1):30–39 93. Mahmoud MF, Mahfouz HM, Mohamed KM (2016) Compatibility of entomopathogenic nematodes with neonctinoids and azadirachtin insecticides for controlling the black cutworm, Agrotis ipsilon (Hufnagel) in canola plants. IJEST 2(1):11–18 94. Ehlers RU (2001) Mass production of entomopathogenic nematodes for plant protection. Appl Microbiol Biotechnol 56 (5–6):623–633 95. Kondo E, Ishibashi N (1986) Nictating behavior and infectivity of entomogenous nematodes, Steinernema spp., to the larvae of common cutworm, Spodoptera litura (Lepidoptera: Noctuidae), on the soil surface. Appl Entomol Zool 21(4):553–560 96. Strauch E, Arnold W, Alijah R, Wohlleben W, Puhler A, Eckes P, Wengenmayer F (1994) U.S. Patent No. 5,276,268. U.S. Patent and Trademark Office, Washington, DC 97. Griffin CT, Boemare NE, Lewis EE (2005) Biology and behavior. In: Grewal PS, Ehler R-U, Shapiro-Ilan DI (eds) Nematodes as biocontrol agents. CABI Publishing, Wallingford, UK, pp 47–64 98. Kaya HK, Koppenho¨fer AM (1999) Biology and ecology of insecticidal nematodes. In: Optimal use of insecticidal nematodes in pest management. IntechOpen, Croatia, pp 1–8 99. Sivaramakrishnan S, Razia M (2015) Sustainability of entomopathogenic nematodes against crop pests. In: Biocontrol of lepidopteran pests. Springer, Cham, pp 315–328 100. Chen MJ (1996) Competitor analysis and interfirm rivalry: toward a theoretical integration. Acad Manag Rev 21:100–134 101. Li J, Chen G, Webster JM (1997) Nematophin, a novel antimicrobial substance produced by Xenorhabdus nematophilus (Enterobactereaceae). Can J Microbiol 43:770–773 102. McInerney BV, Taylor WC, Lacey MJ, Akhurst RJ, Gregson RP (1991) Biologically active metabolites from Xenorhabdus spp. Part 2. Benzopyran-1-one derivatives with gastroprotective activity. J Nat Prod 54:785–795 103. Webster SP, Alexeev D, Campopiano DJ, Watt RM, Alexeeva M, Sawyer L, Baxter RL (2000) Mechanism of 8-amino-7-oxononanoate synthase: spectroscopic, kinetic, and crystallographic studies. Biochemistry 39 (3):516–528 104. Razia M, Karthik Raja R, Padmanaban K, Chellapandi P, Sivaramakrishnan S (2010) A phylogenetic approach for assigning function

168

References

of b-ketoacyl synthase I in entomopathogenic bacteria (Photorhabdus luminescens subsp. laumondii TT01). J Comput Sci Syst Biol 3:21–29 105. Akhurst RJ (1982) Antibiotic activity of Xenorhabdus spp., bacteria symbiotically associated with insect pathogenic nematodes of the families Heterorhabditidae and Steinernematidae. J Gen Microbiol 128:3061–3065 106. Khush RS, Lemaitre B (2000) Genes that fight infection: what the Drosophila genome says about animal immunity. Trends Genet 16:442–449 107. Chaston JM, Suen G, Tucker SL, Andersen AW, Bhasin A, Bode E (2011) The entomopathogenic bacterial endosymbionts Xenorhabdus and Photorhabdus: convergent lifestyles from divergent genomes. PLoS One 6(11):e27909 108. Ogier J, Calteau A, Forst SJ, Blair H, Roche D, Rouy Z, Suen G, Zumbihl R, Givaudan A, Tailliez P, Me´digue C, Gaudriault S (2010) Units of plasticity in bacterial genomes: new insight from the comparative genomics of two bacteria interacting with invertebrates, Photorhabdus and Xenorhabdus. BMC Genomics 11:568 109. Boemare NE, Boyer-Giglio MH, Thaler JO, Akhurst RJ, Brehelin M (1992) Lysogeny and bacteriocinogeny in Xenorhabdus nematophilus, and other Xenorhabdus spp. Appl Environ Microbiol 58:3032–3037 110. Bo¨szo¨rme´nyi E, Ersek T, Fodor A, Fodor AM, Fo¨ldes LS, Hevesi M, Hogan JS, Katona Z, Klein MG, Korma´ny A, Peka´r S, Szentirmai A, Sztaricskai F, Taylor RA (2009) Isolation and activity of Xenorhabdus antimicrobial compounds against the plant pathogens Erwinia amylovora and Phytophthora nicotianae. J Appl Microbiol 107:746–759 111. Xiao Q, Komori H, Lee CY (2012) Klumpfuss distinguishes stem cells from progenitor cells during asymmetric neuroblast division. Development 139(15):2670–2680 112. Fuchs MA, Sato K, Niedzwiecki D, Ye X, Saltz LB, Mayer RJ et al (2014) Sugar-sweetened beverage intake and cancer recurrence and survival in CALGB 89803 (Alliance). PLoS One 9(6):e99816 113. Tobias NJ, Heinrich AK, Eresmann H, Wright PR, Neubacher N, Backofen R, Bode HB (2017) Photorhabdus-nematode symbiosis is dependent on hfq-mediated regulation of secondary metabolites. Environ Microbiol 19:119–129 114. Zhou Y, Chen C, Johansson MJ (2013) The pre-mRNA retention and splicing complex

controls tRNA maturation by promoting TAN1 expression. Nucleic Acids Res 41 (11):5669–5678 115. Bode GH, Coue´ G, Freese C, Pickl KE, Sanchez-Purra` M, Albaiges B, Borro´s S, van Winden EC, Tziveleka LA, Sideratou Z, Engbersen JFJ, Singh S, Albrecht K, Groll J, Mo¨ller M, Po¨tgens AJG, Schmitz C, Fro¨hlich E, Grandfils C, Sinner FM, Kirkpatrick CJ, Steinbusch HWM, Frank HG, Unger RE, Martinez-Martinez P (2017) An in vitro and in vivo study of peptidefunctionalized nanoparticles for brain targeting: the importance of selective blood–brain barrier uptake. Nanomed Nanotechnol Biol Med 13:1289–1300 116. Reimer D, Luxenburger E, Brachmann AO, Bode HB (2009) A new type of pyrrolidine biosynthesis is involved in the late steps of xenocoumacin production in Xenorhabdus nematophila. Chembiochem 10:1997–2001 117. Sugar DR, Murfin KE, Chaston JM, Andersen AW, Richards GR, deLe´on L (2012) Phenotypic variation and host interactions of Xenorhabdus bovienii SS-2004, the entomopathogenic symbiont of Steinernema jollieti nematodes. Environ Microbiol 14:924–939 118. Park HB, Perez CE, Perry EK, Crawford JM (2016) Activating and attenuating the amicoumacin antibiotics. Molecules 21:E824 119. Reimer D, Nollmann FI, Schultz K, Kaiser M, Bode HB (2014) Xenortide biosynthesis by entomopathogenic Xenorhabdus nematophila. J Nat Prod 77:1976–1980 120. Hu MCT, Qiu WR, Wang X, Meyer CF, Tan T-H (1996) Human HPK1, a novel human hematopoietic progenitor kinase that activates the JNK/SAPK kinase cascade. Genes Dev 10:2251–2264 ˜ez G (1999) 121. Hu Y, Benedict MA, Ding L, Nu´n Role of cytochrome c and dATP/ATP hydrolysis in Apaf-1-mediated caspase-9 activation and apoptosis. EMBO J 18:3586–3595 122. Webster MS, Marra PP, Haig SM, Bensch S, Holmes RT (2002) Links between worlds: unraveling migratory connectivity. Trends Ecol Evol 17:76–83 123. Gaugler R, Kaya HK (1990) Entomopathogenic nematodes in biological control. CRC Press, Boca Raton, FL 124. Cutler GC, Webster JM (2003) Host-finding ability of three entomopathogenic nematode isolates in the presence of plant roots. Nematology 5(4):601–608 125. Glaser RW, Farrell CC (1935) Field experiments with the Japanese beetle and its nematode parasite. J N Y Entomol Soc. 43p

References 126. Prasad BC, Reed RR (1999) Chemosensation: molecular mechanisms in worms and mammals. Trends Genet 15(4):150–153 127. Anderson OD, Greene FC (1997) The α-gliadin gene family. II. DNA and protein sequence variation, subfamily structure, and origins of pseudogenes. Theor Appl Genet 95 (1–2):59–65 128. Hui E, Webster JM (2000) Influence of insect larvae and seedling roots on the host-finding ability of Steinernema feltiae (Nematoda: Steinernematidae). J Invertebr Pathol 75 (2):152–162 129. Kaya H, Campbell J (2000) Influence of insect associated cues on the jumping behavior of entomopathogenic nematodes (Steinernema spp.). Behaviour 137(5):591–609 130. Gaugler R, Campbell JF (1993) Nictation behaviour and its ecological implications in the host search strategies of entomopathogenic nematodes (Heterorhabditidae and Steinernematidae). Behaviour 126 (3–4):155–169 131. Volk J (1950) Die Nematoden der Regenwu¨rmer und der aasbesuchenden K€afer. Zool Jb Syst 79:1 132. Lewis EE (2002) 10 behavioural ecology. In: Entomopathogenic nematology. CABI Publishing, Wallingford, UK, p 205 133. Croll NA (1970) The behaviour of nematodes: their activity, senses and responses. Edward Arnold, London 134. Koppenho¨fer AM, Fuzy EM (2003) Steinernema scarabaei for the control of white grubs. Biol Control 28(1):47–59 135. Ali JG, Alborn HT, Stelinski LL (2010) Subterranean herbivore-induced volatiles released by citrus roots upon feeding by Diaprepes abbreviatus recruit entomopathogenic nematodes. J Chem Ecol 36(4):361–368 136. Rasmann S, Ko¨llner TG, Degenhardt J, Hiltpold I, Toepfer S, Kuhlmann U, Turlings TC (2005) Recruitment of entomopathogenic nematodes by insect-damaged maize roots. Nature 434(7034):732–737 137. Gulcu B, Hazir S, Kaya HK (2012) Scavenger deterrent factor (SDF) from symbiotic bacteria of entomopathogenic nematodes. J Invertebr Pathol 110(3):326–333 138. Lu JL, Luo W, Li XY, Yang SQ, Cao JX, Gong XG, Xiang HJ (2017) Two-dimensional node-line semimetals in a honeycombkagome lattice. Chin Phys Lett 34(5):057302 139. Turlings TC, Hiltpold I, Rasmann S (2012) The importance of root-produced volatiles as foraging cues for entomopathogenic nematodes. Plant Soil 358(1–2):51–60

169

140. Gang SS, Hallem EA (2016) Mechanisms of host seeking by parasitic nematodes. Mol Biochem Parasitol 208(1):23–32 141. Hallem EA, Dillman AR, Hong AV, Zhang Y, Yano JM, SF DM, Sternberg PW (2011) A sensory code for host seeking in parasitic nematodes. Curr Biol 21(5):377–383 142. Gaugler R, Lebeck L, Nakagaki B, Boush GM (1980) Orientation of the entomogenous nematode Neoaplectana carpocapsae to carbon dioxide. Environ Entomol 9(5):649–652 ˇ , Trdan S (2016) Attraction beha143. Laznik Z viors of entomopathogenic nematodes (Steinernematidae and Heterorhabditidae) to synthetic volatiles emitted by insect-damaged carrot roots. J Pest Sci 89(4):977–984 144. Robinson AF (1995) Optimal release rates for attracting MeloIDogyne incognita, Rotylenchulus reniformis, and other nematodes to carbon dioxIDe in sand. J Nematol 27:42–50 145. Ali MS, Kim GD, Seo HW, Jung EY, Kim BW, Yang HS, Joo ST (2011) Possibility of making low-fat sausages from duck meat with addition of rice flour. Asian Australas J Anim Sci 24(3):421–428 146. O’Halloran DM, Burnell AM (2003) An investigation of chemotaxis in the insect parasitic nematode Heterorhabditis bacteriophora. Parasitology 127(4):375 147. Ko¨llner TG, Held M, Lenk C, Hiltpold I, Turlings TC, Gershenzon J, Degenhardt J (2008) A maize (E)-β-caryophyllene synthase implicated in indirect defense responses against herbivores is not expressed in most American maize varieties. Plant Cell 20 (2):482–494 148. Helms AM, Ray S, Matulis NL, Kuzemchak MC, Grisales W, Tooker JF, Ali JG (2019) Chemical cues linked to risk: cues from below-ground natural enemies enhance plant defences and influence herbivore behaviour and performance. Funct Ecol 33(5):798–808 149. Dillman AR, Sternberg PW (2012) Entomopathogenic nematodes. Curr Biol 22(11): R430–R431 150. Campbell JF, Kaya HK (1999) How and why a parasitic nematode jumps. Nature 397 (6719):485–486 151. Laznik Z, Trdan S (2013) An investigation on the chemotactic responses of different entomopathogenic nematode strains to mechanically damaged maize root volatile compounds. Exp Parasitol 134(3):349–355 152. Ward S (1973) Chemotaxis by the nematode Caenorhabditis elegans: identification of attractants and analysis of the response by

170

References

use of mutants. Proc Natl Acad Sci U S A 70:817–821 153. Bargmann CI, Horvitz HR (1991) Chemosensory neurons with overlapping functions direct chemotaxis to multiple chemicals in C. elegans. Neuron 7(5):729–742 154. Baiocchi T, Lee G, Choe DH, Dillman AR (2017) Host seeking parasitic nematodes use specific odors to assess host resources. Sci Rep 7(1):1–13 155. Dillman AR, Chaston JM, Adams BJ, Ciche TA, Goodrich-Blair H, Stock SP, Sternberg PW (2012) An entomopathogenic nematode by any other name. PLoS Pathog 8(3): e1002527 156. Baiocchi T, Dillman AR (2015) Chemotaxis and jumping assays in nematodes. Bio-protocol 5:e1587 157. Stilwell RL, Samaniego AC, Atkinson B (2017) U.S. Patent No. 9,801,975. U.S. Patent and Trademark Office, Washington, DC 158. Southey JF (1986) Laboratory methods for work with plant and soil nematodes. HMSO, London 159. Jenkins WRB (1964) A rapid centrifugalflotation technique for separating nematodes from soil. Pl Dis Rep 48(9) 160. Bedding RA, Akhurst RJ (1975) A simple technique for the detection of insect paristic rhabditid nematodes in soil. Nematologica 21 (1):109–110 161. Fan X, Hominick WM (1991) Efficiency of the Galleria (wax month) baiting technique for recovering infective stages of entomopathogenic rhabditids (Steinernematidae and heterorhabditidae) from sand and soil. Rev Nematol 14(3):381–387 162. White GF (1927) A method for obtaining infective nematode larvae from cultures. Science 66(1709):302–303 163. Kaya HK, Stock SP (1997) Techniques in insect nematology. In: Lacey LA (ed) Manual of techniques in insect pathology. Academic Press, London, pp 281–324 164. Woodring JL, Kaya HK (1988) Steinernematid and Heterorhabditid nematodes: a handbook of techniques. South Co-operative Serv Bull 331(1):30 165. Southey JF (1970) Principles of sampling for mematodes. Technical bulletin. Ministry of Agriculture, Fisheries and Food (5th edn; 2), pp 1–4 166. Seinhorst JW (1959) A rapid method for the transfer of nematodes from fixative to anhydrous glycerin. Nematologica 4(1):67–69

167. Brenner S (1974) The genetics of Caenorhabditis elegans. Genetics 77(1):71–94 168. Poinar GO Jr (1967) Description and taxonomic position of the DD-136 nematode (Steinernematidae, Rhabditoidea) and its relationship to Neoaplectana carpocapsae Weiser. Proc Helminthol Soc Wash 34:199–209 169. Mayr E (1978) Origin and history of some terms in systematic and evolutionary biology. Syst Zool 27(1):83–88 170. Poinar GO Jr, Thomas GM (1966) Significance of Achromobacter nematophilus (Achromobactericeae Eubacteriales) in the development of the nematode, DD-136 (Neoaplectana sp. Steinernematidae). Parasitology 56:385–390 171. Dix DR, Bridgham JT, Broderius MA, Byersdorfer CA, Eide DJ (1994) The FET4 gene encodes the low affinity Fe(II) transport protein of Saccharomyces cerevisiae. J Biol Chem 269(42):26092–26099 172. Joyce CM, Steitz TA (1994) Function and structure relationships in DNA polymerases. Annu Rev Biochem 63(1):777–822 173. Reid MB, Stokic´ DS, Koch SM, Khawli FA, Leis AA (1994) N-acetylcysteine inhibits muscle fatigue in humans. J Clin Invest 94 (6):2468–2474 174. Hashmi G, Gaugler R (1998) Genetic diversity in insect-parasitic nematodes (Rhabditida: Heterorhabditidae). J Invertebr Pathol 72 (3):185–189 175. Reid IN, Gizis JE (1997) Low-mass binaries in the Hyades-A scarcity of brown dwarfs. arXiv preprint astro-ph/9709226 176. Reid AP, Homonick WM (1998) Molecular taxonomy of Steinernema by RFLP analysis of the ITS region of the ribosomal DNA repeat unit. In: Abad P, Bernell A, Laumond C, Boemare N, Coudert F (eds) COST 819 entomopathogenic nematodes—genetic and molecular biology of entomopathogenic nematodes. Luxembourg EUR 18261, Brussels, pp 87–93 177. Sneath PH, Sokal RR (1973) Numerical taxonomy. The principles and practice of numerical classification. CABI Publishing, Wallingford, UK 178. SAS Institute (1992) SAS/STAT software: changes and enhancements. SAS Institute, Cary, NC 179. Yamamoto S, Harayama S (1995) PCR amplification and direct sequencing of gyrB genes with universal primers and their application to the detection and taxonomic analysis of Pseudomonas putida strains. Appl Environ Microbiol 61(3):1104–1109

References 180. Wheeler WC, Gladstein DS (1994) MALIGN: a multiple sequence alignment program. J Hered 85(5):417–418 181. Nelson G, Platnick N (1981) Systematics and biogeography, cladistics and vicariance. Colombia University Press, New York 182. Tamura K (2007) MEGA4: molecular evolutionary genetics analysis (MEGA) software version 4.0. Mol Biol Evol 24:1596–1599 183. Brodsky LI (1992) GeneBee: the program package for biopolymer structure analysis. Dimacs 8:127–139 184. Brodsky LI (1995) GeneBee-NET: an internet-based server for analyzing biopolymers structure. Biochemist 60:1221–1230 185. Felsenstein J (1993) Phylogeny inference package. Department of Genetics, University of Washington, Seattle 186. Glazer I, Lewis EE (2000) Bioassays for entomopathogenic nematodes. In: Bioassays of entomopathogenic microbes and nematodes. CABI Publishing, Wallingford, UK, pp 229–247 187. Miller AD, Rosman GJ (1989) Improved retroviral vectors for gene transfer and expression. Biotechniques 7(9):980 188. Grewal D, Gotlieb J, Marmorstein H (1994) The moderating effects of message framing and source credibility on the price-perceived risk relationship. J Consum Res 21 (1):145–153 189. Morris DR (1995) Growth control of translation in mammalian cells. In: Progress in nucleic acid research and molecular biology, vol 51. Academic Press, London, pp 339–363 190. Somasekhar N, Grewal PS, De Nardo EA, Stinner BR (2002) Non-target effects of entomopathogenic nematodes on the soil nematode community. J Appl Ecol 39 (5):735–744 191. Shapiro-Ilan DI, Mbata GN, Nguyen KB, Peat SM, Blackburn D, Adams BJ (2009a) Characterization of biocontrol traits in the entomopathogenic nematode Heterorhabditis georgiana (Kesha strain), and phylogenetic analysis of the nematode’s symbiotic bacteria. Biol Control 51(3):377–387 192. Shapiro-Ilan DI, Reilly CC, Hotchkiss MW (2009b) Suppressive effects of metabolites from Photorhabdus and Xenorhabdus spp. on phytopathogens of peach and pecan. Arch. Phytopathol. Plant Prot 42:715–728 193. Shapiro SL, Schwartz GE, Bonner G (1998) Effects of mindfulness-based stress reduction on medical and premedical students. J Behav Med 21(6):581–599

171

194. Kaya HK, Koppenho¨fer AM (1996) Effects of microbial and other antagonistic organism and competition on entomopathogenic nematodes. Biocontrol Sci Tech 6 (3):357–372 195. Wharton R, Lewith G (1986) Complementary medicine and the general practitioner. Br Med J (Clin Res Ed) 292(6534):1498–1500 196. Koppenho¨fer AM, Kaya HK (1995) Densitydependent effects on Steinernema glaseri (Nematoda: Steinernematidae) within an insect host. J Parasitol 81(5):797–799 197. Wouts WM (1981) Mass production of the entomogenous nematode Heterorhabditisheliothidis (Nematode: Heterorhabditidae) on artificial media. J Nematol 13:467–469 198. Dye DW (1968) A taxonomic study of the genus Erwinia I. The ‘amylovora’ group. N Z J Sci 11:590–607 199. Strauch O, Ehlers RU (1998) Food signal production of Photorhabdus luminescens inducing the recovery of entomopathogenic nematodes Heterorhabditis spp. in liquid culture. Appl Microbiol Biotechnol 50 (3):369–374 200. Abbott WS (1925) A method of computing the effectiveness of an insecticide. J Econ Entomol 18:265–267 201. Akhurst RJ, Boemare NE (1988) A numerical taxonomic study of the genus Xenorhabdus (Enterobacteriaceae) and proposed elevation of the subspecies of X. nematophilus to species. J Gen Microbiol 134:1835–1845. Printed in Great Britain 202. Bergey’s Manual of Determinative Bacteriology (2000) Actinomycetales, 9th edn 203. Sambrook J, Fritschi EF, Maniatis T (1989) Molecular cloning: a laboratory manual. Cold Spring Harbor Laboratory Press, New York 204. Bauer AW, Kirby WMM, Sherris JC, Turck M (1966) Antibiotic susceptibility testing by a standardized single disk method. Am J Clin Pathol 45:493–496 205. Velikova N, Kavanagh K, Jerry M (2016) Evaluation of Galleria mellonella larvae for studying the virulence of Streptococcus suis. BMC Microbiol 16:291 206. Mahar AN, Munir M, Elawad S, Gowen SR, Hague NGM (2005) Pathogenicity of bacterium, Xenorhabdus nematophila isolated from entomopathogenic nematode (Steinernema carpocapsae) and its secretion against Galleria mellonella larvae. J Zhejiang Univ Sci B 6 (6):457–463 207. Lewis P, Abbeduto L, Murphy M, Richmond E, Giles N, Bruno L, Orsmond G (2006) Psychological well-being of mothers

172

References

of youth with fragile X syndrome: syndrome specificity and within-syndrome variability. J Intellect Disabil Res 50(12):894–904 208. Aliyu A, Kariim I, Abdulkareem SA (2017) Effects of aspect ratio of multi-walled carbon nanotubes on coal washery waste water treatment. J Environ Manag 202:84–93 209. Grewal SI, Elgin SC (2002) Heterochromatin: new possibilities for the inheritance of structure. Curr Opin Genet Dev 12 (2):178–187 210. Yukawa T, Pitt JM (1985) Nematode storage and transport. Int. Patent WO85/03412 211. Kaya HK, Nelson CE (1985) Encapsulation of steinernematid and heterorhabditid nematodes with calcium alginate: a new approach for insect control and other applications. Environ Entomol 14:572–574 212. Grewal PS, Peters A (2005) Formulation and quality. In: Nematodes as biocontrol agents. CABI Publishing, Wallingford, UK, pp 79–90 213. Bedding R (1990) Logistics and strategies for introducing entomopathogenic nematode technology into developing countries. In: Gaugler R, Kaya HK (eds) Entomopathogenic nematodes in biological control. CRC Press, Boca Raton, FL, pp 233–246 214. Capinera JL, Hibbard BE (1987) Bait formulations of chemical and microbial insecticides for suppression of crop-feeding grasshoppers. J Agric Entomol 4(4):337–340 215. Geoffrey J, Ilham S, Turlings T (2019) Encapsulated entomopathogenic nematodes can protect maize plants from Diabrotica balteata larvae. Insects 11:27 216. Georgis R (1990) Formulation and application technology. In: Gaugler R, Kaya HK (eds) Entomopathogenic nematodes in biological control. CRC Press, Boca Raton, FL, pp 173–194 217. Koppenho¨fer AM, Brown IM, Gaugler R, Grewal PS, Kaya HK, Klein MG (2000) Synergism of entomopathogenic nematodes and imidacloprid against white grubs: greenhouse and field evaluation. Biol Control 19 (3):245–251 218. Koppenhofer AM, Grewal PS (2005) 20 compatibility and interactions with agrochemicals and other biocontrol agents. In: Nematodes as biocontrol agents. CABI Publishing, Wallingford, UK, p 363 219. Alumai A, Grewal PS (2004) Tank-mix compatibility of the entomopathogenic nematodes, Heterorhabditis bacteriophora and Steinernema carpocapsae, with selected chemical pesticides used in turfgrass. Biocontrol Sci Technol 14(7):725–730

220. Nishimatsu T, Jackson JJ (1998) Interaction of insecticides, entomopathogenic nematodes, and larvae of the western corn root worm (Coleoptera: Chrysomelidae). J Econ Entomol 91(2):410–418 221. Anes KM, Ganguly S (2016) Pesticide compatibility with entomopathogenic nematode, Steinernema thermophilum (Nematoda: Rhabditida). Indian J Nematol 46(1):20–26 222. Shetlar DJ, Suleman PE, Georgis R (1988) Irrigation and use of entomogenous nematodes, Neoaplectana spp. and Heterorhabditis heliothidis (Rhabditida: Steinernematidae and Heterorhabditidae), for control of Japanese beetle (Coleoptera: Scarabaeidae) grubs in turfgrass. J Econ Entomol 81(5):1318–1322 223. Georgis JF, Chauzy S, Coquillat S (1995) Computed conditions of corona emission from two interacting raindrops. Q J R Meteorol Soc 121(528):1853–1866 224. Scher HI, Yagoda A, Herr HW, Sternberg CN, Bosl G, Morse MJ, Geller N (1988) Neoadjuvant M-VAC (methotrexate, vinblastine, doxorubicin and cisplatin) effect on the primary bladder lesion. J Urol 139 (3):470–474 225. McCoy CW, Shapiro DI, Duncan LW, Nguyen K (2000a) Entomopathogenic nematodes and other natural enemies as mortality factors for larvae of Diaprepes abbreviatus (Coleoptera: Curculionidae). Biol Control 19:182–190 226. Gouge DH, Reaves LL, Stoltman MM, Van Berkum JR, Burke RA, Jech LJ, Henneberry TJ (1996) Control of pink bollworm Pectinophora gossypiella (Saunders) larvae in Arizona and Texas cotton fields using the entomopathogenic nematode Steinernema riobravis (Cabanillas, Poinar, & Raulston) (Rhabditida: Steinernematidae). In: Richter DA, Armour J (eds) Proceedings of’ the Beltwide cotton production research conference. National Cotton Council of America, Memphis, pp 1078–1082 227. Wright RJ, Witkowski JF, Echtenkamp G, Georgis R (1993) Efficacy and persistence of Steinernema carpocapsae (Rhabditida: Steinernematidae) applied through a centerpivot irrigation system against larval corn rootworms (Coleoptera: Chrysomelidae). J Econ Entomol 86:1348–1354 228. Curran J (1992) Influence of application method and pest population size on weld efficacy of entomopathogenic nematodes. J Nematol 24:631–636 229. Reed DK, Reed GL, Creighton CS (1986) Introduction of entomogenous nematodes into trickle irrigation systems to control

References striped cucumber beetle, Acalymma vittatum (Coleoptera: Chrysomelidae). J Econ Entomol 79:1330–1333 230. McCoy CW, Shapiro DI, Duncan LW (2000b) Application and evaluation of entomopatho gens for citrus pest control. In: Lacey L, Kaya HK (eds) Manual of techniques in insect pathology: field techniques. Kluwer Academic Publishers, Dordrecht, Holland, pp 577–597 231. Kaya HK (1990) Soil ecology. In: Gaugler R, Kaya HK (eds) Entomopathogenic nematodes in biological control. CRC Press, Boca Raton, FL, pp 93–116 232. Klein MG, Georgis R (1992) Persistence of control of Japanese beetle (Coleoptera: Scarabaeidae) larvae with steinernematid and heterorhabditid nematodes. J Econ Entomol 85:727–730 233. Shapiro DI, Lewis EE, Paramasivam S, McCoy CW (2000) Nitrogen partitioning in Heterorhabditis bacteriophora-infected hosts and the eVects of nitrogen on attraction/ repulsion. J Invertebr Pathol 76:43–48 234. Shapiro-Ilan DI, Gouge DH, Koppenho¨fer AM (2002) Factors affecting commercial success: case studies in cotton, turf and citrus. In: Gaugler R (ed) Entomopathogenic nematology. CABI Publishing, Wallingford, UK, pp 333–356 235. Majic´ I, Sarajlic´ A, Lakatos T, To´th T, Raspudic´ E, Pusˇkadija Z, Kanizˇai SˇG, Laznik ˇ (2019) Virulence of new strain of HeteroZ rhabditis bacteriophora from Croatia against Lasioptera rubi. Plant Protect Sci 55:134–141 236. Yuksel E, Canhilal R (2018) Evaluation of local isolates of entomopathogenic nematodes for the management of black cutworm, Agrotis ipsilon Hufnagel (Lepidoptera: Noctuidae). Egypt J Biol Pest Control 28(1):82 237. Begley JW (1990) Efficacy against insects in habitats other than soil. In: Gaugler R, Kaya HK (eds) Entomopathogenic nematodes in biological control. CRC Press, Boca Raton, FL, pp 215–232 238. Georgis R, Koppenho¨fer AM, Lacey LA, Be´lair G, Duncan LW, Grewal PS, Van Tol RWHM (2006) Successes and failures in the use of parasitic nematodes for pest control. Biol Control 38(1):103–123 239. Lacey LA, Arthurs SP, Unruh TR, Headrick H, Fritts R Jr (2006a) Entomopathogenic nematodes for control of codling moth (Lepidoptera: Tortricidae) in apple and pear orchards: effect of nematode species and seasonal temperatures, adjuvants, application equipment and post-application irrigation. Biol Control 37:214–223

173

240. Lacey LA, Granatstein D, Arthurs SP, Headrick H, Fritts R Jr (2006b) Use of entomopathogenic nematodes (Steinernematidae) in conjunction with mulches for control of overwintering codling moth (Lepidoptera: Tortricidae). J Entomol Sci 41:107–119 241. Mracek Z (2002) Use of entomoparasitic nematodes (EPANs) in biological control. In: Advances in microbial control of insect pests. Springer, Boston, MA, pp 235–264 242. Kaya HK, Joos JL, Falcon LA, Berlowitz A (1984) Suppression of the codling moth (Lepidoptera, Olethreutidae) with the entomogenous nematode, Steinernema feltiae (Rhabditida, Steinernematidae). J Econ Entomol 77:1240–1244 243. Unruh TR, Lacey LA (2001) Control of codling moth, Cydia pomonella (Lepidoptera: Tortricidae), with Steinernema carpocapsae: effects of supplemental wetting and pupation site on infection rate. Biol Control 20 (1):48–56 244. Siegel J, Lacey LA, Fritts R Jr, Higbee BS, Noble P (2004) Use of steinernematid nematodes for post-harvest control of navel orangeworm (Lepidoptera: Pyralidae, Amyelois transitella) in fallen pistachios. Biol Control 30:410–417 245. Broadbent AB, Olthof THA (1995) Foliar application of Steinernema carpocapsae (Rhabditida: Steinernematidae) to control Liriomyza trifolii (Diptera: Agromyzidae) larvae in chrysanthemums. Environ Entomol 24:431–435 246. Jagdale GB, Casey ML, Grewal PS, Lindquist RK (2004) Effects of application rate and timing, potting medium and host plant on efficacy of Steinernema feltiae against fungus gnat, Bradysia coprophila in floriculture. Biol Control 29:296–305 247. Qiu BL, Mandour NS, Xu CX, Ren SX (2008) Evaluation of the entomopathogenic nematode Steinernema feltiae as a biological control agent of the whitefly, Bemisia tabaci. Int J Pest Manag 54(3):247–253 248. Trdan S, Znidarcic D, Vidrih M (2007) Control of Frankliniella occidentalis on glasshouse-grown cucumbers: an efficacy comparison of foliar application of Steinernema feltiae and spraying with abamectin. Russ J Nematol 15(1):25–34 249. Choi WI, Lee EH, Choi BR, Park HM, Ahn YJ (2003) Toxicity of plant essential oils to Trialeurodes vaporariorum (Homoptera: Aleyrodidae). J Econ Entomol 96:1479–1484

174

References

250. Hamdi F, Fargues J, Ridray G, Jeannequin B, Bonato O (2011) Compatibility among entomopathogenic hyphocreales and two beneficial insects used to control Trialeurodes vaporariorum (Hemiptera: Aleurodidae) in Mediterranean greenhouses. J Invertebr Pathol 108:22–29 251. Quesada-Moraga E, Maranhao EAA, Valverde-Garcıa P, Santiago-Alvarez C (2006) Selection of Beauveria bassiana isolates for control of the whiteflies Bemisia tabaci and Trialeurodes vaporariorum on the basis of their virulence, thermal requirements, and toxicogenic activity. Biol Control 36:274–287 252. Rezaei N, Karimi J, Hosseini M, Goldani M, Campos-Herrera R (2015) Pathogenicity of two species of entomopathogenic nematodes against the greenhouse whitefly, Trialeurodes vaporariorum (Hemiptera: Aleyrodidae), in laboratory and greenhouse experiments. J Nematol 47(1):60–66 253. Cuthbertson A, Walters K, Northing P, Luo W (2007) Efficacy of the entomopathogenic nematode, Steinernema feltiae, against sweetpotato whitefly Bemisia tabaci (Homoptera: Aleyrodidae) under laboratory and glasshouse conditions. Bull Entomol Res 97:9–14 254. Cuthbertson AG, Mathers JJ, Northing P, Prickett AJ, Walters KF (2008) The integrated use of chemical insecticides and the entomopathogenic nematode, Steinernema carpocapsae (Nematoda: Steinernematidae), for the control of sweetpotato whitefly, Bemisia tabaci (Hemiptera: Aleyrodidae). Insect Sci 15:447–453 255. Ansari NA, Rasheed Z (2009) Non-enzymatic glycation of proteins: from diabetes to cancer. Biochem (Mosc) Suppl Ser B Biomed Chem 3(4):335

256. Ansari MA, Butt TM (2011) Effects of successive subculturing on stability, virulence, conidial yield, germination and shelf-life of entomopathogenic fungi. J Appl Microbiol 110(6):1460–1469 257. Bruno TJ, Svoronos PD (2020) CRC handbook of basic tables for chemical analysis: data-driven methods and interpretation. CRC Press, Boca Raton, FL 258. Acharya S, Dogra K (2020) Understanding HANK: insights from a PRANK. Econometrica 88(3):1113–1158 259. Shehata IE, Hammam MMA, El-Borai FE, Duncan LW, Abd-Elgawad MMM (2020) Traits of the entomopathogenic nematode, Heterorhabditis bacteriophora (Hb-EG strain), for potential biocontrol in strawberry fields. Egypt J Biol Pest Control 30:40 260. Skowronek M, Sajnaga E, Pleszczyn´ska M, Kazimierczak W, Lis M, Wiater A (2020) Bacteria from the midgut of common cockchafer (Melolontha melolontha L.) larvae exhibiting antagonistic activity against bacterial symbionts of entomopathogenic nematodes: isolation and molecular identification. Int J Mol Sci 21(2):580 261. Dziedziech A, Shivankar S, Theopold U (2020) Drosophila melanogaster responses against entomopathogenic nematodes: focus on hemolymph clots. Insects 11(1):62 262. Kaplan SC (2020) Body mass index and social anxiety: effects of implicit weight bias and body salience in undergraduate women. Doctoral dissertation, Temple University. Libraries 263. Kaya HK (2002) Natural enemies and other antagonists. In: Gaugler R (ed) Entomopathogenic nematology. CABI Publishing, Wallingford, UK, pp 189–203

INDEX A

F

Abiotic ........................................................................., 161 Adsorbents.......................................................... , 147, 148 Aerosols ........................................................................... , 3 Ambushers ........................................................, 20, 28, 29 Analysis ......... , 1, 36, 68, 84, 86, 91, 95, 111, 131, 138, 139, 155 Antibiotics ................ , 9, 23, 25, 26, 118, 119, 140, 141 Antimycotic ............................................................ , 23, 25 Avoid....................., 2, 44, 50, 68, 71, 81, 138, 146, 151

Fertilizers ........................................................, 146, 150–1 Foragers .................................................................. , 28, 29 Formulation...................................................., 146–9, 161

G Galleria.................... , 9, 29, 36, 37, 47–50, 61, 114, 149 Glassware ....................................................................., 2, 3 Greenhouses ..................................................., 153, 155–8

H

B Behaviour............................................... , 27–30, 40–1, 98 Bioactive .............................. , 22, 25, 137, 139, 160, 161 Bioassay..................................., 36, 96, 99, 108, 111, 149 Biochemical .............................., 116, 122, 123, 130, 131 Biocontrol........ , 9, 19, 24, 68, 145, 153, 155, 160, 161 Biotic................................................................... , 151, 161

C

Habitats ..................., 5, 10, 55–7, 101, 153–8, 160, 161 Hazards................................................................... , 1, 161 Hermaphrodite.............................................................. , 21 Hosts......... , 5, 6, 8, 16, 19–21, 24, 27–9, 31, 36–9, 41, 43, 47–55, 57, 65, 76, 80, 96–8, 102–4, 111, 145, 149, 151–5, 160, 161 Hypoxia .................................................................. , 96, 99

I

Cadavers........................, 9, 20, 21, 32, 35, 36, 61–5, 73, 80, 86, 87, 96, 102, 105, 106, 113, 156–7 Carbon dioxide........................................ , 28, 33, 34, 124 Characterization ................ , 68, 86, 88, 96, 116–23, 131 Chemoreceptor ............................................................. , 24 Chemotaxis....................................................... , 32, 39–45 Commercially ............................. , 104, 130, 145, 149–50 Compatibility...................................................... , 150–151 Cruisers ................................................................... , 28, 29 Cryptic habitats ......................................................., 153–5 Cues ......................................................... , 27, 31–2, 35–8 Cuticle........................................... , 20, 41, 72–4, 79, 114

D Damaged ..................................................................., 2, 31 Desiccation ........................................................... , 96, 100 Dispose ............................................................................ , 3

E

Infective juveniles (IJs) ............. , 5, 61–6, 68, 71, 73, 76, 88, 90, 102, 115, 146, 148, 156–7 Insect carcass ................................................................. , 21 Insecticides ...................., 16, 19, 24, 146, 150, 151, 161 Isolation ...... , 15, 57, 59–68, 88, 105, 106, 113–7, 131, 132, 160

L Label .......................................................... , 3, 63, 68, 104 Lethal ................................................ , 15, 20, 24, 99, 145

M Metabolites ...................., 5, 20, 22–5, 31, 126, 159, 160 Microfluidics.............................................................., 43–5 Mutualistic ................................................................., 9, 19

N Nematicidal ......................................................, 20, 23, 25

Ecology ....................................... , 6, 31–45, 93, 145, 160 Emulsifiers ..................................................................., 147 Extinguishers ................................................................... , 2

O Odorants....................................... , 28, 32, 36, 39, 42, 43

S. Sivaramakrishnan and M. Razia, Entomopathogenic Nematodes and Their Symbiotic Bacteria, Springer Protocols Handbooks, https://doi.org/10.1007/978-1-0716-1445-7, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2021

175

ENTOMOPATHOGENIC NEMATODES

176 Index

AND

THEIR SYMBIOTIC BACTERIA

Olfactometer ............................................................., 38–9 Oviposition .............................................. , 36, 50, 55, 156

P Parasitic..........................................................., 20, 32, 149 Pathogenicity................................................, 64, 102, 139 Pathogens ........................................................................ , 3 Penetration ....................................... , 21, 22, 96, 97, 159 Pests ......... , 5, 6, 9, 24, 27, 28, 48, 50, 145–55, 159–61 Pheromones................................................................... , 31 Polymerase chain reaction (PCR) ......, 4, 86, 89–93, 137 Potential.............., 9, 24, 29, 62, 65, 102, 103, 160, 161

Q

S Safety..........................................................., 1–4, 145, 146 Secondary ........................................................., 20, 22, 95 Sedentary .........................................................., 28, 29, 51 Septicemia...................................................................... , 20 Spiracles ......................................................................... , 20 Storage ........................................................, 59–70, 147–9 Surfactants ..................................................................., 147 Symbionts ................, 5, 6, 15–8, 113, 119, 128–30, 160 Symbiosis ............................................., 15–7, 19, 21, 160 Symbiotic ........ , 5, 8, 9, 15, 17, 20, 22, 73, 76, 79, 105, 106, 113–43, 159–61 Systematics..........................................., 12–3, 86, 93, 131

Quality ................................................., 93, 104, 135, 149

T

R

Techniques.............................. , 47–111, 113–43, 145–52 Tritrophic..................................................................., 32–6

Randomly amplified polymorphic DNA (RAPD) ....................................................... , 91, 92 Rearing.............................................................. , 40, 47–55 Repellant........................................................................ , 44 Restriction fragment length polymorphism (RFLP) ........................................................ , 86, 91 Rhabditida .............................................................. , 12, 76

V Vermiculite ......................................................... , 147, 148 Virulence........................, 6, 96, 108, 109, 139, 146, 151 Volatiles....................................., 27, 28, 31, 35–8, 40, 41